*R represents any carbon-containing group. In a molecule with more than   one R group, the groups may be the same or different.    Cover the Structure column and draw the structure for each compound listed on the left. Do the same for each functional group  or linkage.   FIG. 1 -5 Replication through complementarity. In this simple case, a polymer  serves as a template for the assembly of a complementary molecule, which, because of  intramolecular complementarity, is an exact copy of the original.    illustrated in Fig. 1-5. A similar phenomenon is central to the function of  DNA, where the sequence of bases on one strand (e.g., A-C-G-T) absolutely  specifies the sequence of bases on the strand to which it is paired (T-G-C-A).  When DNA replicates, the two strands separate and direct the synthesis of  complementary daughter strands. Complementarity is also the basis for tran¬  scribing DNA into RNA and for translating RNA into protein.   A critical moment in chemical evolution was the transition from systems  of randomly generated molecules to systems in which molecules were organ¬  ized and specifically replicated. Once macromolecules gained the ability to  self-perpetuate, the primordial environment would have become enriched in  molecules that were best able to survive and multiply The first replicating  systems were no doubt somewhat sloppy, with progeny molecules imperfectly  complementary to their parents. Over time, natural selection would have  favored molecules that made more accurate copies of themselves.   • Explain why complementarity would have  been necessary for the development of  self-replicating molecules.    • Compartmentation of cells promotes efficiency by maintaining high local  concentrations of reactants.   • Eukaryotes are characterized by numerous membrane-bounded organelles,  including a nucleus.   • The phylogenetic tree of life includes three domains: bacteria, archaea, and eukarya.   • Evolution occurs as natural selection acts on randomly occurring variations among  individuals.   The types of systems described so far would have had to compete with all the  other components of the primordial earth for the available resources. A selec¬  tive advantage would have accrued to a system that was sequestered and pro¬  tected by boundaries of some sort. How these boundaries first arose, or even  what they were made from, is obscure. One theory is that membranous vesicles  (fluid-filled sacs) first attached to and then enclosed self-replicating systems.  These vesicles would have become the first cells.   The advantages of compartmentation are several. In addition to receiving  some protection from adverse environmental forces, an enclosed system can  maintain high local concentrations of components that would otherwise dif¬  fuse away. More concentrated substances can react more readily, leading to  increased efficiency in polymerization and other types of chemical reactions.   A membrane-bounded compartment that protected its contents would  gradually become quite different in composition from its surroundings.  Modern cells contain high concentrations of ions, small molecules, and large  molecular aggregates that are found in only traces—if at all—outside the cell.  For example, the Escherichia coli {E. colt) cell contains millions of molecules  representing some 3000 to 6000 different compounds (Fig. 1-6). A typical  animal cell may contain 100,000 different types of molecules.   Early cells depended on the environment to supply building materials. As  some of the essential components in the prebiotic soup became scarce, natu¬  ral selection favored organisms that developed mechanisms for synthesizing the  required compounds from simpler but more abundant precursors. The first  metabolic reactions may have used metal or clay catalysts (a catalyst is a sub¬  stance that promotes a chemical reaction without itself being changed). In fact,  metal ions are still at the heart of many chemical reactions in modern cells.  Some catalysts may also have arisen from polymeric molecules that had the  appropriate functional groups.   In general, biosynthetic reactions require energy; hence the first cellular  reactions also needed an energy source. The eventual depletion of preexisting  energy-rich substances in the prebiotic environment would have favored the  development of energy-producing metabolic pathways. For example, photo¬  synthesis evolved relatively early to take advantage of a practically inexhaustible  energy supply, the sun. However, the accumulation of 0 2 generated from H 2 0    FIG. 1 -6 Cross-section through an E. coli  cell. The cytoplasm is packed with  macromolecules. At this magnification  (~ 1,000,000x), individual atoms are too small to  resolve. The green structures on the right include  the inner and outer membrane components along  with a portion of a flagellum. Inside the cell,  various proteins are shown in blue, and  ribosomes are purple. The gold and orange  structures represent DNA and DNA-binding  proteins, respectively. In a living cell, the  remaining spaces would be crowded with water  and small molecules. [From Goodsell, D.S.,   The Machinery of Life (2nd ed.), Springer (2009).  Reproduced with permission.]   tional challenge to organisms adapted to life in an oxygen-poor atmosphere.   Metabolic refinements eventually permitted organisms not only to avoid ox¬  idative damage but to use 0 2 for oxidative metabolism, a much more efficient  form of energy metabolism than anaerobic metabolism. Vestiges of ancient life  can be seen in the anaerobic metabolism of certain modern organisms.   Early organisms that developed metabolic strategies to synthesize biological  molecules, conserve and utilize energy in a controlled fashion, and replicate within  a protective compartment were able to propagate in an ever-widening range of  habitats. Adaptation of cells to different external conditions ultimately led to  the present diversity of species. Specialization of individual cells also made it  possible for groups of differentiated cells to work together in multicellular  organisms.   All modern organisms are based on the same morphological unit, the cell.   There are two major classifications of cells: the eukaryotes (Greek: eu, good  or true + karyon, kernel or nut), which have a membrane-enclosed nucleus  encapsulating their DNA; and the prokaryotes (Greek: pro, before), which  lack a nucleus. Prokaryotes, comprising the various types of bacteria, have rela¬  tively simple structures and are almost all unicellular (although they may form  filaments or colonies of independent cells). Eukaryotes, which are multicellular  as well as unicellular, are vastly more complex than prokaryotes. (Viruses are  much simpler entities than cells and are not classified as living because they  lack the metabolic apparatus to reproduce outside their host cells.)   Prokaryotes are the most numerous and widespread organisms on the  earth. This is because their varied and often highly adaptable metabolisms suit  them to an enormous variety of habitats. Prokaryotes range in size from 1 to  10 fxm and have one of three basic shapes (Fig. 1-7): spheroidal (cocci), rod¬  like (bacilli), and helically coiled (spirilla). Except for an outer cell membrane,  which in most cases is surrounded by a protective cell wall, nearly all prokary¬  otes lack cellular membranes. However, the prokaryotic cytoplasm (cell con¬  tents) is by no means a homogeneous soup. Different metabolic functions are  believed to be carried out in different regions of the cytoplasm (Fig. 1-6). The  best characterized prokaryote is Escherichia coli, a 2 (Jim by 1 |Jim rodlike  bacterium that inhabits the mammalian colon.    FIG. 1 -8 Diagram of a typical animal cell with electron  micrographs of its organelles. Membrane-bounded organelles include  the nucleus, endoplasmic reticulum, lysosome, peroxisome (not pictured),  mitochondrion, vacuole, and Golgi apparatus. The nucleus contains  chromatin (a complex of DNA and protein) and the nucleolus (the site  of ribosome synthesis). The rough endoplasmic reticulum is studded with  ribosomes; the smooth endoplasmic reticulum is not. A pair of centrioles  help organize cytoskeletal elements. A typical plant cell differs mainly  by the presence of an outer cell wall and chloroplasts in the cytosol.   Inc./Phototake; Rough endoplasmic reticulum © Pietro M. Motta &  Tomonori Naguro/Photo Researchers, Inc.; Nucleus © Tektoff-RM,  CNRI/Photo Researchers; Mitochondrion © CNRI/Photo  Researchers; Golgi apparatus © Secchi-Lecaque/Roussel-UCLAF/  CNRI/Photo Researchers; Lysosome © Biophoto Associates/Photo  Researchers.]    Eukaryotic cells are generally 10 to 100 |xm in diameter and thus have a  thousand to a million times the volume of typical prokaryotes. It is not size,  however, but a profusion of membrane-enclosed organelles that best charac¬  terizes eukaryotic cells (Fig. 1-8). In addition to a nucleus, eukaryotes have  an endoplasmic reticulum, the site of synthesis of many cellular components,  some of which are subsequently modified in the Golgi apparatus. The bulk  of aerobic metabolism takes place in mitochondria in almost all eukaryotes,  and photosynthetic cells contain chloroplasts. Other organelles, such as lyso-  somes and peroxisomes, perform specialized functions. Vacuoles, which are  more prominent in plant than in animal cells, usually function as storage  depots. The cytosol (the cytoplasm minus its membrane-bounded organelles)  is organized by the cytoskeleton, an extensive array of filaments that also gives  the cell its shape and the ability to move.   The various organelles that compartmentalize eukaryotic cells represent a level  of complexity that is largely lacking in prokaryotic cells. Nevertheless, prokary¬  otes are more efficient than eukaryotes in many respects. Prokaryotes have  exploited the advantages of simplicity and miniaturization. Their rapid growth rates  permit them to occupy ecological niches in which there may be drastic fluctua¬  tions of the available nutrients. In contrast, the complexity of eukaryotes, which  renders them larger and more slowly growing than prokaryotes, gives them the  competitive advantage in stable environments with limited resources. It is there¬  fore erroneous to consider prokaryotes as evolutionarily primitive compared to  eukaryotes. Both types of organisms are well adapted to their respective lifestyles.   The practice of lumping all prokaryotes in a single category based on what  they lack—a nucleus—obscures their metabolic diversity and evolutionary  history. Conversely, the remarkable morphological diversity of eukaryotic  organisms (consider the anatomical differences among, say, an amoeba, an oak  tree, and a human being) masks their fundamental similarity at the cellular  level. Traditional taxonomic schemes (taxonomy is the science of biological  classification), which are based on gross morphology, have proved inadequate  to describe the actual relationships between organisms as revealed by their evo¬  lutionary history (phylogeny).   Biological classification schemes based on reproductive or developmental  strategies more accurately reflect evolutionary history than those based solely  on adult morphology. But phylogenetic relationships are best deduced by compar¬  ing polymeric molecules — RNA, DNA , or protein—-from dijferent organisms. For  example, analysis of RNA led Carl Woese to group all organisms into three  domains (Fig. 1-9). The archaea (also known as archaebacteria) are a group  of prokaryotes that are as distantly related to other prokaryotes (the bacteria,  sometimes called eubacteria) as both groups are to eukaryotes (eukarya). The  archaea include some unusual organisms: the methanogens (which produce  CH 4 ), the halobacteria (which thrive in concentrated brine solutions), and  certain thermophiles (which inhabit hot springs). The pattern of branches in  Woeses diagram indicates the divergence of different types of organisms (each  branch point represents a common ancestor). The three-domain scheme also  shows that animals, plants, and fungi constitute only a small portion of all  life-forms. Such phylogenetic trees supplement the fossil record, which pro¬  vides a patchy record of life prior to about 600 million years before the pres¬  ent (multicellular organisms arose about 700-900 million years ago).   It is unlikely that eukaryotes are descended from a single prokaryote,  because the differences between bacteria and eukaryotes are so profound.  Instead, eukaryotes probably evolved from the association of archaebacterial  and eubacterial cells. The eukaryotic genetic material includes features that  suggest an archaebacterial origin. In addition, the mitochondria and chloro-  plasts of modern eukaryotic cells resemble bacteria in size and shape, and both  types of organelles contain their own genetic material and protein synthetic  machinery. Evidently, as Lynn Margulis proposed, mitochondria and chloro-  plasts evolved from free-living bacteria that formed symbiotic (mutually ben¬  eficial) relationships with a primordial eukaryotic cell (Box 1-1). In fact,  certain eukaryotes that lack mitochondria or chloroplasts permanently harbor  symbiotic bacteria.    FIG. 1 -9 Phylogenetic tree showing three  domains of organisms. The branches indicate  the pattern of divergence from a common  ancestor. The archaea are prokaryotes, like  bacteria, but share some features with eukaryotes.  [After Wheelis, M.L, Kandler, 0., and Woese, C.R.,  Proc. Natl. Acad Sci. 89, 2931 (1992).]   Lynn Margulis (1938-) After growing up  in Chicago and enrolling in the University of  Chicago at age 16, Lynn Margulis intended  to be a writer. Her interest in biology was  sparked by a required science course for which  she read Gregor Mendel’s accounts  of his experiments with the genetics of pea  plants. Margulis continued her studies at the  University of Wisconsin Madison and at the  University of California Berkeley, earning a doctorate in 1963. Her  careful consideration of cellular structures led her to hypothesize  that eukaryotic cells originated from a series of endosymbiotic  events involving multiple prokaryotes. The term endo (Greek: within)  refers to an arrangement in which one cell comes to reside inside  another. This idea was considered outrageous at the time (1967),  but many of Margulis’s ideas have since become widely accepted.   Endosymbiosis as an explanation for the origin of mitochondria  had been proposed by Ivan Wallin in 1927, who noted the similarity  between mitochondria and bacteria in size, shape, and cytological  staining. Wallin’s hypothesis was rejected as being too fantastic  and was ignored until it was taken up again by Margulis. By  the 1960s, much more was known about mitochondria (and  chloroplasts), including the fact that they contained DNA and  reproduced by division. Margulis did not focus all her attention on  the origin of individual organelles but instead sought to explain the  origin of the entire eukaryotic cell, which also includes centrioles,  another possible bacterial relic. Her paper, “On the origin of  mitosing cells,” was initially rejected by several journals before  being accepted by the Journal of Theoretical Biology. The notion  that a complex eukaryotic cell could arise from a consortium of  mutually dependent prokaryotic cells was incompatible with the  prevailing view that evolution occurred as a series of small steps.  Evolutionary theory of the time had no room for the dramatic  amalgamation of cells-and their genetic material-that Margulis had  proposed. Nevertheless, the outspoken Margulis persisted, and by    the time she published Symbiosis in Cell Evolution in 1981, much  of the biological community had come on board.   Two main tenets of Margulis’s theory, that mitochondria are the  descendants of oxygen-respiring bacteria and chloroplasts were  originally photosynthetic bacteria, are almost universally accepted.  The idea that the eukaryotic cytoplasm is the remnant of an  archaebacterial cell is still questioned by some biologists. Margulis  is in the process of collecting evidence to support a fourth idea,  that cilia and flagella and some sensory structures such as the  light-sensing cells of the eye are descendants of free-living  spirochete bacteria. Margulis’s original prediction that organelles  such as mitochondria could be isolated and cultured has not been  fulfilled. However, there is ample evidence for the transfer of  genetic material between organelles and the nucleus, consistent  with Margulis’s theory of endosymbiosis. In fact, current theories  of evolution include the movement of genetic material among  organisms, as predicted by Margulis, in addition to small random  mutations as agents of change.   Perhaps as an extension of her work on bacterial  endosymbiosis, Margulis came to recognize that the interactions  among many different types of organisms as well as their  interactions with their physical environment constitute a single  self-regulating system. This notion is part of the Gaia hypothesis  proposed by James Lovelock, which views the entire earth as  one living entity (Gaia was a Greek earth goddess). However,  Margulis has no patience with those who seek to build a modern  mythology based on Gaia. She is adamant about the importance  of using scientific tools and reasoning to discover the truth and is  irritated by the popular belief that humans are the center of life  on earth. Margulis understands that human survival depends on  our relationships with waste-recycling, water-purifying, and oxygen-  producing bacteria, with whom we have been evolving, sometimes  endosymbiotically, for billions of years.    Sagan, L, On the origin of mitosing cells, J. Theor. Biol. 14 , 255-274 (1967).   The natural selection that guided prebiotic evolution continues to direct the  evolution of organisms. Richard Dawkins has likened evolution to a blind  watchmaker capable of producing intricacy by accident, although such an  image fails to convey the vast expanse of time and the incremental, trial-and-  error manner in which complex organisms emerge. Small mutations (changes  in an individuals genetic material) arise at random as the result of chemical  damage or inherent errors in the replication process. A mutation that increases  the chances of survival of the individual increases the likelihood that the muta¬  tion will be passed on to the next generation. Beneficial mutations tend to spread  rapidly through a population; deleterious changes tend to die along with the  organisms that harbor them.   The theory of evolution by natural selection, which was first articulated  by Charles Darwin in the 1860s, has been confirmed through observation and  experimentation. It is therefore useful to highlight several important—and often  misunderstood—principles of evolution:   1. Evolution is not directed toward a particular goal. It proceeds by  random changes that may affect the ability of an organism to reproduce    under the prevailing conditions. An organism that is well adapted to its  environment may fare better or worse when conditions change.   2 . Variation among individuals allows organisms to adapt to unexpected  changes. This is one reason why genetically homogeneous populations  (e.g., a corn crop) are so susceptible to a single challenge (e.g., a fun¬  gal blight). A more heterogeneous population is more likely to include  individuals that can resist the adversity and recover.   3. The past determines the future . New structures and metabolic func¬  tions emerge from preexisting elements. For example, insect wings did  not erupt spontaneously but appear to have developed gradually from  small heat-exchange structures.   4. Evolution is ongoing although it does not proceed exclusively toward  complexity. An anthropocentric view places human beings at the pin¬  nacle of an evolutionary scheme, but a quick survey of life’s diversity  reveals that simpler species have not died out or stopped evolving.    • Energy must be conserved, but it can take different forms.   • Entropy, a measure of a system’s disorder, tends to increase.   • The free energy change for a process is determined by the change in enthalpy  and the change in entropy.   • A spontaneous process occurs with a decrease in free energy.   • The free energy change for a reaction can be calculated from the temperature  and the concentrations and stoichiometry of the reactants and products.   • Biochemists define standard state conditions as a temperature of 25°C, a pres¬  sure of 1 atm, and a pH of 7.0.   • Organisms are nonequilibrium, open systems that constantly exchange matter and  energy with their surroundings.    The normal activities of living organisms—moving, growing, reproducing—  demand an almost constant input of energy. Even at rest, organisms devote a  considerable portion of their biochemical apparatus to the acquisition and uti¬  lization of energy. The study of energy and its effects on matter falls under the  purview of thermodynamics (Greek: therme, heat + dynamis , power). Although  living systems present some practical challenges to thermodynamic analysis, life  obeys the laws of thermodynamics. Understanding thermodynamics is important  not only for describing a particular process—such as a biochemical reaction—  in terms that can be quantified, but also for predicting whether that process can  actually occur, that is, whether the process is spontaneous. To begin, we will re¬  view the fundamental laws of thermodynamics. We will then turn our attention  to free energy and how it relates to chemical reactions. Finally, we will look at  how biological systems deal with the laws of thermodynamics.    A The First Law of Thermodynamics States That  Energy Is Conserved   In thermodynamics, a system is defined as the part of the universe that is of  interest, such as a reaction vessel or an organism; the rest of the universe is  known as the surroundings. The system has a certain amount of energy, U.  The first law of thermodynamics states that energy is conserved; it can be neither  created nor destroyed. However, when the system undergoes a change, some  of its energy can be used to perform work. The energy change of the system  is defined as the difference between the heat (q) absorbed by the system from   • Explain the selective advantages of com-  partmentation and metabolic pathways.   • Make a list of the major eukaryotic  organelles and their functions.   • Explain why a taxonomy based on molecu¬  lar sequences is more accurate than one  based on morphology.   • Which of the three domains are prokary¬  otic? Which domain is most similar to  eukaryotes?   • Why is evolutionary change constrained  by its past but impossible to predict?   Chapter 1 Introduction to the Chemistry of Life th e surroundings and work ( w ) done by the system on the surroundings. The   Heat is a reflection of random molecular motion, whereas work, which is de¬  fined as force times the distance moved under its influence, is associated with  organized motion. Force may assume many different forms, including the grav¬  itational force exerted by one mass on another, the expansional force exerted by  a gas, the tensional force exerted by a spring or muscle fiber, the electrical force  of one charge on another, and the dissipative forces of friction and viscosity.  Because energy can be used to perform different kinds of work, it is sometimes  useful to speak of energy taking different forms, such as mechanical energy, elec¬  trical energy, or chemical energy—all of which are relevant to biological systems.   Most biological processes take place at constant pressure. Under such con¬  ditions, the work done by the expansion of a gas (pressure—volume work) is  PA V Consequently, it is useful to define a new thermodynamic quantity, the  enthalpy (Greek: enthalpein , to warm in), abbreviated Hi   AH = AU + PAV= q P - w+ PAV [1-3]   where q P is defined as the heat at constant pressure. Since we already know  that in this system w — PAV,   In other words, the change in enthalpy is equivalent to heat. Moreover, the  volume changes in most biochemical reactions are insignificant {PAV ~ 0),  so the differences between their AU and AH values are negligible, and hence  the energy change for the reacting system is equivalent to its enthalpy change.  Enthalpy, like energy, heat, and work, is given units of joules. (Some com¬  monly used units and biochemical constants and other conventions are given  in Box 1-2.)    Modern biochemistry generally uses Systeme International (SI) units,  including meters (m), kilograms (kg), and seconds (s) and their  derived units, for various thermodynamic and other measurements.  The following lists the commonly used biochemical units, some  useful biochemical constants, and a few conversion factors.    Throughout this text, molecular masses of particles are expressed  in units of daltons (D), which are defined as 1/12th the mass of  a 12 C atom (1000 D = 1 kilodalton, kD). Biochemists also use  molecular weight, a dimensionless quantity defined as the ratio  of the particle mass to 1/12th the mass of a 12 C atom, which is  symbolized M r (for relative molecular mass).   Thermodynamics is useful for indicating the spontaneity of a process. A spon- Section 3 Thermodynamics   taneous process occurs without the input of additional energy from outside the  system. (Thermodynamic spontaneity has nothing to do with how quickly a  process occurs.) The first law of thermodynamics, however, cannot by itself deter¬  mine whether a process is spontaneous. Consider two objects of different temper¬  atures that are brought together. Heat spontaneously flows from the warmer object  to the cooler one, never vice versa. Yet either process would be consistent with the  first law of thermodynamics since the aggregate energy of the two objects does not  change. Therefore, an additional criterion of spontaneity is needed.    B The Second Law of Thermodynamics States  That Entropy Tends to Increase    According to the second law of thermodynamics, spontaneous processes are charac¬  terized by the conversion of order to disorder. In this context, disorder is defined  as the number of energetically equivalent ways, W, of arranging the compo¬  nents of a system. To make this concept concrete, consider a system consist¬  ing of two bulbs of equal volume, one of which contains molecules of an ideal  gas (Fig. 1-10). When the stopcock connecting the bulbs is open, the mole¬  cules become randomly but equally distributed between the two bulbs. The  equal number of gas molecules in each bulb is not the result of any law of  motion; it is because the probabilities of all other distributions of the mole¬  cules are so overwhelmingly small. Thus, the probability of all the molecules  in the system spontaneously rushing into the left bulb (the initial condition)  is nil, even though the energy and enthalpy of that arrangement are exactly  the same as those of the evenly distributed molecules.   The degree of randomness of a system is indicated by its entropy (Greek:  en, in + trope , turning), abbreviated S:   where is the Boltzmann constant. The units of S are J • K 1 (absolute  temperature, in units of kelvins, is a factor because entropy varies with tem¬  perature; e.g., a system becomes more disordered as its temperature rises). The  most probable arrangement of a system is the one that maximizes W and hence  S. Thus, if a spontaneous process, such as the one shown in Fig. 1-10, has  overall energy and enthalpy changes (A U and A H) of zero, its entropy change  (A S) must be greater than zero; that is, the number of equivalent ways of ar¬  ranging the final state must be greater than the number of ways of arranging  the initial state. Furthermore, because    FIG. 1-10 Illustration of entropy. In (a), a  gas occupies the leftmost of two equal-sized  bulbs and hence the entropy is low. When the  stopcock is opened ( b ), the entropy increases as  the gas molecules diffuse back and forth  between the bulbs and eventually become  distributed evenly, half in each bulb.   In chemical and biological systems, it is impractical, if not impossible, to  determine the entropy of a system by counting all the equivalent arrangements  of its components ( W). However, there is an entirely equivalent expression for  entropy that applies to the constant-temperature conditions typical of biolog¬  ical systems: for a spontaneous process,   Thus, the entropy change in a process can be experimentally determined from  measurements of heat.    C The Free Energy Change Determines the Spontaneity of a Process   The spontaneity of a process cannot be predicted from a knowledge of the  systems entropy change alone. For example, 2 mol of H 2 and 1 mol of 0 2 ,  when sparked, react to form 2 mol of H 2 0. Yet two water molecules, each of  whose three atoms are constrained to stay together, are more ordered than are    the three diatomic molecules from which they formed. Thus, the reaction oc¬  curs with a decrease in the systems entropy.   What, then, is the thermodynamic criterion for a spontaneous process?  Equations 1-4 and 1-7 indicate that at constant temperature and pressure    This is the true criterion for spontaneity as formulated, in 1878, by J. Willard Gibbs.  He defined the Gibbs free energy (G, usually called just free energy) as    The change in free energy for a process is AG. Consequently, spontaneous  processes at constant temperature and pressure have    Such processes are said to be exergonic (Greek: ergon , work). Processes that  are not spontaneous have positive AG values (AG > 0) and are said to be  endergonic; they must be driven by the input of free energy. If a process is  exergonic, the reverse of that process is endergonic and vice versa. Thus, the  AG value for a process indicates whether the process can occur spontaneously  in the direction written (see Sample Calculation 1-1). Processes at equilibrium,  those in which the forward and reverse reactions are exactly balanced, are char¬  acterized by AG = 0. For the most part, only changes in free energy, enthalpy,  and entropy (AG, AH. , and AS) can be measured, not their absolute values  (G, H, and S).   A process that is accompanied by an increase in enthalpy (A H> 0), which  opposes the process, can nevertheless proceed spontaneously if the entropy  change is sufficiently positive (AS > 0; Table 1-4). Conversely, a process that  is accompanied by a decrease in entropy (AS < 0) can proceed if its enthalpy  change is sufficiently negative (AH < 0). It is important to emphasize that a  large negative value of A G does not ensure that a process such as a chemical reac¬  tion will proceed at a measurable rate. The rate depends on the detailed mechanism  of the reaction, which is independent of AG.   Free energy as well as energy, enthalpy, and entropy are state functions.  In other words, their values depend only on the current state or properties of  the system, not on how the system reached that state. Therefore, thermody¬  namic measurements can be made by considering only the initial and final states   Variation of Reaction Spontaneity (Sign of AG) with the Signs  of AH and AS   The reaction is both enthalpically favored (exothermic) and  entropically favored. It is spontaneous (exergonic) at all  temperatures.   The reaction is enthalpically opposed (endothermic) but  entropically favored. It is spontaneous only at temperatures  above T = AH/AS.   The reaction is both enthalpically and entropically opposed. It is  nonspontaneous (endergonic) at all temperatures.    The enthalpy and entropy of the initial and final states  shown in the table.    a. Calculate the change in enthalpy and change in entropy for the reaction.   b. Calculate the change in free energy for the reaction when the temperature is 4°C.  Is the reaction spontaneous?   AS = S final - S initial = AS = 43 J • K" 1 • moP 1 - 22 J-KP 1 • mol" 1  = 21 J • K 1 • mol -1   b. First, convert temperature from °C to K: 4 + 273 = 277 K. Then use Eq. 1-11.   A G = (6000 J • mol -1 ) - (277 K)(21 J • K -1 • mol -1 )   The value for AC is greater than zero, so this is an endergonic (nonspontaneous)  reaction at 4°C.   AC = (6000 J • mol -1 ) - (310 K)(21 J • K -1 • mol -1 )   The value for AC is less than zero, so the reaction is spontaneous (exergonic)  at 37°C.    of the system and ignoring all the stepwise changes in enthalpy and entropy that oc¬  cur in between. For example, it is impossible to directly measure the energy change  for the reaction of glucose with 0 2 in a living organism because of the numer¬  ous other simultaneously occurring chemical reactions. But since A G depends on  only the initial and final states, the combustion of glucose can be analyzed in  any convenient apparatus, using the same starting materials (glucose and 0 2 )  and end products (C0 2 and H 2 0) that would be obtained in vivo.   The entropy (disorder) of a substance increases with its volume. For example,  a collection of gas molecules, in occupying all of the volume available to it,  maximizes its entropy. Similarly, dissolved molecules become uniformly dis¬  tributed throughout their solution volume. Entropy is therefore a function of  concentration.   If entropy varies with concentration, so must free energy. Thus, the free  energy change of a chemical reaction depends on the concentrations of both its  reacting substances ( reactants) and its reaction products. This phenomenon has  great significance because many biochemical reactions operate spontaneously  in either direction depending on the relative concentrations of their reactants  and products.    The standard free energy change for the re¬  action A —> B is — 15 kj • mol 1 . What is  the equilibrium constant for the reaction?   Since A G° is known, Eq. 1-17 can be used  to calculate A^ q . Assume the temperature is  25°C (298 K):    Equilibrium Constants Are Related to AG. The relationship between the  concentration and the free energy of a substance A is approximately   where G A is known as the partial molar free energy m* the chemical poten¬  tial of A (the bar indicates the quantity per mole), G A is the partial molar  free energy of A in its standard state, R is the gas constant, and [A] is the  molar concentration of A. Thus, for the general reaction   AG° = cG° c + dG& - aG° A - bGl [1-14]    because free energies are additive and the free energy change of a reaction is  the sum of the free energies of the products less those of the reactants.  Substituting these relationships into Eq. 1-12 yields    where AG° is the free energy change of the reaction when all of its reactants  and products are in their standard states (see below). Thus, the expression for  the free energy change of a reaction consists of two parts: (1) a constant term  whose value depends only on the reaction taking place and (2) a variable term  that depends on the concentrations of the reactants and the products, the stoi¬  chiometry of the reaction, and the temperature.   For a reaction at equilibrium, there is no net change because the free en¬  ergy change of the forward reaction exactly balances that of the reverse reac¬  tion. Consequently, AG = 0, so Eq. 1-15 becomes    The subscript “eq” denotes reactant and product concentrations at equilib¬  rium. (The equilibrium condition is usually clear from the context of the sit¬  uation, so equilibrium concentrations are usually expressed without this  subscript.) The equilibrium constant of a reaction can therefore be calculated from  standard free energy data and vice versa (see Sample Calculation 1-2). The actual  free energy change for a reaction can be calculated from the standard free  energy change (A G°') and the actual concentrations of the reactants and prod¬  ucts (see Sample Calculation 1-3).    Using the data provided in Sample Calculation 1-2, what is the actual free energy  change for the reaction A —> B at 37°C when [A] = 10 mM and [B] =0.1 mM?   A G= -15,000 J-mol -1 + (8.314 J• mol -1 • K _1 )(37 + 273 K) ln(0.1/10)  = -15,000 J-moF 1 - 11,900 J• mol -1  = -26,900 J-moF 1    Equations 1-15 through 1-17 indicate that when the reactants in a process  are in excess of their equilibrium concentrations, the net reaction will proceed  in the forward direction until the excess reactants have been converted to prod¬  ucts and equilibrium is attained. Conversely, when products are in excess, the  net reaction proceeds in the reverse direction. Thus, as Le Chatelier’s principle  states, any deviation from equilibrium stimulates a process that tends to restore  the system to equilibrium. In cells, many metabolic reactions are freely re¬  versible, and the direction of the reaction can shift as reactants and products  are added to or removed from the cell. Some metabolic reactions, however,  proceed in only one direction, so the cell can therefore maintain reactant and  product concentrations far from their equilibrium values.    K Depends on Temperature. The manner in which the equilibrium constant  varies with temperature can be seen by substituting Eq. 1-11 into Eq. 1-16  and rearranging:    where H° and 5° represent enthalpy and entropy in the standard state. Equation  1-18 has the form y = mx + b , the equation for a straight line. A plot of In  K eq versus \IT, known as a van’t Hoff plot, permits the values of A H° and A 5°  (and hence A G°) to be determined from measurements of IQ q at two (or more)  different temperatures. This method is often more practical than directly meas¬  uring AH and A S by calorimetry (which measures the heat, q P , of a process).    Biochemists Have Defined Standard-State Conventions. In order to com¬  pare free energy changes for different reactions, it is necessary to express AG  values relative to some standard state (likewise, we refer the elevations of ge¬  ographic locations to sea level, which is arbitrarily assigned the height of zero).  According to the convention used in physical chemistry, a solute is in its stan¬  dard state when the temperature is 25°C, the pressure is 1 atm, and the solute  has an activity of 1 (activity of a substance is its concentration corrected for  its nonideal behavior at concentrations higher than infinite dilution).   The concentrations of reactants and products in most biochemical reac¬  tions are usually so low (on the order of millimolar or less) that their activi¬  ties are closely approximated by their molar concentrations. Furthermore,  because biochemical reactions occur near neutral pH, biochemists have  adopted a somewhat different standard-state convention:   1. The activity of pure water is assigned a value of 1, even though its con¬  centration is 55.5 M. This practice simplifies the free energy expres¬  sions for reactions in dilute solutions involving water as a reactant,  because the [H 2 0] term can then be ignored.   2. The hydrogen ion (H + ) activity is assigned a value of 1 at the physi¬  ologically relevant pH of 7. Thus, the biochemical standard state is  pH 7.0 (neutral pH, where [H + ] = 10 1 2 3 * * * 7 M) rather than pH 0  ([H + ] = 1 M), the physical chemical standard state, where many bi¬  ological substances are unstable.   3. The standard state of a substance that can undergo an acid-base reac¬   tion is defined in terms of the total concentration of its naturally oc¬   curring ion mixture at pH 7. In contrast, the physical chemistry  convention refers to a pure species whether or not it actually exists at   pH 0. The advantage of the biochemistry convention is that the total  concentration of a substance with multiple ionization states, such as  most biological molecules, is usually easier to measure than the concen¬  tration of one of its ionic species. Since the ionic composition of an  acid or base varies with pH, however, the standard free energies calcu¬   lated according to the biochemical convention are valid only at pH 7.   Chapter 1 Introduction to the Chemistry of Life Under the biochemistry convention, the standard free energy changes of reac¬  tions are customarily symbolized by A G°' to distinguish them from physical  chemistry standard free energy changes, AG°. If a reaction includes neither  H 2 0, H + , nor an ionizable species, then AG°' = AG°.   At one time, many scientists believed that life, with its inherent complexity and  order, somehow evaded the laws of thermodynamics. However, elaborate meas¬  urements on living animals are consistent with the conservation of energy pre¬  dicted by the first law. Unfortunately, experimental verification of the second  law is not practicable, since it requires dismantling an organism to its compo¬  nent molecules, which would result in its irreversible death. Consequently, it is  possible to assert only that the entropy of living matter is less than that of the  products to which it decays. Life persists, however, because a system (a living or¬  ganism) can be ordered at the expense of disordering its surroundings to an even  greater extent. In other words, the total entropy of the system plus its surround¬  ings increases, as required by the second law. Living organisms achieve order  by disordering (breaking down) the nutrients they consume. Thus, the entropy  content of food is as important as its energy content.   Living Organisms Are Open Systems. Classical thermodynamics applies pri¬  marily to reversible processes in isolated systems (which cannot exchange mat¬  ter or energy with their surroundings) or in closed systems (which can only  exchange energy). An isolated system inevitably reaches equilibrium. For exam¬  ple, if its reactants are in excess, the forward reaction will proceed faster than  the reverse reaction until equilibrium is attained (AG = 0), at which point the  forward and reverse reactions exactly balance each other. In contrast, open sys¬  tems, which exchange both matter and energy with their surroundings, can  reach equilibrium only after the flow of matter and energy has stopped.   Living organisms, which take up nutrients, release waste products, and gener¬  ate work and heat, are open systems and therefore can never be at equilibrium. They  continuously ingest high-enthalpy, low-entropy nutrients, which they convert to  low-enthalpy, high-entropy waste products. The free energy released in this  process powers the cellular activities that produce the high degree of organiza¬  tion characteristic of life. If this process is interrupted, the system ultimately  reaches equilibrium, which for living things is synonymous with death. An ex¬  ample of energy flow in an open system is illustrated in Fig. 1-11. Through  photosynthesis, plants convert radiant energy from the sun, the primary energy   Plants use the sun’s radiant energy to synthesize  carbohydrates from C0 2 and H 2 0. Plants or the  animals that eat them eventually metabolize the  carbohydrates to release their stored free energy  and thereby return C0 2 and H 2 0 to the  environment.    source for life on the earth, to the chemical energy of carbohydrates and other  organic substances. The plants, or the animals that eat them, then metabolize  these substances to power such functions as the synthesis of biomolecules, the  maintenance of intracellular ion concentrations, and cellular movements.   Living Things Maintain a Steady State. Even in a system that is not at equi¬  librium, matter and energy flow according to the laws of thermodynamics. For  example, materials tend to move from areas of high concentration to areas of  low concentration. This is why blood takes up 0 2 in the lungs, where 0 2 is  abundant, and releases it to the tissues, where 0 2 is scarce.   Living systems are characterized by being in a steady state. This means  that all flows in the system are constant so that the system does not change  with time. Energy flow in the biosphere (Fig. 1-11) is an example of a system  in a steady state. Slight perturbations from the steady state give rise to changes  in flows that restore the system to the steady state. In all living systems, en¬  ergy flow is exclusively “downhill” (A G < 0). In addition, nature is inherently  dissipative, so the recovery of free energy from a biochemical process is never  total and some energy is always lost to the surroundings.   Enzymes Catalyze Biochemical Reactions. Nearly all the molecular compo¬  nents of an organism can potentially react with each other, and many of these  reactions are thermodynamically favored (spontaneous). Yet only a subset of  all possible reactions actually occur to a significant extent in a living organism.  The rate of a particular reaction depends not on the free energy difference  between the initial and final states but on the actual path through which the  reactants are transformed to products. Living organisms take advantage of cat¬  alysts, substances that increase the rate at which the reaction approaches equi¬  librium without affecting the reactions AG. Biological catalysts are referred to  as enzymes, most of which are proteins.   Enzymes accelerate biochemical reactions by physically interacting with the  reactants and products to provide a more favorable pathway for the transformation  of one to the other. Enzymes increase the rates of reactions by increasing the  likelihood that the reactants can interact productively. Enzymes cannot, how¬  ever, promote reactions whose A G values are positive.   A multitude of enzymes mediate the flow of energy in every cell. As free  energy is harvested, stored, or used to perform cellular work, it may be trans¬  ferred to other molecules. And although it is tempting to think of free energy  as something that is stored in chemical bonds, chemical energy can be trans¬  formed into heat, electrical work, or mechanical work, according to the needs  of the organism and the biochemical machinery with which it has been  equipped through evolution.   • Summarize the relationship between  energy (I J ), heat (g), and work (w).   • Use the analogy of a china cabinet to  describe a system with low entropy or  high entropy.   • Explain why changes in both enthalpy  (A H) and entropy (AS) determine the  spontaneity of a process.   • What is the relationship between the rate  of a process and its thermodynamic  spontaneity?   • What is the free energy change for a  reaction at equilibrium?   • Write the equation showing the relation¬  ship between AG° and K eq .   • Write the equation showing the relation¬  ship between AG, AG°, and the concen¬  trations of the reactants and products.   • Explain how biochemists define the  standard state of a solute. Why do  biochemists and chemists use different  conventions?   • Explain how organisms avoid reaching  equilibrium while maintaining a steady  state.   • How do enzymes affect the rate and free  energy change of a reaction?   • A model for the origin of life proposes that organisms ultimately  arose from simple organic molecules that polymerized to form  more complex molecules capable of replicating themselves.   • Compartmentation gave rise to cells that developed metabolic re¬  actions for synthesizing biological molecules and generating energy.   • All cells are either prokaryotic or eukaryotic. Eukaryotic cells con¬  tain a variety of membrane-bounded organelles.   • Phylogenetic evidence groups organisms into three domains:  archaea, bacteria, and eukarya.   • The first law of thermodynamics (energy is conserved) and the  second law (spontaneous processes increase the disorder of the uni¬  verse) apply to biochemical processes. The spontaneity of a process  is determined by its free energy change (AG = A H — TAS):  spontaneous reactions have A G < 0 and nonspontaneous reactions  have AG > 0.   • The equilibrium constant for a process is related to the standard  free energy change for that process.   • Living organisms are open systems that maintain a steady state.   endoplasmic reticulum 8  Golgi apparatus 8  mitochondrion 8  chloroplast 8  lysosome 8  peroxisome 8  vacuole 8  cytosol 8  cytoskeleton 8  taxonomy 9  phylogeny 9  archaea 9  bacteria 9  eukarya 9  methanogens 9  halobacteria 9   standard state 1 6  equilibrium constant 1 6  Le Chatelier’s principle 1 7  van’t Hoff plot 1 7  activity 17  isolated system 1 8  closed system 1 8  open system 1 8  steady state 1 9  enzyme 19    1. Identify the circled functional groups and linkages in the com¬  pound below.    2. Why is the cell membrane not an absolute barrier between the  cytoplasm and the external environment?   3. A spheroidal bacterium with a diameter of 1 p>m contains two  molecules of a particular protein. What is the molar concentra¬  tion of the protein?   4. How many glucose molecules does the cell in Problem 3 contain  when its internal glucose concentration is 1.0 mM?   3. (a) Which has greater entropy, liquid water at 0°C or ice at 0°C?   (b) How does the entropy of ice at — 3°C differ, if at all, from  its entropy at — 50°C?   7. Consider a reaction with A H = 15 kj and A S = 50 J • K \ Is  the reaction spontaneous (a) at 10°C, (b) at 80°C?   8. For the reaction A —> B at 298 K, the change in enthalpy is  —7 kj • mol 1 and the change in entropy is — 25 J * K 1 • mol \  Is the reaction spontaneous? If not, should the temperature be  increased or decreased to make the reaction spontaneous?   9. For the conversion of reactant A to product B, the change in en¬  thalpy is 7 kj • mol 1 and the change is entropy is 20 J • K 1 •  mol 1 . Above what temperature does the reaction become  spontaneous?   (a) A reaction is said to be spontaneous when it can proceed in  either the forward or reverse direction.   (c) A nonspontaneous reaction will proceed spontaneously in the  reverse direction.   (d) A spontaneous process can occur with a large decrease in  entropy.    11. When the reaction A + B v C is at equilibrium, the concen¬  trations of reactants are as follows: [A] = 2 mM, [B] = 3 mM,  and [C] = 9 mM. What is the standard free energy change for  the reaction?   12 . Calculate AG 0 ' for the reaction A + B v C + D at 23 °C  when the equilibrium concentrations are [A] = 10 pM, [B] = 13  pM, [C] = 3 pM, and [D] = 3 pM. Is the reaction exergonic  or endergonic under standard conditions?   glucose- 1 -phosphate + H 2 0 —> glucose + H 2 PO 4  at pH 7.0 and 25°C (A G°' = —20.9 kj • mol l ).   16. At 10°C, K eq for a reaction is 100. At 30°C, K eq = 10. Does  enthalpy increase or decrease during the reaction?   17. Two biochemical reactions have the same LQ q = 5 X 10 8 at tem¬  perature 7"i = 298 K. However, Reaction 1 has A H° = — 28 kj  • mol 1 and Reaction 2 has A H° = + 28 kj • mol \ The two  reactions utilize the same reactants. Your lab partner has proposed  that you can get more of the reactants to proceed via Reaction 2  rather than Reaction 1 by lowering the temperature of the reac¬  tion. Will this strategy work? Why or why not? How much would  the temperature have to be raised or lowered in order to change  the value of K 2 /K x from 1 to 10 ?   is —7.1 kj • mol \ Calculate the equilibrium ratio of [G 1 P]  to [G 6 P] at 25°C.   (a) If 50 pM of Q is mixed with 50 pM of R, which way will  the reaction proceed: to generate more Q or more R?    Look up methanogen and  methanotroph. Where do such organisms occur? Summarize how  they obtain matter and energy from their surroundings. Draw a dia¬  gram to illustrate the metabolic interdependence of methanogens  and methanotrophs.   Anet, F.A.L., The place of metabolism in the origin of life, Curr. Opin.  Chem. Biol. 8, 654-659 (2004). [Discusses various hypotheses pro¬  posing that life originated as a self-replicating system or as a set of  catalytic polymers.]   Bada, J.L. and Lazcano, A., Prebiotic soup—revisiting the Miller experi¬  ment, Science 300, 745-756 (2003).   McNichol, J., Primordial soup, fool’s gold, and spontaneous generation,  Biochem. Mol. Biol. Ed. 36, 255-261 (2008). [A brief introduction to  the theory, history, and philosophy of the search for the origin of life.]   Nisbet, E.G. and Sleep, N.H., The habitat and nature of early life, Nature  409, 1083-1091 (2001). [Explains some of the hypotheses regarding  the early earth and the origin of life, including the possibility that life  originated at hydrothermal vents.]   Campbell, N.A. and Reece, J.B., Biology ( 8 th ed.), Benjamin/Cummings  (2008). [This and other comprehensive general biology texts provide  details about the structures of prokaryotes and eukaryotes.]    DeLong, E.E and Pace, N.R., Environmental diversity of bacteria and  archaea, Syst. Biol. 593, 470-478 (2001). [Describes some of the  challenges of classifying microbial organisms among the three  domains.]   Lodish, H., Berk, A., Kaiser, C.A., Krieger, M., Scott, M.P., Bretscher A.,  Ploegh, H., and Matsudaria, P., Molecular Cell Biology ( 6 th ed.),  Chapter 5, W.H. Freeman (2008). [This and other cell biology text¬  books offer thorough reviews of cellular structure.]   Tinoco, I., Jr., Sauer, K., Wang, J.C., and Puglisi, J.C., Physical Chemistry.  Principles and Applications in Biological Sciences (4th ed.), Chapters 2-5,  Prentice-Hall ( 2002 ). [Most physical chemistry texts treat thermo¬  dynamics in some detail.]   van Holde, K.E., Johnson, W.C., and Ho, P.S., Principles of Physical  Biochemistry (2nd ed.), Chapters 2 and 3, Prentice-Hall (2006).   A Water Is a Polar Molecule  B Hydrophilic Substances Dissolve in Water  C The Hydrophobic Effect Causes Nonpolar  Substances to Aggregate in Water  D Water Moves by Osmosis and Solutes  Move by Diffusion   A Water Ionizes to Form H + and OH -  B Acids and Bases Alter the pH  C Buffers Resist Changes in pH   The splash caused by a droplet falling onto a water surface is a manifestation of the  internal cohesion of water due mainly to hydrogen bonding between water molecules.  Such weak bonds also play a central role in the structures and functions of biological  molecules. [StudioSource/Alamy]    Any study of the chemistry of life must include a study of water. Biological  molecules and the reactions they undergo can be best understood in the con¬  text of their aqueous environment. Not only are organisms made mostly of  water (about 70% of the mass of the human body is water), they are sur¬  rounded by water on this, the “blue planet.” Aside from its sheer abundance,  water is central to biochemistry for the following reasons:   1. Nearly all biological molecules assume their shapes (and therefore their  functions) in response to the physical and chemical properties of the  surrounding water.   2. The medium for the majority of biochemical reactions is water.  Reactants and products of metabolic reactions, nutrients as well as waste  products, depend on water for transport within and between cells.   3. Water itself actively participates in many chemical reactions that sup¬  port life. Frequently, the ionic components of water, the H + and OH  ions, are the true reactants. In fact, the reactivity of many functional  groups on biological molecules depends on the relative concentrations  of H + and OH in the surrounding medium.   All organisms require water, from the marine creatures who spend their  entire lives in an aqueous environment to terrestrial organisms who must guard  their watery interiors with protective skins. Not surprisingly, living organisms  can be found wherever there is liquid water—in hydrothermal vents as hot as  121°C and in the cracks and crevices between rocks hundreds of meters beneath  the surface of the earth. Organisms that survive desiccation do so only by  becoming dormant, as seeds or spores.   An examination of water from a biochemical point of view requires a look  at the physical properties of water, its powers as a solvent, and its chemical  behavior—that is, the nature of aqueous acids and bases.   • Water molecules, which are polar, can form hydrogen bonds with other molecules.   • In ice, water molecules are hydrogen bonded in a crystalline array, but in liquid  water, hydrogen bonds rapidly break and re-form in irregular networks.   • The attractive forces acting on biological molecules include ionic interactions,  hydrogen bonds, and van der Waals interactions.   • The hydrophobic effect explains the exclusion of nonpolar groups as a way to  maximize the entropy of water molecules.   • Amphiphilic substances form micelles or bilayers that hide their hydrophobic  groups while exposing their hydrophilic groups to water.   • Molecules diffuse across membranes which are permeable to them from regions  of higher concentration to regions of lower concentration.   • In dialysis, solutes diffuse across a semipermeable membrane from regions of  higher concentration to regions of lower concentration.   The colorless, odorless, and tasteless nature of water belies its fundamental im¬  portance to living organisms. Despite its bland appearance to our senses, wa¬  ter is anything but inert. Its physical properties—unique among molecules of  similar size—give it unparalleled strength as a solvent. And yet its limitations  as a solvent also have important implications for the structures and functions  of biological molecules.   A water molecule consists of two hydrogen atoms bonded to an oxygen atom.  The O—H bond distance is 0.958A(lA = 10 -10 m), and the angle formed  by the three atoms is 104.5° (Fig. 2-1). The hydrogen atoms are not arranged  linearly, because the oxygen atoms four sp 3 hybrid orbitals extend roughly to¬  ward the corners of a tetrahedron. Hydrogen atoms occupy two corners of the  tetrahedron, and the nonbonding electron pairs of the oxygen atom occupy  the other two corners (in a perfectly tetrahedral molecule, such as methane,  CH 4 , the bond angles are 109.5°).   Water Molecules Form Hydrogen Bonds. The angular geometry of the wa¬  ter molecule has enormous implications for living systems. Water is a polar  molecule: the oxygen atom with its unshared electrons carries a partial nega¬  tive charge (8 ) of ~0.66e, and the hydrogen atoms each carry a partial pos¬  itive charge (8 + ) of +0.33^, where eis the charge of the electron. Electrostatic  attractions between the dipoles of water molecules are crucial to the proper¬  ties of water itself and to its role as a biochemical solvent. Neighboring water  molecules tend to orient themselves so that the O—H bond of one water mol¬  ecule (the positive end) points toward one of the electron pairs of the other  water molecule (the negative end). The resulting directional intermolecular  association is known as a hydrogen bond (Fig. 2-2).   In general\ a hydrogen bond can be represented as D — H---A, where D—H  is a weakly acidic 0 donor” group such as O — H, N — H y or sometimes S — H y  and A is a weakly basic “acceptor” atom such as O, N, or occasionally S. Hydrogen  bonds are structurally characterized by an H***A distance that is at least 0.5 A  shorter than the calculated van der Waals distance (the distance of closest  approach between two nonbonded atoms). In water, for example, the 0***H  hydrogen bond distance is ~1.8 A versus 2.6 A for the corresponding van der  Waals distance.   A single water molecule contains two hydrogen atoms that can be “do¬  nated” and two unshared electron pairs that can act as “acceptors,” so each  molecule can participate in a maximum of four hydrogen bonds with other  water molecules. Although the energy of an individual hydrogen bond  (~20 kj • mol *) is relatively small (e.g., the energy of an O—H covalent   (a) The shaded outline represents the van der  Waals envelope, the effective “surface” of the  molecule, (b) The oxygen atom’s sp 3 orbitals are  arranged tetrahedrally. Two orbitals contain  nonbonding electron pairs.    FIG. 2-2 A hydrogen bond between  two water molecules. The strength of the  interaction is maximal when the 0—H covalent  bond of one molecule points directly toward the  lone-pair electron cloud of the other.    FIG. 2-3 The structure of ice. Each water  molecule interacts tetrahedrally with four other  water molecules. Oxygen atoms are red and  hydrogen atoms are white. Hydrogen bonds are  represented by dashed lines. [After Pauling, L,  The Nature of the Chemical Bond (3rd ed.),  p. 465, Cornell University Press (I960).]    FIG. 2-4 Rings of water molecules. These  models, containing three, four, or five molecules,  are based on theoretical predictions and  spectroscopic data. [After Liu, K., Cruzan, J.D.,  and Saykally, R.J., Science 271, 929 (1996).]    bond is 460 kj • mol x ), the sheer number of hydrogen bonds in a sample of  water is the key to its remarkable properties.   Ice Is a Crystal of Hydrogen-Bonded Water Molecules. The structure of ice  provides a striking example of the cumulative strength of many hydrogen  bonds. X-Ray and neutron diffraction studies have established that water mol¬  ecules in ice are arranged in an unusually open structure. Each water mole¬  cule is tetrahedrally surrounded by four nearest neighbors to which it is  hydrogen bonded (Fig. 2-3). As a consequence of its open structure, water is  one of the very few substances that expands on freezing (at 0°C, liquid water  has a density of 1.00 g • mL x , whereas ice has a density of 0.92 g • mL x ).   The expansion of water on freezing has overwhelming consequences for  life on the earth. Suppose that water contracted on freezing, that is, became  more dense rather than less dense. Ice would then sink to the bottoms of lakes  and oceans rather than float. This ice would be insulated from the sun so that  oceans, with the exception of a thin surface layer of liquid in warm weather,  would be permanently frozen solid (the water at great depths, even in tropi¬  cal oceans, is close to 4°C, its temperature of maximum density). Thus, the  earth would be locked in a permanent ice age and life might never have arisen.   The melting of ice represents the collapse of the strictly tetrahedral orien¬  tation of hydrogen-bonded water molecules, although hydrogen bonds between  water molecules persist in the liquid state. In fact, liquid water is only ~15%  less hydrogen bonded than ice at 0°C. Indeed, the boiling point of water is  264°C higher than that of methane, a substance with nearly the same molec¬  ular mass as H 2 0 but which is incapable of hydrogen bonding (substances with  similar intermolecular associations and equal molecular masses should have  similar boiling points). This difference reflects the extraordinary internal cohe¬  siveness of liquid water resulting from its intermolecular hydrogen bonding.   The Structure of Liquid Water Is Irregular. Because each molecule of liquid  water reorients about once every 10 12 s, very few experimental techniques  can explore the instantaneous arrangement of these water molecules.  Theoretical considerations and spectroscopic evidence suggest that molecules  in liquid water are each hydrogen bonded to four nearest neighbors, as they  are in ice. These hydrogen bonds are distorted, however, so the networks of  linked molecules are irregular and varied. For example, three- to seven-  membered rings of hydrogen-bonded molecules commonly occur in liquid  water (Fig. 2-4), in contrast to the six-membered rings characteristic of ice  (Fig. 2-3). Moreover, these networks continually break up and re-form every  2X10 11 s or so. Liquid water therefore consists of a rapidly fluctuating, three-  dimensional network of hydro gen-bonded H 2 0 molecules.   Hydrogen Bonds and Other Weak Interactions Influence Biological  Molecules. Biochemists are concerned not just with the strong covalent bonds  that define chemical structure but with the weak forces that act under rela¬  tively mild physical conditions. The structures of most biological molecules  are determined by the collective influence of many individually weak interac¬  tions. The weak electrostatic forces that interest biochemists include ionic in¬  teractions, hydrogen bonds, and van der Waals forces.   The strength of association of ionic groups of opposite charge depends on  the chemical nature of the ions, the distance between them, and the polarity  of the medium. In general, the strength of the interaction between two charged  groups (i.e., the energy required to completely separate them in the medium  of interest) is less than the energy of a covalent bond but greater than the en¬  ergy of a hydrogen bond (Table 2-1).   The noncovalent associations between neutral molecules, collectively  known as van der Waals forces, arise from electrostatic interactions among   permanent or induced dipoles (the hydrogen bond is a special kind of di¬  polar interaction). Interactions among permanent dipoles such as carbonyl groups  (Fig. 2-5 a) are much weaker than ionic interactions. A permanent dipole also  induces a dipole moment in a neighboring group by electrostatically distort¬  ing its electron distribution (Fig. 2-5 b). Such dipole-induced dipole interac¬  tions are generally much weaker than dipole—dipole interactions.   At any instant, nonpolar molecules have a small, randomly oriented di¬  pole moment resulting from the rapid fluctuating motion of their electrons.  This transient dipole moment can polarize the electrons in a neighboring  group (Fig. 2-5 c), so that the groups are attracted to each other. These so-  called London dispersion forces are extremely weak and fall off so rapidly  with distance that they are significant only for groups in close contact. They  are, nevertheless, extremely important in determining the structures of biolog¬  ical molecules, whose interiors contain many closely packed groups.   Solubility depends on the ability of a solvent to interact with a solute more  strongly than solute particles interact with each other. Water is said to be the  “universal solvent.” Although this statement cannot literally be true, water cer¬  tainly dissolves more types of substances and in greater amounts than any other  solvent. In particular, the polar character of water makes it an excellent sol¬  vent for polar and ionic materials, which are said to be hydrophilic (Greek:  hydro, water + philos , loving). On the other hand, nonpolar substances are  virtually insoluble in water (“oil and water don’t mix”) and are consequently  described as hydrophobic (Greek: phobos , fear). Nonpolar substances, how¬  ever, are soluble in nonpolar solvents such as CCI 4 and hexane. This informa¬  tion is summarized by another maxim, “like dissolves like.”   Why do salts such as NaCl dissolve in water? Polar solvents, such as wa¬  ter, weaken the attractive forces between oppositely charged ions (such as Na +  and Cl ) and can therefore hold the ions apart. (In nonpolar solvents, ions  of opposite charge attract each other so strongly that they coalesce to form a  solid salt.) An ion immersed in a polar solvent such as water attracts the op¬  positely charged ends of the solvent dipoles (Fig. 2-6). The ion is thereby sur¬  rounded by one or more concentric shells of oriented solvent molecules. Such  ions are said to be solvated or, when water is the solvent, to be hydrated.   The water molecules in the hydration shell around an ion move more  slowly than water molecules that are not involved in solvating the ion. In bulk   The strength of each dipole is indicated by the  thickness of the accompanying arrow.   FIG. 2-6 Solvation of ions. The dipoles of  the surrounding water molecules are oriented  according to the charge of the ion. Only one  layer of solvent molecules is shown.    FIG. 2-7 Hydrogen bonding by functional  groups. Water forms hydrogen bonds with (a)  hydroxyl groups, (b) keto groups, (c) carboxylate  ions, and (d) ammonium ions.    water, the energetic cost of breaking a hydrogen bond is low because another  hydrogen bond is likely to be forming at the same time. The cost is higher  for the relatively ordered water molecules of the hydration shell. The energet¬  ics of solvation also play a role in chemical reactions, since a reacting group  must shed its waters of hydration (the molecules in its hydration shell) in or¬  der to closely approach another group.   The bond dipoles of uncharged polar molecules make them soluble in  aqueous solutions for the same reasons that ionic substances are water soluble.  The solubilities of polar and ionic substances are enhanced when they carry  functional groups, such as hydroxyl (OH), carbonyl (C=0), carboxylate  (COO ), or ammonium (NH 3 ) groups, that can form hydrogen bonds with  water as illustrated in Fig. 2 -7. Indeed, water-soluble biomolecules such as  proteins, nucleic acids, and carbohydrates bristle with just such groups.  Nonpolar substances, in contrast, lack hydrogen-bonding donor and acceptor  groups.   C The Hydrophobic Effect Causes Nonpolar Substances  to Aggregate in Water   When a nonpolar substance is added to an aqueous solution, it does not dis¬  solve but instead is excluded by the water. The tendency of water to minimize  its contacts with hydrophobic molecules is termed the hydrophobic effect. Many  large molecules and molecular aggregates, such as proteins, nucleic acids, and  cellular membranes, assume their shapes at least partially in response to the  hydrophobic effect.   Consider the thermodynamics of transferring a nonpolar molecule from  an aqueous solution to a nonpolar solvent. In all cases, the free energy change  is negative, which indicates that such transfers are spontaneous processes  (Table 2-2). Interestingly, these transfer processes are either endothermic (pos¬  itive A H) or isothermic (A H = 0 ); that is, it is enthalpically more or less  equally favorable for nonpolar molecules to dissolve in water as in nonpolar  media. In contrast, the entropy change (expressed as — TAS) is large and neg¬  ative in all cases. Clearly, the transfer of a hydrocarbon from an aqueous  medium to a nonpolar medium is entropically driven (i.e., the free energy  change is mostly due to an entropy change).   Entropy, or “randomness,” is a measure of the order of a system (Sec¬  tion 1-3B). If entropy increases when a nonpolar molecule leaves an aqueous    TABLE 2-2 Thermodynamic Changes for Transferring Hydrocarbons from Water to Nonpolar Solvents at 25°C   solution, entropy must decrease when the molecule enters water. This decrease  in entropy when a nonpolar molecule is solvated by water is an experimental  observation, not a theoretical conclusion. Yet the entropy changes are too large  to reflect only the changes in the conformations of the hydrocarbons. Thus  the entropy changes must arise mainly from some sort of ordering of the  water itself. What is the nature of this ordering?   The extensive hydrogen-bonding network of liquid water molecules is  disrupted when a nonpolar group intrudes. A nonpolar group can neither  accept nor donate hydrogen bonds, so the water molecules at the surface of  the cavity occupied by the nonpolar group cannot hydrogen bond to other  molecules in their usual fashion. In order to maximize their hydrogen-bond¬  ing ability, these surface water molecules orient themselves to form a hydro¬  gen-bonded network enclosing the cavity (Fig. 2-8). This orientation  constitutes an ordering of the water structure since the number of ways that  water molecules can form hydrogen bonds around the surface of a nonpo¬  lar group is fewer than the number of ways they can form hydrogen bonds  in bulk water.   Unfortunately, the ever-fluctuating nature of liquid water s basic structure  has not yet allowed a detailed description of this ordering process. One model  proposes that water forms icelike hydrogen-bonded “cages” around the non¬  polar groups. The water molecules of the cages are tetrahedrally hydrogen  bonded to other water molecules, and the ordering of water molecules extends  several layers beyond the first hydration shell of the nonpolar solute.   The unfavorable free energy of hydration of a nonpolar substance caused  by its ordering of the surrounding water molecules has the net result that the  nonpolar substance tends to be excluded from the aqueous phase. This is be¬  cause the surface area of a cavity containing an aggregate of nonpolar mole¬  cules is less than the sum of the surface areas of the cavities that each of these  molecules would individually occupy (Fig. 2-9). The aggregation of the non¬  polar groups thereby minimizes the surface area of the cavity and therefore max¬  imizes the entropy of the entire system. In a sense, the nonpolar groups are  squeezed out of the aqueous phase.   Amphiphiles Form Micelles and Bilayers. Most biological molecules have both  polar (or charged) and nonpolar segments and are therefore simultaneously hy¬  drophilic and hydrophobic. Such molecules, for example, fatty acid ions (soaps;  Fig. 2-10) , are said to be amphiphilic or amphipathic (Greek: amp hi , both;  pathos, suffering). How do amphiphiles interact with an aqueous solvent?  Water tends to hydrate the hydrophilic portion of an amphiphile, but it also  tends to exclude the hydrophobic portion. Amphiphiles consequently tend to  form structurally ordered aggregates. For example, micelles are globules of up  to several thousand amphiphilic molecules arranged so that the hydrophilic  groups at the globule surface can interact with the aqueous solvent while the   ch 3 ch 2 ch 2 ch 2 ch 2 ch 2 ch 2 ch 2 ch 2 ch 2 ch 2 ch 2 ch 2 ch 2 ch 2 —c—cr   ch 3 ch 2 ch 2 ch 2 ch 2 ch 2 ch 2 ch 2 -c=C-CH 2 CH 2 CH 2 CH 2 CH 2 CH 2 CH 2 -C -0“   FIG. 2-10 Fatty acid anions (soaps). Palmitate and oleate are amphiphilic  compounds; each has a polar carboxylate group and a long nonpolar hydrocarbon chain.   FIG. 2-8 Orientation of water molecules  around a nonpolar solute. In order to maximize  their number of hydrogen bonds, water  molecules form a “cage” around the solute.  Dashed lines represent hydrogen bonds.   FIG. 2-9 Aggregation of nonpolar mole¬  cules in water, (a) The individual hydration of  dispersed nonpolar molecules (brown) decreases  the entropy of the system because their hydrat¬  ing water molecules (dark blue) are not as free  to form hydrogen bonds, (b) Aggregation of the  nonpolar molecules increases the entropy of the  system, since the number of water molecules  required to hydrate the aggregated solutes is less  than the number of water molecules required to  hydrate the dispersed solute molecules. This  increase in entropy accounts for the spontaneous  aggregation of nonpolar substances in water.    FIG. 2-1 1 Structures of micelles and  bilayers. In aqueous solution, the polar head  groups of amphipathic molecules are hydrated  while the nonpolar tails aggregate by exclusion  from water, (a) A micelle is a spheroidal  aggregate, (b) A bilayer is an extended planar  aggregate.   hydrophobic groups associate at the center, away from the solvent  (Fig. 2-11 a). Of course, the model presented in Fig. 2-11/2 is an oversimpli¬  fication, since it is geometrically impossible for all the hydrophobic groups to  occupy the center of the micelle. Instead, the amphipathic molecules pack in  a more disorganized fashion that buries most of the hydrophobic groups and  leaves the polar groups exposed (Fig. 2-12).   Alternatively, amphiphilic molecules may arrange themselves to form  bilayered sheets or vesicles in which the polar groups face the aqueous phase  (Fig. 2-11 b). In both micelles and bilayers, the aggregate is stabilized by the  hydrophobic effect, the tendency of water to exclude hydrophobic groups.   The consequences of the hydrophobic effect are often called hydrophobic  forces or hydrophobic “bonds.” However, the term bond implies a discrete di¬  rectional relationship between two entities. The hydrophobic effect acts indi¬  rectly on nonpolar groups and lacks directionality. Despite the temptation to  attribute some mutual attraction to a collection of nonpolar groups excluded  from water, their exclusion is largely a function of the entropy of the surround¬  ing water molecules, not some “hydrophobic force” among them (the London  dispersion forces between the nonpolar groups are relatively weak).   The fluid inside cells and surrounding cells in multicellular organisms is full of  dissolved substances ranging from small inorganic ions to huge molecular aggre¬  gates. The concentrations of these solutes affect waters colligative properties,  the physical properties that depend on the concentration of dissolved substances  rather than on their chemical features. For example, solutes depress the freezing  point and elevate the boiling point of water by making it more difficult for wa¬  ter molecules to crystallize as ice or to escape from solution into the gas phase.   Osmotic pressure also depends on the solute concentration. When a solu¬  tion is separated from pure water by a semipermeable membrane that permits  the passage of water molecules but not solutes, water tends to move into the  solution in order to equalize its concentration on both sides of the membrane.  Osmosis is the movement of solvent across the membrane from a region of  high concentration (here, pure water) to a region of relatively low concentra¬  tion (water containing dissolved solute). The osmotic pressure of a solution is  the pressure that must be applied to the solution to prevent the inward flow  of water; it is proportional to the concentration of the solute (Fig. 2-13). For  a 1 M solution, the osmotic pressure is 22.4 atm. Consider the implications of  osmotic pressure for living cells, which are essentially semipermeable sacs of  aqueous solution. In order to minimize osmotic influx of water, which would  burst the relatively weak cell membrane, many animal cells are surrounded by  a solution of similar osmotic pressure (so there is no net flow of water). Another  strategy, used by most plants and bacteria, is to enclose the cell with a rigid  cell wall that can withstand the osmotic pressure generated within.   When an aqueous solution is separated from pure water by a membrane  that is permeable to both water and solutes, solutes move out of the solution  even as water moves in. The molecules move randomly, or diffuse, until the  concentration of the solute is the same on both sides of the membrane. At    FIG. 2-1 2 Model of a micelle. Twenty molecules of octyl glucoside (an eight-carbon  chain with a sugar head group) are shown in space-filling form in this computer-generated  model. The polar 0 atoms of the glucoside groups are red and C atoms are gray. H atoms  have been omitted for clarity. Computer simulations indicate that such micelles have an  irregular, rapidly fluctuating structure (unlike the symmetric aggregate pictured in Fig. 2-1 la)  such that portions of the hydrophobic tails are exposed on the micelle surface at any given  instant. [Courtesy of Michael Garavito and Shelagh Ferguson-Miller, Michigan State University.]   FIG. 2-13 Osmotic pressure, (a) A water-permeable membrane separates a tube of  concentrated solution from pure water, (b) As water moves into the solution by osmosis, the  height of the solution in the tube increases, (c) The pressure that prevents the influx of  water is the osmotic pressure (22.4 atm for a 1 M solution).    this point, equilibrium is established; that is, there is no further net flow of  water or solute (although molecules continue to move in and out through the  membrane). Note that the tendency for solutes to diffuse from an area of high  concentration to an area of low concentration (i.e., down a concentration gra¬  dient) is thermodynamically favored because it is accompanied by an increase  in entropy.   Diffusion of solutes is the basis for the laboratory technique of dialysis.  In this process, solutes smaller than the pore size of the dialysis membrane  freely exchange between the sample and the bulk solution until equilibrium is  reached (Fig. 2-14). Larger substances cannot cross the membrane and remain  where they are. Dialysis is particularly useful for separating large molecules,  such as proteins or nucleic acids, from smaller molecules. Because small solutes  (and water) move freely between the sample and the surrounding medium,  dialysis can be repeated several times to replace the sample medium with an¬  other solution.    FIG. 2-14 Dialysis, (a) A concentrated solution is separated from a large volume of  solvent by a dialysis membrane (shown here as a tube knotted at both ends). Only small  molecules can diffuse through the pores in the membrane, (b) At equilibrium, the concen¬  trations of small molecules are nearly the same on either side of the membrane, whereas  the macromolecules remain inside the dialysis bag.    • Sketch a diagram of a water molecule  and indicate the ends that bear partial  positive and negative charges.   • Compare the structures of ice and water  with respect to the number and geometry  of hydrogen bonds.   • Which of the functional groups listed in  Table 1-2 can function as hydrogen bond  donors? As hydrogen bond acceptors?   • Describe the nature and relative strength  of covalent bonds, ionic interactions, and  van der Waals interactions (hydrogen  bonds, dipole-dipole interactions, and  London dispersion forces).   • Explain why polar substances dissolve in  water while nonpolar substances do not.   • What is the role of entropy in the  hydrophobic effect?   • How does osmosis differ from  diffusion? Which process occurs  during dialysis?   • Describe the osmotic challenges facing a  cell placed in pure water or in a high-salt  solution.   FIG. 2-1 5 Proton jumping. Proton jumps  occur more rapidly than direct molecular  migration, accounting for the observed high  ionic mobilities of hydronium ions (and  hydroxide ions) in aqueous solutions.    Draw a similar diagram to show how a  hydroxide ion jumps.    T Individuals with kidney failure can undergo a dialysis procedure in which  the blood is pumped through a machine containing a semipermeable mem¬  brane. As blood flows along one side of the membrane, a fluid called the  dialysate flows in the opposite direction on the other side. This countercur¬  rent arrangement maximizes the concentration differences between the two so¬  lutions so that waste materials such as urea and creatinine (present at high  concentration in the blood) will efficiently diffuse through the membrane into  the dialysate (where their concentrations are low). Excess water can also be  eliminated, as it moves into the dialysate by osmosis. The “cleansed” blood is  then returned to the patient. Some challenges of clinical dialysis are the re¬  quirement for ultrapure water to prepare the dialysate and the need to mon¬  itor the patients salt and water balance over the long term.   • A water molecule dissociates to form H + and OH - ions, with a dissociation con¬  stant of 10 -14 .   • The acidity of a solution is expressed as a pH value, where pH = -log [H + ].   • An acid is a compound that can donate a proton, and a base is a compound that  can accept a proton.   • A dissociation constant varies with the strength of an acid.   • The Henderson-Hasselbalch equation relates the pH of a solution of a weak acid  to the p K and the concentrations of the acid and its conjugate base.   • A titration curve demonstrates that if the concentrations of an acid and its conju¬  gate base are close, the solution is buffered against changes in pH when acid or  base is added.   • Many biological molecules contain ionizable groups so that they are sensitive to  changes in pH.   Water is not just a passive component of the cell or extracellular environment.   By virtue of its physical properties, water defines the solubilities of other sub¬  stances. Similarly, waters chemical properties determine the behavior of other   Water is a neutral molecule with a very slight tendency to ionize. We express   There is actually no such thing as a free proton (H + ) in solution. Rather, the  proton is associated with a water molecule as a hydronium ion, H 3 0 + . The  association of a proton with a cluster of water molecules also gives rise to struc¬  tures with the formulas H 5 Oj, H 7 Oj, and so on. For simplicity, however, we  often represent these ions by H + . The other product of waters ionization is  the hydroxide ion, OH - .   The proton of a hydronium ion can jump rapidly to another water mol¬  ecule and then to another (Fig. 2-15). For this reason, the mobilities of H +  and OH ions in solution are much higher than for other ions, which must  move through the bulk water carrying their waters of hydration. Proton jumping   is also responsible for the observation that acid—base reactions are among the fastest  reactions that take place in aqueous solution.   The ionization (dissociation) of water is described by an equilibrium ex¬  pression in which the concentration of the parent substance is in the denom¬  inator and the concentrations of the dissociated products are in the numerator:    K is the dissociation constant (here and throughout the text, quantities in  square brackets symbolize the molar concentrations of the indicated sub¬  stances, which in many cases are only negligibly different from their activities;  Section 1-3D). Because the concentration of the undissociated H 2 0 ([H 2 0])  is so much larger than the concentrations of its component ions, it can be  considered constant and incorporated into K to yield an expression for the  ionization of water,   The value of K w ^ the ionization constant of water, is 10 14 at 25°C.   Pure water must contain equimolar amounts of H + and OH , so [H + ] =  [OH - ] = (K^ 1 ^ 2 = 10 -7 M. Since [H + ] and [OH - ] are reciprocally  related by Eq. 2-2, when [H + ] is greater than 10 7 M, [OH ] must be cor¬  respondingly less and vice versa. Solutions with [H + ] = 10 7 M are said to  be neutral, those with [H + ] > 10 7 M are said to be acidic, and those with  [H + ] <10 7 M are said to be basic. Most physiological solutions have  hydrogen ion concentrations near neutrality. For example, human blood is  normally slightly basic with [H + ] = 4.0 X 10 8 M.   The values of [H + ] for most solutions are inconveniently small and thus  impractical to compare. A more practical quantity, which was devised in 1909  by Soren Sorenson, is known as the pH:   The higher the pH, the lower is the H + concentration; the lower the pH, the  higher is the H + concentration (Fig. 2-16). The pH of pure water is 7.0,  whereas acidic solutions have pH <7.0 and basic solutions have pH >7.0  (see Sample Calculation 2-1). Note that solutions that differ by one pH unit  differ in [H + ] by a factor of 10. The pH values of some common substances  are given in Table 2-3.    10 4 mole of H + (as HC1) is added to 1 liter  of pure water. Determine the final pH of the  solution.    Pure water has a pH of 7, so its [H + ] =  10 7 M. The added H + has a concentration  of 10 4 M, which overwhelms the [H + ]  already present. The total [H + ] is therefore  1.0 X 10 -4 M, so that the pH is equal to  —log[H + ] = —log(1.0 X 10 -4 ) = 4.   FIG. 2-16 Relationship of pH and the  concentrations of H + and OH - in water.   Because the product of [H + ] and [OH - ] is a  constant (10 -14 ), [H + ] and [OH - ] are  reciprocally related. Solutions with relatively more  H + are acidic (pH < 7), solutions with relatively  more OH - are basic (pH > 7), and solutions in  which [H + ] = [OH - ] = 10 -7 M are neutral  (pH = 7). Note the logarithmic scale for ion  concentration.   H + and OH ions derived from water are fundamental to the biochemical re¬  actions we will encounter later in this book. Biological molecules, such as pro¬  teins and nucleic acids, have numerous functional groups that act as acids or  bases, for example, carboxyl and amino groups. These molecules influence the  pH of the surrounding aqueous medium, and their structures and reactivities are  in turn influenced by the ambient pH. An appreciation of acid-base chemistry  is therefore essential for understanding the chemical context of many biological  processes. Box 2-1 describes the effect of ocean acidification on marine life.   An Acid Can Donate a Proton. According to a definition formulated in 1923  by Johannes Bronsted and Thomas Lowry, an acid is a substance that can do¬  nate a proton ,, and a base is a substance that can accept a proton. Under the  Bronsted-Lowry definition, an acid-base reaction can be written as   HA + H 2 0 H 3 0 + + A -   An acid (HA) reacts with a base (H 2 0) to form the conjugate base of the  acid (A ) and the conjugate acid of the base (H 3 0 + ). Accordingly, the ac¬  etate ion (CH 3 COO ) is the conjugate base of acetic acid (CH 3 COOH), and  the ammonium ion (NH 4 ) is the conjugate acid of ammonia (NH 3 ). The  acid-base reaction is frequently abbreviated    The human-generated increase in atmospheric carbon dioxide that  is contributing to climate change through global warming is also  impacting the chemistry of the world’s oceans. Atmospheric C0 2  dissolves in water and reacts with it to generate carbonic acid,  which immediately dissociates to form protons and bicarbonate:   C0 2 + H 2 0 H 2 C0 3 H + + HCO 3   The addition of hydrogen ions from C0 2 -derived carbonic acid  therefore leads to a decrease in the pH. Currently, the earth’s  oceans are slightly basic, with a pH of approximately 8.0. It has  been estimated that over the next 100 years, the ocean pH could  drop to about 7.8. Although the oceans act as a C0 2 “sink” that  helps mitigate the increase in atmospheric C0 2 , the increase in  acidity in the marine environment represents an enormous chal¬  lenge to organisms that must adapt to the new conditions.   A variety of marine organisms, including mollusks, many corals,  and some plankton, use dissolved carbonate ions to construct  protective shells of calcium carbonate (CaC0 3 ). However, carbon¬  ate ions can combine with H + to form bicarbonate:   Consequently, the increase in ocean acidity could decrease the  availability of carbonate and thereby slow the growth of shell¬  building organisms. In fact, experiments have shown that calcifica¬  tion is reduced in organisms such as sea urchins and corals under  acidic conditions. It is also possible that ocean acidification could  dissolve existing carbonate-based coral reefs, which are species-  rich ecosystems and important components of marine food chains.   Interestingly, not all shell-building organisms respond to high levels  of C0 2 in the same way. Experiments with coccolithophores    (single-celled eukaryotes that are encased in calcium carbonate  plates; see photo) indicate that at least under some conditions,  increased C0 2 leads to increased bicarbonate that actually con¬  tributes to increased calcification:   Ca 2+ + 2 HCO 3 -> CaC0 3 + C0 2 + H 2 0   These results suggest that the impact of increased C0 2 on marine  organisms may not be a simple matter of decreased pH but may  be a more complicated function of the relative amounts of all the  carbon species, which include dissolved C0 2 , HC0 3 , and C0 3 “.    The Strength of an Acid Is Specified by Its Dissociation Constant. The   equilibrium constant for an acid-base reaction is expressed as a dissociation  constant with the concentrations of the “reactants” in the denominator and  the concentrations of the “products” in the numerator:    In dilute solutions, the water concentration is essentially constant, 55.5 M (1000  g • L _1 /18.015 g • mol -1 = 55.5 M). Therefore, the term [H 2 OJ is custom¬  arily combined with the dissociation constant, which then takes the form   The dissociation constants of some common acids are listed in Table 2-4.  Because acid dissociation constants, like [H + ] values, can be cumbersome to work  with, they are transformed to p K values by the formula    TABLE 2-4 Dissociation Constants and pK Values at 25°C of Some Acids    ^The pi6for the overall reaction C0 2 + H 2 0 v H 2 C0 3 v H + + HC0 3 ; see Box 2-2.  Source: Dawson, R.M.C., Elliott, D.C., Elliott, W.H., and Jones, K.M., Data for Biochemical  Research (3rd ed.), pp. 424-423, Oxford Science Publications (1986); and Good, N.E., Winget,  G.D., Winter, W., Connolly, T.N., Izawa, S., and Singh, Biochemistry 5, 467 (1966).   Chapter 2 Water Acids can be classified according to their relative strengths, that is, their   abilities to transfer a proton to water. The acids listed in Table 2-4 are known  as weak acids because they are only partially ionized in aqueous solution  {K < 1 ). Many of the so-called mineral acids, such as HCIO 4 , HNO 3 , and  HC1, are strong acids (K 1 ). Since strong acids rapidly transfer all their  protons to H 2 0, the strongest acid that can stably exist in aqueous solutions is H 3 0 + .  Likewise, there can be no stronger base in aqueous solutions than OH . Virtually  all the acid-base reactions that occur in biological systems involve H 3 0 + (and  OH ) and weak acids (and their conjugate bases).    The pH of a Solution Is Determined by the Relative Concentrations of  Acids and Bases. The relationship between the pH of a solution and the  concentrations of an acid and its conjugate base is easily derived. Equation 2-5  can be rearranged to    Calculate the pH of a 2 L solution contain¬  ing 10 mL of 5 M acetic acid and 10 mL  of 1 M sodium acetate.    First, calculate the concentrations of the  acid and conjugate base, expressing all con¬  centrations in units of moles per liter.   Substitute the concentrations of the acid  and conjugate base into the Henderson-  Hasselbalch equation. Find the p K for  acetic acid in Table 2-4.   pH = p K+ log([acetate]/[acetic acid])  pH = 4.76 + log(0.005/0.025)  pH = 4.76 — 0.70  pH = 4.06    This relationship is known as the Henderson-Hasselbalch equation. When  the molar concentrations of an acid (HA) and its conjugate base (A ) are equal '  log ([A ]/ [HA]) = log 1 = 0, and the pH of the solution is numerically equiv¬  alent to the pK of the acid. The Henderson-Hasselbalch equation is invaluable  for calculating, for example, the pH of a solution containing known quanti¬  ties of a weak acid and its conjugate base (see Sample Calculation 2 - 2 ).  However, since the Henderson-Hasselbalch equation does not account for  the ionization of water itself, it is not useful for calculating the pH of solu¬  tions of strong acids or bases. For example, in a 1 M solution of a strong  acid, [H + ] = 1 M and the pH is 0. In a 1 M solution of a strong base,  [OH - ] = 1 M, so [H + ] = [OH~]/A^ = 1 X 10 “ 14 M and the pH is 14.   Adding a 0 . 01 -mL droplet of 1 M HC1 to 1 L of pure water changes the pH  of the water from 7 to 5, which represents a 100-fold increase in [H + ]. Such  a huge change in pH would be intolerable to most biological systems, since  even small changes in pH can dramatically affect the structures and functions  of biological molecules. Maintaining a relatively constant pH is therefore of  paramount importance for living systems. To understand how this is possible,  consider the titration of a weak acid with a strong base.   Figure 2-17 shows how the pH values of solutions of acetic acid, H 2 PC >4 ,  and ammonium ion (NH^) vary as OH is added. Titration curves such as  these can be constructed from experimental observation or by using the  Henderson-Hasselbalch equation to calculate points along the curve (see  Sample Calculation 2-3). When OH reacts with HA, the products are A  and water:   FIG. 2-1 7 Titration curves for acetic acid, phosphate, and ammonia. At the start¬  ing point, the acid form predominates. As strong base (e.g., NaOH) is added, the acid is  converted to its conjugate base. At the midpoint of the titration, where pH = p K, the  concentrations of the acid and the conjugate base are equal. At the end point (equivalence  point), the conjugate base predominates, and the total amount of OH - that has been  added is equivalent to the amount of acid that was present at the starting point. The  shaded bands indicate the pH ranges over which the corresponding solution can function  as a buffer. Q See the Animated Figures.    Several details about the titration curves in Fig. 2-17 should be noted:   1 . The curves have similar shapes but are shifted vertically along the pH  axis.   2 . The pH at the midpoint of each titration is numerically equivalent to  the p K of its corresponding acid; at this point, [HA] = [A ].   3. The slope of each titration curve is much lower near its midpoint than  near its wings. This indicates that when [HA] ~ [A ], the pH of the so¬  lution is relatively insensitive to the addition of strong base or strong acid.  Such a solution, which is known as an acid-base buffer, resists pH  changes because small amounts of added H + or OH react with A or  HA, respectively, without greatly changing the value of log([A ]/[HA]).    Calculate the pH of a 1 L solution contain¬  ing 0.1 M formic acid and 0.1 M sodium  formate before and after the addition of 1  mL of 5 M NaOH. How much would the  pH change if the NaOH were added to 1  L of pure water?    According to Table 2-4, the p K for formic  acid is 3.75. Since [formate] = [formic  acid], the log ([A - ]/[HA]) term of the  Henderson-Hasselbalch equation is 0 and  pH = p K= 3.75. The addition of 1 mL  of NaOH does not significantly change the  volume of the solution, so the [NaOH] is  (0.001 L)(5 M)/(l L) = 0.005 M.   Since NaOH is a strong base, it com¬  pletely dissociates, and [OH - ] = [NaOH]  = 0.005 M. This OH - reacts with formic  acid to produce formate and H 2 0.  Consequently, the concentration of formic  acid decreases and the concentration of for¬  mate increases by 0.005 M.   The new formic acid concentration is  0.1 M — 0.005 M = 0.095 M, and the new  formate concentration is 0.1 M + 0.005 M  = 0.105 M. Substituting these values into  the Henderson-Hasselbalch equation gives   pH = p K+ log ([formate]/[formic acid])  pH = 3.75 + log(0.105/0.095)  pH = 3.75 + 0.04  pH = 3.79   In the absence of the formic acid buffering  system, the [H + ] and therefore the pH can  be calculated directly from K w . Since K w =   [OH - ] ~~ (0.005)  pH = —log[H + ] = —log(2 X 10 -12 ) = 11.7   The first and second equivalence points for  titration of H 3 P0 4 occur at the steepest parts of  the curve. The pH at the midpoint of each stage  provides the p K value of the corresponding  ionization. Q See the Animated Figures.    Sketch the titration curve for the  diprotic acid succinic acid and label the  midpoints and equivalence points.   Substances that can lose more than one proton, or undergo more than  one ionization, such as H 3 PO 4 or H 2 C0 3 , are known as polyprotic acids.  The titration curves of such molecules, as illustrated in Fig. 2-18 for H 3 PO 4 ,  are more complicated than the titration curves of monoprotic acids such as  acetic acid. A polyprotic acid has multiple pK values, one for each ionization  step. H 3 PO 4 , for example, has three dissociation constants because the ionic  charge resulting from one proton dissociation electrostatically inhibits further  proton dissociation, thereby increasing the corresponding piTvalues. Similarly,  a molecule with more than one ionizable group has a discrete p K value for  each group. In a biomolecule that contains numerous ionizable groups with  different p K values, the many dissociation events may yield a titration curve  without any clear “plateaus.”   Biological fluids, both intracellular and extracellular, are heavily buffered.  For example, the pH of the blood in healthy individuals is closely controlled  at pH 7.4 (see Box 2 - 2 ). The phosphate and bicarbonate ions in most biolog¬  ical fluids are important buffering agents because they have pAs in this range  (Table 2-4). Moreover, many biological molecules, such as proteins and some  lipids, as well as numerous small organic molecules, bear multiple acid-base  groups that are effective buffer components in the physiological pH range.   The concept that the properties of biological molecules vary with the acid¬  ity of the solution in which they are dissolved was not fully appreciated before  the twentieth century. Many early biochemical experiments were undertaken  without controlling the acidity of the sample, so the results were often poorly  reproducible. Nowadays, biochemical preparations are routinely buffered to  simulate the properties of naturally occurring biological fluids. A number of  synthetic compounds have been developed for use as buffers; some of these are  included in Table 2-4. The buffering capacity of these weak acids (their abil¬  ity to resist pH changes on addition of acid or base) is maximal when pH =  pAT It is helpful to remember that a weak acid is in its useful buffer range within  one pH unit of its p K (e.g., the shaded regions of Fig. 2-17). Above this range,    Bicarbonate is the most significant buffer compound in human  blood; other buffering agents, including proteins and organic acids,  are present at much lower concentrations. The buffering capacity of  blood depends primarily on two equilibria: ( 1 ) between gaseous C0 2  dissolved in the blood and carbonic acid formed by the reaction   and ( 2 ) between carbonic acid and bicarbonate formed by the  dissociation of H +   The overall p K for these two sequential reactions is 6.35. (The  further dissociation of HC0 3 to CO§ - , p K = 10.33, is not signifi¬  cant at physiological pH.)   When the pH of the blood falls due to metabolic production of  H + , the bicarbonate-carbonic acid equilibrium shifts toward more  carbonic acid. At the same time, carbonic acid loses water to  become C0 2 , which is then expired in the lungs as gaseous C0 2 .  Conversely, when the blood pH rises, relatively more HC0 3 forms.  Breathing is adjusted so that increased amounts of C0 2 in the lungs    can be reintroduced into the blood for conversion to carbonic acid.  In this manner, a near-constant hydrogen ion concentration can be  maintained. The kidneys also play a role in acid-base balance by  excreting HC0 3 and NH 4 .   Disturbances in the blood buffer system can lead to conditions  known as acidosis, with a pH as low as 7.1, or alkalosis, with a pH  as high as 7.6. (Deviations of less than 0.05 pH unit from the “nor¬  mal” value of 7.4 are not significant.) For example, obstructive lung  diseases that prevent efficient expiration of C0 2 can cause respira¬  tory acidosis. Hyperventilation accelerates the loss of C0 2 and  causes respiratory alkalosis. Overproduction of organic acids from  dietary precursors or sudden surges in lactic acid levels during  exercise can lead to metabolic acidosis.   Acid-base imbalances are best alleviated by correcting the un¬  derlying physiological problem. In the short term, acidosis is com¬  monly treated by administering NaHC0 3 intravenously. Alkalosis is  more difficult to treat. Metabolic alkalosis sometimes responds to  KCI or NaCI (the additional CP helps minimize the secretion of H +  by the kidneys), and respiratory alkalosis can be ameliorated by  breathing an atmosphere enriched in C0 2 .    where the ratio [A ]/[HA] >10, the pH of the solution changes rapidly with  added strong base. A buffer is similarly impotent with the addition of strong  acid when its p K exceeds the pH by more than one unit.   In the laboratory, the desired pH of the buffered solution determines  which buffering compound is selected. Typically, the acid form of the com¬  pound and one of its soluble salts are dissolved in the (nearly equal) molar  ratio necessary to provide the desired pH and, with the aid of a pH meter,  the resulting solution is fine-tuned by titration with strong acid or base (see  Sample Calculation 2-4).    How many milliliters of a 2.0 M solution of boric acid must be added to 600 mL  of a solution of 10 mM sodium borate in order for the pH to be 9.45?    Rearrange the Henderson-Hasselbalch equation to isolate the [A ] / [HA] term:    Substitute the known p K (from Table 2-4) and the desired pH:    The starting solution contains (0.6 L)(0.01 mol • L : ) = 0.006 mole of borate (A ).  The amount of boric acid (HA) needed is 0.006 mol/1.62 = 0.0037 mol.   Since the stock boric acid is 2.0 M, the volume of boric acid to be added is  (0.0037 mol)/(2.0 mol • L l ) = 0.0019 L or 1.9 mL.   • What are the products of water’s ioniza¬  tion? How are their concentrations related?   • Predict the pH of a sample of water if K w  were 10 -10 or 10 -20 .   • Describe how to calculate pH from the  concentration of H + or OH - .   • What is the relationship between the  strength of an acid and its p K value?   • Explain why it is more complicated to calcu¬  late the pH of a solution of weak acid or  base than to calculate the pH of a solution  of strong acid or base.   • Be able to sketch a titration curve, and label  its parts, for a monoprotic and a polyprotic  acid.   • What must a buffer solution include in order  to resist changes in pH on addition of acid  or base?   • Why is it important to maintain biological  molecules in a buffered solution?   • Water molecules can form hydrogen bonds with other molecules  because they have two H atoms that can be donated and two un¬  shared electron pairs that can act as acceptors.   • Liquid water is an irregular network of water molecules that  each form up to four hydrogen bonds with neighboring water  molecules.   • Hydrophilic substances such as ions and polar molecules dissolve  readily in water.   • The hydrophobic effect is the tendency of water to minimize its  contacts with nonpolar substances.   • Water molecules move through semipermeable membranes from  regions of high concentration to regions of low concentration by   hydrogen bond 23  van der Waals distance 23  van der Waals forces 24  London dispersion forces 25  hydrophilic 25  hydrophobic 25  solvation 25  hydration 25  waters of hydration 26  hydrophobic effect 26    1 . Identify the potential hydrogen bond donors and acceptors in the  following molecules:    2 . Occasionally, a C—H group can form a hydrogen bond. Why  would such a group be more likely to be a hydrogen bond donor  group when the C is next to N?    osmosis; solutes move from regions of high concentration to regions  of low concentration by diffusion.   • Water ionizes to H + (which represents the hydronium ion,  H 3 0 + ) and OH - .   • The concentration of H + in solutions is expressed as a pH value;  in acidic solutions pH < 7, in basic solutions pH > 7, and in  neutral solutions pH = 7.   • Acids can donate protons and bases can accept protons. The  strength of an acid is expressed as its p K   • The Henderson-Hasselbalch equation relates the pH of a solution  to the p K and concentrations of an acid and its conjugate base.   • Buffered solutions resist changes in pH within about one pH unit  of the p K of the buffering species.    p K 33  weak acid 34  strong acid 34  Henderson-Hasselbalch  equation 34  titration curve 34  buffer 35  polyprotic acid 3 6  buffering capacity 36  acidosis 37  alkalosis 37   7. Describe what happens when a dialysis bag containing pure wa¬  ter is suspended in a beaker of seawater. What would happen if  the dialysis membrane were permeable to water but not solutes?   8 . A red blood cell has an internal salt concentration of —130 mM.  The cell is placed in a beaker of 500 mM salt, (a) Assuming the  cell membrane is permeable to water but not to ions, describe  what will happen to the cell in terms of osmosis, (b) If the mem¬  brane were permeable to ions, in which direction would solutes  diffuse: into or out of the cell?   9. Many foods must be refrigerated to prevent spoiling (microbial  growth). Explain why honey (which is —82% carbohydrate by  weight) resists microbial growth even at room temperature.   10. You have a 5 mL sample of a protein in 0.5 M NaCl. You place  the protein/salt sample inside dialysis tubing (see Fig. 2-14) and  place the bag in a large beaker of distilled water. If your goal is to  remove as much NaCl from the sample as possible, which would  be more effective: (1) placing the dialysis bag in 4 L of distilled wa¬  ter for 12 h, or (2) placing the bag in 1 L of distilled water for 6 h  and then in another 1 L of fresh distilled water for another 6 h?   11. Patients with kidney failure frequently develop metabolic acidosis.  If such patients undergo dialysis, the dialysate includes sodium  bicarbonate at a concentration higher than that of the blood.  Explain why this would benefit the patient.   12 . The kidneys function to eliminate ammonia from the blood. Based  on ammonias pK value, what is the molecular form that predomi¬  nates in the blood? Could this molecule easily diffuse through the  hydrophobic lipid membrane of a kidney cell? Explain.   13. Draw the structures of the conjugate bases of the following acids:    14. Draw the structures of the conjugate bases of the following acids:   13. Indicate the ionic species of ammonia that predominates at pH 4,  8 , and 11 .   16. Indicate the ionic species of phosphoric acid that predominates  at pH 4, 8 , and 11.   17. Calculate the pH of a 200 mL solution of pure water to which  has been added 30 mL of 1 mM HC1.   18. Calculate the pH of a 1 L solution containing (a) 10 mL of 5 M  NaOH, (b) 10 mL of 100 mM glycine and 20 mL of 5 M HC1,  and (c) 10 mL of 2 M acetic acid and 5 g of sodium acetate  (formula weight 82 g • mol 1 ).   19. A solution is made by mixing 50 mL of 2.0 M K 2 HPO 4 and 25  mL of 2.0 M KH 2 PO 4 . The solution is diluted to a final volume  of 200 mL. What is the pH of the final solution?   20. What is the piTof the weak acid HA if a solution containing 0.1  M HA and 0.2 M A“ has a pH of 6.5?   21 . How many grams of sodium succinate (formula weight 140 g •  mol x ) and disodium succinate (formula weight 162 g • mol x )  must be added to 1 L of water to produce a solution with pH 6.0  and a total solute concentration of 50 mM?   22 . Estimate the volume of a solution of 5 M NaOH that must be  added to adjust the pH from 4 to 9 in 100 mL of a 100 mM  solution of phosphoric acid.   23. Calculate the standard free energy change for the dissociation of  HEPES.   24. (a) Would phosphoric acid or succinic acid be a better buffer at   (b) Would ammonia or piperidine be a better buffer at pH 9?   (c) Would HEPES or Tris be a better buffer at pH 7.5?   25. You need a buffer at pH 7.5 for use in purifying a protein at 4°C.  You have chosen Tris, p K 8.08, A H° = 50 kj • mol 1 . You care¬  fully make up 0.01 M Tris buffer, pH 7.5 at 25°C, and store it in  the cold box to equilibrate it to the temperature of the purification.  When you measure the pH of the temperature-equilibrated buffer  it has increased to 8.1. What is the explanation for this increase?  How can you avoid this problem?   26. Glycine hydrochloride (Cl H 3 N + CH 2 COOH) is a diprotic acid  that contains a carboxylic acid group and an ammonium group  and is therefore called an amino acid. It is often used in biochem¬  ical buffers.   (a) Which proton would you expect to dissociate at a lower pH,  the proton of the carboxylic acid group or the ammonium  group?   (b) Write the chemical equations describing the dissociation of  the first and second protons of Cl H 3 N + CH 2 COOH.   (c) A solution containing 0.01 M Cl H 3 N + CH 2 COOH and  0.02 M of the monodissociated species has pH = 2.65. What  is the p K of this dissociation?   (d) In analogy with figure 2-18, sketch the titration curve of this  diprotic acid.   Case 1 Acute Aspirin Overdose: Relationship to the  Blood Buffering System   Focus concept: The carbonic acid-bicarbonate buffering system  responds to an overdose of aspirin.   • Principles of acids and bases, including p K and the  Henderson-Hasselbalch equation.    Compare and contrast the  strategies for eliminating waste nitrogen (as ammonia) and carbon  dioxide in (a) a terrestrial mammal, (b) a freshwater fish, and (c) a  saltwater fish. Be sure to consider osmotic effects as well as  acid-base balance.    Finney, J.L., Water? Whats so special about it? Philos. Trans. R. Soc. London  B Biol. Sci. 29, 1145-1163 (2004). [Includes discussions of the struc¬  ture of water molecules, hydrogen bonding, structures of ice and liq¬  uid water, and how these relate to biological function.]   Gerstein, M. and Levitt, M., Simulating water and the molecules of life,  Sci. Am. 279(11), 101-105 (1998). [Describes the structure of water  and how water interacts with other molecules.]   Good, N.E., Winget, G.D., Winter, W., Connolly, T.N., Izawa, S., and  Singh, R.M.M., Hydrogen ion buffers for biological research, Bio¬  chemistry 5, 467-477 (1966). [A classic paper on laboratory buffers.]  Halperin, M.L. and Goldstein, M.B., Fluid, Electrolyte, and Acid—Base  Physiology: A Problem-Based Approach (3rd ed.), W.B. Saunders  (1999). [Includes extensive problem sets with explanations of basic  science as well as clinical effects of acid-base disorders.]    Jeffrey, G.A. and Saenger, W., Hydrogen Bonding in Biological Structures,  Chapters 1, 2 , and 21 , Springer (1994). [Reviews hydrogen bond  chemistry and its importance in small molecules and macromolecules.]  Lynden-Bell, R.M., Morris, S.C., Barrow, J.D., Finney, J.L., and Harper,  C.L., Jr., Water and Life. The Unique Properties of H 2 0, CRC Press  ( 2010 ).   Segel, I.H., Biochemical Calculations (2nd ed.), Chapter 1 , Wiley (1976).  [An intermediate level discussion of acid-base equilibria with  worked-out problems.]   Tanford, C., The Hydrophobic Effect: Formation of Micelles and Biological  Membranes (2nd ed.), Chapters 5 and 6 , Wiley-Interscience (1980).  [Discusses the structures of water and micelles.]   A Nucleic Acids Are Polymers of Nucleotides  B DNA Forms a Double Helix  C RNA Is a Single-Stranded Nucleic Acid   C Traditional DNA Sequencing Uses the  Chain-Terminator Method  D Entire Genomes Have Been Sequenced  E Evolution Results from Sequence Mutations   A Cloned DNA Is an Amplified Copy  B DNA Libraries Are Collections of Cloned DNA  C DNA Is Amplified by the Polymerase  Chain Reaction    DNA recovered from these Neandertal bone fragments yielded nucleotide sequence  data that suggest-contrary to previous findings—that Neandertals and modern humans  did interbreed to a limited extent. This likely occurred sometime after modern humans  emigrated from Africa and before the Neandertals became extinct some 30,000 years  ago. [Max Planck Institute for Evolutionary Anthropology, Leipzig, Germany.]    Despite obvious differences in lifestyle and macroscopic appearance, organisms  exhibit striking similarity at the molecular level. The structures and metabolic  activities of all cells rely on a common set of molecules that includes amino  acids, carbohydrates, lipids, and nucleotides, as well as their polymeric forms.  Each type of compound can be described in terms of its chemical makeup, its  interactions with other molecules, and its physiological function. We begin  our survey of biomolecules with a discussion of the nucleotides and their poly¬  mers, the nucleic acids.   Nucleotides are involved in nearly every facet of cellular life. Specifically,  they participate in oxidation—reduction reactions, energy transfer, intracellular  signaling, and biosynthetic reactions. Their polymers, the nucleic acids DNA  and RNA, are the primary players in the storage and decoding of genetic  information. Nucleotides and nucleic acids also perform structural and catalytic  roles in cells. No other class of molecules participates in such varied functions  or in so many functions that are essential for life.   Evolutionists postulate that the appearance of nucleotides permitted the  evolution of organisms that could harvest and store energy from their sur¬  roundings and, most importantly, could make copies of themselves. Although  the chemical and biological details of early life-forms are the subject of spec¬  ulation, it is incontrovertible that life as we know it is inextricably linked to  the chemistry of nucleotides and nucleic acids.   In this chapter, we briefly examine the structures of nucleotides and the  nucleic acids DNA and RNA. We also consider how the chemistry of these  molecules allows them to carry biological information in the form of a   sequence of nucleotides. This information is expressed by the transcription of Section 1 Nucleotides   a segment of DNA to yield RNA, which is then translated to form protein.   Because a cells structure and function ultimately depend on its genetic  makeup, we discuss how genomic sequences provide information about evo¬  lution, metabolism, and disease. Finally, we consider some of the techniques  used in manipulating DNA in the laboratory. In later chapters, we will exam¬  ine in greater detail the participation of nucleotides and nucleic acids in meta¬  bolic processes. Chapter 24 includes additional information about nucleic acid  structures, DNAs interactions with proteins, and DNA packaging in cells, as  a prelude to several chapters discussing the roles of nucleic acids in the stor¬  age and expression of genetic information.   • The nitrogenous bases of nucleotides include two types of purines and three  types of pyrimidines.   • A nucleotide consists of a nitrogenous base, a ribose or deoxyribose sugar, and  one or more phosphate groups.   • DNA contains adenine, guanine, cytosine, and thymine deoxyribonucleotides,  whereas RNA contains adenine, guanine, cytosine, and uracil ribonucleotides.   Nucleotides are ubiquitous molecules with considerable structural diversity.  There are eight common varieties of nucleotides, each composed of a nitrogenous  base linked to a sugar to which at least one phosphate group is also attached. The  bases of nucleotides are planar, aromatic, heterocyclic molecules that are struc¬  tural derivatives of either purine or pyrimidine (although they are not syn¬  thesized in vivo from either of these organic compounds).    The most common purines are adenine (A) and guanine (G), and the  major pyrimidines are cytosine (C), uracil (U), and thymine (T). The purines  form bonds to a five-carbon sugar (a pentose) via their N9 atoms, whereas  pyrimidines do so through their N1 atoms (Table 3-1).   In ribonucleotides, the pentose is ribose, while in deoxyribonucleotides  (or just deoxynucleotides), the sugar is 2'-deoxyribose (i.e., the carbon at  position 2' lacks a hydroxyl group).    Note that the “primed” numbers refer to the atoms of the pentose; “unprimed”  numbers refer to the atoms of the nitrogenous base.   In a ribonucleotide or a deoxyribonucleotide, one or more phosphate  groups are bonded to atom C3' or atom C5' of the pentose to form a  3'-nucleotide or a 5'-nucleotide, respectively (Fig. 3-1). When the phosphate  group is absent, the compound is known as a nucleoside. A 5'-nucleotide can  therefore be called a nucleoside-5'-phosphate. Nucleotides most commonly   FIG. 3-1 Chemical structures of  nucleotides, (a) A 5'-ribonucleotide and ( b ) a  3'-deoxynucleotide. The purine or pyrimidine base  is linked to Cl' of the pentose and at least one  phosphate ( red) is also attached. A nucleoside  consists only of a base and a pentose.    The presence of a 2'-deoxyribose unit in place of ribose, as occurs in DNA, is implied by the prefixes “deoxy” or “d.” For example, the deoxynucleoside of  adenine is deoxyadenosine or dA. However, for thymine-containing residues, which rarely occur in RNA, the prefix is redundant and may be dropped. The  presence of a ribose unit may be explicitly implied by the prefix “ribo.”   Hhe position of the phosphate group in a nucleotide may be explicitly specified as in, for example, 3'-AMP and 5 , -GMP.    Without looking at the table, give the name of each base and its corresponding nucleoside.    contain one to three phosphate groups at the C5' position and are called nu¬  cleoside monophosphates, diphosphates, and triphosphates.   The structures, names, and abbreviations of the common bases, nucleo¬  sides, and nucleotides are given in Table 3-1. Ribonucleotides are components  of RNA (ribonucleic acid), whereas deoxynucleotides are components of  DNA (deoxyribonucleic acid). Adenine, guanine, and cytosine occur in both  ribonucleotides and deoxynucleotides (accounting for six of the eight common  nucleotides), but uracil primarily occurs in ribonucleotides and thymine oc¬  curs in deoxynucleotides. Free nucleotides, which are anionic, are almost al¬  ways associated with the counterion Mg 2+ in cells.   Nucleotides Participate in Metabolic Reactions. The bulk of the nucleotides  in any cell are found in polymeric forms, as either DNA or RNA, whose pri¬  mary functions are information storage and transfer. However, free nucleotides   Perhaps the best known nucleotide is adenosine triphosphate (ATP), a  nucleotide containing adenine, ribose, and a triphosphate group. ATP is often  mistakenly referred to as an energy-storage molecule, but it is more accurately  termed an energy carrier or energy transfer agent. The process of photosynthe¬  sis or the breakdown of metabolic fuels such as carbohydrates and fatty acids  leads to the formation of ATP from adenosine diphosphate (ADP):    ATP diffuses throughout the cell to provide energy for other cellular work,  such as biosynthetic reactions, ion transport, and cell movement. The chemi¬  cal potential energy of ATP is made available when it transfers one (or two)  of its phosphate groups to another molecule. This process can be represented  by the reverse of the preceding reaction, namely, the hydrolysis of ATP to ADP.  (As we will see in later chapters, the interconversion of ATP and ADP in the  cell is not freely reversible, and free phosphate groups are seldom released di¬  rectly from ATP.) The degree to which ATP participates in routine cellular ac¬  tivities is illustrated by calculations indicating that while the concentration of  cellular ATP is relatively moderate (~5 mM), humans typically recycle their  own weight of ATP each day.   Nucleotide derivatives participate in a wide variety of metabolic processes.  For example, starch synthesis in plants proceeds by repeated additions of glu¬  cose units donated by ADP-glucose (Fig. 3-2). Other nucleotide derivatives,  as we will see in later chapters, carry groups that undergo oxidation-reduction  reactions. The attached group, which may be a small molecule such as glucose  (Fig. 3-2) or even another nucleotide, is typically linked to the nucleotide  through a mono- or diphosphate group.   • Identify the purines and pyrimidines  commonly found in nucleic acids.   • Describe the chemical differences  between a ribonucleoside triphosphate  and a deoxyribonucleoside monophosphate.   FIG. 3-2 ADP-glucose. In this nucleotide derivative, glucose (blue) is attached to  adenosine (black) by a diphosphate group (red).   • In the DNA double helix, two antiparallel polynucleotide strands wind around each  other, interacting via hydrogen bonding between bases in opposite strands.   • RNA molecules are usually single-stranded and may form intramolecular base pairs.   Nucleotides can be joined to each other to form the polymers that are familiar  to us as RNA and DNA. In this section, we describe the general features of  these nucleic acids. Nucleic acid structure is considered further in Chapter 24.   The nucleic acids are chains of nucleotides whose phosphates bridge the 3'  and 5' positions of neighboring ribose units (Fig. 3-3). The phosphates of  these polynucleotides are acidic, so at physiological pH, nucleic acids are  polyanions. The linkage between individual nucleotides is known as a phos¬  phodiester bond, so named because the phosphate is esterified to two ribose  units. Each nucleotide that has been incorporated into the polynucleotide is  known as a nucleotide residue. The terminal residue whose C5' is not linked  to another nucleotide is called the 5 ' end, and the terminal residue whose C3'  is not linked to another nucleotide is called the 3' end. By convention, the  sequence of nucleotide residues in a nucleic acid is written, left to right, from  the 5' end to the 3' end.   tetraribonucleotide adenylyl-3',5'-uridylyl-3',5'-cytidylyl-3',5'-guany-  late is shown. The sugar atoms are primed to distinguish them  from the atoms of the bases. By convention, a polynucleotide  sequence is written with the 5' end at the left and the 3' end at  the right. Thus, reading left to right, the phosphodiester bond  links neighboring ribose residues in the 5' —> 3' direction. The  sequence shown here can be abbreviated pApUpCpG or just  pAUCG (the “p” to the left of a nucleoside symbol indicates a 5'  phosphoryl group). The corresponding deoxytetranucleotide, in  which the 2'-OH groups are replaced by H and the uracil (U) is  replaced by thymine (T), is abbreviated d(pApTpCpG) or  d(pATCG). (b) Schematic representation of pAUCG. A vertical line  denotes a ribose residue, the attached base is indicated by a  single letter, and a diagonal line flanking an optional “p” repre¬  sents a phosphodiester bond. The atom numbers for the ribose  residue may be omitted. The equivalent representation of  d(pATCG) differs only by the absence of the 2'-OH group and  the replacement of U by T.    FIG. 3-4 Tautomeric forms of bases. Some of the possible tautomeric forms of (a)  thymine and (b) guanine are shown. Cytosine and adenine can undergo similar proton shifts.    The properties of a polymer such as a nucleic acid may be very different  from the properties of the individual units, or monomers, before polymeriza¬  tion. As the size of the polymer increases from dimer, trimer, tetramer, and  so on through oligomer (Greek: oligo, few), physical properties such as charge  and solubility may change. In addition, a polymer of nonidentical residues has  a property that its component monomers lack — namely ; it contains information in  the form of its sequence of residues.   Chargaff’s Rules Describe the Base Composition of DNA. Although there  appear to be no rules governing the nucleotide composition of typical RNA  molecules, DNA has equal numbers of adenine and thymine residues (A = T)  and equal numbers of guanine and cytosine residues (G = C). These relation¬  ships, known as ChargafFs rules, were discovered in the late 1940s by Erwin  Chargaff, who devised the first reliable quantitative methods for the composi¬  tional analysis of DNA.   DNAs base composition varies widely among different organisms. It  ranges from ~25 to 75 mol % G + C in different species of bacteria.  However, it is more or less constant among related species; for example, in  mammals G + C ranges from 39 to 46%. The significance of Chargaff s rules  was not immediately appreciated, but we now know that the structural basis  for the rules derives from DNAs double-stranded nature.   The determination of the structure of DNA by James Watson and Francis  Crick in 1953 is often said to mark the birth of modern molecular biology.  The Watson-Crick structure of DNA not only provided a model of what is  arguably the central molecule of life, it also suggested the molecular mecha¬  nism of heredity. Watson and Cricks accomplishment, which is ranked as one  of sciences major intellectual achievements, was based in part on two pieces  of evidence in addition to Chargaff s rules: the correct tautomeric forms of  the bases and indications that DNA is a helical molecule.   The purine and pyrimidine bases of nucleic acids can assume different  tautomeric forms (tautomers are easily converted isomers that differ only in  hydrogen positions; Fig. 3-4). X-Ray, nuclear magnetic resonance (NMR), and  spectroscopic investigations have firmly established that the nucleic acid bases  are overwhelmingly in the keto tautomeric forms shown in Fig. 3-3. In 1953,  however, this was not generally appreciated. Information about the dominant  tautomeric forms was provided by Jerry Donohue, an office mate of Watson  and Crick and an expert on the X-ray structures of small organic molecules.   Evidence that DNA is a helical molecule was provided by an X-ray dif¬  fraction photograph of a DNA fiber taken by Rosalind Franklin (Fig. 3-5).  The appearance of the photograph enabled Crick, an X-ray crystallographer  by training, to deduce (a) that DNA is a helical molecule and (b) that its pla¬  nar aromatic bases form a stack that is parallel to the fiber axis.    FIG. 3-5 An X-ray diffraction photograph  of a vertically oriented DNA fiber. This photo¬  graph, taken by Rosalind Franklin, provided key  evidence for the elucidation of the Watson-Crick  structure. The central X-shaped pattern indicates  a helix, whereas the heavy black arcs at the top  and bottom of the diffraction pattern reveal the  spacing of the stacked bases (3.4 A). [Courtesy  of Maurice Wilkins, King’s College, London.]   FIG. 3-6 Three-dimensional structure of  DNA. The repeating helix is based on the  structure of the self-complementary dodecamer  d(CGCGAATTCGCG) determined by Richard  Dickerson and Horace Drew. The view in this  ball-and-stick model is perpendicular to the helix  axis. The sugar-phosphate backbones (blue, with  green ribbon outlines ) wind around the periphery  of the molecule. The bases (red) form  hydrogen-bonded pairs that occupy the core. H  atoms have been omitted for clarity. The two  strands run in opposite directions. [Illustration,  Irving Geis. Image from the Irving Geis  Collection/Howard Hughes Medical Institute.  Rights owned by HHMI. Reproduction by  permission only.] O See Interactive Exercise 1  and Kinemage Exercise 2-1.    FIG. 3-7 Diagrams of left- and right-  handed helices. In each case, the fingers curl  in the direction the helix turns when the thumb  points in the direction the helix rises. Note that  the handedness is retained when the helices are  turned upside down.   The limited structural information, along with Chargaff’s rules, provided  few clues to the structure of DNA; Watson and Cricks model sprang mostly  from their imaginations and model-building studies. Once the Watson-Crick  model had been published, however, its basic simplicity combined with its ob¬  vious biological relevance led to its rapid acceptance. Later investigations have  confirmed the general validity of the Watson-Crick model, although its de¬  tails have been modified.   1. Two polynucleotide chains wind around a common axis to form a dou¬  ble helix (Fig. 3-6).   2. The two strands of DNA are antiparallel (run in opposite directions),  but each forms a right-handed helix. (The difference between a right-  handed and a left-handed helix is shown in Fig. 3-7.)   FIG. 3-8 Complementary base pairing in DNA. Adenine in one strand pairs with  thymine in the other strand by forming specific hydrogen bonds (dashed lines). Similarly,  guanine pairs with cytosine. This base pairing between polynucleotide chains is responsible  for the double-stranded nature of DNA. Q See Kinemage Exercise 2-2.   Indicate the 5'—>3' sequence of the strand that would be complementary to the  nucleic acid shown in Fig. 3-3a.    3. The bases occupy the core of the helix and sugar—phosphate chains run  along the periphery, thereby minimizing the repulsions between  charged phosphate groups. The surface of the double helix contains  two grooves of unequal width: the major and minor grooves.   4. Each base is hydrogen bonded to a base in the opposite strand to form  a planar base pair. The Watson—Crick structure can accommodate only  two types of base pairs. Each adenine residue must pair with a thymine  residue and vice versa, and each guanine residue must pair with a cy¬  tosine residue and vice versa (Fig. 3-8). These hydrogen-bonding in¬  teractions, a phenomenon known as complementary base pairing,  result in the specific association of the two chains of the double helix.   The Watson—Crick structure can accommodate any sequence of bases on  one polynucleotide strand if the opposite strand has the complementary base  sequence. This immediately accounts for Chargaff s rules. More importantly,  it suggests that each DNA strand can act as a template for the synthesis of its  complementary strand and hence that hereditary information is encoded in the  sequence of bases on either strand.   Most DNA Molecules Are Large. The extremely large size of DNA molecules  is in keeping with their role as the repository of a cells genetic information.  Of course, an organisms genome, its unique DNA content, may be allocated  among several chromosomes (Greek: chromos , color + soma, body), each of  which contains a separate DNA molecule. Note that many organisms are  diploid; that is, they contain two equivalent sets of chromosomes, one from  each parent. Their content of unique (haploid) DNA is half their total DNA.  For example, humans are diploid organisms that carry 46 chromosomes per  cell; their haploid number is therefore 23.   Because of their great lengths, DNA molecules are described in terms of  the number of base pairs (bp) or thousands of base pairs (kilobase pairs, or  kb). Naturally occurring DNAs vary in length from ~5 kb in small DNA-  containing viruses to well over 250,000 kb in the largest mammalian chromo¬  somes. Although DNA molecules are long and relatively stiff, they are not  completely rigid. We will see in Chapter 24 that the DNA double helix forms   FIG. 3-9 Formation of a stem-loop structure. Base pairing between complementary  sequences within an RNA strand allows the polynucleotide to fold back on itself.   • Using Fig. 3-3a as a guide, draw the com¬  plete structure of a nucleoside triphosphate  before and after it becomes incorporated  into a polynucleotide chain. Draw the struc¬  ture that would result if the newly formed  phosphodiester bond were hydrolyzed.   • Using a three-dimensional computer model  of the DNA molecule (such as shown in  Interactive Exercise 1), identify each of the  following structural features: the 3' and 5'  end of each strand, the atoms that make  up the sugar-phosphate backbone, the  major and minor grooves, the bases in  several base pairs, and the atoms that  participate in hydrogen bonding in A-T  and G-C base pairs.    coils and loops when it is packaged inside the cell. Furthermore, depending  on the nucleotide sequence, DNA may adopt slightly different helical confor¬  mations. Finally, in the presence of other cellular components, the DNA may  bend sharply or the two strands may partially unwind.   Single-stranded DNA is rare, occurring mainly as the hereditary material of  certain viruses. In contrast, RNA occurs primarily as single strands, which usu¬  ally form compact structures rather than loose extended chains (double-  stranded RNA is the hereditary material of certain viruses). An RNA  strand—which is identical to a DNA strand except for the presence of 2'-OH  groups and the substitution of uracil for thymine—can base-pair with a com¬  plementary strand of RNA or DNA. As expected, A pairs with U (or T in  DNA), and G with C. Base pairing often occurs intramolecularly, giving rise  to stem-loop structures (Fig. 3-9) or, when loops interact with each other, to  more complex structures.   The intricate structures that can potentially be adopted by single-stranded  RNA molecules provide additional evidence that RNA can do more than just  store and transmit genetic information. Numerous investigations have found  that certain RNA molecules can specifically bind small organic molecules and  can catalyze reactions involving those molecules. These findings provide sub¬  stantial support for theories that many of the processes essential for life began  through the chemical versatility of small polynucleotides (a situation known as the  RNA world). We will further explore RNA structure and function in Section  24-2C.    • DNA carries genetic information in the form of its sequence of nucleotides.   • The nucleotide sequence of DNA is transcribed into the nucleotide sequence of  messenger RNA, which is then translated into a protein, a sequence of amino acids.    DNA is the carrier of genetic information in all cells and in many viruses. Yet  a period of over 75 years passed from the time the laws of inheritance were  discovered by Gregor Mendel until the biological role of DNA was elucidated.  Even now, many details of how genetic information is expressed and transmit¬  ted to future generations remain unclear.   large colonies are virulent pneumococci that  resulted from the transformation of  nonpathogenic pneumococci (smaller colonies)  by DNA extracted from the virulent strain. We  now know that this DNA contained a gene that  was defective in the nonpathogenic strain. [From  Avery, O.T., MacLeod, C.M., and McCarty, M., J.  Exp. Med. 79, 153 (1944). Copyright © 1944 by  Rockefeller University Press.]    Mendels work with garden peas led him to postulate that an individual  plant contains a pair of factors (which we now call genes), one inherited from  each parent. But Mendels theory of inheritance, reported in 1866, was almost  universally ignored by his contemporaries, whose knowledge of anatomy and  physiology provided no basis for its understanding. Eventually, genes were hy¬  pothesized to be part of chromosomes, and the pace of genetic research greatly  accelerated.   Until the 1940s, it was generally assumed that genes were made of protein,  since proteins were the only biochemical entities that, at the time, seemed  complex enough to serve as agents of inheritance. Nucleic acids, which had  first been isolated in 1869 by Friedrich Miescher, were believed to have mo¬  notonously repeating nucleotide sequences and were therefore unlikely candi¬  dates for transmitting genetic information.   It took the efforts of Oswald Avery, Colin MacLeod, and Maclyn McCarty  to demonstrate that DNA carries genetic information. Their experiments, com¬  pleted in 1944, showed that DNA—not protein—extracted from a virulent  (pathogenic) strain of the bacterium Diplococcus pneumoniae was the substance  that transformed (permanently changed) a nonpathogenic strain of the organ¬  ism to the virulent strain (Fig. 3-10). Avery’s discovery was initially greeted  with skepticism, but it influenced Erwin Chargaff, whose rules (Section 3-2A)  led to subsequent models of the structure and function of DNA.   The double-stranded, or duplex, nature of DNA facilitates its replication.  When a cell divides, each DNA strand acts as a template for the assembly of  its complementary strand (Fig. 3-11). Consequently, every progeny cell con¬  tains a complete DNA molecule (or a complete set of DNA molecules in or¬  ganisms whose genomes contain more than one chromosome). Each DNA  molecule consists of one parental strand and one daughter strand. Daughter  strands are synthesized by the stepwise polymerization of nucleotides that  specifically pair with bases on the parental strands. The mechanism of repli¬  cation, while straightforward in principle, is exceedingly complex in the cell,  requiring a multitude of cellular factors to proceed with fidelity and efficiency,  as we will see in Chapter 25.   The question of how sequences of nucleotides control the characteristics of or¬  ganisms took some time to be answered. In experiments with the mold  Neurospora crassa in the 1940s, George Beadle and Edward Tatum found that  there is a specific connection between genes and enzymes , the one gene—one enzyme  theory Beadle and Tatum showed that mutant varieties of Neurospora that were    FIG. 3-1 1 DNA replication. Each strand of  parental DNA (blue) acts as a template for the  synthesis of a complementary daughter strand  (red). Thus, the resulting double-stranded  molecules are identical.   One strand of DNA directs the synthesis of  messenger RNA (mRNA). The base sequence of  the transcribed RNA is complementary to that of  the DNA strand. The message is translated  when transfer RNA (tRNA) molecules align with  the mRNA by complementary base pairing  between three-nucleotide segments known as  codons. Each tRNA carries a specific amino  acid. These amino acids are covalently joined to  form a protein. Thus, the sequence of bases in  DNA specifies the sequence of amino acids in a  protein.    What might happen if a mutation  changed one of the nucleotides in the    generated by irradiation with X-rays required additional nutrients in order to  grow. Presumably, the offspring of the radiation-damaged cells lacked the spe¬  cific enzymes necessary to synthesize those nutrients.   The link between DNA and enzymes (nearly all of which are proteins) is  RNA. The DNA of a gene is transcribed to produce an RNA molecule that is  complementary to the DNA. The RNA sequence is then translated into the cor¬  responding sequence of amino acids to form a protein (Fig. 3-12). These trans¬  fers of biological information are summarized in the so-called central dogma  of molecular biology formulated by Crick in 1958.   In this diagram, arrows represent information transfer when DNA directs its  own replication to produce new DNA molecules; when DNA is transcribed  into RNA; and when RNA is translated into protein.   Just as the daughter strands of DNA are synthesized from free deoxynu-  cleoside triphosphates that pair with bases in the parent DNA strand, RNA  strands are synthesized from free ribonucleoside triphosphates that pair with  the complementary bases in one DNA strand of a gene (transcription is de¬  scribed in greater detail in Chapter 26). The RNA that corresponds to a  protein-coding gene (called messenger RNA, or mRNA) makes its way to a  ribosome, an organelle that is itself composed largely of RNA (ribosomal  RNA, or rRNA). At the ribosome, each set of three nucleotides in the mRNA  pairs with three complementary nucleotides in a small RNA molecule called    FIG. 3-13 Translation. tRNA molecules  with their attached amino acids bind to  complementary three-nucleotide sequences  (codons) on mRNA. The ribosome facilitates the  alignment of the tRNA and the mRNA, and it  catalyzes the joining of amino acids to produce a  protein chain. When a new amino acid is added,  the preceding tRNA is ejected, and the ribosome  proceeds along the mRNA.    a transfer RNA, or tRNA (Fig. 3-13). Attached to each tRNA molecule is its  corresponding amino acid. The ribosome catalyzes the joining of amino acids,  which are the monomeric units of proteins (protein synthesis is described in  detail in Chapter 27). Amino acids are added to the growing protein chain  according to the order in which the tRNA molecules bind to the mRNA. Since  the nucleotide sequence of the mRNA in turn reflects the sequences of nu¬  cleotides in the gene, DNA directs the synthesis of proteins. It follows that al¬  terations to the genetic material of an organism (mutations) may manifest  themselves as proteins with altered structures and functions.   Using techniques that are described in the following sections and in other  parts of this book, researchers can compile a catalog of all the information en¬  coded in an organisms DNA. The study of the genomes size, organization,  and gene content is known as genomics. By analogy, transcriptomics refers  to the study of gene expression, which focuses on the set of mRNA molecules,  or transcriptome, that is transcribed from DNA under any particular set of  circumstances. Finally, proteomics is the study of the proteins (the proteome)  produced as a result of transcription and translation. Although an organisms  genome remains essentially unchanged throughout its lifetime, its transcrip¬  tome and proteome may vary significantly among different types of tissues,  developmental stages, and environmental conditions.   • Explain why the double-stranded nature of  DNA is relevant for copying and transmit¬  ting genetic information when a cell divides.   • Summarize the steps of the central dogma.  What role does RNA play in each?    • In the laboratory, nucleic acids can be cut at specific sequences by restriction  enzymes.   • In the chain-termination method, DNA polymerase generates DNA fragments that  are randomly terminated. The identities of the terminator nucleotides of successive  fragments reveal the original DNA sequence.   • The human genome contains —23,000 genes, corresponding to about 1.2% of its  3 billion nucleotides.    Much of our current understanding of protein structure and function rests  squarely on information gleaned not from the proteins themselves, but indi¬  rectly from their genes. The ability to determine the sequence of nucleotides in   FIG. 3-14 Determining the sequence of  an oligonucleotide using nonspecific  enzymes. The oligonucleotide is partially  digested with snake venom phosphodiesterase,  which breaks the phosphodiester bonds between  nucleotide residues, starting at the 3' end of the  oligonucleotide. The result is a mixture of  fragments of all lengths, which are then  separated. Comparing the base composition of a  pair of fragments that differ in length by one  nucleotide establishes the identity of the  3'-terminal nucleotide in the larger fragment.  Analysis of each pair of fragments reveals the  sequence of the original oligonucleotide.    nucleic acids has made it possible to deduce the amino acid sequences of their en¬  coded proteins and, to some extent, the structures and functions of those proteins.  Nucleic acid sequencing has also revealed information about the regulation of genes.  Certain portions of genes that are not actually transcribed into RNA neverthe¬  less influence how often a gene is transcribed and translated, that is, expressed.  Moreover, efforts to elucidate the sequences in hitherto unmapped regions of  DNA have led to the discovery of new genes and new regulatory elements.  Once in hand, a nucleic acid sequence can be duplicated, modified, and expressed,  making it possible to study proteins that could not otherwise be obtained in useful  quantities. In this section, we describe how nucleic acids are sequenced and  what information the sequences may reveal. In the following section, we dis¬  cuss the manipulation of purified nucleic acid sequences for various purposes.   The overall strategy for sequencing any polymer of nonidentical units is   1. Cleave the polymer into fragments that are small enough to be fully  sequenced.   3. Determine the order of the fragments in the original polymer by align¬  ing fragments that contain overlapping sequences.   The first efforts to sequence RNA used nonspecific enzymes to generate rela¬  tively small fragments whose nucleotide composition was then determined by  partial digestion with an enzyme that selectively removed nucleotides from one  end or the other (Fig. 3-14). Sequencing RNA in this manner was tedious  and time-consuming. Using such methods, it took Robert Holley 7 years to  determine the sequence of a 76-residue tRNA molecule.   After 1975, dramatic progress was made in nucleic acid sequencing tech¬  nology. The advances were made possible by the discovery of enzymes that  could cleave DNA at specific sites and by the development of rapid sequenc¬  ing techniques for DNA. Because most specific DNA sequences are normally  present in a genome in only a single copy, most sequencing projects take ad¬  vantage of methods to amplify segments of DNA by cloning or copying them  (Section 3-5).   Many bacteria are able to resist infection by bacteriophages (viruses that are  specific for bacteria) by virtue of a restriction-modification system. The bac¬  terium modifies certain nucleotides in specific sequences of its own DNA by  adding a methyl (—CH 3 ) group in a reaction catalyzed by a modification  methylase. A restriction endonuclease, which recognizes the same nucleotide  sequence as does the methylase, cleaves any DNA that has not been modified  on at least one of its two strands. (An endonuclease cleaves a nucleic acid  within the polynucleotide strand; an exonuclease cleaves a nucleic acid by re¬  moving one of its terminal residues.) This system destroys foreign (phage)  DNA containing a recognition site that has not been modified by methyla-  tion. The host DNA is always at least half methylated, because although the  daughter strand is not methylated until shortly after it is synthesized, the  parental strand to which it is paired is already modified (and thus protects  both strands of the DNA from cleavage by the restriction enzyme).   Type II restriction endonucleases are particularly useful in the laboratory.  These enzymes cleave DNA within the four- to eight-base sequence that is  recognized by their corresponding modification methylase. (Type I and Type  III restriction endonucleases cleave DNA at sites other than their recognition  sequences.) Nearly 4000 Type II restriction enzymes with over 270 different  recognition sequences have been characterized. Some of the more widely used  restriction enzymes are listed in Table 3-2. A restriction enzyme is named by  the first letter of the genus and the first two letters of the species of the bac¬  terium that produced it, followed by its serotype or strain designation, if any,    The recognition sequence is abbreviated so that only one strand, reading 5' to 3', is given. The  cleavage site is represented by an arrow (4). R and Y represent a purine nucleotide and a pyrimi¬  dine nucleotide, respectively.   Source: Roberts, R.J. and Macelis, D., REBASE—the restriction enzyme database,  http://rebase.neb.com.    and a roman numeral if the bacterium contains more than one type of restric¬  tion enzyme. For example, EcoRI is produced by E. coli strain RY13.   Interestingly, most Type II restriction endonucleases recognize and cleave  palindromic DNA sequences. A palindrome is a word or phrase that reads the  same forward or backward. Two examples are “refer” and “Madam, I’m Adam.”  In a palindromic DNA segment, the sequence of nucleotides is the same in  each strand, and the segment is said to have twofold symmetry (Fig. 3-15).  Most restriction enzymes cleave the two strands of DNA at positions that are  staggered, producing DNA fragments with complementary single-strand exten¬  sions. Restriction fragments with such sticky ends can associate by base  pairing with other restriction fragments generated by the same restriction   recognition sequences for Type II restriction  endonucleases are palindromes, sequences with  a twofold axis of symmetry, (a) Recognition site  for EcoRI, which generates DNA fragments with  sticky ends, (b) Recognition site for EcoRV,  which generates blunt-ended fragments.    FIG. 3-16 Apparatus for gel electrophoresis. Samples are applied in slots at the  top of the gel and electrophoresed in parallel lanes. Negatively charged molecules such  as DNA migrate through the gel matrix toward the anode in response to an applied  electric field. Because smaller molecules move faster, the molecules in each lane are  separated according to size. Following electrophoresis, the separated molecules may be  visualized by staining or fluorescence.    enzyme. Some restriction endonucleases cleave the two strands of DNA at the  symmetry axis to yield restriction fragments with fully base-paired blunt ends.   Treating a DNA molecule with a restriction endonuclease produces a series of  precisely defined fragments that can be separated according to size. Gel elec¬  trophoresis is commonly used for the separation. In principle, a charged mol¬  ecule moves in an electric field with a velocity proportional to its overall charge  density, size, and shape. For molecules with a relatively homogeneous compo¬  sition (such as nucleic acids), shape and charge density are constant, so the  velocity depends primarily on size. Electrophoresis is carried out in a gel-like  matrix, usually made from agarose (carbohydrate polymers that form a loose  mesh) or polyacrylamide (a more rigid cross-linked synthetic polymer). The  gel is typically held between two glass plates (Fig. 3-16) or inside a narrow  capillary tube. The molecules to be separated are applied to one end of the  ABCDE FGH I gel, and the molecules move through the pores in the matrix under the influ¬  ence of an electric field. Smaller molecules move more rapidly through the gel  and therefore migrate farther in a given time.   Following electrophoresis, the separated molecules may be visualized in  the gel by an appropriate technique, such as addition of a stain that binds  tightly to the DNA, by radioactive labeling, or by their fluorescence.  Depending on the dimensions of the gel and the visualization technique used,  samples containing less than a nanogram of material can be separated and de¬  tected by gel electrophoresis. Several samples can be electrophoresed simulta¬  neously. For example, the fragments obtained by digesting a DNA sample with  different restriction endonucleases can be visualized side by side (Fig. 3-17).    FIG. 3-17 Electrophoretogram of restriction digests. The plasmid pAgK84 has  been digested with (A) BamHI, (B) Pstl, (C) Bglll, (D) Haelll, (E) Hindi, (F) Sacl, (G) Xbal,  and (H) Hpal. Lane I contains bacteriophage A digested with Hindi 11 as a standard since  these fragments have known sizes. The restriction fragments in each lane are made visible  by fluorescence against a black background. [From Slota, J.E. and Farrand, S.F., Plasmid 8 ,  180 (1982). Copyright © 1982 by Academic Press.]   The sizes of the various fragments can be determined by comparing their elec- Section 4 Nucleic Acid Sequencing   Until recent years, the most widely used technique for sequencing DNA was  the chain-terminator method, which was devised by Frederick Sanger. The  first step in this procedure is to obtain single polynucleotide strands.  Complementary DNA strands can be separated by heating, which breaks the  hydrogen bonds between bases. Next, polynucleotide fragments that terminate  at positions corresponding to each of the four nucleotides are generated.  Finally, the fragments are separated and detected.    DNA Polymerase Copies a Template Strand. The chain-terminator  method (also called the dideoxy method) uses an E. coli enzyme to make  complementary copies of the single-stranded DNA being sequenced. The en¬  zyme is a fragment of DNA polymerase I, one of the enzymes that partic¬  ipates in replication of bacterial DNA (Section 25-2A). Using the single  DNA strand as a template, DNA polymerase I assembles the four deoxynu-  cleoside triphosphates (dNTPs), dATP, dCTP, dGTP, and dTTP, into a com¬  plementary polynucleotide chain that it elongates in the 5' —> 3' direction  (Fig. 3-18).   DNA polymerase I can sequentially add deoxynucleotides only to the 3'  end of a polynucleotide. Hence, replication is initiated in the presence of a  short polynucleotide (a primer) that is complementary to the 3' end of the  template DNA and thus becomes the 5' end of the new strand. The primer  base-pairs with the template strand, and nucleotides are sequentially added to  the 3' end of the primer. If the DNA being sequenced is a restriction frag¬  ment, as it usually is, it begins and ends with a restriction site. The primer  can therefore be a short DNA segment with the sequence of this restriction  site.   FIG. 3-1 8 Action of DNA polymerase I. Using a single DNA  strand as a template, the enzyme elongates the primer by stepwise  addition of complementary nucleotides. Incoming nucleotides pair with  bases on the template strand and are joined to the growing    polynucleotide strand in the 5' —> 3' direction. The polymerase-  catalyzed reaction requires a free 3'-0H group on the growing strand.  Pyrophosphate (P 2 O 7 - ; PP,) is released with each nucleotide  addition.    DNA Synthesis Terminates after Specific Bases. In the chain-terminator  technique (Fig. 3-19), the DNA to be sequenced is incubated with DNA poly¬  merase I, a suitable primer, and the four dNTP substrates (reactants in enzy¬  matic reactions) for the polymerization reaction. The key component of the  reaction mixture is a small amount of a 2',3'-dideoxynucleoside triphos¬  phate (ddNTP),   which lacks the 3'-OH group of deoxynucleotides. When the dideoxy analog is  incorporated into the growing polynucleotide in place of the corresponding normal  nucleotide, chain growth is terminated because addition of the next nucleotide re¬  quires a free 3'-OH.   By using only a small amount of the ddNTP, a series of truncated chains is  generated, each of which ends with the dideoxy analog at one of the positions  occupied by the corresponding base. Each ddNTP bears a different fluorescent  “tag” so that the products of the polymerase reaction can be readily detected. Gel  electrophoresis separates the newly synthesized DNA segments, which differ in  size by one nucleotide. Thus, the sequence of the replicated strand can be di¬  rectly read from the gel. Note that the sequence obtained by the chain-termi¬  nator method is complementary to the DNA strand being sequenced.   The most advanced sequencing devices that employ the chain-terminator  method identify each DNA fragment as it exits the bottom of a capillary  electrophoresis tube, so the sequence data take the form of a series of peaks    dATP + dCTP + dGTP + dTTP +  ddATP—O + ddCTP—O +  ddGTP—O + ddTTP— •    FIG. 3-19 Automated DNA sequencing  by the chain-terminator method. The reaction  mixture includes the single-stranded DNA to be  sequenced (the template), a primer, the four  deoxynucleotide triphosphates (represented as  dATP, etc.), and small amounts of the four fluo-  rescently labeled dideoxynucleoside  triphosphates (ddATP, etc.). DNA polymerase  extends the primer until a chain-terminating  dideoxy nucleotide is added, generating a set of  DNA fragments that differ by one nucleotide.  The mixture is subjected to gel electrophoresis  in a capillary tube, which separates the  fragments according to size. As each polynu¬  cleotide passes the detector, its 3'-terminal  nucleotide is identified according to its laser-  stimulated fluorescence.    Dye-labeled DNA segments are  subjected to electrophoresis  in a capillary gel    AGTTCTAG AGCGGCCGCC AC CGCGGTGGNAGC TCC AGCT T TTGT TCCCTT T AGT GAGGGTT AAT TTCGAGCTTGGCGT A AT CAT GGTCAT AGCTGT T T  0 110 120 130 140 150 160 4 170 180 190   ICTGTGTGAAATTGTT ATCCGCTCAC AAT TCC ACACAACAT ACG AGC CGG A AG CAT A AAGT GT AAAGC C TGG GG TGC CT A AT G A GTG AG CTj  200 . 210 220 A 230 . 240 . 250 260 270 280    FIG. 3-20 DNA sequence data. Each of the four colored curves  represents the electrophoretic pattern of fragments containing one of  the dideoxynucleotides: Green, red, black, and blue peaks correspond  to fragments ending in ddATP, ddTTP, ddGTP, and ddCTP, respectively.  The 3'-terminal base of each oligonucleotide, identified by the    fluorescence of its gel band, is indicated by a single letter (A, T, G, or  C). This portion of the readout corresponds to nucleotides 100-290  of the DNA segment being sequenced. [Courtesy of Mark Adams, The  Institute for Genomic Research, Rockville, Maryland.]    (Fig. 3-20). Sample preparation and data analysis are fully automated, and  sequences of DNA up to 1000 nucleotides can be obtained from a single re¬  action mixture. Moreover, these systems contain arrays of 96 capillary tubes  and hence can simultaneously sequence 96 DNA segments.   Newer Sequencing Technologies Use Light or Voltage Changes. The value  of DNA sequence information, particularly the enormous datasets obtained by  sequencing entire genomes, has driven the development of novel sequencing  technologies that offer various trade-offs among cost, speed, and accuracy. Like  the older chain-terminator procedure, newer methods take advantage of the  ability of DNA polymers to produce a complementary copy of a template  DNA strand.   In pyrosequencing, molecules of the template DNA are immobilized on  the surfaces of microscopic plastic beads that are deposited in small wells in a  fiber-optic slide with one bead per well. A primer and DNA polymerase are  added, and then a dNTP substrate is introduced. If DNA polymerase adds  that nucleotide to the new DNA strand, pyrophosphate is released and trig¬  gers a chemical reaction involving the firefly enzyme luciferase, which gener¬  ates a flash of light. Solutions of each of the four dNTPs are successively  washed across the immobilized DNA template, and a detector records whether  light is produced in the presence of a particular dNTP. In this way, the se¬  quence of nucleotides complementary to the template strand can be deduced,  and no electrophoretic separation is needed. Pyrosequencing can accurately  “read” stretches of 300—500 nucleotides, somewhat shorter than the sequences  revealed by dideoxy sequencing. The fiber-optic slides contain numerous wells  so that as many as ~400,000 templates can be sequenced simultaneously.  Consequently, the pyrosequencing system is ~300-fold faster than the most  advanced dideoxy sequencing systems.   Other sequencing instruments detect the proton generated on pyrophos¬  phate release, so that a DNA sequence can be deduced from the minute change  in voltage when the correct nucleotide (the one matching the template at that  point) is incorporated into the new chain. All of these devices, like the pyrose¬  quencing system, sequence large numbers of DNA fragments simultaneously  and hence determine the identities of billions of nucleotides in a single run.   Databases Store Nucleotide Sequences. The results of sequencing projects  large and small are customarily deposited in online databases such as GenBank  (see Bioinformatics Project 1). Approximately 300 billion nucleotides represent¬  ing over 200 million sequences have been recorded as of late 2010.    Francis S. Collins (1950-) By the mid¬  twentieth century, the molecular bases of  several human diseases were appreciated. For  example, sickle-cell anemia (Section 7-1 E)  was known to be caused by an abnormal  hemoglobin protein. Studies of sickle-cell  hemoglobin eventually revealed the underlying  genetic defect, a mutation in a hemoglobin  gene. It therefore seemed possible to trace  other diseases to defective genes. But for  many genetic diseases, even those with well-characterized  symptoms, no defective protein had yet been identified. One such  disease was cystic fibrosis, which is characterized mainly by the  secretion of thick mucus that obstructs the airways and creates  an ideal environment for bacterial growth. Cystic fibrosis is the  most common inherited disease in individuals of northern  European descent, striking about 1 in 2500 newborns and leading  to death by early adulthood due to irreversible lung damage. It  was believed that identifying the molecular defect in cystic fibrosis  would lead to better understanding of the disease and to the  ability to design more effective treatments.   Enter Francis Collins, who began his career by earning a  doctorate in physical chemistry but then enrolled in medical school  to take part in the molecular biology revolution. As a physician-  scientist, Collins developed methods for analyzing large stretches  of DNA in order to home in on specific genes, including the one  that, when mutated, causes cystic fibrosis. By analyzing the DNA  of individuals with the disease (who had two copies of the  defective gene) and of family members who were asymptomatic  carriers (with one normal and one defective copy of the gene),  Collins and his team localized the cystic fibrosis gene to the long    arm of chromosome 7. They gradually closed in on a DNA  segment that appears to be present in a number of mammalian  species, which suggests that the segment contains an essential  gene. The cystic fibrosis gene was finally identified in 1989.  Collins had demonstrated the feasibility of identifying a genetic  defect in the absence of other molecular information.   Once the cystic fibrosis gene was in hand, it was a relatively  straightforward process to deduce the probable structure and  function of the encoded protein, which turned out to be a  membrane channel for chloride ions. When functioning normally,  the protein helps regulate the ionic composition and viscosity of  extracellular secretions. Discovery of the cystic fibrosis gene also  made it possible to design tests to identify carriers so that they  could take advantage of genetic counseling.   Throughout Collins’ work on the cystic fibrosis gene and during  subsequent hunts for the genes that cause neurofibromatosis and  Huntington’s disease, he was mindful of the ethical implications of  the new science of molecular genetics. Collins has been a strong  advocate for protecting the privacy of genetic information. At the  same time, he recognizes the potential therapeutic use of such  information. In his tenure as director of the human genome  project, he was committed to making the results freely and  immediately accessible, as a service to researchers and the  individuals who might benefit from new therapies based on  molecular genetics. He is presently the director of the National  Institutes of Health (NIH).   Riordan, J.R., Rommens, J.M., Kerem, B.-S., Alon, N., Rozmahel, R., Grzelczak, Z.,  Zielensky, J., Lok, S., Plavsic, N., Chou, J.-L, Drumm, M.L, lannuzzi, M.C., Collins, F.S.,  and Tsui, L-C., Identification of the cystic fibrosis gene: Cloning and characterization  of complementary DNA, Science 245 , 1066-1073 (1989).   Nucleic acid sequencing has become so routine that directly determining  a proteins amino acid sequence (Section 5-3) is generally far more time-  consuming than determining the base sequence of its corresponding gene. In  fact, nucleic acid sequencing is invaluable for studying genes whose products  have not yet been identified. If the gene can be sequenced, the probable func¬  tion of its protein product may be deduced by comparing the base sequence  to those of genes whose products are already characterized (see Box 3-1).   The advent of large-scale sequencing techniques brought to fruition the dream  of sequencing entire genomes. However, the major technical hurdle in se¬  quencing all the DNA in an organisms genome is not the DNA sequencing  itself but, rather, assembling the tens of thousands to tens of millions of se¬  quenced segments (depending on the size of the genome) into contiguous  blocks and assigning them to their correct chromosomal positions. To do so  required the development of automated sequencing protocols and mathemat¬  ically sophisticated computer algorithms.   The first complete genome sequence to be determined, that of the bacterium  Haemophilus influenzae, was reported in 1995 by Craig Venter. By mid-2011,  the complete genome sequences of over 1600 prokaryotes had been reported  (with many more being determined) as well as over 140 eukaryotes, including  humans, human pathogens, plants, and laboratory organisms (Table 3-3).    In metagenomic sequencing, the DNA sequences of multiple organisms  are analyzed as a single data set. This approach is used to characterize com¬  plex microbial communities, such as those in marine environments, where in¬  dividual species—including many not yet identified—cannot be cultured and  sequenced one by one. Metagenomic sequence data reveal the overall gene  number and an estimate of the collective metabolic capabilities of the com¬  munity. Over 3 million genes have been identified in a metagenomic analysis  of the microorganisms that inhabit the human gut, representing some 1000  bacterial species. While most humans share a common core set of about 60  gut microorganisms, significant differences appear to correlate with metabolic  variables such as body mass.   The Human Genome Contains Relatively Few Genes. The determination  of the 3-billion-nucleotide human genome sequence was a gargantuan under¬  taking involving hundreds of scientists working in two groups, one led by  Venter and the other by Francis Collins (Box 3-1), Eric Lander, and John  Sulston. After over a decade of intense effort, the “rough draft” of the human  genome sequence was reported in early 2001 and the “finished” sequence, cov¬  ering —99% of the genome, was reported in 2004. This stunning achievement  promises to revolutionize the way both biochemistry and medicine are viewed  and practiced, although it is likely to require many years of further effort    before its full significance is understood. Nevertheless, numerous important  conclusions can already be drawn, including these:   1. About half the human genome consists of repeating sequences of  various types.   2. Up to 80% of the genome may be transcribed to RNA.   4. The human genome appears to contain only —23,000 protein-encoding  genes [also known as open reading frames (ORFs)] rather than the  50,000 to 140,000 ORFs that had previously been predicted. This com¬  pares with the —6000 ORFs in yeast, —13,000 in Drosophila. , —19,000  in C. elegans, and —26,000 in Arabidopsis (although these numbers will  almost certainly change as our ability to recognize ORFs improves).   5. Only a small fraction of human proteins are unique to vertebrates;  most occur in other if not all life-forms.   6. Two randomly selected human genomes differ, on average, by only 1  nucleotide per 1000; that is, any two people are likely to be >99.9%  genetically identical.   The obviously greater complexity of humans (vertebrates) relative to inverte¬  brate forms of life is unlikely to be due to the not-much-larger numbers of  ORFs that vertebrates encode. Rather, it appears that vertebrate proteins them¬  selves are more complex than those of invertebrates; that is, vertebrate proteins  tend to have more domains (modules) than invertebrate proteins, and these  modules are more often selectively expressed through alternative gene splic¬  ing (a phenomenon in which a given gene transcript can be processed in  multiple ways so as to yield different proteins when translated; Section 26-3B).  In fact, most vertebrate genes encode several different although similar proteins.   One of the richest rewards of nucleic acid sequencing technology is the infor¬  mation it provides about the mechanisms of evolution. The chemical and  physical properties of DNA, such as its regular three-dimensional shape and  the elegant process of replication, may leave the impression that genetic infor¬  mation is relatively static. In fact, DNA is a dynamic molecule , subject to changes  that alter genetic information. For example, the mispairing of bases during  DNA replication can introduce errors known as point mutations in the  daughter strand. Mutations also result from DNA damage by chemicals or ra¬  diation. More extensive alterations in genetic information are caused by faulty  recombination (exchange of DNA between chromosomes) and the transpo¬  sition of genes within or between chromosomes and, in some cases, from one  organism to another. All these alterations to DNA provide the raw material  for natural selection. When a mutated gene is transcribed and the messenger  RNA is subsequently translated, the resulting protein may have properties that  confer some advantage to the individual. As a beneficial change is passed from  generation to generation, it may become part of the standard genetic makeup  of the species. Of course, many changes occur as a species evolves, not all of  them simple and not all of them gradual.   Phylogenetic relationships can be revealed by comparing the sequences of  similar genes in different organisms. The number of nucleotide differences be¬  tween the corresponding genes in two species roughly indicates the degree to  which the species have diverged through evolution. The regrouping of prokary¬  otes into archaea and bacteria (Section 1-2C) according to rRNA sequences  present in all organisms illustrates the impact of sequence analysis.   Nucleic acid sequencing also reveals that species differing in phenotype  (physical characteristics) are nonetheless remarkably similar at the molecular  level. For example, humans and chimpanzees share nearly 99% of their DNA.   FIG. 3-21 Maize and teosinte. Despite the  large differences in phenotype—maize ( bottom)  has hundreds of easily chewed kernels whereas  teosinte (top) has only a few hard, inedible  kernels—the plants differ in only a few genes.   The ancestor of maize is believed to be a mutant  form of teosinte in which the kernels were more  exposed, [(top) Courtesy John Doebley, University  of Wisconsin; (bottom) Marek Mnich/Stockphoto.]    Studies of corn (maize) and its putative ancestor, teosinte, suggest that the  plants differ in only a handful of genes governing kernel development (teosinte  kernels are encased by an inedible shell; Fig. 3-21).   Small mutations in DNA are apparently responsible for relatively large evo¬  lutionary leaps. This is perhaps not so surprising when the nature of genetic in¬  formation is considered. A mutation in a gene segment that does not encode  protein might interfere with the binding of cellular factors that influence the  timing of transcription. A mutation in a gene encoding an RNA might inter¬  fere with the binding of factors that affect the efficiency of translation. Even a  minor rearrangement of genes could disrupt an entire developmental process,  resulting in the appearance of a novel species. Notwithstanding the high prob¬  ability that most sudden changes would lead to diminished individual fitness or  the inability to reproduce, the capacity for sudden changes in genetic informa¬  tion is consistent with the fossil record. Ironically, the discontinuities in the fos¬  sil record that are probably caused in part by sudden genetic changes once fueled  the adversaries of Charles Darwins theory of evolution by natural selection.   T Sequence Variations Can Be Linked to Human Diseases. Almost 2000  genes have been linked to different human diseases, yet monogenetic dis¬  eases, such as cystic fibrosis (see Box 3-1), are relatively rare. Most diseases re¬  sult from interactions among multiple genes and from environmental factors.  Nevertheless, scientists are hopeful that genomics can lead to a better under¬  standing of how genetic information impacts human health and susceptibility  to illness. One area of success is the screening of individuals who are carriers for  a recessive genetic disorder; that is, they have a normal phenotype but bear one  copy of the defective gene, which may be passed to their children. Clinical tests  are available to detect over a hundred such single-gene defects (Table 3-4).  Preliminary results suggest that as more parents become aware of their carrier  status, fewer children with the disease are being born.   • Explain how restriction enzymes generate  either sticky ends or blunt ends.   • Why do the smallest fragments of DNA  move the farthest during electrophoresis?   • List all the components and explain their  purpose in the reaction mixture used for  the dideoxy DNA sequencing method.   • Summarize what is known about the size  and gene content of the human genome.    Although new sequencing technologies will soon make it feasible to se¬  quence the complete genomes of individuals, for example, members of a fam¬  ily with a poorly characterized genetic disease, most approaches focus on  single-nucleotide polymorphisms (SNPs, instances where the DNA sequence  differs among individuals at one nucleotide). From studies involving thousands  of subjects with and without certain complex diseases such as cancer and type  2 diabetes, researchers have identified a number of SNPs that are associated  with increased risk for these conditions. The SNPs are only proxies for genes  but could provide starting points for further efforts to locate genes or regula¬  tory DNA sequences that might be directly involved in the disease. An ongo¬  ing challenge for this work is that each genetic variant associated with a disease  typically increases risk for the disease by only a few percent, so an individuals  likelihood of developing a particular disease appears to be a complicated func¬  tion of which variants are present. Several commercial enterprises offer indi¬  vidual genome-sequencing services, but until genetic information can be  reliably translated into effective disease-prevention or treatment regimens, the  practical value of “personal genomics” is quite limited.    • Segments of DNA can be cloned, or reproduced, in a host organism.   • A DNA library is a collection of cloned DNA segments that can be screened to  find a particular gene.   • The polymerase chain reaction amplifies a DNA segment by repeatedly synthesizing  complementary strands.   • Recombinant DNA technology can be used to manipulate genes for protein  expression or for the production of transgenic organisms.    Along with nucleic acid sequencing, techniques for manipulating DNA in  vitro and in vivo (in the test tube and in living systems) have produced dra¬  matic advances in biochemistry, cell biology, and genetics. In many cases, this  recombinant DNA technology has made it possible to purify specific DNA  sequences and to prepare them in quantities sufficient for study. Consider the  problem of isolating a unique 1000-bp length of chromosomal DNA from E.  coli. A 10-L culture of cells grown at a density of —10 10 cells • mL 1 con¬  tains only —0.1 mg of the desired DNA, which would be all but impossible  to separate from the rest of the DNA using classical separation techniques  (Sections 5-2 and 24-3). Recombinant DNA technology, also called molecular  cloning or genetic engineering makes it possible to isolate, amplify, and mod¬  ify specific DNA sequences.   The following approach is used to obtain and amplify a segment of DNA:   1. A fragment of DNA of the appropriate size is generated by a restric¬  tion enzyme, by PCR (Section 3-5C), or by chemical synthesis.   2. The fragment is incorporated into another DNA molecule known  as a vector, which contains the sequences necessary to direct DNA  replication.   3. The vector—with the DNA of interest—is introduced into cells, in  which it is replicated.   Cloning refers to the production of multiple identical organisms derived from  a single ancestor. The term clone refers to the collection of cells that contain    FIG. 3-22 The plasmid pUC18. As shown in this diagram, the circular plasmid  contains multiple restriction sites, including a polylinker sequence that contains 13  restriction sites that are not present elsewhere on the plasmid. The three genes  expressed by the plasmid are amp R , which confers resistance to the antibiotic  ampicillin; lacZ, which encodes the enzyme (3-galactosidase; and lad, which  encodes a factor that controls the transcription of lacZ (as described in  Section 28-2A).    the vector carrying the DNA of interest or to the DNA itself.   In a suitable host organism, such as E. coli or yeast, large  amounts of the inserted DNA can be produced.   Cloned DNA can be purified and sequenced (Section 3-  4). Alternatively, if a cloned gene is flanked by the properly  positioned regulatory sequences for RNA and protein synthe¬  sis, the host may also produce large quantities of the RNA and  protein specified by that gene. Thus, cloning provides materials  (nucleic acids and proteins) for other studies and also provides  means for studying gene expression under controlled conditions.    Cloning Vectors Carry Foreign DNA. A variety of small, autonomously repli¬  cating DNA molecules are used as cloning vectors. Plasmids are circular DNA  molecules of 1 to 200 kb found in bacteria or yeast cells. Plasmids can be con¬  sidered molecular parasites, but in many instances they benefit their host by  providing functions, such as resistance to antibiotics, that the host lacks.   Some types of plasmids are present in one or a few copies per cell and  replicate only when the bacterial chromosome replicates. However, the plas¬  mids used for cloning are typically present in hundreds of copies per cell and  can be induced to replicate until the cell contains two or three thousand copies  (representing about half of the cells total DNA). The plasmids that have been  constructed for laboratory use are relatively small, replicate easily, carry genes  specifying resistance to one or more antibiotics, and contain a number of con¬  veniently located restriction endonuclease sites into which foreign DNA can  be inserted. Plasmid vectors can be used to clone DNA segments of no more  than ~10 kb. The E. coli plasmid designated pUC18 (Fig. 3-22) is a repre¬  sentative cloning vector (“pUC” stands for plasmid-Universal Cloning).   Bacteriophage \ (Fig. 3-23) is an alternative cloning vector that can ac¬  commodate DNA inserts up to 16 kb. The central third of the 48.5-kb phage  genome is not required for infection and can therefore be replaced by foreign  DNAs of similar size. The resulting recombinant, or chimera (named after  the mythological monster with a lions head, goats body, and serpents tail), is  packaged into phage particles that can then be introduced into the host cells.  One advantage of using phage vectors is that the recombinant DNA is pro¬  duced in large amounts in easily purified form. Baculoviruses, which infect  insect cells, are similarly used for cloning in cultures of insect cells.   Much larger DNA segments—up to several hundred kilobase pairs—can  be cloned in large vectors known as bacterial artificial chromosomes (BACs)  or yeast artificial chromosomes (YACs). YACs are linear DNA molecules that  contain all the chromosomal structures required for normal replication and  segregation during yeast cell division. BACs, which replicate in E. coli, are de¬  rived from circular plasmids that normally replicate long regions of DNA and  are maintained at the level of approximately one copy per cell (properties sim¬  ilar to those of actual chromosomes).    FIG. 3-23 Bacteriophage A. During phage infection, DNA contained in the “head” of  the phage particle enters the bacterial cell, where it is replicated —100 times and pack¬  aged to form progeny phage. [Electron micrograph courtesy of A.F. Howatson. From Lewin,  B., Gene Expression, Vol. 3, Fig. 5.23, Wiley (1977).]    Ligase Joins Two DNA Segments. A DNA segment to be cloned is often  obtained through the action of restriction endonucleases. Most restriction en¬  zymes cleave DNA to yield sticky ends (Section 3-4A). Therefore, as Janet  Mertz and Ron Davis first demonstrated in 1972, a restriction fragment can  be inserted into a cut made in a cloning vector by the same restriction enzyme  (Fig. 3-24). The complementary ends of the two DNAs form base pairs (an¬  neal) and the sugar-phosphate backbones are covalently ligated, or spliced  together, through the action of an enzyme named DNA ligase. (A ligase pro¬  duced by a bacteriophage can also join blunt-ended restriction fragments.) A  great advantage of using a restriction enzyme to construct a recombinant  DNA molecule is that the DNA insert can later be precisely excised from the  cloned vector by cleaving it with the same restriction enzyme.   Selection Detects the Presence of a Cloned DNA. The expression of a  chimeric plasmid in a bacterial host was first demonstrated in 1973 by Herbert  Boyer and Stanley Cohen. A host bacterium can take up a plasmid when the  two are mixed together, but the vector becomes permanently established in its  bacterial host (transformation) with an efficiency of only —0.1%. However, a  single transformed cell can multiply without limit, producing large quantities  of recombinant DNA. Bacterial cells are typically plated on a semisolid growth  medium at a low enough density that discrete colonies, each arising from a  single cell, are visible.   It is essential to select only those host organisms that have been transformed  and that contain a properly constructed vector. In the case of plasmid transfor¬  mation, selection can be accomplished through the use of antibiotics and/or  chromogenic (color-producing) substances. For example, the lacZ gene in the  pUC18 plasmid (see Fig. 3-22) encodes the enzyme (3-galactosidase, which  cleaves the colorless compound X-gal to a blue product:    Cells of E. coli that have been transformed by an unmodified pUC18 plasmid  form blue colonies. However, if the plasmid contains a foreign DNA insert in  its polylinker region, the colonies are colorless because the insert interrupts the  protein-coding sequence of the lacZ gene and no functional (3-galactosidase is  produced. Bacteria that have failed to take up any plasmid are also colorless  due to the absence of (3-galactosidase, but these cells can be excluded by adding  the antibiotic ampicillin to the growth medium (the plasmid includes the gene  amp R , which confers ampicillin resistance). Thus, successfully transformed cells   The sticky ends of  the vector and the  foreign DNA  fragments anneal  and are covalently  joined by DNA  ligase.   The result is a  chimeric DNA  containing a portion  of the foreign DNA  inserted into the  vector.    FIG. 3-24 Construction of a recombinant DNA molecule. O See the Animated Figures.    form colorless colonies in the presence of ampicillin. Genes such as amp R are  known as selectable markers.   Genetically engineered bacteriophage X vectors contain restriction sites  that flank the dispensable central third of the phage genome. This segment  can be replaced by foreign DNA, but the chimeric DNA is packaged in phage  particles only if its length is from 75 to 105% of the 48.5-kb wild-type X  genome (Fig. 3-25). Consequently, X phage vectors that have failed to acquire  a foreign DNA insert are unable to propagate because they are too short to   Remaining X phage DNA  contains genes required  for infection but is too  small to package.    FIG. 3-25 Cloning with bacteriophage A. Removal of a nonessential portion of the phage genome allows a segment of foreign DNA to  be inserted. The DNA insert can be packaged into an infectious phage particle only if the insert DNA has the appropriate size. Q See the    form infectious phage particles. Of course, the production of infectious phage  particles results not in a growing bacterial colony but in a plaque, a region of  lysed bacterial cells, on a culture plate containing a “lawn” of the host bacte¬  ria. The recombinant DNA—now much amplified—can be recovered from  the phage particles in the plaque.   In order to clone a particular DNA fragment, it must first be obtained in rel¬  atively pure form. The magnitude of this task can be appreciated by consid¬  ering that, for example, a 1-kb fragment of human DNA represents only  0.00003% of the 3 billion-bp human genome. Of course, identifying a par¬  ticular DNA fragment requires knowing something about its nucleotide se¬  quence or its protein product. In practice, it is usually more difficult to identify  a particular DNA fragment from an organism and then clone it than it is to  clone all the organisms DNA that might contain the DNA of interest and  then identify the clones containing the desired sequence.    How many clones must be obtained from  Drosophila in order to be 99% sure that  they include a particular 10-kb fragment?    Use Equation 3-2 and the size of the  Drosophila genome given in Table 3-3.  Here, f = 10 kb/180,000 kb = 3.36 X  1(T 3 .   = log( 1 - 0.99)/log(l - 5.56 X 10 -3 )  = —2/(—2.413 X 1(T 3 )    A Genomic Library Includes All of an Organism’s DNA. The cloned set of  all DNA fragments from a particular organism is known as its genomic  library. Genomic libraries are generated by a procedure known as shotgun  cloning. The chromosomal DNA of the organism is isolated, cleaved to frag¬  ments of cloneable size, and inserted into a cloning vector. The DNA is usu¬  ally fragmented by partial rather than exhaustive restriction digestion so that  the genomic library contains intact representatives of all the organisms genes,  including those that contain restriction sites. DNA in solution can also be me¬  chanically fragmented (sheared) by rapid stirring.   Given the large size of the genome relative to a gene, the shotgun cloning  method is subject to the laws of probability. The number of randomly gener¬  ated fragments that must be cloned to ensure a high probability that a desired  sequence is represented at least once in the genomic library is calculated as  follows: The probability Pthat a set of clones contains a fragment that con¬  stitutes a fraction f, in bp, of the organisms genome is   Thus, in order for P to equal 0.99 for fragments averaging 10 kb in length,  N = 2162 for the 4600-kb E. coli chromosome (see Sample Calculation 3-1).  The use of BAC- or YAC-based genomic libraries with their large fragment  lengths therefore greatly reduces the effort necessary to obtain a given gene seg¬  ment from a large genome. After a BAC- or YAC-based clone containing the  desired DNA has been identified (see below), its large DNA insert can be fur¬  ther fragmented and cloned again (subcloned) to isolate the target DNA.    A cDNA Library Represents Expressed Genes. A different type of DNA  library contains only the expressed sequences from a particular cell type. Such  a cDNA library is constructed by isolating all the cells mRNAs and then  copying them to DNA using a specialized type of DNA polymerase known as  reverse transcriptase because it synthesizes DNA using RNA templates (Box  25-2). The complementary DNA (cDNA) molecules are then inserted into  cloning vectors to form a cDNA library. A cDNA library can also be used to  construct a DNA microarray (DNA chip), in which each different cDNA is  immobilized at a specific position on a slide. A DNA chip can be used for  detecting the presence of mRNA in a biological sample (the mRNA, if pres¬  ent, will bind to its complementary cDNA; Section 14-4C).    FIG. 3-26 Colony (in situ ) hybridization. Colonies are transferred from a “master”  culture plate by replica plating. Clones containing the DNA of interest are identified by the  ability to bind a specific probe. Here, binding is detected by laying X-ray film over the dried  filter. Since the colonies on the master plate and on the filter have the same spatial  distribution, positive colonies are easily retrieved.    A Library Is Screened for the Gene of Interest. Once the requisite number  of clones is obtained, the genomic library must be screened for the presence  of the desired gene. This can be done by a process known as colony or in  situ hybridization (Latin: in situ. , in position; Fig. 3-26). The cloned yeast  colonies, bacterial colonies, or phage plaques to be tested are transferred, by  replica plating, from a master plate to a nitrocellulose filter (replica plating  is also used to transfer colonies to plates containing different growth media).  Next, the filter is treated with NaOH, which lyses the cells or phages and sep¬  arates the DNA into single strands, which preferentially bind to the nitrocel¬  lulose. The filter is then dried to fix the DNA in place and incubated with a  labeled probe. The probe is a short segment of DNA or RNA whose sequence  is complementary to a portion of the DNA of interest. After washing away  unbound probe, the presence of the probe on the nitrocellulose is detected by  a technique appropriate for the label used (e.g., exposure to X-ray film for a  radioactive probe, a process known as autoradiography, or illumination with  an appropriate wavelength for a fluorescent probe). Only those colonies or  plaques containing the desired gene bind the probe and are thereby detected.  The corresponding clones can then be retrieved from the master plate. Using  this technique, a human genomic library of ~ 1 million clones can be readily  screened for the presence of one particular DNA segment.   Choosing a probe for a gene whose sequence is not known requires some  artistry. The corresponding mRNA can be used as a probe if it is available in  sufficient quantities to be isolated. Alternatively, if the amino acid sequence  of the protein encoded by the gene is known, the probe may be a mixture of  the various synthetic oligonucleotides that are complementary to a segment of  the genes inferred base sequence. Several disease-related genes have been iso¬  lated using probes specific for nearby markers, such as repeated DNA se¬  quences, that were already known to be genetically linked to the disease genes.   Although molecular cloning techniques are indispensable to modern biochem¬  ical research, the polymerase chain reaction (PCR) is often a faster and more  convenient method for amplifying a specific DNA. Segments of up to 6 kb can  be amplified by this technique, which was devised by Kary Mullis in 1985. In  PCR, a DNA sample is separated into single strands and incubated with DNA  polymerase, dNTPs, and two oligonucleotide primers whose sequences flank the  DNA segment of interest. The primers direct the DNA polymerase to synthesize com¬  plementary strands of the target DNA (Fig. 3-27). Multiple cycles of this process,  each doubling the amount of the target DNA, geometrically amplify the DNA  starting from as little as a single gene copy. In each cycle, the two strands of  the duplex DNA are separated by heating, then the reaction mixture is cooled  to allow the primers to anneal to their complementary segments on the DNA.  Next, the DNA polymerase directs the synthesis of the complementary strands.  The use of a heat-stable DNA polymerase, such as Taq polymerase isolated  from Thermus aquaticus, a bacterium that thrives at 75°C, eliminates the need  to add fresh enzyme after each round of heating (heat inactivates most en¬  zymes). Hence, in the presence of sufficient quantities of primers and dNTPs,  PCR is carried out simply by cyclically varying the temperature.    Velvet is pressed  to master plate  and transferred to  nitrocellulose filter.    FIG. 3-27 The polymerase chain reaction (PCR). The number of “unit-length”  strands doubles with every cycle after the second cycle. By choosing primers specific for  each end of a gene, the gene can be amplified over a millionfold.    Twenty cycles of PCR increase the amount of the target sequence around a  millionfold (~2 20 ) with high specificity. Indeed, PCR can amplify a target DNA  present only once in a sample of 10 5 cells, so this method can be used without  prior DNA purification. The amplified DNA can then be sequenced or cloned.   PCR amplification has become an indispensable tool. Clinically, it is used to  diagnose infectious diseases and to detect rare pathological events such as muta¬  tions leading to cancer. Forensically, the DNA from a single hair or sperm can be  amplified by PCR so that it can be used to identify the donor (Box 3-2).  Traditional ABO blood-type analysis requires a coin-sized drop of blood; PCR is  effective on pinhead-sized samples of biological fluids. Courts now consider DNA  sequences as unambiguous identifiers of individuals, as are fingerprints, because  the chance of two individuals sharing extended sequences of DNA is typically one  in a million or more. In a few cases, PCR has dramatically restored justice to con¬  victs who were released from prison on the basis of PCR results that proved their  innocence—even many years after the crime-scene evidence had been collected.   The ability to manipulate DNA sequences allows genes to be altered and ex¬  pressed in order to obtain proteins with improved functional properties or to  correct genetic defects.   Cloned Genes Can Be Expressed. The production of large quantities of  scarce or novel proteins is relatively straightforward only for bacterial proteins:  A cloned gene must be inserted into an expression vector, a plasmid that  contains properly positioned transcriptional and translational control se¬  quences. The production of a protein of interest may reach 30% of the host’s  total cellular protein. Such genetically engineered organisms are called over¬  producers. Bacterial cells often sequester large amounts of useless and possi¬  bly toxic (to the bacterium) protein as insoluble inclusions, which sometimes  simplifies the task of purifying the protein.   Bacteria can produce eukaryotic proteins only if the recombinant DNA  that carries the protein-coding sequence also includes bacterial transcriptional  and translational control sequences. Synthesis of eukaryotic proteins in bacte¬  ria also presents other problems. For example, many eukaryotic genes are large  and contain stretches of nucleotides (introns) that are transcribed and excised  before translation (Section 26-3A); bacteria lack the machinery to excise the  introns. In addition, many eukaryotic proteins are posttranslationally modi¬  fied by the addition of carbohydrates or by other reactions. These problems  can be overcome by using expression vectors that propagate in eukaryotic  hosts, such as yeast or cultured insect or animal cells.   Table 3-5 lists some recombinant proteins produced for medical and agri¬  cultural use. In many cases, purification of these proteins directly from human    Forensic DNA testing takes advantage of  DNA sequence variations or polymor¬  phisms that occur among individuals.   Many genetic polymorphisms have no  functional consequences because they  occur in regions of the DNA that contain  many repetitions but do not encode genes  (although if they are located near a “dis¬  ease” gene, they can be used to track and  identify the gene). Modern DNA finger¬  printing methods examine these noncoding repetitive DNA  sequences in samples that have been amplified by PCR.   Tandemly repeated DNA sequences occur throughout the human  genome and include short tandem repeats (STRs), which contain  variable numbers of repeating segments of two to seven base pairs.   The most popular STR sites for forensic use contain tetranucleotide  repeats. The number of repeats at any one site on the DNA varies  between individuals, even within a family. Each different number of  repeats at a site is called an allele, and each individual can have  two alleles, one from each parent (see figure above).   Since PCR is the first step of the fingerprinting process, only a  tiny amount (~1 ng) of DNA is needed. The region of DNA contain¬  ing the STR is amplified by PCR using primers that are complemen¬  tary to the unique (nonrepeating) sequences flanking the repeats.   The amplified products are separated by electrophoresis and  detected by the fluorescent tag on their primers. An STR allele is  small enough (—500 bp) that DNA fragments differing by a four-  base repeat can be readily differentiated. The allele designation  for each STR site is generally the number of times a repeated  unit is present. STR sites that have been selected for forensic use 8  generally have 7 to 30 different alleles. g   In the example shown at right, the upper trace shows the <8   (the set of all possible alleles, each identified by the number of  repeat units, from 13 to 23). The lower trace corresponds to the  sample being tested, which contains two alleles, one with 16  repeats and one with 18 repeats. Several STR sites can be ana¬  lyzed simultaneously by using the appropriate primers and tagging  them with different fluorescent dyes.   The probability of two individuals having matching DNA finger¬  prints depends on the number of STR sites examined and the    number of alleles at each site. For example, if a pair of alleles at  one site occurs in the population with a frequency of 10% (1/10),  and a pair of alleles at a second site occurs with a frequency of  5% (1/20), then the probability that the DNA fingerprints from  two individuals would match at both sites is 1 in 200 (1/10 X  1/20; the probabilities of independent events are multiplied). By  examining multiple STR sites, the probability of obtaining matching  fingerprints by chance becomes exceedingly small.   or animal tissues is unfeasible on ethical or  practical grounds. Expression systems per¬  mit large-scale, efficient preparation of the  proteins while minimizing the risk of con¬  tamination by viruses or other pathogens  from tissue samples.   Site-Directed Mutagenesis Alters a  Gene’s Nucleotide Sequence. After iso¬  lating a gene, it is possible to modify the  nucleotide sequence to alter the amino acid  sequence of the encoded protein. Site-  directed mutagenesis, a technique pio¬  neered by Michael Smith, mimics the natural   A synthetic oligonucleotide primer  incorporating the desired base  changes anneals to the DNA  containing the gene to be altered.   FIG. 3-28 Site-directed mutagenesis. The altered gene can be inserted into a suitable cloning vector to be amplified, expressed, or used  to generate a mutant organism. Q See the Animated Figures.   FIG. 3-29 Transgenic mouse. The gigantic  mouse on the left was grown from a fertilized  ovum that had been microinjected with DNA  containing the rat growth hormone gene. He is  nearly twice the weight of his normal littermate  on the right. [Courtesy of Ralph Brinster,  University of Pennsylvania.]    process of evolution and allows predictions about the structural and functional roles  of particular amino acids in a protein to be rigorously tested in the laboratory.   Synthetic oligonucleotides are required to specifically alter genes through  site-directed mutagenesis. An oligonucleotide whose sequence is identical to a  portion of the gene of interest except for the desired base changes is used to  direct replication of the gene. The oligonucleotide hybridizes to the correspon¬  ding wild-type (naturally occurring) sequence if there are no more than a few  mismatched base pairs. Extension of the oligonucleotide, called a primer, by  DNA polymerase yields the desired altered gene (Fig. 3-28). The altered gene  can then be inserted into an appropriate vector. A mutagenized primer can  also be used to generate altered genes by PCR.   Transgenic Organisms Contain Foreign Genes. For many purposes it is prefer¬  able to tailor an intact organism rather than just a protein—true genetic engi¬  neering. Multicellular organisms expressing a gene from another organism are said  to be transgenic, and the transplanted foreign gene is called a transgene.   For the change to be permanent, that is, heritable, a transgene must be  stably integrated into the organisms germ cells. For mice, this is accomplished  by microinjecting cloned DNA encoding the desired altered characteristics into  a fertilized egg and implanting it into the uterus of a foster mother. A well-  known example of a transgenic mouse contains extra copies of a growth hor¬  mone gene (Fig. 3-29).    Transgenic farm animals have also been developed. Ideally, the genes of  such animals could be tailored to allow the animals to grow faster on less food  or to be resistant to particular diseases. Some transgenic farm animals have  been engineered to secrete medically useful proteins into their milk. Harvesting  such a substance from milk is much more cost-effective than producing the  same substance in bacterial cultures.   One of the most successful transgenic organisms is corn (maize) that has  been genetically modified to produce a protein that is toxic to plant-eating in¬  sects (but harmless to vertebrates). The toxin is synthesized by the soil mi¬  crobe Bacillus thuringiensis. The toxin gene has been cloned into corn in order  to confer protection against the European corn borer, a commercially signifi¬  cant pest that spends much of its life cycle inside the corn plant, where it is  largely inaccessible to chemical insecticides. The use of “Bt corn,” which is  now widely planted in the United States, has greatly reduced the need for such  toxic substances.   Transgenic plants have also been engineered for better nutrition. For ex¬  ample, researchers have developed a strain of rice with foreign genes that en¬  code enzymes necessary to synthesize P-carotene (an orange pigment that is  the precursor of vitamin A) and a gene for the iron-storage protein ferritin.  The genetically modified rice, which is named “golden rice” (Fig. 3-30),  should help alleviate vitamin A deficiencies (which afflict some 400 million  people) and iron deficiencies (an estimated 30% of the worlds population suf¬  fers from iron deficiency). Other transgenic plants include freeze-tolerant  strawberries, slow-ripening tomatoes, and rapidly maturing fruit trees.   There is presently a widely held popular suspicion, particularly in Europe,  that genetically modified or “GM” foods will somehow be harmful. However,  extensive research, as well as considerable consumer experience, has failed to  reveal any deleterious effects caused by GM foods (see Box 3-3).   Transgenic organisms have greatly enhanced our understanding of gene  expression. Animals that have been engineered to contain a defective gene or   FIG. 3-30 Golden rice. The white grains on  the left are the wild type. The grains on the right  have been engineered to store up to three times  more iron and to synthesize |3-carotene, which  gives them their yellow color. [Courtesy Ingo  Potrykus.]    In the early 1970s, when genetic engineering was first discussed,  little was known about the safety of the proposed experiments.  After considerable debate, during which there was a moratorium  on such experiments, regulations for recombinant DNA research  were drawn up. The rules prohibit obviously dangerous experi¬  ments (e.g., introducing the gene for diphtheria toxin into E. coli,  which would convert this human symbiont into a deadly  pathogen). Other precautions limit the risk of accidentally releas¬  ing potentially harmful organisms into the environment. For exam¬  ple, many vectors must be cloned in host organisms with special  nutrient requirements. These organisms are unlikely to survive  outside the laboratory.   The proven value of recombinant DNA technology has silenced  nearly all its early opponents. Certainly, it would not have been  possible to study some pathogens, such as the virus that causes  AIDS, without cloning. The lack of recombinant-induced genetic  catastrophes so far does not guarantee that recombinant organ¬  isms won’t ever adversely affect the environment. Nevertheless,  the techniques used by genetic engineers mimic those used in  nature—that is, mutation and selection—so natural and man-made  organisms are fundamentally similar. In any case, people have  been breeding plants and animals for several millennia already,    and for many of the same purposes that guide experiments with  recombinant DNA.   There are other ethical considerations to be faced as new  genetic engineering techniques become available. Bacterially pro¬  duced human growth hormone is now routinely prescribed to  increase the stature of abnormally short children. However, should  athletes be permitted to use this protein, as some reportedly have,  to increase their size and strength? Few would dispute the use of  gene therapy, if it can be developed, to cure such genetic defects  as sickle-cell anemia (Section 7-1 E) and Lesch-Nyhan syndrome  (Section 23-ID). If, however, it becomes possible to alter complex  (i.e., multigene) traits such as athletic ability and intelligence, which  changes would be considered desirable and who would decide  whether to make them? Should gene therapy be used only to  correct an individual’s defects, or should it also be used to alter  genes in the individual’s germ cells so that succeeding genera¬  tions would not inherit the defect? If it becomes easy to deter¬  mine an individual’s genetic makeup, should this information be  used in evaluating applicants for educational and employment  opportunities or for health insurance? These conundrums have led  to the creation of a branch of philosophy, named bioethics,  designed to deal with them.    that lack a gene entirely (a so-called gene knockout) also serve as experimen¬  tal models for human diseases.   • Summarize the steps required to amplify a  given segment of DNA in vivo and in vitro.   • Compare the properties of cloning vectors  such as pUC 18 , bacteriophage X, and  BACs.   • Describe the activities of the enzymes  required to construct a recombinant DNA  molecule.   • Explain why a cloning vector usually  includes a selectable marker.   • What is a DNA library and how can it be  screened for a particular gene?   • What are some of the challenges of ex¬  pressing a eukaryotic gene in a prokary¬  otic host cell?   • Explain how site-directed mutagenesis can  be used to produce an altered protein in  bacterial cells.   • What is the difference between manipulat¬  ing a gene for gene therapy and for pro¬  ducing a transgenic organism?    T Genetic Defects Can Be Corrected. Gene therapy is the transfer of new  genetic material to the cells of an individual in order to produce a thera¬  peutic effect. Although the potential benefits of this as yet rudimentary tech¬  nology are enormous, there are numerous practical obstacles to overcome. For  example, the retroviral vectors (RNA-containing viruses) commonly used to  directly introduce genes into humans can provoke a fatal immune response.   The first documented success of gene therapy in humans occurred in  children with a form of severe combined immunodeficiency disease  (SCID) known as SCID-X1, which without treatment would have required  their isolation in a sterile environment to prevent fatal infection. SCID-X1  is caused by a defect in the gene encoding yc cytokine receptor, whose ac¬  tion is essential for proper immune system function. Bone marrow cells (the  precursors of white blood cells) were removed from the bodies of SCID-X1  victims, incubated with a vector containing a normal yc cytokine receptor  gene, and returned to their bodies. The transgenic bone marrow cells re¬  stored immune system function. However, because the viral vector integrates  into the genome at random, the location of the transgene may affect the ex¬  pression of other genes, triggering cancer. At least two children have devel¬  oped leukemia (a white blood cell cancer) as a result of gene therapy for  SCID-X1.   Other diseases that have been successfully treated by gene therapy are  Leber’s congenital amaurosis, a rare form of blindness, X-linked  adrenoleukodystrophy, in which a defect in a membrane transport protein  leads to brain damage, and (3-thalassemia, a type of severe anemia.   • Nucleotides consist of a purine or pyrimidine base linked to ri-  bose to which at least one phosphate group is attached. RNA is  made of ribonucleotides; DNA is made of deoxynucleotides  (which contain 2'-deoxyribose).   • In DNA, two antiparallel chains of nucleotides linked by phos-  phodiester bonds form a double helix. Bases in opposite strands  pair: A with T, and G with C.   • Single-stranded nucleic acids, such as RNA, can adopt stem-loop  structures.   • DNA carries genetic information in its sequence of nucleotides.  When DNA is replicated, each strand acts as a template for the  synthesis of a complementary strand.   • According to the central dogma of molecular biology, one strand  of the DNA of a gene is transcribed into mRNA. The RNA is  then translated into protein by the ordered addition of amino  acids that are bound to tRNA molecules that base-pair with the  mRNA at the ribosome.   • Restriction endonucleases that recognize certain sequences of  DNA are used to specifically cleave DNA molecules.    • Gel electrophoresis is used to separate and measure the sizes of  DNA fragments.   • In the chain-terminator method of DNA sequencing, the sequence  of nucleotides in a DNA strand is determined by enzymatically  synthesizing complementary polynucleotides that terminate with  a dideoxy analog of each of the four nucleotides. Polynucleotide  fragments of increasing size are separated by electrophoresis to re¬  construct the original sequence.   • Mutations and other changes to DNA are the basis for the evo¬  lution of organisms.   • In molecular cloning, a fragment of foreign DNA is inserted into  a vector for amplification in a host cell. Transformed cells can be  identified by selectable markers.   • Genomic libraries contain all the DNA of an organism. Clones  harboring particular DNA sequences are identified by screening  procedures.   • Recombinant DNA methods are used to produce wild-type or  selectively mutagenized proteins in cells or entire organisms.   transformation 49  replication 49  transcription 50  translation 50  central dogma of molecular  biology 50  mRNA 50  ribosome 50  rRNA 50  tRNA 50  genomics 51  transcriptomics 51  proteomics 51  gene expression 51  bacteriophage 52  restriction-modification  system 52   3. In many organisms, DNA is modified by methylation. Draw the  structure of 5-methylcytosine, a base that occurs with high fre¬  quency in inactive DNA.   4. When cytosine is treated with bisulfite, the amino group is re¬  placed with a carbonyl group. Identify the resulting base.   3. Kinases are enzymes that transfer a phosphoryl group from a nu¬  cleoside triphosphate. Which of the following are valid kinase-  catalyzed reactions?   6. Kinases are enzymes that transfer a phosphoryl group from a nu¬  cleoside triphosphate. Which of the following are valid kinase-  catalyzed reactions?   7. A diploid organism with a 45,000-kb haploid genome contains  21% G residues. Calculate the number of A, C, G, and T  residues in the DNA of each cell in this organism.   8. A segment of DNA containing 20 base pairs includes 7 guanine  residues. How many adenine residues are in the segment? How  many uracil residues are in the segment?   11. The adenine derivative hypoxanthine can base-pair with cytosine.  Draw the structure of this base pair.   12. Hypoxanthine can also base-pair with adenine. Draw the struc¬  ture of this base pair.   13. Explain why the strands of a DNA molecule can be separated  more easily at pH > 11.   13. An enzyme from the human immunodeficiency virus (HIV, which  causes AIDS) can synthesize DNA from an RNA template. Explain  how this enzyme activity contradicts Cricks central dogma.   16. How many different amino acids could theoretically be encoded  by nucleic acids containing four different nucleotides if (a) each  nucleotide coded for one amino acid; (b) consecutive sequences  of two nucleotides coded for one amino acid; (c) consecutive se¬  quences of three nucleotides coded for one amino acid; (d) con¬  secutive sequences of four nucleotides coded for one amino acid?   17. The recognition sequence for the restriction enzyme TaqI is  T^CGA. Indicate the products of the reaction of TaqI with the  DNA sequence shown.   18. Using the data in Table 3-2, identify restriction enzymes that (a)  produce blunt ends; (b) recognize and cleave the same sequence  (called isoschizomers); (c) produce identical sticky ends.   19. Describe the outcome of a chain-terminator sequencing proce¬  dure in which (a) too little ddNTP is added or (b) too much  ddNTP is added.   20. Describe the outcome of a chain-terminator sequencing proce¬  dure in which (a) too few primers are present or (b) too many  primers are present.   21. The 13-Mb genome of the green alga Ostreococcus tauri contains  — 8000 genes. Compare the gene density in this eukaryote to that  of E. coli (—4300 genes) and that of A. thaliana (—23,300 genes).   22. Describe how to select recombinant clones if a foreign DNA is  inserted into the polylinker site of pUC18 and then introduced  into E. coli cells.   23. Calculate the number of clones required to obtain with a proba¬  bility of 0.99 a specific 5-kb fragment from C. elegans (Table 3-3).   24. You are attempting to clone a 250-kb segment of mouse DNA  in a yeast artificial chromosome. You obtain 5000 similar-sized  clones representing the entire mouse genome. How confident are  you that you have cloned the DNA you are interested in?   25. Why is a genomic library larger than a cDNA library for a given  organism?   26. Why do cDNA libraries derived from different cell types within  the same organism differ from each other?   27. Describe the possible outcome of a PCR experiment in which (a)  one of the primers is inadvertently omitted from the reaction  mixture and (b) one of the primers is complementary to several  sites in the starting DNA sample.    28. Describe the possible outcome of a PCR experiment in which (a)  there is a single-stranded break in the target DNA sequence,  which is present in only one copy in the starting sample, and (b)  there is a double-stranded break in the target DNA sequence,  which is present in only one copy in the starting sample.   29. Write the sequences of the two 12-residue primers that could be  used to amplify the following DNA segment by PCR.   30. A blood stain from a crime scene and blood samples from four  suspects were analyzed by PCR using fluorescent primers associ¬  ated with three STR loci: D3S1358, vWA, and FGA. The result¬  ing electrophoretograms are shown below. The numbers beneath  each peak identify the allele (upper box) and the height of the  peak in relative fluorescence units (lower box).   (a) Since everyone has two copies of each chromosome and  therefore two alleles of each gene, what accounts for the  appearance of only one allele at some loci?   (c) Could the suspect be identified using just one of the three  STR loci?   (d) What can you conclude about the amount of DNA obtained  from Suspect 1 compared to Suspect 4?    [From Thompson, W.C., Ford, S., Doom, T., Raymer, M., and Krane,  D.E., Evaluating forensic DNA evidence: Essential elements of a com¬  petent defense review, The Champion 27 , 16-25 (2003).]   1. Finding Databases. Locate databases for genome sequences  and explore the meaning of terms related to them.   2. The Institute for Genomic Research. Explore a prokaryotic  genome and find listings for eukaryotic genomes.   3. Analyzing a DNA Sequence. Given a DNA sequence, identify  its open reading frame and translate it into a protein sequence.   4. Sequence Homology. Perform a BLAST search for homologs  of a protein sequence.   5. Plasmids and Cloning. Predict the sizes of the fragments pro¬  duced by the action of various restriction enzymes on plasmids.    Look up one of the genetic dis¬  eases listed in Table 3-4. What gene is involved? What is the nor¬  mal function of the protein encoded by the gene, and how does  the gene defect produce the characteristic symptoms? How might  this disease be treated through gene therapy?   Bloomfield, V.A., Crothers, D.M., andTinoco, I., Jr., Nucleic Acids. Struc¬  tures, Properties, and Functions, University Science Books (2000).   Dickerson, R.E., DNA structure from A to Z, Methods Enzymol. 211,  67-111 (1992). [Describes the various crystallographic forms of  DNA.]   Thieffry, D., Forty years under the central dogma, Trends Biochem. Sci.  23, 312-316 (1998). [Traces the origins, acceptance, and shortcom¬  ings of the idea that nucleic acids contain biological information.]   Watson, J.D. and Crick, F.H.C., Molecular structure of nucleic acids, Na¬  ture 171, 737-738 (1933); <m^Genetical implications of the struc¬  ture of deoxyribonucleic acid, Nature 171, 964-967 (1933). [The  seminal papers that are widely held to mark the origin of modern  molecular biology.]   Galperin, M.Y., and Cochrane, G.R. The 2011 Nucleic Acids Research data¬  base issue and online molecular biology database collection, Nucleic  Acids Res. 38, Database issue D1-D6 (2011). [This annually updated  article describes 1330 databases covering various aspects of molecu¬  lar biology, biochemistry, and genetics. Additional articles in the same  issue provide more information on individual databases. Freely avail¬  able at http://nar.oxfordjournals.org.]   Graham, C.A. and Hill, A.J.M. (Eds.), DNA Sequencing Protocols (2nd  ed.), Humana Press (2001).   Higgins, D. and Taylor, W. (Eds.), Bioinformatics. Sequence, Structure and  Databanks, Oxford University Press (2000).   International Human Genome Sequencing Consortium, Initial sequenc¬  ing and analysis of the human genome, Nature 409, 860-921    (2001); and Venter, J.C., et al., the sequence of the human genome,  Science 291, 1304-1351 (2001). [These and other papers in the  same issues of Nature and Science describe the data that constitute  the draft sequence of the human genome and discuss how this in¬  formation can be used in understanding biological function, evolu¬  tion, and human health.]   International Human Genome Sequencing Consortium, Finishing the eu-  chromatic sequence of the human genome, Nature 431, 931-945  (2004). [Describes the “finished” version of the human genome  sequence.]   Ausubel, F.M., Brent, R., Kingston, R.E., Moore, D.D., Seidman, J.G.,  Smith, J.A., and Struhl, K., Short Protocols in Molecular Biology (5th  ed.), Wiley (2002).   Pingoud, A., Fuxreiter, M., Pingoud, V., and Wende, W., Type II restric¬  tion endonucleases: structure and mechanism, Cell. Mol. Life Sci. 62,  685-707 (2005). [Includes an overview of different types of restric¬  tion enzymes.]   Sambrook, J., and Russell, D., Molecular Cloning (3rd ed.), Cold Spring  Harbor Laboratory (2001). [A three-volume “bible” of laboratory  protocols with accompanying background explanations.]   von Bubnoff, A., Next-generation sequencing: The race is on, Cell 132,  721-723 (2008).   Watson, J.D., Meyers, R.M., Caudy, A.A., and Witkowski, J.A., Recom¬  binant DNA. Genes and Genomes—A Short Course (3rd ed.), Free¬  man (2007). [An exposition of the methods, findings, and results of  recombinant DNA technology and research.]   A Amino Acids Are Dipolar Ions  B Peptide Bonds Link Amino Acids  C Amino Acid Side Chains Are Nonpolar, Polar,  or Charged   A Protein Side Chains May Be Modified  B Some Amino Acids Are Biologically Active   The byssal threads by which a mussel attaches itself to rocks are made from adhesive  proteins that contain an unusually high proportion of a hydroxylated form of the amino  acid tyrosine called L-3,4-dihydroxyphenylalanine (l-DOPA). This amino acid forms  extraordinarily strong noncovalent interactions with various types of surfaces. Individual  bonds between the protein and the substrate can break under stress but can also  re-form, giving the byssal threads their characteristic strength and extensibility.  [Photography by Matthew Harrington]    When scientists first turned their attention to nutrition, early in the nineteenth  century, they quickly discovered that natural products containing nitrogen  were essential for the survival of animals. In 1839, the Swedish chemist Jacob  Berzelius coined the term protein (Greek: proteios , primary) for this class of  compounds. The physiological chemists of that time did not realize that pro¬  teins were actually composed of smaller components, amino acids, although  the first amino acids had been isolated in 1830. In fact, for many years, it was  believed that substances from plants—including proteins—were incorporated  whole into animal tissues. This misconception was laid to rest when the  process of digestion came to light. After it became clear that ingested proteins  were broken down to smaller compounds containing amino acids, scientists  began to consider the nutritive qualities of those compounds (Box 4-1).   Modern studies of proteins and amino acids owe a great deal to nineteenth  and early twentieth century experiments. We now understand that nitrogen-  containing amino acids are essential for life and that they are the building  blocks of proteins. The central role of amino acids in biochemistry is perhaps  not surprising: Several amino acids are among the organic compounds believed  to have appeared early in the earths history (Section 1-1 A). Amino acids, as  ancient and ubiquitous molecules, have been co-opted by evolution for a va¬  riety of purposes in living systems. We begin this chapter by discussing the  structures and chemical properties of the common amino acids, including their  stereochemistry, and end with a brief summary of the structures and functions  of some related compounds.    William C. Rose (1887-1985) Identifying  - . \ the amino acid constituents of proteins was a  f j scientific challenge that grew out of studies of   not all foods provided adequate nutrition. For  example, rats fed the corn protein zein as  their only source of nitrogen failed to grow  unless the amino acids tryptophan and lysine were added to their  diet. Knowledge of metabolism at that time was mostly limited to  information gleaned from studies in which intake of particular  foods in experimental subjects (including humans) was linked to  the urinary excretion of various compounds. Results of such  studies were consistent with the idea that compounds could be  transformed into other compounds, but clearly, nutrients were not  wholly interchangeable.   At the University of Illinois, William C. Rose focused his  research on nutritional studies to decipher the metabolic  relationships of nitrogenous compounds. Among other things, his  studies of rat growth and nutrition helped show that purines and  pyrimidines were derived from amino acids but that those  compounds could not replace dietary amino acids.   In order to examine the nutritional requirements for individual  amino acids, Rose hydrolyzed proteins to obtain their component  amino acids and then selectively removed certain amino acids. In  one of his first experiments, he removed arginine and histidine  from a hydrolysate of the milk protein casein. Rats fed on this  preparation lost weight unless the amino acid histidine was added  back to the food. However, adding back arginine did not  compensate for the apparent requirement for histidine. These  results prompted Rose to investigate the requirements for all the    amino acids. Using similar experimental approaches, Rose  demonstrated that cysteine, histidine, and tryptophan could not be  replaced by other amino acids.   From preparations based on hydrolyzed proteins, Rose moved  to mixtures of pure amino acids. Thirteen of the 19 known amino  acids could be purified, and the other six synthesized. However,  rats fed these 19 amino acids as their sole source of dietary  nitrogen lost weight. Although one possible explanation was that  the proportions of the pure amino acids were not optimal, Rose  concluded that there must be an additional essential amino acid,  present in naturally occurring proteins and their hydrolysates but  not in his amino acid mixtures.   After several years of effort, Rose obtained and identified the  missing amino acid. In work published in 1935, Rose showed that  adding this amino acid to the other 19 could support rat growth.  Thus, the twentieth and last amino acid, threonine, was discovered.   Experiments extending over the next 20 years revealed that 10  of the 20 amino acids found in proteins are nutritionally essential, so  that removal of one of these causes growth failure and eventually  death in experimental animals. The other 10 amino acids were  considered “dispensable” since animals could synthesize adequate  amounts of them.   Rose’s subsequent work included verifying the amino acid  requirements of humans, using graduate students as subjects.  Knowing which amino acids were required for normal health—and in  what amounts-made it possible to evaluate the potential nutritive  value of different types of food proteins. Eventually, these findings  helped guide the formulations used for intravenous feeding.   McCoy, R.H., Meyer, C.E., and Rose, W.C., Feeding experiments with mixtures of highly  purified amino acids. VIII. Isolation and identification of a new essential amino acid,    • The 20 standard amino acids share a common structure but differ in their side  chains.   • Some amino acid side chains contain ionizable groups whose p K values may vary.    The analyses of a vast number of proteins from almost every conceivable  source have shown that all proteins are composed of 20 “standard” amino acids.  Not every protein contains all 20 types of amino acids, but most proteins con¬  tain most if not all of the 20 types.   The common amino acids are known as ot-amino acids because they have  a primary amino group (—NH 2 ) as a substituent of the cl carbon atom, the  carbon next to the carboxylic acid group (—COOH; Fig. 4-1). The sole  exception is proline, which has a secondary amino group (—NH—), although  for uniformity we will refer to proline as an a-amino acid. The 20 standard  amino acids differ in the structures of their side chains (R groups). Table 4-1  displays the names and complete chemical structures of the 20 standard amino  acids.   FIG. 4-1 General structure of an ct-amino  acid. The R groups differentiate the 20 standard  amino acids.    The ionic forms shown are those predominating at pH 7.0 (except for that of histidine-^) although residue mass is given for the neutral compound. The  C a atoms, as well as those atoms marked with an asterisk, are chiral centers with configurations as indicated according to Fischer projection formulas  (Section 4-2). The standard organic numbering system is provided for heterocycles.   ^The residue masses are given for the neutral residues. For the molecular masses of the parent amino acids, add 18.0 D, the molecular mass of H 2 0, to  the residue masses. For side chain masses, subtract 56.0 D, the formula mass of a peptide group, from the residue masses.   The average amino acid composition in the complete SWISS-PROT database (http://www.expasy.ch/sprot/relnotes/relstat.html), Release 55.11.   ^Data from Dawson, R.M.C., Elliott, D.C., Elliott, W.H., and Jones, K.M., Data for Biochemical Research (3rd ed.), pp. 1-31, Oxford Science  Publications (1986).    CThe three- and one-letter symbols for asparagine or aspartic acid are Asx and B, whereas for glutamine or glutamic acid they are Glx and Z. The one-letter  symbol for an undetermined or “nonstandard” amino acid is X.   ^Both neutral and protonated forms of histidine are present at pH 7.0, since its pAh is close to 7.0.   physiological pH, the amino group is protonated  and the carboxylic acid group is unprotonated.    FIG. 4-3 Condensation of two amino  acids. Formation of a CO—NH bond with the  elimination of a water molecule produces a  dipeptide. The peptide bond is shown in red.  The residue with a free amino group is the  N-terminus of the peptide, and the residue with  a free carboxylate group is the C-terminus.    Draw a tripeptide resulting from the  condensation of a third amino acid.   The amino and carboxylic acid groups of amino acids readily ionize. The p K  values of the carboxylic acid groups (represented by p K x in Table 4-1) lie in a  small range around 2.2, while the pK values of the a-amino groups (p K 2 ) are  near 9.4. At physiological pH (~ 7.4), the amino groups are protonated and the  carboxylic acid groups are in their conjugate base (carboxylate) form (Fig. 4-2).  An amino acid can therefore act as both an acid and a base. Table 4-1 also lists  the p K values for the seven side chains that contain ionizable groups (pA^).   Molecules such as amino acids, which bear charged groups of opposite po¬  larity, are known as dipolar ions or zwitterions. Amino acids, like other ionic  compounds, are more soluble in polar solvents than in nonpolar solvents. As  we will see, the ionic properties of the side chains influence the physical and  chemical properties of free amino acids and amino acids in proteins.   Amino acids can be polymerized to form chains. This process can be repre¬  sented as a condensation reaction (bond formation with the elimination of  a water molecule), as shown in Fig. 4-3. The resulting CO—NH linkage, an  amide linkage, is known as a peptide bond.   Polymers composed of two, three, a few (3—10), and many amino acid  units are known, respectively, as dipeptides, tripeptides, oligopeptides, and  polypeptides. These substances, however, are often referred to simply as  “peptides.” After they are incorporated into a peptide, the individual amino  acids (the monomeric units) are referred to as amino acid residues.   Polypeptides are linear polymers rather than branched chains; that is, each  amino acid residue participates in two peptide bonds and is linked to its neigh¬  bors in a head-to-tail fashion. The residues at the two ends of the polypep¬  tide each participate in just one peptide bond. The residue with a free amino  group (by convention, the leftmost residue, as shown in Fig. 4-3) is called the  amino terminus or N-terminus. The residue with a free carboxylate group  (at the right) is called the carboxyl terminus or C-terminus.   Proteins are molecules that contain one or more polypeptide chains.  Variations in the length and the amino acid sequence of polypeptides are major  contributors to the diversity in the shapes and biological functions of proteins, as  we will see in succeeding chapters.   The most useful way to classify the 20 standard amino acids is by the polar¬  ities of their side chains. According to the most common classification scheme,  there are three major types of amino acids: (1) those with nonpolar R groups,  (2) those with uncharged polar R groups, and (3) those with charged polar  R groups.   The Nonpolar Amino Acid Side Chains Have a Variety of Shapes and Sizes.   Nine amino acids are classified as having nonpolar side chains. The three-  dimensional shapes of some of these amino acids are shown in Fig. 4-4. Glycine  has the smallest possible side chain, an H atom. Alanine, valine, leucine, and  isoleucine have aliphatic hydrocarbon side chains ranging in size from a methyl  group for alanine to isomeric butyl groups for leucine and isoleucine.  Methionine has a thioether side chain that resembles an /z-butyl group in many  of its physical properties (C and S have nearly equal electronegativities, and S  is about the size of a methylene group). Proline has a cyclic pyrrolidine side  group. Phenylalanine (with its phenyl moiety) and tryptophan (with its in¬  dole group) contain aromatic side groups, which are characterized by bulk as  well as nonpolarity.   FIG. 4-4 Some amino acids with nonpolar side chains. The amino acids are shown  as ball-and-stick models embedded in transparent space-filling models. The atoms are  colored according to type with C green, H white, N blue, and 0 red.   Uncharged Polar Side Chains Have Hydroxyl, Amide, or Thiol Groups. Six   amino acids are commonly classified as having uncharged polar side chains  (Table 4-1 and Fig. 4-5). Serine and threonine bear hydroxylic R groups of  different sizes. Asparagine and glutamine have amide-bearing side chains of  different sizes. Tyrosine has a phenolic group (and, like phenylalanine and  tryptophan, is aromatic). Cysteine is unique among the 20 amino acids in  that it has a thiol group that can form a disulfide bond with another cysteine  through the oxidation of the two thiol groups (Fig. 4-6).    FIG. 4-5 Some amino acids with uncharged polar side chains. Atoms are  represented and colored as in Fig. 4-4. Note the presence of electronegative atoms on  the side chains.    FIG. 4-6 Disulfide-bonded cysteine residues. The disulfide bond forms when  the two thiol groups are oxidized.    FIG. 4-7 Some amino acids with charged  polar side chains. Atoms are represented and  colored as in Fig. 4-4.    In the neutral species, the a-carboxylate  group is deprotonated and the a-amino  group is protonated. Protonation of the a-  carboxylate group or deprotonation of the  (3-carboxylate group would both yield  charged species. Therefore, the p K values  for these groups (1.99 and 3.90; see Table  4-1) should be used with Equation 4-1:    Charged Polar Side Chains Are Positively or Negatively Charged. Five  amino acids have charged side chains (Table 4-1 and Fig. 4-7). The side chains  of the basic amino acids are positively charged at physiological pH values.  Lysine has a butylammonium side chain, and arginine bears a guanidino  group. As shown in Table 4-1, histidine carries an imidazolium moiety. Note  that only histidine, with a pA^ of 6.04, readily ionizes within the physiolog¬  ical pH range. Consequently, both the neutral and cationic forms occur in  proteins. In fact, the protonation-deprotonation of histidine side chains is a  feature of numerous enzymatic reaction mechanisms.   The side chains of the acidic amino acids, aspartic acid and glutamic  acid, are negatively charged above pH 3; in their ionized state, they are often  referred to as aspartate and glutamate. Asparagine and glutamine are, respec¬  tively, the amides of aspartic acid and glutamic acid.   The foregoing allocation of the 20 amino acids among the three different  groups is somewhat arbitrary. For example, glycine and alanine, the smallest  of the amino acids, and tryptophan, with its heterocyclic ring, might just as  well be classified as uncharged polar amino acids. Similarly, tyrosine and cys¬  teine, with their ionizable side chains, might also be thought of as charged po¬  lar amino acids, particularly at higher pH values. In fact, the deprotonated  side chain of cysteine (which contains the thiolate anion, S ) occurs in a va¬  riety of enzymes, where it actively participates in chemical reactions.   Inclusion of a particular amino acid in one group or another reflects not  just the properties of the isolated amino acid, but its behavior when it is part  of a polypeptide. The structures of most polypeptides depend on a tendency  for polar and ionic side chains to be hydrated and for nonpolar side chains to  associate with each other rather than with water. This property of polypep¬  tides is the hydrophobic effect (Section 2-1C) in action. As we will see, the  chemical and physical properties of amino acid side chains also govern the  chemical reactivity of the polypeptide. It is therefore worthwhile studying  the structures of the 20 standard amino acids in order to appreciate how they  vary in polarity, acidity, aromaticity, bulk, conformational flexibility, ability  to cross-link, ability to hydrogen bond, and reactivity toward other groups.   D The p K Values of Ionizable Groups Depend on Nearby Groups   The a-amino acids have two or, for those with ionizable side chains, three  acid-base groups. At very low pH values, these groups are fully protonated,  and at very high pH values, these groups are unprotonated. At intermediate  pH values, the acidic groups tend to be unprotonated, and the basic groups  tend to be protonated. Thus, for the amino acid glycine, below pH 2.35 (the  pK value of its carboxylic acid group), the + H 3 NCH 2 COOH form predom¬  inates. Above pH 2.35, the carboxylic acid is mostly ionized but the amino  group is still mostly protonated ( + H 3 NCH 2 COO _ ). Above pH 9.78 (the pK  value of the amino group), the H 2 NCH 2 COO form predominates. Note  that in aqueous solution ., the un-ionized form (H 2 NCH 2 C00H) is present only  in vanishingly small quantities.   The pH at which a molecule carries no net electric charge is known as its  isoelectric point, p/. For the a-amino acids,   where A^- and Kj are the dissociation constants of the two ionizations involv¬  ing the neutral species. For monoamino, monocarboxylic acids such as glycine,  A!) and Kj represent K x and K 2 . However, for aspartic and glutamic acids, A!)  and Kj are K x and A^, whereas for arginine, histidine, and lysine, these quan¬  tities are K R and K 2 (see Sample Calculation 4-1).   Of course, amino acid residues in the interior of a polypeptide chain do  not have free a-amino and a-carboxyl groups that can ionize (these groups    are joined in peptide bonds; Fig. 4-3). Furthermore, the pK values of all ion-  izable groups, including the N- and C-termini, usually differ from the pAwal-  ues listed in Table 4-1 for free amino acids. For example, the p K values of  a-carboxyl groups in unfolded proteins range from 3.5 to 4.0. In the free  amino acids, the p K values are much lower, because the positively charged am¬  monium group electrostatically stabilizes the COO group, in effect making  it easier for the carboxylic acid group to ionize. Similarly, the p K values for  a-amino groups in proteins range from 7.5 to 8.5. In the free amino acids,  the p K values are higher, due to the electron-withdrawing character of the  nearby carboxylate group, which makes it more difficult for the ammonium  group to become deprotonated. In addition, the three-dimensional structure  of a folded polypeptide chain may bring polar side chains and the N- and C-  termini close together. The resulting electrostatic interactions between these  groups may shift their pK values up to several pH units from the values for  the corresponding free amino acids. For this reason, the pi of a polypeptide,  which is a function of the p K values of its many ionizable groups, is not eas¬  ily predicted and is usually determined experimentally.   The three-letter abbreviations for the 20 standard amino acids given in Table  4-1 are widely used in the biochemical literature. Most of these abbreviations  are taken from the first three letters of the name of the corresponding amino  acid and are pronounced as written. The symbol Glx indicates Glu or Gin,  and similarly, Asx means Asp or Asn. This ambiguous notation stems from  laboratory experience: Asn and Gin are easily hydrolyzed to Asp and Glu, re¬  spectively, under the acidic or basic conditions often used to recover them  from proteins. Without special precautions, it is impossible to tell whether a  detected Glu was originally Glu or Gin, and likewise for Asp and Asn.   The one-letter symbols for the amino acids are also given in Table 4-1.  This more compact code is often used when comparing the amino acid se¬  quences of several similar proteins. Note that the one-letter symbol is usually  the first letter of the amino acids name. However, for sets of residues that  have the same first letter, this is true only of the most abundant residue of  the set.   Amino acid residues in polypeptides are named by dropping the suffix,  usually -ine, in the name of the amino acid and replacing it by -yl. Polypeptide  chains are described by starting at the N-terminus and proceeding to the  C-terminus. The amino acid at the C-terminus is given the name of its parent  amino acid. Thus, the compound   is called alanyltyrosylaspartylglycine. Obviously, such names for polypeptide  chains of more than a few residues are extremely cumbersome. The tetrapep-  tide above can also be written as Ala-Tyr-Asp-Gly using the three-letter abbre¬  viations, or AYDG using the one-letter symbols.   The various atoms of the amino acid side chains are often named in se¬  quence with the Greek alphabet, starting at the carbon atom adjacent to the   • Draw a generic amino acid and identify  the a carbon and its substituents.   • Draw the structures of the 20 standard  amino acids and give their one- and three-  letter abbreviations.   • Draw a Cys-Gly-Asn tripeptide. Identify  the peptide bond and the N- and  C-termini, and determine the peptide’s  net charge at neutral pH.   • Classify the 20 standard amino acids by  polarity, structure, type of functional group,  and acid-base properties.   • Why do the p K values of ionizable groups  differ between free amino acids and amino  acid residues in polypeptides?    FIG. 4-8 Greek nomenclature for amino acids. The carbon atoms are assigned  sequential letters in the Greek alphabet, beginning with the carbon next to the carbonyl group.    peptide carbonyl group. Therefore, as Fig. 4-8 indicates, the Lys residue is said  to have an e-amino group and Glu has a y-carboxyl group. Unfortunately, this  labeling system is ambiguous for several amino acids. Consequently, stan¬  dard numbering schemes for organic molecules are also employed (and are in¬  dicated in Table 4-1 for the heterocyclic side chains).    • Amino acids and many other biological compounds are chiral molecules whose  configurations can be depicted by Fischer projections.   • The amino acids in proteins all have the l stereochemical configuration.    With the exception of glycine, all the amino acids recovered from polypep¬  tides are optically active; that is, they rotate the plane of polarized light. The  direction and angle of rotation can be measured using an instrument known   Optically active molecules are asymmetric; that is, they are not superimpos-  able on their mirror image in the same way that a left hand is not superim-  posable on its mirror image, a right hand. This situation is characteristic of    Optically active  substance in solution  in the tube causes  the plane of the polarized  light to rotate.    Plane of polarization  of the emerging light  is not the same as  that of the entering  polarized light.    FIG. 4-9 Diagram of a polarimeter. This device is used to measure optical rotation.    substances containing tetrahedral carbon atoms that have four different sub¬  stituents. For example, the two molecules depicted in Fig. 4-10 are not su-  perimposable; they are mirror images. The central atoms in such molecules are  known as asymmetric centers or chiral centers and are said to have the prop¬  erty of chirality (Greek: cheir, hand). The C a atoms of the amino acids (ex¬  cept glycine) are asymmetric centers. Glycine, which has two H atoms attached  to its C a atom, is superimposable on its mirror image and is therefore not op¬  tically active. Many biological molecules in addition to amino acids contain  one or more chiral centers.   Chiral Centers Give Rise to Enantiomers. Molecules that are nonsuperim-  posable mirror images are known as enantiomers of one another.  Enantiomeric molecules are physically and chemically indistinguishable by  most techniques. Only when probed asymmetrically ; for example , by plane-  polarized light or by reactants that also contain chiral centers , can they be distin¬  guished or differentially manipulated.   Unfortunately, there is no clear relationship between the structure of a  molecule and the degree or direction to which it rotates the plane of polar¬  ized light. For example, leucine isolated from proteins rotates polarized light  10.4° to the left, whereas arginine rotates polarized light 12.5° to the right.  (The enantiomers of these compounds rotate polarized light to the same de¬  gree but in the opposite direction.) It is not yet possible to predict optical  rotation from the structure of a molecule or to derive the absolute  configuration (spatial arrangement) of chemical groups around a chiral center  from optical rotation measurements.   The Fischer Convention Describes the Configuration of Asymmetric  Centers. Biochemists commonly use the Fischer convention to describe  different forms of chiral molecules. In this system, the configuration of the  groups around an asymmetric center is compared to that of glyceraldehyde,  a molecule with one asymmetric center. In 1891, Emil Fischer proposed that  the spatial isomers, or stereoisomers, of glyceraldehyde be designated D-  glyceraldehyde and L-glyceraldehyde (Fig. 4-11). The prefix L (note the use  of a small uppercase letter) signified rotation of polarized light to the left  (Greek: levo , left), and the prefix D indicated rotation to the right (Greek: dex-  tro, right) by the two forms of glyceraldehyde. Fischer assigned the prefixes to  the structures shown in Fig. 4-11 without knowing whether the structure on  the left and the structure on the right were actually levorotatory or dextro¬  rotatory, respectively. Only in 1949 did experiments confirm that Fischers  guess was indeed correct.   Fischer also proposed a shorthand notation for molecular configurations,  known as Fischer projections, which are also given in Fig. 4-11. In the Fischer  convention, horizontal bonds extend above the plane of the paper and verti¬  cal bonds extend below the plane of the paper.   The configuration of groups around any chiral center can be related to  that of glyceraldehyde by chemically converting the groups to those of glyc¬  eraldehyde. For a-amino acids, the amino, carboxyl, R, and H groups around  the C a atom correspond to the hydroxyl, aldehyde, CH 2 OH, and H groups,  respectively, of glyceraldehyde.   Therefore, L-glyceraldehyde and L-a-amino acids are said to have the same rela¬  tive configuration. All amino acids derived from proteins have the L stereochemical   FIG. 4-1 O The two enantiomers of  fluorochlorobromomethane. The four  substituents are tetrahedrally arranged around the  central carbon atom. A dotted line indicates that  a substituent lies behind the plane of the paper,  a wedged line indicates that it lies above the  plane of the paper, and a thin line indicates that  it lies in the plane of the paper. The mirror plane  relating the enantiomers is represented by a  vertical dashed red line.   enantiomers of glyceraldehyde are shown as  geometric formulas (top) and as Fischer  projections (bottom). In a Fischer projection,  horizontal lines represent bonds that extend  above the page and vertical lines represent  bonds that extend below the page (in some  Fischer projections, the central chiral carbon  atom is not shown explicitly).    Draw Fischer projections for the two  enantiomers of alanine or use a molec¬  ular model kit to prove that the two  isomers are nonsuperimposable mirror  images.    A system to unambiguously describe the configurations of mole¬  cules with more than one asymmetric center was devised in 1956  by Robert Cahn, Christopher Ingold, and Vladimir Prelog. In the  Cahn-Ingold-Prelog or RS system, the four groups surrounding  a chiral center are ranked according to a specific although arbi¬  trary priority scheme: Atoms of higher atomic number rank above  those of lower atomic number (e.g., —OH ranks above —CH 3 ). If  the first substituent atoms are identical, the priority is established  by the next atom outward from the chiral center (e.g., —CH 2 0H  takes precedence over —CH 3 ). The order of priority of some  common functional groups is   SH > OH > NH 2 > COOH > CHO > CH 2 0H > C 6 H 5 > CH 3 > H   The prioritized groups are assigned the letters W, X, Y, Z such that  their order of priority ranking is W > X > Y > Z. To establish the  configuration of the chiral center, it is viewed from the asymmetric  center toward the Z group (lowest priority). If the order of the  groups W —» X —» Y is clockwise, the configuration is designated  R (Latin: rectus, right). If the order of W —» X —» Y is counter¬  clockwise, the configuration is designated S (Latin: sinistrus, left).    L-Glyceraldehyde is (S)-glyceraldehyde because the three  highest priority groups are arranged counterclockwise when the  H atom (dashed lines) is positioned behind the chiral C atom  (large circle).    All the L-amino acids in proteins are (S)-amino acids except  cysteine, which is (R)-cysteine because the S in its side chain  increases its priority. Other closely related compounds with the  same designation under the Fischer dl convention may have  different representations under the RS system. The RS system is  particularly useful for describing the chiralities of compounds with  multiple asymmetric centers. Thus, L-threonine can also be called  (2S,3R)-threonine.   configuration; that is, they all have the same relative configuration around their C a  atoms. Of course, the L or D designation of an amino acid does not indicate  its ability to rotate the plane of polarized light. Many L-amino acids are  dextrorotatory.   The Fischer system has some shortcomings, particularly for molecules with  multiple asymmetric centers. Each asymmetric center can have two possible  configurations, so a molecule with n chiral centers has 2 n different possible  stereoisomers. Threonine and isoleucine, for example, each have two chiral car¬  bon atoms, and therefore each has four stereoisomers, or two pairs of enan¬  tiomers. [The enantiomers (mirror images) of the L forms are the D forms.]  For most purposes, the Fischer system provides an adequate description of bi¬  ological molecules. A more precise nomenclature system is also occasionally  used by biochemists (see Box 4-2).   Life Is Based on Chiral Molecules. Consider the ordinary chemical synthe¬  sis of a chiral molecule, which produces a racemic mixture (containing equal  amounts of each enantiomer). In order to obtain a product with net asymme¬  try, a chiral process must be employed. One of the most striking characteris¬  tics of life is its production of optically active molecules. Biosynthetic processes  almost invariably produce pure stereoisomers. The fact that the amino acid  residues of proteins all have the L configuration is just one example of this  phenomenon. Furthermore, because most biological molecules are chiral, a  given molecule—present in a single enantiomeric form—will bind to or react  with only a single enantiomer of another compound. For example, a protein  made of L-amino acid residues that reacts with a particular L-amino acid does  not readily react with the D form of that amino acid. An otherwise identical  synthetic protein made of D-amino acid residues, however, readily reacts only  with the corresponding D-amino acid.   D-Amino acid residues are components of some relatively short (<20  residues) bacterial polypeptides. These polypeptides are perhaps most widely  distributed as constituents of bacterial cell walls (Section 8-3B). The presence  of the D-amino acids renders bacterial cell walls less susceptible to attack by  the peptidases (enzymes that hydrolyze peptide bonds) that are produced by   FIG. 4-1 2 Ibuprofen. Only the enantiomer shown has anti-inflammatory action. The  chiral carbon is red.   FIG. 4-1 3 Thalidomide. This drug was widely used in Europe as a mild sedative in  the early 1960s. Its inactive enantiomer (not shown), which was present in equal amounts  in the formulations used, causes severe birth defects in humans when taken during the  first trimester of pregnancy. Thalidomide was often prescribed to alleviate the nausea  (morning sickness) that is common during this period. In recent years, however, it was  found that thalidomide is an effective drug for the treatment of the immune system cancer  multiple myeloma.   other organisms to digest bacteria. Likewise, D-amino acids are components  of many bacterially produced peptide antibiotics. Most peptides containing D-  amino acids are not synthesized by the standard protein synthetic machinery,  in which messenger RNA is translated at the ribosome by transfer RNA mol¬  ecules with attached L- amino acids (Chapter 27). Instead, the D-amino acids  are directly joined together by the action of specific bacterial enzymes.   r The importance of stereochemistry in living systems is also a concern of  the pharmaceutical industry. Many drugs are chemically synthesized as  racemic mixtures , although only one enantiomer has biological activity : In most  cases, the opposite enantiomer is biologically inert and is therefore packaged  along with its active counterpart. This is true, for example, of the anti-inflam¬  matory agent ibuprofen, only one enantiomer of which is physiologically  active (Fig. 4-12). Occasionally, the inactive enantiomer of a useful drug  produces harmful effects and must therefore be eliminated from the  racemic mixture. The most striking example of this is the drug thalidomide  (Fig. 4-13), a mild sedative whose inactive enantiomer causes severe birth  defects. Partly because of the unanticipated problems caused by inactive  drug enantiomers, chiral organic synthesis has become an active area of  medicinal chemistry.   • Explain why all the amino acids except for  glycine are chiral.   • Identify all the chiral carbons in the amino  acids shown in Table 4-1.   • Explain how the Fischer convention  describes the absolute configuration  of a chiral molecule.   • Explain why an enzyme can catalyze a  chemical reaction involving just one  enantiomer of a compound.    The side chains of amino acid residues in proteins may be covalently modified.  Some amino acids and amino acid derivatives function as hormones and regula¬  tory molecules.    The 20 common amino acids are by no means the only amino acids that oc¬  cur in biological systems. “Nonstandard” amino acid residues are often impor¬  tant constituents of proteins and biologically active peptides. In addition,    many amino acids are not constituents of polypeptides at all but independ¬  ently play a variety of biological roles.   The “universal” genetic code, which is nearly identical in all known life-forms  (Section 27-1), specifies only the 20 standard amino acids of Table 4-1.  Nevertheless, many other amino acids, some of which are shown in Fig. 4-14,  are components of certain proteins. In almost all cases, these unusual amino acids  result from the specific modification of an amino acid residue after the polypeptide  chain has been synthesized.   Amino acid modifications include the simple addition of small chemical  groups to certain amino acid side chains: hydroxylation, methylation, acetyla¬  tion, carboxylation, and phosphorylation. Larger groups, including lipids and  carbohydrate polymers, are attached to particular amino acid residues of cer¬  tain proteins. The free amino and carboxyl groups at the N- and C-termini  of a polypeptide can also be chemically modified. These modifications are of¬  ten important, if not essential, for the function of the protein. In some cases,  several amino acid side chains together form a novel structure (Box 4-3).   The 20 standard amino acids undergo a bewildering number of chemical  transformations to other amino acids and related compounds as part of their  normal cellular synthesis and degradation. In a few cases, the intermediates of  amino acid metabolism have functions beyond their immediate use as precur¬  sors or degradation products of the 20 standard amino acids. Moreover, many  amino acids are synthesized not to be residues of polypeptides but to function  independently. We will see that many organisms use certain amino acids to  transport nitrogen in the form of amino groups (Section 21-2A). Amino acids  may also be oxidized as metabolic fuels to provide energy (Section 21-4). In    FIG. 4-14 Some modified amino acid residues in proteins. The side chains of  these residues are derived from one of the 20 standard amino acids after the polypeptide  has been synthesized. The standard R groups are red, and the modifying groups are blue.    Explain how each modification alters the polarity of the parent amino acid.    Genetic engineers often link a protein-coding gene to a “reporter  gene,” for example, the gene for an enzyme that yields a colored  reaction product. The intensity of the colored compound can be  used to estimate the level of expression of the engineered gene.  One of the most useful reporter genes is the one that codes for  green fluorescent protein (GFP). This protein, from the biolumi-  nescent jellyfish Aequorea victoria, fluoresces with a peak wave¬  length of 508 nm (green light) when irradiated by ultraviolet or  blue light (optimally 400 nm).   Green fluorescent protein is nontoxic and intrinsically fluores¬  cent; it requires no substrate or small molecule cofactor to fluo¬  resce as do other highly fluorescent proteins. Consequently, when  the gene for green fluorescent protein is linked to another gene,  the level of expression of the fused genes can be measured  noninvasively by fluorescence microscopy.   Green fluorescent protein consists of a chain of 238 amino  acid residues. The light-emitting group is a derivative of three  consecutive amino acids: Ser, Tyr, and Gly. After the protein has    been synthesized, the three amino acids undergo spontaneous  cyclization and oxidation. The carbonyl C atom of Ser forms a  covalent bond to the amino N atom contributed by Gly, followed  by the elimination of water and the oxidation of the C a —C p bond  of Tyr to a double bond. The resulting structure contains a system  of conjugated double bonds that gives the protein its fluorescent  properties.   Cyclization between Ser and Gly is probably rapid, and the oxida¬  tion of the Tyr side chain (by 0 2 ) is probably the rate-limiting step  of fluorophore generation. Genetic engineering has introduced  site-specific mutations that enhance fluorescence intensity and  shift the wavelength of the emitted light to different colors,  thereby making it possible to simultaneously monitor the expres¬  sion of two or more different genes.    addition, amino acids and their derivatives often function as chemical messen¬  gers for communication between cells (Fig. 4-15). For example, glycine, y-  aminobutyric acid (GABA; a glutamate decarboxylation product), and  dopamine (a tyrosine derivative) are neurotransmitters, substances released  by nerve cells to alter the behavior of their neighbors. Histamine (the decar-   FIG. 4-1 5 Some biologically active amino acid derivatives. The remaining portions  of the parent amino acids are black and red, and additional groups are blue.   Chapter 4 Amino Acids boxylation product of histidine) is a potent local mediator of allergic reactions.   Thyroxine (another tyrosine derivative) is an iodine-containing thyroid hor¬  mone that generally stimulates vertebrate metabolism.   Many peptides containing only a few amino acid residues have important  physiological functions as hormones or other regulatory molecules. One nearly  ubiquitous tripeptide called glutathione plays a role in cellular metabolism.  Glutathione is a Glu-Cys-Gly peptide in which the y-carboxylate group of  the glutamate side chain forms an isopeptide bond with the amino group of  the Cys residue (so called because a standard peptide bond is taken to be the  amide bond formed between an a-carboxylate and an a-amino group of two  amino acids). Two of these tripeptides (abbreviated GSH) undergo oxidation  of their SH groups to form a dimeric disulfide-linked structure called glu¬  tathione disulfide (GSSG):   • Cover the labels in Figs. 4-14 and 4-15  and identify each parent amino acid and  the type of chemical modification that has  occurred.   Glutathione helps inactivate oxidative compounds that could potentially dam¬  age cellular structures, since the oxidation of GSH to GSSG is accompanied  by the reduction of another compound (shown as 0 2 above):   • At neutral pH, the amino group of an amino acid is protonated  and its carboxylic acid group is ionized.   • Proteins are polymers of amino acids joined by peptide bonds.   • The 20 standard amino acids can be classified as nonpolar (Gly,  Ala, Val, Leu, lie, Met, Pro, Phe, Trp), uncharged polar (Ser, Thr,  Asn, Gin, Tyr, Cys), and charged (Lys, Arg, His, Asp, Glu).   • The p K values of the ionizable groups of amino acids may be  altered when the amino acid is part of a polypeptide.   • Amino acids are chiral molecules. Only l- amino acids are found  in proteins (some bacterial peptides contain D-amino acids).   • Amino acids may be covalently modified after they have been  incorporated into a polypeptide.   • Individual amino acids and their derivatives have diverse physio¬  logical functions.    1. Identify the amino acids that differ from each other by a single  methyl or methylene group.   2. Glutamate, a 5-carbon amino acid, is the precursor of three other  amino acids that contain a 5-carbon chain. Identify these amino  acids.   3. The 20 standard amino acids are called a-amino acids. Certain  (3-amino acids are found in nature. Draw the structure of  (3-alanine (3-amino-w-propionate).   (a) Explain why this designation is not valid, (b) From which of  the 20 standard amino acids is taurine derived? Describe the  chemical change(s) that occurred.   6. Calculate the number of possible pentapeptides that contain one  residue each of Ala, Gly, His, Lys, and Val.   7. Identify the hydrogen bond donor and acceptor groups in as¬  paragine.   8. In some proteins, the side chain of serine appears to undergo ion¬  ization. Explain why ionization would be facilitated by the pres¬  ence of an aspartate residue nearby.   9. A sample of the amino acid tyrosine is barely soluble in water.  Would a polypeptide containing only Tyr residues, poly(Tyr), be  more or less soluble, assuming the total number of Tyr groups  remains constant?   10. (a) What is the net charge at neutral pH of a tripeptide contain¬  ing only alanine? (b) How does the total number of negative and  positive charges change following hydrolysis of the tripeptide?   11. Determine the net charge of the predominant form of Asp at  (a) pH 1.0, (b) pH 3.0, (c) pH 6.0, and (d) pH 11.0.   12. Determine the net charge of the predominant form of Arg at   (a) pH 1.0, (b) pH 3.0, (c) pH 10.5, and (d) pH 13.5.   13. Calculate the p/of (a) Ala, (b) His, and (c) Glu.   14. Estimate the isoelectric point of a Ser-His dipeptide. Explain  why this value is only an estimate.    16. The protein insulin consists of two polypeptides termed the A  and B chains. Insulins from different organisms have been iso¬  lated and sequenced. Human and duck insulins have the same  amino acid sequence with the exception of six amino acid  residues, as shown below. Is the pi of human insulin lower than  or higher than that of duck insulin?    17. Indicate whether the following familiar objects are chiral or  nonchiral: (a) a glove; (b) a tennis ball; (c) a screw; (d) this page;  (e) a snowflake; (f) a spiral staircase; and (g) a shoe.   20. The two C a H atoms of Gly are said to be prochiral, because  when one of them is replaced by another group, C a becomes chi¬  ral. Draw a Fischer projection of Gly and indicate which H must  be replaced with CH 3 to yield D-Ala.   21. The bacterially produced antibiotic gramicidin A forms channels  in cell membranes that allow the free diffusion of Na + and K +  ions, thereby killing the cell. This peptide consists of a sequence  of D- and L-amino acids. The sequence of a segment of five amino  acids in gramicidin A is R-Gly-L-Ala-D-Leu-L-Ala-D-Val-R'.  Complete the Fischer projection below by adding the correct  group to each vertical bond.   22. Describe isoleucine (as shown in Table 4-1) using the  RS system.   23. Some amino acids are synthesized by replacing the keto group  (C=0) of an organic acid known as an a-keto acid with an  amino group (C—NH^). Identify the amino acids that can be  produced this way from the following a-keto acids:   24. Patients with Parkinson’s disease are sometimes given l-DOPA  (see page 76). What neurotransmitter is produced by the decar¬  boxylation of l-DOPA?   23. Identify the amino acid residue from which the following groups  are synthesized:    26. Describe how each of the amino acid modifications shown in Fig.  4-14 would affect the p/of a peptide containing that modified  amino acid residue.   27. Which amino acids have side chains that are capable of forming  isopeptide bonds?   28. Draw the structure of Lys and Ala linked by an isopeptide bond.   Presumably, the mix of prebi-  otic compounds on early earth included both d- and L-amino acids.  What are some hypotheses that attempt to explain why cells build  proteins only from the l forms?    Barrett, G.C. and Elmore, D.T., Amino Acids and Peptides, Cambridge Solomons, G.T.W. and Fryhle, C., Organic Chemistry (10th ed.), Chap-  University Press (2001). [Includes structures of the common amino ter 3, Wiley (2011). [A discussion of chirality. Most other organic   acids along with a discussion of their chemical reactivities and chemistry textbooks contain similar material.]   Lamzin, V.S., Dauter, Z., and Wilson, K.S., How nature deals with  stereoisomers, Curr. Opin. Struct. Biol. 5 , 830-836 (1995). [Dis¬  cusses proteins synthesized from D-amino acids.]    The great variation in structure and function among proteins reflects the astronomical  variation in the sequences of their component amino acids: There are far more  possible amino acid sequences than there are stars in the universe. [PhotoDisc,  Inc./Getty Images.]    Proteins are at the center of action in biological processes. Nearly all the mo¬  lecular transformations that define cellular metabolism are mediated by pro¬  tein catalysts. Proteins also perform regulatory roles, monitoring extracellular  and intracellular conditions and relaying information to other cellular compo¬  nents. In addition, proteins are essential structural components of cells. A com¬  plete list of known protein functions would contain many thousands of entries,  including proteins that transport other molecules and proteins that generate  mechanical and electrochemical forces. And such a list would not account for  the thousands of proteins whose functions are not yet fully characterized or,  in many cases, are completely unknown.   One of the keys to deciphering the function of a given protein is to un¬  derstand its structure. Like the other major biological macromolecules, the nu¬  cleic acids (Section 3-2) and the polysaccharides (Section 8-2), proteins are  polymers of smaller units. But unlike many nucleic acids, proteins do not have  uniform, regular structures. This is, in part, because the 20 kinds of amino  acid residues from which proteins are made have widely differing chemical and  physical properties (Section 4-1C). The sequence in which these amino acids  are strung together can be analyzed directly, as we describe in this chapter, or  indirectly, via DNA sequencing (Section 3-4). In either case, amino acid se¬  quence information provides insights into the chemical and physical proper¬  ties of proteins, their relationships to other proteins, and ultimately, their  mechanisms of action in living organisms. After a brief introduction to the va¬  riety in protein structure we will examine some methods for purifying and an¬  alyzing proteins, procedures for determining the sequence of amino residues,  and finally, some approaches to understanding protein evolution.   A The First Step Is to Separate Subunits  B The Polypeptide Chains Are Cleaved  C Edman Degradation Removes a Peptide’s First  Amino Acid Residue   B Proteins Evolve by the Duplication of Genes or  Gene Segments    • In theory, the size and composition of a polypeptide chain are unlimited.   • In cells, this potential variety is limited by the efficiency of protein synthesis and  by the ability of the polypeptide to fold into a functional structure.    Like all polymeric molecules, proteins can be described in terms of levels of  organization, in this case, their primary, secondary, tertiary, and quaternary  structures. A proteins primary structure is the amino acid sequence of its  polypeptide chain. , or chains if the protein consists of more than one polypep¬  tide. An example of an amino acid sequence is given in Fig. 5-1. Each residue  is linked to the next via a peptide bond (Fig. 4-3). Higher levels of protein  structure—secondary, tertiary, and quaternary—refer to the three-dimensional  shapes of folded polypeptide chains and will be described in the next chapter.   Proteins are synthesized in vivo by the stepwise polymerization of amino  acids in the order specified by the sequence of nucleotides in a gene. The  direct correspondence between one linear polymer (DNA) and another (a  polypeptide) illustrates the elegant simplicity of living systems and allows us  to extract information from one polymer and apply it to the other.    The Theoretical Possibilities for Polypeptides Are Unlimited. With 20 dif¬  ferent choices available for each amino acid residue in a polypeptide chain, it  is easy to see that a huge number of different protein molecules are possible.  For a protein of n residues, there are 20” possible sequences. A relatively small  protein molecule may consist of a single polypeptide chain of 100 residues.  There are 20 100 ~ 1.27 X 10 130 possible unique polypeptide chains of this  length, a quantity vastly greater than the estimated number of atoms in the  universe (9 X 10 78 ). Clearly, evolution has produced only a tiny fraction of  the theoretical possibilities—a fraction that nevertheless represents an astro¬  nomical number of different polypeptides.   general, proteins contain at least 40 residues or so; polypeptides smaller than  that are simply called peptides. The largest known polypeptide chain belongs  to the 34,350-residue titin, a giant (3816 kD) protein that helps arrange the  repeating structures of muscle fibers (Section 7-2A). However, the vast major¬  ity of polypeptides contain between 100 and 1000 residues (Table 5-1).  Multisubunit proteins contain several identical and/or nonidentical chains  called subunits. Some proteins are synthesized as single polypeptides that are  later cleaved into two or more chains that remain associated; insulin is such  a protein (Fig. 5-1).   The size range in which most polypeptides fall probably reflects the opti¬  mization of several biochemical processes:   1. Forty residues appears to be near the minimum for a polypeptide chain  to fold into a discrete and stable shape that allows it to carry out a  particular function.   2. Polypeptides with well over 1000 residues may approach the limits of  efficiency of the protein synthetic machinery. The longer the polypep-    Val - Glu - Gin - Cys - Cys - Ala - Ser - Val - Cys - Ser - Leu - Tyr - Gin - Leu - Glu - Asn - Tyr - Cys - Asn    FIG. 5-1 The primary structure of bovine insulin. Note the intrachain and interchain  disulfide bond linkages.    tide (and the longer its corresponding mRNA), the greater the likeli¬  hood of introducing errors during transcription and translation.   In addition to these mild constraints on size, polypeptides are subject to  more severe limitations on amino acid composition. The 20 standard amino  acids do not appear with equal frequencies in proteins. (Table 4-1 lists the av¬  erage occurrence of each amino acid residue.) For example, the most abun¬  dant amino acids in proteins are Leu, Ala, Gly, Val, Glu, and Ser; the rarest  are Trp, Cys, Met, and His.   Because each amino acid residue has characteristic chemical and physical  properties, its presence at a particular position in a protein influences the prop¬  erties of that protein. In particular, as we will see, the three-dimensional shape  of a folded polypeptide chain is a consequence of the intramolecular forces  among its various residues. In general, a proteins hydrophobic residues clus¬  ter in its interior, out of contact with water, whereas its hydrophilic side chains  tend to occupy the proteins surface.   The characteristics of an individual protein depend more on its amino  acid sequence than on its amino acid composition per se, for the same reason  that “kitchen” and its anagram “thicken” are quite different words. In addi¬  tion, many proteins consist of more than just amino acid residues. They may  form complexes with metal ions such as Zn 2+ and Ca 2+ , they may covalently  or noncovalently bind certain small organic molecules, and they may be co¬  valently modified by the posttranslational attachment of groups such as phos¬  phates and carbohydrates.   • What factors limit the size and composi¬  tions of polypeptides?    Environmental conditions such as pH and temperature affect a protein’s stability  during purification.   An assay based on a protein’s chemical or binding properties may be used to  quantify a protein during purification.   Fractionation procedures take advantage of a protein’s unique structure and  chemistry in order to separate it from other molecules.   FIG. 5-2 Inclusion body. A genetically  engineered organism that produces large  amounts of a foreign protein often sequesters it  in inclusion bodies. This electron micrograph  shows an inclusion body of the protein prochy-  mosin in an E. coli cell. [Courtsey of Teruhiko  Beppu, Nikon University, Japan.]    • Increasing the salt concentration causes selective “salting out" (precipitation) of  proteins with different solubilities.   • A protein’s ionic charge, polarity, size, and ligand-binding ability influence its  chromatographic behavior.   • Gel electrophoresis and its variations can separate proteins according to charge,  size, and isoelectric point.   • The overall size and shape of macromolecules and larger assemblies can be  assessed through ultracentrifugation.   Purification is an all but mandatory step in studying macromolecules, but it  is not necessarily easy Typically, a substance that makes up <0.1% of a tis¬  sues dry weight must be brought to ~98% purity. Purification problems of  this magnitude would be considered unreasonably difficult by most synthetic  chemists! The following sections outline some of the most common techniques  for purifying and, to some extent, characterizing proteins. Most of these tech¬  niques can be used, sometimes in modified form, for nucleic acids and other  types of biological molecules.   The task of purifying a protein present in only trace amounts was once so  arduous that many of the earliest proteins to be characterized were studied in  part because they are abundant and easily isolated. For example, hemoglobin,  which accounts for about one-third the weight of red blood cells, has histori¬  cally been among the most extensively studied proteins. Most of the enzymes  that mediate basic metabolic processes or that are involved in the expression and  transmission of genetic information are common to all species. For this reason,  a given protein is frequently obtained from a source chosen primarily for con¬  venience, for example, tissues from domesticated animals or easily obtained  microorganisms such as E. coli and Saccharomyces cerevisiae (bakers yeast).   Molecular cloning techniques (Section 3-5) allow almost any protein¬  encoding gene to be isolated from its parent organism, specifically altered (ge¬  netically engineered) if desired, and expressed at high levels in a microorganism.  Indeed, the cloned protein may constitute up to 40% of the microorganisms  total cell protein (Fig. 5-2). This high level of protein production generally ren¬  ders the cloned protein far easier to isolate than it would be from its parent  organism (in which it may occur in vanishingly small amounts).   The first step in the isolation of a protein or other biological molecule is  to get it out of the cell and into solution. Many cells require some sort of me¬  chanical disruption to release their contents. Most of the procedures for lysing  cells use some variation of crushing or grinding followed by filtration or cen¬  trifugation to remove large, insoluble particles. If the target protein is tightly  associated with a lipid membrane, a detergent or organic solvent may be used  to solubilize the lipids and recover the protein.   pH, Temperature, and Other Conditions Must Be Controlled to Keep  Proteins Stable. Once a protein has been removed from its natural environ¬  ment, it becomes exposed to many agents that can irreversibly damage it.  These influences must be carefully controlled at all stages of a purification  process. The following factors should be considered:   1. pH. Biological materials are routinely dissolved in buffer solutions ef¬  fective in the pH range over which the materials are stable (buffers are  described in Section 2-2C). Failure to do so could cause their denat-  uration (structural disruption), if not their chemical degradation.   2. Temperature. The thermal stability of proteins varies. Although some  proteins denature at low temperatures, most proteins denature at high  temperatures, sometimes only a few degrees higher than their native  environment. Protein purification is normally carried out at tempera¬  tures near 0°C.    3. Presence of degmdative enzymes . Destroying tissues to liberate the  molecule of interest also releases degradative enzymes, including pro¬  teases and nucleases. Degradative enzymes can be inhibited by adjust¬  ing the pH or temperature to values that inactivate them (provided this  does not adversely affect the protein of interest) or by adding com¬  pounds that specifically block their action.   4. Adsorption to surfaces . Many proteins are denatured by contact with  the air-water interface or with glass or plastic surfaces. Hence, protein  solutions are handled so as to minimize foaming and are kept relatively  concentrated.   5. Long-term storage . All the factors listed above must be considered  when a purified protein sample is to be kept stable. In addition,  processes such as slow oxidation and microbial contamination must be  prevented. Protein solutions are sometimes stored under nitrogen or  argon gas (rather than under air, which contains ~21% 0 2 ) and/or  are frozen at — 80°C or — 196°C (the temperature of liquid nitrogen).   Proteins Are Quantified by Assays. Purifying a substance requires some  means for quantitatively detecting it. Accordingly, an assay must be devised  that is specific for the target protein, highly sensitive, and convenient to use  (especially if it must be repeated at every stage of the purification process).   Among the most straightforward protein assays are those for enzymes that  catalyze reactions with readily detected products, because the rate of product  formation is proportional to the amount of enzyme present. Substances with col¬  ored or fluorescent products have been developed for just this purpose. If no  such substance is available for the enzyme being assayed, the product of the  enzymatic reaction may be converted, by the action of another enzyme, to an  easily quantified substance. This is known as a coupled enzymatic reaction.  Proteins that are not enzymes can be detected by their ability to specifically  bind certain substances or to produce observable biological effects.   Immunochemical procedures are among the most sensitive of assay tech¬  niques. Immunoassays use antibodies, proteins produced by an animals im¬  mune system in response to the introduction of a foreign substance (an  antigen). Antibodies recovered from the blood serum of an immunized ani¬  mal or from cultures of immortalized antibody-producing cells bind specifi¬  cally to the original protein antigen.   A protein in a complex mixture can be detected by its binding to its cor¬  responding antibodies. In one technique, known as a radioimmunoassay  (RIA), the protein is indirectly detected by determining the degree to which  it competes with a radioactively labeled standard for binding to the antibody.  Another technique, the enzyme-linked immunosorbent assay (ELISA), has  many variations, one of which is diagrammed in Fig. 5-3.   Protein Concentrations Can Be Determined by Spectroscopy. The concen¬  tration of a substance in solution can be measured by absorbance spec¬  troscopy. A solution containing a solute that absorbs light does so according  to the Beer-Lambert law,   where A is the solutes absorbance (alternatively, its optical density), 7 0 is the  intensity of the incident light at a given wavelength X, / is its transmitted in¬  tensity at X, s is the absorptivity (alternatively, the extinction coefficient) of  the solute at X, c is its concentration, and / is the length of the light path in  centimeters. The value of s varies with X; a plot of A or s versus X for the  solute is called its absorption spectrum. If the value of s for a substance is  known, then its concentration can be spectroscopically determined.   FIG. 5-3 Enzyme-linked immunosorbent  assay. (1) An antibody against the protein of  interest is immobilized on an inert solid such as  polystyrene. (2) The solution to be assayed is  applied to the antibody-coated surface. The  antibody binds the protein of interest, and other  proteins are washed away. (3) The protein-  antibody complex is reacted with a second  protein-specific antibody to which an enzyme is  attached. (4) Binding of the second antibody-  enzyme complex is measured by assaying the  activity of the enzyme. The amount of substrate  converted to product indicates the amount of  protein present. Q See the Animated  Figures.   Here the molar absorptivity (s when c is  expressed in mol • L -1 ) for each aromatic amino  acid is displayed on a log scale. [After Wetlaufer,  D.B., Adv. Prot. Chem. 7, 310 (1962).]    Polypeptides absorb strongly in the ultraviolet (UV) region of the spectrum  (X = 200 to 400 nm) largely because their aromatic side chains (those of Phe,  Trp, and Tyr) have particularly large extinction coefficients in this spectral  region (ranging into the tens of thousands when c is expressed in mol • L  Fig. 5-4). However, polypeptides do not absorb visible light (X = 400 to 800  nm) so that they are colorless. Nevertheless, if a protein has a chromophore  that absorbs in the visible region of the spectrum, this absorbance can be used  to assay for the presence of the protein in a mixture of other proteins.   Purification Is a Stepwise Process. Proteins are purified by fractionation  procedures. In a series of independent steps, the various physicochemical  properties of the protein of interest are used to separate it progressively from  other substances. The idea is not necessarily to minimize the loss of the de¬  sired protein, but to eliminate selectively the other components of the mixture so  that only the required substance remains.   Protein purification is considered as much an art as a science, with many  options available at each step. While a trial-and-error approach can work,  knowing something about the target protein (or the proteins it is to be sepa¬  rated from) simplifies the selection of fractionation procedures. Some of the  procedures we discuss and the protein characteristics they depend on are listed  in Table 5-2.    Because a protein contains multiple charged groups, its solubility depends on  the concentrations of dissolved salts, the polarity of the solvent, the pH, and  the temperature. Some or all of these variables can be manipulated to selec¬  tively precipitate certain proteins while others remain soluble.   The solubility of a protein at low ion concentrations increases as salt is  added, a phenomenon called salting in. The additional ions shield the pro¬  teins multiple ionic charges, thereby weakening the attractive forces between  individual protein molecules (such forces can lead to aggregation and precip¬  itation). However, as more salt is added, particularly with sulfate salts, the sol¬  ubility of the protein again decreases. This salting out effect is primarily a  result of the competition between the added salt ions and the other dissolved  solutes for molecules of solvent. At very high salt concentrations, so many of  the added ions are solvated that there is significantly less bulk solvent avail¬  able to dissolve other substances, including proteins.   Since different proteins have different ionic and hydrophobic composi¬  tions and therefore precipitate at different salt concentrations, salting out is  the basis of one of the most commonly used protein purification procedures.  Adjusting the salt concentration in a solution containing a mixture of proteins  to just below the precipitation point of the protein to be purified eliminates  many unwanted proteins from the solution (Fig. 5-5). Then, after removing    FIG. 5-5 Fractionation by salting out. (a) The salt of choice, usually ammonium  sulfate, is added to a solution of macromolecules to a concentration just below the  precipitation point of the protein of interest. ( b ) After centrifugation, the unwanted  precipitated proteins (red spheres ) are discarded and more salt is added to the supernatant  to a concentration sufficient to salt out the desired protein (green spheres), (c) After a  second centrifugation, the protein is recovered as a precipitate, and the supernatant is  discarded.    the precipitated proteins by filtration or centrifugation, the salt concentration  of the remaining solution is increased to precipitate the desired protein. This  procedure results in a significant purification and concentration of large quan¬  tities of protein. Ammonium sulfate, (NH^SC^, is the most commonly used  reagent for salting out proteins because its high solubility (3.9 M in water at  0°C) allows the preparation of solutions with high ionic strength. The pH may  be adjusted to approximate the isoelectric point (p/) of the desired protein  because a protein is least soluble when its net charge is zero. The p/s of some  proteins are listed in Table 5-3.   The process of chromatography (Greek: chroma , color + graphein , to write)  was discovered in 1903 by Mikhail Tswett, who separated solubilized plant  pigments using solid adsorbents. In most modern chromatographic proce¬  dures, a mixture of substances to be fractionated is dissolved in a liquid  (the “mobile” phase) and percolated through a column containing a porous  solid matrix (the “stationary” phase). As solutes flow through the column,  they interact with the stationary phase and are retarded. The retarding force  depends on the properties of each solute. If the column is long enough, sub¬  stances with different rates of migration will be separated. The chromato¬  graphic procedures that are most useful for purifying proteins are classified  according to the nature of the interaction between the protein and the sta¬  tionary phase.   Early chromatographic techniques used strips of filter paper as the sta¬  tionary phase, whereas modern column chromatography uses granular de¬  rivatives of cellulose, agarose, or dextran (all carbohydrate polymers) or  synthetic substances such as cross-linked polyacrylamide or silica. High-  performance liquid chromatography (HPLC) employs automated sys¬  tems with precisely applied samples, controlled flow rates at high pressures  (up to 5000 psi), a chromatographic matrix of specially fabricated 3- to  300-fJim-diameter glass or plastic beads coated with a uniform layer of  chromatographic material, and on-line sample detection. This greatly im¬  proves the speed, resolution, and reproducibility of the separation—fea¬  tures that are particularly desirable when chromatographic separations are  repeated many times or when they are used for analytical rather than  preparative purposes.   Ion Exchange Chromatography Separates Anions and Cations. In ion ex¬  change chromatography, charged molecules bind to oppositely charged  groups that are chemically linked to a matrix such as cellulose or agarose.  Anions bind to cationic groups on anion exchangers, and cations bind to an¬  ionic groups on cation exchangers. Perhaps the most frequently used anion  exchanger is a matrix with attached diethylaminoethyl (DEAE) groups, and  the most frequently used cation exchanger is a matrix bearing carboxymethyl  (CM) groups.   Proteins and other polyelectrolytes (polyionic polymers) that bear both  positive and negative charges can bind to both cation and anion exchangers.  The binding affinity of a particular protein depends on the presence of other ions  that compete with the protein for binding to the ion exchanger and on the pH of  the solution ., which influences the net charge of the protein.   The proteins to be separated are dissolved in a buffer of an appropriate  pH and salt concentration and are applied to a column containing the ion    FIG. 5-6 Ion exchange chromatography. The tan region of the  column represents the ion exchanger and the colored bands represent  proteins, (a) A mixture of proteins dissolved in a small volume of  buffer is applied to the top of the matrix in the column, (b) As elution  progresses, the proteins separate into discrete bands as a result of  their different affinities for the exchanger. In this diagram, the first  protein (red) has passed through the column and has been isolated as    a separate fraction. The other proteins pass through the column more  slowly, (c and d) The salt concentration in the eluant is increased to  elute the remaining proteins, (e) The elution diagram of the protein  mixture from the column. Q See the Animated Figures.   How could ion exchange be used to concentrate a dilute  solution of protein?    exchanger. The column is then washed with the buffer (Fig. 5-6). As the col¬  umn is washed, proteins with relatively low affinities for the ion exchanger  move through the column faster than proteins that bind with higher affini¬  ties. The column effluent is collected in a series of fractions. Proteins that bind  tightly to the ion exchanger can be eluted (washed through the column) by  applying a buffer, called the eluant, that has a higher salt concentration or a  pH that reduces the affinity with which the matrix binds the protein. The col¬  umn effluent can be monitored for the presence of protein by measuring its  absorbance at 280 nm. The eluted fractions can also be tested for the protein  of interest using a more specific assay.   Hydrophobic interactions between proteins and the chromatographic matrix  can be exploited to purify the proteins. In hydrophobic interaction chro¬  matography, the matrix material is lightly substituted with octyl or phenyl  groups. At high salt concentrations, nonpolar groups on the surface of pro¬  teins “interact” with the hydrophobic groups; that is, both types of groups are  excluded by the polar solvent (hydrophobic effects are augmented by increased  ionic strength). The eluant is typically an aqueous buffer with decreasing salt  concentrations, increasing concentrations of detergent (which disrupts hy¬  drophobic interactions), or changes in pH.   Gel Filtration Chromatography Separates Molecules According to Size. In Section 2 Protein Purification and Analysis   gel filtration chromatography (also called size exclusion or molecular sieve  chromatography), molecules are separated according to their size and shape.   The stationary phase consists of gel beads containing pores that span a rela¬  tively narrow size range. The pore size is typically determined by the extent  of cross-linking between the polymers of the gel material. If an aqueous solu¬  tion of molecules of various sizes is passed through a column containing such  “molecular sieves,” the molecules that are too large to pass through the pores  are excluded from the solvent volume inside the gel beads. These large mole¬  cules therefore traverse the column more rapidly than small molecules that pass  through the pores (Fig. 5-7). Because the pore size in any gel varies to some  degree, gel filtration can be used to separate a range of molecules; larger mol¬  ecules with access to fewer pores elute sooner (i.e., in a smaller volume of elu¬  ant) than smaller molecules that have access to more of the gels interior volume.   Within the size range of molecules separated by a particular pore size,  there is a linear relationship between the relative elution volume of a substance  and the logarithm of its molecular mass (assuming the molecules have similar  shapes). If a given gel filtration column is calibrated with several proteins of  known molecular mass, the mass of an unknown protein can be conveniently  estimated by its elution position.   FIG. 5-7 Gel filtration chromatography, (a) A gel bead consists  of a gel matrix (wavy solid lines) that encloses an internal solvent  space. Small molecules (red dots ) can freely enter the internal space  of the gel bead. Large molecules (blue dots ) cannot penetrate the gel  pores, (b) The sample solution is applied to the top of the column (the  gel beads are represented as tan spheres), (c) The small molecules  can penetrate the gel and consequently migrate through the column  more slowly than the large molecules that are excluded from the gel.    (d and e) The large molecules elute first and are collected as fractions.  Small molecules require a larger volume of solvent to elute.  The  elution diagram, or chromatogram, indicating the complete separation    How could gel filtration be used to reduce the salt concen¬  tration of a protein solution?    FIG. 5-8 Affinity chromatography. A ligand  (shown here as black dots) is immobilized by  covalently binding it to the chromatographic  matrix. The orange, blue, and green shapes  represent macromolecules whose cutout regions  symbolize their ligand-binding sites. Only certain  molecules (represented by orange circles )  specifically bind to the ligand. The other  components are washed through the column.    Affinity Chromatography Exploits Specific Binding Behavior. A striking  characteristic of many proteins is their ability to bind specific molecules tightly  but noncovalently. This property can be used to purify such proteins by affin¬  ity chromatography (Fig. 5-8). In this technique, a molecule (a ligand) that  specifically binds to the protein of interest (e.g., a nonreactive analog of an  enzyme’s substrate) is covalently attached to an inert matrix. When an impure  protein solution is passed through this chromatographic material, the desired pro¬  tein binds to the immobilized ligand, whereas other substances are washed through  the column with the buffer. The desired protein can then be recovered in highly  purified form by changing the elution conditions to release the protein from  the matrix. The great advantage of affinity chromatography is its ability to ex¬  ploit the desired proteins unique biochemical properties rather than the small  differences in physicochemical properties between proteins exploited by other  chromatographic methods. Accordingly, the separation power of affinity chro¬  matography for a specific protein is often greater than that of other chromato¬  graphic techniques.   Affinity chromatography columns can be constructed by chemically at¬  taching small molecules or proteins to a chromatographic matrix. In im-  munoaffinity chromatography, an antibody is attached to the matrix in order  to purify the protein against which the antibody was raised. In all cases, the  ligand must have an affinity high enough to capture the protein of interest  but not so high as to prevent the proteins subsequent release without dena¬  turing it. The bound protein can be eluted by washing the column with a so¬  lution containing a high concentration of free ligand or a solution of different  pH or ionic strength.   In metal chelate affinity chromatography, a divalent metal ion such as  Zn 2+ or Ni 2+ is attached to the chromatographic matrix so that proteins bear¬  ing metal-chelating groups (e.g., multiple His side chains) can be retained.  Recombinant DNA techniques (Section 3-5) can be used to append a segment  of six consecutive His residues, known as a His tag, to the N- or C-terminus  of the polypeptide to be isolated. This creates a metal ion-binding site that  allows the recombinant protein to be purified by metal chelate chromatogra¬  phy. After the protein has been eluted, usually by altering the pH, the His tag  can be removed by the action of a specific protease whose recognition sequence  separates the (His)sequence from the rest of the protein.   Electrophoresis, the migration of ions in an electric field, is described in  Section 3-4B. Polyacrylamide gel electrophoresis (PAGE) of proteins is typ¬  ically carried out in polyacrylamide gels with a characteristic pore size, so the  molecular separations are based on gel filtration (size and shape) as well as elec¬  trophoretic mobility (electric charge). However, electrophoresis differs from gel  filtration in that the electrophoretic mobility of smaller molecules is greater  than the mobility of larger molecules with the same charge density. The pH  of the gel is high enough (usually about pH 9) so that nearly all proteins have  net negative charges and move toward the positive electrode when the current  is switched on. Molecules of similar size and charge move as a band through  the gel.   Following electrophoresis, the separated bands may be visualized in the gel  by an appropriate technique, such as soaking the gel in a solution of a stain  that binds tightly to proteins. If the proteins in a sample are radioactive, the  gel can be dried and then clamped over a sheet of X-ray film. After a time,  the film is developed and the resulting autoradiograph shows the positions  of the radioactive components by a blackening of the film. If an antibody to    1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17    FIG. 5-9 SDS-PAGE. Samples of proteins  were electrophoresed from top  to bottom   in parallel lanes in a polyacrylamide slab, which  was then stained to reveal the proteins. Lanes 1,  2, 10, and 17 contain molecular mass standards  whose molecular masses are indicated on the  left. [Image courtesy of Thermo Scientific Pierce  Protein Research Products.]    a protein of interest is available, it can be used to specifically detect the pro¬  tein on a gel in the presence of many other proteins, a process called  immunoblotting or Western blotting that is similar to ELISA (Fig. 5-3).  Depending on the dimensions of the gel and the visualization technique used,  samples containing less than a nanogram of protein can be separated and de¬  tected by gel electrophoresis.   SDS-PAGE Separates Proteins by Mass. In one form of polyacrylamide gel  electrophoresis, the detergent sodium dodecyl sulfate (SDS)   [CH 3 (CH 2 ) 10 CH 2 O SO 3 ]Na +   is added to denature proteins. Amphiphilic molecules (Section 2 - 1 C) such as  SDS interfere with the hydrophobic interactions that normally stabilize pro¬  teins. Proteins assume a rodlike shape in the presence of SDS. Furthermore,  most proteins bind SDS in a ratio of about 1.4 g SDS per gram protein (about  one SDS molecule for every two amino acid residues). The large negative  charge that the SDS imparts masks the proteins’ intrinsic charge. The net re¬  sult is that SDS-treated proteins have similar shapes and charge-to-mass ra¬  tios. SDS-PAGE therefore separates proteins purely by gel filtration effects , that  is, according to molecular mass. Figure 5-9 shows examples of the resolving  power and the reproducibility of SDS-PAGE.   In SDS-PAGE, the relative mobilities of proteins vary approximately lin¬  early with the logarithm of their molecular masses (Fig. 5-10). Consequently,  the molecular mass of a protein can be determined with about 5 to 10 % ac¬  curacy by electrophoresing it together with several “marker” proteins of known  molecular masses that bracket that of the protein of interest. Because SDS dis¬  rupts noncovalent interactions between polypeptides, SDS-PAGE yields the  molecular masses of the subunits of multisubunit proteins. The possibility that  subunits are linked by disulfide bonds can be tested by preparing samples for  SDS-PAGE in the presence and absence of a reducing agent, such as 2 -  mercaptoethanol (HSCH 2 CH 2 OH), that breaks those bonds (Section 5-3A).   Although gel electrophoresis in its various forms is highly effective at separat¬  ing charged molecules, it can require up to several hours and is difficult to  quantitate and automate. These disadvantages are largely overcome through  the use of capillary electrophoresis (CE), a technique in which electrophore¬  sis is carried out in very thin capillary tubes (20- to 100-fJim inner diameter).  Such narrow capillaries rapidly dissipate heat and hence permit the use of very  high electric fields, which reduces separation times to a few minutes. The CE   FIG. 5-10 Logarithmic relationship  between the molecular mass of a protein  and its electrophoretic mobility in SDS-  PAGE. The masses of 37 proteins ranging  from 11 to 70 kD are plotted. [After Weber, K.  and Osborn, M., J. Biol. Chem. 244, 4406  (1969).]    techniques have extremely high resolution and can be automated in much the  same way as is HPLC, that is, with automatic sample loading and on-line sam¬  ple detection. Since CE can separate only small amounts of material, it is  largely limited to use as an analytical tool.   A protein has charged groups of both polarities and therefore has an isoelectric  point, pi, at which it is immobile in an electric field. If a mixture of proteins is  electrophoresed through a solution or gel that has a stable pH gradient in which the  pH smoothly increases from anode to cathode, each protein will migrate to the po¬  sition in the pH gradient corresponding to its pi If a protein molecule diffuses  away from this position, its net charge will change as it moves into a region of  different pH and the resulting electrophoretic forces will move it back to its iso¬  electric position. Each species of protein is thereby “focused” into a narrow band  about its pi. This type of electrophoresis is called isoelectric focusing (IEF).   IEF can be combined with SDS-PAGE in an extremely powerful separation  technique named two-dimensional (2D) gel electrophoresis. First, a sample of  proteins is subjected to IEF in one direction, and then the separated proteins  are subjected to SDS-PAGE in the perpendicular direction. This procedure gen¬  erates an array of spots, each representing a protein (Fig. 5-11). Up to 5000  proteins have been observed on a single two-dimensional electrophoretogram.   Two-dimensional gel electrophoresis is a valuable tool for proteomics, a  field of study that involves cataloguing all of a cells expressed proteins with  emphasis on their quantitation, localization, modifications, interactions, and  activities. Individual protein spots in a stained 2D gel can be excised with a  scalpel, destained, and the protein eluted from the gel fragment for identifi¬  cation and/or characterization, often by mass spectrometry (Section 5-3D). 2D  electrophoretograms can be analyzed by computer after they have been  scanned and digitized. This facilitates the detection of variations in the posi¬  tions and intensities of protein spots in samples obtained from different tis¬  sues or under different growth conditions. Numerous reference 2D gels are  publicly available for this purpose in the Web-accessible databases listed at  http://us.expasy.org/ch2d/. These databases contain images of 2D gels of a va¬  riety of organisms and tissues and identify many of their component proteins.  Their use is illustrated in Bioinformatics Project 2.   Macromolecules in solution do not respond to the earths gravity by settling,  because their random thermal (Brownian) motion keeps them uniformly    FIG. 5-1 1 Two-dimensional gel  electrophoresis. In this example, E. coli proteins  that had been labeled with 14 C-amino acids  were subjected to isoelectric focusing  (horizontally) followed by SDS-PAGE (vertically).  Over 1000 spots can be resolved in the  autoradiogram shown here. [Courtesy of Patrick  O' Farrell, University of California at San  Francisco.]   FIG. 5-1 2 Ultracentrifugation. The sample is layered  onto a preformed sucrose density gradient (left). During centrifu¬  gation (middle), each particle sediments at a rate that depends  largely on its mass. After centrifugation, the tube is punctured  and the separated particles are collected (right). In some cases,  the centrifuge tube is filled with a sample solution containing  CsCI, which forms a density gradient as the tube spins.    distributed throughout the solution. Only when subjected to enormous acceler¬  ations do macromolecules begin to sediment , much like grains of sand in water.   The ultracentrifuge, which was developed around 1923 by the Swedish  biochemist The Svedberg, can attain rotational speeds as high as 80,000 rpm  so as to generate centrifugal fields in excess of 600,000g. Using this instru¬  ment, Svedberg first demonstrated that proteins are macromolecules with ho¬  mogeneous compositions and that many proteins contain subunits.   The rate at which a particle sediments in the ultracentrifuge is related to its  mass (the density of the solution and the shape of the particle also affect the  sedimentation rate). A proteins sedimentation coefficient (its sedimentation  velocity per unit of centrifugal force) is usually expressed in units of 10 13 s,  which are known as Svedbergs (S). The relationship between molecular mass  and sedimentation coefficient is not linear; therefore, values of sedimentation  coefficients are not additive. The sedimentation coefficients of proteins range  from about IS to about 50S; viruses have sedimentation coefficients in the  range of 40S to 1000S. Subcellular particles such as mitochondria have sedi¬  mentation coefficients of tens of thousands.   Today, ultracentrifugation is rarely used as analytical tool, but it is useful  for fractioning macromolecules. Typically, sedimentation is carried out in a so¬  lution of an inert substance in which the concentration, and therefore the den¬  sity, of the solution increases from the top to the bottom of the centrifuge  tube. Such density gradients can be generated by filling the centrifuge tube  with layers of sucrose solutions of decreasing concentrations, with the sample  of macromolecules on the very top. During centrifugation, each species of  macromolecule moves through the preformed gradient at a rate largely deter¬  mined by its sedimentation coefficient and therefore travels as a zone that can  be separated from other such zones (Fig. 5-12). After centrifugation, the tube  is punctured to collect fractions containing the separated macromolecules.  Alternatively, the sample may be dissolved in a relatively concentrated solu¬  tion of a dense, fast-diffusing substance such as CsCI, which forms a density  gradient under the high gravitational field produced at high spin rates. The  sample components form bands at positions where their densities are equal to  that of the solution.   • What are some environmental conditions  that must be controlled while purifying a  protein?   • Describe how a protein may be quantified  by an assay or by absorbance  spectroscopy.   • Explain how an antibody could be useful  for purifying a protein and for determining  its concentration.   • Describe the basis for separating  proteins by ion exchange, hydrophobic  interaction, gel filtration, and affinity  chromatography.   • Describe the processes of gel  electrophoresis, SDS-PAGE, and 2D  gel electrophoresis.    • To be sequenced, a protein must be separated into individual polypeptides that  can be cleaved into sets of overlapping fragments.   • The amino acid sequence can be determined by Edman degradation, a procedure  for removing N-terminal residues one at a time.   • Mass spectrometry can identify amino acid sequences from the mass-to-charge  ratio of gas-phase protein fragments.    Once a pure sample of protein has been obtained, it may be used for a vari¬  ety of purposes. But if the protein has not previously been characterized, the  next step is often determining its sequence of amino acids. Frederick Sanger  determined the first known protein sequence, that of bovine insulin, in 1953,  thereby definitively establishing that proteins have unique covalent structures  (Box 5-1). Since then, many additional proteins have been sequenced, and the  sequences of many more proteins have been inferred from their DNA  sequences. All told, the amino acid sequences of around seven million  polypeptides are now known. Such information is valuable for the following  reasons:    1. Knowledge of a proteins amino acid sequence is prerequisite for deter¬  mining its three-dimensional structure and is essential for understand¬  ing its molecular mechanism of action.   2 . Sequence comparisons among analogous proteins from different species  yield insights into protein function and reveal evolutionary relation¬  ships among the proteins and the organisms that produce them.   3. Many inherited diseases are caused by mutations that result in an  amino acid change in a protein. Amino acid sequence analysis can as¬  sist in the development of diagnostic tests and effective therapies.   Sanger’s determination of the sequence of insulins 51 residues (Fig. 5-1)  took about 10 years and required ~100 g of protein. Procedures for primary  structure determination have since been so refined and automated that most  proteins can be sequenced within a few hours or days using only a few mi¬  crograms of material. Regardless of the technique used, the basic approach for  sequencing proteins is similar to the procedure developed by Sanger. The  protein must be broken down into fragments small enough to be individually se¬  quencerl and the primary structure of the intact protein is then reconstructed from  the sequences of overlapping fragments (Fig. 5-13). Such a procedure, as we have  seen (Section 3-4C), is also used to sequence DNA.   The complete amino acid sequence of a protein includes the sequence of each  of its subunits, if any, so the subunits must be identified and isolated before  sequencing begins.   Each polypeptide chain (if it is not chemically blocked) has an N-terminal  residue. Identifying this “end group” can establish the number of chemically dis¬  tinct polypeptides in a protein. For example, insulin has equal amounts of the  N-terminal residues Gly and Phe, which indicates that it has equal numbers  of two nonidentical polypeptide chains.   The N-terminus of a polypeptide can be determined by several methods.  The fluorescent compound 5-dimethylamino-l-naphthalenesulfonyl chlo¬  ride (dansyl chloride) reacts with primary amines to yield dansylated   Use chemical or enzymatic  methods to break each poly- ( 2a  peptide into smaller peptides, j   I Use different methods to  2b generate a different set  of peptide fragments.   Use the two sets of overlapping  4) peptide sequences to reconstruct  the sequence of each polypeptide.    Repeat fragmentation without  breaking disulfide bonds to identify  the Cys-containing sequence  involved in the disulfide linkages.   Frederick Sanger {1918-) At one time,  many biochemists believed that proteins  were amorphous “colloids” of variable size,  shape, and composition. This view was  largely abandoned by the 1940s, when it  became possible to determine the amino  acid composition of a protein, that is, the  number of each kind of constituent amino  acid. However, such information revealed  nothing about the order in which the amino  acids were combined. In fact, skeptics still  questioned whether proteins even had  unique sequences. One plausible theory was that proteins were  populations of related molecules that were probably assembled  from shorter pieces. Some studies went so far as to describe the  rules governing the relative stoichiometries and spacing of the  various amino acids in proteins.   During the 1940s, the accuracy of amino acid analysis began  to improve as a result of the development of chromatographic  techniques for separating amino acids and the use of the reagent  ninhydrin, which forms colored adducts with amino acids, for  chemically quantifying them (previous methods used cumbersome  biological assays). Frederick Sanger, who began his work on  protein sequencing in 1943, used these new techniques as well  as classic organic chemistry. Sanger did not actually set out to  sequence a protein; his first experiments were directed toward  devising a better method for quantifying free amino groups such  as the amino groups corresponding to the N-terminus of  a polypeptide.   Sanger used the reagent 2,4-dinitrofluorobenzene,  which forms a yellow dinitrophenyl (DNP) derivative  at terminal amino groups without breaking any  peptide bonds.    When the protein is subsequently hydrolyzed to break its peptide  bonds, the N-terminal amino acid retains its DNP label and can be  identified when the hydrolysis products are separated by  chromatography. Sanger found that some DNP-amino acid  derivatives were unstable during hydrolysis, so this step had to be  shortened. In comparing the results of long and short hydrolysis  times, Sanger observed extra yellow spots corresponding to  dipeptides or other small peptides with intact peptide bonds. He  could then isolate the dipeptides and identify the second residue.  Sanger’s genius was to recognize that by determining the sequences  of overlapping small peptides from an incompletely hydrolyzed  protein, the sequence of the intact protein could be determined.   An enormous amount of work was required to turn the basic  principle into a sound laboratory technique for sequencing a protein.  Sanger chose insulin as his subject, because it was one of the  smallest known proteins. Insulin contains 51 amino acids in two  polypeptide chains (called A and B). The sequence of the  30 residues in the B chain was completed in 1951, and the sequence  of the A chain (21 residues) in 1953. Because the two chains are  linked through disulfide bonds, Sanger also endeavored to find the  optimal procedure for cleaving those bonds and then identifying their  positions in the intact protein, a task he completed in 1955.   Sanger’s work of over a decade, combining his expertise in  organic chemistry (the cleavage and derivatization reactions) with  the development of analytical tools for separating and identifying  the reaction products, led to his winning a Nobel prize in 1958  [he won a second Nobel prize in 1980, for his invention of the  chain terminator method of nucleic acid sequencing (Section  3-4C)]. A testament to Sanger’s brilliance is that his basic  approaches to sequencing polypeptides and nucleic acids are still  widely used.   Publication of the sequence of insulin in 1955  convinced skeptics that a protein has a unique amino  acid sequence. Sanger’s work also helped catalyze  thinking about the existence of a genetic code that would  link the amino acid sequence of a protein to the  nucleotide sequence of DNA, a molecule whose structure  had just been elucidated in 1953.   57, 1-28 (1988). [A scientific autobiography that provides a glimpse  of the early difficulties in sequencing proteins.]   Sanger, F., Thompson, E.O.P., and Kitai, R., The amide groups of insulin,  Biochem. J. 59, 509-518 (1955).    polypeptides (Fig. 5-14). The treatment of a dansylated polypeptide with  aqueous acid at high temperature hydrolyzes its peptide bonds. This liberates  the dansylated N-terminal residue, which can then be separated chromato-  graphically from the other amino acids and identified by its intense yellow  fluorescence. The N-terminal residue can also be identified by performing the  first step of Edman degradation (Section 5-3C), a procedure that liberates  amino acids one at a time from the N-terminus of a polypeptide.   Disulfide Bonds between and within Polypeptides Are Cleaved. Disulfide  bonds between Cys residues must be cleaved to separate polypeptide chains—  if they are disulfide-linked—and to ensure that polypeptide chains are fully   FIG. 5-14 The dansyl chloride reaction. The reaction of dansyl chloride with primary  amino groups is used for end group analysis.    linear (residues in polypeptides that are “knotted” with disulfide bonds may  not be accessible to all the enzymes and reagents for sequencing). Disulfide  bonds can be reductively cleaved by treating them with 2 -mercaptoethanol or  another mercaptan (compounds that contain an —SH group):   The resulting free sulfhydryl groups are then alkylated, usually by treatment  with iodoacetate, to prevent the re-formation of disulfide bonds through ox¬  idation by 0 2 :   Polypeptides that are longer than 40 to 100 residues cannot be directly se¬  quenced and must therefore be cleaved, either enzymatically or chemically, to  specific fragments that are small enough to be sequenced. Various endopep-  tidases (enzymes that catalyze the hydrolysis of internal peptide bonds, as op¬  posed to exopeptidases, which catalyze the hydrolysis of N- or C-terminal  residues) can be used to fragment polypeptides. Both endopeptidases and ex¬  opeptidases (which collectively are called proteases) have side chain require¬  ments for the residues flanking the scissile peptide bond (i.e., the bond that  is to be cleaved; Table 5-4). The digestive enzyme trypsin has the greatest  specificity and is therefore the most valuable member of the arsenal of  endopeptidases used to fragment polypeptides. It cleaves peptide bonds on the  C side (toward the carboxyl terminus) of the positively charged residues Arg  and Lys if the next residue is not Pro.    The other endopeptidases listed in Table 5-4 exhibit broader side chain  specificities than trypsin and often yield a series of peptide fragments with  overlapping sequences. However, through limited proteolysis, that is, by ad¬  justing reaction conditions and limiting reaction times, these less specific en¬  dopeptidases can yield a set of discrete, nonoverlapping fragments.   Several chemical reagents promote peptide bond cleavage at specific  residues. The most useful of these, cyanogen bromide (CNBr), cleaves on the  C side of Met residues (Fig. 5-15).   Once the peptide fragments formed through specific cleavage reactions have  been isolated, their amino acid sequences can be determined. This can be ac¬  complished through repeated cycles of Edman degradation. In this process  (named after its inventor, Pehr Edman), phenylisothiocyanate (PITC; also  known as Edman’s reagent) reacts with the N-terminal amino group of a  polypeptide under mildly alkaline conditions to form a phenylthiocarbamyl  (PTC) adduct (Fig. 5-16). This product is treated with anhydrous    FIG. 5-15 Cyanogen bromide cleavage of a polypeptide. CNBr reacts specifically  with Met residues, resulting in cleavage of the peptide bond on their C-terminal side. The  newly formed C-terminal residue forms a cyclic structure known as a peptidyl homoser¬  ine lactone.   R n -i = bulky hydrophobic residues: Phe,  Trp, Tyr; R n A Pro   Occasionally cleaves at R n = Ala,  Asp, His, Thr; heat stable   R n = Leu, Phe, Trp, Tyr; R n -i A Pro    FIG. 5-16 Edman degradation. The reaction  occurs in three stages, each requiring different  conditions. Amino acid residues can therefore be  sequentially removed from the N-terminus of a  polypeptide in a controlled, stepwise fashion.    Many proteins include an acetyl group at  the N-terminus. How would this affect the  Edman degradation process?   Chapter 5 Proteins: Primary Structure trifluoroacetic acid, which cleaves the N-terminal residue as a thiazolinone   derivative but does not hydrolyze other peptide bonds. Edman degradation  therefore releases the N-terminal amino acid residue but leaves intact the rest  of the polypeptide chain. The thiazolinone-amino acid is selectively extracted  into an organic solvent and is converted to the more stable phenylthiohydan-  toin (PTH) derivative by treatment with aqueous acid. This PTH-amino acid  can later be identified by chromatography. Thus, it is possible to determine the  amino acid sequence of a polypeptide chain from the N-terminus inward by sub¬  jecting the polypeptide to repeated cycles of Edman degradation and, after every  cycle, identifying the newly liberated PTH-amino acid.   The Edman degradation technique has been automated and refined, re¬  sulting in great savings of time and material. In modern instruments, the pep¬  tide sample is dried onto a disk of glass fiber paper, and accurately measured  quantities of reagents are delivered and products removed as vapors in a stream  of argon at programmed intervals. Up to 100 residues can be identified be¬  fore the cumulative effects of incomplete reactions, side reactions, and peptide  loss make further amino acid identification unreliable. Since less than a pico-  mole of a PTH-amino acid can be detected and identified, sequence analysis  can be carried out on as little as 5 to 10 pmol of a peptide (<0.1 (mg—an  invisibly small amount).    FIG. 5-1 7 Electrospray ionization mass  spectrometry (ESI), (a) Dry N 2 gas promotes the  evaporation of solvent from charged droplets  containing the protein of interest, leaving gas-phase  ions, whose charge is due to the protonation of Arg  and Lys residues. The mass spectrometer then  determines the mass-to-charge ratio of these ions.  The resulting mass spectrum consists of a series of  peaks corresponding to ions that differ by a single  ionic charge and the mass of one proton). [After  Fitzgerald, M.C. and Siuzdak, G., Chem. Biol. 3, 708  (1996).] (b) The ESI mass spectrum of horse heart  apomyoglobin (myoglobin that lacks its Fe ion).   The measured m/z ratios and the inferred charges  for most of the peaks are indicated. The data  provided by this spectrum permit the mass of the  original molecule to be calculated (see Sample  Calculation 5-1). [After Yates, J.R., Methods EnzymoL  271, 353 (1996).]   Mass spectrometry has emerged as an important technique for characterizing  and sequencing polypeptides. Mass spectrometry accurately measures the mass-  to-charge (m/z) ratio for ions in the gas phase (where m is the ions mass and  z is its charge). Until about 1985, macromolecules such as proteins and nu¬  cleic acids could not be analyzed by mass spectrometry. This was because  macromolecules were destroyed during the production of gas-phase ions,  which required vaporization by heating followed by ionization via bombard¬  ment with electrons.   Newer techniques have addressed this shortcoming. For example, in the  electrospray ionization (ESI) technique, a solution of a macromolecule such  as a peptide is sprayed from a narrow capillary tube maintained at high volt¬  age (~4000 V), forming fine, highly charged droplets from which the solvent  rapidly evaporates (Fig. 5-17a). This yields a series of gas-phase macromolec-  ular ions that typically have ionic charges in the range + 0.5 to +2 per kilo-  dalton. The charges result from the protonation of basic side chains such as  Arg and Lys. The ions are directed into the mass spectrometer, which meas¬  ures their mlz values with an accuracy of ~0.01% (Fig. 5-17^). Consequently,  determining an ions z permits its molecular mass to be determined with far  greater accuracy than by any other method.   Short polypeptides (<25 residues) can be directly sequenced though the  use of a tandem mass spectrometer (Fig. 5-18; two mass spectrometers cou¬  pled in series). The first mass spectrometer functions to select and separate  the peptide ion of interest from peptide ions of different masses as well as  any contaminants that may be present. The selected peptide ion is then passed  into a collision cell, where it collides with chemically inert atoms such as he¬  lium. The energy thereby imparted to the peptide ion causes it to fragment  predominantly at only one of its several peptide bonds, thereby yielding one  or two charged fragments per original ion. The molecular masses of the nu¬  merous charged fragments so produced are then determined by the second  mass spectrometer.   By comparing the molecular masses of successively larger members of a family  of fragments, the molecular masses and therefore the identities of the corresponding  amino acid residues can be determined. The sequence of an entire polypeptide  can thus be elucidated (although mass spectrometry cannot distinguish the iso¬  meric residues lie and Leu because they have exactly the same mass, and it   FIG. 5-1 8 Tandem mass spectrometry in peptide sequencing. Electrospray  ionization (ESI), the ion source, generates gas-phase peptide ions, labeled P 2 , etc., from a  digest of the protein to be sequenced. These peptides are separated by the first mass spec¬  trometer (MS-1) according to their m/z values, and one of them (here, P 3 ) is directed into the  collision cell, where it collides with helium atoms. This treatment breaks the peptide into frag¬  ments (F-|, F 2 , etc.), which are directed into the second mass spectrometer (MS-2) for deter¬  mination of their m/z values. [After Biemann, K. and Scoble, H.A., Science 237, 992 (1987).]    An ESI mass spectrum such as that of  apomyoglobin (Fig. 5-17^) contains a  series of peaks, each corresponding to the  m/z ratio of an (M + »H)” + ion. Two suc¬  cessive peaks in this mass spectrum have  measured m/z ratios of 1414.0 and  1542.3. What is the molecular mass of the  original apomyoglobin molecule, how does  it compare with the value given for it in  Table 5-1, and what are the charges of the  ions causing these peaks?   The first peak (p x = 1414.0) arises from  an ion with charge z and mass M + z,  where M is the molecular mass of the  original protein. Then the adjacent peak  (p 2 = 1542.3), which is due to an ion  with one less proton, has charge z — 1  and mass M + z — 1. The mlz ratios for  these ions, p x and p 2 , are therefore given  by the following expressions.   These two linear equations can readily be  solved for their unknowns, M and z. Solve  the first equation for M.   M = (p2 ~ l)(p\ ~ l)/(p2 ~ P\)  Plugging in the values for p x and p 2 ,   which is only 0.14% larger than the  16,951 D for horse apomyoglobin given in  Table 5-1. For the charge on ion 1,    FIG. 5-19 Generating overlapping fragments to determine the amino acid  sequence of a polypeptide. In this example, two sets of overlapping peptide fragments  are made by using trypsin to cleave the polypeptide after all its Arg and Lys residues and,  in a separate reaction, using CNBr to cleave it after all its Met residues. O See the    cannot always reliably distinguish Gin and Lys residues because their molec¬  ular masses differ by only 0.036 D).   Computerization of the mass-comparison process has reduced the time re¬  quired to sequence a short polypeptide to only a few minutes (one cycle of  Edman degradation may take an hour). The reliability of this process has been  increased through the computerized matching of a measured mass spectrum with  those of peptides of known sequence as maintained in databases. Mass spectrom¬  etry can also be used to sequence peptides with chemically blocked N-termini  (which prevents Edman degradation) and to characterize other posttranslational  modifications such as the addition of phosphate or carbohydrate groups.   After individual peptide fragments have been sequenced, their order in the  original polypeptide must be elucidated. This is accomplished by conducting  a second round of protein cleavage with a reagent of different specificity and  then comparing the amino acid sequences of the overlapping sets of peptide  fragments (Fig. 5-19).   The final step in an amino acid sequence analysis is to determine the po¬  sitions (if any) of the disulfide bonds. This can be done by cleaving a sample  of the protein, with its disulfide bonds intact, to yield pairs of peptide frag¬  ments, each containing a single Cys, that are linked by a disulfide bond. After  isolating a disulfide-linked polypeptide fragment, the disulfide bond is cleaved  and alkylated (Section 5-3A), and the sequences of the two peptides are de¬  termined (Fig. 5-20). The various pairs of such polypeptide fragments are  identified by comparing their sequences with that of the protein, thereby es¬  tablishing the locations of the disulfide bonds.   Sequences Are Recorded in Databases. After a proteins amino acid se¬  quence has been determined, the information is customarily deposited in a  public database. Databases for proteins as well as DNA sequences are accessi¬  ble via the Internet (Table 5-5). Electronic links between databases allow rapid  updates and cross-checking of sequence information.   Most sequence databases use similar conventions. For example, the protein  may be given an ID code that includes information about its source (e.g., HU¬  MAN or ECOLI). This is accompanied by an accession number, which is as¬  signed by the database as a way of identifying an entry even if its ID code must  be changed (for example, see Fig. 5-21). The entry includes a description of the  protein, its function (if known), its sequence, and features such as disulfide bonds,  posttranslational modifications, and binding sites. The entry ends with a list of  pertinent references (which are linked to PubMed) and links to other databases.    TABLE 5-5 Internet Addresses for the Major Protein and DNA  Sequence Data Banks   Data Banks Containing Gene Sequences  GenBank: http: //www. ncbi. nlm. nih. gov/genbank/    FIG. 5-21 The initial portion of a UniProt  entry. This information pertains to the protein  adiponectin, which is produced by adipose (fat)  tissue and helps regulate fuel metabolism  (Section 22-3B). The complete entry includes  additional information and references as well as  the protein’s sequence. [From  http://www.uniprot.org/.]    Armed with the appropriate software (which is often publicly available at  the database sites), a researcher can search a database to find proteins with  similar sequences in various organisms. The sequence of even a short peptide  fragment may be sufficient to “fish out” the parent protein or its counterpart  from another species (see the Bioinformatics Projects).   Amino acid sequence information is no less valuable when the nucleotide  sequence of the corresponding gene is also known, because the protein se¬  quence provides information about protein structure that is not revealed by  nucleic acid sequencing (see Section 3-4). For example, only direct protein se¬  quencing can reveal the locations of disulfide bonds in proteins. In addition,  many proteins are modified after they are synthesized. For example, certain  residues may be excised to produce the “mature” protein (insulin, shown in  Fig. 5-1, is actually synthesized as an 84-residue polypeptide that is proteolyt-  ically processed to its smaller, two-chain form). Amino acid side chains may  also be modified by the addition of carbohydrates, phosphate groups, or acetyl  groups, to name only a few. Although some of these modifications occur at  characteristic amino acid sequences and are therefore identifiable in nucleotide  sequences, only the actual protein sequence can confirm whether and where  they occur.   • Why is it important to identify the  N-terminal residue(s) of a protein?   • What are some advantages of sequencing  peptides by mass spectrometry rather  than by Edman degradation?   • Explain why long polypeptides must be  broken into at least two different sets of  peptide fragments for sequencing.   • What types of information can be retrieved  from a protein sequence database?   TABLE 5-6 Amino Acid Sequences of Cytochromes c from 38 Species 9    ^he amino acid side chains have been shaded according to their polarity characteristics so that an invariant or conservatively substituted residue is  identified by a vertical band of a single color. The letter a at the beginning of the chain indicates that the N-terminal amino group is acetylated; an h  indicates that the acetyl group is absent.   Source: After Dickerson, R.E., Sci. Am. 226(4); 58-72 (1972), with corrections from Dickerson, R.E., and Timkovich, R., in Boyer, P.D. (Ed.), The  Enzymes (3rd ed.), Vol. 11, pp. 421-422, Academic Press (1975). Illustration, Irving Geis. Image from the Irving Geis Collection/Howard Hughes Medical  Institute. Rights owned by HHMI. Reproduction by permission only.    Sequence comparisons reveal the evolutionary relationships between proteins.  Protein families evolve by the duplication and divergence of genes encoding  protein domains.    Because an organisms genetic material specifies the amino acid sequences of  all its proteins, changes in genes due to random mutation can alter a proteins  primary structure. A mutation in a protein is propagated only if it somehow  increases, or at least does not decrease, the probability that its owner will sur¬  vive to reproduce. Many mutations are deleterious or produce lethal effects  and therefore rapidly die out. On rare occasions, however, a mutation arises  that improves the fitness of its host under the prevailing conditions. This is  the essence of evolution by natural selection.    45 50 55 60 65 70 75 80 85 90 95 100 104    |~D~| Asp [T] Glu  | H | His | K | Lys  |~B~| Asn or Asp   [g] Gly [n] Asn [q] Gin  |~W~| Trp [V] Tyr [z] Gin or Glu   The primary structures of a given protein from related species closely resemble  one another. Consider cytochrome c, a protein found in nearly all eukaryotes.  Cytochrome c is a component of the mitochondrial electron-transport system  (Section 18-2), which is believed to have taken its present form between 1.5  and 2 billion years ago, when organisms developed mechanisms for aerobic res¬  piration. Emanuel Margoliash, Emil Smith, and others have elucidated the  amino acid sequences of the cytochromes c from over 100 eukaryotic species  ranging in complexity from yeast to humans. The cytochromes c from differ¬  ent species are single polypeptides of 104 to 112 residues. The sequences of 38  of these proteins are arranged in Table 5-6 to show the similarities between   Chapter 5 Proteins: Primary Structure vertically aligned residues (the residues have been color-coded according to their   physical properties). A survey of the aligned sequences (bottom line of Table  5-6) shows that at 38 positions (23 positions in the complete set of >100 se¬  quences), the same amino acid appears in all species. Most of the remaining  positions are occupied by chemically similar residues in different organisms. In  only eight positions does the sequence accommodate six or more different  residues.   According to evolutionary theory, related species have evolved from a com¬  mon ancestor, so it follows that the genes specifying each of their proteins must  likewise have evolved from the corresponding gene in that ancestor. The sequence  of the ancestral cytochrome c is accessible only indirectly, by examining the  sequences of extant proteins.   Sequence Comparisons Provide Information on Protein Structure and  Function. In general , comparisons of the primary structures of homologous  proteins (evolutionarily related proteins) indicate which of the proteins residues  are essential to its function , which are less significant , and which have little spe¬  cific function. Finding the same residue at a particular position in the amino  acid sequence of a series of related proteins suggests that the chemical or  structural properties of that so-called invariant residue uniquely suit it to  some essential function of the protein. For example, its side chain may be  necessary for binding another molecule or for participating in a catalytic re¬  action. Other amino acid positions may have less stringent side chain re¬  quirements and can therefore accommodate residues with similar  characteristics (e.g., Asp or Glu, Ser or Thr, etc.); such positions are said to  be conservatively substituted. On the other hand, a particular amino acid  position may tolerate many different amino acid residues, indicating that the  functional requirements of that position are rather nonspecific. Such a po¬  sition is said to be hypervariable.   Why is cytochrome c —an ancient and essential protein—not identical in  all species? Even a protein that is well adapted to its function, that is, one that  is not subject to physiological improvement, nevertheless continues evolving.  The random nature of mutational processes will, in time, change such a protein  in ways that do not significantly affect its function, a process called neutral drift  (deleterious mutations are, of course, rapidly rejected through natural selection).  Hypervariable residues are apparently particularly subject to neutral drift.   Phylogenetic Trees Depict Evolutionary History. Far-reaching conclusions  about evolutionary relationships can be drawn by comparing the amino acid  sequences of homologous proteins. The simplest way to assess evolutionary  differences is to count the amino acid differences between proteins. For exam¬  ple, the data in Table 5-6 show that primate cytochromes c more nearly re¬  semble those of other mammals than they do those of insects (8-12 differences  among mammals versus 26-31 differences between mammals and insects).  Similarly, the cytochromes c of fungi differ as much from those of mammals  (45-51 differences) as they do from those of insects (41-47) or higher plants  (47-54). The order of these differences largely parallels that expected from  classical taxonomy, which is based primarily on morphological rather than mo¬  lecular characteristics.   The sequences of homologous proteins can be analyzed by computer to  construct a phylogenetic tree, a diagram that indicates the ancestral relation¬  ships among organisms that produce the protein. The phylogenetic tree for  cytochrome c is sketched in Fig. 5-22. Similar trees have been derived for  other proteins. Each branch point of the tree represents a putative common  ancestor for all the organisms above it. The distances between branch points  are expressed as the number of amino acid differences per 100 residues of the  protein. Such trees present a more quantitative measure of the degree of re¬  latedness of the various species than macroscopic taxonomy can provide.   FIG. 5-22 Phylogenetic tree of cytochrome c. Each branch  point represents an organism ancestral to the species connected  above it. The number beside each branch indicates the number of  inferred differences per 100 residues between the cytochromes c of    the flanking branch points or species. [After Dayhoff, M.O., Park, C.M.,  and McLaughlin, P.J., in Dayhoff, M.O. (Ed.), Atlas of Protein Sequence  and Structure, p. 8, National Biomedical Research Foundation (1972).]    Note that the evolutionary distances from all modern cytochromes c to  the lowest point, the earliest common ancestor producing this protein, are ap¬  proximately the same. Thus, “lower” organisms do not represent life-forms that  appeared early in history and ceased to evolve further. The cytochromes c of  all the species included in Fig. 5-22—whether called “primitive” or  “advanced”—have evolved to about the same extent.   B Proteins Evolve by the Duplication of Genes or Gene Segments   The effort to characterize protein evolution and create databases of related  proteins was pioneered by Margaret Dayhoff beginning in the 1960s. Since  then, around seven million protein sequences have been catalogued as the result  of direct protein sequencing or DNA sequencing projects. This has led to the  development of mathematically sophisticated computer algorithms to identify  similarities between sequences. These search protocols can detect similarities be¬  tween proteins that have evolved to the extent that their amino acid sequences  are <20% identical and have acquired several sequence insertions and/or    deletions of various lengths. The use of such sequence alignment programs,  which are publicly available, is demonstrated in Bioinformatics Project 2.   Analysis of a large number of proteins indicates that evolutionarily con¬  served sequences are often segments of about 40 to 200 residues, called  domains. Originally, this term referred to a portion of discrete protein structure,  but the usage has expanded to include the corresponding amino acid sequence.  As we shall see in Section 6-2D, structural similarities between two domains may  be apparent even when the sequences have few residues in common.   Protein domains can be grouped into an estimated 1000-1400 different  families, but about half of all known domains fall into just 200 families. Most  protein families have only a few members, and 10-20% of proteins appear to  be unique, although it is also possible that they represent family members whose  sequences have simply diverged beyond recognition. Domains whose sequences  are more than about 40% identical usually have the same function; domains  with less than about 25% sequence identity usually perform different roles.   Each branch point represents a gene duplication  event Myoglobin is a single-chain protein. The  globins identified by Greek letters are subunits  of hemoglobins.    Protein Families Can Arise through Gene Duplication. It is not surprising  that proteins with similar functions have similar sequences; such proteins pre¬  sumably evolved from a common ancestor. Homologous proteins with the same  function in different species (e.g., the cytochromes c shown in Table 5-6) are  said to be orthologous.   Within a species, similar proteins arise through gene duplication, an aber¬  rant genetic recombination event in which one member of a chromosome pair  acquires both copies of the primordial gene (genetic recombination is discussed  in Section 25-6). Following duplication, the sequences may diverge as muta¬  tions occur over time. Gene duplication is a particularly efficient mode of evo¬  lution because one copy of the gene evolves a new function through natural selection  while its counterpart continues to direct the synthesis of the original protein. Two  independently evolving genes that are derived from a duplication event are  said to be paralogous. In prokaryotes, approximately 60% of protein domains  have been duplicated; in many eukaryotes the figure is ~90%, and in  humans, ~98%.   r The globin family of proteins provides an excellent example of evolution  through gene duplication and divergence. Hemoglobin, which transports  0 2 from the lungs (or gills or skin) to the tissues, is a tetramer with the sub¬  unit composition a 2 (3 2 (i.e., two a polypeptides and two (3 polypeptides). The  sequences of the a and (3 subunits are similar to each other and to the se¬  quence of the protein myoglobin, which facilitates oxygen diffusion through  muscle tissue (hemoglobin and myoglobin are discussed in more detail in  Chapter 7). The primordial globin probably functioned simply as an oxygen-  storage protein. Gene duplication allowed one globin to evolve into a  monomeric hemoglobin a chain. Duplication of the a chain gene gave rise  to the paralogous gene for the (3 chain. Other members of the globin family  include the (3-like y chain that is present in fetal hemoglobin, an a 2 y 2  tetramer, and the (3-like s and a-like £ chains that appear together early in  embryogenesis as £ 2 8 2 hemoglobin. Primates contain a relatively recently du¬  plicated globin, the (3-like 8 chain, which appears as a minor component  (~1%) of adult hemoglobin. Although the a 2 8 2 hemoglobin has no known  unique function, perhaps it may eventually evolve one. The genealogy of the  members of the globin family is diagrammed in Fig. 5-23. The human  genome also contains the relics of globin genes that are not expressed. These  pseudogenes can be considered the dead ends of protein evolution. Note that  a duplicated and therefore initially superfluous gene has only a limited time  to evolve a new functionality that provides a selective advantage to its host be¬  fore it is inactivated through mutation, that is, becomes a pseudogene.   The Rate of Sequence Divergence Varies. The sequence differences between  orthologous proteins can be plotted against the time when, according to the   FIG. 5-24 Rates of evolution  of four proteins. The graph was  constructed by plotting the number  of different amino acid residues in  the proteins on two sides of a  branch point of a phylogenetic tree  versus the time, according to the  fossil record, since the correspon¬  ding species diverged from their  common ancestor. [Illustration, Irving  Geis/Geis Archives Trust Copyright  Howard Hughes Medical Institute.  Reproduced with permission.]    fossil record, the species producing the proteins diverged. The plot for a given  protein is essentially linear, indicating that its mutations accumulate at a con¬  stant rate over a geological time scale. However, rates of evolution vary among  proteins (Fig. 5-24). This does not imply that the rates of mutation of the  DNAs specifying those proteins differ, but rather that the rate at which muta¬  tions are accepted into a protein varies.   A major contributor to the rate at which a protein evolves is the effect of  amino acid changes on the proteins function. For example, Fig. 5-24 shows  that histone H4, a protein that binds to DNA in eukaryotes (Section 24-5A),  is among the most highly conserved proteins (the histones H4 from peas and  cows, species that diverged 1.2 billion years ago, differ by only two conserva¬  tive changes in their 102 residues). Evidently, histone H4 is so finely tuned  to its function of packaging DNA in cells that it is extremely intolerant of any  mutations. Cytochrome c is only slightly more tolerant. It is a relatively small  protein that binds to several other proteins. Hence, any changes in its amino  acid sequence must be compatible with all its binding partners (or they would  have to simultaneously mutate to accommodate the altered cytochrome c, an  unlikely event). Hemoglobin, which functions as a free-floating molecule, is  subject to less selective pressure than histone H4 or cytochrome c, so its surface  residues are more easily substituted by other amino acids. The fibrinopeptides  are ~20-residue fragments that are cleaved from the vertebrate protein  fibrinogen to induce blood clotting (Box 11-4). Once they have been removed,  the fibrinopep tides are discarded, so that they are subject to little selective  pressure to maintain their amino acid sequences.   The rate of protein evolution also depends on the proteins structural sta¬  bility. For example, a mutation that slowed the rate at which a newly synthe¬  sized polypeptide chain folds into its functional three-dimensional shape could  affect the cells survival, even if the protein ultimately functioned normally.  Such mutations would be especially critical for proteins that are produced at  high levels, since the not-yet-folded proteins could swamp the cells protein¬  folding mechanisms. In fact, the genes for highly expressed proteins appear to  evolve more slowly than the genes for rarely expressed proteins.   Mutational changes in proteins do not account for all evolutionary  changes among organisms. The DNA sequences that control the expression of  proteins (Chapters 26, 27, and 28) are also subject to mutation. These se¬  quences control where, when, and how much of the corresponding protein is  made. Thus, although the proteins of humans and chimpanzees are >99%  identical on average (e.g., their cytochromes c are identical), their anatomical    A Fibronectin domain 1  ■ Fibronectin domain 2  Fibronectin domain 3  # y-Carboxyglutamate domain  A Epidermal growth factor domain  | Serine protease domain  ▼ Kringle domain  Unique domain    FIG. 5-25 Construction of some multidomain proteins. Each  shape represents a segment of —40 to 100 residues that appears,  with some sequence variation, several times in the same protein or in  a number of related proteins, (a) Fibronectin, an —500-kD protein of  the extracellular matrix, is composed mostly of repeated domains of    three types, (b) Some of the proteins that participate in blood clotting  are built from a small set of domains. (The epidermal growth factor  domain is so named because it was first observed as a component of  epidermal growth factor.) [After Baron, M., Norman, D.G., and  Campbell, I.D., Trends Biochem. Sci. 16, 14 (1991).]   • How can sequence comparisons reveal  which amino acid residues are essential  for a protein’s function?   • Using Table 5-6, identify some invariant,  conservative, and hypervariable positions.  What types of amino acids appear at  conservatively substituted sites?   • Explain how the number of amino acid  differences between homologous proteins  can be used to construct a phylogenetic  tree.   • Explain the origin of orthologous proteins,  paralogous proteins, and multidomain  proteins.   • Why do different proteins appear to  evolve at different rates?    and behavioral differences are so great that they are classified as belonging to  different families.   Many Proteins Contain Domains That Occur in Other Proteins. Gene du¬  plication is not the only mechanism that generates new proteins. Analysis of  protein sequences has revealed that many proteins, particularly those made by  eukaryotes, are mosaics of sequence motifs or domains of about 40-100 amino  acid residues. These domains occur in several other proteins in the same organ¬  ism and may be repeated numerous times within a given protein (Fig. 5-25 a).  For example, most of the proteins involved in blood clotting are composed of  sets of smaller domains (Fig. 5-25 £). The sequence identity between homolo¬  gous domains is imperfect since each domain evolves independently. The func¬  tions of individual domains are not always known: Some appear to have discrete  activities, such as catalyzing a certain chemical reaction or binding a particular  molecule, but others may merely be spacers or scaffolding for other domains.   We will see in Section 25-6C how gene segments encoding protein do¬  mains are copied and inserted into other positions in a genome to generate  new genes that encode proteins with novel sequences. Such domain shuffling  is a much faster process than the duplication of an entire gene followed by its  evolution of a new functionality. Nevertheless, both mechanisms have played  important roles in the evolution of proteins.   • The properties of proteins depend largely on the sizes and  sequences of their component polypeptides.   • Protein purification requires controlled conditions such as pH and  temperature, and a means to quantify the protein (an assay).   • Fractionation procedures are used to purify proteins on the basis  of solubility, charge, polarity, size, and binding specificity.   • Differences in solubility permit proteins to be concentrated and  purified by salting out.   • Chromatography, the separation of soluble substances by their rate  of movement through an insoluble matrix, is a technique for pu¬  rifying molecules by charge (ion exchange chromatography),    hydrophobicity (hydrophobic interaction chromatography), size  (gel filtration chromatography), and binding specificity (affinity  chromatography). Binding and elution often depend on the salt  concentration and pH.   • Electrophoresis separates molecules by charge and size; SDS-  PAGE separates them primarily by size. 2D electrophoresis can  resolve thousands of proteins.   • A macromolecules rate of sedimentation in an ultracentrifuge is  related to its mass.   • Analysis of a protein’s sequence begins with end group analysis, to  determine the number of different subunits, and the cleavage of  disulfide bonds.    • Polypeptides are cleaved into fragments suitable for sequencing by  Edman degradation, in which residues are removed, one at a time,  from the N-terminus. Peptides can also be sequenced by mass  spectrometry.   • A protein’s sequence is reconstructed from the sequences of over¬  lapping peptide fragments and from information about the loca¬  tions of disulfide bonds. The sequences of numerous proteins are  archived in publicly available databases.   • Proteins evolve through changes in primary structure. Protein  sequences can be compared to construct phylogenetic trees and to  identify essential amino acid residues.   • New proteins arise as a result of the duplication of genes or gene  segments specifying protein domains, followed by their divergence.   1. It has been hypothesized that early life-forms used only eight dif¬  ferent amino acids to build small peptides. How many different  10-residue peptides could be constructed from eight amino acids?   2. If insulin (Fig. 5-1) were treated with mercaptoethanol to break its  disulfide bonds, and then the mercaptoethanol were removed so that  disulfide bonds could re-form, how many different ways could the  two polypeptide chains become linked to each other (assuming that  one or two disulfide bonds can form between the two chains)?   4. Protein X has an absorptivity of 0.4 mL • mg 1 2 3 4 * 6 • cm 1 at 280  nm. What is the absorbance at 280 nm of a 2.0 mg • mL 1  solution of protein X? (Assume the light path is 1 cm.)   3. You are using ammonium sulfate to purify protein Q (p/ = 5.0)  by salting out from a solution at pH 7.0. How should you ad¬  just the pH of the mixture to maximize the amount of protein  Q that precipitates?   6. You are trying to purify a protein that is soluble in a solution of  2 M ammonium sulfate. After centrifugation to remove other  proteins that have precipitated at this high salt concentration, you    recover the supernatant to assay the target protein’s activity in a  cell culture system, (a) Explain why the cells die when incubated  with the supernatant, (b) What procedure could you perform to  correct the problem? (Hint See Section 2-ID).   7. (a) In what order would the amino acids Arg, His, and Leu be  eluted from a carboxymethyl column at pH 6? (b) In what or¬  der would Glu, Lys, and Val be eluted from a diethylaminoethyl  column at pH 8?   8. Explain how you could use a column containing carboxymethyl  groups to separate serum albumin and ribonuclease A (see Table   9. Explain why a certain protein has an apparent molecular mass of  90 kD when determined by gel filtration and 60 kD when deter¬  mined by SDS-PAGE in the presence or absence of 2-mercap-  toethanol. Which molecular mass determination is more accurate?   Molecular mass by gel filtration: 200 kD  Molecular mass by SDS-PAGE: 100 kD   11. What fractionation procedure could be used to purify protein 1  from a mixture of three proteins whose amino acid compositions  are as follows?   1. 25% Ala, 20% Gfy, 20% Ser, 10% He, 10% Val, 5% Asn,  5% Gin, 5% Pro   2. 30% Gin, 25% Glu, 20% Lys, 15% Ser, 10% Cys   3. 25% Asn, 20% Gly, 20% Asp, 20% Ser, 10% Lys, 5% Tyr   All three proteins are similar in size and p/, and there is no  antibody available for protein 1.   12. Consult Table 5-1 to complete the following: (a) On a plot of  absorbance at 280 nm versus elution volume, sketch the results  of gel filtration of a mixture containing human cytochrome c and  bacteriophage T7 RNA polymerase and identify each peak,   (b) Sketch the results of SDS-PAGE of the same protein mixture  showing the direction of migration and identifying each band.   13. Explain why a protein, which has a sedimentation coefficient of  2.6S when ultracentrifuged in a solution containing 0.1 M NaCl,  has a sedimentation coefficient of 4.3S in a solution containing  1 M NaCl.   14. A protein has an apparent mass of 800 kD by gel filtration chro¬  matography, but SDS-PAGE shows a single band at a position  corresponding to 200 kD. In an ultracentrifuge, will the protein  exhibit a sedimentation coefficient corresponding to 200 kD or  800 kD?   15. Purification of Myoglobin (Mb). Purification tables are often used  to keep track of the yield and purification of a protein. The spe¬  cific activity is a ratio of the amount of the protein of interest, in  this case Mb, obtained at a given step (fimol or enzyme units) di¬  vided by the amount (mg) of total protein. The yield is the ratio  of the amount of the protein of interest obtained at a given step  (panol or enzyme units) divided by the original amount present  in the crude extract, often converted to percent yield by multiply¬  ing by 100. The fold purification is the ratio of the specific activ¬  ity of the purified protein to that of the crude preparation.   (a) For the purification table below, calculate the specific activ¬  ity, % yield, and fold purification for the empty cells.   (d) If you wanted to use only one purification step, which tech¬  nique would you choose?   16. A 50-mL crude extract of skeletal muscle contains 32 mg of pro¬  tein per mL. Ten jjlL of the extract catalyzes a reaction at a rate of  0.14 |JLmol product formed per minute. The extract was frac¬  tionated by ammonium sulfate precipitation, and the fraction    precipitating between 20% and 40% saturation was redissolved in  10 mL. This solution contains 50 mg/mL protein. Ten jjlL of this  purified fraction catalyzes the reaction at a rate of 0.65 pmol/min.  (a) What is the degree of purification (fold purification)? (b) What  is the percent yield of the enzyme recovered in the purified fraction?   17. Explain why the dansyl chloride treatment of a single polypep¬  tide chain followed by its complete acid hydrolysis yields several  dansylated amino acids.   18. Identify the first residue obtained by Edman degradation of cy¬  tochrome c from (a) Drosophila , (b) baker’s yeast, and (c) wheat  germ (see Table 5-6).   19. You must cleave the following peptide into smaller fragments.  Which of the proteases listed in Table 5-4 would be likely to yield  the most fragments? The fewest?   20. You wish to determine the sequence of a polypeptide that has the  following amino acid composition.   1 Ala 4 Arg 2 Asn 3 Asp 4 Cys 3 Gly 1 Gin 4 Glu  1 His 1 Lys 1 Met 1 Phe 2 Pro 4 Ser 2 Tyr 1 Trp   (a) What is the maximum number of peptides you can expect if  you cleave the polypeptide with cyanogen bromide?   (b) What is the maximum number of peptides you can expect if  you cleave the polypeptide with chymotrypsin?   (c) Analysis of the intact polypeptide reveals that there are no  free sulfhydryl groups. How many disulfide bonds are likely  to be present?   21. Electrospray ionization mass spectrometry (ESI-MS) of proteins  involves creating positively charged ions of the protein and sep-   (a) What causes the different positive charges on different  particles of the protein?   (b) The amino acid composition (in numbers of residues per  chain) of hen egg-white lysozyme (HEWL) is as follows:    22. (a) The ESI-MS spectrum below was obtained for HEWL. Using  peaks 5 and 6, calculate the molecular mass of HEWL (see  Sample Calculation 5-1).   http: //www. astbury. leeds. ac.uk/ facil / M Stut/mstutorial. htm]  (b) What is the charge on the ion that makes peak 5?   23. You wish to determine the sequence of a short peptide.  Cleavage with trypsin yields three smaller peptides with the  sequences Leu-Glu, Gly-Tyr-Asn-Arg, and Gln-Ala-Phe-  Val-Lys. Cleavage with chymotrypsin yields three peptides  with the sequences Gln-Ala-Phe, Asn-Arg-Leu-Glu, and  Val-Lys-Gly-Tyr. What is the sequence of the intact peptide?   24. Separate cleavage reactions of a polypeptide by CNBr and chy-  motrypsin yield fragments with the following amino acid se¬  quences. What is the sequence of the intact polypeptide?   Indicate the positions of the disulfide bonds in the intact  polypeptide.   26. You wish to sequence the light chain of a protease inhibitor from  the Brassica nigra plant. Cleavage of the light chain by trypsin   and chymotrypsin yields the following fragments. What is the  sequence of the light chain?   27. In site-directed mutagenesis experiments, Gly is often successfully  substituted for Val, but Val can rarely substitute for Gly. Explain.   28. Below is a list of the first 10 residues of the B helix in myoglo¬  bin from different organisms.    Based on this information, which positions (a) appear unable to  tolerate substitutions, (b) can tolerate conservative substitution,  and (c) are highly variable?   29. Sketch a phylogenetic tree for the family of homologous proteins  whose partial sequences are given below.   30. Draw a different phylogenetic tree based on the data in Problem 29.   1. Obtaining Sequences from BLAST. Using a known protein se¬  quence, find and retrieve the sequences of related proteins from  other organisms.    3. Phylogenetic Trees. Set up and interpret phylogenetic trees to  explore the evolutionary relationships between related protein  sequences.   4. One-Dimensional Electrophoresis. Perform an SDS-PAGE  electrophoresis simulation with known and unknown proteins.   5. Two-Dimensional Electrophoresis. Explore the predicted  and observed electrophoretic parameters (p/, molecular mass,  and fragmentation pattern) for a known protein.   Focus concept: A histidine-proline-rich glycoprotein may serve as  a plasma sensor and regulate local pH in extracellular fluid during  ischemia or metabolic acidosis.    Select one of the proteins  listed in Table 5-1 or 5-3. How is this protein purified? What is the  source of the protein? What types of chromatography are typically  used? What chemical or biological feature is used to detect it or  quantify it? Is there anything unusual about its amino acid se¬  quence? Does it contain any disulfide bonds? Is it present only in  animals or does it also occur in prokaryotes?   Boyer, R.F., Biochemistry Laboratory: Modern Theory and Techniques ,  Benjamin Cummings (2006).   Burgess, R.R. and Deutscher, M.P. (Eds.), Guide to Protein Purification  (2nd ed.), Methods Enzymol. 463 (2009).   Janson, J.-C. (Ed.), Protein Purification: Principles, High Resolution Meth¬  ods, and Applications (3rd ed.), Wiley (2007). [Contains detailed dis¬  cussions of a variety of chromatographic and electrophoretic  separation techniques.]   Ninfa, A.J., Ballou, D.R, and Benore, M., Fundamental Laboratory  Approaches for Biochemistry and Biotechnology (2nd ed.), Wiley (2010).   Simpson, R.J., Adams, P.D., and Golemis, E.A. (Eds.), Basic Methods in  Protein Purification and Analysis. A Laboratory Manual, Cold Spring  Harbor Laboratory Press (2009).   Tanford, C. and Reynolds, J., Nature’s Robots: A History of Proteins,  Oxford University Press (2001). [Descriptions of some early discoveries  related to the nature of proteins and their purification and analysis.]   Aebersold, R. and Mann, M., Mass spectrometry-based proteomics,  Nature 422, 198-207 (2003). [Describes some of the methods of mass  spectrometric analysis of proteins as well as current and potential  applications.]   Findlay, J.B.C. and Geisow, M.J. (Eds.), Protein Sequencing. A Practical  Approach, IRL Press (1989).    Galperin, M.Y. and Cochrane, G.R. The 2011 Nucleic Acids Research data¬  base issue and online database collection: a community of data  resources, Nucleic Acids Res. 38, Database issue D1-D6 (2011). [This  and other articles in the Database Issue describe the features and  potential uses of various protein and DNA sequence databases. Freely  available at http://nar.oxfordjournals.org/.]   Baxevanis, A.D. and Ouellette, B.F.F. (Eds.), Bioinformatics, A Practical  Guide to the Analysis of Genes and Proteins (3rd ed.), Wiley-  Interscience (2003).   Doolittle, R.F., Feng, D.-F., Tsang, S., Cho, G., and Little, E., Determin¬  ing divergence times of the major kingdoms of living organisms with  a protein clock, Science 271, 470-477 (1996). [Demonstrates how  protein sequences can be used to draw phylogenetic trees.]   Lesk, A.M., Lntroduction to Bioinformatics (3rd ed.), Oxford University  Press (2008).   Mount, D.W., Bioinformatics: Sequence and Genome Analysis (2nd ed.),  Cold Spring Harbor Laboratory Press (2004).   Pal, C., Papp, B., and Lercher, M.J., An integrated view of protein evo¬  lution, Nature Reviews Genetics! , 337-348 (2006). [Discusses some  of the factors that contribute to the variable rate of evolution among  proteins, including protein dispensability and expression level.]   The outer shell of the rhinovirus, cause of the common cold, consists of 60 copies  each of three proteins (colored red, blue, and green), which self-assemble to form a  300-A-diameter capsule that encloses the viral genome. [Image generated by the Virus  Particle Explorer (http://viperdb.scripps.edu); based on an X-ray structure by Verdaguer N.,  Blaas D., and Fita I.; Institut de Biologia de Barcelona, Spain. PDBid 1 FPN.]    For many years, it was thought that proteins were colloids of random structure  and that the enzymatic activities of certain crystallized proteins were due to un¬  known entities associated with an inert protein carrier. In 1934, J.D. Bernal and  Dorothy Crowfoot Hodgkin showed that a crystal of the protein pepsin yielded  a discrete diffraction pattern when placed in an X-ray beam. This result pro¬  vided convincing evidence that pepsin was not a random colloid but an ordered  array of atoms organized into a large yet uniquely structured molecule.   Even relatively small proteins contain thousands of atoms, almost all of  which occupy definite positions in space. The first X-ray structure of a protein,  that of sperm whale myoglobin, was reported in 1958 by John Kendrew and  co-workers. At the time—only 5 years after James Watson and Francis Crick  had elucidated the simple and elegant structure of DNA (Section 3-2B)—  protein chemists were chagrined by the complexity and apparent lack of  regularity in the structure of myoglobin. In retrospect, such irregularity seems  essential for proteins to fulfill their diverse biological roles. However, compar¬  isons of the nearly 80,000 protein structures now known have revealed that  proteins actually exhibit a remarkable degree of structural regularity.   As we saw in Section 5-1, the primary structure of a protein is its linear  sequence of amino acids. In discussing protein structure, three further levels  of structural complexity are customarily invoked:   • Secondary structure is the local spatial arrangement of a polypeptides  backbone atoms without regard to the conformations of its side chains.   • Tertiary structure refers to the three-dimensional structure of an entire  polypeptide, including its side chains.   B The Most Common Regular Secondary  Structures Are the a Helix and the (3 Sheet  C Fibrous Proteins Have Repeating Secondary  Structures   A Most Protein Structures Are Determined by  X-Ray Crystallography or Nuclear Magnetic  Resonance   B Side Chain Location Varies with Polarity  C Tertiary Structures Contain Combinations of  Secondary Structure   D Structure Is Conserved More than Sequence  E Structural Bioinformatics Provides Tools for  Storing, Visualizing, and Comparing Protein  Structural Information   A Proteins Are Stabilized by Several Forces  B Proteins Can Undergo Denaturation  and Renaturation  C Proteins Are Dynamic   A Proteins Follow Folding Pathways  B Molecular Chaperones Assist Protein Folding  C Some Diseases Are Caused by  Protein Misfolding   Chapter 6 Proteins: Three-Dimensional Structure • Many proteins are composed of two or more polypeptide chains, loosely   referred to as subunits. A proteins quaternary structure refers to the  spatial arrangement of its subunits.   The four levels of protein structure are summarized in Fig. 6-1.   In this chapter, we explore secondary through quaternary structure,  including examples of proteins that illustrate each of these levels. We also dis¬  cuss the process of protein folding and the forces that stabilize folded proteins.    • The planar character of the peptide group limits the conformational flexibility of  the polypeptide chain.   • The a helix and the p sheet allow the polypeptide chain to adopt favorable 4> and  i|/ angles and to form hydrogen bonds.   • Fibrous proteins contain long stretches of regular secondary structure, such as  the coiled coils in a keratin and the triple helix in collagen.   • Not all polypeptide segments form regular secondary structure such as a helices  or p sheets.    Protein secondary structure includes the regular polypeptide folding patterns  such as helices, sheets, and turns. However, before we discuss these basic struc¬  tural elements, we must consider the geometric properties of peptide groups,  which underlie all higher order structures.   Recall from Section 4-IB that a polypeptide is a polymer of amino acid  residues linked by amide (peptide) bonds. In the 1930s and 1940s, Linus    (a) - Lys - Ala - His - Gly - Lys - Lys - Val - Leu - Gly - Ala -    FIG. 6-1 Levels of protein structure, (a) Primary structure, (b) secondary structure,  (c) tertiary structure, and (c/) quaternary structure. [Illustration, Irving Geis. Image from the  Irving Geis Collection/Howard Hughes Medical Institute. Rights owned by HHMI.  Reproduction by permission only.]   bond lengths (in angstroms) and angles (in  degrees) are derived from X-ray crystal  structures. [After Marsh, R.E. and Donohue, J.,  Adv. Protein Chem. 22, 249 (1967).] O See    Pauling and Robert Corey determined the X-ray structures of several amino  acids and dipeptides in an effort to elucidate the conformational constraints  on a polypeptide chain. These studies indicated that the peptide group has a  rigid , planar structure as a consequence of resonance interactions that give the  peptide bond ~ 40% double-bond character:    This explanation is supported by the observations that a peptide group’s C—N  bond is 0.13 A shorter than its N—C a single bond and that its C=0 bond  is 0.02 A longer than that of aldehydes and ketones. The planar conforma¬  tion maximizes TT-bonding overlap, which accounts for the peptide groups  rigidity.   Peptide groups, with few exceptions, assume the trans conformation, in  which successive C a atoms are on opposite sides of the peptide bond joining  them (Fig. 6-2). The cis conformation, in which successive C a atoms are on  the same side of the peptide bond, is ~8 kj • mol 1 less stable than the trans  conformation because of steric interference between neighboring side chains.  However, this steric interference is reduced in peptide bonds to Pro residues,  so ~ 10% of the Pro residues in proteins follow a cis peptide bond.    Torsion Angles between Peptide Groups Describe Polypeptide Chain  Conformations. The backbone or main chain of a protein refers to the  atoms that participate in peptide bonds, ignoring the side chains of the amino  acid residues. The backbone can be drawn as a linked sequence of rigid pla¬  nar peptide groups (Fig. 6-3). The conformation of the backbone can therefore  be described by the torsion angles (also called dihedral angles or rotation  angles) around the C a —N bond (<fi) and the C a —C bond (ifr) of each residue   FIG. 6-3 Extended conformation of a polypeptide. The backbone is shown as a  series of planar peptide groups. [Illustration, Irving Geis. Image from the Irving Geis  Collection/Howard Hughes Medical Institute. Rights owned by HHMI. Reproduction by  permission only.]    FIG. 6-4 Torsion angles of the  polypeptide backbone. Two planar peptide  groups are shown. The only reasonably free  movements are rotations around the C a —N  bond (measured as c|>) and the C a —C bond  (measured as i|/). By convention, both cf> and i|/  are 180° in the conformation shown and  increase, as indicated, when the peptide plane is  rotated in the clockwise direction as viewed from  C a . [Illustration, Irving Geis. Image from the Irving  Geis Collection/Howard Hughes Medical  Institute. Rights owned by HHMI. Reproduction  by permission only.] Q See Kinemage  Exercise 3-1.   (Fig. 6 -4). These angles, <J) and i(j, are both defined as 180° when the  polypeptide chain is in its fully extended conformation and increase clock¬  wise when viewed from C a .   The conformational freedom and therefore the torsion angles of a  polypeptide backbone are sterically constrained. Rotation around the C a —N  and C a —C bonds to form certain combinations of 4> and i(i angles will cause  the amide hydrogen, the carbonyl oxygen, or the substituents of C a of adja¬  cent residues to collide (e.g., Fig. 6-5). Certain conformations of longer  polypeptides can similarly produce collisions between residues that are far apart  in sequence.   The Ramachandran Diagram Indicates Allowed Conformations of  Polypeptides. The sterically allowed values of c and v(i can be calculated.  Sterically forbidden conformations, such as the one shown in Fig. 6-5, have  4> and ij/ values that would bring atoms closer than the corresponding van der  Waals distance (the distance of closest contact between nonbonded atoms).  Such information is summarized in a Ramachandran diagram (Fig. 6-6),  which is named after its inventor, G. N. Ramachandran.    FIG. 6-5 Steric interference between adjacent peptide groups. Rotation can result  in a conformation in which the amide hydrogen of one residue and the carbonyl oxygen of  the next are closer than their van der Waals distance. [Illustration, Irving Geis. Image from  the Irving Geis Collection/Howard Hughes Medical Institute. Rights owned by HHMI.  Reproduction by permission only.] O See Kinemage Exercise 3-1.   FIG. 6-6 The Ramachandran diagram. The blue-shaded regions indicate the sterically  allowed $ and i|i angles for all residues except Gly and Pro. The green-shaded regions  indicate the more crowded (outer limit) cf> and i|i angles. The yellow circles represent  conformational angles of several secondary structures: a, right-handed a helix; TT, parallel p  sheet; 1\f antiparallel p sheet; C, collagen helix; a L , left-handed a helix.    Most areas of the Ramachandran diagram (most combinations of 4> and  vji) represent forbidden conformations of a polypeptide chain. Only three small  regions of the diagram are physically accessible to most residues. The observed  4> and vji values of accurately determined structures nearly always fall within  these allowed regions of the Ramachandran plot. There are, however, some  notable exceptions:   1. The cyclic side chain of Pro limits its range of 4> values to angles of  around —60°, making it, not surprisingly, the most conformationally  restricted amino acid residue.   2. Gly, the only residue without a Cp atom, is much less sterically hin¬  dered than the other amino acid residues. Hence, its permissible range  of 4> and i(i covers a larger area of the Ramachandran diagram. At Gly  residues, polypeptide chains often assume conformations that are for¬  bidden to other residues.    B The Most Common Regular Secondary Structures Are the  a Helix and the (3 Sheet    A few elements of protein secondary structure are so widespread that they are  immediately recognizable in proteins with widely differing amino acid  sequences. Both the o i helix and the (5 sheet are such elements; they are called  regular secondary structures because they are composed of sequences of  residues with repeating 4> and vji values.    The a Helix Is a Coil. Only one polypeptide helix has both a favorable  hydrogen bonding pattern and 4> and vji values that fall within the fully allowed  regions of the Ramachandran diagram: the a helix. Its discovery by Linus  Pauling in 1951, through model building, ranks as one of the landmarks of  structural biochemistry (Box 6-1).    Linus Pauling (1901-1994) Linus  Pauling, the only person to have been  awarded two unshared Nobel prizes, is  clearly the dominant figure in twentieth-  century chemistry and one of the  greatest scientific figures of all time. He  received his B.Sc. in chemical engineering  from Oregon Agricultural College (now  Oregon State University) in 1922 and his  Ph.D. in chemistry from the California  Institute of Technology in 1925, where he  spent most of his career.   The major theme throughout Pauling’s long scientific life was  the study of molecular structures and the nature of the chemical  bond. He began this career by using the then recently invented  technique of X-ray crystallography to determine the structures of  simple minerals and inorganic salts. At that time, methods for  solving the phase problem (Box 7-2) were unknown, so X-ray  structures could only be determined using trial-and-error  techniques. This limited the possible molecules that could be  effectively studied to those with few atoms and high symmetry  such that their atomic coordinates could be fully described by only  a few parameters (rather than the three-dimensional coordinates  of each of its atoms). Pauling realized that the positions of atoms  in molecules were governed by fixed atomic radii, bond distances,  and bond angles and used this information to make educated  guesses about molecular structures. This greatly extended the  complexity of the molecules whose structures could be  determined.   In his next major contribution, occurring in 1931, Pauling  revolutionized the way that chemists viewed molecules by applying  the then infant field of quantum mechanics to chemistry. Pauling  formulated the theories of orbital hybridization, electron-pair  bonding, and resonance and thereby explained the nature of  covalent bonds. This work was summarized in his highly influential  monograph, The Nature of the Chemical Bond, which was first  published in 1938.   In the mid-1930s, Pauling turned his attention to biological  chemistry. He began these studies in collaboration with his  colleague, Robert Corey, by determining the X-ray structures of  several amino acids and dipeptides. At that time, the X-ray  structural determination of even such small molecules required  around a year of intense effort, largely because the numerous  calculations required to solve a structure had to be made by hand  (electronic computers had yet to be invented). Nevertheless, these  studies led Pauling and Corey to the conclusions that the peptide  bond is planar, which Pauling explained from resonance  considerations (Section 6-1 A), and that hydrogen bonding plays a  central role in maintaining macromolecular structures.   In the 1940s, Pauling made several unsuccessful attempts to  determine whether polypeptides have any preferred conformations.  Then, in 1948, while visiting Oxford University, he was confined to  bed by a cold. He eventually tired of reading detective stories and  science fiction and again turned his attention to proteins. By  folding drawings of polypeptides in various ways, he discovered  the a helix, whose existence was rapidly confirmed by X-ray    studies of a keratin (Section 6-1C). This work was reported in  1951, and later that year Pauling and Corey also proposed both  the parallel and antiparallel p pleated sheets. For these ground¬  breaking insights, Pauling received the Nobel Prize in Chemistry  in 1954, although a helices and p sheets were not actually  visualized until the first X-ray structures of proteins were  determined, five to ten years later.   Pauling made numerous additional pioneering contributions to  biological chemistry, most notably that the heme group in  hemoglobin changes its electronic state on binding oxygen  (Section 7-1 A), that vertebrate hemoglobins are a 2 p 2  heterotetramers (Section 7-1 B), that the denaturation of proteins  is caused by the unfolding of their polypeptide chains, that sickle¬  cell anemia is caused by a mutation in the p chain of normal  adult hemoglobin (the first so-called molecular disease to be  characterized; Section 7-1 E), that molecular complementarity plays  an important role in antibody-antigen interactions (Section 7-3B)  and by extension all macromolecular interactions, that enzymes  catalyze reactions by preferentially binding their transition states  (Section 11-3E), and that the comparison of the sequences of the  corresponding proteins in different organisms yields evolutionary  insights (Section 5-4).   Pauling was also a lively and stimulating lecturer who for many  years taught a general chemistry course [which one of the authors  of this textbook (DV) had the privilege of taking]. His textbook,  General Chemistry, revolutionized the way that introductory  chemistry was taught by presenting it as a subject that could be  understood in terms of atomic physics and molecular structure.   For a book of such generality, an astounding portion of its subject  matter had been elucidated by its author. Pauling’s amazing grasp  of chemistry was demonstrated by the fact that he dictated each  chapter of the textbook in a single sitting.   By the late 1940s, Pauling became convinced that the  possibility of nuclear war posed an enormous danger to humanity  and calculated that the radioactive fallout from each aboveground  test of a nuclear bomb would ultimately cause cancer in thousands  of people. He therefore began a campaign to educate the public  about the hazards of bomb testing and nuclear war. The political  climate in the United States at the time was such that the  government considered Pauling to be subversive and his passport  was revoked (and only returned two weeks before he was to leave  for Sweden to receive his first Nobel prize). Nevertheless, Pauling  persisted in this campaign, which culminated, in 1962, with the  signing of the first Nuclear Test Ban Treaty. For his efforts, Pauling  was awarded the 1962 Nobel Peace Prize.   Pauling saw science as the search for the truth, which included  politics and social causes. In his later years, he became a  vociferous promoter of what he called orthomolecular medicine,  the notion that large doses of vitamins could ward off and cure  many human diseases, including cancer. In the best known  manifestation of this concept, Pauling advocated taking large  doses of vitamin C to prevent the common cold and lessen its  symptoms, advice still followed by millions of people, although the  medical evidence supporting this notion is scant. It should be  noted, however, that Pauling, who followed his own advice,  remained active until he died in 1994 at the age of 93.   FIG. 6-7 The ol helix. This right-handed helical conformation has 3.6 residues  per turn. Dashed lines indicate hydrogen bonds between C=0 groups and N—H  groups that are four residues farther along the polypeptide chain. [Illustration, Irving  Geis. Image from the Irving Geis Collection/Howard Hughes Medical Institute.  Rights owned by HHMI. Reproduction by permission only.] Q See Kinemage  Exercise 3-2 and the Animated Figures.    How many amino acid residues are in this helix? How many intrachain    The a helix (Fig. 6-7) is right-handed; that is, it turns in the direction  that the fingers of a right hand curl when its thumb points in the direction  that the helix rises (Fig. 3-7). The a helix has 3.6 residues per turn and a  pitch (the distance the helix rises along its axis per turn) of 5.4 A. The a he¬  lices of proteins have an average length of —12 residues, which corresponds  to over three helical turns, and a length of — 18 A.   In the a helix, the backbone hydrogen bonds are arranged such that the pep¬  tide C=0 bond of the nth residue points along the helix axis toward the peptide  N—H group of the (n + 4)th residue. This results in a strong hydrogen bond  that has the nearly optimum N***0 distance of 2.8 A. Amino acid side chains  project outward and downward from the helix (Fig. 6-8), thereby avoiding  steric interference with the polypeptide backbone and with each other. The core  of the helix is tightly packed; that is, its atoms are in van der Waals contact.    FIG. 6-8 Space-filling model of an ol  helix. The backbone atoms are colored according  to type with C green, N blue, 0 red, and H  white. The side chains (gold) project away from  the helix. This a helix is a segment of sperm  whale myoglobin. [Based on an X-ray structure  by lime Schlichting, Max Planck Institut fur  Molekulare Physiologie, Dortmund, Germany.  PDBid 1A6M (for the definition of “PDBid” see  Section 6-2E).]   FIG. 6-9 p Sheets. Dashed lines indicate  hydrogen bonds between polypeptide strands.  Side chains are omitted for clarity, (a) An  antiparallel p sheet, (b) A parallel p sheet.  [Illustration, Irving Geis. Image from the Irving  Geis Collection/Howard Hughes Medical  Institute. Rights owned by HHMI. Reproduction  by permission only.] Q See Kinemage  Exercise 3-3 and the Animated Figures.   P Sheets Are Formed from Extended Chains. In 1951, the same year  Pauling proposed the a helix, Pauling and Corey postulated the existence of  a different polypeptide secondary structure, the P sheet. Like the a helix, the  P sheet uses the full hydrogen-bonding capacity of the polypeptide backbone.  In (3 sheets, however, hydrogen bonding occurs between neighboring polypeptide  chains rather than within one as in an a helix.    1. The antiparallel P sheet, in which neighboring hydrogen-bonded  polypeptide chains run in opposite directions (Fig. 6-9 d).   2. The parallel P sheet, in which the hydrogen-bonded chains extend in  the same direction (Fig. 6-9 £).   The conformations in which these p structures are optimally hydrogen bonded  vary somewhat from that of the fully extended polypeptide shown in Fig. 6-3.  They therefore have a rippled or pleated edge-on appearance (Fig. 6-10) and  for that reason are sometimes called “pleated sheets.” Successive side chains of  a polypeptide chain in a P sheet extend to opposite sides of the sheet with a  two-residue repeat distance of 7.0 A.   P Sheets in proteins contain 2 to as many as 22 polypeptide strands, with  an average of 6 strands. Each strand may contain up to 15 residues, the average  being 6 residues. A seven-stranded antiparallel P sheet is shown in Fig. 6-11.   Parallel P sheets containing fewer than five strands are rare. This observa¬  tion suggests that parallel p sheets are less stable than antiparallel p sheets,  possibly because the hydrogen bonds of parallel sheets are distorted compared  to those of the antiparallel sheets (Fig. 6-9). P Sheets containing mixtures of  parallel and antiparallel strands frequently occur.    FIG. 6-10 Pleated appearance of a p sheet. Dashed lines indicate hydrogen bonds. The R groups  ( purple ) on each polypeptide chain alternately extend to opposite sides of the sheet and are in register  on adjacent chains. [Illustration, Irving Geis. Image from the Irving Geis Collection/Howard Hughes  Medical Institute. Rights owned by HHMI. Reproduction by permission only.]    How many residues are in this p sheet? How many interchain hydrogen bonds?    (3 Sheets almost invariably exhibit a pronounced right-handed twist when  viewed along their polypeptide strands (Fig. 6-12). Conformational energy  calculations indicate that the twist is a consequence of interactions between  chiral L- amino acid residues in the extended polypeptide chains. The twist dis¬  torts and weakens the (3 sheets interchain hydrogen bonds. The geometry of  a particular (3 sheet is thus a compromise between optimizing the conforma¬  tional energies of its polypeptide chains and preserving its hydrogen bonding.   The topology (connectivity) of the polypeptide strands in a (3 sheet can  be quite complex. The connection between two antiparallel strands may be    FIG. 6-1 1 Space-filling model of a p sheet. The backbone  atoms are colored according to type with C green, N blue, 0 red, and  H white. The R groups are represented by large magenta spheres.   This seven-stranded antiparallel p sheet, which is shown with its  polypeptide strands approximately horizontal, is from the jack bean  protein concanavalin A. [Based on an X-ray structure by Gerald  Edelman, The Rockefeller University. PDBid 2CNA.] Q See Kinemage  Exercise 3-3.   polypeptide backbone is drawn in ribbon form with a helices depicted as cyan  coils, the strands of the p sheet represented by green arrows pointing toward  the C-terminus, and its remaining portions portrayed by orange worms. Side  chains are not shown. The eight-stranded p sheet forms a saddle-shaped  curved surface with a right-handed twist. [Based on an X-ray structure by William  Lipscomb, Harvard University. PDBid 3CPA.]   FIG. 6-13 Connections between adjacent  strands in p sheets, (a) Antiparallel strands  may be connected by a small loop, (b) Parallel  strands require a more extensive crossover  connection. [After Richardson, J.S., Adv. Protein  Chem. 34, 196 (1981).]    just a small loop (Fig. 6-13tf), but the link between tandem parallel strands  must be a crossover connection that is out of the plane of the (3 sheet  (Fig. 6-\3b). The connecting link in either case can be extensive, often con¬  taining helices (e.g., Fig. 6-12).   Turns Connect Some Units of Secondary Structure. Polypeptide segments  with regular secondary structure such as a helices or the strands of (3 sheets are  often joined by stretches of polypeptide that abruptly change direction. Such  reverse turns or (3 bends (so named because they often connect successive  strands of antiparallel (3 sheets; Fig. 6-13a) almost always occur at protein sur¬  faces. They usually involve four successive amino acid residues arranged in one  of two ways, Type I and Type II, that differ by a 180° flip of the peptide unit  linking residues 2 and 3 (Fig. 6-14). Both types of turns are stabilized by a  hydrogen bond, although deviations from these ideal conformations often dis¬  rupt this hydrogen bond. In Type II turns, the oxygen atom of residue 2 crowds  the Cp atom of residue 3, which is therefore usually Gly. Residue 2 of either  type of turn is often Pro since it can assume the required conformation.   Proteins have historically been classified as either fibrous or globular, depend¬  ing on their overall morphology. This dichotomy predates methods for deter¬  mining protein structure on an atomic scale and does not do justice to proteins  that contain both stiff, elongated, fibrous regions as well as more compact,  highly folded, globular regions. Nevertheless, the division helps emphasize the  properties of fibrous proteins, which often have a protective, connective, or  supportive role in living organisms. The two well-characterized fibrous pro¬  teins we discuss here—keratin and collagen—are highly elongated molecules  whose shapes are dominated by a single type of secondary structure. They are  therefore useful examples of these structural elements.   a Keratin Is a Coiled Coil. Keratin is a mechanically durable and relatively  unreactive protein that occurs in all higher vertebrates. It is the principal com¬  ponent of their horny outer epidermal layer and its related appendages such  as hair, horn, nails, and feathers. Keratins have been classified as either a ker¬  atins, which occur in mammals, or (3 keratins, which occur in birds and reptiles.    FIG. 6-14 Reverse turns in polypeptide  chains. Dashed lines represent hydrogen bonds,  (a) Type I. ( b ) Type II. [Illustration, Irving Geis.  Image from the Irving Geis Collection/Howard  Hughes Medical Institute. Rights owned by  HHMI. Reproduction by permission only.] Q See  Kinemage Exercise 3-4.    FIG. 6-1 5 A coiled coil, (a) View down the coil axis showing the alignment of  nonpolar residues along one side of each a helix. The helices have the pseudorepeating  sequence a-b-c-d-e-f-g in which residues a and d are predominately nonpolar. [After  McLachlan, A.D. and Stewart, M., J. Mol. Biol. 98, 295 (1975).] ( b ) Side view of the  polypeptide backbones in stick form (left) and of the entire polypeptides in space-filling  form (right). The atoms are colored according to type with C green in one chain and cyan  in the other, N blue, 0 red, and S yellow. The 81-residue chains are parallel with their  N-terminal ends above. Note that in the space-filling model the side chains of the two  polypeptides contact each other. This coiled coil is a portion of the muscle protein  tropomyosin (Section 7-2A). [Based on an X-ray structure by Carolyn Cohen, Brandeis  University. PDBid 11C2J Q See Kinemage Exercises 4-1 and 4-2.   Humans have over 50 keratin genes that are expressed in a tissue-specific    The X-ray diffraction pattern of a keratin resembles that expected for an  a helix (hence the name a keratin). However, a keratin exhibits a 5.1 -A spac¬  ing rather than the 5.4-A distance corresponding to the pitch of the a helix.  This discrepancy is the result of two a keratin polypeptides, each of which forms  an a helix, twisting around each other to form a left-handed coil. The normal  5.4-A repeat distance of each a helix in the pair is thereby tilted relative to the  axis of this assembly, yielding the observed 5.1 -A spacing. The assembly is said  to have a coiled coil structure because each a helix itself follows a helical path.   The conformation of a keratins coiled coil is a consequence of its pri¬  mary structure: The central ~310-residue segment of each polypeptide chain  has a 7-residue pseudorepeat, a-b-c-d-e-fg, with nonpolar residues predomi¬  nating at positions a and d. Since an a helix has 3.6 residues per turn, a ker¬  atins a and d residues line up along one side of each a helix (Fig. 6-15^).  The hydrophobic strip along one helix associates with the hydrophobic strip  on another helix. Because the 3.5-residue repeat in a keratin is slightly smaller  than the 3.6 residues per turn of a standard a helix, the two keratin helices  are inclined about 18° relative to one another, resulting in the coiled coil  arrangement (Fig. 6-15 b). Coiled coils also occur in numerous other proteins,  some of which are globular rather than fibrous.   The higher order structure of a keratin is not well understood. The  N- and C-terminal domains of each polypeptide facilitate the assembly of  coiled coils (dimers) into protofilaments, two of which constitute a protofib¬  ril (Fig. 6-16). Four protofibrils constitute a microfibril, which associates with  other microfibrils to form a macrofibril. A single mammalian  hair consists of layers of dead cells, each of which is packed  with parallel macrofibrils.   a Keratin is rich in Cys residues, which form disulfide  bonds that cross-link adjacent polypeptide chains. The a ker¬  atins are classified as “hard” or “soft” according to whether  they have a high or low sulfur content. Hard keratins, such as  those of hair, horn, and nail, are less pliable than soft keratins,  such as those of skin and callus, because the disulfide bonds  resist deformation. The disulfide bonds can be reductively  cleaved by disulfide interchange with mercaptans (Section 5-  3A). Hair so treated can be curled and set in a “permanent  wave” by applying an oxidizing agent that reestablishes the  disulfide bonds in the new “curled” conformation. Conversely,  curly hair can be straightened by the same process.   FIG. 6-16 Higher order ol keratin structure, (a) Two keratin polypeptides form a  dimeric coiled coil, (b) Protofilaments are formed from two staggered rows of head-to-tail  associated coiled coils, (c) Protofilaments dimerize to form a protofibril, four of which form  a microfibril. The structures of the latter assemblies are poorly characterized.   Chapter 6 Proteins: Three-Dimensional Structure The springiness of hair and wool fibers is a consequence of the coiled coils   tendency to recover its original conformation after being untwisted by stretch¬  ing. If some of its disulfide bonds have been cleaved, however, an a keratin  fiber can be stretched to over twice its original length. At this point, the  polypeptide chains assume a (3 sheet conformation. (3 Keratin, such as that in  feathers, exhibits a (3-like pattern in its native state.   Left-handed polypeptide helices are twisted  together to form a right-handed superhelical  structure. [Illustration, Irving Geis. Image from the  Irving Geis Collection/Howard Hughes Medical  Institute. Rights owned by HHMI. Reproduction  by permission only.]    Collagen Is a Triple Helix. Collagen, which occurs in all multicellular ani¬  mals, is the most abundant vertebrate protein. Its strong, insoluble fibers are  the major stress-bearing components of connective tissues such as bone, teeth,  cartilage, tendon, and the fibrous matrices of skin and blood vessels. A single  collagen molecule consists of three polypeptide chains. Vertebrates have 46 ge¬  netically distinct polypeptide chains that are assembled into 28 collagen vari¬  eties found in different tissues in the same individual. One of the most  common collagens, called Type I, consists of two 0^(1) chains and one a 2 (I)  chain. It has a molecular mass of —285 kD, a width of —14 A, and a length  of -3000 A.   Collagen has a distinctive amino acid composition: Nearly one-third of its  residues are Gly; another 15 to 30% of its residues are Pro and 4-hydroxypro-  lyl (Hyp). 3-Hydroxyprolyl and 5-hydroxylysyl (Hyl) residues also occur in  collagen, but in smaller amounts.    These nonstandard residues are formed after the collagen polypeptides are syn¬  thesized. For example, Pro residues are converted to Hyp in a reaction cat¬  alyzed by prolyl hydroxylase. This enzyme requires ascorbic acid (vitamin C)  to maintain its activity. The disease scurvy results from the dietary deficiency  of vitamin C (Box 6-2).   The amino acid sequence of a typical collagen polypeptide consists of  monotonously repeating triplets of sequence Gly-X-Y over a segment of — 1000  residues, where X is often Pro and Y is often Hyp. Hyl sometimes appears at  the Y position. Collagens Pro residues prevent it from forming an a helix (Pro  residues cannot assume the a-helical backbone conformation and lack the  backbone N—H groups that form the intrahelical hydrogen bonds shown in  Fig. 6-7). Instead, the collagen polypeptide assumes a left-handed helical confor¬  mation with about three residues per turn. Three parallel chains wind around  each other with a gentle, right-handed, ropelike twist to form the triple-helical  structure of a collagen molecule (Fig. 6-17).   This model of the collagen structure has been confirmed by Barbara  Brodsky and Helen Berman, who determined the X-ray crystal structure of a  collagen-like model polypeptide. Every third residue of each polypeptide chain    Some collagen diseases have dietary causes. In scurvy (caused by  vitamin C deficiency), Hyp production decreases because prolyl  hydroxylase requires vitamin C. Thus, in the absence of vitamin C,  newly synthesized collagen cannot form fibers properly, resulting in  skin lesions, fragile blood vessels, poor wound healing, and, ulti¬  mately, death. Scurvy was common in sailors whose diets were  devoid of fresh foods on long voyages. The introduction of limes  to the diet of the British navy by the renowned explorer Captain  James Cook alleviated scurvy and led to the nickname “limey” for  the British sailor.   The disease lathyrism is caused by regular ingestion of the  seeds from the sweet pea Lathyrus odoratus, which contain a  compound that specifically inactivates lysyl oxidase (see below).  The resulting reduced cross-linking of collagen fibers produces  serious abnormalities of the bones, joints, and large blood vessels.   Several rare heritable disorders of collagen are known. Muta¬  tions of Type I collagen, which constitutes the major structural  protein in most human tissues, usually result in osteogenesis  imperfecta (brittle bone disease). The severity of this disease    varies with the nature and position of the mutation: Even a single  amino acid change can have lethal consequences. For example,  the central Gly —» Ala substitution in the model polypeptide  shown in Fig. 6-18 locally distorts the already internally crowded  collagen helix. This ruptures the hydrogen bond from the  backbone N—H of each Ala (normally Gly) to the carbonyl group  of the adjacent Pro in a neighboring chain, thereby reducing the  stability of the collagen structure. Mutations may affect the struc¬  ture of the collagen molecule or how it forms fibrils. These muta¬  tions tend to be dominant because they affect either the folding  of the triple helix or fibril formation even when normal chains are  also involved.   Many collagen disorders are characterized by deficiencies in the  amount of a particular collagen type synthesized, or by abnormal  activities of collagen-processing enzymes such as lysyl hydroxylase  and lysyl oxidase. One group of at least 10 different collagen-  deficiency diseases, the Ehlers-Danlos syndromes, are all char¬  acterized by the hyperextensibility of the joints and skin. The  “India-rubber man” of circus fame had an Ehlers-Danlos syndrome.    passes through the center of the triple helix, which is so crowded that only a  Gly side chain can fit there. This crowding explains the absolute requirement  for a Gly at every third position of a collagen polypeptide chain. The three  polypeptide chains are staggered so that a Gly, X, and Y residue occurs at each  level along the triple helix axis (Fig. 6-18#). The peptide groups are oriented  such that the N—H of each Gly makes a strong hydrogen bond with the car¬  bonyl oxygen of an X (Pro) residue on a neighboring chain (Fig. 6-18^). The  bulky and relatively inflexible Pro and Hyp residues confer rigidity on the en¬  tire assembly.   FIG. 6-1 8 Structure of a collagen model  peptide. In this X-ray structure of (Pro-Hyp-  Gly) 10 , the fifth Gly of each peptide has been  replaced by Ala. (a) A stick model of the middle  portion of the triple helix oriented with its  N-termini at the bottom. The C atoms of the  three chains are colored orange, magenta, and  gray. The N and 0 atoms on all chains are blue  and red. Note how the replacement of Gly with  the bulkier Ala (C atoms green) distorts the  triple helix, (b) This view from the N-terminus  down the helix axis shows the interchain  hydrogen-bonding associations. Three  consecutive residues from each chain are shown  in stick form (C atoms green). Hydrogen bonds  are represented by dashed lines from Gly N  atoms to Pro 0 atoms in adjacent chains. Dots  represent the van der Waals surfaces of the  backbone atoms of the central residue in each  chain. Note the close packing of the atoms  along the triple helix axis. [Based on an X-ray  structure by Helen Berman, Rutgers University,  and Barbara Brodsky, UMDNJ-Robert Wood  Johnson Medical School. PDBid 1CAGJ ©See  Kinemage Exercises 4-3 and 4-4.   • Describe the four levels of protein struc¬  ture. Do all proteins exhibit all four levels?   • Without looking at Fig. 6-3, draw the  structure of a polypeptide backbone.   • Explain why the conformational freedom of  peptide bonds is limited.   • Summarize the features of an a helix and  a parallel and antiparallel p sheet.   • What properties do fibrous proteins confer  on substances such as hair and bones?   • Describe the features of the amino acid  sequences that are necessary to form a  coiled coil or a left-handed triple helix.    Collagens well-packed, rigid, triple-helical structure is responsible for its  characteristic tensile strength. The twist in the helix cannot be pulled out un¬  der tension because its component polypeptide chains are twisted in the op¬  posite direction (Fig. 6-17). Successive levels of fiber bundles in high-quality  ropes and cables, as well as in other proteins such as keratin (Fig. 6-16), are  likewise oppositely twisted.   Several types of collagen molecules assemble to form loose networks or  thick fibrils arranged in bundles or sheets, depending on the tissue. The colla¬  gen molecules in fibrils are organized in staggered arrays that are stabilized by  hydrophobic interactions resulting from the close packing of triple-helical units.  Collagen is also covalently cross-linked, which accounts for its poor solubility.  The cross-links cannot be disulfide bonds, as in keratin, because collagen is al¬  most devoid of Cys residues. Instead, the cross-links are derived from Lys and  His side chains in reactions such as those shown in Fig. 6-19. Lysyl oxidase,  the enzyme that converts Lys residues to those of the aldehyde allysine, is the  only enzyme implicated in this cross-linking process. Up to four side chains  can be covalently bonded to each other. The cross-links do not form at ran¬  dom but tend to occur near the N- and C-termini of the collagen molecules.  The degree of cross-linking in a particular tissue increases with age. This is why  meat from older animals is tougher than meat from younger animals.   The majority of proteins are globular proteins that, unlike the fibrous proteins  discussed above, may contain several types of regular secondary structure, includ¬  ing a helices, (3 sheets, and other recognizable elements. A significant portion of  a proteins structure may also be irregular or unique. Segments of polypeptide  chains whose successive residues do not have similar cj) and i(i values are some¬  times called coils. However, you should not confuse this term with the appella¬  tion random coil, which refers to the totally disordered and rapidly fluctuating  conformations assumed by denatured (fully unfolded) proteins in solution. In  native (folded) proteins, nonrepetitive structures are no less ordered than are helices  or /3 sheets; they are simply irregular and hence more difficult to describe.   Sequence Affects Secondary Structure. Variations in amino acid sequence as  well as the overall structure of the folded protein can distort the regular confor¬  mations of secondary structural elements. For example, the a helix frequently de¬  viates from its ideal conformation in the initial and final turns of the helix.  Similarly, a strand of polypeptide in a (3 sheet may contain an “extra” residue  that is not hydrogen bonded to a neighboring strand, producing a distortion  known as a (3 bulge.   Many of the limits on amino acid composition and sequence (Section 5-1)  may be due in part to conformational constraints in the three-dimensional struc¬  ture of proteins. For example, a Pro residue produces a kink in an a helix or (3  sheet. Similarly, steric clashes between several sequential amino acid residues with  large branched side chains (e.g., lie and Tyr) can destabilize a helices.   Analysis of known protein structures by Peter Chou and Gerald Fasman  revealed the propensity P of a residue to occur in an a helix or a (3 sheet  (Table 6-1). Chou and Fasman also discovered that certain residues not only  have a high propensity for a particular secondary structure but they tend to  disrupt or break other secondary structures. Such data are useful for predict¬  ing the secondary structures of proteins with known amino acid sequences.   The presence of certain residues outside of a helices or (3 sheets may also  be nonrandom. For example, a helices are often flanked by residues such as  Asn and Gin, whose side chains can fold back to form hydrogen bonds with  one of the four terminal residues of the helix, a phenomenon termed helix  capping. Recall that the four residues at each end of an a helix are not fully  hydrogen bonded to neighboring backbone segments (Fig. 6-7).    TABLE 6-1 Propensities of Amino Acid  Residues for a Helical and  (3 Sheet Conformations    Source: Chou, P.Y. and Fasman, G.D., Annu. Rev.  Biochem. 47, 258 (1978).    FIG. 6-19 A reaction pathway for cross-linking side chains in collagen. The first  step is the lysyl oxidase-catalyzed oxidative deamination of Lys to form the aldehyde  allysine. Two allysines then undergo an aldol condensation to form allysine aldol. This  product can react with His to form aldol histidine, which can in turn react with 5-  hydroxylysine to form a Schiff base (an imine bond), thereby cross-linking four side chains.   • X-Ray crystallography and NMR spectroscopy are used to determine the positions  of atoms in proteins.   • Nonpolar residues tend to occur in the protein interior and polar residues on the  exterior.   • A protein’s tertiary structure consists of secondary structural elements that  combine to form motifs and domains.   • Over time, a protein’s structure is more highly conserved than its sequence.   • Bioinformatics databases store macromolecular structure coordinates. Software  makes it possible to visualize proteins and compare their structural features.   The tertiary structure of a protein describes the folding of its secondary struc¬  tural elements and specifies the positions of each atom in the protein, includ¬  ing its side chains. This information is deposited in a database and is readily  available via the Internet, which allows the tertiary structures of a variety of  proteins to be analyzed and compared. The common features of protein ter¬  tiary structures reveal much about the biological functions of proteins and  their evolutionary origins.   A Most Protein Structures Are Determined by X-Ray Crystallography  or Nuclear Magnetic Resonance   X-Ray crystallography is a technique that directly images molecules. X-Rays  must be used to do so because, according to optical principles, the uncertainty  in locating an object is approximately equal to the wavelength of the radia¬  tion used to observe it (covalent bond distances and the wavelengths of the  X-rays used in structural studies are both ~ 1.5 A; individual molecules can¬  not be seen in a light microscope because visible light has a minimum wave¬  length of 4000 A). There is, however, no such thing as an X-ray microscope  because there are no X-ray lenses. Rather, a crystal of the molecule to be im¬  aged (e.g., Fig. 6-20) is exposed to a collimated beam of X-rays and the re¬  sulting diffraction pattern, which arises from the regularly repeating positions  of atoms in the crystal, is recorded by a radiation detector or, now infrequently,  on photographic film (Fig. 6-21). The X-rays used in structural studies are  produced by laboratory X-ray generators or, now commonly, by synchrotrons,  particle accelerators that produce X-rays of far greater intensity. The intensi¬  ties of the diffraction maxima (darkness of the spots on a film) are then used  to construct mathematically the three-dimensional image of the crystal struc¬  ture through methods that are beyond the scope of this text. In what follows,  we discuss some of the special problems associated with interpreting the X-ray  crystal structures of proteins.   X-Rays interact almost exclusively with the electrons in matter, not the  nuclei. An X-ray structure is therefore an image of the electron density of the    FIG. 6-20 Protein crystals, (a) Azurin from  Pseudomonas aeruginosa, ( b ) flavodoxin from  Desulfovibrio vulgaris, and (c) rubredoxin from  Clostridium pasteurianum. These crystals are  colored because the proteins contain light¬  absorbing groups; proteins are colorless in the  absence of such groups. [Courtesy of Larry  Siecker, University of Washington.]   FIG. 6-21 An X-ray diffraction photograph  of a crystal of sperm whale myoglobin. The   intensity of each diffraction maximum (the  darkness of each spot) is a function of the  crystal’s electron density. [Courtesy of John  Kendrew, Cambridge University, U.K.]    object under study. Such electron density maps are usually presented with  the aid of computer graphics as one or more sets of contours, in which a con¬  tour represents a specific level of electron density in the same way that a con¬  tour on a topographic map indicates locations that have a particular altitude.  A portion of an electron density map of a protein is shown in Fig. 6-22.   molecules in protein crystals, as in other crystalline substances, are arranged  in regularly repeating three-dimensional lattices. Protein crystals, however, dif¬  fer from those of most small organic and inorganic molecules in being highly  hydrated; they are typically 40 to 60% water by volume. The aqueous solvent  of crystallization is necessary for the structural integrity of the protein crys¬  tals, because water is required for the structural integrity of native proteins  themselves (Section 6-4).   The large solvent content of protein crystals gives them a soft, jellylike  consistency so that their molecules usually lack the rigid order characteristic  of crystals of small molecules such as NaCl or glycine. The molecules in a pro¬  tein crystal are typically disordered by more than an angstrom, so the corre¬  sponding electron density map lacks information concerning structural details  of smaller size. The crystal is therefore said to have a resolution limit of that  size. Protein crystals typically have resolution limits in the range 1.5 to 3.0 A,  although some are better ordered (have higher resolution, that is, a lesser res¬  olution limit) and many are less ordered (have lower resolution).   FIG. 6-22 A thin section through a  1.5-A-resolution electron density map of a  protein that is contoured in three  dimensions. Only a single contour level (cyan)  is shown, together with a ball-and-stick model of  the corresponding polypeptide segments colored  according to atom type with C yellow, N blue,  and 0 red. A water molecule is represented  by a red sphere. [Courtesy of Xinhua Ji,    FIG. 6-23 Electron density maps of  diketopiperazine at different resolution levels.   obtain crystals of sufficiently high resolution is a major limiting factor in de¬  termining the X-ray crystal structure of a protein or other macromolecule.  Figure 6-23 indicates how the quality (degree of focus) of an electron density  map varies with its resolution limit. At 6-A resolution, the presence of a mol¬  ecule the size of diketopiperazine is difficult to discern. At 2.0-A resolution,  its individual atoms cannot yet be distinguished, although its molecular shape  has become reasonably evident. At 1.5-A resolution, which roughly corre¬  sponds to a bond distance, individual atoms become partially resolved. At 1.1-A  resolution, atoms are clearly visible.   Most protein crystal structures are too poorly resolved for their electron  density maps to reveal clearly the positions of individual atoms (e.g., Fig. 6-23).  Nevertheless, the distinctive shape of the polypeptide backbone usually permits  it to be traced, which, in turn, allows the positions and orientations of its side  chains to be deduced (e.g., Fig. 6-22). Yet side chains of comparable size and  shape, such as those of Leu, lie, Thr, and Val, cannot always be differentiated  (hydrogen atoms, having only one electron, are visible only if the resolution  limit is less than ~1.2 A). Consequently, a protein structure cannot be eluci¬  dated from its electron density map alone, but knowing the primary structure  of the protein permits the sequence of amino acid residues to be fitted to the  electron density map. Mathematical refinement can then reduce the uncertainty  in the crystal structures atomic positions to as little as 0.1 A.   Most Crystalline Proteins Maintain Their Native Conformations. Does the  structure of a protein in a crystal accurately reflect the structure of the pro¬  tein in solution, where globular proteins normally function? Several lines of  evidence indicate that crystalline proteins assume very nearly the same structures  that they have in solution:   1. A protein molecule in a crystal is essentially in solution because it is  bathed by solvent of crystallization over all of its surface except for the  few, generally small patches that contact neighboring protein molecules.   2. In cases where different crystal forms of a protein have been analyzed,  or when a crystal structure has been compared to a solution structure  (determined by NMR; see below), the molecules have virtually identi¬  cal conformations. Evidently, crystal packing forces do not greatly per¬  turb the structures of protein molecules.   3. Many enzymes are catalytically active in the crystalline state. Since  the activity of an enzyme is very sensitive to the positions of the  groups involved in binding and catalysis (Chapter 11), the crystalline  enzymes must have conformations that closely resemble their solu¬  tion conformations.    FIG. 6-24 NOESY spectrum of a protein. The diagonal  represents the conventional one-dimensional NMR spectrum presented  as a contour plot Note that it is too crowded with peaks to be directly  interpretable (even a small protein has hundreds of protons). The cross  (off-diagonal) peaks each arise from the interaction of two protons  that are <5 A apart in space (their one-dimensional NMR peaks are  located where horizontal and vertical lines intersect the diagonal). The  line to the left of the spectrum represents the extended polypeptide  chain with its N- and C-termini labeled N and C and the positions of  four protons labeled a to d. The dashed arrows indicate the diagonal  NMR peaks to which these protons give rise. Cross peaks, such as i, j,  and k, each located at the intersection of the corresponding horizontal  and vertical lines, show that two protons are <5 A apart. These  distance relationships are schematically drawn as three looped  structures of the polypeptide chain below the spectrum. The  assignment of a distance relationship between two protons in a  polypeptide requires that the NMR peaks to which they give rise and  their positions in the polypeptide be known, which requires that the  polypeptide’s amino acid sequence has been previously determined.  [After Wuthrich, K., Science 243, 45 (1989).]    Protein Structures Can Be Determined by NMR. The basis of nuclear mag¬  netic resonance (NMR) is the observation that an atomic nucleus, such as a  proton (a hydrogen nucleus), resonates in an applied magnetic field in a way  that is sensitive to its electronic environment and its interactions with nearby  nuclei. The development of NMR techniques, since the mid-1980s, in large  part by Kurt Wuthrich, has made it possible to determine the three-dimensional  structures of small globular proteins in aqueous solution.   A proteins conventional (one-dimensional) NMR spectrum is crowded  with overlapping peaks, since even a small protein has hundreds of protons.  This problem is addressed by two-dimensional (2D) NMR spectroscopy,  which yields additional peaks arising from the interactions of protons that are  less than 5 A apart. Correlation spectroscopy (COSY) provides interatomic  distances between protons that are covalently connected through one or two  other atoms, such as the H atoms attached to the N and C a of the same amino  acid (corresponding to the 4> torsion angle). Nuclear Overhauser spectroscopy  (NOESY) provides interatomic distances for protons that are close in space al¬  though they may be far apart in the protein sequence. An example of a  NOESY spectrum is shown in Fig. 6-24.   Interatomic distance measurements, along with knowledge of the proteins  sequence and known geometric constraints such as covalent bond distances  and angles, group planarity, chirality, and van der Waals radii, are used to com¬  pute the proteins three-dimensional structure. However, since interproton dis¬  tance measurements are imprecise, they cannot imply a unique structure but  rather are consistent with an ensemble of closely related structures.  Consequently, an NMR structure of a protein (or another macromolecule) is  often presented as a sample of structures that are consistent with the data (e.g.,  Fig. 6-25). The “tightness” of a bundle of such structures is indicative both    FIG.6-25 The NMR structure of a protein. The drawing represents 20 superimposed  structures of a 64-residue polypeptide comprising the Src protein SH3 domain (Section  13-2B). The polypeptide backbone (its connected C a atoms) is white, and its Phe, Tyr, and  Trp side chains are yellow, red, and blue, respectively. The polypeptide backbone folds into  two 3-stranded antiparallel |3 sheets that form a sandwich. [Courtesy of Stuart Schreiber,  Harvard University.]   Chapter 6 Proteins: Three-Dimensional Structure G f the accuracy with which the structure is known, which in the most favor¬  able cases is roughly comparable to that of an X-ray crystal structure with a  resolution of 2 to 2.5 A, and of the conformational fluctuations that the pro¬  tein undergoes (Section 6-4A). Although present NMR methods are limited  to determining the structures of macromolecules with molecular masses no  greater than —100 kD, recent advances in NMR technology suggest that this  limit may soon increase to ~1000 kD or more.   NMR methods, besides validating the structures of proteins analyzed by  X-ray crystallography (or in some cases identifying protein residues that are  perturbed by crystallization), can determine the structures of proteins and  other macromolecules that fail to crystallize. Moreover, since NMR can probe  motions over time scales spanning 10 orders of magnitude, it can also be used  to study protein folding and dynamics (Sections 6-4 and 6-5).   Proteins Can Be Depicted in Different Ways. The huge number of atoms  in proteins makes it difficult to visualize them using the same sorts of mod¬  els employed for small organic molecules. Ball-and-stick representations show¬  ing all or most atoms in a protein (as in Figs. 6-7 and 6-10) are exceedingly  cluttered, and space-filling models (as in Figs. 6-8 and 6-11) obscure the in¬  ternal details of the protein. Accordingly, computer-generated or artistic ren¬  ditions (e.g., Fig. 6-12) are often more useful for representing protein  structures. The course of the polypeptide chain can be followed by tracing the  positions of its C a atoms or by representing helices as helical ribbons or cylin¬  ders and (3 sheets as sets of flat arrows pointing from the N- to the C-termini.   In the years since Kendrew solved the structure of myoglobin, nearly 80,000  protein structures have been reported. No two are exactly alike, but they ex¬  hibit remarkable consistencies. The primary structures of globular proteins  generally lack the repeating sequences that support the regular conformations  seen in fibrous proteins. However, the amino acid side chains in globular pro¬  teins are spatially distributed according to their polarities:   1. The nonpolar residues Val, Leu, lie, Met, and Phe occur mostly in the  interior of a protein, out of contact with the aqueous solvent. The hy¬  drophobic effects that promote this distribution are largely responsible  for the three-dimensional structure of native proteins.   2. The charged polar residues Arg, His, Lys, Asp, and Glu are usually lo¬  cated on the surface of a protein in contact with the aqueous solvent.  This is because immersing an ion in the virtually anhydrous interior  of a protein is energetically unfavorable.   3. The uncharged polar groups Ser, Thr, Asn, Gin, and Tyr are usually  on the protein surface but also occur in the interior of the molecule.  When buried in the protein, these residues are almost always hydro¬  gen bonded to other groups; in a sense, the formation of a hydrogen  bond “neutralizes” their polarity. This is also the case with the polypep¬  tide backbone.   These general principles of side chain distribution are evident in individ¬  ual elements of secondary structure (Fig. 6-26) as well as in whole proteins  (Fig. 6 -27). Polar side chains tend to extend toward—and thereby help  form—the proteins surface, whereas nonpolar side chains largely extend  toward—and thereby occupy—its interior. Turns and loops joining second¬  ary structural elements usually occur at the protein surface.   Most proteins are quite compact, with their interior atoms packed together  even more efficiently than the atoms in a crystal of small organic molecules.  Nevertheless, the atoms of protein side chains almost invariably have low-  energy arrangements. Evidently, interior side chains adopt relaxed conformations    FIG.6-26 Side chain locations in an a  helix and a p sheet. In these space-filling  models, the main chain is gray, nonpolar side  chains are gold, and polar side chains are purple,  (a) An a helix from sperm whale myoglobin.   Note that the nonpolar residues are primarily on  one side of the helix, (b) An antiparallel |3 sheet  from con can aval in A (side view). The protein  interior is to the right and the exterior is to the  left. [Based on X-ray structures by lime  Schlichting, Max Planck Institut fur Molekulare  Physiologie, Dortmund, Germany, and Gerald  Edelman, The Rockefeller University. PDBids  1A6M and 2CNA.]   c. In these paintings, based on an X-ray structure determined by  Richard Dickerson, the protein is illuminated by its single iron atom  centered in a heme group. Hydrogen atoms are not shown. In (a) the   hydrophilic side chains are green, and in ( b ) the hydrophobic side  chains are orange. [Illustration, Irving Geis. Image from the Irving Geis  Collection/Howard Hughes Medical Institute. Rights owned by HHMI.  Reproduction by permission only.] O See Kinemage Exercise 5.   Chapter 6 Proteins: Three-Dimensional Structure despite the profusion of intramolecular interactions. Closely packed protein   interiors generally exclude water. When water molecules are present, they of¬  ten occupy specific positions where they can form hydrogen bonds, sometimes  acting as a bridge between two hydrogen-bonding protein groups.   Globular proteins—each with a unique tertiary structure—are built from com¬  binations of secondary structural elements. The proportions of a helices and  (3 sheets and the order in which they are connected provide an informative  way of classifying and analyzing protein structure.   Certain Combinations of Secondary Structure Form Motifs. Groupings of  secondary structural elements, called supersecondary structures or motifs,  occur in many unrelated globular proteins:   1. The most common form of supersecondary structure is the (3ot(3 mo¬  tif, in which an a helix connects two parallel strands of a (3 sheet  (Fig. 6-28^).   2. Another common supersecondary structure, the P hairpin motif, con¬  sists of antiparallel strands connected by relatively tight reverse turns  (Fig. 6-28 £).   3. In an olol motif, two successive antiparallel a helices pack against each  other with their axes inclined. This permits energetically favorable in¬  termeshing of their contacting side chains (Fig. 6-28 c). Similar associ¬  ations stabilize the coiled coil conformation of a keratin and  tropomyosin (Fig. 6-15^), although their helices are parallel rather than  antiparallel.   4. In the Greek key motif (Fig. 6-2 Sd; named after an ornamental de¬  sign commonly used in ancient Greece; see inset), a (3 hairpin is folded  over to form a 4-stranded antiparallel (3 sheet.   Most Proteins Can Be Classified as a, p, or a/p. The major types of second¬  ary structural elements occur in globular proteins in varying proportions and  combinations. Some proteins, such as E. coli cytochrome b^ 2 (Kg« 6-29 d),  consist only of a helices spanned by short connecting links and are therefore   (a) A pap motif, (b) a p hairpin motif, (c) an aa motif, and ( d ) a  Greek key motif, showing how it is constructed from a folded-over p    hairpin. The polypeptide backbones are drawn as ribbons, with p  strands shown as flat arrows pointing from N- to C-terminus, and a  helices represented by cylinders.    FIG. 6-29 A selection of protein structures. The proteins are represented by their  peptide backbones, drawn in ribbon form, with p strands shown as flat arrows pointing from  N- to C-terminus and a helices depicted as coils. The polypeptide chain is colored, from  N- to C-terminus, in rainbow order from red to blue. Below each drawing is the corresponding  topological diagram indicating the connectivity of its helices (cylinders) and p strands (flat  arrows), (a) E. coli cytochrome b 562 (106 residues), which forms an up-down-up-down  4-helix bundle. Its bound heme group is shown in ball-and-stick form with C magenta, N blue,  0 red, and Fe orange, (b) The N-terminal domain of the human immunoglobulin fragment  Fab New (103 residues) showing its immunoglobulin fold. The polypeptide chain is folded  into a sandwich of 3- and 4-stranded antiparallel p sheets, (c) The N-terminal domain of  dogfish lactate dehydrogenase (163 residues). It contains a 6-stranded parallel p sheet in  which the crossovers between p strands all contain an a helix that forms a right-handed  helical turn with its flanking p strands. [Based on X-ray structures by (a) F. Scott Matthews,  Washington University School of Medicine; (b) Roberto Poljak, The Johns Hopkins School  of Medicine; and (c) Michael Rossmann, Purdue University. PDBids (a) 256B, (b) 7FAB, and  (c) 6LDH.]    classified as ol proteins. Others, such as immunoglobulins, contain the im¬  munoglobulin fold (Fig. 6-29 b), and are called P proteins because they have  a large proportion of p sheets and are devoid of a helices. Most proteins, how¬  ever, including lactate dehydrogenase (Fig. 6-29 c) and carboxypeptidase A  (Fig. 6-12), are known as ct/fl proteins because they largely consist of mixtures  of both types of secondary structure (proteins, on average, contain —31%  a helix and —28% p sheet).   The a, p, and a/p classes of proteins can be further subdivided by their  topology, that is, according to how their secondary structural elements are con¬  nected. For example, extended P sheets often roll up to form P barrels. Three  different types of 8-stranded P barrels, each with a different topology, are    FIG. 6-30 X-Ray structures of p barrels. Each polypeptide is drawn and colored and  accompanied by its corresponding topological diagram as is described in the legend to Fig.  6-29. (a) Human retinol binding protein showing its 8-stranded up-and-down p barrel  (residues 1-142 of the 182-residue protein). Note that each p strand is linked via a short  loop to its clockwise-adjacent strand as seen from the top. The protein’s bound retinol  molecule is represented by a gray ball-and-stick model, (b) Peptide-yv 4 -(W-acetyl-p-D-  glucosaminyl)asparagine amidase F from Flavobacterium meningosepticum (residues  1-140 of the 340-residue enzyme). Note how its 8-stranded p barrel is formed by rolling  up a 4-segment p hairpin. Here the two p strands in each segment of the p hairpin are  colored alike with strands 1 and 8 (the N- and C-terminal strands) red, strands 2 and 7  orange, strands 3 and 6 cyan, and strands 4 and 5 blue. This motif, which is known as a  jelly roll or Swiss roll barrel, is so named because of its topological resemblance to the  rolled-up pastries, (c) Chicken muscle triose phosphate isomerase (TIM; 247 residues)  forms a so-called a/p barrel in which 8 pairs of alternating p strands and a helices roll up  to form an inner barrel of 8 parallel p strands surrounded by an outer barrel of 8 parallel a  helices. The protein is viewed approximately along the axis of the a/p barrel. Note that the  a/p barrel is essentially a series of linked pap motifs. [Based on X-ray structures by (a) T.  Alwyn Jones, Biomedical Center, Uppsala, Sweden; (b) Patrick Van Roey, New York State  Department of Health, Albany, New York; and (c) David Phillips, Oxford University, Oxford,  U.K. PDBids (a) 1 RBP, (b) 1 PNG, and (c) 1TIMJ    shown in Fig. 6-30. Two of these (Fig. 6-30 a,B) are all-(3 structures contain¬  ing multiple (3 hairpin motifs. The third, known as an a/|3 barrel (Fig. 6-30 c),  can be considered as a set of overlapping (3a(3 motifs (and is a member of the  a/(3 class of proteins).   Large Polypeptides Form Domains. Polypeptide chains containing more  than ~200 residues usually fold into two or more globular clusters known as  domains, which give these proteins a bi- or multilobal appearance. Each subunit   The N-terminal domain (light blue) binds NAD + (drawn in stick form and colored according  to atom type with C green, N blue, 0 red, and P magenta), and the C-terminal domain  ( orange ) binds glyceraldehyde-3-phosphate (not shown). [Based on an X-ray structure by  Alan Wonacott, Imperial College, London, U.K. PDBid 1GD1.] O See Interactive Exercise 2.    of the enzyme glyceraldehyde-3-phosphate dehydrogenase, for example, has  two distinct domains (Fig. 6-31). Most domains consist of 40 to 200 amino  acid residues and have an average diameter of ~25 A. An inspection of the  various protein structures diagrammed in this chapter reveals that domains  consist of two or more layers of secondary structural elements. The reason for  this is clear: At least two such layers are required to seal off a domains  hydrophobic core from its aqueous environment.   A polypeptide chain wanders back and forth within a domain, but neigh¬  boring domains are usually connected by only one or two polypeptide seg¬  ments. Consequently, many domains are structurally independent units that have  the characteristics of small globular proteins. Nevertheless, the domain structure  of a protein is not necessarily obvious since its domains may make such  extensive contacts with each other that the protein appears to be a single glob¬  ular entity.   Domains often have a specific function such as the binding of a small  molecule. In Fig. 6-31, for example, the dinucleotide NAD + (nicotinamide  adenine dinucleotide; Fig. 11-4) binds to the N-terminal domain of glycer-  aldehyde-3-phosphate dehydrogenase. Michael Rossmann has shown that a  (3a(3a|3 unit, in which the |3 strands form a parallel sheet with a helical con¬  nections, often acts as a nucleotide-binding site. Two of these (3a|3a(3 units  combine to form a domain known as a dinucleotide-binding fold, or  Rossmann fold. Glyceraldehyde-3-phosphate dehydrogenases N-terminal do¬  main contains such a fold, as does lactate dehydrogenase (Fig. 6-29 e). In some  multidomain proteins, binding sites occupy the clefts between domains; that  is, small molecules are bound by groups from two domains. In such cases, the  relatively pliant covalent connection between the domains allows flexible in¬  teractions between the protein and the small molecule.   The many thousands of known protein structures, comprising an even greater  number of separate domains, can be grouped into families by examining the  overall paths followed by their polypeptide chains. Although it is estimated  that there are as many as 1400 different protein domain families, approxi¬  mately 200 different folding patterns account for about half of all known pro¬  tein structures. As described in Section 5-4B, the domain is the fundamental  unit of protein evolution. Apparently, the most common protein domains are  evolutionary sinks—domains that arose and persisted because of their ability  (1) to form stable folding patterns; (2) to tolerate amino acid deletions, sub¬  stitutions, and insertions, thereby making them more likely to survive evolu¬  tionary changes; and/or (3) to support essential biological functions.   Polypeptides with similar sequences tend to adopt similar backbone con¬  formations. This is certainly true for evolutionarily related proteins that carry  out similar functions. For example, the cytochromes c of different species are  highly conserved proteins with closely similar sequences (Table 5-6) and three-  dimensional structures.   Cytochrome c occurs only in eukaryotes, but prokaryotes contain pro¬  teins, known as c-type cytochromes, which perform the same general func¬  tion (that of an electron carrier). The c-type cytochromes from different  species exhibit only low degrees of sequence similarity to each other and to  eukaryotic cytochromes c. Yet their X-ray structures are clearly similar,   134 amino acid residues 112 amino acid residues 103 amino acid residues   FIG. 6-32 Three-dimensional structures of c-type  cytochromes. The polypeptide backbones (blue) are shown in  analogous orientations such that their heme groups (red) are viewed  edge-on. The Cys, Met, and His side chains that covalently link the  heme to the protein are also shown, (a) Cytochrome c 550 from  Paracoccus denitrificans, ( b ) cytochrome c 2 from Rhodospirillum    rubrum, (c) cytochrome c from tuna, and (d) cytochrome c 555 from  Chlorobium limicola. [Illustration, Irving Geis. Image from the Irving  Geis Collection/Howard Hughes Medical Institute. Rights owned by  HHMI. Reproduction by permission only.]    particularly in polypeptide chain folding and side chain packing in the pro¬  tein interior (Fig. 6-32). The major structural differences among c-type cy¬  tochromes lie in the various polypeptide loops on their surfaces. The  sequences of the c-type cytochromes have diverged so far from one another  that, in the absence of their X-ray structures, they can be properly aligned  only through the use of mathematically sophisticated computer programs.  Thus, it appears that the essential structural and functional elements of proteins,  rather than their amino acid residues , are conserved during evolution.   E Structural Bioinformatics Provides Tools for Storing, Visualizing,  and Comparing Protein Structural Information   The data obtained by X-ray crystallography, NMR spectroscopy, and certain  other techniques take the form of three-dimensional coordinates describing the  spatial positions of atoms in molecules. This kind of information can be eas¬  ily stored, displayed, and compared, much like sequence information obtained  by nucleotide or protein sequencing methods (see Sections 3-4 and 5-3).  Bioinformatics is the rapidly growing discipline that deals with the burgeon¬  ing amount of information related to molecular sequences and structures.  Structural bioinformatics is a branch of bioinformatics that is concerned with  how macromolecular structures are displayed and compared. Some of the data¬  bases and analytical tools that are used in structural bioinformatics are listed  in Table 6-2 and described in Bioinformatics Project 3.   The Protein Data Bank Is the Repository for Structural Information. The   atomic coordinates of nearly 80,000 macromolecular structures, including pro¬  teins, nucleic acids, and carbohydrates, are archived in the Protein Data Bank  (PDB). Indeed, most scientific journals that publish macromolecular struc¬  tures require that authors deposit their structures coordinates in the PDB.   Each independently determined structure in the PDB is assigned a unique  four-character identifier (its PDBid). For example, the PDBid for the struc¬  ture of sperm whale myoglobin is 1MBO. A coordinate file includes the  macromolecules source (the organism from which it was obtained), the au¬  thor (s) who determined the structure, key journal references, information  about how the structure was determined, and indicators of its accuracy. The  sequences of the structure’s various chains are then listed together with  the descriptions and formulas of its so-called HET (for heterogen) groups, which   Molecular Modeling Database (MMDB): http://www.ncbi.nlm.nih.gov/sites/entrez?db=structure  Most Representative NMR Structure in an Ensemble: http://www.ebi.ac.uk/msd-srv/olderado   CATH (Class, Architecture, Topology, and //omologous superfamily): http://www.cathdb.info/index.html  CE (Combinatorial Extension of optimal pathway): http://cl.sdsc.edu/    are molecular entities that are not among the “standard” amino acid or nucleotide  residues (for example, organic molecules, nonstandard residues such as Hyp,  metal ions, and bound water molecules). The positions of the structures sec¬  ondary structural elements and its disulfide bonds are then provided.   The bulk of a PDB file consists of a series of records (lines), each of which  provides the three-dimensional (x, y, z) coordinates in angstroms of one atom  in the structure. Each atom is identified by a serial number, an atom name (for  example, C and O for an amino acid residues carbonyl C and O atoms, CA  and CB for C a and Cp atoms), the name of the residue, and a letter to iden¬  tify the chain to which it belongs (for structures that have more than one chain).  For NMR-based structures, the PDB file contains a full set of records for each  member of the ensemble of structures (the most representative member of such  a coordinate set can be obtained from another database; see Table 6-2).   A particular PDB file may be located according to its PDBid or, if this is  unknown, by searching with a variety of criteria including a proteins name,  its source, or the author(s). Selecting a particular macromolecule in the PDB  initially displays a summary page with options for viewing the structure  (either statically or interactively), for viewing or downloading the coordinate file,  and for classifying or analyzing the structure in terms of its geometric prop¬  erties and sequence. The Nucleic Acid Database (NDB) archives the atomic  coordinates of structures that contain nucleic acids, using roughly the same  format as PDB files.   Molecular Graphics Programs Interactively Show Macromolecules in  Three Dimensions. The most informative way to examine a macromolecular  structure is through the use of molecular graphics programs that permit the  user to interactively rotate a macromolecule and thereby perceive its three-  dimensional structure. This impression may be further enhanced by simultane¬  ously viewing the macromolecule in stereo. Most molecular graphics programs  use PDB files as input. The programs described here can be downloaded from   Chapter 6 Proteins: Three-Dimensional Structure t h e Internet addresses listed in Table 6-2, some of which also provide instruc¬  tions for the programs use.   Jmol, which functions as both a Web browser-based applet or as a stand¬  alone program, allows the user to display user-selected macromolecules in a  variety of colors and formats (e.g., wire frame, ball-and-stick, backbone, space¬  filling, and cartoons). The Interactive Exercises on the website that accom¬  panies this textbook (http://www.wiley.com/college/voet/) all use Jmol.  FirstGlance uses Jmol to display macromolecules via a user-friendly interface.  KiNG, which also has Web browser-based and standalone versions, displays  the so-called Kinemages on this textbooks accompanying website. KiNG  provides a generally more author-directed user environment than does Jmol.  Macromolecules can be displayed directly from their corresponding PDB page  using either Jmol or KiNG. The Swiss-PDB Viewer (also called Deep View),  in addition to displaying molecular structures, provides tools for basic model  building, homology modeling, energy minimization, and multiple sequence  alignment. One advantage of the Swiss-PDB Viewer is that it allows users to  easily superimpose two or more models. Proteopedia is a 3D interactive en¬  cyclopedia of proteins and other macromolecules that resembles Wikipedia in  that it is user edited. It uses mainly Jmol as a viewer.   Structure Comparisons Reveal Evolutionary Relationships. Most proteins  are structurally related to other proteins, since evolution tends to conserve the  structures of proteins rather than their sequences. The computational tools de¬  scribed below facilitate the classification and comparison of protein structures.  These programs can be accessed directly via their Internet addresses and in  some cases through the PDB. Studies using these programs yield functional  insights, reveal distant evolutionary relationships that are not apparent from  sequence comparisons, generate libraries of unique folds for structure predic¬  tion, and provide indications as to why certain types of structures are preferred  over others.   1. CATH (for Class, Architecture, Topology, and Pfomologous superfam¬  ily), as its name suggests, categorizes proteins in a four-level structural  hierarchy: (1) Class, the highest level, places the selected protein in one  of four levels of gross secondary structure (Mainly a, Mainly (3, al (3,  and Few Secondary Structures); (2) Architecture, the gross arrange¬  ment of secondary structure; (3) Topology, which depends on both  the overall shape of the protein domain and the connectivity of its  secondary structural elements; and (4) Homologous Superfamily,  which identifies the protein as a member of a group that shares a com¬  mon ancestor.   2. CE (for Combinatorial Extension of the optimal path) finds all proteins  in the PDB that can be structurally aligned with the query structure  to within user-specified geometric criteria. CE can optimally align and  display two user-selected structures.   3. Pfam (for Protein families) is a database of nearly 11,000 multiple se¬  quence alignments of protein domains (called Pfam families). Using  Pfam, one can analyze a protein for Pfam matches (74% of proteins  have at least one match in Pfam), determine the domain organization  of a protein based on its sequence or its structure, examine the phylo¬  genetic tree of a Pfam family, and view the occurrence of a proteins  domains across different species.   4. SCOP (Structural Classification Of Proteins) classifies protein struc¬  tures based mainly on manually generated topological considerations  according to a six-level hierarchy: Class (e.g., all-a, all-(3, a/(3), Fold  (based on the arrangement of secondary structural elements),  Superfamily (indicative of distant evolutionary relationships based on  structural criteria and functional features), Family (indicative of near    evolutionary relationships based on sequence as well as on structure),  Protein, and Species. SCOP permits the user to navigate through its  treelike hierarchical organization and lists the known members of any  particular branch.   5. VAST (Vector Alignment Search Tool), a component of the National  Center for Biotechnology Information (NCBI) Entrez system, reports a  precomputed list of proteins of known structure that structurally resem¬  ble the query protein (“structure neighbors”). The VAST system uses the  Molecular Modeling Database (MMDB), an NCBI-generated database  that is derived from PDB coordinates but in which molecules are rep¬  resented by connectivity graphs rather than sets of atomic coordinates.  VAST displays the superposition of the query protein in its structural  alignment with up to five other proteins using the molecular graphics  program Cn3D. VAST also reports a precomputed list of proteins that  are similar to the query protein in sequence (“sequence neighbors”).    Most proteins, particularly those with molecular masses >100 kD, consist of  more than one polypeptide chain. These polypeptide subunits associate with  a specific geometry. The spatial arrangement of these subunits is known as a  proteins quaternary structure.   There are several reasons why multisubunit proteins are so common. In  large assemblies of proteins, such as collagen fibrils, the advantages of subunit  construction over the synthesis of one huge polypeptide chain are analogous  to those of using prefabricated components in constructing a building: Defects  can be repaired by simply replacing the flawed subunit; the site of subunit  manufacture can be different from the site of assembly into the final product;  and the only genetic information necessary to specify the entire edifice is the  information specifying its few different self-assembling subunits. In the case  of enzymes, increasing a proteins size tends to better fix the three-dimensional  positions of its reacting groups. Increasing the size of an enzyme through the  association of identical subunits is more efficient than increasing the length of its  polypeptide chain since each subunit has an active site. More importantly ; the sub¬  unit construction of many enzymes provides the structural basis for the regulation  of their activities (Sections 7-ID and 12-3).    Subunits Usually Associate Noncovalently. A multisubunit protein may con¬  sist of identical or nonidentical polypeptide chains. Hemoglobin, for example,  has the subunit composition a 2 (3 2 (Fig. 6-33). Proteins with more than one  subunit are called oligomers, and their identical units are called protomers. A  protomer may therefore consist of one polypeptide chain or several unlike  polypeptide chains. In this sense, hemoglobin is a dimer of a(3 protomers.   The contact regions between subunits resemble the interior of a single-subunit  protein: They contain closely packed nonpolar side chains, hydrogen bonds in¬  volving the polypeptide backbones and their side chains, and, in some cases,  interchain disulfide bonds. However, the subunit interfaces of proteins that  dissociate in vivo have lesser hydrophobicities than do permanent interfaces.    Subunits Are Symmetrically Arranged. In the vast majority of oligomeric  proteins, the protomers are symmetrically arranged; that is, each protomer oc¬  cupies a geometrically equivalent position in the oligomer. Proteins cannot   • List some of the relative advantages and  disadvantages of using X-ray crystallogra¬  phy and NMR spectroscopy to determine  the structure of a protein.   • Explain why knowing a protein’s amino  acid sequence is required to determine its  tertiary structure.   • Why do turns and loops most often occur  on the protein surface?   • Cover up the bottom portions of Figs. 6-29  and 6-30 and sketch a connectivity dia¬  gram for the secondary structures in each  protein.   • Why does a protein domain consist of at  least 40 amino acid residues?   • Explain why a protein’s sequence evolves  faster than its structure.   • Summarize the types of information pro¬  vided in a PDB file.   • Why is it useful to compare protein struc¬  tures in addition to protein sequences?   FIG. 6-33 Quaternary structure of  hemoglobin. In this space-filling model, the a 1(  a 2 , pi, and p 2 subunits are colored yellow,  green, cyan, and blue, respectively. Heme groups  are red. [Based on an X-ray structure by Max  Perutz, MRC Laboratory of Molecular Biology,  Cambridge, U.K. PDBid 2DHB.]    FIG. 6-34 Symmetries of oligomeric  proteins. The oval, the triangle, the square, and  the pentagon at the ends of the dashed green  lines indicate, respectively, the unique twofold,  threefold, fourfold, and fivefold rotational axes of  the objects shown, (a) Assemblies with cyclic  symmetry (b) Assemblies with dihedral symmetry  In these objects, a twofold axis is perpendicular  to another rotational axis, (c) Assemblies with  the rotational symmetries of a tetrahedron, a  cube or octahedron, and an icosahedron.  [Illustration, Irving Geis. Image from the Irving  Geis Collection/Howard Hughes Medical  Institute. Rights owned by HHMI. Reproduction  by permission only.] ^ See the Animated  Figures.    have inversion or mirror symmetry, however, because bringing the protomers  into coincidence would require converting chiral L residues to D residues.  Thus, proteins can have only rotational symmetry .   In the simplest type of rotational symmetry, cyclic symmetry, protomers  are related by a single axis of rotation (Fig. 6-34#). Objects with 2-, 3-, or  ^-fold rotational axes are said to have C 2 , C 3 , or C n symmetry, respectively.  C 2 symmetry is the most common; higher cyclic symmetries are relatively rare.   Dihedral symmetry (DJ, a more complicated type of rotational symme¬  try, is generated when an ^-fold rotation axis intersects a 2-fold rotation axis  at right angles (Fig. 6-34 £). An oligomer with D n symmetry consists of 2 n  protomers. D 2 symmetry is the most common type of dihedral symmetry in  proteins.   Other possible types of rotational symmetry are those of a tetrahedron,  cube, and icosahedron (Fig. 6-34 e). Some multienzyme complexes and spher¬  ical viruses (see page 127) are built on these geometric plans.    • Protein stability depends primarily on hydrophobic effects and secondarily on  electrostatic interactions.    Incredible as it may seem, thermodynamic measurements indicate that native  proteins are only marginally stable under physiological conditions. The free energy  required to denature them is ~0.4 kj • mol 1 per amino acid residue, so  a fully folded 100-residue protein is only about 40 kj • mol 1 more stable than    its unfolded form (for comparison, the energy required to break a typical hy¬  drogen bond is ~20 kj • mol X ). The various noncovalent influences on  proteins—hydrophobic effects, electrostatic interactions, and hydrogen  bonding—each have energies that may total thousands of kilojoules per mole  over an entire protein molecule. Consequently, a protein structure is the  result of a delicate balance among powerful countervailing forces.   Protein structures are governed primarily by hydrophobic effects and, to a lesser  extent, by interactions between polar residues, and by other types of associations.   The hydrophobic effect , which causes nonpolar substances to minimize their con¬  tacts with water (Section 2-1C), is the major determinant of native protein struc¬  ture. The aggregation of nonpolar side chains in the interior of a protein is  favored by the increase in entropy of the water molecules that would other¬  wise form ordered “cages” around the hydrophobic groups. The combined hy¬  drophobic and hydrophilic tendencies of individual amino acid residues in  proteins can be expressed as hydropathies (Table 6-3). The greater a side  chains hydropathy, the more likely it is to occupy the interior of a protein  and vice versa. Hydropathies are good predictors of which portions of a  polypeptide chain are inside a protein, out of contact with the aqueous sol¬  vent, and which portions are outside (Fig. 6-35).   Site-directed mutagenesis experiments in which individual interior residues  have been replaced by a number of others suggest that the factors that affect  stability are, in order, the hydrophobicity of the substituted residue, its steric  compatibility, and, last, the volume of its side chain.   Electrostatic Interactions Contribute to Protein Stability. In the closely  packed interiors of native proteins, van der Waals forces, which are relatively  weak (Section 2-1 A), are nevertheless an important stabilizing influence. This  is because these forces only act over short distances and hence are lost when  the protein is unfolded.   Perhaps surprisingly, hydrogen bonds , which are central features of protein  structures , make only minor contributions to protein stability. This is because  hydrogen-bonding groups in an unfolded protein form hydrogen bonds with  water molecules. Thus the contribution of a hydrogen bond to the stability of  a native protein is the small difference in hydrogen bonding free energies  between the native and unfolded states ( — 2 to 8 kj • mol 1 as determined by  site-directed mutagenesis studies). Nevertheless, hydrogen bonds are important  determinants of native protein structures, because if a protein folded in a way  that prevented a hydrogen bond from forming, the stabilizing energy of that  hydrogen bond would be lost. Hydrogen bonding therefore fine-tunes tertiary  structure by “selecting” the unique native structure of a protein from among  a relatively small number of hydrophobically stabilized conformations.    Source: Kyte, J. and Doolittle, R.F., J. Mol. Biol.  157, 110 ( 1982 ).    FIG. 6-35 A hydropathic index plot for  bovine chymotrypsinogen. The sum of the   hydropathies of nine consecutive residues is  plotted versus residue sequence number. A  large positive hydropathic index indicates a  hydrophobic region of the polypeptide, whereas  a large negative value indicates a hydrophilic  region. The upper bars denote the protein’s  interior regions, as determined by X-ray  crystallography, and the lower bars denote the  protein’s exterior regions. [After Kyte, J. and  Doolittle, R.F., J. Mol. Biol. 157, 111 (1982).]    0 20 40 60 80 100 120 140 160 180 200 220 240    FIG. 6-36 Examples of ion pairs in  myoglobin. In each case, oppositely charged  side chain groups from residues far apart in  sequence closely approach each other through  the formation of ion pairs.    The association of two ionic protein groups of opposite charge (e.g., Lys  and Asp) is known as an ion pair or salt bridge. About 75% of the charged  residues in proteins are members of ion pairs that are located mostly on the  protein surface (Fig. 6-36). Despite the strong electrostatic attraction between  the oppositely charged members of an ion pair, these interactions contribute  little to the stability of a native protein. This is because the free energy of an  ion pairs charge-charge interactions usually fails to compensate for the loss of  entropy of the side chains and the loss of solvation free energy when the  charged groups form an ion pair. This accounts for the observation that ion  pairs are poorly conserved among homologous proteins.   Disulfide Bonds Cross-Link Extracellular Proteins. Disulfide bonds  (Fig. 4-6) within and between polypeptide chains form as a protein folds to  its native conformation. Some polypeptides whose Cys residues have been  derivatized or mutagenically replaced to prevent disulfide bond formation can  still assume their fully active conformations, suggesting that disulfide bonds  are not essential stabilizing forces. They may, however, be important for “lock¬  ing in” a particular backbone folding pattern as the protein proceeds from its  fully extended state to its mature form.   Disulfide bonds are rare in intracellular proteins because the cytoplasm is  a reducing environment. Most disulfide bonds occur in proteins that are se¬  creted from the cell into the more oxidizing extracellular environment. The  relatively hostile extracellular world (e.g., uncontrolled temperature and pH)  apparently requires the additional structural constraints conferred by disulfide  bonds.    Metal Ions Stabilize Some Small Domains. Metal ions may also function  to internally cross-link proteins. For example, at least ten motifs collectively  known as zinc fingers have been described in nucleic acid-binding proteins.  These structures contain about 25-60 residues arranged around one or two  Zn 2+ ions that are tetrahedrally coordinated by the side chains of Cys, His,  and occasionally Asp or Glu (Fig. 6-37). The Zn 2+ ion allows relatively short  stretches of polypeptide chain to fold into stable units that can interact with  nucleic acids. Zinc fingers are too small to be stable in the absence of Zn 2+ .  Zinc is ideally suited to its structural role in intracellular proteins: Its filled d  electron shell permits it to interact strongly with a variety of ligands (e.g.,  sulfur, nitrogen, or oxygen) from different amino acid residues. In addition,  zinc has only one stable oxidation state (unlike, for example, copper and iron),  so it does not undergo oxidation-reduction reactions in the cell.    FIG. 6-37 X-Ray structure of a zinc finger  motif of the DNA-binding protein Zif268. The   polypeptide chain is drawn in ribbon form and  colored in rainbow order from its N-terminus  (blue) to its C-terminus (red). The side chains of  the Cys and His residues that tetrahedrally  ligand a Zn 2+ ion (cyan sphere) are drawn in  stick form with C green, N blue, and S yellow.  [Based on an X-ray structure by Carl Pabo, MIT  PDBid 1ZAAJ   The low conformational stabilities of native proteins make them easily suscep¬  tible to denaturation by altering the balance of the weak nonbonding forces  that maintain the native conformation. Proteins can be denatured by a vari¬  ety of conditions and substances:   1. Heating causes a proteins conformationally sensitive properties, such  as optical rotation (Section 4-2), viscosity, and UV absorption, to  change abruptly over a narrow temperature range. Such a sharp tran¬  sition indicates that the entire polypeptide unfolds or “melts” cooper¬  atively, that is, nearly simultaneously. Most proteins have melting  temperatures that are well below 100°C. Among the exceptions are the  proteins of thermophilic bacteria (Box 6-3).   2. pH variations alter the ionization states of amino acid side chains,  thereby changing protein charge distributions and hydrogen-bonding  requirements.   3. Detergents associate with the nonpolar residues of a protein, thereby  interfering with the hydrophobic interactions responsible for the pro¬  teins native structure.   in concentrations in the range 5 to 10 M, are the most commonly used  protein denaturants. Chaotropic agents are ions or small organic mol¬  ecules that increase the solubility of nonpolar substances in water. Their    Certain species of bacteria known as hyperthermophiles grow at  temperatures near 100°C. They live in such places as hot springs  and submarine hydrothermal vents, with the most extreme, the  archaebacterium Pyrolobus fumarii, able to grow at temperatures  as high as 1 13°C. These organisms have many of the same  metabolic pathways as do mesophiles (organisms that grow at  “normal” temperatures). Yet most mesophilic proteins denature at  temperatures where hyperthermophiles thrive. What is the struc¬  tural basis for the thermostability of hyperthermophilic proteins?   The difference in the thermal stabilities of the corresponding  (hyper)thermophilic and mesophilic proteins does not exceed  — 100 kJ • mol -1 , the equivalent of a few noncovalent interac¬  tions. This is probably why comparisons of the X-ray structures of  hyperthermophilic enzymes with their mesophilic counterparts have  failed to reveal any striking differences between them. These  proteins exhibit some variations in secondary structure but no  more than would be expected for homologous proteins from dis¬  tantly related mesophiles. However, several of these thermostable  enzymes have a superabundance of salt bridges on their surfaces,  many of which are arranged in extensive networks containing up  to 18 side chains.   The idea that salt bridges can stabilize a protein structure  appears to contradict the conclusion of Section 6-4A that ion  pairs are, at best, marginally stable. The key to this apparent para¬    dox is that the salt bridges in thermostable proteins form networks.  Thus, the gain in charge-charge free energy on associating a  third charged group with an ion pair is comparable to that  between the members of this ion pair, whereas the free energy  lost on desolvating and immobilizing the third side chain is only  about half that lost in bringing together the first two side chains.  The same, of course, is true for the addition of a fourth, fifth, etc.,  side chain to a salt bridge network.   Not all thermostable proteins have such a high incidence of  salt bridges. Structural comparisons suggest that these proteins  are stabilized by a combination of small effects, the most impor¬  tant of which are an increased size of the protein’s hydrophobic  core, an increased size of the interface between its domains  and/or subunits, and a more tightly packed core as evidenced by  a reduced surface-to-volume ratio.   The fact that the proteins of hyperthermophiles and mesophiles  are homologous and carry out much the same functions indicates  that mesophilic proteins are by no means maximally stable. This, in  turn, strongly suggests that the marginal stability of most proteins  under physiological conditions (averaging — 0.4 kJ • mor 1 of amino  acid residues) is an essential property that has arisen through natu¬  ral selection. Perhaps this marginal stability helps confer the struc¬  tural flexibility that many proteins require to carry out their  physiological functions.   Chapter 6 Proteins: Three-Dimensional Structure effectiveness as denaturants stems from their ability to disrupt   hydrophobic interactions, although their mechanism of action is not  well understood.   Many Denatured Proteins Can Be Renatured. In 1957, the elegant exper¬  iments of Christian Anfinsen on ribonuclease A (RNase A) showed that pro¬  teins can be denatured reversibly. RNase A, a 124-residue single-chain  protein, is completely unfolded and its four disulfide bonds reductively  cleaved in an 8 M urea solution containing 2-mercaptoethanol. Dialyzing  away the urea and reductant and exposing the resulting solution to 0 2 at  pH 8 (which oxidizes the SH groups to form disulfides) yields a protein that is  virtually 100% enzymatically active and physically indistinguishable from na¬  tive RNase A (Fig. 6-38). The protein must therefore renature spontaneously.   The renaturation of RNase A demands that its four disulfide bonds re¬  form. The probability of one of the eight Cys residues randomly forming a  disulfide bond with its proper mate among the other seven Cys residues is 1/7;   8 M urea denatures the  protein, and mercaptoethanol  cleaves its disulfide bonds.   Removal of only the  mercaptoethanol allows the  disulfide bonds to reform in'  the denatured protein.    Subsequent removal of the urea  generates an enyzmatically inactive  protein in which the disulfide bonds  have formed at random.   Removal of the denaturant (urea) and  x reductant (mercaptoethanol) allows the  ^'protein to renature and re-form disulfide  bonds in the presence of oxygen.    .Adding a small amount of mercaptoethanol to  5 The scrambled protein in the absence of 0 2  catalyzes its conversion to the active enzyme  through disulfide interchange reactions that  allow the native disulfide bonds to form.    FIG. 6-38 Denaturation and renaturation of RNase A. The polypeptide is represented by a blue line, with its disulfide bonds in yellow.   that of one of the remaining six Cys residues then randomly forming its proper  disulfide bond is 1/5; etc. Thus the overall probability of RNase A re-forming  its four native disulfide links at random is    Clearly, the disulfide bonds do not randomly re-form under renaturing con¬  ditions, since, if they did, only 1% of the refolded protein would be catalyt-  ically active. Indeed, if the RNase A is reoxidized in 8 M urea so that its  disulfide bonds re-form while the polypeptide chain is a random coil, then  after removal of the urea, the RNase A is, as expected, only ~1% active  (Fig. 6-38, Steps 3-4). This “scrambled” protein can be made fully active by  exposing it to a trace of 2-mercaptoethanol, which breaks the improper disul¬  fide bonds and allows the proper bonds to form. Anfinsens work demonstrated  that proteins can fold spontaneously into their native conformations under physi¬  ological conditions. This implies that a proteins primary structure dictates its three-  dimensional structure.   The static way that protein structures are usually portrayed may leave the false  impression that proteins have fixed and rigid structures. In fact, proteins are  flexible and rapidly fluctuating molecules whose structural mobilities are func¬  tionally significant. Groups ranging in size from individual side chains to en¬  tire domains or subunits may be displaced by up to several angstroms through  random intramolecular movements or in response to a trigger such as the bind¬  ing of a small molecule. Extended side chains, such as Lys, and the N- and  C-termini of polypeptide chains are especially prone to wave around in solu¬  tion because there are few forces holding them in place.   Theoretical calculations by Martin Karplus indicate that a proteins native  structure probably consists of a large collection of rapidly interconverting con¬  formations that have essentially equal stabilities (Fig. 6-39). Conformational  flexibility, or breathing, with structural displacement of up to ~2 A, allows  small molecules to diffuse in and out of the interior of certain proteins. In  some cases, a proteins conformational flexibility includes two stable alterna¬  tives in dynamic equilibrium. A change in cellular conditions, such as pH or  oxidation state, or the presence of a binding partner can tip the balance  toward one conformation or the other.    Some Proteins Contain Unfolded Regions. An entire protein or a long  polypeptide segment (>30 residues) may lack defined structure in its native  state. Such intrinsically disordered proteins are characterized by sequences  rich in certain polar and charged amino acids (Gin, Ser, Pro, Glu, Lys, Gly,  and Ala) and lacking in bulky hydrophobic groups (Val, Leu, lie, Met, Phe,  Trp, and Tyr). Sequence analysis suggests that approximately 33% of eukary¬  otic proteins may contain long disordered segments, whereas only a few per¬  cent of prokaryotic proteins do.   Most intrinsically disordered proteins adopt a specific secondary or terti¬  ary structure when they bind to some other molecule. For example, the tran¬  scription factor known as CREB (cyclic AMP response element—binding  protein) is disordered when free in solution but folds to an ordered confor-    FIG. 6-39 Molecular dynamics of myoglobin. Several “snapshots” of the protein  calculated at intervals of 5 X 10 -12 s are superimposed. The backbone is blue, the heme  group is yellow, and the His side chain linking the heme to the protein is orange. [Courtesy  of Martin Karplus, Harvard University.]   backbone of the kinase-inducible domain (KID) of CREB is drawn as a  pink worm. It is unstructured when free in solution (left) but becomes two  perpendicular helices when it forms a complex with the KID-binding  domain of another protein (shown with its solvent-accessible surface in  gray, right). The KID residues Ser 133, which has a covalently attached  phosphoryl group, and Leu 141 are drawn in ball-and-stick form with C  green, 0 red, and P yellow. [Courtesy of Peter Wright, Scripps Research  Institute, La Jolla, California. PDBid 1 KDX.]   • Describe the hydropathic index plot for a  fibrous protein such as collagen or keratin.   • Describe the forces that stabilize proteins,  and rank their relative importance.   • Why would it be advantageous for a pro¬  tein or a segment of a protein to lack  defined secondary or tertiary structure?    mation when it interacts with the CREB-binding protein (Fig. 6-40).  Apparently, the increased flexibility of disordered protein segments enables  them to perform a relatively unhindered conformational search when binding  to their target molecules. It has also been suggested that a structured globular  protein would have to be two to three times larger than a disordered protein  to provide the same size intermolecular interface, and hence the use of disor¬  dered proteins provides genetic economy and reduces intracellular crowding.    • A folding protein follows a pathway from high energy and high entropy to low  energy and low entropy.   • A variety of molecular chaperones assist protein folding via an ATP-dependent  bind-and-release mechanism.    Studies of protein stability and renaturation suggest that protein folding is di¬  rected largely by the residues that occupy the interior of the folded protein.  But how does a protein fold to its native conformation? One might guess that  this process occurs through the proteins random exploration of all the con¬  formations available to it until it eventually stumbles onto the correct one. A  simple calculation first made by Cyrus Levinthal, however, convincingly  demonstrates that this cannot possibly be the case: Assume that an ^-residue  proteins 2" torsion angles, 4> and i]each have three stable conformations.  This yields 3 2 " ~ 10" possible conformations for the protein (a gross under¬  estimate because we have completely neglected its side chains). Then, if the  protein could explore a new conformation every 10 13 s (the rate at which  single bonds reorient), the time t , in seconds, required for the protein to ex¬  plore all the conformations available to it is   For a small protein of 100 residues, t — 10 87 s, which is immensely greater  than the apparent age of the universe (—13-7 billion years = 4.3 X 10 17 s).  Clearly, proteins must fold more rapidly than this.   Experiments have shown that many proteins fold to their native conformations  in less than a few seconds. This is because proteins fold to their native confor¬  mations via directed pathways rather than stumbling on them through random   FIG. 6-41 Hypothetical protein folding pathway. This example shows a linear  pathway for folding a two-domain protein. [After Goldberg, M.E., Trends Biochem. Sci. 10,  389 (1985).]    conformational searches. Thus, as a protein folds, its conformational stability in¬  creases sharply (i.e., its free energy decreases sharply), which makes folding a  one-way process. A hypothetical folding pathway is diagrammed in Fig. 6-41.   Experimental observations indicate that protein folding begins with the for¬  mation of local segments of secondary structure (a helices and (3 sheets). This  early stage of protein folding is extremely rapid, with much of the native sec¬  ondary structure in small proteins appearing within 5 ms of the initiation of  folding. Since native proteins contain compact hydrophobic cores, it is likely  that the driving force in protein folding is what has been termed a hydropho¬  bic collapse. The collapsed state is known as a molten globule, a species that  has much of the secondary structure of the native protein but little of its ter¬  tiary structure. Theoretical studies suggest that helices and sheets form in part  because they are particularly compact ways of folding a polypeptide chain.   Over the next 5 to 1000 ms, the secondary structure becomes stabilized  and tertiary structure begins to form. During this intermediate stage, the na¬  tivelike elements are thought to take the form of subdomains that are not yet  properly docked to form domains. In the final stage of folding, which for  small, single-domain proteins occurs over the next few seconds, the protein  undergoes a series of complex rearrangements in which it attains its relatively  stable internal side chain packing and hydrogen bonding while it expels the  remaining water molecules from its hydrophobic core.   In multidomain and multisubunit proteins, the respective units then as¬  semble in a similar manner, with a few slight conformational adjustments re¬  quired to produce the proteins native tertiary or quaternary structure. Thus,  proteins appear to fold in a hierarchical manner, with small local elements of struc¬  ture forming and then coalescing to yield larger elements, which coalesce with other  such elements to form yet larger elements, etc.   Folding, like denaturation, appears to be a cooperative process, with small  elements of structure accelerating the formation of additional structures. A  folding protein must proceed from a high-energy, high-entropy state to a low-  energy, low-entropy state. This energy-entropy relationship, which is dia¬  grammed in Fig. 6-42, is known as a folding funnel. An unfolded  polypeptide has many possible conformations (high entropy). As it folds into  an ever-decreasing number of possible conformations, its entropy and free en¬  ergy decrease. The energy-entropy diagram is not a smooth valley but a jagged    FIG. 6-42 Energy-entropy diagram for protein folding. The width of the diagram  represents entropy, and the depth, the energy. The unfolded polypeptide proceeds from a  high-entropy, disordered state (wide) to a single low-entropy (narrow), low-energy native  conformation. [After Onuchic, J.N., Wolynes, P.G., Luthey-Schulten, Z., and Socci, N.D.,  Proc. Natl. Acad Sci. 92, 3626 (1995).]   Chapter 6 Proteins: Three-Dimensional Structure landscape. Minor clefts and gullies represent conformations that are temporar¬  ily trapped until, through random thermal activation, they overcome a slight  “uphill” free energy barrier and can then proceed to a lower energy conforma¬  tion. Evidently, proteins have evolved to have efficient folding pathways as well  as stable native conformations.   Understanding the process of protein folding as well as the forces that sta¬  bilize folded proteins is essential for elucidating the rules that govern the re¬  lationship between a proteins amino acid sequence and its three-dimensional  structure. Such information will prove useful in predicting the structures of  the millions of proteins that are known only from their sequences (Box 6-4).   Protein Disulfide Isomerase Acts during Protein Folding. Even under op¬  timal experimental conditions, proteins often fold more slowly in vitro than  they fold in vivo. One reason is that folding proteins often form disulfide  bonds not present in the native proteins, and then slowly form native disul¬  fide bonds through the process of disulfide interchange. Protein disulfide iso¬  merase (PDI) catalyzes this process. Indeed, the observation that RNase A  folds so much faster in vivo than in vitro led Anfinsen to discover this enzyme.   PDI binds to a wide variety of unfolded polypeptides via a hydrophobic  patch on its surface. A Cys—SH group on reduced (SH-containing) PDI re¬  acts with a disulfide group on the polypeptide to form a mixed disulfide and  a Cys—SH group on the polypeptide (Fig. 6-43 d). Another disulfide group   (a) Reduced (SH-containing) PDI catalyzes the rearrangement of a  polypeptide’s non-native disulfide bonds via disulfide interchange  reactions to yield native disulfide bonds, (b) Oxidized (disulfide-   containing) PDI catalyzes the initial formation of a polypeptide’s  disulfide bonds through the formation of a mixed disulfide. Reduced  PDI can then react with a cellular oxidizing agent to regenerate  oxidized PDI. Q See the Animated Figures.    Around 7 million polypeptide sequences are known, yet the struc¬  tures of only —80,000 proteins have been determined. Consequently,  there is a need to develop robust techniques for predicting a  protein’s structure from its amino acid sequence. This represents  a formidable challenge but promises great rewards in terms of  understanding protein function, identifying diseases related to  abnormal protein sequences, and designing drugs to alter protein  structure or function.   There are several major approaches to protein structure predic¬  tion. The simplest and most reliable approach, homology modeling,  aligns the sequence of interest with the sequence of a homolo¬  gous protein or domain of known structure—compensating for  amino acid substitutions, insertions, and deletions—through model¬  ing and energy minimization calculations. This method yields reli¬  able models for proteins that have as little as 25% sequence  identity with a protein of known structure, although, of course, the  accuracy of the model increases with the degree of sequence  identity. The emerging field of structural genomics, which seeks  to determine the X-ray structures of all representative domains, is  aimed at expanding this predictive technique. The identification of  structural homology is likely to provide clues as to a protein’s  function even with imperfect structure prediction.   Distantly related proteins may be structurally similar even  though they have diverged to such an extent that their sequences  show no obvious resemblance. Threading is a computational  technique that attempts to determine the unknown structure of a  protein by ascertaining whether it is consistent with a known  protein structure. It does so by placing (threading) the unknown  protein’s residues along the backbone of a known protein struc¬  ture and then determining whether the amino acid side chains of  the unknown protein are stable in that arrangement. This method  is not yet reliable, although it has yielded encouraging results.   Empirical methods based on experimentally determined statisti¬  cal information such as the a helix and p sheet propensities de¬  duced by Chou and Fasman (Table 6-1) have been moderately  successful in predicting the secondary structures of proteins. Their  main drawback is that neighboring residues in a polypeptide  sometimes exert strong influence on a given residue’s tendency to  form a particular secondary structure.   Since the native structure of a protein ultimately depends on  its amino acid sequence, it should be possible, in principle, to  predict the structure of a protein based only on its chemical and  physical properties (e.g., the hydrophobicity, size, hydrogen-bonding  propensity, and charge of each of its amino acid residues). The  most successful of these ab initio (from the beginning) methods  is the Rosetta program formulated by David Baker. To satisfy the  program’s computational needs, a volunteer network of —100,000  computers, known as Rosetta@home, provides the 500,000 or so  hours of processing time required to generate a structure. Two  examples of successful protein structure prediction by Rosetta  appear above, marked (a) and (fa). The predicted model (gray) for  each of these bacterial proteins is superimposed on the experi¬  mentally determined X-ray structure, colored in rainbow order from  N-terminus (blue) to C-terminus (red) with core side chains drawn  as sticks.   Protein design, the experimental inverse of protein structure  prediction, has provided insights into protein folding and stability.  Protein design attempts to construct an amino acid sequence that  will form a structure such as a sandwich of p sheets or a bundle   of a helices. The designed polypeptide is then chemically or bio¬  logically synthesized, and its structure is determined. Experimental  results suggest that the greatest challenge of protein design may  lie not in getting the polypeptide to fold to the desired conforma¬  tion but in preventing it from folding into other unwanted confor¬  mations. In this respect, science lags far behind nature.   The first wholly successful de novo (beginning anew) protein  design, accomplished by Stephen Mayo, was for a 28-residue ppa  motif that has a backbone conformation designed to resemble a  zinc finger (Fig. 6-37) but that contains no stabilizing metal ions.   A computational design process considered the interactions  among side chain and backbone atoms, screened all possible  amino acid sequences, and, in order to take into account side  chain flexibility, tested all sets of energetically allowed torsion  angles for each side chain. The number of amino acid sequences  to be tested was limited to 1.9 X 10 27 , representing 1.1 X 10 62  possible conformations! The design process yielded an optimal  sequence of 28 residues, which was chemically synthesized and  its structure determined by NMR spectroscopy. The designed  protein, called FSD-1, closely resembled its predicted structure,  and its backbone conformation (blue) was nearly superimposable  on that of a known zinc finger motif (red). Although FSD-1 is  relatively small, it folds into a unique stable structure, thereby  demonstrating the power of protein design techniques.   [Figures courtesy of Gautam Dantas, Washington University School of  Medicine; PDBids 1WHZ and 2HH6; and Stephen Mayo, California  Institute of Technology.]   Chapter 6 Proteins: Three-Dimensional Structure Q n the polypeptide, brought into proximity by the spontaneous folding of the   polypeptide, is attacked by this Cys—SH group. The newly liberated Cys—SH  group then repeats this process with another disulfide bond, and so on, ulti¬  mately yielding the polypeptide containing only native disulfide bonds, along  with regenerated PDI.   Oxidized (disulfide-containing) PDI also catalyzes the initial formation of  a polypeptides disulfide bonds by a similar mechanism (Fig. 6-43£). In this  case, the reduced PDI reaction product must be reoxidized by cellular oxidiz¬  ing agents in order to repeat the process.   Proteins begin to fold as they are being synthesized, so the renaturation of a  denatured protein in vitro may not entirely mimic the folding of a protein  in vivo. In addition, proteins fold in vivo in the presence of extremely high  concentrations of other proteins with which they can potentially interact.  Molecular chaperones are essential proteins that bind to unfolded and partially  folded polypeptide chains to prevent the improper association of exposed hydropho¬  bic segments that might lead to nonnative folding as well as polypeptide aggrega¬  tion and precipitation. This is especially important for multidomain and  multisubunit proteins, whose components must fold fully before they can  properly associate with each other. Molecular chaperones also induce misfolded  proteins to refold to their native conformations.   Many molecular chaperones were first described as heat shock proteins  (Hsp) because their rate of synthesis is increased at elevated temperatures.  Presumably, the additional chaperones are required to recover heat-denatured  proteins or to prevent misfolding under conditions of environmental stress.   Most Chaperones Require ATP. There are several classes of molecular chap¬  erones in both prokaryotes and eukaryotes, including the following:   1. The Hsp70 family of proteins are highly conserved 70-kD proteins in  both prokaryotes and eukaryotes. In association with the cochaperone  protein Hsp40, they facilitate the folding of newly synthesized pro¬  teins and reverse the denaturation and aggregation of proteins. Hsp70  proteins also function to unfold proteins in preparation for their trans¬  port through membranes (Section 9-4D) and to subsequently refold  them.   2. Trigger factor is a ribosome-associated chaperone in prokaryotes that  prevents the aggregation of polypeptides as they emerge from the ri¬  bosome (Section 27-5A). Trigger factor and Hsp70 are the first chap¬  erones a newly made prokaryotic protein encounters. Subsequently,  many partially folded proteins are handed off to other chaperones to  complete the folding process. Eukaryotes lack trigger factor but con¬  tain other small chaperones that have similar functions.   3. The chaperonins form large, multisubunit, cagelike assemblies in both  prokaryotes and eukaryotes. They bind improperly folded proteins and  induce them to refold inside an internal cavity (see below).   4. The Hsp90 proteins are eukaryotic proteins that mainly facilitate the  late stages of folding of proteins involved in cellular signaling  (Chapter 13). Hsp90 proteins are among the most abundant proteins  in eukaryotes, accounting for up to 6% of cellular protein under stress¬  ful conditions that destabilize proteins.   All of these molecular chaperones operate by binding to an unfolded or ag¬  gregated polypeptides solvent-exposed hydrophobic surface and subsequently  releasing it, often repeatedly, in a manner that facilitates its proper folding.  Most molecular chaperones are ATPases, that is, enzymes that catalyze the    hydrolysis of ATP (adenosine triphosphate) to ADP (adenosine diphosphate) Section 5 Protein Folding   The favorable free energy change of ATP hydrolysis drives the chaperones  bind-and-release reaction cycle.   The GroEL/ES Chaperonin Forms Closed Chambers in Which Proteins  Fold. The chaperonins in E. coli consist of two types of subunits named  GroEL and GroES. The X-ray structure of a GroEL-GroES-(ADP) 7 complex  (Fig. 6-44), determined by Arthur Horwich and Paul Sigler, reveals fourteen  identical 549-residue GroEL subunits arranged in two stacked rings of seven  subunits each. This complex is capped at one end by a domelike heptameric  ring of 97-residue GroES subunits to form a bullet-shaped complex with C 7  symmetry. The two GroEL rings each enclose a central chamber with a diam¬  eter of ~45 A in which partially folded proteins fold to their native confor¬  mations. A barrier in the center of the complex (Fig. 6-44 c) prevents a folding  protein from passing between the two GroEL chambers. The GroEL ring that  contacts the GroES heptamer is called the cis ring; the opposing GroEL ring  is known as the trans ring.   ATP Binding and Hydrolysis Drive the Conformational Changes in  GroEL/ES. Each GroEL subunit has a binding pocket for ATP that catalyzes  the hydrolysis of its bound ATP to ADP + P z . When the cis ring subunits  hydrolyze their bound ATP molecules and release the product P z , the protein  undergoes a conformational change that widens and elongates the cis inner  cavity so as to more than double its volume from 85,000 A 3 to 175,000 A 3 . (In  the structure shown in Fig. 6-44, the cis ring has already hydrolyzed its seven  molecules of ATP to ADP.) The expanded cavity can enclose a partially folded  substrate protein of at least 70 kD. All seven subunits of the GroEL ring act   FIG. 6-44 X-Ray structure of the GroEL-GroES-(ADP) 7  complex, (a) A space-filling drawing as viewed perpendicularly to the  complex’s sevenfold axis with the GroES ring orange, the cis ring of  GroEL green, and the trans ring of GroEL red with one subunit of  each ring shaded more brightly. The dimensions of the complex are  indicated. Note the different conformations of the two GroEL rings.  The ADPs, whose binding sites are in the base of each cis ring  GroEL subunit, are not seen because they are surrounded by protein.    (b) As in Part a but viewed along the sevenfold axis, (c) As in Part a  but with the two GroEL subunits closest to the viewer in both the cis  and trans rings removed to expose the interior of the complex. The  level of fog increases with the distance from the viewer. Note the  much larger size of the cavity formed by the cis ring and GroES in  comparison to that of the trans ring. [Based on an X-ray structure by  Paul Sigler, Yale University. PDBid 1AON.]   Chapter 6 Proteins: Three-Dimensional Structure { n concert; that is , they are mechanically linked such that they change their   The cis and trans GroEL rings undergo conformadonal changes in a recip¬  rocating fashion, with events in one ring influencing events in the other ring.  The entire GroEL/ES chaperonin complex functions as follows (Fig. 6-45):   1. One GroEL ring that has bound 7 ATP also binds an improperly folded  substrate protein, which associates with hydrophobic patches that line the  inner wall of the GroEL chamber. The GroES cap then binds to the  GroEL ring like a lid on a pot, inducing a conformational change in the  resulting cis ring that buries the hydrophobic patches, thereby depriving  the substrate protein of its binding sites. This releases the substrate pro¬  tein into the now enlarged and closed cavity, where it commences fold¬  ing. The cavity, which is now lined only with hydrophilic groups, provides  the substrate protein with an isolated microenvironment that prevents it  from nonspecifically aggregating with other misfolded proteins. Moreover,  the conformational change that buries GroELs hydrophobic patches  stretches and thereby partially unfolds the improperly folded substrate  protein before it is released. This rescues the substrate protein from a lo¬  cal energy minimum in which it had become trapped (Fig. 6-42), thereby  permitting it to continue its conformational journey down the folding  funnel toward its native state (the state of lowest free energy).   2. Within, —10 s (the time the substrate protein has to fold), the cis ring  catalyzes the hydrolysis of its 7 bound ATPs to ADP + P z and the P z   The trans ring with its  bound ATP now  becomes the cis ring.    GroES cap binds to the GroEL ring. This induces a  conformational change in the now cis ring, which  releases the improperly folded protein into the  enlarged cavity where it commences to refold.    Within ~10 s the cis ring  catalyzes the hydrolysis of its  7 bound ATPs and releases 2  the resulting P,-, which  weakens the interactions  binding GroES to GroEL.    A second molecule of improperly  folded substrate protein binds to  the trans ring, followed by 7 ATP.   FIG. 6-45 Reaction cycle of the GroEL/ES chaperonin. The protein complex is colored as in Fig. 6-44. See the text for an explanation.   3. A second molecule of improperly folded substrate protein binds to the  trans ring followed by 7 ATP. Conformational linkages between the cis  and trans rings prevent the binding of both substrate protein and ATP  to the trans ring until the ATP in the cis ring has been hydrolyzed.   4. The binding of substrate protein and ATP to the trans ring conforma-  tionally induces the cis ring to release its bound GroES, 7 ADP, and  the presumably now better-folded substrate protein. This leaves ATP  and substrate protein bound only to the trans ring of GroEL, which  now becomes the cis ring as it binds GroES.    Steps 1 through 4 are then repeated. The GroEL/ES system expends 7 ATPs  per folding cycle. If the released substrate protein has not achieved its native  state, it may subsequently rebind to GroEL (a substrate protein that has  achieved its native fold lacks exposed hydrophobic groups and hence cannot  rebind to GroEL). Typically, only ~5% of substrate proteins fold to their na¬  tive state in each reaction cycle. Thus, to fold half the substrate protein pres¬  ent would require log(l — 0.5)/log(l — 0.05) ~ 14 reaction cycles and hence  7 X 14 = 98 ATPs (which appears to be a profligate use of ATP but consti¬  tutes only a small fraction of the thousands of ATPs that must be hydrolyzed  to synthesize a typical polypeptide and its component amino acids). Because  protein folding occurs alternately in the two GroEL rings, the proper func¬  tioning of the chaperonin requires both GroEL rings, even though their two  cavities are unconnected.   Experiments indicate that the GroEL/ES system interacts with only a sub¬  set of E. coli proteins, most with molecular masses in the range 20 to 60 kD.  These proteins tend to contain two or more a/(3 domains that mainly consist  of open (3 sheets. Such proteins are expected to fold only slowly to their na¬  tive state because the formation of hydrophobic sheets requires a large num¬  ber of specific long-range interactions. Proteins dissociate from GroEL/ES after  folding, but some frequently revisit the chaperonin, apparently because they  are structurally labile or prone to aggregate and must return to GroEL for pe¬  riodic maintenance.   Eukaryotic cells contain the chaperonin TRiC, with double rings of eight  nonidentical subunits, each of which resembles a GroEL subunit. However,  the TRiC proteins contain an additional segment that acts as a built-in lid, so  the complex encloses a polypeptide chain and mediates protein folding with¬  out the assistance of a GroES-like cochaperone. Like its bacterial counterpart,  TRiC operates in an ATP-dependent fashion. Around 10% of eukaryotic  proteins transiently interact with TRiC.   r Most proteins in the body maintain their native conformations or, if they  become partially denatured, are either renatured through the auspices of  molecular chaperones or are proteolytically degraded (Section 21-1). However,  at least 35 different—and usually fatal—human diseases are associated with  the extracellular deposition of normally soluble proteins in certain tissues in  the form of insoluble fibrous aggregates (Table 6-4). The aggregates are known  as amyloids, a term that means starchlike because it was originally thought  that the material resembled starch.   The diseases known as amyloidoses are a set of relatively rare inherited  diseases in which mutant forms of normally occurring proteins [e.g.,  lysozyme, an enzyme that hydrolyzes bacterial cell walls (Section 11-4), and  fibrinogen, a blood plasma protein that is the precursor of fibrin, which forms   FIG. 6-46 Brain tissue from an individual  with Alzheimer’s disease. The two circular  objects in this photomicrograph are plaques that  consist of amyloid deposits of Ap protein  surrounded by a halo of neurites (axons and  dendrites) from dead and dying neurons.  [Courtesy of Dennis Selkoe and Marcia Podlisny,  Harvard University Medical School.]   FIG. 6-47 Electron micrograph of a  cluster of partially proteolyzed prion rods.   The black dots are colloidal gold beads that are  coupled to anti-PrP antibodies adhering to the  PrP. [Courtesy of Stanley Prusiner, University of  California at San Francisco Medical Center.]    blood clots (Box 11-4)] accumulate in a variety of tissues as amyloids. The  symptoms of amyloidoses usually do not become apparent until the third to  seventh decade of life and typically progress over 5 to 15 years, ending in  death.   Amyloid-p Protein Accumulates in Alzheimer’s Disease. Alzheimer’s dis¬  ease, a neurodegenerative condition that strikes mainly the elderly, causes dev¬  astating mental deterioration and eventual death (it affects —10% of those  over 65 and —50% of those over 85). It is characterized by brain tissue con¬  taining abundant amyloid plaques (deposits) surrounded by dead and dying  neurons (Fig. 6-46). The amyloid plaques consist mainly of fibrils of a 40- to  42-residue protein named amyloid-P protein (AP). Ap is a fragment of a  770-residue membrane protein called the Ap precursor protein (PPP), whose  normal function is unknown. Ap is excised from PPP in a multistep process  through the actions of two proteolytic enzymes dubbed P- and 7 -secretases.  The neurotoxic effects of Ap begin even before significant amyloid deposits  appear (see below).   The age dependence of Alzheimer’s disease suggests that Ap deposition is  an ongoing process. Indeed, several rare mutations in the PPP gene that in¬  crease the rate of Ap production result in the onset of Alzheimer’s disease as  early as the fourth decade of life. A similar phenomenon occurs in individu¬  als with Down’s syndrome, a condition characterized by mental retardation  and a distinctive physical appearance caused by the trisomy (3 copies per cell)  of chromosome 21 rather than the normal two copies. These individuals in¬  variably develop Alzheimer’s disease by their 40th year because the gene  encoding PPP is located on chromosome 21 and hence individuals with  Down’s syndrome produce pPP and presumably Ap at an accelerated rate.  Consequently, a promising strategy for halting the progression of Alzheimer’s  disease is to develop drugs that inhibit the action of the p- and/or y-secretases  so as to decrease the rate of Ap production.   Prion Diseases Are Infectious. Certain diseases that affect the mammalian  central nervous system were originally thought to be caused by “slow viruses”  because they take months, years, or even decades to develop. Among them are  scrapie (a neurological disorder of sheep and goats), bovine spongiform  encephalopathy (BSE or mad cow disease), and kuru (a degenerative brain  disease in humans that was transmitted by ritual cannibalism among the Fore  people of Papua New Guinea; kuru means “trembling”). There is also a  sporadic (spontaneously arising) human disease with similar symptoms,  Creutzfeldt-Jakob disease (CJD), which strikes one person per million per  year and which may be identical to kuru. In all of these invariably fatal dis¬  eases, neurons develop large vacuoles that give brain tissue a spongelike mi¬  croscopic appearance. Hence the diseases are collectively known as  transmissible spongiform encephalopathies (TSEs).   Unlike other infectious diseases, the TSEs are not caused by a virus or mi¬  croorganism. Indeed, extensive investigations have failed to show that they are  associated with any nucleic acid. Instead, as Stanley Prusiner demonstrated for  scrapie, the infectious agent is a protein called a prion (for proteinaceous  infectious particle that lacks nucleic acid) and hence TSEs are alternatively called  prion diseases. The scrapie prion, which is named PrP (for Prion Protein),  consists of 208 mostly hydrophobic residues. This hydrophobicity causes par¬  tially proteolyzed PrP to aggregate as clusters of rodlike particles that closely  resemble the amyloid fibrils seen on electron microscopic examination of  prion-infected brain tissue (Fig. 6-47). These fibrils presumably form the amy¬  loid plaques that accompany the neuronal degeneration in TSEs.   How are prion diseases transmitted? PrP is the product of a normal cel¬  lular gene that has no known function (genetically engineered mice that fail  to express PrP appear to be normal). Infection of cells by prions somehow   form of PrP (PrP Sc ) is identical to normal cellular PrP (PrP C ) in sequence but  differs in secondary and/or tertiary structure. This suggests that PrP Sc induces  PrP C to adopt the conformation ofPrP S \ that is, a small amount of PrP Sc trig¬  gers the formation of additional PrP 5c from PrP , which triggers more PrP 5c  to form, and so on. This accounts for the observation that mice that do not  express the gene encoding PrP cannot be infected with scrapie.   Human PrP C consists of a disordered (and hence unseen) 98-residue  N-terminal “tail” and a 110-residue C-terminal globular domain containing  three a helices and a short two-stranded antiparallel (3 sheet (Fig. 6-48^).   Unfortunately, the insolubility of PrP Sc has precluded its structural determi¬  nation, but spectroscopic methods indicate that it has a lower a helix content  and a higher (3 sheet content than PrP C . This suggests that the protein has re¬  folded (Fig. 6-48/?). The high (3 sheet content of PrP Sc presumably facilitates  the aggregation of PrP Sc as amyloid fibrils (see below).   Prion diseases can be transmitted by the consumption of nerve tissue from  infected individuals, as illustrated by the incidence of BSE. This disease was  unknown before 1985 but reached epidemic proportions among cattle in the  U.K. in 1993. The rise in BSE reflects the practice, beginning in the 1970s,  of feeding cattle preparations of meat and bone meal that were derived from  other animals by a method that failed to inactivate prions. The BSE epidemic  abated due to the banning of such feeding in 1988, together with the slaugh¬  ter of a large number of animals at risk for having BSE. However, it is now  clear that BSE was transmitted to humans who ate meat from BSE-infected  cattle: Some 200 cases of so-called new variant CJD have been reported to  date, almost entirely in the U.K., many of which occurred in teenagers and  young adults. Yet before 1994, CJD under the age of 40 was extremely rare.   It should be noted that the transmission of BSE from cattle to humans was  unexpected: Scrapie-infected sheep have long been consumed worldwide and  yet the incidence of CJD in mainly meat-eating countries such as the U.K.   (in which sheep are particularly abundant) was no greater than that in largely  vegetarian countries such as India.   FIG. 6-48 Prion protein conformations, (a) The NMR structure of human prion  protein (PrP c ). The protein, missing its first 23 residues, is drawn in ribbon form with  helices red, |3 sheets green, and other segments orange. Its disulfide bond is shown in yellow.  (b) A plausible model for the structure of PrP Sc represented as in Part a. Note the  formation of structure in what was the flexibly disordered N-terminal region. [Courtesy of  Fred Cohen, University of California at San Francisco. Part a based on an NMR structure  by Kurt Wuthrich, Eidqenossische Technische Hochschule, Zurich, Switzerland. PDBid  1QLXJ   • Describe the energy and entropy changes  that occur during protein folding.   • Explain why it is important for protein  disulfide isomerase to catalyze both the  breaking and formation of disulfide bonds.   • How does protein renaturation in vitro  differ from protein folding in vivo ?   • Explain why a protein such as RNase A  can be easily denatured and renatured  in vitro , whereas most proteins that are  denatured do not refold property in vitro.   • Explain the role of ATP in the action of  Hsp70 and GroEL/ES.   • Why would cells need more than one type  of chaperone?   • What are amyloid fibrils, what is their  origin, and why are they harmful?   • Explain what must happen in order for a  folded globular protein to form an amyloid  fiber.    FIG. 6-49 Model of an amyloid fibril, (a) The model, based on X-ray fiber diffraction  measurements, is viewed normal to the fibril axis (above) and along the fibril axis (below).  The arrowheads indicate the path but not necessarily the direction of the |3 strands.   (b) A single p sheet, which is shown for clarity. The loop regions connecting the p strands  have unknown structure. [Courtesy of Colin Blake, Oxford University, Oxford, U.K., and  Louise Serpell, University of Cambridge, U.K.]    Amyloid Fibrils Are p Sheet Structures. The amyloid fibers that charac¬  terize the amyloidoses, Alzheimer’s disease, and the TSEs are built from pro¬  teins that exhibit no structural or functional similarities in their native states.  In contrast, the appearance of their fibrillar forms is strikingly similar.  Spectroscopic analysis of amyloid fibrils indicates that they are rich in  P structure, with individual P strands oriented perpendicular to the fiber axis  (Fig. 6-49). Furthermore, the ability to form amyloid fibrils is not unique  to the small set of proteins associated with specific diseases. Under the ap¬  propriate conditions, almost any protein can be induced to aggregate. Thus,  the ability to form amyloid may be an intrinsic property of all polypeptide chains.   A variety of experiments indicate that amyloidogenic mutant proteins are  significantly less stable than their wild-type counterparts (e.g., they have sig¬  nificantly lower melting temperatures). This suggests that the partially un¬  folded, aggregation-prone forms are in equilibrium with the native  conformation even under conditions in which the native state is thermody¬  namically stable [keep in mind that the equilibrium ratio of unfolded (U)  to native (N) protein molecules in the reaction N U is governed by  Eq. 1-17: = [U]/[N] = e AG ^ RT , where A G°' is the standard free energy of   unfolding, so that as A G°' decreases, the equilibrium proportion of U in¬  creases]. It is therefore likely that fibril formation is initiated by the associa¬  tion of the P domains of two or more partially unfolded amyloidogenic  proteins to form a more extensive P sheet. This would provide a template or  nucleus for the recruitment of additional polypeptide chains to form the grow¬  ing fibril. Since most amyloid diseases require several decades to become symp¬  tomatic, the development of an amyloid nucleus must be a rare event. Once  an amyloid fiber begins to grow, however, its development is more rapid.   The factors that trigger amyloid formation remain obscure, even when  mutations (in the case of hereditary amyloidoses) or infection (in the case of  TSEs) appear to be the cause. After it has formed, an amyloid fibril is virtu¬  ally indestructible under physiological conditions, possibly due to the large  number of main-chain hydrogen bonds that must be broken in order to sep¬  arate the individual polypeptide strands (side chain interactions are less im¬  portant in stabilizing P sheets). It seems likely that protein folding pathways  have evolved not only to allow polypeptides to assume stable native structures  but also to avoid forming interchain hydrogen bonds that would lead to fib¬  ril formation.   Are fibrillar deposits directly responsible for the neurodegeneration seen  in many amyloid diseases? A growing body of evidence suggests that cellular  damage begins when the misfolded proteins first aggregate but are still solu¬  ble. For example, in mouse models of Alzheimer s disease, cognitive impair¬  ment is evident before amyloid plaques develop. Other experiments show that  the most infectious prion preparations contain just 14—28 PrP Sc molecules,  that is, a nucleus for a fibril, not the fibril itself. Even a modest number of  misfolded protein molecules could be toxic if they prevented the cells chap¬  erones from assisting other more critical proteins to fold. The appearance of  extracellular—and sometimes intracellular—amyloid fibrils may simply repre¬  sent the accumulation of protein that has overwhelmed the cellular mecha¬  nisms that govern protein folding or the disposal of misfolded proteins.   • Four levels of structural complexity are used to describe the three-  dimensional shapes of proteins.   • The conformational flexibility of the peptide group is described  by its 4> and v|i torsion angles.   • The a helix is a regular secondary structure in which hydrogen  bonds form between backbone groups four residues apart. In the  |3 sheet, hydrogen bonds form between the backbones of separate  polypeptide segments.   • Fibrous proteins are characterized by a single type of secondary  structure: a keratin is a left-handed coil of two a helices, and col¬  lagen is a left-handed triple helix with three residues per turn.   • The structures of proteins have been determined mainly by X-ray  crystallography and NMR spectroscopy.   • The nonpolar side chains of a globular protein tend to occupy the  protein’s interior; the polar side chains tend to define its surface.   • Protein structures can be classified on the basis of motifs, second¬  ary structure content, topology, or domain architecture. Structural  elements are more likely to be evolutionarily conserved than are  amino acid sequences.   • Structural bioinformatics is concerned with the storage, visualiza¬  tion, analysis, and comparison of macromolecular structures.   • The individual subunits of multisubunit proteins are usually sym¬  metrically arranged.   • Native protein structures are only slightly more stable than their  denatured forms. The hydrophobic effect is the primary determi¬  nant of protein stability. Hydrogen bonding and ion pairing con¬  tribute relatively little to a protein’s stability.   • Studies of protein denaturation and renaturation indicate that the  primary structure of a protein determines its three-dimensional  structure.   • Proteins fold to their native conformations via directed pathways  in which small elements of structure coalesce into larger structures.   • Molecular chaperones facilitate protein folding in vivo by repeat¬  edly binding and releasing a polypeptide in an ATP-dependent  manner and providing it with an isolated microenvironment in  which to fold.   • Diseases caused by protein misfolding include the amyloidoses,  Alzheimer’s disease, and the transmissible spongiform en¬  cephalopathies (TSEs).   (Rossmann) fold 150  structural bioinformatics 152  oligomer 155  protomer 155  rotational symmetry 156  cyclic symmetry 156  dihedral symmetry 156  hydropathy 157  ion pair (salt bridge) 158  zinc finger 158  cooperativity 159  chaotropic agent 159  renaturation 160  breathing 161   hydrophobic collapse 163  molten globule 163  homology modeling 165  threading 165  ab initio 165  molecular chaperone 166  heat shock protein 166  chaperonin 166  ATPase 166  amyloid 169  Alzheimer’s disease 170  Creutzfeldt-Jakob disease 170  transmissible spongiform  encephalopathies  (TSEs) 170  prion 170    1. Draw a cis peptide bond and identify the groups that experience  steric interference.   2. How many peptide bonds are shown in the structure drawn in   3. Helices can be described by the notation n mi where n is the num¬  ber of residues per helical turn and m is the number of atoms,    including H, in the ring that is closed by the hydrogen bond,  (a) What is this notation for the a helix? (b) Is the 3io helix  steeper or shallower than the a helix?   4. Why would you be unlikely to see an a helix containing only  the following amino acids: Arg, Lys, Met, Phe, Trp, Tyr, Val?   5. Calculate the length in angstroms of a 100-residue segment of  the a keratin coiled coil.   6. Hydrophobic residues usually appear at the first and fourth  positions in the seven-residue repeats of polypeptides that form  coiled coils, (a) Why do polar or charged residues usually appear  in the remaining five positions? (b) Why is the sequence  Ile-Gln-Glu-Val-Glu-Arg-Asp more likely than the sequence  Trp-Gln-Glu-Tyr-Glu-Arg-Asp to appear in a coiled coil?   7. The digestive tract of the larvae of clothes moths is a strongly  reducing environment. Why is this beneficial to the larvae?   8. Collagen IV, which occurs in basement membranes, contains a  sulfilimine bond (colored red in the structure below) that cross¬  links two collagen triple helices. Identify the parent amino acid  residues that participate in this linkage.   9. Describe the primary, secondary, tertiary, and quaternary struc¬  tures of collagen.   10. Explain why gelatin, which is mostly collagen, is nutritionally in¬  ferior to other types of protein.   11. Globular proteins are typically constructed from several layers of  secondary structure, with a hydrophobic core and a hydrophilic  surface. Is this true for a fibrous protein such as a keratin?   12. Is it possible for a native protein to be entirely irregular, that is,  without a helices, (3 sheets, or other repetitive secondary structure?   13. Which of the following polypeptides is most likely to form an  a helix?   14. Which of the peptides in Problem 13 is least likely to form a  (3 strand?   13. The X-ray crystallographic analysis of a protein often fails to  reveal the positions of the first few and/or the last few residues  of a polypeptide chain. Explain.   16. (a) Is Trp or Gin more likely to be on a protein’s surface? (b) Is  Ser or Val less likely to be in a protein’s interior? (c) Is Leu or lie  less likely to be found in a middle of an a helix? (d) Is Cys or  Ser more likely to be in a (3 sheet?   17. What types of rotational symmetry are possible for a protein with   18. Bacterial glutamate synthetase consists of 12 identical subunits  arranged in two stacked rings of six subunits. How would you  describe this protein’s symmetry?   19. You are performing site-directed mutagenesis to test predictions  about which residues are essential for a protein’s function. Which  of each pair of amino acid substitutions listed below would you  expect to disrupt protein structure the most? Explain.   22. Describe the intra- and intermolecular bonds or interactions that  are broken or retained when collagen is heated to produce gelatin.   23. Under physiological conditions, polylysine assumes a random coil  conformation. Under what conditions might it form an a helix?   23. It is often stated that proteins are quite large compared to the  molecules they bind. However, what constitutes a large number  depends on your point of view. Calculate the ratio of the volume  of a hemoglobin molecule (65 kD) to that of the four 0 2 mol¬  ecules that it binds and the ratio of the volume of a typical  office (4 X 4 X 3 m) to that of the typical (70-kg) office worker  that occupies it. Assume that the molecular volumes of hemoglo¬  bin and 0 2 are in equal proportions to their molecular masses  and that the office worker has a density of 1.0 g/cm 3 . Compare  these ratios. Is this the result you expected?   26. In prokaryotes, the error rate in protein synthesis may be as high  as 5 X 10 4 per codon. What fraction of polypeptides contain¬  ing (a) 500 residues or (b) 2000 residues would you expect to  contain at least one amino acid substitution?   27. Not all heat shock proteins are chaperones; some are proteins that  facilitate the degradation rather than the refolding of other pro¬  teins. Explain why the rate of protein degradation would increase  during heat shock.   28. Protein denaturation can be triggered by a variety of environmen¬  tal insults, including high temperature, covalent modification,  and oxidation. Explain why researchers have observed a correla¬  tion between the level of heat shock proteins and the ratio of ox¬  idized to reduced glutathione (see Section 4-3B) in cells subjected  to oxidative stress.   29. Researchers introduced prions into normal mice and mice that were  genetically predisposed to develop a disease resembling Alzheimer’s.  Explain why the Alzheimer’s-prone mice displayed symptoms of the  prion disease much sooner than did the normal mice.   30. The genetically engineered proteins that accumulate in bacterial  inclusion bodies (Fig. 5-2) form amyloid structures. Such pro¬  teins are often difficult to recover in functional form from the  bacteria. Explain.    Project 3 Visualizing Three-Dimensional Protein  Structures Using the Molecular Visualization  Programs Jmol and PyMOL   There are two halves to Project 3. The first half contains the exercises  using Jmol and the second half contains the same exercises using  PyMOL   1. Obtaining Structural Information. Compare different second¬  ary structure predictions for a given protein sequence, then  inspect its X-ray crystallographic structure.   2. Exploring the Protein Data Bank. Learn how to locate and  download specific protein structure files, sequences, and im¬  ages. Explore additional educational resources such as Molecule  of the Month and links to additional structural biology resources.   3. Examining Protein Structures. Examine a protein structure file  and use molecular modeling programs to visualize the protein  and highlight selected features.   1. Alignment of small molecules. Use Marvin Sketch software to  draw small molecules - ethane and the amino acid, lysine.   2. Alignment of peptides. Construct and optimize a small peptide  with MarvinSketch, then align its 3D structure with the structure  of the same peptide sequence found in a protein.   3. CASP and protein structure prediction. Researchers spend up to  two years predicting the structure of a protein based on its amino  acid sequence, then meet at the CASP conference, where the  newly determined structure is released and teams are evaluated on  how closely their predictions align with the new structures. Learn  more about CASP, explore some of the literature, and then compare  3D alignments of proteins using the Protein Data Bank web site.   Focus concept: The primary structure of insulin is examined, and  the sequences of various animal insulins are compared.    Case 5 Characterization of Subtilisin from the  Antarctic Psychrophile Bacillus TA41  Focus concept: The structural features involved in protein adapta¬  tion to cold temperatures are explored.    A number of human diseases  are caused by point mutations that alter a single amino acid in a  polypeptide, (a) Explain how such a small change in a collagen sub¬  unit can destabilize the entire collagen structure, (b) Investigate  how an amino acid substitution in an enzyme can render the en¬  zyme inactive, (c) Investigate how an amino acid substitution can  affect a protein’s folding efficiency without affecting its ultimate  structure or function.   Branden, C. and Tooze, J., Introduction to Protein Structure (2nd ed.),  Garland Publishing (1999). [A well-illustrated book with chapters  introducing amino acids and protein structure, plus chapters on spe¬  cific proteins categorized by their structure and function.]   Goodsell, D.S., Visual methods from atoms to cells, Structure 13,  347-454 (2005). [Discusses several ways of depicting different fea¬  tures of molecular structures.]   Goodsell, D.S. and Olson, J., Structural symmetry and protein function,  Annu. Rev. Biophys. Biomol. Struct. 29, 105-153 (2000).   Kessel, A. and Ben-Tal, N., Introduction to Proteins. Structure, Function,  and Motion, CRC Press (2011).   Petsko, G.A. and Ringe, D., Protein Structure and Function, New Science  Press (2004).   Brodsky, B. and Persikov, A.V., Molecular structure of the collagen triple  helix, Adv. Protein Chem. 70, 301-339 (2005).   Shoulders, M.D. and Raines, R.T., Collagen structure and stability, Annu.  Rev. Biochem. 78, 929-958 (2009).   Rhodes, G., Crystallography Made Crystal Clear: A Guide for Users of  Macromolecular Models (3rd ed.), Academic Press (2006). [Includes  overviews, methods, and discussions of model quality.]   Rupp, B., Biomolecular Crystallography. Principles, Practice, and Applica¬  tion to Structural Biology, Garland Science (2010).   Wider, G. and Wiithrich, K., NMR spectroscopy of large molecules and  multimolecular assemblies in solution, Curr. Opin. Struct. Biol. 9,  594-601 (1999).   Bolen, D.W. and Rose, G.D., Structure and energetics of the hydrogen-  bonded backbone in protein folding, Annu. Rev. Biochem. 77,  339-362 (2008).    Fersht, A., Structure and Mechanism in Protein Science, Chapter 11,  Freeman (1999).   Karplus, M. and McCammon, J.A., Molecular dynamics simulations of  biomolecules, Nature Struct. Biol. 9, 646-652 (2002).   Das, R. and Baker, D., Macromolecular modeling with Rosetta, Annu.  Rev. Biochem. 77, 363-382 (2008).   Dill, K.A., Ozkan, S.B., Shell, M.S., and Weikl, T.R., The protein fold¬  ing problem, Annu. Rev. Biophys. 37, 289-316 (2008).   Dunker, A.K., Silman, I., Uversky, V.N., and Sussman, J.L., Function and  structure of inherently disordered proteins, Curr. Opin. Struct. Biol.  18, 756-764 (2008).   Fitzkee, N.C., Fleming, P.J., Gong, H., Panasik, N., Jr., Street, T.O., and  Rose, G.D., Are proteins made from a limited parts list? Trends  Biochem. Sci. 30, 73-80 (2005).   Horwich, A.R., Farr, G.W., and Fenton, W.A., GroEL-GroES-mediated  protein folding, Chem. Rev. 106, 1917-1930 (2006)   Onuchic, J.N. and Wolynes, P.G., Theory of protein folding, Curr. Opin.  Struct. Biol. 14, 70-75 (2004).   Schueler-Furman, O., Wang, C., Bradley, P., Misura, K., and Baker, D.,  Progress in modeling of protein structures and interactions, Science  310, 638-642 (2005).   Caughey, B., Baron, G.S., Chesebro, B., and Jeffrey, M., Getting a grip  on prions: oligomers, amyloids, and pathological membrane interac¬  tions, Annu. Rev. Biochem. 78, 177-204 (2009).   Chiti, F. and Dobson, C.M., Protein misfolding, functional amyloid, and  human disease, Annu. Rev. Biochem. 75, 333-366 (2006).   Joyama, B.H. and Weissman, G.S., Amyloid structure: conformational di¬  versity and consequences, Annu. Rev. Biochem. 80, 557-585 (2011).   Moore, R.A., Taubner, L.M., and Priola, S.A., Prion misfolding and dis¬  ease, Curr. Opin. Struct. Biol. 19, 14-22 (2009).   Selkoe, D.J., Cell biology of protein misfolding: the examples of  Alzheimer’s and Parkinson’s diseases, Nature Cell Biol. 6, 1054-1061  (2004). [Describes the relationship between protein misfolding and  cellular pathology.]   B Muscle Contraction Occurs when Myosin Heads  Walk up Thin Filaments    The nearly colorless icefish is the only adult vertebrate that lacks hemoglobin. It can  survive without this otherwise essential oxygen-binding protein because in the cold  (-1.9°C) Antarctic waters where it lives, the fish consumes little oxygen and the  solubility of oxygen is relatively high. [© Doug Allan/Peter Arnold, Inc.]    The preceding two chapters have painted a broad picture of the chemical and  physical properties of proteins but have not delved deeply into their physio¬  logical functions. Nevertheless, it should come as no surprise that the struc¬  tural complexity and variety of proteins allow them to carry out an enormous  array of specialized biological tasks. For example, the enzyme catalysts of vir¬  tually all metabolic reactions are proteins (we consider enzymes in detail in  Chapters 11 and 12). Genetic information would remain locked in DNA were  it not for the proteins that participate in decoding and transmitting that in¬  formation. Remarkably, the thousands of proteins that participate in building,  supporting, recognizing, transporting, and transforming cellular components  act with incredible speed and accuracy and in many cases are subject to mul¬  tiple regulatory mechanisms.   The specialized functions of proteins, from the fibrous proteins we exam¬  ined in Section 6-1C to the precisely regulated metabolic enzymes we discuss  in later chapters, can all be understood in terms of how proteins bind to and  interact with other components of living systems. In this chapter, we focus on  three sets of proteins: the oxygen-binding proteins myoglobin and hemoglo¬  bin, the actin and myosin proteins responsible for muscle contraction, and an¬  tibody molecules. The molecular structures and physiological roles of these  proteins are known in detail, and their proper functioning is vital for human  health. In addition, these proteins serve as models for many of the proteins  we will examine later when we discuss metabolism and the management of  genetic information.    • Myoglobin, with its single heme prosthetic group, exhibits a hyperbolic 0 2 -binding  curve.   • Hemoglobin can adopt the deoxy (T) or oxy (R) conformation, which differ in  0 2 -binding affinity.   • Oxygen binding triggers conformational changes in hemoglobin so that oxygen  binds to the protein cooperatively, yielding a sigmoidal binding curve.    We begin our study of protein function with two proteins that reversibly bind  molecular oxygen (0 2 ). Myoglobin, the first protein whose structure was  determined by X-ray crystallography is a small protein with relatively simple  oxygen-binding behavior. Hemoglobin, a tetramer of myoglobin-like polypep¬  tides, is a more complicated protein that functions as a sophisticated system  for delivering oxygen to tissues throughout the body. The efficiency with  which hemoglobin binds and releases 0 2 is reminiscent of the specificity and  efficiency of metabolic enzymes. It is worthwhile to study hemoglobins struc¬  ture and function because many of the theories formulated to explain 0 2 bind¬  ing to hemoglobin also explain the control of enzyme activity.   Myoglobin is a small intracellular protein in vertebrate muscle. Its X-ray struc¬  ture, determined by John Kendrew in 1959, revealed that most of myoglobins  153 residues are members of eight a helices (traditionally labeled A through H)  that are arranged to form a globular protein with approximate dimensions   FIG. 7-1 Structure of sperm whale myoglobin. This 153-residue monomeric protein  consists of eight a helices, labeled A through H, that are connected by short polypeptide  links (the last half of what was originally thought to be the EF corner has been shown to  form a short helix that is designated the F' helix). The heme group is shown in red.  [Illustration, Irving Geis. Image from the Irving Geis Collection/Howard Hughes Medical  Institute. Rights owned by HHMI. Reproduction by permission only.] Q See Kinemage  Exercise 6-1.   Chapter 7 Protein Function: Myoglobin and  Hemoglobin, Muscle Contraction, and Antibodies   FIG. 7-2 The heme group. The central  Fe(ll) atom is shown liganded to the four N atoms  of the porphyrin ring, whose pyrrole groups are  labeled A-D. The heme is a conjugated system,  so all the Fe—N bonds are equivalent The Fe(ll)  is also liganded to a His side chain and, when it  is present, to 0 2 . The six ligands are arranged at  the corners of an octahedron centered on the Fe  ion (octahedral geometry).    Myoglobin Contains a Heme Prosthetic Group. Myoglobin, other members  of the globin family of proteins (Section 5-4B), and a variety of other proteins  such as cytochrome c (Sections 5-4A and 6-2D) all contain a single heme group  (Fig. 7 -2). The heme is tightly wedged in a hydrophobic pocket between the E  and F helices in myoglobin. The heterocyclic ring system of heme is a porphyrin  derivative containing four pyrrole groups (labeled A-D) linked by methene  bridges (other porphyrins vary in the substituents attached to rings A—D). The  Fe(II) atom at the center of heme is coordinated by the four porphyrin N atoms  and one N from a His side chain (called, in a nomenclature peculiar to myo¬  globin and hemoglobin, His F8 because it is the eighth residue of the F helix).  A molecule of oxygen (0 2 ) can act as a sixth ligand to the iron atom. His E7  (the seventh residue of helix E) hydrogen bonds to the 0 2 with the geometry  shown in Fig. 7-3. Two hydrophobic side chains on the 0 2 -binding side of the  heme, Val Ell and Phe CD1 (the first residue in the segment between helices  C and D), help hold the heme in place. These side chains presumably swing  aside as the protein “breathes” (Section 6-4A), allowing 0 2 to enter and exit.   When exposed to oxygen, the Fe(II) atom of isolated heme is irreversibly  oxidized to Fe(III), a form that cannot bind 0 2 . The protein portion of myo¬  globin (and of hemoglobin, which contains four heme groups in four globin  chains) prevents this oxidation and makes it possible for 0 2 to bind reversibly  to the heme group. Oxygenation alters the electronic state of the Fe(II)—heme  complex, as indicated by its color change from dark purple (the color of hemo¬  globin in venous blood) to brilliant scarlet (the color of hemoglobin in arterial  blood). Under some conditions, the Fe(II) of myoglobin or hemoglobin becomes  oxidized to Fe(III) to form metmyoglobin or methemoglobin, respectively;  these proteins are responsible for the brown color of old meat and dried blood.   In addition to 0 2 , certain other small molecules such as CO, NO, and  H 2 S can bind to heme groups in proteins. These other compounds bind with    FIG. 7-3 The heme complex in myoglobin. In the upper drawing, atoms are  represented in space-filling form (H atoms are not shown). The lower drawing shows the  corresponding skeletal model with a dashed line representing the hydrogen bond between  His E7 and the bound 0 2 . [Based on an X-ray structure by Simon Phillips, MRC Laboratory  of Molecular Biology, Cambridge, U.K. PDBid 1MB0.] Q See Kinemage Exercise 6-1.   much higher affinity than 0 2 , which accounts for their toxicity. CO, for ex¬  ample, has 200-fold greater affinity for hemoglobin than does 0 2 .   Myoglobin Binds 0 2 to Facilitate Its Diffusion. Although myoglobin was  originally thought to be only an oxygen-storage protein, it is now apparent  that its major physiological role is to facilitate oxygen diffusion in muscle (the most  rapidly respiring tissue under conditions of high exertion). The rate at which  0 2 can diffuse from the capillaries to the tissues is limited by its low solubil¬  ity in aqueous solution (—10 4 M in blood). Myoglobin increases the  effective solubility of 0 2 in muscle cells, acting as a kind of molecular bucket  brigade to boost the 0 2 diffusion rate. The oxygen-storage function of myo¬  globin is probably significant only in aquatic mammals such as seals and  whales, whose muscle myoglobin concentrations are around 10-fold greater  than those in terrestrial mammals (which is one reason why Kendrew chose  the sperm whale as a source of myoglobin for his X-ray crystallographic stud¬  ies). Nevertheless, mice in which the gene for myoglobin has been “knocked  out” appear to be normal, although their muscles are lighter in color than  those of wild-type mice. However, closer scrutiny revealed several compensa¬  tory adaptations in these mice, including a greater concentration of red blood  cells and increased capillary density in their muscles. Moreover, many of the  mutant embryos died in utero due to cardiovascular defects. Vertebrates also  express two recently discovered globins: neuroglobin, which is present  mainly in brain, retina, and endocrine tissues, and cytoglobin, which occurs  in most tissues. Neuroglobin protects neurons (nerve cells) from damage  under conditions of ischemia (inadequate blood flow, such as in a stroke),  most likely by preventing reperfusion injury (the damage caused by the oxy¬  gen radicals generated when blood flow is restored). Cytoglobin may have  similar functions.    Myoglobin’s Oxygen-Binding Curve Is Hyperbolic. The reversible binding  of 0 2 to myoglobin (Mb) is described by a simple equilibrium reaction:    Note that biochemists usually express equilibria in terms of dissociation con¬  stants, the reciprocal of the association constants favored by chemists. The 0 2  dissociation of myoglobin can be characterized by its fractional saturation,  Yq 2 , which is defined as the fraction of 0 2 -binding sites occupied by 0 2 :    Yq ranges from zero (when no 0 2 is bound to the myoglobin molecules) to  one (when the binding sites of all the myoglobin molecules are occupied).  Equation 7-1 can be rearranged to    When this expression for [MbOJ is substituted into Eq. 7-2, the fractional  saturation becomes   Chapter 7 Protein Function: Myoglobin and  Hemoglobin, Muscle Contraction, and Antibodies    Factoring out the [Mb] / K term in the numerator and denominator gives    Since 0 2 is a gas, its concentration is conveniently expressed by its partial  pressure, p0 2 (also called the oxygen tension). Equation 7-5 can therefore be  expressed as    This equation describes a rectangular hyperbola and is identical in form to  the equations that describe a hormone binding to its cell-surface receptor or a small  molecular substrate binding to the active site of an enzyme. This hyperbolic func¬  tion can be represented graphically as shown in Fig. 7-4. At low p0 2 , very  little 0 2 binds to myoglobin (}q 2 is very small). As the p0 2 increases, more  0 2 binds to myoglobin. At very high p0 2 , virtually all the 0 2 -binding sites  are occupied and myoglobin is said to be saturated with 0 2 .   The steepness of the hyperbola for a simple binding event, such as 0 2  binding to myoglobin, increases as the value of K decreases. This means that  the lower the value of K, the tighter is the binding. K is equivalent to the con¬  centration of ligand at which half of the binding sites are occupied. In other  words, when p0 2 — K, myoglobin is half-saturated with oxygen. This can be  shown algebraically by substituting p0 2 for K in Eq. 7-6:    At what oxygen concentration will myo¬  globin be 75% saturated with oxygen?   pQ 2 = Yo 2 (p50 + pO 2 )  pT> 2 = Yo 2 p5o + Y Oz p 0 2  pT> 2 — Y Oz p 0 2 = Y 0 j > 5 0  ^ _ Y o 2 p5 0    Thus, K can be operationally defined as the value of p0 2 at which Y — 0.5  (Fig. 7-4).   It is convenient to define K as /> 50 , that is, the oxygen pressure at which  myoglobin is 50% saturated. The p^ 0 for myoglobin is 2.8 torr (760 torr =  1 atm). Over the physiological range of p0 2 in the blood (100 torr in arte¬  rial blood and 30 torr in venous blood), myoglobin is almost fully saturated  with oxygen; for example, Y 0z = 0.97 at p0 2 = 100 torr and 0.91 at 30 torr  (see Sample Calculation 7-1). Consequently, myoglobin efficiently relays oxygen  from the capillaries to muscle cells.   Myoglobin, a single polypeptide chain with one heme group and hence  one oxygen-binding site, is a useful model for other binding proteins. Even  proteins with multiple binding sites for the same small molecule, or ligand,  may generate hyperbolic binding curves like myoglobins. A hyperbolic bind¬  ing curve occurs when ligands interact independently with their binding sites. In  practice, the affinity of a ligand for its binding protein may not be known.  Constructing a binding curve such as the one shown in Fig. 7-4 may provide  this information.    FIG. 7-4 Oxygen-binding curve of myoglobin. Myoglobin is half-saturated  with 0 2 (Vq 2 = 0.5) at an oxygen partial pressure (p0 2 ) of 2.8 torr (dashed  lines). The hyperbolic shape of myoglobin’s binding curve is typical of the simple  binding of a small molecule to a protein. The background is shaded to indicate  the color change that myoglobin undergoes as it binds 0 2 .    How much does fractional saturation change between p0 2 values of  5 and 15, and 15 and 25 torr?   Hemoglobin, the intracellular protein that gives red blood cells their color, is  one of the best-characterized proteins and was one of the first proteins to be  associated with a specific physiological function (oxygen transport). Animals  that are too large (>1 mm thick) for simple diffusion to deliver sufficient oxy¬  gen to their tissues have circulatory systems containing hemoglobin or a pro¬  tein of similar function that does so (Box 7-1).   Mammalian hemoglobin, as we saw in Fig. 6-33, is an a 2 (3 2 tetramer (a  dimer of a(3 protomers). The a and (3 subunits are structurally and evolution-  arily related to each other and to myoglobin. The structure of hemoglobin was  determined by Max Perutz (Box 7-2). Only about 18% of the residues are  identical in myoglobin and in the a and (3 subunits of hemoglobin, but the  three polypeptides have remarkably similar tertiary structures (hemoglobin  subunits follow the myoglobin helix-labeling system, although the a chain has  no D helix). The a(3 protomers of hemoglobin are symmetrically related by a  2-fold rotation (i.e., a rotation of 180° brings the protomers into coincidence).   In addition, hemoglobins structurally similar a and (3 subunits are related by  an approximate 2-fold rotation (pseudosymmetry) whose axis is perpendicu¬  lar to that of the exact 2-fold rotation. Thus, hemoglobin has exact C 2 sym¬  metry and pseudo-Z) 2 symmetry (Section 6-3; objects with D 2 symmetry have    The presence of 0 2 in the earth’s atmosphere and its utility in the  oxidation of metabolic fuels have driven the evolution of various  mechanisms for storing and transporting oxygen. Small organisms  rely on diffusion to supply their respiratory oxygen needs.   However, since the rate at which a substance diffuses varies  inversely with the square of the distance it must diffuse, organ¬  isms of >1-mm thickness overcome the constraints of diffusion  with circulatory systems and boost the limited solubility of 0 2 in  water with specific 0 2 -transport proteins.   Many invertebrates, and even some plants and bacteria, contain  heme-based 0 2 -binding proteins. Single-subunit and multimeric  hemoglobins are found both as intracellular proteins and as extra¬  cellular components of blood and other body fluids. The existence  of hemoglobin-like proteins in some species of bacteria is evi¬  dence of gene transfer from animals to bacteria at one or more  points during evolution. In bacteria, these proteins may function as  sensors of environmental conditions such as local 0 2 concentra¬  tion. In some leguminous plants, the so-called leghemoglobins  bind 0 2 that would otherwise interfere with nitrogen fixation car¬  ried out by bacteria that colonize plant root nodules (Section  21-7). The chlorocruorins, which occur in some annelids (e.g.,  earthworms), contain a somewhat differently derivatized porphyrin  than that in hemoglobin, which accounts for the green color of  chlorocruorins.   The two other types of 0 2 -binding proteins, hemerythrin and  hemocyanin (neither of which contains heme groups), occur only  in invertebrate animals. Hemerythrin, which occurs in only a few  species of marine worms, is an intracellular protein with a subunit  mass of —13 kD. It contains two Fe atoms liganded by His and  acidic residues. It is violet-pink when oxygenated and colorless  when deoxygenated.   Hemocyanins, which are exclusively extracellular, transport 0 2 in  mollusks and arthropods. The molluscan and arthropod hemocyanins    are large multimeric proteins that differ in their primary through  quaternary structures. However, their oxygen-binding sites are highly  similar, consisting of a pair of copper atoms, each liganded by three  His residues.    In this model of the 0 2 -binding site of hemocyanin from the  horseshoe crab Limulus polyphemus, atoms are colored according  to type with C gray, N blue, 0 red, and Cu purple. The otherwise  colorless complex turns blue when it binds 0 2 .   Hemocyanins must be present at high concentrations in order  to function efficiently as oxygen carriers. For example, octopus  hemolymph (its equivalent of blood) contains about 100 mg/mL  hemocyanin. In order to minimize the osmotic pressure of so much  protein, hemocyanins form multimeric structures with masses as  great as 9 X 10 6 D in some species. Hemocyanins are often the  predominant extracellular protein and may therefore have addi¬  tional functions as buffers against pH changes and osmotic fluctu¬  ations. In some invertebrates, hemocyanins may serve as a  nutritional reserve, for example, during metamorphosis or molting.    [Figure based on an X-ray structure by Wim Hoi, University of  Washington School of Medicine. PDBid 10XYJ   determination of the three-dimensional  structures of proteins has become so  commonplace that it is difficult to  appreciate the challenges that faced the  first protein crystallographers. Max Perutz  was a pioneer in this area, spending  many years determining the structure of  hemoglobin at atomic resolution and then  using this information to explain the  physiological function of the protein.   In 1934, two years before Perutz  began his doctoral studies in Cambridge, J.D. Bernal and Dorothy  Crowfoot Hodgkin had placed a crystal of the protein pepsin in an  X-ray beam and obtained a diffraction pattern. Perutz tried the  same experiment with hemoglobin, chosen because of its  abundance, ease of crystallization, and obvious physiological  importance. Hemoglobin crystals yielded diffraction patterns with  thousands of diffraction maxima (called reflections), the result of  X-ray scattering by the thousands of atoms in each protein  molecule. At the time, X-ray crystallography had been used to  determine the structures of molecules containing no more than  around 40 atoms, so the prospect of using the technique to  determine the atomic structure of hemoglobin seemed impossible.  Nevertheless, Perutz took on the challenge and spent the rest of  his long career working with hemoglobin.   In X-ray crystallography, the intensities and the positions of the  reflections can be readily determined, but the values of their  phases (the relative positions of the wave peaks, the knowledge  of which is as important as wave amplitude for image  reconstruction) cannot be directly measured. Although  computational techniques for determining the values of the phases  had been developed for small molecules, methods for solving this  so-called phase problem for such complex entities as proteins  seemed hopelessly out of reach. In 1952, Perutz realized that the  method of isomorphous replacement might suffice to solve the  phase problem for hemoglobin. In this method, a heavy atom such  as a Hg 2+ ion, which is rich in electrons (the particles that scatter  X-rays), must bind to specific sites on the protein without  significantly disturbing its structure (which would change the    positions of the reflections). If this causes measurable changes in  the intensities of the reflections, these differences would provide  the information to determine their phases. With trepidation  followed by jubilation, Perutz observed that Hg-doped hemoglobin  crystals indeed yielded reflections with measurable changes in  intensity but no changes in position. Still, it took another 5 years  to obtain the three-dimensional structure of hemoglobin at low  (5.5-A) resolution and it was not until 1968, some 30 years after  he began the project, that he determined the structure of  hemoglobin at near atomic (2.8-A) resolution. In the meantime,  Perutz’s colleague John Kendrew used the method of  isomorphous replacement to solve the structure of myoglobin, a  smaller and simpler relative of hemoglobin. For their  groundbreaking work, Perutz and Kendrew were awarded the  1962 Nobel Prize in Chemistry.   For Perutz, obtaining the structure of hemoglobin was only part  of his goal of understanding hemoglobin. For example, functional  studies indicated that the four oxygen-binding sites of hemoglobin  interacted, as if they were in close contact, but Perutz’s structure  showed that the binding sites lay in deep and widely separated  pockets. Perutz was also intrigued by the fact that crystals of  hemoglobin prepared in the absence of oxygen would crack when  they were exposed to air (the result, it turns out, of a dramatic  conformational change). Although many other researchers also  turned their attention to hemoglobin, Perutz was foremost among  them in ascribing oxygen-binding behavior to protein structural  features. He also devoted considerable effort to relating functional  abnormalities in mutant hemoglobins to structural changes.   Perutz’s groundbreaking work on the X-ray crystallography of  proteins paved the way for other studies. For example, the first  X-ray structure of an enzyme, lysozyme, was determined in 1965.  The nearly 80,000 macromolecular structures that have been  obtained since then owe a debt to Perutz and his decision to  pursue an “impossible” task and to follow through on his structural  work to the point where he could use his results to explain  biological phenomena.   Perutz, M.F., Rossmann, M.G., Cullis, A.F., Muirhead, H., Will, G., and North, A.C.T.,  Structure of haemoglobin: A three-dimensional Fourier synthesis at 5.5 A resolution,  obtained by X-ray analysis. Nature 185 , 416-422 (1960).   the rotational symmetry of a tetrahedron). The hemoglobin molecule has over¬  all dimensions of about 64 X 55 X 50 A.   Oxygen binding alters the structure of the entire hemoglobin tetramer, so  the structures of deoxyhemoglobin (Fig. 7-5 a) and oxyhemoglobin (Fig. 7-5 £)  are noticeably different. In both forms of hemoglobin, the a and (3 subunits form  extensive contacts: Those at the interface (and its a 2 -|3 2 symmetry equiv¬   alent) involve 35 residues, and those at the a 1 -(3 2 (and cx 2 —(3 X ) interface involve  19 residues. These associations are predominantly hydrophobic, although numer¬  ous hydrogen bonds and several ion pairs are also involved. Note, however, that  the a x -a 2 and |3 1 -(3 2 interactions are tenuous at best because these subunit pairs  are separated by an ~20-A-diameter solvent-filled channel that parallels the  50-A length of hemoglobins exact 2-fold axis (Fig. 7-5).   When oxygen binds to hemoglobin, the a 1 -(3 2 (and cx 2 —(3 X ) contacts shift,  producing a change in quaternary structure. Oxygenation rotates one a(3   Deoxyhemoglobin and (b) oxyhemoglobin. The  protomer is related to the a 2 p 2 protomer  by a 2-fold axis of symmetry (lenticular symbol),  which is perpendicular to the page. Oxygenation  causes one protomer to rotate ~ 15° relative to  the other, bringing the p chains closer together  (compare the lengths of the double-headed  arrows) and shifting the contacts between  subunits at the a 1 -p 2 and e^-p! interfaces  (some of the relevant side chains are drawn in  black). The large gray arrows in Part b indicate  the molecular movements that accompany  oxygenation. [Illustration, Irving Geis. Image from  the Irving Geis Collection/Howard Hughes  Medical Institute. Rights owned by HHMI.  Reproduction by permission only.] Q See  Kinemage Exercises 6-2 and 6-3.    dimer —15° with respect to the other ap dimer (gray arrows in Fig. 7-5 b),  which brings the p subunits closer together and narrows the solvent-filled cen¬  tral channel (Fig. 7-5). Some atoms in the a x -p 2 and a 2 -P x interfaces shift  by as much as 6 A (oxygenation causes such extensive quaternary structural  changes that crystals of deoxyhemoglobin shatter on exposure to 0 2 ). This  structural rearrangement is a crucial element of hemoglobins oxygen-binding  behavior.   Chapter 7 Protein Function: Myoglobin and  Hemoglobin, Muscle Contraction, and Antibodies   FIG. 7-6 Oxygen-binding curve of  hemoglobin. In whole blood, hemoglobin is  half-saturated at an oxygen pressure of 26 torr.  The normal sea level values of human arterial  and venous p0 2 are indicated (atmospheric p0 2  is 160 torr at sea level). The 0 2 -binding curve  for myoglobin is included for comparison. The  dashed line is a hyperbolic 0 2 -binding curve with  the same p 50 as hemoglobin. The background is  shaded to indicate the color change that  hemoglobin undergoes as it binds 0 2 . © See  the Animated Figures.   Hemoglobin has a p 50 of 26 torr (i.e., hemoglobin is half-saturated with 0 2  at an oxygen partial pressure of 26 torr), which is nearly 10 times greater than  the y? 5 o of myoglobin. Moreover, hemoglobin does not exhibit a myo¬  globin-like hyperbolic oxygen-binding curve. Instead, 0 2 binding to  hemoglobin is described by a sigmoidal (S-shaped) curve  (Fig. 7-6). This permits the blood to deliver much more 0 2 to the tis¬  sues than if hemoglobin had a hyperbolic curve with the same p 50  (dashed curve in Fig. 7-6). For example, hemoglobin is nearly fully  saturated with 0 2 at arterial oxygen pressures (Yq 2 — 0.95 at 100  torr) but only about half-saturated at venous oxygen pressures  (Yq 2 = 0.55 at 30 torr). This 0.40 difference in oxygen saturation, a  measure of hemoglobins ability to deliver 0 2 from the lungs to  the tissues, would be only 0.25 if hemoglobin exhibited hyperbolic  binding behavior.   In any binding system, a sigmoidal curve is diagnostic of a cooper¬  ative interaction between binding sites. This means that the binding  of a ligand to one site affects the binding of additional ligands to the  other sites. In the case of hemoglobin, 0 2 binding to one subunit  increases the 0 2 affinity of the remaining subunits. The initial slope  of the oxygen-binding curve (Fig. 7-6) is low, as hemoglobin subunits inde¬  pendently compete for the first 0 2 . However, an 0 2 molecule bound to one of  hemoglobins subunits increases the 0 2 -binding affinity of its other subunits,  thereby accounting for the increasing slope of the middle portion of the  sigmoidal curve.   The Hill Equation Describes Hemoglobin’s 0 2 -Binding Curve. The earliest  attempt to analyze hemoglobins sigmoidal 0 2 dissociation curve was formu¬  lated by Archibald Hill in 1910. Hill assumed that hemoglobin (Hb) bound  n molecules of 0 2 in a single step,   that is, with infinite cooperativity. Thus, in analogy with the derivation of  Eq. 7-6,    which is known as the Hill equation. Like Eq. 7-6, it describes the degree of  saturation of hemoglobin as a function of p0 2 (see Sample Calculation 7-2).   Infinite 0 2 binding cooperativity, as Hill assumed, is a physical impossi¬  bility. Nevertheless, n may be taken to be a nonintegral parameter related to  the degree of cooperativity among interacting hemoglobin subunits rather than  the number of subunits that bind 0 2 in one step. The Hill equation can then  be taken as a useful empirical curve-fitting relationship rather than as an in¬  dicator of a particular model of ligand binding.    Calculate the fractional saturation of hemoglobin at p0 2 — 50 torr and n = 3.   The quantity n, the Hill constant , increases with the degree of cooperativity  of a reaction and therefore provides a convenient although simplistic characteriza¬  tion of a ligand-binding reaction. If n = 1, Eq. 7-8 describes a hyperbola as  does Eq. 7-6 for myoglobin, and the 0 2 -binding reaction is said to be non-  cooperative. If n > 1, the reaction is described as being positively coopera¬  tive, because 0 2 binding increases the affinity of hemoglobin for further 0 2  binding (cooperativity is infinite in the limit that n = 4, the number of 0 2  binding sites in hemoglobin). Conversely, if n < 1, the reaction is said to be  negatively cooperative, because 0 2 binding would then reduce the affinity of  hemoglobin for subsequent 0 2 binding.   The Hill coefficient, n, and the value of p 50 that best describe hemoglo¬  bins saturation curve can be graphically determined by rearranging Eq. 7-8.  First, divide both sides by 1 — Y 0 y.   Y Oi = (>50)” + (/>Q 2 )” = (/ > 5 o)” + (/ > Q 2 )” r 7 q-,   Factoring out the [(/> 5 o)” + (/>0 2 )”] term gives   1 - ^o 2 [{p^Y + (po 2 y] - (po 2 y (p 50 y    The linear plot of log[lo 2 /(l — ^o 2 )] versus log y>0 2 , the Hill plot, has a  slope of n and an intercept on the log p0 2 axis of log y? 50 (recall that the lin¬  ear equation y = mx + b describes a line with a slope of m and an x intercept  of — blni).   Figure 7-7 shows the Hill plots for myoglobin and purified hemoglobin.  For myoglobin, the plot is linear with a slope of 1, as expected. Although all  subunits of hemoglobin do not bind 0 2 in a single step as was assumed in    FIG. 7-7 Hill plots for myoglobin and  purified hemoglobin. Note that this is a log-log  plot. At p0 2 = P 5 o, ho 2 /(1 ~ >o 2 ) = E [The p 50  for hemoglobin in vivo is higher than the p 50 of  purified hemoglobin due to its binding of certain  substances present in the red cell (see below).]   Chapter 7 Protein Function: Myoglobin and  Hemoglobin, Muscle Contraction, and Antibodies    deriving the Hill equation, its Hill plot is essentially linear for values of Yq 2  between 0.1 and 0.9. When p0 2 — p^ Yq 2 = 0.5, and    As can be seen in Fig. 7-7, this is the region of maximum slope, whose value  is customarily taken to be the Hill coefficient, n. For normal human hemo¬  globin, the Hill coefficient is between 2.8 and 3.0; that is, hemoglobins oxy¬  gen binding is highly, but not infinitely, cooperative. Many abnormal  hemoglobins exhibit smaller Hill coefficients (Section 7-IE), indicating that  they have a less than normal degree of cooperativity.   At Yq 2 values near zero, when few hemoglobin molecules have bound even  one 0 2 molecule, the Hill plot for hemoglobin assumes a slope of 1 (Fig. 7-7,  lower asymptote) because the hemoglobin subunits independently compete for  0 2 as do molecules of myoglobin. At Yq 2 values near 1, when at least three of  hemoglobins four 0 2 -binding sites are occupied, the Hill plot also assumes a  slope of 1 (Fig. 7-7, upper asymptote) because the few remaining unoccupied  sites are on different molecules and therefore bind 0 2 independently.   Extrapolating the lower asymptote in Fig. 7-7 to the horizontal axis in¬  dicates, according to Eq. 7-11, that p 50 = 30 torr for binding the first 0 2  to purified hemoglobin. Likewise, extrapolating the upper asymptote yields  p 50 = 0.3 torr for binding hemoglobins fourth 0 2 . Thus, the fourth 0 2 binds  to hemoglobin with 100-fold greater affinity than the first. This difference, as  we will see below, is entirely due to the influence of the globin chain on the  0 2 affinity of heme.   FIG. 7-8 Movements of the heme and F  helix during the T —» R transition in  hemoglobin. In the T form (blue), the Fe is 0.6  A above the center of the domed porphyrin ring.  On assuming the R form (red), the Fe moves  into the plane of the now undomed porphyrin,  where it can more tightly bind 0 2 , and, in doing  so, pulls His F8 and its attached F helix with  it. O See Kinemage Exercise 6-4 and the  Animated Figures.   The cooperativity of oxygen binding to hemoglobin arises from the effect of  the ligand-binding state of one heme group on the ligand-binding affinity of  another. Yet the hemes are 25 to 37 A apart—too far to interact electronically.  Instead, information about the 0 2 -binding status of a heme group is mechan¬  ically transmitted to the other heme groups by motions of the protein. These  movements are responsible for the different quaternary structures of oxy- and  deoxyhemoglobin depicted in Fig. 7-5.   to R. On the basis of the X-ray structures of oxy- and deoxyhemoglobin, Perutz  formulated a model for hemoglobin oxygenation. In the Perutz mechanism ,  hemoglobin has two stable conformational states , the T state (the conformation of  de oxyhemoglobin) and the R state (the conformation of oxyhemoglobin). The con¬  formations of all four subunits in T-state hemoglobin differ from those in the  R state. Oxygen binding initiates a series of coordinated movements that result  in a shift from the T state to the R state within a few microseconds:   1. In the T state, the Fe(II) in each of the four hemes is situated ~0.6 A  out of the heme plane because of a pyramidal doming of the porphyrin  group toward His F8 and because the Fe—Np orp hy r in bonds are too  long to allow the Fe to lie in the porphyrin plane (Fig. 7-8). 0 2 bind¬  ing changes the hemes electronic state, which shortens the  Fe—N porphyrin bonds by ~0.1 A and causes the porphyrin doming to  subside. Consequently, during the T —> R transition, the Fe(II) moves  into the center of the heme plane.   2. The Fe(II) drags the covalently linked His F8 along with it. However,  the direct movement of His F8 by 0.6 A toward the heme plane would  cause it to collide with the heme. To avoid this steric clash, the  attached F helix tilts and translates by ~1 A across the heme plane.   3. The changes in tertiary structure are coupled to a shift in the arrange- Section 1 Oxygen Binding to   the T —» R transition is the result of movements of residues at the  ai-(3 2 and a 2 -|3i interfaces, that is, at the interface between the two  protomeric units of hemoglobin. In the T state, His 97 in the (3 chain  contacts Thr 41 in the a chain (Fig. 7-9 d). In the R state, His 97 con¬  tacts Thr 38, which is positioned one turn back along the C helix  (Fig. 7-9 £). In both conformations, the “knobs” on one subunit mesh  nicely with the “grooves” on the other. An intermediate position would   FIG. 7-9 Changes at the ct 1 -p 2 interface during the T —» R  transition in hemoglobin, (a) The T state and (b) the R state. In the  upper drawings, the C helix is represented by a purple ribbon, the  contacting residues forming the o^C-p^FG contact are shown in ball-  and-stick form colored by atom type (C green, N blue, and 0 red),  and their van der Waals surfaces are outlined by like-colored dots.   The lower drawings are the corresponding schematic diagrams of the  a 1 C-|3 2 FG contact. Upon a T —» R transformation, the (3 2 FG region  shifts by one turn along the c^C helix with no stable intermediate   (note how in both conformations, the knobs formed by the side chains  of His 9713 and Asp 99|3 fit between the grooves on the C helix  formed by the side chains of Thr 38a, Thr 41a, and Pro 44a). The  subunits are joined by different hydrogen bonds in the two quaternary  states. Figure 7-5 provides another view of these interactions. [Based  on X-ray structures by Giulio Fermi, Max Perutz, and Boaz Shaanan,  MRC Laboratory of Molecular Biology, Cambridge, U.K. PDBids (a)  2HHB and ( b ) 1HHOJ O See Kinemage Exercise 6-5.   FIG. 7-10 Networks of ion pairs and hydrogen bonds in  deoxyhemoglobin. These bonds, which involve the last two  residues of (a) the a chains and (b) the p chains, are ruptured in the  T —» R transition. Two groups that become partially deprotonated in   the R state (part of the Bohr effect) are indicated by white plus signs.  [Illustration, Irving Geis. Image from the Irving Geis Collection/Howard  Hughes Medical Institute. Rights owned by HHMI. Reproduction by  permission only.]    be severely strained because it would bring His 97 and Thr 41 too  close together (i.e., knobs on knobs).   4. The C-terminal residues of each subunit (Arg 14la and His l46p) in  T-state hemoglobin each participate in a network of intra- and intersub¬  unit ion pairs (Fig. 7-10) that stabilize the T state. However, the con¬  formational shift in the T —» R transition tears away these ion pairs in  a process that is driven by the energy of formation of the Fe—0 2 bonds.   The essential feature of hemoglobins T —> R transition is that its subunits  are so tightly coupled that large tertiary structural changes within one subunit can¬  not occur without quaternary structural changes in the entire tetrameric protein.  Hemoglobin is limited to only two quaternary forms, T and R, because the  intersubunit contacts shown in Fig. 7-9 act as a binary switch that permits  only two stable positions of the subunits relative to each other. The inflexibil¬  ity of the a^Px and a 2 ~P 2 interfaces requires that the T —» R shift occur si¬  multaneously at both the a x —P 2 and a 2 —p x interfaces. No one subunit or  dimer can greatly change its conformation independently of the others.   We are now in a position to structurally rationalize the cooperativity of oxy¬  gen binding to hemoglobin. The T state of hemoglobin has low 0 2 affinity,  mostly because of the 0.1 A greater length of its Fe—0 2 bond relative to that  of the R state (see the blue structure shown in Fig. 7-8). Experimental evidence  indicates that when at least one 0 2 has bound to each ap dimer, the strain in  the T-state hemoglobin molecule is sufficient to tear away the C-terminal ion  pairs, thereby snapping the protein into the R state. All the subunits are thereby  simultaneously converted to the R-state conformation whether or not they have  bound 0 2 . Unliganded subunits in the R-state conformation have increased oxy¬  gen affinity because they are already in the 0 2 -binding conformation. This ac¬  counts for the high 0 2 affinity of nearly saturated hemoglobin.   The Bohr Effect Enhances Oxygen Transport. The conformational changes  in hemoglobin that occur on oxygen binding decrease the piCs of several  groups. Recall that the tendency for a group to ionize depends on its micro¬  environment, which may include other ionizable groups. For example, in T-state  hemoglobin, the N-terminal amino groups of the a subunits and the    C-terminal His of the (3 subunits are positively charged and participate in ion  pairs (see Fig. 7 - 10 ). The formation of ion pairs increases the p K values of  these groups (makes them less acidic and therefore less likely to give up their  protons). In R-state hemoglobin, these ion pairings are absent, and the p K’s  of the groups decrease (making them more acidic and more likely to  give up protons). Consequently, under physiological conditions, hemo¬  globin releases ~ 0.6 protons for each 0 2 it binds. Conversely, increas¬  ing the pH, that is, removing protons, stimulates hemoglobin to bind  more 0 2 (Fig- 7-11). This phenomenon is known as the Bohr effect  after Christian Bohr (father of the physicist Niels Bohr), who first Y  reported it in 1904. ° 2   The Bohr effect has important physiological functions in trans¬  porting 0 2 from the lungs to respiring tissue and in transporting the  C0 2 produced by respiration back to the lungs (Fig. 7-12). The C0 2  produced by respiring tissues diffuses from the tissues to the capillar¬  ies. This dissolved C 0 2 forms bicarbonate (HCO^) on ty ver y slowly,  by the reaction    C0 2 + H 2 0 H + + HCO 3 "   However, in the erythrocyte (red blood cell; Greek: erythrose, red + kytos , a  hollow vessel), the enzyme carbonic anhydrase greatly accelerates this reac¬  tion. Accordingly, most of the C0 2 in the blood is carried in the form of bi¬  carbonate (in the absence of carbonic anhydrase, bubbles of C0 2 would form  in the blood).   In the capillaries, where p0 2 is low, the H + generated by bicarbonate for¬  mation is taken up by hemoglobin in forming the ion pairs of the T state,  thereby inducing hemoglobin to unload its bound 0 2 . This H + uptake, more¬  over, facilitates C0 2 transport by stimulating bicarbonate formation.  Conversely, in the lungs, where p0 2 is high, 0 2 binding by hemoglobin dis¬  rupts the T-state ion pairs to form the R state, thereby releasing the Bohr pro¬  tons, which recombine with bicarbonate to drive off C0 2 . These reactions are  closely matched, so they cause very little change in blood pH (see Box 2-2).   The Bohr effect provides a mechanism whereby additional oxygen can be  supplied to highly active muscles, where the p0 2 may be <20 torr. Such mus¬  cles generate lactic acid (Section 15-3A) so fast that they lower the pH of the  blood passing through them from 7.4 to 7 . 2 . At a p0 2 of 20 torr, hemoglo¬  bin releases —10% more 0 2 at pH 7.2 than it does at pH 7.4 (Fig. 7-11).    FIG. 7-1 1 The Bohr effect. The 0 2 affinity  of hemoglobin increases with increasing pH.   The dashed line indicates the p0 2 in actively  respiring muscle. [After Benesch, R.E. and  Benesch, R., Adv. Protein Chem. 28, 212  (1974).] Q See the Animated Figures.    FIG. 7-1 2 The roles of hemoglobin and myoglobin in 0 2 and  C0 2 transport. Oxygen is inhaled into the lungs at high p0 2 , where it  binds to hemoglobin in the blood. The 0 2 is then transported to  respiring tissue, where the p0 2 is low. The 0 2 therefore dissociates  from the Hb and diffuses into the tissues, where it is used to oxidize  metabolic fuels to C0 2 and H 2 0. In rapidly respiring muscle tissue,  the 0 2 first binds to myoglobin (whose oxygen affinity is higher than    that of hemoglobin). This increases the rate at which 0 2 can diffuse  from the capillaries to the tissues by, in effect, increasing its solubility.  The Hb and C0 2 (mostly as HCO 3 ") are then returned to the lungs,  where the C0 2 is exhaled.   Stripped hemoglobin (left) has higher 0 2 affinity  than whole blood (red curve). Adding BPG or  C0 2 or both to hemoglobin shifts the  dissociation curve back to the right (lowers  hemoglobin’s 0 2 affinity). [After Kilmartin, J.V.  and Rossi-Bernardi, L, Physiol. Rev. 53, 884  (1973).] O See the Animated Figures.    C0 2 also modulates 0 2 binding to hemoglobin by combining reversibly  with the N-terminal amino groups of blood proteins to form carbamates:   The T (deoxy) form of hemoglobin binds more C0 2 as carbamate than does  the R (oxy) form. When the C0 2 concentration is high, as it is in the capil¬  laries, the T state is favored, stimulating hemoglobin to release its bound 0 2 .  The protons released by carbamate formation further promote 0 2 release  through the Bohr effect. Although the difference in C0 2 binding between the  oxy and deoxy states of hemoglobin accounts for only ~5% of the total blood  C0 2 , it is nevertheless responsible for around half the C0 2 transported by the  blood. This is because only ~10% of the total blood C0 2 is lost through the  lungs in each circulatory cycle.    Bisphosphoglycerate Binds to Deoxyhemoglobin. Highly purified  (“stripped”) hemoglobin has a much greater oxygen affinity than hemoglobin in  whole blood (Fig. 7-13). This observation led Joseph Barcroft, in 1921, to spec¬  ulate that blood contains some other substance besides C0 2 that affects oxygen  binding to hemoglobin. This compound is D-2,3-bisphosphoglycerate (BPG).   BPG binds tightly to deoxyhemoglobin but only weakly to oxy¬  hemoglobin. The presence of BPG in mammalian erythrocytes there¬  fore decreases hemoglobins oxygen affinity by keeping it in the deoxy  conformation. In other vertebrates, different phosphorylated com¬  pounds elicit the same effect.   BPG has an indispensable physiological function: In arterial  blood, where p0 2 is ~100 torr, hemoglobin is ~95% saturated  with 0 2 , but in venous blood, where p0 2 is ~30 torr, it is only  55% saturated (Fig. 7-6). Consequently, in passing through the  capillaries, hemoglobin unloads ~40% of its bound 0 2 . In the ab¬  sence of BPG, little of this bound 0 2 would be released since he¬  moglobins 0 2 affinity is increased, thus shifting its 0 2 dissociation  curve significantly toward lower p0 2 (Fig. 7-13, left). BPG also  plays an important role in adaptation to high altitudes (Box 7-3).   The X-ray structure of a BPG—deoxyhemoglobin complex  shows that BPG binds in the central cavity of deoxyhemoglobin  (Fig. 7-14). The anionic groups of BPG are within hydrogen¬  bonding and ion-pairing distances of the N-terminal amino  groups of both (3 subunits. The T —» R transformation brings the  two (3H helices together, which narrows the central cavity (com¬  pare Figs. 7-5a and 7-5 b) and expels the BPG. It also widens the    FIG. 7-14 Binding of BPG to deoxyhemoglobin. BPG (red) binds in  hemoglobin’s central cavity. The BPG, which has a charge of -5 under  physiological conditions, is surrounded by eight cationic groups (blue) extending  from the two p subunits. In the R state, the central cavity is too narrow to  contain BPG. Some of the ion pairs and hydrogen bonds that help stabilize the  T state (Fig. 7-10b) are indicated at the lower right. [Illustration, Irving Geis.  Image from the Irving Geis Collection/Howard Hughes Medical Institute. Rights  owned by HHMI. Reproduction by permission only.] Q See Kinemage  Exercise 6-3.    Atmospheric pressure decreases with altitude, so that the oxygen  pressure at 3000 m (10,000 feet) is only —110 torr, 70% of its  sea-level pressure. A variety of physiological responses are re¬  quired to maintain normal oxygen delivery (without adaptation, p0 2  levels of 85 torr or less result in mental impairment).   High-altitude adaptation is a complex process that involves  increases in the number of erythrocytes and the amount of hemo¬  globin per erythrocyte. It normally requires several weeks to com¬  plete. Yet, as is clear to anyone who has climbed to high altitude,  even a 1-day stay there results in a noticeable degree of adapta¬  tion. This effect results from a rapid increase in the amount of  BPG synthesized in erythrocytes (from —4 mM to —8 mM; BPG  cannot cross the erythrocyte membrane). As illustrated by plots of  Yq 2 versus p0 2 , the high altitude-induced increase in BPG causes  the 0 2 -binding curve of hemoglobin to shift from its sea-level  position (black line ) to a lower affinity position (red line).   At sea level, the difference between arterial and venous p0 2 is  70 torr (100 torr - 30 torr), and hemoglobin unloads 38% of its  bound 0 2 . However, when the arterial p0 2 drops to 55 torr, as it  does at an altitude of 4500 m, hemoglobin would be able to un¬  load only 30% of its 0 2 . High-altitude adaptation (which decreases  the amount of 0 2 that hemoglobin can bind in the lungs but, to a  greater extent, increases the amount of 0 2 it releases at the tis¬  sues) allows hemoglobin to deliver a near-normal 37% of its bound  0 2 . BPG concentrations also increase in individuals suffering from  disorders that limit the oxygenation of the blood (hypoxia), such  as various anemias and cardiopulmonary insufficiency.   The BPG concentration in erythrocytes can be adjusted more  rapidly than hemoglobin can be synthesized (Box 15-2; erythro¬  cytes lack nuclei and therefore cannot synthesize proteins). An  altered BPG level is also a more sensitive regulator of oxygen  delivery than an altered respiratory rate. Hyperventilation, another  early response to high altitude, may lead to respiratory alkalosis  (Box 2-2). Interestingly, individuals in long-established Andean and  Himalayan populations exhibit high lung capacity, along with high    hemoglobin levels and, often, enlarged right ventricles (reflecting  increased cardiac output), compared to individuals from low-  altitude populations.   In contrast to the mechanism of human adaptation to high  altitude, most mammals that normally live at high altitudes (e.g., the  llama) have genetically altered hemoglobins that have higher 0 2 -  binding affinities than do their sea-level cousins. Thus, both raising  and lowering hemoglobin’s p 50 can provide high-altitude adaptation.   distance between the (3 N-terminal amino groups from 16 to 20 A, which  prevents their simultaneous hydrogen bonding with BPG s phosphate groups.  BPG therefore binds to and stabilizes only the T conformation of hemoglo¬  bin by cross-linking its (3 subunits. This shifts the T R equilibrium to¬  ward the T state, which lowers hemoglobins 0 2 affinity.   Fetal Hemoglobin Has Low BPG Affinity. The effects of BPG also help sup¬  ply the fetus with oxygen. A fetus obtains its 0 2 from the maternal circula¬  tion via the placenta. The concentration of BPG is the same in adult and fetal  erythrocytes, but BPG binds more tightly to adult hemoglobin than to fetal  hemoglobin. The higher oxygen affinity of fetal hemoglobin facilitates the  transfer of 0 2 to the fetus.   Fetal hemoglobin has the subunit composition a 2 y 2 in which the y sub¬  unit is a variant of the (3 chain (Section 5-4B). Residue 143 of the (3 chain  of adult hemoglobin has a cationic His residue, whereas the y chain has an  uncharged Ser residue. The absence of this His eliminates a pair of interac¬  tions that stabilize the BPG—deoxyhemoglobin complex (Fig. 7-14).   Hemoglobin Is a Model Allosteric Protein. The cooperativity of oxygen  binding to hemoglobin is a classic model for the behavior of many other   Chapter 7 Protein Function: Myoglobin and  Hemoglobin, Muscle Contraction, and Antibodies   multisubunit proteins (including many enzymes) that bind small molecules.  In some cases, binding of a ligand to one site increases the affinity of other  binding sites on the same protein (as in 0 2 binding to hemoglobin). In other  cases, a ligand decreases the affinity of other binding sites (as when BPG bind¬  ing decreases the 0 2 affinity of hemoglobin). All these effects are the result of  allosteric interactions (Greek: alios, other + stereos, solid or space). Allosteric  effects, in which the binding of a ligand at one site affects the binding of another  ligand at another site, generally require interactions among subunits of oligomeric  proteins. The T —» R transition in hemoglobin subunits explains the difference  in the oxygen affinities of oxy- and deoxyhemoglobin. Other proteins exhibit  similar conformational shifts, although the molecular mechanisms that underlie  these phenomena are not completely understood.   Two models that account for cooperative ligand binding have received the  most attention. One of them, the symmetry model of allosterism, formulated  in 1965 by Jacques Monod, Jeffries Wyman, and Jean-Pierre Changeux, is de¬  fined by the following rules:   1. An allosteric protein is an oligomer of symmetrically related subunits  (although the a and p subunits of hemoglobin are only pseudosym-  metrically related).   2. Each oligomer can exist in two conformational states, designated R and  T; these states are in equilibrium.   3. The ligand can bind to a subunit in either conformation. Only the con¬  formational change alters the affinity for the ligand.   4. The molecular symmetry of the protein is conserved during the conforma¬  tional change. The subunits must therefore change conformation in a  concerted manner; in other words, there are no oligomers that simul¬  taneously contain R- and T-state subunits.   The symmetry model is diagrammed for a tetrameric binding protein in  Fig. 7-15. If a ligand binds more tightly to the R state than to the T state,  ligand binding will promote the T —» R shift, thereby increasing the affin¬  ity of the unliganded subunits for the ligand.   One major objection to the symmetry model is that it is difficult to be¬  lieve that oligomeric symmetry is perfectly preserved in all proteins, that is, that  the T —» R shift occurs simultaneously in all subunits regardless of the num¬  ber of ligands bound. In addition, the symmetry model can account only for  positive cooperativity, although some proteins exhibit negative cooperativity.   An alternative to the symmetry model is the sequential model of alloster¬  ism, proposed by Daniel Koshland. According to this model, ligand binding  induces a conformational change in the subunit to which it binds, and coop¬  erative interactions arise through the influence of those conformational changes  on neighboring subunits. The conformational changes occur sequentially as  more ligand-binding sites are occupied (Fig. 7-16). The ligand-binding affin¬  ity of a subunit varies with its conformation and may be higher or lower than  that of the subunits in the ligand-free protein. Thus, proteins that follow the  sequential model of allosterism may be positively or negatively cooperative.   If the mechanical coupling between subunits in the sequential model is  particularly strong, the conformational changes occur simultaneously and the  oligomer retains its symmetry, as in the symmetry model. Thus, the symme¬  try model of allosterism may be considered to be an extreme case of the more  general sequential model.   FIG. 7-1 5 The symmetry model of allosterism. Squares and circles represent  T- and R-state subunits, respectively, of a tetrameric protein. The T and R states are in  equilibrium regardless of the number of ligands (represented by S) that have bound to the  protein. All the subunits must be in either the T or the R form; the model does not allow  combinations of T- and R-state subunits in the same protein.   FIG. 7-16 The sequential model of allosterism. Ligand binding progressively  induces conformational changes in the subunits, with the greatest changes occurring in  those subunits that have bound ligand. The symmetry of the oligomeric protein is not  preserved in this process as it is in the symmetry model.    Oxygen binding to hemoglobin exhibits features of both models. The quater¬  nary T —> R conformational change is concerted, as the symmetry model requires.  Yet ligand binding to the T state does cause small tertiary structural changes, as  the sequential model predicts. These minor conformational shifts are undoubtedly  responsible for the buildup of strain that eventually triggers the T —» R transition.  It therefore appears that the complexity of ligand—protein interactions in hemo¬  globin and other proteins allows binding processes to be fine-tuned to the needs  of the organism under changing internal and external conditions. We will revisit  allosteric effects when we discuss enzymes in Chapter 12.   Before the advent of recombinant DNA techniques, mutant hemoglobins pro¬  vided what was an almost unique opportunity to study structure-function re¬  lationships in proteins. This is because, for many years, hemoglobin was the  only protein of known structure that had a large number of well-characterized  naturally occurring variants. The examination of individuals with physiolog¬  ical disabilities, together with the routine electrophoretic screening of human  blood samples, has led to the discovery of over 1000 variant hemoglobins,  >90% of which result from single amino acid substitutions in a globin  polypeptide chain. Indeed, about 5% of the worlds human population are car¬  riers of an inherited variant hemoglobin.   Not all hemoglobin variants produce clinical symptoms, but some abnor¬  mal hemoglobin molecules do cause debilitating diseases (~300,000 individu¬  als with serious hemoglobin disorders are born every year; naturally occurring  hemoglobin variants that are lethal are, of course, never observed). Table 7-1  lists several hemoglobin variants. Mutations that destabilize hemoglobins terti¬  ary or quaternary structure alter hemoglobins oxygen-binding affinity and    ^Hemoglobin variants are usually named after the place where they were discovered (e.g., hemoglobin Boston).   Chapter 7 Protein Function: Myoglobin and reduce its cooperativity (Hill coefficient). Moreover, the unstable hemoglobins   the erythrocytes to lyse (break open). The resulting hemolytic anemia (ane¬  mia is a deficiency of red blood cells) compromises 0 2 delivery to tissues.   Certain mutations at the 0 2 -binding site of either the a or p chain  favor the oxidation of Fe(II) to Fe(III). Individuals carrying the resulting  methemoglobin subunit exhibit cyanosis, a bluish skin color, due to the pres¬  ence of methemoglobin in their arterial blood. These hemoglobins have re¬  duced cooperativity (Hill coefficient —1.2 compared to a maximum value of  2, since only two subunits in each of these methemoglobins can bind oxygen).   Mutations that increase hemoglobins oxygen affinity lead to increased  numbers of erythrocytes in order to compensate for the less than normal  amount of oxygen released in the tissues. Individuals with this condition,  which is named polycythemia, often have a ruddy complexion.    A Single Amino Acid Change Causes Sickle-Cell Anemia. Most harmful  hemoglobin variants occur in only a few individuals, in many of whom the  mutation apparently originated. However, —10% of African-Americans and as  many as 25% of black Africans carry a single copy of (are heterozygous for)  the gene for sickle-cell hemoglobin (hemoglobin S). Individuals who carry  two copies of (are homozygous for) the gene for hemoglobin S suffer from  sickle-cell anemia, in which deoxyhemoglobin S forms insoluble filaments  that deform erythrocytes (Fig. 7-17). In this painful, debilitating, and often  fatal disease, the rigid, sickle-shaped cells cannot easily pass through the cap¬  illaries. Consequently, in a sickle-cell “crisis,” the blood flow to some tissues  may be completely blocked, resulting in tissue death. In addition, the mechan¬  ical fragility of the misshapen cells results in hemolytic anemia. Heterozygotes,  whose hemoglobin is —40% hemoglobin S, usually lead a normal life, al¬  though their erythrocytes have a shorter than normal lifetime.   In 1945, Linus Pauling hypothesized that sickle-cell anemia was the result  of a mutant hemoglobin, and in 1949 he showed that the mutant hemoglobin  had a less negative ionic charge than normal adult hemoglobin. This was the  first evidence that a disease could result from an alteration in the molecular struc¬  ture of a protein. Furthermore, since sickle-cell anemia is an inherited disease,  a defective gene must be responsible for the abnormal protein. Nevertheless, the  molecular defect in sickle-cell hemoglobin was not identified until 1956, when  Vernon Ingram showed that hemoglobin S contains Val rather than Glu at the  sixth position of each (3 chain. This was the first time an inherited disease was  shown to arise from a specific amino acid change in a protein.   The X-ray structure of deoxyhemoglobin S has revealed that one mutant  Val side chain in each hemoglobin S tetramer nestles into a hydrophobic pocket  on the surface of a (3 subunit in another hemoglobin tetramer (Fig. 7-18). This  intermolecular contact allows hemoglobin S tetramers to form linear polymers.  Aggregates of 14 strands that wind around each other form fibers extending  throughout the length of the erythrocyte (Fig. 7-19). The hydrophobic pocket  on the (3 subunit cannot accommodate the normally occurring Glu side chain,  and the pocket is absent in oxyhemoglobin. Consequently, neither normal he¬  moglobin nor oxyhemoglobin S can polymerize. In fact, hemoglobin S fibers  dissolve essentially instantaneously on oxygenation, so none are present in ar¬  terial blood. The danger of sickling is greatest when erythrocytes pass through    FIG. 7-1 7 Scanning electron micrographs of human erythrocytes, (a) Normal  erythrocytes are flexible, biconcave disks that can tolerate slight distortions as they pass  through the capillaries (many of which have smaller diameters than erythrocytes).   (b) Sickled erythrocytes from an individual with sickle-cell anemia are elongated and rigid  and cannot easily pass through capillaries, [(a) © Susumu Nishinaga/Photo Researchers, Inc.;  (b) Bill Longcore/Photo Researchers, Inc.]    FIG. 7-1 8 Structure of a deoxyhemoglobin S fiber, (a) The arrangement of  deoxyhemoglobin S molecules in the fiber. Only three subunits of each deoxyhemoglobin S  molecule are shown, (b) The side chain of the mutant Val 6 in the p 2 chain of one  hemoglobin S molecule (yellow knob in Part a) binds to a hydrophobic pocket on the p!  subunit of a neighboring deoxyhemoglobin S molecule. [Illustration, Irving Geis. Image from  the Irving Geis Collection/Howard Hughes Medical Institute. Rights owned by HHMI.  Reproduction by permission only.]    the capillaries, where deoxygenation occurs. The polymerization of hemoglobin  S molecules is time and concentration dependent, which explains why blood  flow blockage occurs only sporadically (in a sickle-cell “crisis”).   Interestingly, many hemoglobin S homozygotes have only a mild form of  sickle-cell anemia because they express relatively high levels of fetal hemoglo¬  bin, which contains y chains rather than the defective (3 chains. The fetal he¬  moglobin dilutes the hemoglobin S, making it more difficult for hemoglobin  S to aggregate during the 10—20 s it takes for an erythrocyte to travel from  the tissues to the lungs for reoxygenation. The administration of hydroxyurea,   the first and as yet the only effective treatment for sickle-cell anemia, amelio¬  rates the symptoms of sickle-cell anemia by increasing the fraction of cells con¬  taining fetal hemoglobin (although the mechanism whereby hydroxyurea acts  is unknown).   FIG. 7-19 Electron micrograph of  deoxyhemoglobin S fibers spilling out of a  ruptured erythrocyte. [Courtesy of Robert  Josephs, University of Chicago.]    Hemoglobin S Protects Against Malaria. Before the advent of modern pal¬  liative therapies, individuals with sickle-cell anemia rarely survived to maturity.  Natural selection has not minimized the prevalence of the hemoglobin S vari¬  ant, however, because heterozygotes are more resistant to malaria, an often  lethal infectious disease. Of the 2.5 billion people living within malaria-endemic  areas, 100 million are clinically ill with the disease at any given time and around   Chapter 7 Protein Function: Myoglobin and  Hemoglobin, Muscle Contraction, and Antibodies    FIG. 7-20 Correspondence between  malaria and the sickle-cell gene. The blue  areas of the map indicate regions where malaria  is or was prevalent. The pink areas represent the  distribution of the gene for hemoglobin S. Note  the overlap ( purple ) of the distributions.   • Explain the structural basis for cooperative  oxygen binding to hemoglobin.   • Sketch a binding curve (% bound ligand  versus ligand concentration) for coopera¬  tive and noncooperative binding.   • Explain why the 0 2 -binding behavior of  myoglobin and hemoglobin can be  summed up by a single number (the p 50 ).   • Could a binding protein have a Hill con¬  stant of zero?   • Describe how myoglobin and hemoglobin  function in delivering 0 2 from the lungs to  respiring tissues.   • What is the physiological relevance of the  Bohr effect and BPG?   • Explain why mutations can increase or  decrease the oxygen affinity and coopera-  tivity of hemoglobin. How can the body  compensate for these changes?    1 million, mostly very young children, die from it each year. Malaria is caused  by the mosquito-borne protozoan Plasmodium falciparum , which resides within  an erythrocyte during much of its 48-h life cycle. Infected erythrocytes adhere  to capillary walls, causing death when cells impede blood flow to a vital organ.   The regions of equatorial Africa where malaria is a major cause of  death coincide closely with those areas where the sickle-cell gene is preva¬  lent (Fig. 7-20), thereby suggesting that the sickle-cell gene confers resistance  to malaria. How does it do so? Plasmodia increase the acidity of infected  erythrocytes by ~0.4 pH units. The lower pH favors the formation of de¬  oxyhemoglobin via the Bohr effect, thereby increasing the likelihood of  sickling in erythrocytes that contain hemoglobin S. Erythrocytes damaged  by sickling are normally removed from the circulation by the spleen.  During the early stages of a malarial infection, parasite-enhanced sickling  probably allows the spleen to preferentially remove infected erythrocytes.  In the later stages of infection, when the parasitized erythrocytes attach to  the capillary walls (presumably to prevent the spleen from removing them  from the circulation), sickling may mechanically disrupt the parasite.  Consequently, heterozygous carriers of hemoglobin S in a malarial region  have an adaptive advantage: They are more likely to survive to maturity  than individuals who are homozygous for normal hemoglobin. Thus, in  malarial regions, the fraction of the population who are heterozygotes for  the sickle-cell gene increases until their reproductive advantage is balanced  by the correspondingly increased proportion of homozygotes (who, with¬  out modern medical treatment, die in childhood).    Myosin is a motor protein that undergoes conformational changes as it hydrolyzes  ATP.    • The sliding filament model of muscle contraction describes the movement of thick  filaments relative to thin filaments.   • The globular protein actin can form structures such as microfilaments and the thin  filaments of muscle.    One of the most striking characteristics of living things is their capacity for  organized movement. Such phenomena occur at all structural levels and in¬  clude such diverse vectorial processes as the separation of replicated chromo¬  somes during cell division, the beating of flagella and cilia, and, most  obviously, muscle contraction. In this section, we consider the structural and  chemical basis of movement in striated muscle, one of the best understood  mobility systems.    The voluntary muscles, which include the skeletal muscles, have a striated  (striped) appearance when viewed by light microscopy (Fig. 7-21). Such mus¬  cles consist of long multinucleated cells (the muscle fibers) that contain par¬  allel bundles of myofibrils (Greek: my os, muscle; Fig. 7-22). Electron  micrographs show that muscle striations arise from the banded structure of  multiple in-register myofibrils. The bands are formed by alternating regions of  greater and lesser electron density called A bands and I bands, respectively  (Fig. 7-23). The myofibrils repeating unit, the sarcomere (Greek: sarkos ,  flesh), is bounded by Z disks at the center of each I band. The A band is cen¬  tered on the H zone, which in turn is centered on the M disk. The A band  contains 150-A-diameter thick filaments, and the I band contains 70-  A-diameter thin filaments. The two sets of filaments are linked by cross¬  bridges where they overlap.   A contracted muscle can be as much as one-third shorter than its fully ex¬  tended length. The contraction results from a decrease in the length of the  sarcomere, caused by reductions in the lengths of the I band and the H zone    FIG. 7-21 Photomicrograph of a muscle fiber. The longitudinal axis of the fiber is  horizontal (perpendicular to the striations). The alternating pattern of dark A bands and light  I bands from multiple in-register myofibrils is clearly visible. [J.C. Revy, CNRI/Photo  Researchers.]   FIG. 7-22 Skeletal muscle organization. A muscle (a) consists of bundles of muscle  fibers (b), each of which is a long, thin, multinucleated cell (c) that may run the length of  the muscle. Muscle fibers contain bundles of laterally aligned myofibrils (c/), which in turn  consist of bundles of alternating thick and thin filaments.   Chapter 7 Protein Function: Myoglobin and  Hemoglobin, Muscle Contraction, and Antibodies   electron micrograph shows parts of three  myofibrils, which are separated by horizontal  gaps. The accompanying interpretive drawing  shows the major features of the myofibril: the  light I band, which contains only thin filaments;  the A band, whose dark H zone contains only  thick filaments and whose darker outer  segments contain overlapping thick and thin  filaments; the Z disk, to which the thin filaments  are anchored; and the M disk, which arises from  a bulge at the center of each thick filament. The  myofibril’s functional unit, the sarcomere, is the  region between two successive Z disks.  [Courtesy of Hugh Huxley, Brandeis University.]   (Fig. 7-24 d). These observations, made by Hugh Huxley in 1954 (Box 7-4),  are explained by the sliding filament model in which interdigitated thick and  thin filaments slide past each other (Fig. 7-24 b). Thus, during a contraction,  a muscle becomes shorter, and because its total volume does not change, it  also becomes thicker.   Thick Filaments Consist Mainly of Myosin. Vertebrate thick filaments are  composed almost entirely of a single type of protein, myosin, which consists  of six polypeptide chains: two 220-kD heavy chains and two pairs of differ¬  ent light chains, the so-called essential and regulatory light chains (ELC and  RLC) that vary in size between 15 and 22 kD, depending on their source. X-  Ray structure determinations by Ivan Rayment and Hazel Holden of the N-  terminal half of the myosin heavy chain, the so-called myosin head, reveals   FIG. 7-24 Myofibril contraction, (a) Electron micrographs  showing myofibrils in progressively more contracted states. The  lengths of the I band and H zone decrease on contraction, whereas  the lengths of the thick and thin filaments remain constant.    (b) Interpretive drawings showing interpenetrating sets of thick and  thin filaments sliding past each other. [Courtesy of Hugh Huxley,  Brandeis University.]   Hugh Huxley (1924-) The mechanism  of muscle action has fascinated scientists  for hundreds if not thousands of years. The  first close look at muscle fibers came in  1682, when Antoni van Leeuwenhoek’s  early microscope revealed a pattern of thin  longitudinal fibers. In the modern era,  research on muscle has followed one of  two approaches. First, it is possible to study  muscle as an energy-transducing system, in  which metabolic energy is generated and  consumed. This line of research received a  tremendous boost in the 1930s with the discovery that ATP is the  energy source for muscle contraction. The second approach  involves treating muscle as a mechanical system, that is, sorting  out its rods and levers. Ultimately, a molecular approach united the  mechanical and energetic aspects of muscle research. The  insights of Hugh Huxley made this possible.   The molecular characterization of muscle did not occur  overnight. In 1859, Willi Kuhne isolated a proteinaceous  substance from muscle tissue that he named “myosin” (almost  certainly a mixture of many proteins), but it tended to aggregate  and was therefore not as popular a study subject as the more  soluble proteins such as hemoglobin. A major breakthrough in  muscle protein chemistry came in 1941, when the Hungarian  biochemist Albert Szent-Gyorgyi showed that two types of protein  could be extracted from ground muscle by a solution with high  salt concentration (Szent-Gyorgyi also contributed to the  elucidation of the citric acid cycle; Box 17-1). Extraction for 20  minutes yielded a protein he named myosin A but which is now  called myosin. However, extraction overnight yielded a second  protein which he named myosin B but is now called actomyosin.   It soon became apparent that myosin B was a mixture of two  proteins, myosin and a new protein which was named actin.  Further work showed that threads of actomyosin contracted to  — 10% of their original length in the presence of ATP. Since actin  and myosin alone do not contract in the presence of ATP, the  contraction must have resulted from their interaction. However, it  took another decade to develop a realistic model of how myosin  and actin interact.   A number of theories had been advanced to explain muscle  contraction. According to one theory, the cytoplasm of muscle  cells moved like that of an amoeba. Other theories proposed that  muscle fibers took up and gave off water or repelled and  attracted other fibers electrostatically. Linus Pauling, who had  recently discovered the structures of the a helix and p sheet  (Box 6-1), ventured that myosin could change its length by  shifting between the two protein conformations. Huxley formulated  an elegant—and correct—explanation in his sliding filament model  for muscle contraction.   In 1948, Huxley began his doctoral research at Cambridge  University in the United Kingdom, in the laboratory of John  Kendrew (who 10 years later determined the first X-ray structure  of a protein, that of myoglobin; Section 7-1 A). There, through  X-ray studies on frog muscle fibers, Huxley established that the  X-ray diffraction pattern changes with the muscle’s physiological    state. Furthermore, he showed that muscle contained two sets of  parallel fibers, rather than one, and that these fibers were linked  together by multiple cross-links. These observations became the  germ for further research, which he carried out at MIT in 1953  and 1954. He teamed up with Jean Hanson, a Briton who was  also working at MIT. Hanson made good use of her knowledge of  muscle physiology and her expertise in phase-contrast microscopy,  a technique that could visualize the banded patterns of muscle  fibers. Huxley and Hanson observed rabbit muscle fibers under  different experimental conditions, making precise measurements of  the width of the A and I bands in sarcomeres (Fig. 7-23). In one  experiment, they extracted myosin from the muscle fiber, noted  the loss of the dark A band, and concluded that the A band  consists of myosin. When they extracted both actin and myosin, all  identifiable structure was lost, and they concluded that actin is  present throughout the sarcomere.   When ATP was added, the muscle slowly contracted, and  Huxley and Hanson were able to measure the shortening of the  I band. The A band maintained a constant length but became  darker. A muscle fiber under the microscope could also be  stretched by pulling on the coverslip. As the muscle “relaxed,” the  I band increased in width and the A band became less dense.  Measurements were made for muscle fibers contracted to 60% of  their original length and stretched to 120% of their original length.   The key to the sliding filament model that Huxley described  and subsequently refined is that the individual molecules (that is,  their observable fibrous forms) do not shrink or extend but instead  slide past each other. During contraction, actin filaments (thin  filaments) in the I band are drawn into the A band, which consists  of stationary myosin-containing filaments (thick filaments). During  stretching, the actin filaments withdraw from the A band. Similar  conclusions were reached by the team of Andrew Huxley (no  relation to Hugh) and Rolf Niedergerke, who examined the  contraction of living frog muscle fibers. Both groups published  their work in back-to-back papers in Nature in 1954.   Hugh Huxley went on to supply additional details to his sliding  filament model. For example, he showed that myosin forms cross¬  bridges with actin fibers. However, these bridges are asymmetric,  pointing in opposite directions in the two halves of the sarcomere.  This arrangement allows myosin to pull thin filaments in opposite  directions toward the center of the sarcomere (Fig. 7-24).   While Huxley was describing the mechanism of muscle  contraction, Watson and Crick discovered the structure of DNA,  and Max Perutz made a decisive breakthrough in the use of heavy  metal atoms to solve the phase problem in his X-ray studies of  hemoglobin (Box 7-2). Collectively, these discoveries indicated the  tremendous potential for describing biological phenomena in  molecular terms. Subsequent studies of muscle contraction have  used electron microscopy, X-ray crystallography, and enzymology to  probe the fine details of the sliding filament model, including the  structure of myosin’s lever arm, the composition of the thin  filament, and the exact role of ATP in triggering conformational  changes that generate mechanical force.   Huxley, H.E. and Hanson, J., Changes in the cross-striations of muscle during con¬  traction and stretch and their structural interpretation, Nature 173, 973-976 (1954).   FIG. 7-25 Structure of myosin, (a) A ribbon diagram of the myosin head from  chicken muscle. The heavy chain is represented by a ribbon diagram embedded in its semi¬  transparent molecular surface with different portions green, blue, and red. The essential and  regulatory light chains, RLC and ELC, are drawn in worm form with each colored in rainbow  order from its N-terminus (blue) to its C-terminus (red). A sulfate ion, shown in space-filling  form (C red and S yellow), occupies the binding site of ATP’s (3-phosphate group. An RLC-  bound Ca 2+ ion (lower left) is represented by a cyan sphere. [Based on an X-ray structure  by Ivan Rayment and Hazel Holden, University of Wisconsin. PDBid 2MYS.] (b) Diagram of  the myosin molecule. Its two identical heavy chains (green and orange) each have an  N-terminal globular head and an a-helical tail. Between the head and tail is an a helix, the  lever arm, that associates with the two kinds of light chains (magenta and yellow). The tails  wind around each other to form a 1600-A-long parallel coiled coil.    that it forms an elongated (55 X 165 A) globular head to which one subunit  each of ELC and RLC bind (Fig. 7-25 d). The C-terminal half of the heavy  chain forms a long fibrous a-helical tail, two of which associate to form a left-  handed coiled coil. Thus, myosin consists of a 1600-A-long rodlike segment with  two globular heads (Fig. 7-25 b). The amino acid sequence of myosins a-helical  tail is characteristic of coils such as those in keratin (Section 6-1C): It has a  seven-residue pseudorepeat, a-b-c-d-e-f-g, with nonpolar residues predominat¬  ing at positions a and d.   FIG. 7-26 Structure of the thick filament, (a) Electron ( b) Drawing of a thick filament, in which several hundred myosin   micrograph showing the myosin heads projecting from the thick molecules form a staggered array with their globular heads pointing   filament. [From Trinick, J. and Elliott, A., J. Mol. Biol. 131, 135 (1977).] away from the filament.    Under physiological conditions, several hundred myosin molecules aggre¬  gate to form a thick filament. The rodlike tails pack end to end in a regular  staggered array, leaving the globular heads projecting to the sides on both ends  (Fig. 7-26). These myosin heads form the cross-bridges to thin filaments in  intact myofibrils. The myosin head, which is an ATPase (ATP-hydrolyzing en¬  zyme), has its ATP-binding site located in a 13-A-deep V-shaped pocket.   Thin Filaments Consist Mainly of Actin. Thin filaments consist mainly of  polymers of actin, the most abundant cytosolic protein in eukaryotes. In its  monomeric form, this ~375-residue protein is known as G-actin (G for glob¬  ular); when polymerized, it is called F-actin (F for fibrous). Each actin sub¬  unit has binding sites for ATP and a Ca 2+ or Mg 2+ ion that are located in a  deep cleft (Fig. 7-27). ATP hydrolysis to ADP + P z is not required for actin  polymerization but occurs afterward (Section 7-2C).   The fibrous nature of F-actin and its variable fiber lengths have thwarted  its crystallization in a manner suitable for X-ray crystallographic analysis.  Consequently, our current understanding of the atomic structure of F-actin is  based on electron micrographs (Fig. 7-28 d) together with low resolution mod¬  els based on X-ray studies of oriented gels of F-actin into which high resolu¬  tion atomic models of G-actin have been fitted (Fig. 7-28 £). These models  indicate that the actin polymer is a double-stranded helix of subunits in which  each subunit contacts four others. Each actin subunit has the same head-to-   FIG. 7-27 X-Ray structure of rabbit  muscle G-actin in complex with ATP and a  Ca 2+ ion. The four domains of the protein are  colored cyan, magenta, orange, and yellow, and  the N- and C-termini are labeled. The ATP, which  is drawn in space-filling form with C green, N  blue, 0 red, and P orange, binds at the bottom  of a deep cleft between the domains. The Ca 2+  ion is represented by a light green sphere.   [Based on an X-ray structure by Leslie Burtnik,  University of British Columbia, Vancouver, British  Columbia. PDBid 3HBT.]   (a) Cryoelectron microscopy-based image. The  tropomyosin binding sites (see below) are blue.  [Courtesy of Daniel Safer, University of  Pennsylvania, and Ronald Milligan, The Scripps  Research Institute, La Jolla, California.] ( b ) Model  based on fitting a known X-ray structure of  G-actin to a cryoEM-based image of F-actin,  represented by five consecutive actin • ADP  subunits drawn in ribbon form, each with a  different color, embedded in their semitransparent  molecular surface. The ADPs are drawn in  space-filling form with C green, N blue, 0 red,  and P orange. The central subunit ( magenta ) is  oriented as is the G-actin in Fig. 7-27. [Based on  a structure by Keiichi Namba, Osaka University,  Japan. PDBid 3MFP.]   Chapter 7 Protein Function: Myoglobin and  Hemoglobin, Muscle Contraction, and Antibodies    FIG. 7-29 Model of the myosin-actin  interaction. This space-filling model was  constructed from the X-ray structures of actin  and the myosin head and electron micrographs  of their complex. The actin filament is at the top.  The myosin heavy chain is colored as in Fig.   7-25 and the light chains are yellow and  magenta. The coiled-coil tail is not shown. An  ATP-binding site is located in a cleft in the blue  domain of the myosin head. In a myofibril, every  actin monomer has the potential to bind a  myosin head, and the thick filament has many  myosin heads projecting from it. [Modified from a  drawing by Ivan Rayment and Hazel Holden,  University of Wisconsin.]    tail orientation (e.g., all the nucleotide-binding clefts open upward in Fig. 7-28#),  so the assembled fiber has a distinct polarity. The end of the fiber toward  which the nucleotide-binding sites open is known as the  end, and its  opposite end is the  end. The  ends of the thin filaments bind to the  Z disk (Fig. 7-23).   Each of muscle F-actin s monomeric units can bind a single myosin head  (Fig. 7-29), probably by ion pairing and by the association of hydrophobic  patches on each protein. Electron micrographs indicate that the myosin heads  bound to an F-actin filament all have the same orientation (Fig. 7-30) and  that in thin filaments that are still attached to the Z disk, the myosin heads  all point away from the Z disk.   Tropomyosin and Troponin Are Thin Filament Components. Myosin and  actin, the major components of muscle, account for 60 to 70% and 20 to  25% of total muscle protein, respectively. Of the remainder, two proteins that  are associated with the thin filaments are particularly prominent:   1. Tropomyosin, a homodimer whose two 284-residue a-helical subunits  wrap around each other to form a parallel coiled coil that extends  nearly the entire 400-A length of the molecule (a portion of which  is shown in Fig. 6-15#). Multiple copies of these rod-shaped proteins  are joined head-to-tail to form cables wound in the grooves of the  F-actin helix such that each tropomyosin molecule contacts seven con¬  secutive actin subunits in a quasi-equivalent manner (Fig. 7-30).    FIG. 7-30 Cryoelectron microscopy-based image at <25-A resolution of a thin  filament decorated with myosin heads. F-actin is red, tropomyosin is blue, the myosin  motor domain is yellow, and the essential light chain is green. The helical filament has a  pitch (rise per turn) of 370 A. [Courtesy of Ronald Milligan, The Scripps Research Institute,  La Jolla, California.]   2. Troponin, which consists of three subunits: TnC, a Ca 2+ -binding pro¬  tein; Tnl, which binds to actin; and TnT, an elongated molecule,  which binds to tropomyosin at its head-to-tail junctions. The X-ray  structure of troponin in complex with four Ca 2+ ions (Fig. 7-31), de¬  termined by Robert Fletterick, reveals that Tnl closely resembles the  myosin light chains and that the inhibitory segment of Tnl binds to  TnC’s rigid central helix in this Ca 2+ -activated state.   The tropomyosin—troponin complex, as we will see below, regulates muscle contrac¬  tion by controlling the access of the myosin heads to their binding sites on actin.   Other proteins serve to form the Z disk and the M disk and to organize the  arrays of thick and thin filaments. For instance, a-actinin, a rodlike homo-  dimeric protein that cross-links F-actin filaments, is localized in the Z disks  interior and is therefore thought to attach oppositely oriented thin filaments  to the Z disk.   One of the more unusual muscle proteins, titin, the longest known polypep¬  tide chain (34,350 residues), is composed of ~300 repeating globular domains.  Three to six titin molecules associate with each thick filament, spanning the 1-  |Jim distance between the M and Z disks. Titin is believed to function as a mo¬  lecular bungee cord to keep the thick filament centered on the sarcomere:  During muscle contraction, it compresses as the sarcomere shortens, but when  the muscle relaxes, titin resists sarcomere extension past the starting point.   Nebulin, which is also extremely large (6669 residues), is a mainly a-helical  protein that is associated with the thin filament. It is thought to set the length  of the thin filament by acting as a template for actin polymerization. This  length is held constant by tropomodulin, which caps the  end of the thin  filament (the end not attached to the Z disk), thereby preventing further actin  polymerization and depolymerization. CapZ (also called (5-actinin) is an  a-actinin-associated heterodimer that similarly caps the  end of F-actin.   The M disk (Fig. 7-23) arises from the local enlargement of in-register  thick filaments. Two proteins that are associated with this structure,  myomensin and M-protein, bind to titin and are therefore likely to partici¬  pate in thick filament assembly, as does the thick filament-associated myosin¬  binding protein C.   r Duchenne muscular dystrophy (DMD) and the less severe Becker mus¬  cular dystrophy (BMD) are both sex-linked muscle-wasting diseases. In  DMD, which has an onset age of 2 to 5 years, muscle degeneration exceeds  muscle regeneration, causing progressive muscle weakness and ultimately  death, typically due to respiratory disorders or heart failure, usually by age 25.  In BMD, the onset age is 5 to 10 years and there is an overall less progres¬  sive course of muscle degeneration and a longer (sometimes normal) life span  than in individuals with DMD.   The gene responsible for DMD/BMD encodes a 3685-residue protein  named dystrophin, which has a normal abundance in muscle tissue of  0.002%. Individuals with DMD usually have no detectable dystrophin in their  muscles, whereas those with BMD mostly have dystrophins of altered sizes.  Evidently, the dystrophins of individuals with DMD are rapidly degraded,  whereas those of individuals with BMD are semifunctional.   Dystrophin is a member of a family of flexible rod-shaped proteins that  includes other actin-binding cytoskeletal components. Dystrophin associates  on the inner surface of the muscle plasma membrane with a transmembrane  glycoprotein complex, where it helps anchor F-actin to the extracellular ma¬  trix and thereby protects the plasma membrane from being torn by the me¬  chanical stress of muscle contraction. Although such small tears are common  in muscle cells, they occur much more frequently in dystrophic cells, leading  to a greatly increased rate of cell death.    FIG. 7-31 X-Ray structure of chicken  skeletal muscle troponin. TnC is red, Tnl is  blue, and TnT is gold. The four Ca 2+ ions bound  by TnC are represented by cyan spheres. [Based  on an X-ray structure by Robert Fletterick,  University of California at San Francisco. PDBid  1YTZJ   Chapter 7 Protein Function: Myoglobin and  Hemoglobin, Muscle Contraction, and Antibodies    B Muscle Contraction Occurs when Myosin Heads Walk  up Thin Filaments   In order to complete our description of muscle contraction we must deter¬  mine how ATP hydrolysis is coupled to the sliding filament model. If the slid¬  ing filament model is correct then it would be impossible for a myosin  cross-bridge to remain attached to the same point on a thin filament during  muscle contraction. Rather, it must repeatedly detach and then reattach itself  at a new site further along the thin filament toward the Z disk. This, in turn,  suggests that muscular tension is generated through the interaction of myosin cross¬  bridges with thin filaments. The actual contractile force is provided by ATP hy¬  drolysis. Thus, myosin is a motor protein that converts the chemical energy  of ATP hydrolysis to the mechanical energy of movement. Edwin Taylor for¬  mulated a model for myosin-mediated ATP hydrolysis, which has been refined  by the structural studies of Rayment, Holden, and Ronald Milligan as follows  (Fig. 7-32):   1. ATP binds to a myosin head in a manner that causes myosins actin-  binding site to open up and release its bound actin.   2. Myosins active site (distinct from its actin-binding site) closes around  the ATP. The resulting hydrolysis of the ATP to ADP + P z “cocks” the  myosin head, that is, puts it into its “high energy” conformation in  which it is approximately perpendicular to the thick filament.   3. The myosin head binds weakly to an actin monomer that is closer to  the Z disk than the one to which it had been bound previously.   4. Myosin releases P z , which causes its actin-binding site to close, thereby  increasing its affinity for actin.   5. The resulting transient state is immediately followed by the power  stroke, a conformational shift that sweeps the myosin head s C-terminal  tail by an estimated —100 A toward the Z disk relative to the actin-  binding site on its head, thus translating the attached thin filament by  this distance toward the M disk.   Because the reaction cycle involves several steps, some of which are irre¬  versible (e.g., ATP hydrolysis and P z release), the entire cycle is unidirectional.  The —500 myosin heads on every thick filament asynchronously cycle through  this reaction sequence about five times each per second during a strong mus¬  cular contraction. The myosin heads thereby “walk” or “row” up adjacent thin  filaments toward the Z disk with the concomitant contraction of the muscle.  Although myosin is dimeric, its two heads function independently.   Calcium Triggers Muscle Contraction. Highly purified actin and myosin can  contract regardless of the Ca 2+ concentration, but preparations containing in¬  tact thin filaments contract only in the presence of Ca 2+ , due to the regula¬  tory action of troponin C (Fig. 7-31). Stimulation of a myofibril by a nerve  impulse results in the release of Ca 2+ from the sarcoplasmic reticulum (a sys¬  tem of flattened vesicles derived from the endoplasmic reticulum). As a result,  the intracellular [Ca 2+ ] increases from —10 7 to—10 3 M. The higher    FIG. 7-32 Mechanism of force generation in muscle. (Opposite) The myosin head  “walks” up the actin thin filament through a unidirectional cyclic process that is driven by  ATP hydrolysis to ADP and P,. Only one myosin head is shown. The actin monomer to  which the myosin head is bound at the beginning of the cycle is more darkly colored for  reference. [After Rayment, I. and Holden, H., Curr. Opin. Struct. Biol. 3, 949 (1993).] O  See the Animated Figures.    ATP binds to myosin  heads; actin binding  site opens; myosin  head releases actin.   FIG. 7-33 Comparison of the positions of tropomyosin on the thin filament in  the absence and presence of Ca 2+ . In this superposition of cryoelectron microscopy-based  images, the F-actin filament is gold, the tropomyosin in the absence of Ca 2+ is red, and  that in the presence of Ca 2+ is green. [Courtesy of William Lehman, Boston University  School of Medicine.]    calcium concentration triggers the conformational change in the troponin-  tropomyosin complex that exposes the site on actin where the myosin head  binds (Fig. 7-33). When the myofibril [Ca 2+ ] is low (Ca 2+ is rapidly pumped  back into the sarcoplasmic reticulum by ATP-requiring Ca 2+ pumps; Section  10-3B), the troponin—tropomyosin complex assumes its resting conformation,  blocking myosin binding to actin and causing the muscle to relax.   Although actin and myosin are most prominent in muscle, they also occur in  other tissues. In fact, actin is ubiquitous and is usually the most abundant cy¬  toplasmic protein in eukaryotic cells, typically accounting for 5 to 10% of their  total protein. Nonmuscle actin forms ~70-A-diameter fibers known as micro¬  filaments that can be visualized by immunofluorescence microscopy (in which  a fluorescent-tagged antibody is used to “stain” the actin to which it binds; Fig.  7-34). In nonmuscle cells, actin plays an essential role in many processes, in¬  cluding changes in cell shape, cell division, endocytosis, and organelle transport.   Microfilament Treadmilling Can Mediate Locomotion. ATP-G-actin binds  to both ends of an F-actin filament but with a greater affinity for its  end  (hence its name). This polymerization activates F-actin subunits to hydrolyze  their bound ATP to ADP + P z with the subsequent dissociation of P r The  resulting conformation change reduces the affinity of an ADP-F-actin subunit  for its neighboring subunits relative to that of ATP-F-actin. Since F-  actin—catalyzed ATP hydrolysis occurs more slowly than actin polymerization  and F-actin s bound nucleotide does not exchange with those in solution (its  nucleotide-binding site is blocked by its associated subunits), F-actins more  recently polymerized and hence predominantly ATP-containing subunits  occur largely at its  end, whereas its  end consists mainly of less re¬  cently polymerized and hence predominantly ADP-containing subunits. The  lesser affinity of ADP-containing actin subunits for F-actin results in their net  dissociation from the  end of the polymer.   The steady state (when the microfilament maintains a constant length)  occurs when the net rate of addition of subunits at the  end matches the  net rate of dissociation of subunits at the  end. Then, subunits that have   microfilaments in a fibroblast resting on the  surface of a culture dish are revealed by  immunofluorescence microscopy using a  fluorescently labeled antibody to actin. When  the cell begins to move, the filaments  disassemble. [Courtesy of John Victor Small,  Austrian Academy of Sciences, Salzburg.]   FIG. 7-35 Microfilament treadmilling. In the steady state, actin monomers continually  add to the  end of the filament (left) but dissociate at the same rate from the  end  (right). The filament thereby maintains a constant length while its component monomers  translocate from left to right.   FIG. 7-36 Scanning electron micrograph of a crawling macrophage. The leading  edge of this white blood cell (top) is ruffled where it has become detached from the  surface and is in the process of extending. The cell’s trailing edge or tail (bottom), which  is still attached to the surface, is gradually being pulled toward the leading edge. The  rate of actin polymerization is greatest at the leading edge. The macrophage is in the  course of engulfing Staphyloccus sp. bacteria (orange globules) by a process known as  phagocytosis. [© Eye of Science/Photo Researchers, Inc.]    added to the  end move toward the  end where they dissociate, a process  called treadmilling (Fig. 7-35). Thus, a fluorescenriy labeled actin monomer  is seen to move from the  end of the micro filament toward its  end.  Treadmilling is driven by the free energy of ATP hydrolysis and hence is not  at equilibrium.   The directional growth of actin filaments exerts force against the plasma  membrane, allowing a cell to extend its cytoplasm in one direction. If the cy¬  toplasmic protrusion anchors itself to the underlying surface, then the cell can  use the adhesion point for traction to advance further. In order for the cell to  crawl, however, the trailing edge of the cell must release its contacts with the  surface while newer contacts are being made at the leading edge (Fig. 7-36).  In addition, as microfilament polymerization proceeds at the leading edge, de¬  polymerization must occur elsewhere in the cell, since the pool of G-actin is  limited. A variety of actin-binding proteins modulate the rate of actin depoly¬  merization and repolymerization in vivo.   Actin-mediated cell locomotion, that is, amoeboid motion, is the most  primitive mechanism of cell movement. Nevertheless, virtually all eukaryotic  cells undertake some version of it, even if it involves just a small patch of actin  near the cell surface. More extensive microfilament rearrangements are essen¬  tial for cells such as neutrophils (a type of white blood cell) that travel rela¬  tively long distances to sites of infection or inflammation.   • Explain how myosin structure relates to its  function as a motor protein.   • Draw a diagram of the components of a  sarcomere, including the locations of all  the proteins mentioned in the text. Which  components are globular and which are  fibrous?   • Explain the molecular basis of the sliding  filament model of muscle contraction.   • What are the roles of Ca 2+ and ATP in  muscle contraction?   • Is it possible to construct a fibrous protein  from globular subunits?   • How does a microfilament differ from the  thin filament in a myofibril?   • Describe the process of actin polymerization  and depolymerization during cell crawling.   Chapter 7 Protein Function: Myoglobin and  Hemoglobin, Muscle Contraction, and Antibodies    • The humoral immune response is mediated by antibodies, which include constant  domains as well as variable domains that recognize specific antigens.    All organisms are continually subject to attack by other organisms, including  disease-causing microorganisms and viruses. In higher animals, these  pathogens may penetrate the physical barrier presented by the skin and mu¬  cous membranes (a first line of defense) only to be identified as foreign in¬  vaders and destroyed by the immune system. Two types of immunity have  been distinguished:   1. Cellular immunity, which plays a role in fighting most pathogens and  is particularly effective at eliminating virally infected cells, is mediated by  T lymphocytes or T cells, so called because they mature in the thymus.   2. Humoral immunity {humor is an archaic term for fluid), which is most  effective against bacterial infections and the extracellular phases of viral  infections, is mediated by an enormously diverse collection of related  proteins known as antibodies or immunoglobulins. Antibodies are  produced by B lymphocytes or B cells, which in mammals mature in  the bone marrow.   In this section we focus on the structure, function, and generation of antibodies.   The immune response is triggered by the presence of a foreign macromolecule,  often a protein or carbohydrate, known as an antigen . B cells display immunoglob¬  ulins on their surfaces. If a B cell encounters an antigen that binds to its par¬  ticular immunoglobulin, it engulfs the antigen-antibody complex, degrades it,  and displays the antigen fragments on the cell surface. T cells then stimulate the  B cell to proliferate. Most of the B cell progeny are circulating cells that secrete  large amounts of the antigen-specific antibody. These antibodies can bind to ad¬  ditional antigen molecules, thereby marking them for destruction by other com¬  ponents of the immune system. Although most B cells live only a few weeks  unless stimulated by their corresponding antigen, a few long-lived memory B  cells can recognize antigen several months or even many years later and can  mount a more rapid and massive immune response (called a secondary response)  than B cells that have not yet encountered their antigen (Fig. 7-37).   The immunoglobulins form a related but enormously diverse group of  proteins. All immunoglobulins contain at least four subunits: two identical    FIG. 7-37 Primary and secondary  immune responses. Antibodies to antigen A  appear in the blood following primary  immunization on day 0 and secondary  immunization on day 28. Antigen B is included in  the secondary immunization to demonstrate the  specificity of immunological memory for antigen  A. The secondary response to antigen A is both  faster and greater than the primary response.   ~23-kD light chains (L) and two identical 53- to 75-kD heavy chains Section 3 Antibodies   (H). These subunits associate by disulfide bonds and by noncovalent inter¬  actions to form a roughly Y-shaped symmetric molecule with the formula  (LH) 2 (Fig. 7-38).   The five classes of immunoglobulin (Ig) differ in the type of heavy chain  they contain and, in some cases, in their subunit structure (Table 7-2). For  example, IgM consists of five Y-shaped molecules arranged around a central  J subunit; IgA occurs as monomers, dimers, trimers, and tetramers. The var¬  ious immunoglobulin classes also have different physiological functions. IgM  is most effective against microorganisms and is the first immunoglobulin to  be secreted in response to an antigen. IgG, the most common immunoglob¬  ulin, is equally distributed between the blood and the extravascular fluid.   IgA occurs predominantly in the intestinal tract and defends against  pathogens by adhering to their antigenic sites so as to block their attach¬  ment to epithelial (outer) surfaces. IgE, which is normally present in the  blood in minute concentrations, protects against parasites and has been im¬  plicated in allergic reactions. IgD, which is also present in small amounts,  has no clearly known function. Our discussion of antibody structure will fo¬  cus on IgG.   IgG can be cleaved through limited proteolysis with the enzyme papain into  three ~50-kD fragments: two identical Fab fragments and one Fc fragment.   The protein is shown in ribbon form with its two  heavy chains gold and cyan and its two light  chains both magenta. Its two identical  carbohydrate chains are drawn in space-filling  form with C green, N blue, and 0 red. The  antigen-binding sites are located at the ends of  the two approximately horizontal Fab arms  formed by the association of the light chains  with the heavy chains. This particular antibody  recognizes canine lymphoma (a type of cancer)  and is therapeutically useful against it. [Based on  an X-ray structure by Alexander McPherson,  University of California at Irvine. PDBid  11GT.] © See Interactive Exercise 3.   (0i 2 K 2 )n] a Or (a 2 \2)J a   (ja 2 k 2 ) 5 J or (ja 2 X 2 ) 5 J   ^gG has four subclasses, IgGl, IgG 2 , IgG3, and IgG4, which differ in their 7 chains.   Chapter 7 Protein Function: Myoglobin and  Hemoglobin, Muscle Contraction, and Antibodies    FIG. 7-39 Diagram of human  immunoglobulin G (IgG). Each light chain  contains a variable (V L ) and a constant (C L )  region, and each heavy chain contains one  variable (V H ) and three constant (C H 1, C H 2, and  C h 3) regions. Each of the variable and constant  domains contains a disulfide bond, and the four  polypeptide chains are linked by disulfide bonds.  Hypervariable loops, three in each variable  domain (fuzzy lines), determine antigen specificity.  The proteolytic enzyme papain cleaves IgG at  the hinge region to yield two Fab fragments and  one Fc fragment. CHO represents carbohydrate  chains. [Illustration, Irving Geis. Image from Irving  Geis Collection/Howard Hughes Medical  Institute. Rights owned by HHMI. Reproduction  by permission only.]   tire L chain and the N-terminal half of an H chain (Fig. 7-39). These fragments  contain IgGs antigen-binding sites (the “ab” in Fab stands for antigen binding).  The Fc portion (“c” because it crystallizes easily) derives from the “stem” of the  antibody and consists of the C-terminal halves of two H chains. The arms of the  Y are connected to the stem by a flexible hinge region. The hinge angles may  vary, so an antibody molecule may not be perfectly symmetrical (e.g., Fig. 7-38).   Although all IgG molecules have the same overall structure, IgGs that rec¬  ognize different antigens have different amino acid sequences. The light chains  of different antibodies differ mostly in their N-terminal halves. These polypep¬  tides are therefore said to have a variable region, V L (residues 1 to —108), and  a constant region, C L (residues 109 to 214). Comparisons of H chains, which  have —446 residues, reveal that H chains also have a variable region, V H , and  a constant region, C H . As indicated in Fig. 7-39, the C H region consists of  three —110-residue segments, C H 1, C H 2, and C H 3, which are homologous to  each other and to C L . In fact, all the constant and variable regions resemble  each other in sequence and in disulfide-bonding pattern. These similarities sug¬  gest that the six different homology units of an IgG evolved through the du¬  plication of a primordial gene encoding an —110-residue protein.   The immunoglobulin homology units all have the same characteristic  immunoglobulin fold: a sandwich composed of three- and four-stranded an¬  tiparallel (3 sheets that are linked by a disulfide bond (Fig. 6-29 b). Nevertheless,  the basic immunoglobulin structure must accommodate an enormous variety of  antigens. The ability to recognize antigens resides in three loops in each variable  domain (Fig. 7-40). Most of the amino acid variation among antibodies is con¬  centrated in these three short segments, called hypervariable sequences. As hy¬  pothesized by Elvin Kabat, the hypervariable sequences line an immunoglobulins  antigen-binding site, so that their amino acids determine its binding specificity.   Scientists have determined the X-ray structures of Fab fragments from  monoclonal antibodies (Box 7-5) and monospecific antibodies isolated from    FIG. 7-40 Immunoglobulin folds in a light  chain. Both the constant and variable domains  consist of a sandwich of a four-stranded  antiparallel p sheet (blue) and a three-stranded  antiparallel p sheet ( orange ) that are linked by a  disulfide bond (yellow). The positions of the  three hypervariable sequences in the variable  domain are indicated. [After Schiffer, M., Girling,  R.L., Ely, K.R., and Edmundson, A.B., Biochemistry  12 , 4628 (1973).]    patients with multiple myeloma (a disease in which a cancerous B cell pro¬  liferates and produces massive amounts of a single immunoglobulin;  immunoglobulins purified from ordinary blood are heterogeneous and hence  cannot be used for detailed structural studies). As predicted by the positions  of the hypervariable sequences, the antigen-binding site is located at the tip  of each Fab fragment in a crevice between its V L and V H domains.   The association between antibodies and their antigens involves van der  Waals, hydrophobic, hydrogen bonding, and ionic interactions. Their dissoci¬  ation constants range from 10 4 to 10 10 M, comparable (or even greater)  in strength to the associations between enzymes and their substrates. The speci¬  ficity and strength of an antigen—antibody complex are a function of the  exquisite structural complementarity between the antigen and the antibody  (e.g., Fig. 7-41). These are also the features that make antibodies such useful  laboratory reagents (Fig. 5-3, for example).   Most immunoglobulins are divalent molecules; that is, they can bind two  identical antigens simultaneously (IgM and IgA are multivalent). A foreign  substance or organism usually has multiple antigenic regions, and a typical im¬  mune response generates a mixture of antibodies with different specificities.  Divalent binding allows antibodies to cross-link antigens to form an extended  lattice (Fig. 7-42), which hastens the removal of the antigen and triggers B  cell proliferation.   novel antigen does not direct a B cell to begin manufacturing a new im¬  munoglobulin to which it can bind. Rather, an antigen stimulates the prolifer¬  ation of a preexisting B cell whose antibodies happen to recognize the antigen. The  immune system has the potential to produce an enormous number of differ¬  ent antibodies, probably >10 18 . Even though this number is so large that an  individual can synthesize only a small fraction of its potential immunoglobu¬  lin repertoire during its lifetime, this fraction is still sufficient to react with  almost any antigen the individual might encounter. Yet the number of  immunoglobulin genes is far too small to account for the observed level of an¬  tibody diversity. The diversity in antibody sequences arises instead from ge¬  netic changes during B lymphocyte development.   Each chain of an immunoglobulin molecule is encoded by multiple DNA  segments: V, J, and C segments for the light chains, and V, D, J, and C seg¬  ments for heavy chains. These segments are joined together by somatic   FIG. 7-41 Interaction between an antigen  and an antibody. This X-ray structure shows a  portion of the solvent-accessible surface of a  monoclonal antibody Fab fragment (green) with a  stick model of a bound nine-residue fragment of  its peptide antigen (lavender). [Courtesy of Ian  Wilson, The Scripps Research Institute, La Jolla,  California. PDBid 1HMMJ    FIG. 7-42 Antigen cross-linking by antibodies. A mixture of divalent antibodies that  recognizes the several different antigenic regions of an intruding particle such as a toxin  molecule or a bacterium can form an extensive lattice of antigen and antibody molecules.    Introducing a foreign molecule into an animal induces the synthe¬  sis of large amounts of antigen-specific but heterogeneous anti¬  bodies. One might expect that a single lymphocyte from such an  animal could be cloned (allowed to reproduce) to yield a harvest  of homogeneous immunoglobulin molecules. Unfortunately, lym¬  phocytes do not grow continuously in culture. In the late 1970s,  however, Cesar Milstein and Georges Kohler developed a tech¬  nique for immortalizing such cells so that they can grow continu¬  ously and secrete virtually unlimited quantities of a specific  antibody. Typically, lymphocytes from a mouse that has been im¬  munized with a particular antigen are harvested and fused with  mouse myeloma cells (a type of blood system cancer), which can  multiply indefinitely (see figure). The cells are then incubated in a  selective medium that inhibits the synthesis of purines, which are  essential for myeloma growth [the myeloma cells lack the enzyme  hypoxanthine phosphoribosyl transferase (HPRT), which could  otherwise participate in a purine nucleotide salvage pathway;  Section 23-ID]. The only cells that can grow in the selective  medium are fused cells, known as hybridoma cells, that combine  the missing HPRT (it is supplied by the lymphocyte) with the  immortal attributes of the myeloma cells. Clones derived from  single fused cells are then screened for the presence of antibod¬  ies to the original antigen. Antibody-producing cells can be grown  in large quantities in tissue culture or as semisolid tumors in  mouse hosts.   Monoclonal antibodies are used to purify macromolecules  (Section 5-2), to identify infectious diseases, and to test for the  presence of drugs and other substances in body tissues. Because  of their purity and specificity and, to some extent, their biocompat¬  ibility, monoclonal antibodies also hold considerable promise as  therapeutic agents against cancer and other diseases. In fact, the  monoclonal antibody known as Herceptin binds specifically to  the growth factor receptor HER2 that is overexpressed in about  one-quarter of breast cancers. Herceptin binding to HER2 blocks  its growth-signaling activity, thereby causing the tumor to stop  growing or even regress.   Other monoclonal antibodies have been developed to bind to  and interfere with components of the inflammatory response in  order to treat the symptoms of diseases such as rheumatoid  arthritis. In some cases, these “biologic” drugs have been geneti¬  cally engineered to make them less mouselike and more human¬  like and therefore less likely to be recognized and rejected as  foreign by the patient’s own immune system.    recombination during B cell development before being transcribed and trans¬  lated into protein. The process is called somatic (Greek: soma ., body) to dis¬  tinguish it from the recombination that occurs in reproductive cells. Because  there are multiple versions of the V, D, J, and C segments in the genome, the  combinatorial possibilities are enormous. In addition, the recombination  process sometimes adds or deletes nucleotides at the junctions between gene  segments, further contributing to the diversity of the encoded protein. The  generation of antibody diversity is further discussed in Section 28-3D.   Additional changes can occur after a B cell has encountered its antigen and  begun secreting antibody molecules. As the antibody-producing B cells divide,  their rate of immunoglobulin gene mutation increases dramatically, favoring the  substitution of one nucleotide for another and leading to an average of one   Low blood glucose, muscle weakness, Na + loss,  K + retention, increased susceptibility to stress   Oversecretion of thyroid hormone resulting in  increased appetite accompanied by weight loss    amino acid change for every cell generation. This process, which is called  somatic hypermutation, permits the antigen specificity of the antibody to be  fine-tuned over many cell generations, because the rate of B cell proliferation  increases with the antigen-binding affinity of the antibody it produces.   Another remarkable property of the immune system is that its power is un¬  leashed only against foreign substances and not against any of the tens of thou¬  sands of endogenous (self) molecules of various sorts. Virtually all  macromolecules are potentially antigenic, as can be demonstrated by trans¬  planting tissues from one individual to another, even within a species. This  incompatibility presents obvious challenges for therapies ranging from routine  blood transfusions to organ transplants.   The mechanism whereby an individuals immune system distinguishes self  from non-self is not completely understood but includes the elimination of self¬  reactive B and T cells before they are fully mature. Self-tolerance begins to de¬  velop around the time of birth and must be ongoing, since new lymphocytes  arise throughout an individuals lifetime. Occasionally, the immune system loses  tolerance to some of its self-antigens, resulting in an autoimmune disease.   All the body’s organ systems are theoretically susceptible to attack by an  immune system that has lost its self-tolerance, but some tissues are attacked  more often than others. Some of the most common antibody-mediated  autoimmune diseases are listed in Table 7-3 (other autoimmune diseases re¬  sult mainly from inappropriate T cell activation). The symptoms of a particu¬  lar disease reflect the type of tissue with which the autoantibodies react or, in  the case of systemic diseases, the accumulation of antigen—antibody complexes  in multiple locations. In general, autoimmune diseases are chronic, often with  periods of remission, and their clinical severity may differ among individuals.   The loss of tolerance to ones own antigens may result from an innate mal¬  functioning of the mechanism by which the immune system distinguishes self  from non-self, possibly precipitated by an event, such as trauma or infection,  in which tissues that are normally sequestered from the immune system are  exposed to lymphocytes. For example, breaching the blood—brain barrier may  allow lymphocytes access to the brain or spinal cord, and injury may allow ac¬  cess to the spaces at joints, which are not normally served by blood vessels.  There is also evidence that some autoimmune diseases are caused by antibod¬  ies to certain viral or bacterial antigens that cross-react with endogenous sub¬  stances because of chance antigenic similarities. Some diseases, such as systemic  lupus erythematosus, represent a more generalized breakdown of the immune  system, so that antibodies to many endogenous substances (e.g., DNA and  phospholipids) may be generated.   • Without looking at Fig. 7-39, draw a diagram  of an IgG molecule and identify the heavy  and light chains, the constant and variable  domains, and the antigen-binding site(s).   • Explain how a single human can poten¬  tially generate billions of different antibody  molecules.   • What are some factors that might con¬  tribute to the development of an auto¬  immune disease?   • Why do the symptoms of an autoimmune  disease often differ between individuals?   • Myoglobin, a monomeric heme-containing muscle protein,  reversibly binds a single 0 2 molecule.   • Hemoglobin, a tetramer with pseudo-Z) 2 symmetry, has distinctly  different conformations in its oxy and deoxy states.   • Oxygen binds to hemoglobin in a sigmoidal fashion, indicating  cooperative binding.   • 0 2 binding to a heme group induces a conformational change in  the entire hemoglobin molecule that includes movements at the  subunit interfaces and the disruption of ion pairs. The result is a  shift from the T to the R state.   • C0 2 promotes 0 2 dissociation from hemoglobin through the  Bohr effect. BPG decreases hemoglobin’s 0 2 affinity by binding  to deoxyhemoglobin.   • The symmetry and sequential models of allosterism explain how  binding of a ligand at one site affects binding of another ligand  at a different site.   • Hemoglobin variants have revealed structure-function relation¬  ships. Hemoglobin S produces the symptoms of sickle-cell anemia  by forming rigid fibers in its deoxy form.   • The thick filaments of a sarcomere are composed of the motor pro¬  tein myosin and the thin filaments are composed mainly of actin.   • The heads of myosin molecules in thick filaments form bridges to  actin in thin filaments such that the detachment and reattachment  of the myosin heads cause the thick and thin filaments to slide  past each other during muscle contraction. The contractile force  derives from conformational changes in myosin that are triggered  by ATP hydrolysis.   • In nonmuscle cells, actin forms microfilaments, which are com¬  ponents of the cytoskeleton. Microfilaments are dynamic struc¬  tures whose growth and regression are responsible for certain types  of cell movement.   • The immune system responds to foreign macromolecules through  the production of antibodies (immunoglobulins).   • The Y-shaped IgG molecule consists of two heavy and two light  chains. The two antigen-binding sites are formed by the hyper¬  variable sequences in the variable domains at the ends of a heavy  and a light chain.   • Antibody diversity results from somatic recombination during B  cell development and from somatic hypermutation.    myofibril 197  sarcomere 197  thick filament 197  thin filament 197  sliding filament model 198   end 202   end 202  motor protein 204  microfilament 206  treadmilling 207  pathogen 208  immune system 208  cellular immunity 208  lymphocyte 208  humoral immunity 208  immunoglobulin (Ig) 208    1. Estimate K from the following data describing ligand binding to  a protein.    2. Which set of binding data is likely to represent cooperative  ligand binding to an oligomeric protein?    3. In active muscles, the p0 2 may be 10 torr at the cell surface and  1 torr at the mitochondria (the organelles where oxidative  metabolism occurs). Use Eq. 7-6 to show how myoglobin  (/? 50 = 2.8 torr) facilitates the diffusion of 0 2 through these cells.   4. If myoglobin had the same p 50 value as hemoglobin, how well  would it facilitate 0 2 diffusion under the conditions described in  Problem 3?   3. Use Eq. 7-8 to estimate the fractional saturation of hemoglobin  when p0 2 is (a) 20 torr, (b) 40 torr, and (c) 60 torr.   6. Calculate the y? 50 value for hemoglobin if Yq 2 = 0.82 when  p0 2 = 23 torr.   7. Drinking a few drops of a commercial preparation called  “vitamin O,” which consists of oxygen and sodium chloride dis¬  solved in water, is claimed to increase the concentration of oxy¬  gen in the body, (a) Use your knowledge of oxygen transport to  evaluate this claim, (b) Would vitamin O be more or less effec¬  tive if it were infused directly into the bloodstream?   8. Explain why long-distance runners prefer to train at high altitude  even when the race is to be held at sea level. Why must the run¬  ners spend more than a day or two at the higher elevation?   9. Some primitive animals have a hemoglobin that consists of two  identical subunits. Sketch an oxygen-binding curve for this protein.   10. What is the likely range of the Hill constant for the hemoglobin  described in Problem 9?   11. In humans, the urge to breathe results from high concentrations  of C0 2 in the blood; there are no direct physiological sensors of  blood p0 2 . Skindivers often hyperventilate (breathe rapidly and  deeply for several minutes) just before making a dive in the be¬  lief that this will increase the 0 2 content of their blood, (a) Does  it do so? (b) Use your knowledge of hemoglobin function to eval¬  uate whether this practice is useful.    12. The crocodile, which can remain under water without breathing  for up to 1 h, drowns its air-breathing prey and then dines at its  leisure. An adaptation that aids the crocodile in doing so is that  it can utilize virtually 100% of the 0 2 in its blood whereas hu¬  mans, for example, can extract only ~65% of the 0 2 in their  blood. Crocodile Hb does not bind BPG. However, crocodile  deoxyHb preferentially binds HC0 3 . How does this help the  crocodile obtain its dinner?   13. Is the y> 5 o higher or lower than normal in (a) hemoglobin Yakima  and (b) hemoglobin Kansas? Explain.   14. In hemoglobin Rainier, Tyr 145(3 is replaced by Cys, which forms  a disulfide bond with another Cys residue in the same subunit.  This prevents the formation of ion pairs that normally stabilize  the T state. How does hemoglobin Rainier differ from normal  hemoglobin with respect to (a) oxygen affinity, (b) the Bohr  effect, and (c) the Hill constant?   15. Hemoglobin S homozygotes who are severely anemic often have el¬  evated levels of BPG in their erythrocytes. Is this a beneficial effect?   16. In the variant hemoglobin C, glutamate at position 6 of the  (3 chain has been replaced with lysine, (a) Would you expect this  mutant hemoglobin to polymerize as hemoglobin S does? (b) Red  blood cells containing hemoglobin C have a shorter lifespan than  red blood cells containing normal hemoglobin. How might this  affect a person’s resistance to malaria?   17. Is myosin a fibrous protein or a globular protein? Explain.   18. In striated muscle, cells undergo mitosis (nuclear division) with¬  out cytokinesis (cellular division), giving rise to large multinucle-  ate cells. Explain why muscle cells would be less effective if  cytokinesis occurred with every round of mitosis.   19. A myosin head can undergo five ATP hydrolysis cycles per sec¬  ond, each of which moves an actin monomer by ~100 A. How  is it possible for an entire sarcomere to shorten by 1000 A in this  same period?   20. Rigor mortis, the stiffening of muscles after death, is caused by  depletion of cellular ATP. Describe the molecular basis of rigor.   21. Explain why a microfilament is polar whereas a filament of keratin  is not.   22. Cells contain an assortment of proteins that promote microfila-  ment disassembly during cell shape changes. How can such pro¬  teins distinguish newly synthesized microfilaments from older  microfilaments?   23. Give the approximate molecular masses of an immunoglobulin G  molecule analyzed by (a) gel filtration chromatography, (b) SDS-  PAGE, and (c) SDS-PAGE in the presence of 2-mercaptoethanol.   24. Many fish produce a tetrameric IgM. If each H chain has a mass  of 70 kD, each light chain has a mass of 25 kD, and each J chain  has a mass of 15 kD, what is the approximate mass of the IgM  molecule?   25. Explain why the variation in V L and V H domains of  immunoglobulins is largely confined to the hypervariable loops.   26. How many hypervariable loops are present in (a) IgG and (b) IgM?   27. Why do antibodies raised against a native protein sometimes fail  to bind to the corresponding denatured protein?   28. Antibodies raised against a macromolecular antigen usually produce  an antigen-antibody precipitate when mixed with that antigen.  Explain why no precipitate forms when (a) Fab fragments from    those antibodies are mixed with the antigen; (b) antibodies raised  against a small antigen are mixed with that small antigen; and (c)  the antibody is in great excess over the antigen and vice versa.   29. Some bacteria produce proteases that can cleave the hinge region  of IgA molecules without affecting antigen binding. Explain why  these proteases would give the bacteria a better chance of start¬  ing an infection.   30. Individuals with systemic lupus erythematosus (SLE) produce an¬  tibodies to DNA and phospholipids, (a) Explain why normal in¬  dividuals do not make antibodies to these substances, (b) During  a normal response to a viral or bacterial infection, the immune sys¬  tem produces large amounts of antigen-specific antibodies, and the  resulting antigen-antibody complexes are subsequently removed  from the circulation and degraded. Explain why antigen-antibody  complexes accumulate in the tissues of individuals with SLE.   Case 8 Hemoglobin, the Oxygen Carrier  Focus concept: A mutation in the gene for hemoglobin results in  an altered protein responsible for the disease sickle-cell anemia.  An understanding of the biochemistry of the disease may suggest  possible treatments.   Focus concept: The effect of allosteric modulators on oxygen affin¬  ity for crocodile hemoglobin differs from that of other species.   Case 10 The Biological Roles of Nitric Oxide  Focus concept: Nitric oxide, a small lipophilic molecule, acts as a  second messenger in blood vessels.    (a) Look up information about  hemoglobin variants. Why don’t they generate the same symp¬  toms? How do the symptoms of thalassemias differ? Which hemo¬  globin variants appear to offer a selective advantage under certain  conditions? (b) In addition to myosin, which interacts with actin fil¬  aments, cells contain several other motor protein systems. Describe  the structure and activity of the motor proteins kinesin and dynein.  Against what fibrous proteins do they exert force? How do these  systems differ from the actin-myosin system? (c) Explain how im¬  munological memory is exploited in the development of vaccines  for viral and bacterial infections. What factors make a vaccine most  effective in preventing disease?   Ackers, G.K. and Holt, J.M., Asymmetric cooperativity in a symmetric  tetramer: human hemoglobin, J. Biol. Chem. 281, 11441-11443  (2006). [A brief review of hemoglobins allosteric behavior.]   Allison, A.C., The discovery of resistance to malaria of sickle-cell het¬  erozygotes, Biochem. Mol. Biol. Educ. 30, 279-287 (2002).  Dickerson, R.E. and Geis, I., Hemoglobin, Benjamin/Cummings (1983).  [A beautifully written and lavishly illustrated treatise on the struc¬  ture, function, and evolution of hemoglobin.]   Ferry, G., Max Perutz and the Secret of Life, Cold Spring Harbor Labora¬  tory Press (2007). [A definitive biography.]   Hsia, C.C.W., Respiratory function of hemoglobin, New Engl. J. Med.  338, 239-247 (1998). [A short review of hemoglobin’s physiologi¬  cal role.]   Judson, H.F., The Eighth Day of Creation (Expanded edition), Chapters  9 and 10, Cold Spring Harbor Laboratory Press (1996). [Includes a  fascinating historical account of how our present perception of he¬  moglobin structure and function came about.]   Nagel, R.L., Haemoglobinopathies due to structural mutations, in  Provan, D. and Gribben, J. (Eds.), Molecular Haematology, pp.  121-133, Blackwell Science (2000).   Ordway, G.A. and Garry, D.J., Myoglobin: an essential hemoprotein in  striated muscle, J. Exp. Biol. 20 7, 341-3446 (2004).   Perutz, M.F., Wilkinson, A.J., Paoli, M., and Dodson, G.G., The stereo¬  chemical mechanism of the cooperative effects in hemoglobin revis¬  ited, Annu. Rev. Biophys. Biomol. Struct. 27, 1-34 (1998).   Schechter, A.N., Hemoglobin research and the origins of molecular med¬  icine, Blood 112, 3927-3938 (2008). [Reviews hemoglobin struc¬  ture and function, including hemoglobin S.]   Strasser, B.J., Sickle-cell anemia, a molecular disease, Science 286,  1488-1490 (1999). [A short history of Pauling’s characterization of  sickle-cell anemia.]   Cooper, J.A. and Schafer, D.A., Control of actin assembly and disassem¬  bly at filament ends, Curr. Opin. Cell Biol. 12, 97-103 (2000). [Pro¬  vides an overview of the principles of microfilament dynamics and  some of the key protein players.]   Craig, R. and Woodhead, J.L., Structure and function of myosin fila¬  ments, Curr. Opin. Struct. Biol. 16, 204-212 (2006). [Includes de¬  tails of myosin and thick filament structure, including its  arrangement in the sarcomere.]   Dominguez, R. and Holmes, K.C., Actin structure and function, Annu.  Rev. Biophys. 40, 169-186 (2011).   Reisler, E. and Egelman, E.H., Actin structure and function: what we still  do not understand,/. Biol. Chem. 282, 36133-36137 (2007). [Sum¬  marizes some areas of research on actin structure and its assembly  into filaments.]   Schliwa, M. and Woehlke, G., Molecular motors, Nature 422, 739-763  (2003). [Includes reviews of myosin and other motor proteins.]   Spudich, J.A., The myosin swinging cross-bridge model, Nature Rev. Mol.  Cell Biol. 2, 387-391 (2001). [Summarizes the history and current  state of models for myosin action.]   Davies, D.R. and Cohen, G.H., Interactions of protein antigens with an¬  tibodies, Proc. Natl. Acad. Sci. 93, 7-12 (1996).   Harris, L.J., Larson, S.B., Hasel, K.W., Day, J., Greenwood, A., and  McPherson, A., The three-dimensional structure of an intact mon¬  oclonal antibody for canine lymphoma, Nature 360, 369-372  (1992). [The first high-resolution X-ray structure of an intact IgG.]   Marrack, P., Kappler, J., and Kotzin, B.L., Autoimmune disease: why and  where it occurs, Nature Med. 7, 899-905 (2001).   Approximately half the dry mass of corn (maize) kernels is starch, a polysaccharide  that is broken down to glucose during digestion. Starch, along with the indigestible  polysaccharide cellulose, is also the starting material for production of the biofuel  ethanol. [© Sharon Dominick/iStockphoto]   A Monosaccharides Are Aldoses or Ketoses  B Monosaccharides Vary in Configuration and  Conformation   A Lactose and Sucrose Are Disaccharides  B Cellulose and Chitin Are Structural  Polysaccharides   A Proteoglycans Contain Glycosaminoglycans  B Bacterial Cell Walls Are Made of Peptidoglycan  C Many Eukaryotic Proteins Are Glycosylated  D Oligosaccharides May Determine Glycoprotein  Structure, Function, and Recognition    Carbohydrates or saccharides (Greek: sakcharon, sugar) are the most abun¬  dant biological molecules. They are chemically simpler than nucleotides or  amino acids, containing just three elements—carbon, hydrogen, and  oxygen—combined according to the formula (C • H 2 0)„, where n > 3. The  basic carbohydrate units are called monosaccharides. There are numerous dif¬  ferent types of monosaccharides, which, as we discuss below, differ in their  number of carbon atoms and in the arrangement of the H and O atoms at¬  tached to the carbons. Furthermore, monosaccharides can be strung together  in almost limitless ways to form polysaccharides.   Until the 1960s, carbohydrates were thought to have only passive roles as  energy sources (e.g., glucose and starch) and as structural materials (e.g., cellu¬  lose). Carbohydrates, as we will see, do not catalyze complex chemical reactions  as do proteins, nor do carbohydrates replicate themselves as do nucleic acids.  And because polysaccharides are not built according to a genetic “blueprint,” as  are nucleic acids and proteins, they tend to be much more heterogeneous—both  in size and in composition—than other biological molecules.   However, it has become clear that the innate structural variation in car¬  bohydrates is fundamental to their biological activity. The apparently haphaz¬  ard arrangements of carbohydrates on proteins and on the surfaces of cells are  the key to many recognition events between proteins and between cells. An  understanding of carbohydrate structure, from the simplest monosaccharides  to the most complex branched polysaccharides, is essential for appreciating the  varied functions of carbohydrates in biological systems.    • The smallest sugars are aldoses or ketoses with the formula (C • H 2 0) n .   • The derivatives of monosaccharides include the aldonic acids, uronic acids,  alditols, deoxy sugars, and amino sugars.   • Monosaccharides can be linked to each other or to other molecules by glycosidic  bonds.   • Monosaccharides, or simple sugars, are synthesized from smaller precursors that  are ultimately derived from C0 2 and H 2 0 by photosynthesis.   Monosaccharides are aldehyde or ketone derivatives of straight-chain poly hydroxy  alcohols containing at least three carbon atoms. They are classified according to  the chemical nature of their carbonyl group and the number of their C atoms.  If the carbonyl group is an aldehyde, the sugar is an aldose. If the carbonyl    FIG. 8-1 The D-aldoses with three to six carbon atoms. The   arrows indicate stereochemical relationships (not biosynthetic  pathways). The configuration around C2 (red) distinguishes the    members of each pair of monosaccharides. The l counterparts of  these 15 sugars are their mirror images. The biologically most  common aldoses are boxed.    group is a ketone, the sugar is a ketose. The smallest monosaccharides, those  with three carbon atoms, are trioses. Those with four, five, six, seven, etc.  C atoms are, respectively, tetroses, pentoses, hexoses, heptoses, etc.    All but two of its six C atoms, Cl and C6, are chiral centers, so D-glu¬  cose is one of 2 4 = 16 possible stereoisomers. The stereochemistry and  nomenclature of the D-aldoses are presented in Fig. 8-1. The assignment  of D or L is made according to the Fischer convention (Section 4-2): D  sugars have the same absolute configuration at the asymmetric center farthest  from their carbonyl group as does D-glyceraldehyde (i.e., the —OH at C5 of  D-glucose is on the right in a Fischer projection). The L sugars are the mir¬  ror images of their D counterparts. Because L sugars are biologically much  less abundant than D sugars, the D prefix is often omitted.   Sugars that differ only by the configuration around one C atom  are known as epimers of one another. Thus, D-glucose and D-man-  nose are epimers with respect to C2. The most common aldoses in¬  clude the six-carbon sugars glucose, mannose, and galactose. The  pentose ribose is a component of the ribonucleotide residues of  RNA. The triose glyceraldehyde occurs in several metabolic path¬  ways.   The most common ketoses are those with their ketone function  at C2 (Fig. 8-2). The position of their carbonyl group gives ketoses one  less asymmetric center than their isomeric aldoses, so a ketohexose has  only 2 3 = 8 possible stereoisomers (4 D sugars and 4 L sugars).   The most common ketoses are dihydroxyacetone, ribulose, and  fructose, which we will encounter in our studies of metabolism.   Alcohols react with the carbonyl groups of aldehydes and ketones to form    FIG. 8-2 The D-ketoses with three to six  carbon atoms. The configuration around C3  (red) distinguishes the members of each pair.  The biologically most common ketoses are  boxed.    The hydroxyl and either the aldehyde or the ketone functions of monosaccha¬  rides can likewise react intramolecularly to form cyclic hemiacetals and   FIG. 8-3 Cyclization of glucose and fructose, (a) The linear form of D-glucose  yielding the cyclic hemiacetal p-D-glucopyranose. ( b ) The linear form of D-fructose yielding  the hemiketal p-D-fructofuranose. The cyclic sugars are shown as both Haworth projections  and in stick form embedded in their semitransparent space-filling models with C green, H  white, and 0 red.    In a Haworth projection, which hydroxyl groups correspond to the hydroxyl  groups on the left in a Fischer projection?    hemiketals (Fig. 8 -3). The configurations of the substituents of each carbon  atom in these sugar rings are conveniently represented by their Haworth pro¬  jections, in which the heavier ring bonds project in front of the plane of the  paper and the lighter ring bonds project behind it.   A sugar with a six-membered ring is known as a pyranose in analogy with  pyran (at left), the simplest compound containing such a ring. Similarly sug¬  ars with five-membered rings are designated furanoses in analogy with furan  (at left). The cyclic forms of glucose and fructose with six- and five-membered  rings are therefore known as glucopyranose and fructofuranose, respectively.   Cyclic Sugars Have Two Anomeric Forms. When a monosaccharide cy-  clizes, the carbonyl carbon, called the anomeric carbon, becomes a chiral  center with two possible configurations. The pair of stereoisomers that dif¬  fer in configuration at the anomeric carbon are called anomers. In the  Oi anomer, the OH substituent of the anomeric carbon is on the opposite  side of the sugar ring from the CH 2 OH group at the chiral center that desig¬  nates the D or L configuration (C5 in hexoses). The other form is known as  the P anomer (Fig. 8-4).   The two anomers of D-glucose have slightly different physical and chem¬  ical properties, including different optical rotations (Section 4-2). The anomers  freely interconvert in aqueous solution ., so at equilibrium, D-glucose is a mixture  of the P anomer (63.6%) and the a anomer (36.4%). The linear form is nor¬  mally present in only minute amounts.   Sugars Can Adopt Different Conformations. A given hexose or pentose can  assume pyranose or furanose forms. In principle, hexoses and larger sugars can   FIG. 8-4 ol and p anomers. The monosaccharides a-D-glucopyranose and  p-D-glucopyranose, drawn as Haworth projections and ball-and-stick models, interconvert  through the linear form. They differ only by their configuration about the anomeric carbon, Cl.    form rings of seven or more atoms, but such rings are rarely observed because  of the greater stabilities of the five- and six-membered rings. The internal strain  of three- and four-membered rings makes them less stable than the linear forms.   The use of Haworth formulas may lead to the erroneous impression that  furanose and pyranose rings are planar. This cannot be the case, however, be¬  cause all the atomic orbitals in the ring atoms are tetrahedrally ( sp 3 ) hybridized.   The pyranose ring, like the cyclohexane ring, can assume a chair conforma¬  tion, in which the substituents of each atom are arranged tetrahedrally. Of the  two possible chair conformations, the one that predominates is the one in  which the bulkiest ring substituents occupy equatorial positions rather than  the more crowded axial positions (Fig. 8-5). Only (3-D-glucose can simulta¬  neously have all five of its non-H substituents in equatorial positions. Perhaps  this is why glucose is the most abundant monosaccharide in nature.   Furanose rings can also adopt different conformations, whose stabilities  depend on the arrangements of bulky substituents. Note that a monosaccha¬  ride can readily shift its conformation ,, because no bonds are broken in the  process. The shift in configuration between the a and p anomeric forms or be¬  tween the pyranose and furanose forms, which requires breaking and re-forming  bonds, occurs slowly in aqueous solution. Other changes in configuration,  such as epimerization, do not occur under physiological conditions without  the appropriate enzyme.    Because the cyclic and linear forms of aldoses and ketoses do interconvert, HO —C—H   1. Oxidation of an aldose converts its aldehyde group to a carboxylic acid H—C—OH   group, thereby yielding an aldonic acid such as gluconic acid (at 6 I   right). Aldonic acids are named by appending the suffix -onic acid to 2   FIG. 8-5 The two chair conformations of p-D-glucopyranose. In the conformation  on the left, which predominates, the relatively bulky OH and CH 2 OH substituents all occupy  equatorial positions, where they extend alternately above and below the ring. In the confor¬  mation on the right (drawn in ball-and-stick form in Fig. 8-4, right), the bulky groups occupy  the more crowded axial (vertical) positions. O See Kinemage Exercise 7-1.    2. Oxidation of the primary alcohol group of aldoses yields uronic acids,  which are named by appending -uronic acid to the root name of the  parent aldose, for example, D-glucuronic acid (at left). Uronic acids  can assume the pyranose, furanose, and linear forms.   3. Aldoses and ketoses can be reduced under mild conditions, for exam¬  ple, by treatment with NaBH 4 , to yield polyhydroxy alcohols known  as alditols, which are named by appending the suffix -itol to the root  name of the parent aldose. Ribitol is a component of flavin coenzymes  (Fig. 14-13), and glycerol and the cyclic polyhydroxy alcohol myo¬  inositol are important lipid components (Section 9-1). Xylitol is a  sweetener that is used in “sugarless” gum and candies:    4. Monosaccharide units in which an OH group is replaced by H are  known as deoxy sugars. The biologically most important of these is  [J-D-2-deoxyribose, the sugar component of DNAs sugar—phosphate  backbone (Section 3-2B). L-Fucose is one of the few L sugar compo¬  nents of polysaccharides.   5. In amino sugars, one or more OH groups have been replaced by an  amino group, which is often acetylated. D-Glucosamine and  D-galactosamine (at left) are the most common. AUVcetylneuraminic  acid, which is derived from A^acetylmannosamine and pyruvic acid (Fig.  8-6), is an important constituent of glycoproteins and glycolipids (pro¬  teins and lipids with covalently attached carbohydrate). A^Acetylneuraminic  acid and its derivatives are often referred to as sialic acids.   FIG. 8-6 A/-Acetylneuraminic acid. In the cyclic form of this nine-carbon  monosaccharide, the pyranose ring incorporates the pyruvic acid residue (blue) and  part of the mannose moiety.   FIG. 8-7 Formation of glycosides. The acid-catalyzed condensation of a-D-glucose  with methanol yields an anomeric pair of methyl-D-glucosides.   anomeric group of a sugar can condense with an alcohol to form o i- and  (5-glycosides (Greek: glykys , sweet; Fig. 8-7). The bond connecting the anomeric  carbon to the alcohol oxygen is termed a glycosidic bond. TV-Glycosidic  bonds, which form between the anomeric carbon and an amine, are the bonds  that link D-ribose to purines and pyrimidines in nucleic acids:   Like peptide bonds, glycosidic bonds hydrolyze extremely slowly under  physiological conditions in the absence of appropriate hydrolytic enzymes.  Consequently, an anomeric carbon that is involved in a glycosidic bond can¬  not freely convert between its a and (3 anomeric forms. Saccharides bearing  anomeric carbons that have not formed glycosides are termed reducing sugars,  because the free aldehyde group that is in equilibrium with the cyclic form of  the sugar reduces mild oxidizing agents. Identification of a sugar as nonre¬  ducing is evidence that it is a glycoside.   • Draw a Fischer projection of D-glucose.  Draw two stereoisomers of this molecule,  including one that is an epimer.   • Show how aldoses and ketoses can form  five- and six-membered rings.   • Draw a Haworth projection of D-glucose  and identify it as an a or (3 anomer.   • Explain why anomers of a monosaccharide  can readily interconvert whereas epimers  do not.   • Describe aldonic acids, uronic acids,  alditols, deoxy sugars, and amino sugars.   • Explain why a sugar can form at least two  different glycosides.    • Disaccharides such as lactose and sucrose consist of two sugars linked by  specific glycosidic bonds.   • Cellulose and chitin are polymers of p(1 —>4)-linked glucose residues.   • In starch and glycogen, glucose residues are linked mainly by a(1^4) bonds.   • Glycosaminoglycans and other large heteropolysaccharides typically have a gel¬  like structure.    Polysaccharides, which are also known as glycans, consist of monosaccharides linked  together by glycosidic bonds. They are classified as homopolysaccharides or  heteropolysaccharides if they consist of one type or more than one type of mono¬  saccharide. Although the monosaccharide sequences of heteropolysaccharides can,  in principle, be even more varied than those of proteins, many are composed of  only a few types of monosaccharides that alternate in a repetitive sequence.   Polysaccharides, in contrast to proteins and nucleic acids, form branched as  well as linear polymers. This is because glycosidic linkages can be made to any  of the hydroxyl groups of a monosaccharide. Fortunately for structural bio¬  chemists, most polysaccharides are linear and those that branch do so in only  a few well-defined ways.   A complete description of an oligosaccharide or polysaccharide includes  the identities, anomeric forms, and linkages of all its component monosaccha¬  ride units. Some of this information can be gathered through the use of spe¬  cific exoglycosidases and endoglycosidases, enzymes that hydrolyze  monosaccharide units in much the same way that exopeptidases and endopep-  tidases cleave amino acid residues from polypeptides (Section 5-3B). NMR  measurements are also invaluable in determining both sequences and confor¬  mations of polysaccharides.    Oligosaccharides containing three or more residues are relatively rare, occur¬  ring almost entirely in plants. Disaccharides, the simplest polysaccharides, are  more common. Many occur as the hydrolysis products of larger molecules.  However, two disaccharides are notable in their own right. Lactose (at left),  for example, occurs naturally only in milk, where its concentration ranges from  0 to 7% depending on the species (Box 8-1). The systematic name for lac¬  tose, 0-|3-D-galactopyranosyl-(l—>4)-D-glucopyranose, specifies its monosac¬  charides, their ring types, and how they are linked together. The symbol  (1—>4) combined with the (3 in the prefix indicates that the glycosidic bond  links Cl of the (3 anomer of galactose to 04 of glucose. Note that lactose has  a free anomeric carbon on its glucose residue and is therefore a reducing sugar.    In infants, lactose (also known as milk sugar) is hydrolyzed by the  intestinal enzyme p-D-galactosidase (or lactase) to its compo¬  nent monosaccharides for absorption into the bloodstream. The  galactose is enzymatically converted (epimerized) to glucose,  which is the primary metabolic fuel of many tissues.   Since mammals are unlikely to encounter lactose after they  have been weaned, most adult mammals have low levels of  p-galactosidase. Consequently, much of the lactose they might  ingest moves through their digestive tract to the colon, where  bacterial fermentation generates large quantities of C0 2 , H 2 , and    irritating organic acids. These products cause the embarrassing  and often painful digestive upset known as lactose intolerance.   Lactose intolerance, which was once considered a metabolic  disturbance, is actually the norm in adult humans, particularly  those of African and Asian descent. Interestingly, however,  p-galactosidase levels decrease only mildly with age in descendants  of populations that have historically relied on dairy products for  nutrition throughout life. Modern food technology has come to the  aid of milk lovers who develop lactose intolerance: Milk in which the  lactose has been hydrolyzed enzymatically is widely available.    The most abundant disaccharide is sucrose (at right), the major form in  which carbohydrates are transported in plants. Sucrose is familiar to us as com¬  mon table sugar (see Kinemage Exercise 7-2). The systematic name for su¬  crose, 0-a-D-glucopyranosyl-(l->2)-|3-D-fructofuranoside, indicates that the  anomeric carbon of each sugar (Cl in glucose and C2 in fructose) participates  in the glycosidic bond, and hence sucrose is not a reducing sugar.  Noncarbohydrate molecules that mimic the taste of sucrose are used as sweet¬  ening agents in foods and beverages (Box 8-2).    Artificial sweeteners are added to processed foods and beverages  to impart a sweet taste without adding calories. This is possible  because the compounds mimic sucrose in its interactions with  taste receptors but either are not metabolized or contribute very  little to energy metabolism because they are used at such low  concentrations.   Naturally occurring saccharides, such as fructose, are slightly  sweeter than sucrose. Honey, which contains primarily fructose,  glucose, and maltose (a glucose disaccharide), is about 1.5 times as  sweet as sucrose. How is sweetness measured? There is no substi¬  tute for the human sense of taste, so a panel of individuals sample  solutions of a compound and compare them to a reference solution  containing sucrose. The very sweet compounds listed below must  be diluted significantly before testing in this manner.    One of the oldest artificial sweeteners  is saccharin, discovered in 1879 and  commonly consumed as Sweet’N Low®. In  the 1970s, extremely high doses of sac¬  charin were found to cause cancer in  laboratory rats. Such doses are now con¬  sidered to be so far outside of the range  used for sweetening as to be of insignificant concern to users.   Aspartame, the active ingredient in NutraSweet® and Equal®, was  approved for human use in 1981 and is currently the market leader:    Unlike saccharin, which is not metabolized by the human body,  aspartame is broken down into its components: aspartate  (green), phenylalanine (red), and methanol (blue). The Asp and  Phe, like all amino acids, can be metabolized, so aspartame is  not calorie-free. Methanol in large amounts is toxic; however, the  amount derived from an aspartame-sweetened drink is compara¬  ble to the amount naturally present in the same volume of fruit  juice. Individuals with the genetic disease phenylketonuria, who  are unable to metabolize phenylalanine, are advised to avoid  ingesting excess Phe in the form of aspartame (or any other  polypeptide). The greatest drawback of aspartame may be its  instability to heat, which makes it unsuitable for baking. In  addition, aspartame in soft drinks hydrolyzes over a period of  months and hence loses its flavor.   Acesulfame is sometimes used in combination with aspar¬  tame, since the two compounds act synergistically (i.e., their  sweetness when combined is greater than the sum of their  individual sweetnesses).   Other artificial sweeteners are derivatives of sugars, such as su-  cralose (Splenda®; see Problem 8-12), or of aspartame (e.g.,  alitame). Some plant extracts (e.g., Stevia) are also used as artifi¬  cial sweeteners.   The market for artificial sweeteners is worth several billion  dollars annually. But surprisingly, the most successful sweetening  agents have not been the result of dedicated research efforts.  Instead, they were discovered by chance or mishap. For example,  aspartame was discovered in 1965 by a synthetic chemist who  unknowingly got a small amount of the compound on his fingers  and happened to lick them. Sucralose came to light in 1975 when  a student was asked to “test” a compound and misunderstood the  directions as “taste” the compound.   FIG. 8-8 Electron micrograph of cellulose  fibers. The cellulose fibers in this sample of cell  wall from the alga Chaetomorpha are arranged  in layers. [Biophoto Associates/ Photo  Researchers.]    Plants have rigid cell walls that can withstand osmotic pressure differences be¬  tween the extracellular and intracellular spaces of up to 20 atm. In large plants,  such as trees, the cell walls also have a load-bearing function. Cellulose, the  primary structural component of plant cell walls (Fig. 8-8), accounts for over  half of the carbon in the biosphere: Approximately 10 15 kg of cellulose is es¬  timated to be synthesized and degraded annually.   Cellulose is a linear polymer of up to 15,000 D-glucose residues linked by  (3(1 —>4) glycosidic bonds:   X-Ray and other studies of cellulose fibers reveal that cellulose chains are flat  ribbons in which successive glucose rings are turned over 180° with respect to  each other. This permits the C3—OH group of each glucose residue to form a  hydrogen bond with the ring oxygen (05) of the next residue. Parallel cellulose  chains form sheets with interchain hydrogen bonds, including 02—H---06 and  06—H---03 bonds (Fig. 8-9). Stacks of these sheets are held together by hy¬  drogen bonds and van der Waals interactions. This highly cohesive structure  gives cellulose fibers exceptional strength and makes them water insoluble de¬  spite their hydrophilicity. In plant cell walls, the cellulose fibers are embedded  in and cross-linked by a matrix containing other polysaccharides and lignin, a  plasticlike phenolic polymer. The resulting composite material can withstand  large stresses because the matrix evenly distributes the stresses among the cellu¬  lose reinforcing elements. The difficulty of removing these other substances,    FIG. 8-9 Model of cellulose. Cellulose  fibers consist of —40 parallel, extended glycan  chains. Each of the p(1—»4)-l inked glucose units  in a chain is rotated 180° with respect to its  neighboring residues and is held in this position  by intrachain hydrogen bonds (dashed lines). The  glycan chains line up laterally to form sheets,  and these sheets stack vertically so that they are  staggered by half the length of a glucose unit.  The entire assembly is stabilized by  intermolecular hydrogen bonds. Hydrogen atoms  not participating in hydrogen bonds have been  omitted for clarity. [Illustration, Irving Geis. Image  from the Irving Geis Collection, Howard Hughes  Medical Institute. Reprinted with permission.]    however, is one of the main reasons why the cellulose in wood and agricultural  waste, despite its abundance, cannot be easily converted to biofuels.   Although vertebrates themselves do not possess an enzyme capable of hy¬  drolyzing the (3(1—>4) linkages of cellulose, the digestive tracts of herbivores  contain symbiotic microorganisms that secrete a series of enzymes, collectively  known as cellulases, that do so. The same is true of termites. Nevertheless,  the degradation of cellulose is a slow process because its tightly packed and  hydrogen-bonded glycan chains are not easily accessible to cellulase and do  not separate readily even after many of their glycosidic bonds have been  hydrolyzed. Thus, cows and other ruminants must chew their cud.   Chitin is the principal structural component of the exoskeletons of inverte¬  brates such as crustaceans, insects, and spiders and is also present in the cell walls  of most fungi and many algae. It is therefore almost as abundant as cellulose.  Chitin is a homopolymer of (3(1 —>4)-linked 7V-acetyl-D-glucosamine residues:   It differs chemically from cellulose only in that each C2—OH group is re¬  placed by an acetamido function. X-Ray analysis indicates that chitin and cel¬  lulose have similar structures.   Starch is a mixture of glycans that plants synthesize as their principal energy  reserve. It is deposited in the chloroplasts of plant cells as insoluble granules  composed of ot-amylose and amylopectin. a-Amylose is a linear polymer of  several thousand glucose residues linked by a(l—>4) bonds.    FIG. 8-10 a-Amylose. This regularly repeating polymer forms a left-handed helix.   Note the great differences in structure and properties that result from changing a-amylose’s  a(1—>4) linkages to the p(1 —>4) linkages of cellulose (Fig. 8-9). [Illustration, Irving  Geis/Geis Archives Trust. Copyright Howard Hughes Medical Institute. Reproduced with  permission.]    Note that although a-amylose is an isomer of cellulose, it has very  different structural properties. While celluloses |3-glycosidic link¬  ages cause it to assume a tightly packed, fully extended conforma¬  tion (Fig. 8-9), a-amylose’s a-glycosidic bonds cause it to adopt an  irregularly aggregating helically coiled conformation (Fig. 8-10).   FIG. 8-1 1 Glycogen granules in a liver  cell. In this photomicrograph, glycogen granules  are pink, the greenish objects are mitochondria,  and the yellow object is a fat globule. The  glycogen content of liver may reach 10% of its  net weight. [CNRI/Science Photo Library/Photo  Researchers.]    Amylopectin (at left) consists mainly of a( 1—>4)'-linked glucose residues  but is a branched molecule with a(l—>6) branch points every 24 to 30 glu¬  cose residues on average. Amylopectin molecules contain up to 10 6 glucose  residues, making them some of the largest molecules in nature. The storage of  glucose as starch greatly reduces the large intracellular osmotic pressure that  would result from its storage in monomeric form, because osmotic pressure is  proportional to the number of solute molecules in a given volume (Section 2-  1D). Starch is a reducing sugar, although it has only one residue, called the  reducing end, that lacks a glycosidic bond.   The digestion of starch, the main carbohydrate source in the human diet,  begins in the mouth. Saliva contains an amylase, which randomly hydrolyzes  the a(l—>4) glycosidic bonds of starch. Starch digestion continues in the small  intestine under the influence of pancreatic amylase, which degrades starch to  a mixture of small oligosaccharides. Further hydrolysis by an ot-glucosidase,  which removes one glucose residue at a time, and by a debranching enzyme,  which hydrolyzes a(l—>6) as well as a(l—>4) bonds, produces monosaccha¬  rides that are absorbed by the intestine and transported to the bloodstream.   Glycogen, the storage polysaccharide of animals, is present in all cells but  is most prevalent in skeletal muscle and in liver, where it occurs as cytoplas¬  mic granules (Fig. 8-11). The primary structure of glycogen resembles that of  amylopectin, but glycogen is more highly branched, with branch points oc¬  curring every 8 to 14 glucose residues. In the cell, glycogen is degraded for  metabolic use by glycogen phosphorylase, which phosphorolytically cleaves  glycogens a(l—>4) bonds sequentially inward from its nonreducing ends.  Glycogens highly branched structure , which has many nonreducing ends , permits  the rapid mobilization of glucose in times of metabolic need. The a(l—>6)  branches of glycogen are cleaved by glycogen debranching enzyme (glycogen  breakdown is discussed further in Section 16-1).   The extracellular spaces, particularly those of connective tissues such as cartilage,  tendon, skin, and blood vessel walls, contain collagen (Section 6-1C) and other  proteins embedded in a gel-like matrix that is composed largely of  glycosaminoglycans. These unbranched polysaccharides consist of alternating  uronic acid and hexosamine residues. Solutions of glycosaminoglycans have a  slimy, mucuslike consistency that results from their high viscosity and elasticity.   Hyaluronate Acts as a Shock Absorber and Lubricant. Hyaluronic acid  (hyaluronate) is an important glycosaminoglycan component of connective  tissue, synovial fluid (the fluid that lubricates joints), and the vitreous humor  of the eye. Hyaluronate molecules are composed of 250 to 25,000 (3(1—>4)-  linked disaccharide units that consist of D-glucuronic acid and Af-acetyl-D-  glucosamine (GlcNAc) linked by a (3(1—>3) bond (Fig. 8-12). Hyaluronate  is an extended, rigid molecule whose numerous repelling anionic groups bind  cations and water molecules. In solution, hyaluronate occupies a volume  — 1000 times that in its dry state.   Hyaluronate solutions have a viscosity that is shear dependent (an object  under shear stress has equal and opposite forces applied across its opposite  faces). At low shear rates, hyaluronate molecules form tangled masses that  greatly impede flow; that is, the solution is quite viscous. As the shear stress  increases, the stiff hyaluronate molecules tend to line up with the flow and  thus offer less resistance to it. This viscoelastic behavior makes hyaluronate so¬  lutions excellent biological shock absorbers and lubricants.   Some Glycosaminoglycans Are Sulfated. The other common glycosamino¬  glycans shown in Fig. 8-12 consist of 50 to 1000 sulfated disaccharide units.  Chondroitin-4-sulfate and chondroitin-6-sulfate differ only in the sulfation    FIG. 8-1 2 Repeating disaccharide units of some glycosaminoglycans. The anionic  groups are shown in red and the A/-acetylamido groups are shown in blue. O See    of their acetylgalactosamine (GalNAc) residues. Dermatan sulfate is de¬  rived from chondroitin by enzymatic epimerization of the C5 of glucuronate  residues to form iduronate residues. Keratan sulfate (not to be confused with  the fibrous protein keratin; Section 6-1C) is the most heterogeneous of the  major glycosaminoglycans in that its sulfate content is variable and it con¬  tains small amounts of fucose, mannose, GlcNAc, and sialic acid. Heparin  is also variably sulfated, with an average of 2.5 sulfate residues per disaccha¬  ride unit, which makes it the most highly charged polymer in mammalian  tissues (Fig. 8-13).   In contrast to the other glycosaminoglycans, heparin is not a constituent  of connective tissue but occurs almost exclusively in the intracellular granules  of the mast cells that occur in arterial walls. It inhibits the clotting of blood,  and its release, through injury, is thought to prevent runaway clot formation.  Heparin is therefore in wide clinical use to inhibit blood clotting, for exam¬  ple, in postsurgical patients.   FIG. 8-13 NMR structure of heparin. Three consecutive disaccharide units of this  helical polymer are drawn in stick form. Atoms are colored according to type with  glucosamine C green, iduronate C cyan, H white, N blue, 0 red, and S yellow. The helical  repeat unit is two disaccharides with a pitch (rise per turn) of 17.5 A. Note the high density  of anionic sulfate groups. [Based on an NMR structure by Barbara Mulloy and Mark  Forster, National Institute for Biological Standards and Control, Herts, UK. PDBid 1HPN.]   FIG. 8-14 A Pseudomonas aeruginosa  biofilm. Bacterial colonies growing on the  surface of an agar plate form a biofilm with  complex architecture. [Courtesy of Roberto  Kolter, Harvard Medical School.]   • Describe the monosaccharide units and  their linkages in the common disaccha¬  rides and polysaccharides.   • Explain why the systematic name of an  oligosaccharide must include more than  just the names of the component  monosaccharides.   • Compare and contrast the structures and  functions of cellulose, chitin, starch, and  glycogen.   • How do the physical properties of  glycosaminoglycans and similar molecules  relate to their biological roles?    Heparan sulfate, a ubiquitous cell-surface component as well as an extra¬  cellular substance in blood vessel walls and brain, resembles heparin but has  a far more variable composition with fewer TV- and O-sulfate groups and more  TV-acetyl groups. Heparan sulfate plays a critical role in development and in  wound healing. Various growth factors bind to heparan sulfate, and the for¬  mation of complexes of the glycosaminoglycan, the growth factor, and the  growth factor receptor is required to initiate cell differentiation and prolifera¬  tion. Specific sulfation patterns on heparan sulfate are required for the forma¬  tion of these ternary complexes.   Plants Produce Pectin. Plants do not synthesize glycosaminoglycans, but the  pectins, which are major components of cell walls, may function similarly as  shock absorbers. Pectins are heterogeneous polysaccharides with a core of  a(l—>4)-linked galacturonate residues interspersed with the hexose rhamnose.    The galacturonate residues may be modified by the addition of methyl and  acetyl groups. Other polysaccharide chains, some containing the pentoses ara-  binose and xylose and other sugars, are attached to the galacturonate. The  aggregation of pectin molecules to form bundles requires divalent cations  (usually Ca 2+ ), which form cross-links between the anionic carboxylate groups  of neighboring galacturonate residues. The tendency for pectin to form highly  hydrated gels is exploited in the manufacture of jams and jellies, to which  pectin is often added to augment the endogenous pectin content of the fruit.   Bacterial Biofilms Are a Type of Extracellular Matrix. Outside the labora¬  tory, bacteria are most often found growing on surfaces as a biofilm, an as¬  sociation of cells in a semisolid matrix (Fig. 8-14). The extracellular material  of the biofilm consists mostly of highly hydrated polysaccharides such as an¬  ionic poly-D-glucuronate, poly-TV-acetylglucosamine, cellulose-like molecules,  and acetylated glycans. A biofilm is difficult to characterize, as it typically  houses a mixture of species, and the proportions of its component polysaccha¬  rides can vary over time and space.   The gel-like consistency of a biofilm, for example, the plaque that forms  on teeth, prevents bacterial cells from being washed away and protects them  from desiccation. Biofilms that develop on medical apparatus, such as  catheters, are problematic because they offer a foothold for pathogenic organ¬  isms and create a barrier to soluble antimicrobial agents.   • Bacterial cell walls consist of glycan chains cross-linked by peptides.   • The oligosaccharide chains covalently attached to eukaryotic glycoproteins may  play a role in protein structure and recognition.    Many proteins are actually glycoproteins, with carbohydrate contents varying  from <1% to >90% by weight. Glycoproteins occur in all forms of life and  have functions that span the entire spectrum of protein activities, including   those of enzymes, transport proteins, receptors, hormones, and structural pro¬  teins. The polypeptide chains of glycoproteins, like those of all proteins, are  synthesized under genetic control. Their carbohydrate chains, in contrast, are  enzymatically generated and covalently linked to the polypeptide without the  rigid guidance of nucleic acid templates. For this reason, glycoproteins tend  to have variable carbohydrate composition, a phenomenon known as micro¬  heterogeneity. Characterizing the structures of carbohydrates—and their  variations—is one goal of the field of glycomics, which complements the studies  of genomics (for DNA) and proteomics (for proteins).   Proteins and glycosaminoglycans in the extracellular matrix aggregate cova¬  lently and noncovalently to form a diverse group of macromolecules known  as proteoglycans. Electron micrographs (Fig. 8-15^) and other evidence indi¬  cate that proteoglycans have a bottlebrush-like molecular architecture, with  “bristles” noncovalently attached to a filamentous hyaluronate “backbone.”  The bristles consist of a core protein to which glycosaminoglycans, most  often keratan sulfate and chondroitin sulfate, are covalently linked (Fig. 8-15^).  The interaction between the core protein and the hyaluronate is stabilized by  a link protein. Smaller oligosaccharides are usually attached to the core pro¬  tein near its site of attachment to hyaluronate. These oligosaccharides are gly-  cosidically linked to the protein via the amide N of specific Asn residues (and  are therefore known as A^linked oligosaccharides; Section 8-3C). The ker¬  atan sulfate and chondroitin sulfate chains are glycosidically linked to the core  protein via oligosaccharides that are covalently bonded to side chain O atoms  of specific Ser or Thr residues (i.e., O -linked oligosaccharides).   Altogether, a central strand of hyaluronate, which varies in length from  4000 to 40,000 A, can have up to 100 associated core proteins, each of which  binds ~50 keratan sulfate chains of up to 250 disaccharide units and --TOO  chondroitin sulfate chains of up to 1000 disaccharide units each. This accounts  for the enormous molecular masses of many proteoglycans, which range up to  tens of millions of daltons.   The extended brushlike structure of proteoglycans, together with the poly an¬  ionic character of their keratan sulfate and chondroitin sulfate components, cause  these complexes to be highly hydrated. Cartilage, which consists of a meshwork  of collagen fibrils that is filled in by proteoglycans, is characterized by its high  resilience: The application of pressure on cartilage squeezes water away from  the charged regions of its proteoglycans until charge—charge repulsions prevent  further compression. When the pressure is released, the water returns. Indeed,  the cartilage in the joints, which lacks blood vessels, is nourished by this flow  of liquid brought about by body movements. This explains why long periods  of inactivity cause cartilage to become thin and fragile.   Bacteria are surrounded by rigid cell walls that give them their  characteristic shapes (Fig. 1-7) and permit them to live in hypo¬  tonic (less than intracellular salt concentration) environments  that would otherwise cause them to swell osmotically until their   FIG. 8-1 5 A proteoglycan, (a) Electron micrograph showing a central  strand of hyaluronate, which supports numerous projections. [From Caplan, A.I., Sci.   Am. 251(4), 87 (1984). Copyright © Scientific American, Inc. Used by permission.]   (b) Bottlebrush model of the proteoglycan shown in Part a. Numerous core proteins are  noncovalently linked to the central hyaluronate strand. Each core protein has three  saccharide-binding regions. (b)   able medical significance because they are, in part, responsible for bacterial  virulence (disease-evoking power). In fact, the symptoms of many bacterial  diseases can be elicited in animals merely by injecting bacterial cell walls.  Furthermore, bacterial cell wall components are antigenic (Section 7-3B), so  such injections often invoke immunity against these bacteria.   Bacteria are classified as gram-positive or gram-negative according to  whether or not they take up Gram stain (a procedure developed in 1884 by  Christian Gram in which heat-fixed cells are successively treated with the dye  crystal violet and iodine and then destained by ethanol or acetone). Gram¬  positive bacteria (Fig. 8-16^) have a thick cell wall (—250 A) surrounding  their plasma membrane, whereas gram-negative bacteria (Fig. 8-1 6 b) have a  thin cell wall (—30 A) covered by a complex outer membrane. This outer  membrane functions, in part, to exclude substances toxic to the bacterium, in¬  cluding Gram stain. This accounts for the observation that gram-negative bac¬  teria are more resistant to antibiotics than are gram-positive bacteria.   The cell walls of bacteria consist of covalently linked polysaccharide and  polypeptide chains, which form a baglike macromolecule that completely en¬  cases the cell. This framework, whose structure was elucidated in large part by  Jack Strominger, is known as a peptidoglycan. Its polysaccharide component  consists of linear chains of alternating (3(1—>4)-linked GlcNAc and  iV-acetylmuramic acid (Latin: murus , wall). The lactic acid group of  TV-acetylmuramic acid forms an amide bond with a D-amino acid-containing  tetrapeptide to form the peptidoglycan repeating unit (Fig. 8-17). Neighboring  parallel peptidoglycan chains are covalently cross-linked through their tetrapep¬  tide side chains, although only —40% of possible cross-links are made.   In the bacterium Staphylococcus aureus , whose tetrapeptide has the  sequence L-Ala-D-isoglutamyl-L-Lys-D-Ala, the cross-link consists of a pen-  taglycine chain that extends from the terminal carboxyl group of one  tetrapeptide to the 8-amino group of the Lys in a neighboring tetrapeptide.  To explore this structure, Simon Foster has used atomic force microscopy  (AFM), an imaging technique that reports the variation in the force between  a probe that is several nanometers in diameter and a surface of interest as the  probe is scanned over the surface; its resolution is as little as several angstroms.  Foster s model of the cell wall of the gram-negative bacterium Bacillus subtilis  is shown in Figure 8-18. Several glycan chains are cross-linked much as de¬  scribed above to form a peptidoglycan “rope,” which due to its natural twist,  forms an — 50-nm-diameter helical cable up to 50 |Jim in length that coils  around the long axis of the bacterium to form its cell wall. This structure is  presumably stabilized by the formation of covalent cross-links between neigh¬  boring segments of the coil. The cell walls of gram-negative bacteria appear to    FIG. 8-16 Bacterial cell walls. This diagram compares the cell envelopes of  (a) gram-positive bacteria and (b) gram-negative bacteria.    FIG. 8-1 7 Peptidoglycan. (a) The repeating unit of peptidoglycan is an  A/-acetylglucosamine-A/-acetylmuramic acid disaccharide whose lactyl side chain forms an  amide bond with a tetrapeptide. The tetrapeptide of S. aureus is shown. The isoglutamyl  residue is so designated because it forms a peptide link via its y-carboxyl group, (b) The  S. aureus bacterial cell wall peptidoglycan, showing its pentaglycine connecting bridges  (purple).    be only one layer thick, whereas as those of gram-positive bacteria are postu¬  lated to consist of several such layers. How the peptidoglycan imposes cell  shape is unknown.   The D-amino acids of peptidoglycans render them resistant to proteases,  which are mostly specific for L- amino acids. However, lysozyme, an enzyme  that is present in tears, mucus, and other vertebrate body secretions, as well  as in egg whites, catalyzes the hydrolysis of the (3(1—>4) glycosidic linkage be¬  tween N -acetylmuramic acid and TV-acetylglucosamine (the structure and  mechanism of lysozyme are examined in detail in Section 11-4). The cell wall  is also compromised by antibiotics that inhibit its biosynthesis (Box 8-3).    FIG. 8-18 Model of the B. subtilis cell wall. The cell wall consists of a right-handed  helical cable composed of several peptidoglycan strands that wraps about the bacterium’s  plasma membrane. The cell is ~3 |±m long. [Courtesy of Simon Foster, University of  Sheffield, U.K.]    In 1928, Alexander Fleming noticed  that the chance contamination of a  bacterial culture plate with the mold  Penicillium notatum resulted in the  lysis of the bacteria in the vicinity of  the mold. This was caused by the  presence of penicillin, an antibiotic  secreted by the mold. Penicillin con¬  tains a thiazolidine ring (red) fused to  a (3-lactam ring (blue). A variable  R group is bonded to the (3-lactam  ring via a peptide link.   Penicillin specifically binds to and inactivates enzymes that  cross-link the peptidoglycan strands of bacterial cell walls.  Since cell wall expansion in growing cells requires that their  rigid cell walls be opened up for the insertion of new cell wall  material, exposure of growing bacteria to penicillin results in  cell lysis. However, since no human enzyme binds penicillin  specifically, it is not toxic to humans and is therefore therapeu¬  tically useful.    Most bacteria that are resistant to penicillin secrete the enzyme  penicillinase (also called (3-lactamase), which inactivates peni¬  cillin by cleaving the amide bond of its (3-lactam ring. Attempts to  overcome this resistance have led to the development of (3-lacta-  mase inhibitors such as sulbactam that are often administered in  combination with penicillin derivatives.   Multiple drug resistant bacteria are a growing problem. For  many years, vancomycin, the so-called antibiotic of last resort,  has been used to treat bacterial infections that do not succumb  to other antibiotics. Vancomycin inhibits the transpeptidation  (cross-linking) reaction of bacterial cell wall synthesis by binding  to the peptidoglycan precursor. However, bacteria can become  resistant to vancomycin by acquiring a gene that allows cell wall  synthesis from a slightly different precursor sequence, to which  vancomycin binds much less effectively.   One limitation of drugs such as vancomycin and penicillin,  particularly for slow-growing bacteria, is that the drug may halt  bacterial growth without actually killing the cells. For this reason,  effective antibacterial treatments may require combinations of  antibiotics over a course of several weeks.   Almost all the secreted and membrane-associated proteins of eukaryotic cells  are glycosylated. Oligosaccharides are covalently attached to proteins by  either TV-glycosidic or O-glycosidic bonds.   A/-Linked Oligosaccharides Are Attached to Asparagine. In N-linked  oligosaccharides, GlcNAc is invariably -linked to the amide nitrogen of an Asn  residue in the sequence Asn-X-Ser or Asn-X- Thr, where X is any amino acid ex¬  cept Pro and only rarely Asp, Glu, Leu, or Trp.   TV-Glycosylation occurs cotranslationally, that is, while the polypeptide is be¬  ing synthesized. Proteins containing TV-linked oligosaccharides typically are  glycosylated and then processed as elucidated, in large part, by Stuart Kornfeld   1. An oligosaccharide containing 9 mannose residues, 3 glucose residues,  and 2 GlcNAc residues is attached to the Asn of a growing polypep¬  tide chain that is being synthesized by a ribosome associated with the  endoplasmic reticulum (Section 9-4D).   2. Some of the sugars are removed during processing, which begins in the  lumen (internal space) of the endoplasmic reticulum and continues in  the Golgi apparatus (Fig. 1-8). Enzymatic trimming is accomplished  by glucosidases and mannosidases.   Removal of monosaccharide  units produces a  (mannose) 3 (GlcNAc) 2  oligosaccharide. This  (mannose) 3 (GlcNAc) 2 core  is found in all /V-linked  oligosaccharides.    Further trimming and  addition of other sugars  yields a variety of /V-linked  oligosaccharides. ,   addition of a (mannose) 9 (glucose) 3 (GlcNAc) 2 oligosaccharide is  followed by removal of monosaccharides as catalyzed by glycosidases,  and the addition of other monosaccharides as catalyzed by    glycosyltransferases. The core pentasaccharide occurs in all AHinked  oligosaccharides. [Adapted from Kornfeld, R. and Kornfeld, S., Annu.  Rev Biochem. 54, 640 (1985).] O See Kinemage Exercise 7-4.    3. Additional monosaccharide residues, including GlcNAc, galactose,  fucose, and sialic acid, are added by the action of specific glycosyl¬  transferases in the Golgi apparatus.   The exact steps of TV-linked oligosaccharide processing vary with the iden¬  tity of the glycoprotein and the battery of endoglycosidases in the cell, but all  TV-linked oligosaccharides have a common core pentasaccharide with the fol¬  lowing structure:   In some glycoproteins, processing is limited, leaving “high-mannose”  oligosaccharides; in other glycoproteins, extensive processing generates large  oligosaccharides containing several kinds of sugar residues. There is enormous di¬  versity among the oligosaccharides ofN-linked glycoproteins. Indeed, even glycopro¬  teins with a given polypeptide chain exhibit considerable microheterogeneity,    presumably as a consequence of incomplete glycosylation and lack of absolute  specificity on the part of glycosidases and glycosyltransferases.    O-Linked Oligosaccharides Are Attached to Serine or Threonine. The most  common O-glycosidic attachment involves the disaccharide core f-galactosyl-  (l^3)-oi -N-acetylgalactosamine linked to the OH group of either Ser or Thr:    Less commonly galactose, mannose, and xylose form O-glycosides with Ser or  Thr. Galactose also forms O-glycosidic bonds to the 5-hydroxylysyl residues  of collagen (Section 6-1C). O-Linked oligosaccharides vary in size from a sin¬  gle galactose residue in collagen to the chains of up to 1000 disaccharide units  in proteoglycans.   O-Linked oligosaccharides are synthesized in the Golgi apparatus by the  serial addition of monosaccharide units to a completed polypeptide chain.  Synthesis starts with the transfer of GalNAc to a Ser or Thr residue on the  polypeptide. N- Linked oligosaccharides are transferred to an Asn in a specific  amino acid sequence, but O-glycosylated Ser and Thr residues are not mem¬  bers of any common sequence. Instead, the locations of glycosylation sites  are specified only by the secondary or tertiary structure of the polypeptide.  O-Glycosylation continues with stepwise addition of sugars by the correspon¬  ding glycosyltransferases. The energetics and enzymology of oligosaccharide  synthesis are discussed further in Section 16-5.   A single protein may contain several N- and O-linked oligosaccharide chains,  although different molecules of the same glycoprotein may differ in the se¬  quences, locations, and numbers of covalently attached carbohydrates (the vari¬  ant species of a glycoprotein are known as its glycoforms). This heterogeneity  makes it difficult to assign discrete biological functions to oligosaccharide  chains. In fact, certain glycoproteins synthesized by cells that lack particular  oligosaccharide-processing enzymes appear to function normally despite ab¬  normal or absent glycosylation. In other cases, however, glycosylation may  affect a proteins structure, stability, or activity.   Oligosaccharides Help Define Protein Structure. Oligosaccharides are usu¬  ally attached to proteins at sequences that form surface loops or turns. Since  sugars are hydrophilic, the oligosaccharides tend to project away from the pro¬  tein surface. Because carbohydrate chains are often conformationally mobile,  oligosaccharides attached to proteins can occupy time-averaged volumes of  considerable size (Fig. 8-20). In this way, an oligosaccharide can shield a pro¬  teins surface, possibly modifying its activity or protecting it from proteolysis.    FIG. 8-20 Model of oligosaccharide dynamics. The allowed conformations of a  (GlcNAc) 2 (mannose) 5 _g oligosaccharide ( yellow ) attached to the bovine pancreatic enzyme  ribonuclease B ( purple ) are shown in superimposed “snapshots.” [Courtesy of Raymond  Dwek, Oxford University, U.K.]   In addition, some oligosaccharides may play structural roles by limiting Section 3 Glycoproteins   the conformational freedom of their attached polypeptide chains. Since  TV-linked oligosaccharides are added as the protein is being synthesized, the  attachment of an oligosaccharide may help determine how the protein folds.   In addition, the oligosaccharide may help stabilize the folded conformation of  a polypeptide by reducing backbone flexibility. In particular, O-linked  oligosaccharides, which are usually clustered in heavily glycosylated segments  of a protein, may help stiffen and extend the polypeptide chain.    Oligosaccharides Mediate Recognition Events. The many possible ways  that carbohydrates can be linked together to form branched structures gives  them the potential to carry more biological information than either nucleic  acids or proteins of similar size. For example, two different nucleotides can  make only two distinct dinucleotides, but two different hexoses can combine in  36 different ways (although not all possibilities are necessarily realized in nature).   The first evidence that unique combinations of carbohydrates might be  involved in intercellular communication came with the discovery that all cells  are coated with sugars in the form of glycoconjugates such as glycoproteins  and glycolipids. The oligosaccharides of glyco conjugates form a fuzzy layer up  to 1400 A thick in some cells (Fig. 8-21).   Additional evidence that cell-surface carbohydrates have recognition func¬  tions comes from lectins (proteins that bind carbohydrates), which are ubiq¬  uitous in nature and frequently appear on the surfaces of cells. Lectins are  exquisitely specific: They can recognize individual monosaccharides in partic¬  ular linkages to other sugars in an oligosaccharide (this property also makes  lectins useful laboratory tools for isolating glycoproteins and oligosaccharides).  Protein-carbohydrate interactions are typically characterized by extensive hydro¬  gen bonding (often including bridging water molecules) and the van der Waals  packing of hydrophobic sugar faces against aromatic side chains (Fig. 8-22).   Proteins known as selectins mediate the attachment between leukocytes  (circulating white blood cells) and the surfaces of endothelial cells (the cells  that line cavities, in this case blood vessels). Leukocytes constitutively (contin¬  ually) express selectins on their surface; endothelial cells transiently display  their own selectins in response to tissue damage from infection or mechanical  injury. The selectins recognize and bind specific oligosaccharides on cell-  surface glycoproteins. Reciprocal selectin-oligosaccharide interactions between  the two cell types allow the endothelial cells to “capture” circulating leukocytes,  which then crawl past the endothelial cells on their way to eliminate the  infection or help repair damaged tissues.   Other cell-cell recognition phenomena depend on oligosaccharides. For  example, proteins on the surface of mammalian spermatozoa recognize  GlcNAc or galactose residues on the glycoproteins of the ovum as part of the  binding and activation events during fertilization. Many viruses, bacteria, and  eukaryotic parasites invade their target tissues by first binding to cell-surface  carbohydrates.   T Oligosaccharides Are Antigenic Determinants. The carbohydrates on  cell surfaces are some of the best known immunochemical markers. For ex¬  ample, the ABO blood group antigens are oligosaccharide components of  glycoproteins and glycolipids on the surfaces of an individuals cells (not just    FIG. 8-22 Carbohydrate binding by a lectin. Human galectin-2 binds p-galactosides,  such as lactose, primarily through their galactose residue. The galactose and glucose  residues are shown with C green and 0 red, and the lectin amino acid side chains  are shown in violet. Hydrogen bonds between the side chains and the sugar residues are  shown as dashed yellow lines. [Courtesy of Hakon Leffler, University of California at  San Francisco. PDBid 1HLCJ   FIG. 8-21 Electron micrograph of the  erythrocyte surface. Its thick (up to 1400 A)   carbohydrate coat, which is called the  glycocalyx, consists of closely packed  oligosaccharides attached to cell-surface proteins  and lipids. [Courtesy of Harrison Latta, UCLA.]   Chapter 8 Carbohydrates TABLE 8-1 Structures of the A, B, and H Antigenic Determinants in   • Describe the general structures of proteo¬  glycans, peptidoglycans, and glycosylated  proteins.   • Why are gram-positive bacteria more  susceptible to antibiotics such as peni¬  cillin?   • Explain why the oligosaccharides of glyco¬  proteins are more complicated to describe  than amino acids or nucleotide  sequences.    red blood cells). Individuals with type A cells have A antigens on their cell  surfaces and carry anti-B antibodies in their blood; those with type B cells,  which bear B antigens, carry anti-A antibodies; those with type AB cells, which  have both A and B antigens, carry neither anti-A nor anti-B antibodies; and  type O individuals, whose cells bear neither antigen, carry both anti-A and  anti-B antibodies. Consequently, the transfusion of type A blood into a type  B individual, for example, results in an anti-A antibody-A antigen reaction,  which agglutinates (clumps together) the transfused erythrocytes, resulting in  an often fatal blockage of blood vessels.   Table 8-1 lists the oligosaccharides found in the A, B, and H antigens  (type O individuals have the H antigen). These occur at the nonreducing ends  of the oligosaccharides. The H antigen is the precursor oligosaccharide of  A and B antigens. Type A individuals have a 303-residue glycosyltransferase that  specifically adds a GalNAc residue to the terminal position of the H antigen.  In type B individuals, this enzyme, which differs by four amino acid residues  from that of type A individuals, instead adds a galactose residue. In type O  individuals, the enzyme is inactive because its synthesis terminates after its  115th residue.   • Monosaccharides, the simplest carbohydrates, are classified as  aldoses or ketoses.   • The cyclic hemiacetal and hemiketal forms of monosaccharides  have either the a or (3 configuration at their anomeric carbon but  are conformationally variable.   • Monosaccharide derivatives include aldonic acids, uronic acids,  alditols, deoxy sugars, amino sugars, and a- and (3-glycosides.   • Cellulose and chitin are polysaccharides whose (3(1—>4) linkages  cause them to adopt rigid and extended structures.   • The storage polysaccharides starch and glycogen consist of  a-glycosidically linked glucose residues.   • Glycosaminoglycans are unbranched polysaccharides containing  uronic acid and amino sugars that are often sulfated.   • Proteoglycans are enormous molecules consisting of hyaluronate  with attached core proteins that bear numerous glycosaminogly¬  cans and oligosaccharides.   • Bacterial cell walls are made of peptidoglycan, a network of poly¬  saccharide and polypeptide chains.   • Glycosylated proteins may contain A-linked oligosaccharides  (attached to Asn) or O-linked oligosaccharides (attached to Ser or  Thr) or both. Different molecules of a glycoprotein may  contain different sequences and locations of oligosaccharides.   • Oligosaccharides play important roles in determining protein  structure and in cell-surface recognition phenomena.   1. How many stereoisomers are possible for (a) a ketopentose,  (b) a ketohexose, and (c) a ketoheptose?   2. Are (a) D-glucitol, (b) D-galactitol, and (c) D-glycerol optically  active?   3. Which of the following pairs of sugars are epimers of each other?   4. Which of the following pairs of sugars are epimers of each other?   6. The sucrose substitute tagatose (Fig. 8-2) is produced by hy¬  drolyzing lactose and then chemically converting one of the two  resulting aldoses to a ketose. Which residue of lactose gives rise  to tagatose?   7. Draw a Fischer projection of L-fucose. L-Fucose is the 6-deoxy  form of which L-hexose?   10. In addition to lactose, how many other heterodisaccharides can  D-galactose and D-glucose form?   11. (a) Deduce the structure of the disaccharide trehalose from the  following information: Complete hydrolysis yields only  D-glucose; it is hydrolyzed by a-glucosidase but not P-glucosidase;  and it does not reduce Cu 2+ to Cu + . (b) When exposed to  dehydrating conditions, many plants and invertebrates synthesize  large amounts of trehalose, which enables them to survive pro¬  longed desiccation. What properties of the trehalose molecule  might allow it to act as a water substitute?   12. The artificial sweetener sucralose is a derivative of sucrose with  the formal name l,6-dichloro-l,6-dideoxy-P-D-fructofuranosyl-  4-chloro-4-deoxy-a-D-galactopyranoside. Draw its structure.   13. Draw the structure of the mannose trisaccharide that is part of  the core AHinked oligosaccharide.    14. Identify the monosaccharides and their linkages in the following  disaccharide.   15. Cellulose is treated with methanol, which methylates free  anomeric carbons, (a) How many methyl groups would be incor¬  porated per cellulose chain? (b) Cellulose is treated with dimethyl  sulfate, which adds a methyl group to all free hydroxyl groups.  The cellulose is then hydrolyzed to release all its monosaccha¬  rides. Draw their structure.   16. Glycogen is treated with dimethyl sulfate, which adds a methyl  group to every free OH group. Next, the molecule is hydrolyzed  to break all the glycosidic bonds between glucose residues. The  reaction products are then chemically analyzed.   (a) How many different types of methylated glucose molecules  are obtained?   (b) Draw the structure of the one that is most abundant.   17. How many reducing ends are in a molecule of glycogen that con¬  tains 10,000 residues with a branch every 10 residues?   18. Is amylose or amylopectin more likely to be a long-term storage  polysaccharide in plants?   19. “Nutraceuticals” are products that are believed to have some ben¬  eficial effect but are not strictly defined as either food or drug.  Why might an individual suffering from osteoarthritis be  tempted to consume the nutraceutical glucosamine?   20. The core of pectin molecules is a polymer of a( 1—>4)-linked  D-galacturonate. Draw one of its residues.   21. Calculate the net charge of a chondroitin-4-sulfate molecule con¬  taining 100 disaccharide units.   22. Basic fibroblast growth factor binds to glycosaminoglycans in the  extracellular matrix. Which amino acids would you expect to be  abundant in this protein?   23. Following their synthesis, many proteoglycans are stored in a  highly condensed state in intracellular compartments. These  compartments also contain many Ca 2+ ions, which are rapidly  pumped out as the proteoglycans are released extracellularly.  Explain the function of the Ca 2+ ions.   24. The peptidoglycan cell walls of bacteria contain pores that allow  the passage of substances with diameters less than 200 A. Why  is it important that the cell wall be porous rather than solid?   23. Draw the structure of the O-type oligosaccharide (the H antigen,  described in Table 8-1).   26. During gel electrophoresis, glycoproteins migrate as relatively dif¬  fuse bands, whereas nonglycosylated proteins typically migrate as  narrow, well-defined bands. Explain the difference in elec¬  trophoretic behavior.   1. Looking at branched carbohydrates. Explore some well-known  glycans using MarvinSketch, PubChem, and the KEGG N-glycan  synthesis reference pathway.   2. The role of glycans in swine flu. Review the structures of two  proteins (the H and N in H1N1) that are critical to swine flu  infections and vaccine development, including the glycans that  are attached to these proteins. Then explore the glycan targets  for swine flu.    Chitosan is a substance pro¬  duced from the chitin of crustacean shells (mainly from shrimp and  crab). How is the chitin chemically modified, and how does this  treatment affect its physical and chemical properties? What are  some industrial and medical uses for chitosan? Are all the adver¬  tised uses for chitosan supported by scientific data?    Aoki-Kinoshita, K.F., An introduction to bioinformatics for glycomics  research, PLoS Comput. Biol 4, el000073 (2008). [Describes some  of the approaches used to characterize carbohydrate structures.]  Branda, S.S., Vik, A., Friedman, L., and Kolter, R., Biofilms: the matrix  revisited, Trends Microbiol. 13, 20-26 (2005). [Summarizes the gen¬  eral features of biofilms and ways of studying them.]   Esko, J.D. and Lindahl, U., Molecular diversity of heparan sulfate,  J. Clin. Invest. 108, 169-173 (2001). [Reviews the structure, function,  and biosynthesis of heparan sulfate-containing proteoglycans.]  Meroueh, S.O., Bencze, K.Z., Hesek, D., Lee, M., Fisher, J.F., Stemmier,  T.L., and Mobashery, S., Three-dimensional structure of the bac¬  terial cell wall peptidoglycan, Proc. Natl. Acad. Sci. 103, 4404-4409  (2006).   Mitra, N., Sinha, S., Ramya, T.N.C., and Surolia, A., TV-Linked oligosac¬  charides as outfitters for glycoprotein folding, form and function,    Trends Biochem. Sci. 31, 156-163 and 251 (2006). [Summarizes the  ways in which oligosaccharides can influence glycoprotein structure.]  Sharon, N. and Lis, H., History of lectins: from hemagglutinins to bio¬  logical recognition molecules, Glycobiology 14, 53R-62R (2004).  [A historical account of lectin research and applications.]   Spiro, R.G., Protein glycosylation: nature, distribution, enzymatic forma¬  tion, and disease implications of glycopeptide bonds, Glycobiology  12 , 43R-56R (2002). [Catalogs the various ways in which saccha¬  rides are linked to proteins, and describes the enzymes involved in  glycoprotein synthesis.]   Taylor, M.E. and Drickamer, K., Introduction to Glycobiology, Oxford  University Press (2011).   Varki, A., Cummings, R.D., Esko, J.D., Freeze, H.H., Stanley,  S., Bertozzi, C.R., Hart, G.W., and Etzler, M.E. (Eds.), Essentials of  Glycobiology (2nd ed.), Cold Spring Harbor Laboratory Press (2009).    This fluorescence micrograph shows the spherical bilayered structures, or vesicles, that  form from mixtures of lipids. In some cases, the lipids in the bilayer do not mix uni¬  formly but segregate into domains that are visible as bright areas (where there are  fluorescently labeled lipids) against a darker background (unlabeled lipids). [Courtesy  Aurelia R. Honerkamp-Smith and Sarah L. Keller, University of Washington, Seattle.]   A The Properties of Fatty Acids Depend on Their  Hydrocarbon Chains   C Glycerophospholipids Are Amphiphilic  D Sphingolipids Are Amino Alcohol Derivatives  E Steroids Contain Four Fused Rings  F Other Lipids Perform a Variety of Metabolic  Roles    Lipids (Greek: lip os , fat) are the fourth major group of molecules found in all  cells. Unlike nucleic acids, proteins, and polysaccharides, lipids are not poly¬  meric. However, they do aggregate, and it is in this state that they perform  their central function as the structural matrix of biological membranes.   Lipids exhibit greater structural variety than the other classes of biologi¬  cal molecules. To a certain extent, lipids constitute a catchall category of sub¬  stances that are similar only in that they are largely hydrophobic and only  sparingly soluble in water. In general, lipids perform three biological functions  (although certain lipids apparently serve more than one purpose in some cells):   C Membrane Lipids Are Distributed Asymmetrically  D The Secretory Pathway Generates Secreted and  Transmembrane Proteins  E Intracellular Vesicles Transport Proteins  F Proteins Mediate Vesicle Fusion    1. Lipid molecules in the form of lipid bilayers are essential components  of biological membranes.    In this chapter we examine the structures and physical properties of the  most common types of lipids. Next, we look at the properties of the lipid  bilayer and the proteins that are situated within it. Finally, we explore current  models of membrane structure. The following chapter examines membrane  transport phenomena. The participation of lipids in intracellular signaling is  discussed in Section 13-4.   • The length and saturation of a fatty acid chain determine its physical properties.   • Steroids, isoprenoids, and other lipids perform a wide variety of functions.   Lipids are substances of biological origin that are soluble in organic solvents such  as chloroform and methanol. Hence, they are easily separated from other bio¬  logical materials by extraction into organic solvents. Fats, oils, certain vitamins  and hormones, and most nonprotein membrane components are lipids. In this  section, we discuss the structures and physical properties of the major classes  of lipids.   A The Properties of Fatty Acids Depend on Their  Hydrocarbon Chains   Fatty acids are carboxylic acids with long-chain hydrocarbon side groups (Fig. 9-1).  They usually occur in esterified form as major components of the various lipids  described in this chapter. The more common biological fatty acids are listed  in Table 9-1. In higher plants and animals, the predominant fatty acid residues  are those of the C 1( 5 and C 18 species: palmitic, oleic, linoleic, and stearic  acids. Fatty acids with <14 or >20 carbon atoms are uncommon. Most fatty  acids have an even number of carbon atoms because they are biosynthesized  by the concatenation of C 2 units (Section 20-4).   Over half of the fatty acid residues of plant and animal lipids are unsat¬  urated (contain double bonds) and are often polyunsaturated (contain two    FIG. 9-1 The structural formulas of some  C 18 fatty acids. The double bonds all have the  cis configuration.   CH 3 (CH 2 ) 5 CH=CH(CH 2 ) 7 COOH   CH 3 (CH 2 ) 7 CH=CH(CH 2 ) 7 COOH   CH 3 (CH 2 ) 4 (CH=CHCH 2 ) 2 (CH 2 ) 6 COOH   CH 3 CH 2 (CH=CHCH 2 ) 3 (CH 2 ) 6 COOH   CH 3 (CH 2 ) 4 (CH=CHCH 2 ) 3 (CH 2 ) 3 COOH   CH 3 (CH 2 ) 4 (CH=CHCH 2 ) 4 (CH 2 ) 2 COOH   CH 3 CH 2 (CH=CHCH 2 ) 5 (CH 2 ) 2 COOH   CH 3 CH 2 (CH=CHCH 2 ) 6 CH 2 COOH   CH 3 (CH 2 ) 7 CH=CH(CH 2 ) 13 COOH    ^Number of carbon atoms: Number of double bonds. For unsaturated fatty acids, the quantity “ n-x indicates the position of the last double bond in the  fatty acid, where n is its number of C atoms, and x is the position of the last double-bonded C atom counting from the methyl-terminal (w) end.    or more double bonds). Bacterial fatty acids are rarely polyunsaturated but are  commonly branched, hydroxylated, or contain cyclopropane rings.   Table 9-1 indicates that the first double bond of an unsaturated fatty  acid commonly occurs between its C9 and CIO atoms counting from the  carboxyl C atom. This bond is called a A 9 - or 9-double bond. In polyun¬  saturated fatty acids, the double bonds tend to occur at every third carbon  atom (e.g., —CH=CH—CH 2 —CH=CH—) and so are not conjugated (as  in —CH=CH—CH=CH—). Two important classes of polyunsaturated  fatty acids are designated as u>-3 or o>-6 fatty acids, a nomenclature that iden¬  tifies the last double-bonded carbon atom as counted from the methyl termi¬  nal ( 00 ) end of the chain. ot-Linolenic acid and linoleic acid (Fig. 9-1) are  examples of such fatty acids.   Saturated fatty acids (which are fully reduced or “saturated” with hydro¬  gen) are highly flexible molecules that can assume a wide range of conforma¬  tions because there is relatively free rotation around each of their C—C bonds.  Nevertheless, their lowest energy conformation is the fully extended confor¬  mation, which has the least amount of steric interference between neighbor¬  ing methylene groups. The melting points (mp) of saturated fatty acids, like  those of most substances, increase with their molecular mass (Table 9-1).   Fatty acid double bonds almost always have the cis configuration (Fig. 9-1).  This puts a rigid 30° bend in the hydrocarbon chain. Consequently, unsatu¬  rated fatty acids pack together less efficiently than saturated fatty acids. The  reduced van der Waals interactions of unsaturated fatty acids cause their melt¬  ing points to decrease with the degree of unsaturation. The fluidity of lipids  containing fatty acid residues likewise increases with the degree of unsaturation   Chapter 9 Lipids and Biological Membranes G f th e fatty acids. This phenomenon, as we will see, has important conse¬    FIG. 9-2 Scanning electron micrograph of  adipocytes. Each adipocyte contains a fat  globule that occupies nearly the entire cell.   The fats and oils that occur in plants and animals consist largely of mixtures  of triacylglycerols (also called triglycerides). These nonpolar, water-insoluble  substances are fatty acid triesters of glycerol ( at left). Triacylglycerols function  as energy reservoirs in animals and are therefore their most abundant class of  lipids even though they are not components of cellular membranes.   Triacylglycerols differ according to the identity and placement of their three  fatty acid residues. Most triacylglycerols contain two or three different types of  fatty acid residues and are named according to their placement on the glycerol  moiety, for example, l-palmitoleoyl-2-linoleoyl-3-stearoylglycerol ( at left).  Note that the - ate ending of the name of the fatty acid becomes -oyl in the  fatty acid ester. Fats and oils (which differ only in that fats are solid and oils  are liquid at room temperature) are complex mixtures of triacylglycerols whose  fatty acid compositions vary with the organism that produced them. Plant oils  are usually richer in unsaturated fatty acid residues than animal fats, as the  lower melting points of oils imply.   Triacylglycerols Function as Energy Reserves. Fats are a highly efficient  form in which to store metabolic energy. This is because triacylglycerols are  less oxidized than carbohydrates or proteins and hence yield significantly more  energy per unit mass on complete oxidation. Furthermore, triacylglycerols,  which are nonpolar, are stored in anhydrous form, whereas glycogen (Section  8-2C), for example, binds about twice its weight of water under physiologi¬  cal conditions. Fats therefore provide about six times the metabolic energy of an  equal weight of hydrated glycogen.   In animals, adipocytes (fat cells; Fig. 9-2) are specialized for the synthe¬  sis and storage of triacylglycerols. Whereas other types of cells have only a few  small droplets of fat dispersed in their cytosol, adipocytes may be almost en¬  tirely filled with fat globules. Adipose tissue is most abundant in a subcuta¬  neous layer and in the abdominal cavity. The fat content of normal humans  (21% for men, 26% for women) allows them to survive starvation for 2 or  3 months. In contrast, the body’s glycogen supply, which functions as a short¬  term energy store, can provide for the body’s energy needs for less than a day.  The subcutaneous fat layer also provides thermal insulation, which is particularly    FIG. 9-3 Structure of glycerophospholipids. (a) The backbone, L-glycerol-3-phosphate.  (b) The general formula of the glycerophospholipids. and R 2 are the long-chain  hydrocarbon tails of fatty acids, and X is derived from a polar alcohol (Table 9-2). Note that  glycerol-3-phosphate and glycerophospholipid are chiral compounds.    important for warm-blooded aquatic animals, such as whales, seals, geese, and  penguins, which are routinely exposed to low temperatures.   Glycerophospholipids (or phosphoglycerides) are the major lipid compo¬  nents of biological membranes. They consist of glycerol-3-phosphate whose  Cl and C2 positions are esterified with fatty acids. In addition, the phospho-  ryl group is linked to another usually polar group, X (Fig. 9-3).  Glycerophospholipids are therefore amphiphilic molecules with nonpolar aliphatic  (hydrocarbon) “tails” and polar phosphoryl-X “heads. ”   The simplest glycerophospholipids, in which X = H, are phosphatidic  acids; they are present in only small amounts in biological membranes. In the  glycerophospholipids that commonly occur in biological membranes, the head  groups are derived from polar alcohols (Table 9-2). Saturated or C 18 fatty  acids usually occur at the Cl position of the glycerophospholipids, and the  C2 position is often occupied by an unsaturated C 1( 5 to C 2 o fatty acid.  Individual glycerophospholipids are named according to the identities of these   C green, H white, N blue, 0 red, and P orange.  Note how the unsaturated oleoyl chain (left) is  bent compared to the saturated stearoyl chain.  [Based on coordinates provided by Richard  Venable and Richard Pastor, NIH, Bethesda,  Maryland.]    fatty acid residues (e.g., Fig. 9-4). A glycerophospholipid containing two  palmitoyl chains is an important component of lung surfactant (Box 9-1).   Phospholipases Hydrolyze Glycerophospholipids. The chemical structures—  including fatty acyl chains and head groups—of glycerophospholipids can be  determined from the products of the hydrolytic reactions catalyzed by enzymes  known as phospholipases. For example, phospholipase A 2 hydrolytically ex¬  cises the fatty acid residue at C2, leaving a lysophospholipid (Fig. 9-5).  Lysophospholipids, as their name implies, are powerful detergents that disrupt  cell membranes, thereby lysing cells. Bee and snake venoms are rich sources  of phospholipase A 2 . Other types of phospholipases act at different sites in  glycerophospholipids, as shown in Fig. 9-5.   Enzymes that act on lipids have fascinated biochemists because the en¬  zymes must gain access to portions of the lipids that are buried in a nonaque-  ous environment. Phospholipases A 2 , which constitute some of the best    Dipalmitoylphosphatidylcholine (DPPC) is the major lipid of  lung surfactant, the protein-lipid mixture that is essential for normal  pulmonary function. The surfaces of the cells that form the alveoli  (small air spaces of the lung) are coated with surfactant, which  decreases the alveolar surface tension. Lung surfactant contains  80 to 90% phospholipid by weight, and 70 to 80% of the phos¬  pholipid is phosphatidylcholine, mostly the dipalmitoyl species.   Because the palmitoyl chains of DPPC are saturated, they tend  to extend straight out without bending. This allows close packing  of DPPC molecules, which are oriented in a single layer with their  nonpolar tails toward the air and their polar heads toward the  alveolar cells. When air is expired from the lungs, the volume and  surface area of the alveoli decrease. The collapse of the alveolar    space is prevented by the surfactant, because the closely packed  DPPC molecules resist compression. Reopening a collapsed air  space requires a much greater force than expanding an already  open air space.   Lung surfactant is continuously synthesized, secreted, and recy¬  cled by alveolar cells. Because surfactant production is low until  just before birth, premature infants are at risk of developing respi¬  ratory distress syndrome, which is characterized by difficulty in  breathing due to alveolar collapse. The syndrome can be treated by  introducing exogenous surfactant into the lungs. A related condition  in adults (adult respiratory distress syndrome) is characterized  by insufficient surfactant, usually secondary to other lung injury.   FIG. 9-5 Action of phospholipases. Phospholipase A 2 hydrolytically excises the  C2 fatty acid residue from a phospholipid to yield the corresponding lysophospholipid. The  bonds hydrolyzed by other types of phospholipases, which are named according to their  specificities, are also indicated.    understood lipid-specific enzymes, are relatively small proteins (~14 kD,  —125 amino acid residues). The X-ray structure of phospholipase A 2 from  cobra venom suggests that the enzyme binds a glycerophospholipid molecule  such that its polar head group fits into the enzymes active site, whereas the  hydrophobic tails, which extend beyond the active site, interact with several  aromatic side chains (Fig. 9-6).   Lipases specific for triacylglycerols and membrane lipids catalyze their  degradation in vivo. Occasionally, the hydrolysis products are not destined for  further degradation but instead serve as intra- and extracellular signal mole¬  cules. For example, lysophosphatidic acid (l-acyl-glycerol-3-phosphate),  which is not actually lytic since it has a small head group (an unsubstituted  phosphate group), is produced by hydrolysis of membrane lipids in blood  platelets and injured cells and stimulates cell growth as part of the wound-  repair process. 1,2-Diacylglycerol, derived from membrane lipids by the ac¬  tion of phospholipase C, is an intracellular signal molecule that activates a  protein kinase (Section 13-4C; kinases catalyze ATP-dependent phosphoryl-  transfer reactions).   Plasmalogens Contain an Ether Linkage. Plasmalogens are glycerophos-  pholipids in which the Cl substituent of the glycerol moiety is linked via an  a, (3-unsaturated ether linkage in the cis configuration rather than through an  ester linkage. x   FIG. 9-6 Model of phospholipase A 2  and a glycerophospholipid. The X-ray  structure of the enzyme from cobra venom is  shown with a space-filling model of  dimyristoylphosphatidylethanolamine in its active  site as located by NMR methods. A Ca 2+ ion in  the active site is shown in magenta. [Courtesy of  Edward A. Dennis, University of California at San  Diego.] Q See Interactive Exercise 4.   Ethanolamine, choline, and serine (Table 9-2) form the most common plas¬  malogen head groups. The functions of most plasmalogens are not well un¬  derstood. Because the vinyl ether group is easily oxidized, plasmalogens may  react with oxygen free radicals, by-products of normal metabolism, thereby  preventing free-radical damage to other cell constituents.    FIG. 9-7 A sphingomyelin, (a) Molecular formula, (b) Energy-minimized space-filling  model with C green, H white, N blue, 0 red, and P orange. [Based on coordinates provided  by Richard Venable and Richard Pastor, NIH, Bethesda, Maryland.]   Sphingolipids are also major membrane components. They were named after  the Sphinx because their function in cells was at first mysterious. Most sphin¬  golipids are derivatives of the C 18 amino alcohol sphingosine, whose double  bond has the trans configuration. The TV-acyl fatty acid derivatives of sphin-  gosine are known as ceramides:   FIG. 9-8 Electron micrograph of  myelinated nerve fibers. This cross-sectional  view shows the spirally wrapped membranes  around each nerve axon. The myelin sheath may  be 10-15 layers thick. Its high lipid content  makes it an electrical insulator. [Courtesy of  Cedric S. Raine, Albert Einstein College of  Medicine.]   1. Sphingomyelins, the most common sphingolipids, are ceramides bear¬  ing either a phosphocholine (Fig. 9-7) or a phosphoethanolamine head  group, so they can also be classified as sphingophospholipids. They  typically make up 10 to 20 mol % of plasma membrane lipids.  Although sphingomyelins differ chemically from phosphatidylcholine and  phosphatidylethanolamine , their conformations and charge distributions  are quite similar (compare Figs. 9-4 and 9-7). The membranous myelin  sheath that surrounds and electrically insulates many nerve cell axons  is particularly rich in sphingomyelins (Fig. 9-8).   2. Cerebrosides are ceramides with head groups that consist of a single  sugar residue. These lipids are therefore glycosphingolipids.  Galactocerebrosides and glucocerebrosides are the most prevalent.  Cerebrosides, in contrast to phospholipids, lack phosphate groups and  hence are nonionic.   ch 2 oh ch 2 oh ch 2 oh ch 2 oh   FIG. 9-9 Gangliosides. (a) Structural formula of gangliosides  G M1 , G M2 , and G M3 . Gangliosides G M2 and G M3 differ from G M1 only  by the sequential absences of the terminal D-galactose and  /V-acetyl-D-galactosamine residues. Other gangliosides have different    oligosaccharide head groups, (b) Energy-minimized space-filling model  of G M1 with C green, H white, N blue, and 0 red. [Based on  coordinates provided by Richard Venable and Richard Pastor, NIH,  Bethesda, Maryland.]    3. Gangliosides are the most complex glycosphingolipids. They are cer-  amides with attached oligosaccharides that include at least one sialic  acid residue. The structures of gangliosides G M1 , G M2 , and G M3 ,  three of the hundreds that are known, are shown in Fig. 9-9.  Gangliosides are primarily components of cell-surface membranes and  constitute a significant fraction (6%) of brain lipids.   Gangliosides have considerable physiological and medical significance.  Their complex carbohydrate head groups, which extend beyond the surfaces  of cell membranes, act as specific receptors for certain pituitary glycoprotein  hormones that regulate a number of important physiological functions.  Gangliosides are also receptors for certain bacterial protein toxins such as  cholera toxin. There is considerable evidence that gangliosides are specific de¬  terminants of cell-cell recognition, so they probably have an important role  in the growth and differentiation of tissues as well as in carcinogenesis.  Disorders of ganglioside breakdown are responsible for several hereditary  sphingolipid storage diseases, such as Tay-Sachs disease (Box 20-4), which are  characterized by an invariably fatal neurological deterioration in early childhood.   Sphingolipids, like glycerophospholipids, are a source of smaller lipids that  have discrete signaling activity. Sphingomyelin itself, as well as the ceramide  portions of more complex sphingolipids, appear to specifically modulate the  activities of protein kinases and protein phosphatases (enzymes that remove  phosphoryl groups from proteins) that are involved in regulating cell growth  and differentiation.   Steroids, which are mostly of eukaryotic origin, are derivatives of cyclopen-  tanoperhydrophenanthrene (at right), a compound that consists of four  fused, nonplanar rings (labeled A-D). The much maligned cholesterol, which   FIG. 9-10 Cholesterol, (a) Structural  formula with the standard numbering system.   (b) Energy-minimized space-filling model with  C green, H white, and 0 red. [Based on  coordinates provided by Richard Venable and  Richard Pastor, NIH, Bethesda, Maryland.]   ? Compare the overall size and shape of  cholesterol, a glycerophospholipid, a  sphingolipid, and a ganglioside.   is the most abundant steroid in animals, is further classified as a sterol be¬  cause of its C3-OH group (Fig. 9-10). Cholesterol is a major component of  animal plasma membranes, typically constituting 30 to 40 mol % of plasma  membrane lipids. Its polar OH group gives it a weak amphiphilic character,  whereas its fused ring system provides it with greater rigidity than other mem¬  brane lipids. Cholesterol can also be esterified to long-chain fatty acids to form  cholesteryl esters, such as cholesteryl stearate.   Plants contain little cholesterol but synthesize other sterols. Yeast and fungi  also synthesize sterols, which differ from cholesterol in their aliphatic side  chains and number of double bonds. Prokaryotes contain little, if any, sterol.   In mammals, cholesterol is the metabolic precursor of steroid hormones,  substances that regulate a great variety of physiological functions. The struc¬  tures of some steroid hormones are shown in Fig. 9-11. Steroid hormones are  classified according to the physiological responses they evoke:   1. The glucocorticoids, such as cortisol (a C 2 i compound), affect car¬  bohydrate, protein, and lipid metabolism and influence a wide variety  of other vital functions, including inflammatory reactions and the ca¬  pacity to cope with stress.   2. Aldosterone and other mineralocorticoids regulate the excretion of  salt and water by the kidneys.   3. The androgens and estrogens affect sexual development and function.  Testosterone, a C 19 compound, is the prototypic androgen (male sex  hormone), whereas [5-estradiol, a C 18 compound, is an estrogen (fe¬  male sex hormone).   Glucocorticoids and mineralocorticoids are synthesized by the cortex  (outer layer) of the adrenal gland. Both androgens and estrogens are synthe¬  sized by testes and ovaries (although androgens predominate in testes and es¬  trogens predominate in ovaries) and, to a lesser extent, by the adrenal cortex.  Because steroid hormones are water insoluble, they bind to proteins for trans¬  port through the blood to their target tissues.    T Impaired adrenocortical function, either through disease or trauma, results  in Addison’s disease, which is characterized by hypoglycemia (decreased  amounts of glucose in the blood), muscle weakness, Na + loss, K + retention,  impaired cardiac function, and greatly increased susceptibility to stress. The  victim, unless treated by the administration of glucocorticoids and mineralo-  corticoids, slowly languishes and dies without any particular pain or distress.  Conversely, adrenocortical hyperfunction, which is often caused by a tumor  of the adrenal cortex, results in Cushing’s syndrome, which is characterized  by fatigue, hyperglycemia (increased amount of glucose in the blood), edema  (water retention), and a redistribution of body fat to yield a characteristic  “moon face.”    Vitamin D Regulates Ca 2+ Metabolism. The various forms of vitamin D,   which are really hormones, are sterol derivatives in which the steroid B ring  is disrupted between C9 and CIO:    R = X Vitamin D 3 (cholecalciferol)  R = Y Vitamin D 2 (ergocalciferol)    Vitamin D 2 (ergocalciferol) is nonenzymatically formed in the skin of ani¬  mals through the photolytic action of UV light on the plant sterol ergosterol,  a common milk additive, whereas the closely related vitamin D 3 (cholecal-  ciferol) is similarly derived from 7-dehydrocholesterol (hence the saying that  sunlight provides vitamin D).   Vitamins D 2 and D 3 are inactive; the active forms are produced through  their enzymatic hydroxylation (addition of an OH group) carried out by the liver  (at C25) and by the kidney (at Cl) to yield 1 ct,25-dihydroxycholecalciferol (at left).  Active vitamin D increases serum [Ca 2+ ] by promoting the intestinal absorption  of dietary Ca 2+ . This increases the deposition of Ca 2+ in bones and teeth.   r Vitamin D deficiency in children produces rickets, a disease characterized  by stunted growth and deformed bones caused by insufficient bone min¬  eralization. Although rickets was first described in 1645, it was not until the  early twentieth century that eating animal fats, particularly fish liver oils, was  shown to prevent this deficiency disease. Rickets can also be prevented by ex¬  posing children to sunlight or just UV light in the wavelength range 230 to  313 nm, regardless of their diets.   Since vitamin D is water insoluble, it can accumulate in fatty tissues.  Excessive intake of vitamin D over long periods results in vitamin D intoxi¬  cation. The consequent high serum [Ca 2+ ] results in aberrant calcification of  soft tissues and in the development of kidney stones, which can cause kidney  failure. The observation that the level of skin pigmentation in indigenous hu¬  man populations tends to increase with their proximity to the equator is ex¬  plained by the hypothesis that skin pigmentation functions to prevent vitamin  D intoxication by filtering out excessive solar radiation.   In addition to the well-characterized lipids that are found in large amounts in  cellular membranes, many organisms synthesize compounds that are not mem¬  brane components but are classified as lipids on the basis of their physical  properties. For example, lipids occur in the waxy coatings of plants, where  they protect cells from desiccation by creating a water-impermeable barrier.    Isoprenoids Are Built from Five-Carbon Units. Among the compounds that  are not structural components of membranes—although they are soluble in  the lipid bilayer—are the isoprenoids, which are built from five-carbon units  with the same carbon skeleton as isoprene.    For example, the isoprenoid ubiquinone (also known as coenzyme Q) is re¬  versibly reduced and oxidized in the mitochondrial membrane (its activity is  described in more detail in Section 18-2C). Mammalian ubiquinone consists  of 10 isoprenoid units.    The plant kingdom is rich in isoprenoid compounds, which serve as pigments,  molecular signals (hormones and pheromones), and defensive agents. Indeed,  over 25,000 isoprenoids (also known as terpenoids), which are mostly of    plant, fungal, and bacterial origin, have been characterized. During the course  of evolution, vertebrate metabolism has co-opted several of these compounds  for other purposes. Some of these compounds (e.g., vitamin D) are known as  fat-soluble vitamins (vitamins are organic substances that an animal requires  in small amounts but cannot synthesize and hence must acquire in its diet).   Vitamin A, or retinol, is derived mainly from plant products such as  P-carotene [a red pigment that is present in green vegetables as well as in car¬  rots (after which it is named) and tomatoes; Section 19-1B]. Retinol is oxidized  to its corresponding aldehyde, retinal (at right), which functions as the eyes  photoreceptor at low light intensities. Light causes the retinal to isomerize, trig¬  gering, via a complex signaling pathway, an impulse through the optic nerve.  A severe deficiency of vitamin A can lead to blindness. Retinoic acid also has  hormonelike properties in that it stimulates tissue repair. It is used to treat se¬  vere acne and skin ulcers and is also used cosmetically to eliminate wrinkles.   Vitamin K is a lipid synthesized by plants (as phylloquinone) and bac¬  teria (as menaquinone):    About half of the daily requirement for humans is supplied by intestinal bac¬  teria. Vitamin K participates in the carboxylation of Glu residues in some of  the proteins involved in blood clotting (vitamin K is named for the Danish  word Koagulatiori ). Vitamin K deficiency prevents this carboxylation, and the  resulting inactive clotting proteins lead to excessive bleeding. Compounds that  interfere with vitamin K function are the active ingredients in some rodent  poisons.   Vitamin E is actually a group of compounds whose most abundant mem¬  ber is (^-tocopherol:   This highly hydrophobic molecule is incorporated into cell membranes, where  it functions as an antioxidant that prevents oxidative damage to membrane  proteins and lipids. A deficiency of vitamin E elicits a variety of nonspecific  symptoms, which makes the deficiency difficult to detect. The popularity of  vitamin E supplements rests on the hypothesis that vitamin E protects against  oxidative damage to cells and hence reduces the effects of aging, although clin¬  ical trials have shown that this is not the case.   X = CH 2 OH Retinol (vitamin A)  X = CHO Retinal    Eicosanoids Are Derived from Arachidonic Acid. Other less common lipids  are derived from relatively abundant membrane lipids. Prostaglandins (e.g.,   FIG. 9-1 2 Eicosanoids. Arachidonate is the precursor of  prostaglandins (PG), prostacyclins, thromboxanes (Tx), and lipoxins  (LX). Arachidonate also leads to leukotrienes. Although only a single    example of each type of eicosanoid is shown, each has numerous  physiologically significant derivatives, which are designated by letters  and subscripts (e.g., PGH 2 for prostaglandin H 2 ).    How do lipids differ from the three other  major classes of biological molecules?    • Explain the trend in melting point with  increasing fatty acid chain length. Explain  the trend with increasing number of dou¬  ble bonds.    • Explain why fats contain —38 kJ of energy  per gram whereas carbohydrates contain  —17 kJ per gram.   • Summarize the structures and physical  properties of fatty acids, triacylglycerols,  glycerophospholipids, sphingolipids, and  steroids.    • Which lipids can be considered to be  glycolipids? Phospholipids? Which are  ionic?    Fig. 9-12) were discovered in the 1930s by Ulf von Euler, who thought they  were produced by the prostate gland. Prostaglandins and related compounds—  prostacyclins, thromboxanes, leukotrienes, and lipoxins—are known collec¬  tively as eicosanoids because they are all C 2 o compounds (Greek: eikosi,  twenty). The eicosanoids act at very low concentrations and arc involved in the  production of pain and fever ; and in the regulation of blood pressure, blood coag¬  ulation, and reproduction. Unlike most other types of hormones, eicosanoids  are not transported by the bloodstream to their sites of action but tend to act  locally, close to the cells that produced them. In fact, most eicosanoids de¬  compose within seconds or minutes, which limits their effects on nearby tis¬  sues. The synthesis of the eicosanoids is discussed in Section 20-6C.   In humans, the most important eicosanoid precursor is arachidonic acid,  a polyunsaturated fatty acid with four double bonds (Table 9-1). Arachidonate  is stored in cell membranes as the C2 ester of phosphatidylinositol (Table 9-2)  and other phospholipids. The fatty acid residue is released by the action of  phospholipase A 2 (Fig. 9-5).   The specific products of arachidonate metabolism are tissue-dependent.  For example, platelets produce thromboxanes almost exclusively, but endothe¬  lial cells that line the walls of blood vessels predominantly synthesize prosta¬  cyclins. Interestingly, thromboxanes stimulate vasoconstriction and platelet  aggregation (which helps initiate blood clotting), while prostacyclins elicit the  opposite effects. Thus, the two substances act in opposition to maintain a bal¬  ance in the cardiovascular system.   FIG. 9-13 Aggregates of single-tailed lipids. The tapered van  der Waals envelope of these lipids (a) permits them to pack efficiently  to form a spheroidal micelle (b). The diameter of the micelles depends  on the length of the tails. Spheroidal micelles composed of many    more lipid molecules than the optimal number (c) would have an  unfavorable water-filled center (blue). Such micelles could flatten out  to collapse the hollow center, but as these ellipsoidal micelles become  elongated (of), they also develop water-filled spaces.   The bilayer is a fluid structure in which lipids rapidly diffuse laterally.    In living systems, lipids are seldom found as free molecules but instead asso¬  ciate with other molecules, usually other lipids. In this section, we discuss how  lipids aggregate to form micelles and bilayers. We are concerned with the phys¬  ical properties of lipid bilayers because these aggregates form the structural  basis for biological membranes.    In aqueous solutions, amphiphilic molecules such as soaps and detergents form  micelles (globular aggregates whose hydrocarbon groups are out of contact  with water; Section 2-1C). This molecular arrangement eliminates unfavorable  contacts between water and the hydrophobic tails of the amphiphiles and yet  permits the solvation of the polar head groups.   The approximate size and shape of a micelle can be predicted from geo¬  metrical considerations. Single-tailed amphiphiles, such as soap anions, form  spheroidal or ellipsoidal micelles because of their tapered shapes (their hy¬  drated head groups are wider than their tails; Fig. 9-13a,b). The number of  molecules in such a micelle depends on the amphiphile, but for many sub¬  stances it is on the order of several hundred. Too few lipid molecules would  expose the hydrophobic core of the micelle to water, whereas too many would  give the micelle an energetically unfavorable hollow center (Fig. 9-13c). Of  course, a large micelle could flatten out to eliminate this hollow center, but  the resulting decrease of curvature at the flattened surfaces would also gener¬  ate empty spaces (Fig. 9-13^).   The two hydrocarbon tails of glycerophospholipids and sphingolipids give  these amphiphiles a somewhat rectangular cross section (Fig. 9-14 a). The steric  requirements of packing such molecules together yields large disklike micelles  (Fig. 9-1 4 b) that are really extended bimolecular leaflets. These lipid bilayers  are ~60 A thick, as measured by electron microscopy and X-ray diffraction  techniques, the value expected for more or less fully extended hydrocarbon tails.   A suspension of phospholipids (glycerophospholipids or sphingomyelins)  can form liposomes—closed, self-sealing solvent-filled vesicles that are   FIG. 9-14 Bilayer formation by  phospholipids. The cylindrical van der Waals  envelope of these lipids (a) causes them to form  extended disklike micelles (b) that are better  described as lipid bilayers.   FIG. 9-1 5 Electron micrograph of a liposome. Its wall, as the accompanying  diagram indicates, consists of a lipid bilayer. [Courtesy of Walther Stoeckenius, University of  California at San Francisco.]    bounded by only a single bilayer (Fig. 9-15 and page 241). They typically  have diameters of several hundred angstroms and, in a given preparation, are  rather uniform in size. Once formed, liposomes are quite stable and can be  purified by dialysis, gel filtration chromatography, or centrifugation.  Liposomes whose internal environment differs from the surrounding solution  can therefore be readily prepared. Liposomes serve as models of biological  membranes and also hold promise as vehicles for drug delivery since they are  absorbed by many cells through fusion with the plasma membrane.   FIG. 9-16 Phospholipid diffusion in a  lipid bilayer, (a) Transverse diffusion (a flip-flop)  is defined as the transfer of a phospholipid  molecule from one bilayer leaflet to the other.   (b) Lateral diffusion is defined as the pairwise  exchange of neighboring phospholipid molecules  in the same bilayer leaflet.   The transfer of a lipid molecule across a bilayer (Fig. 9- 16a), a process termed  transverse diffusion or a flip-flop y is an extremely rare event. This is because  a flip-flop requires the hydrated, polar head group of the lipid to pass through  the anhydrous hydrocarbon core of the bilayer. The flip-flop rates of phospho¬  lipids have half-times of several days or more. In contrast to their low flip-  flop rates, lipids are highly mobile in the plane of the bilayer (lateral diffusion;  Fig. 9-16£). It has been estimated that lipids in a membrane can diffuse the  l-|mm length of a bacterial cell in s. Because of the mobilities of the lipids,  the lipid bilayer can be considered to be a two-dimensional fluid.   The interior of the lipid bilayer is in constant motion due to rotations  around the C—C bonds of the lipid tails. Various physical measurements sug¬  gest that the interior of the bilayer has the viscosity of light machine oil. This  feature of the bilayer core is evident in molecular dynamics simulations, in  which the time-dependent positions of atoms are predicted from calculations  of the forces acting on them (Fig. 9-17). The viscosity of the bilayer increases  dramatically closer to the lipid head groups, whose rotation is limited and  whose lateral mobility is more constrained by interactions between other po¬  lar or charged head groups.   FIG. 9-1 7 Model (snapshot) of a lipid bilayer at an instant in time. The   conformations of dipalmitoylphosphatidylcholine molecules in a bilayer surrounded by water  were modeled by computer. Atom colors are chain and glycerol C gray except terminal  methyl C yellow, ester 0 red, phosphate P and 0 green, and choline C and N magenta.  Water molecules are represented by translucent blue spheres (those near the bilayer  appear dark because they overlap head group atoms). [Courtesy of Richard Pastor and  Richard Venable, NIH, Bethesda, Maryland.]   FIG. 9-1 8 Phase transition in a lipid bilayer, (a) Above the  transition temperature, both the lipid molecules as a whole and  their nonpolar tails are highly mobile in the plane of the bilayer.   (b) Below the transition temperature, the lipid molecules form a  much more orderly array to yield a gel-like solid. [After Robertson,    How would changes in fatty acid chain length and degree  of saturation affect the bilayer’s transition temperature?    Note that the hydrophobic tails of the lipids shown in Fig. 9-17 are not  stiffly regimented as Fig. 9-16 might suggest, but instead bend and interdigitate.  A typical biological membrane includes many different lipid molecules, some  of whose tails are of different lengths or are kinked due to the presence of dou¬  ble bonds. Under physiological conditions, highly mobile chains fill any gaps  that might form between lipids in the bilayer interior.   The model bilayer shown in Fig. 9-17 indicates that the phospholipids  bob up and down to some degree. This is also the case in naturally occurring  membranes, which contain a variety of different lipid head groups that must  nestle among each other. The polar nature of the outer surface of the bilayer  extends from the head groups to the carbonyl groups of the ester and amide  bonds that link the fatty acyl chains. Consequently, water molecules penetrate  a lipid bilayer to a depth of up to 15 A, avoiding only the central ~30 A  hydrocarbon core.    The Fluidity of a Lipid Bilayer Is Temperature-Dependent. As a lipid bilayer  cools below a characteristic transition temperature, it undergoes a sort of phase  change in which it becomes a gel-like solid; that is, it loses its fluidity (Fig. 9-18).  Above the transition temperature, the highly mobile lipids are in a state known  as a liquid crystal because they are ordered in some directions but not in oth¬  ers. The bilayer is thicker in the gel state than in the liquid crystal state due  to the stiffening of the hydrocarbon tails at lower temperatures.   The transition temperature of a bilayer increases with the chain length and  the degree of saturation of its component fatty acid residues for the same reasons  that the melting points of fatty acids increase with these quantities. The transi¬  tion temperatures of most biological membranes are in the range 10 to 40°C.  Bacteria and cold-blooded animals such as fish modify (through lipid synthe¬  sis and degradation) the fatty acid compositions of their membrane lipids with  ambient temperature so as to maintain a constant level of fluidity. Thus, the flu¬  idity of biological membranes is one of their important physiological attributes.   Cholesterol, which by itself does not form a bilayer, decreases membrane flu¬  idity because its rigid steroid ring system interferes with the motions of the fatty  acid side chains in other membrane lipids. It also broadens the temperature range  of the phase transition. This is because cholesterol inhibits the ordering of fatty  acid side chains by fitting in between them. Thus, cholesterol functions as a  kind of membrane plasticizer.   • Explain why lateral diffusion of membrane  lipids is faster than transverse diffusion.   • Integral membrane proteins contain a transmembrane structure consisting of  a helices or a p barrel with a hydrophobic surface.   • Lipid-linked proteins have a covalently attached prenyl group, fatty acyl group, or  glycosylphosphatidylinositol group.   • Peripheral membrane proteins interact noncovalently with proteins or lipids at the  membrane surface.   Biological membranes contain proteins as well as lipids. The exact lipid and  protein components and the ratio of protein to lipid varies with the identity  of the membrane. For example, the lipid-rich myelinated membranes that sur¬  round and insulate certain nerve axons (Fig. 9-8) are only 19% protein by  mass, whereas the protein-rich inner membrane of mitochondria, which me¬  diates numerous chemical reactions, is 76% protein by mass. Eukaryotic  plasma membranes are typically ~50% protein by mass.   Membrane proteins catalyze chemical reactions, mediate the flow of nu¬  trients and wastes across the membrane, and participate in relaying informa¬  tion about the extracellular environment to various intracellular components.  Such proteins carry out their functions in association with the lipid bilayer.  They must therefore interact to some degree with the hydrophobic core and/or  the polar surface of the bilayer. In this section, we examine the structures of  some membrane proteins, which are classified by their mode of interaction  with the membrane.   Integral or intrinsic proteins (Fig. 9-19) associate tightly with membranes  through hydrophobic interactions and can be separated from membranes only  by treatment with agents that disrupt membranes. For example, detergents  such as sodium dodecyl sulfate (Section 5-2D) solubilize membrane proteins  by replacing the membrane lipids that normally surround the protein. The hy¬  drophobic portions of the detergent molecules coat the hydrophobic regions  of the protein, and the polar head groups render the detergent—protein com¬  plex soluble in water. Chaotropic agents such as guanidinium ion and urea  (Section 6-4B) disrupt water structure, thereby reducing the hydrophobic ef¬  fect, the primary force stabilizing the association of the protein with the mem¬  brane. Some integral proteins bind lipids so tenaciously that they can be freed  from them only under denaturing conditions.   Once they have been solubilized, integral proteins can be purified by many  of the protein fractionation methods described in Section 5-2. Since these pro-    FIG. 9-19 X-Ray structure of the integral membrane protein  aquaporin-0 (AQPO) in association with lipids. The protein is  represented by its surface diagram, which is colored according to charge  (red negative, blue positive, and white uncharged). Tightly bound molecules  of dimyristoylphosphatidylcholine are drawn in space-filling form with  C green, 0 red, and P orange. Note how the lipid tails closely conform to  the nonpolar surface of the protein, thereby solvating it. The arrangement  of the two rows of lipid molecules, with phosphate-phosphate distances of  —35 A, matches the dimensions of a lipid bilayer. [Based on an electron  crystallographic structure by Stephen Harrison and Thomas Walz, Harvard  Medical School. PDBid 2B60.]   quently requires the presence of detergents or water-miscible organic solvents  such as butanol or glycerol.   Integral Proteins Are Asymmetrically Oriented Amphiphiles. Integral pro¬  teins are amphiphiles; the protein segments immersed in a membrane's nonpolar  interior have predominantly hydrophobic surface residues , whereas those portions  that extend into the aqueous environment are by and large sheathed with polar  residues. This was first demonstrated through surface labeling, a technique  employing agents that react with proteins but cannot penetrate membranes.   For example, the extracellular domain of an integral protein binds antibodies  elicited against it, but its cytoplasmic domain will do so only if the membrane  has been ruptured. Membrane-impermeable protein-specific reagents that are  fluorescent or radioactively labeled can be similarly employed. Alternatively,  proteases, which digest only the solvent-exposed portions of an integral pro¬  tein, may be used to identify the membrane-immersed portions of the pro¬  tein. These techniques revealed, for example, that the erythrocyte membrane  protein glycophorin A has three domains (Fig. 9-20): (1) a 72-residue externally  located N-terminal domain that bears 16 carbohydrate chains; (2) a 19-residue  sequence, consisting almost entirely of hydrophobic residues, that spans the  erythrocyte cell membrane; and (3) a 40-residue cytoplasmic C-terminal domain  that has a high proportion of charged and polar residues. Thus, glycophorin A  is a transmembrane (TM) protein; that is, it completely spans the membrane.   FIG. 9-20 Human erythrocyte glycophorin A. The protein bears  15 0-linked oligosaccharides ( green diamonds) and one that is  AHinked ( dark green hexagon) on its extracellular domain. The  predominant sequence of the 0-linked oligosaccharides is also shown  (NeuNAc = A/-acetylneuraminic acid). The protein’s transmembrane  portion ( brown and purple) consists of 19 sequential predominantly    hydrophobic residues. Its C-terminal portion, which is located on the  membrane’s cytoplasmic face, is rich in anionic (pink), cationic (blue),  and polar (blue-gray) residues. There are two common genetic variants  of glycophorin A: Glycophorin A M has Ser and Gly at positions 1 and  5, whereas glycophorin A N has Leu and Glu at these positions. [After  Marchesi, V.T., Semin. Hematol. 16, 8 (1979).]    FIG. 9-21 Identification of glycophorin  A’s transmembrane domain. The calculated  free energy change in transferring 20-residue-  long a-helical segments from the interior of a  membrane to water is plotted against the  position of the segment’s first residue. Peaks  higher than +85 kJ • mol -1 indicate a  transmembrane helix. [After Engelman, D.M.,  Steitz, T.A., and Goldman, A., Annu. Rev. Biophys.  Biophys. Chem. 15, 343 (1986).]    Studies of a variety of biological membranes have established that biolog¬  ical membranes are asymmetric in that a particular membrane protein is invari¬  ably located on only one particular face of a membrane , or in the case of a  transmembrane protein ., oriented in only one direction with respect to the mem¬  brane. However, no protein is known to be completely buried in a membrane;  that is, all membrane-associated proteins are at least partially exposed to the  aqueous environment.   Transmembrane Proteins May Contain a Helices. In order for a polypeptide  chain to penetrate or span the lipid bilayer, it must have hydrophobic side chains  that contact the lipid tails and it must shield its polar backbone groups. This  second requirement is met by the formation of secondary structure that satisfies  the hydrogen-bonding capabilities of the polypeptide backbone. Consequently,  all known TM segments of integral membrane proteins consist of either a he¬  lices or (3 sheets. For example, glycophorin As 19-residue TM sequence almost  certainly forms an a helix. The existence of such a TM helix can be predicted  by comparing the free energy change in transferring a-helical polypeptide seg¬  ments from the nonpolar interior of a membrane to water (Fig. 9-21).  Alternatively, a potential TM sequence can be identified by reference to hydropa¬  thy indices such as those in Table 6-3. Methods for predicting the position of  a TM a helix are useful because the difficulty in crystallizing integral membrane  proteins has permitted relatively few of their X-ray structures to be determined.   Nigel Unwin and Richard Henderson used electron crystallography to de¬  termine the structure of the integral membrane protein bacteriorhodopsin (Box  9-2). This 247-residue homotrimeric protein, which is produced by the  halophilic (salt-loving) archaebacterium Halobacterium salinarum (it grows best  in 4.3 M NaCl), is a light-driven proton pump: It generates a proton concen¬  tration gradient across the cell membrane that powers ATP synthesis by a mech¬  anism discussed in Section 18-3B. A covalently bound retinal (Section 9-IF) is  the proteins light-absorbing group. Bacteriorhodopsin consists largely of a bun¬  dle of seven ~25-residue a-helical rods that span the lipid bilayer in directions  almost perpendicular to the bilayer plane (Fig. 9-22). As expected, the amino  acid side chains that contact the lipid tails are highly hydrophobic. Successive  membrane-spanning helices are connected in head-to-tail fashion by hydrophilic  loops of varying size. This arrangement places the proteins charged residues near  the surfaces of the membrane in contact with the aqueous environment.    FIG. 9-22 The structure of  bacteriorhodopsin. The protein is shown in  ribbon form as viewed from within the  membrane plane and colored in rainbow order  from its N-terminus (blue) to its C-terminus  (red). Its covalently bound retinal is drawn in  stick form (magenta). [Based on an X-ray  structure by Nikolaus Grigorieff and Richard  Henderson, MRC Laboratory of Molecular  Biology, Cambridge, U.K. PDBid 2BRD.] O  See Kinemage Exercise 8-1.    Richard Henderson (1945-) Richard  Henderson, like a number of other  pioneering structural biologists, began  his career as a physicist. He turned his  attention to membrane proteins because  of their importance in cellular metabolic,  transport, and signaling phenomena.  Unlike globular proteins, which are  soluble in aqueous solution from which  they can often be crystallized, integral  membrane proteins aggregate in  aqueous solution and hence can only be  kept in solution by the presence of a suitable detergent. In spite  of this, such solubilized proteins rarely crystallize in a manner  suitable for X-ray analysis (and none had been made to do so at  the time that Henderson began his studies). To circumvent this  obstacle, Henderson adapted the technique of electron  crystallography to macromolecules, using the membrane protein  bacteriorhodopsin as his subject.   Bacteriorhodopsin was discovered in 1967, and its function as  a proton pump was described shortly thereafter. The protein is  synthesized by halophilic archaebacteria such as H. salinarum  (formerly known as H. halobium). In addition to bacteriorhodopsin,  the archaeal rhodopsin family includes chloride-pumping and  sensory proteins. Similar proteins have also been identified in  eubacteria and in unicellular eukaryotes. What made  bacteriorhodopsin attractive as a research subject is its relatively  small size (248 residues), its stability, and-most importantly-its  unusual proclivity to form ordered two-dimensional arrays in the  bacterial cell membrane. In H. salinarum, such arrays, which occur  as 0.5-fxm-wide patches, are known as purple membranes due to  the color of the protein’s bound retinal molecule. Each purple  membrane patch, which is essentially a two-dimensional crystal,  consists of 75% bacteriorhodopsin and 25% lipid.   At the time that Henderson began his structural studies of  bacteriorhodopsin, the X-ray structures of only around a dozen  different globular proteins had been reported. Henderson, working  with Nigel Unwin, adapted the principles of X-ray crystallography  to the two-dimensional bacteriorhodopsin crystals by measuring  the diffraction intensities generated by the electron beam from an  electron microscope impinging on a purple membrane patch (as  explained by the wave-particle duality, electrons, like all particles,  have wavelike properties with a wavelength, X = h/mv, where h is  Planck’s constant, m is the particle mass, and v is its velocity). It  was necessary to use an electron beam rather than X-rays  because the electron microscope can focus its electron beam on  the microscopically small purple membrane patches. Nevertheless,    only very low electron beam intensities could be used because  otherwise the resulting radiation damage would destroy the purple  membrane. Consequently, to obtain diffraction data of sufficiently  high signal-to-noise ratio, the diffraction patterns of around one  hundred or more purple membrane samples had to be averaged.   Working at the limits of the available technology, Henderson  and Unwin, in 1975, published a low-resolution model for the  structure of bacteriorhodopsin. This was the first glimpse of an  integral membrane protein. Its seven transmembrane helices were  clearly visible as columns of electron density that were  approximately perpendicular to the plane of the membrane.  However, to obtain three-dimensional diffraction data, a two-  dimensional crystal must be systematically tilted relative to the  electron beam. Mechanical limitations that prevented the sample  from being tilted to the degree necessary to obtain a full three-  dimensional data set as well as other technical difficulties  therefore yielded a model that had a resolution of 7 A in the  plane of the membrane but only 14 A in a perpendicular direction.  Consequently, the polypeptide loops connecting the seven helices  could not be discerned, nor were the protein’s associated retinal  molecule or any of its side chains visible. However, over the next  15 years, developments in electron microscopy, such as the use of  better electron sources and liquid-helium temperatures to minimize  radiation damage to the sample, permitted Henderson to extend  the resolution of the bacteriorhodopsin structure to 3.5 A in the  plane of the membrane and 10 A in the perpendicular direction.  The resulting electron density map clearly revealed the positions  of several bulky aromatic residues and the bound retinal and  permitted the loops connecting the transmembrane helices to be  visualized. Thus, electron crystallography has become a useful tool  for determining the structures of a variety of proteins that can be  induced to form two-dimensional arrays as well as those that form  very thin three-dimensional crystals.   Henderson’s groundbreaking work on bacteriorhodopsin made  a seminal contribution to the growing body of biochemical,  genetic, spectroscopic, and structural studies of bacteriorhodopsin.  This information, together with more recent high-resolution X-ray  structures derived from three-dimensional crystals of  bacteriorhodopsin, obtained by crystallizing the protein in a lipid  matrix, have led to a detailed understanding of the mechanism of  the light-induced structural changes through which  bacteriorhodopsin pumps protons out of the bacterial cell.   Henderson, R. and Unwin, P.N., Three-dimensional model of purple membrane  obtained by electron microscopy, Nature 257 , 28-32 (1975).   Henderson, R., Baldwin, J.M., Ceska, T.A., Zemlin, R, Beckmann, E., and Downing, K.H.,  Model for the structure of bacteriorhodopsin based on high-resolution electron  cryomicroscopy, J. Mol. Biol. 213 , 899-929 (1990).    Hydrophobic effects, as we saw in Section 6-4A, are the dominant forces  stabilizing the three-dimensional structures of water-soluble globular proteins.  However, since the TM regions of integral membrane proteins are immersed  in nonpolar environments, what stabilizes their structures? Analysis of integral  protein structures indicates that their interior residues have hydrophobicities  comparable to those of water-soluble proteins. However, the membrane-  exposed residues of these proteins, on average, are even more hydrophobic than  their interior residues. Evidently, the structures of both integral and water-soluble   Chapter 9 Lipids and Biological Membranes proteins are stabilized by the exclusion of their interior residues from the surround¬   ing solvent , but in the case of integral proteins , the solvent is the lipid bilayer. In  addition, the low polarity and anhydrous environments of transmembrane  proteins are likely to strengthen their hydrogen bonds relative to those of sol¬  uble proteins.   Some Transmembrane Proteins Contain p Barrels. A protein segment im¬  mersed in the nonpolar interior of a membrane must fold so that it satisfies  the hydrogen-bonding potential of its polypeptide backbone. An a helix can  do so as can an antiparallel P sheet that rolls up to form a barrel (a (3 barrel;  Section 6-2C). Transmembrane p barrels of known structure consist of 8 to  22 strands. The number of strands must be even to permit the P sheet to close  up on itself with all strands antiparallel.   P Barrels occur in porins, which are channel-forming proteins in the outer  membrane of gram-negative bacteria (Section 8-3B). The outer membrane  protects the bacteria from hostile environments while the porins permit the  entry of small polar solutes such as nutrients. Porins also occur in eukaryotes  in the outer membranes of mitochondria and chloroplasts (consistent with the  descent of these organelles from free-living gram-negative bacteria; Section 1-2C).   Bacterial porins are monomers or trimers of identical 30- to 50-kD sub¬  units. X-Ray structural studies show that most porin subunits consist largely   (a) A ribbon diagram of the 16-stranded monomer colored in rainbow  order from its N-terminus (blue) to its C-terminus (red), (b) Ribbon  diagram of the trimer embedded in its semitransparent surface and  viewed along its 3-fold axis of symmetry from the cell’s exterior,  showing the pore through each subunit Each subunit is differently  colored. Adjacent |3 strands in adjoining subunits extend essentially  perpendicularly to each other, (c) A space-filling model of the trimer  viewed perpendicular to its 3-fold axis. N atoms are blue, 0 atoms are  red, and C atoms are green, except those of Trp and Tyr side chains,  which are white. These latter groups delimit an ~25-A-high  hydrophobic band (scale at right) that is immersed in the nonpolar  portion of the bacterial outer membrane (with the cell’s exterior at the  tops of Parts a and c). [Based on an X-ray structure by Johan  Jansonius, University of Basel, Switzerland. PDBid 10PF.] Q See  Kinemage Exercise 8-3.    of at least a 16-stranded antiparallel (3 barrel that forms a solvent-accessible  central channel with a length of ~55 A and a minimum diameter of ~7 A  (Fig. 9-23). As expected, the side chains of the proteins membrane-exposed  surface are nonpolar, thereby forming an ~25-A-high hydrophobic band  encircling the trimer (Fig. 9-23 c). This band is flanked by more polar aromatic  side chains (Table 6-3) that form interfaces with the head groups of the lipid  bilayer (Fig. 9-23 c). In contrast, the side chains at the solvent-exposed surface  of the protein, including those lining the walls of the aqueous channel, are  polar. Possible mechanisms for the solute selectivity of porins are discussed in  Section 10 - 2 B.   Some membrane-associated proteins contain covalently attached lipids that an¬  chor the protein to the membrane. The lipid group, like any modifying group,  may also mediate protein—protein interactions or modify the structure and ac¬  tivity of the protein to which it is attached. Lipid-linked proteins come in  three varieties: prenylated proteins, fatty acylated proteins, and glycosylphos-  phatidylinositol-linked proteins. A single protein may contain more than one  covalently linked lipid group.   Prenylated proteins have covalently attached lipids that are built from  isoprene units (Section 9 -IF). The most common isoprenoid groups are the  Ql 5 farnesyl and C 2 o geranylgeranyl residues.    The most common prenylation site in proteins is the C-terminal tetrapeptide  C-X-X-Y, where C is Cys and X is often an aliphatic amino acid residue.  Residue Y influences the type of prenylation: Proteins are farnesylated when  Y is Ala, Met, or Ser and geranylgeranylated when Y is Leu. In both cases, the  prenyl group is enzymatically linked to the Cys sulfur atom via a thioether  linkage. The X-X-Y tripeptide is then proteolytically excised, and the newly ex¬  posed terminal carboxyl group is esterified with a methyl group, producing a  C-terminus with the structure shown.   Two kinds of fatty acids, myristic acid and palmitic acid, are linked to  membrane proteins. Myristic acid, a biologically rare saturated C 14 fatty acid,  is appended to a protein via an amide linkage to the a-amino group of an N-  terminal Gly residue. Myristoylation is stable: The fatty acyl group remains  attached to the protein throughout its lifetime. Myristoylated proteins are lo¬  cated in a number of subcellular compartments, including the cytosol, endo¬  plasmic reticulum, plasma membrane, and the nucleus.   represent fatty acid residues whose identities vary with the protein. The tetrasaccharide  may have a variety of attached sugar residues whose identities also vary.    In palmitoylation, the saturated C 1( 5 fatty acid palmitic acid is joined in  thioester linkage to a specific Cys residue. Palmitoylated proteins occur almost  exclusively on the cytoplasmic face of the plasma membrane, where many par¬  ticipate in transmembrane signaling. The palmitoyl group can be removed by  the action of palmitoyl thioesterases, suggesting that reversible palmitoylation  may regulate the association of the protein with the membrane and thereby  modulate the signaling processes.   occur in all eukaryotes but are particularly abundant in some parasitic proto¬  zoa, which contain relatively few membrane proteins anchored by transmem¬  brane polypeptide segments. Like glycoproteins and glycolipids, GPI-linked  proteins are located only on the exterior surface of the plasma membrane.   The core structure of the GPI group consists of phosphatidylinositol  (Table 9-2) glycosidically linked to a linear tetrasaccharide composed of three  mannose residues and one glucosaminyl residue (Fig. 9-24). The mannose at  the nonreducing end of this assembly forms a phosphodiester bond with a  phosphoethanolamine residue that is amide-linked to the proteins C-terminal  carboxyl group. The core tetrasaccharide is generally substituted with a variety  of sugar residues that vary with the identity of the protein. There is likewise  considerable diversity in the fatty acid residues of the phosphatidylinositol group.   • What are the two types of secondary  structures that occur in transmembrane  proteins?   • Describe the covalent modifications of  lipid-linked proteins. Why might some of  these modifications be reversible?    Peripheral or extrinsic proteins, unlike integral membrane proteins or lipid-  linked proteins, can be dissociated from membranes by relatively mild proce¬  dures that leave the membrane intact, such as exposure to high ionic strength  salt solutions or pH changes. Peripheral proteins do not bind lipid and, once  purified, behave like water-soluble proteins. They associate with membranes  by binding at their surfaces, most likely to certain lipids or integral proteins,  through electrostatic and hydrogen-bonding interactions. Cytochrome c  (Sections 5-4A, 6-2D, and 18-2E) is a peripheral membrane protein that is  associated with the outer surface of the inner mitochondrial membrane. At  physiological pH, cytochrome c is cationic and can interact with negatively  charged phospholipids such as phosphatidylserine and phosphatidylglycerol.   • The dynamic arrangement and interactions of membrane lipids and proteins are  described by the fluid mosaic model.   • The membrane skeleton gives the cell shape yet is flexible.   • Lipids are not distributed uniformly throughout a membrane and may form rafts.   • The secretory pathway describes the transmembrane passage of membrane and  secreted proteins.   • Different types of coated vesicles transport proteins between cellular compartments.   Membranes were once thought to consist of a phospholipid bilayer sandwiched  between two layers of unfolded polypeptide. This sandwich model, which is  improbable on thermodynamic grounds, was further discredited by electron  microscopic visualization of membranes and other experimental approaches.  More recent studies have revealed insights into the fine structure of mem¬  branes, including a surprising degree of heterogeneity. In this section, we con¬  sider the arrangement of membrane proteins and lipids and examine some of  the mechanisms by which these components move throughout the cell.   The demonstrated fluidity of artificial lipid bilayers (Section 9-2B) suggests that  biological membranes have similar properties. This idea was proposed in 1972  by S. Jonathan Singer and Garth Nicolson in their unifying theory of mem¬  brane structure known as the fluid mosaic model. In this model, integral pro¬  teins are visualized as “icebergs” floating in a two-dimensional lipid “sea” in a  random or mosaic distribution (Fig. 9-25). A key element of the model is that  integral proteins can diffuse laterally in the lipid matrix unless their move¬  ments are restricted by association with other cell components. This model of    FIG. 9-25 Diagram of a plasma membrane. Integral proteins  (purple ) are embedded in a bilayer composed of phospholipids ( blue  head groups attached to wiggly tails ) and cholesterol (yellow). The  carbohydrate components ( green and brown beads) of glycoproteins    and glycolipids occur on only the external face of the membrane.  Most membranes contain a higher proportion of protein than is  depicted here.   Chapter 9 Lipids and Biological Membranes membrane fluidity explained the earlier experimental results of Michael   Edidin, who fused cultured cells and observed the intermingling of their dif¬  ferently labeled cell-surface proteins (Fig. 9-26).   The rates of diffusion of proteins in membranes can be determined from  measurements of fluorescence recovery after photobleaching (FRAP). In  this technique, a fluorophore (fluorescent group) is specifically attached to a  membrane component in an immobilized cell or in an artificial membrane  system. An intense laser pulse focused on a very small area (~3 |Jim 2 ) destroys  (bleaches) the fluorophore there (Fig. 9-27). The rate at which the bleached  area recovers its fluorescence, as monitored by fluorescence microscopy, indi¬  cates the rate at which unbleached and bleached fluorophore-labeled molecules  laterally diffuse into and out of the bleached area.    The cell-surface  proteins of cultured  mouse and human  cells are labeled (l  with green and red  fluorescent  markers.   accompanying photomicrographs (in square boxes) were taken  through filters that allowed only red or green light to reach the    camera. [Immunofluorescence photomicrographs courtesy of Michael  Edidin, The Johns Hopkins University.]   FIG. 9-27 The fluorescence recovery after photobleaching  (FRAP) technique, (a) An intense laser light pulse bleaches the  fluorescent markers (green) from a small region of an immobilized cell  that has a fluorophore-labeled membrane component, (b) The    fluorescence of the bleached area recovers as the bleached  molecules laterally diffuse out of it and intact fluorescent molecules  diffuse into it. (c) The fluorescence recovery rate depends on the  diffusion rate of the labeled molecule.    FRAP measurements demonstrate that membrane proteins vary in their  lateral diffusion rates. Some 30 to 90% of these proteins are freely mobile;  they diffuse at rates only an order of magnitude or so slower than those of  the much smaller lipids, so they can diffuse the 20-|Jim length of a eukaryotic  cell within an hour. Other proteins diffuse more slowly, and some are essen¬  tially immobile due to submembrane attachments.   Studies of membrane structure and composition often make use of erythro¬  cyte membranes, since these are relatively simple and easily isolated. A mature  mammalian erythrocyte lacks organelles and carries out few metabolic  processes; it is essentially a membranous bag of hemoglobin. Erythrocyte mem¬  branes can be obtained by osmotic lysis, which causes the cell contents to leak  out. The resulting membranous particles are known as erythrocyte ghosts  because, on return to physiological conditions, they reseal to form colorless  particles that retain their original shape but are devoid of cytoplasm.   A normal erythrocytes biconcave disklike shape (Fig. 7-1 Id) ensures the rapid  diffusion of 0 2 to its hemoglobin molecules by placing them no farther than 1  [xm from the cell surface. However, the rim and the dimple regions of an ery¬  throcyte do not occupy fixed positions on the cell membrane. This can be demon¬  strated by anchoring an erythrocyte to a microscope slide by a small portion of  its surface and inducing the cell to move laterally with a gentle flow of buffer. A  point originally on the rim of the erythrocyte will move across the dimple to the  rim on the opposite side of the cell. Evidently, the membrane rolls across the cell  while maintaining its shape, much like the tread of a tractor. This remarkable  mechanical property of the erythrocyte membrane results from the presence of a  sub membranous network of proteins that function as a membrane “skeleton.”   The fluidity and flexibility imparted to an erythrocyte by its membrane  skeleton have important physiological consequences. A slurry of solid particles  of a size and concentration equal to that of red cells in blood has flow char¬  acteristics approximating those of sand. Consequently, in order for blood to  flow at all, much less for its erythrocytes to squeeze through capillary blood  vessels smaller in diameter than they are, erythrocyte membranes, with their  membrane skeletons, must be fluidlike and easily deformable.    FIG. 9-28 Structure of spectrin, (a) Structure of an ap dimer. Both of these anti¬  parallel polypeptides contain multiple 106-residue repeats, which are thought to form flexibly  connected triple-helical bundles. Two of these heterodimers join, head to head, to form an  (ap) 2 heterotetramer. [After Speicher, D.W. and Marchesi, V. Nature 311, 177 (1984).]   (b) The X-ray structure of two consecutive repeats of chicken brain a-spectrin. Each of  these 106-residue repeats consists of an up-down-up triple-helical bundle in which the  C-terminal helix of the first repeat (red) is continuous, via a 5-residue helical linker (green),  with the N-terminal helix of the second repeat (blue). The helices within each triple-helical  bundle wrap around each other in a gentle left-handed supercoil. [Courtesy of Alfonso  Mondragon, Northwestern University. PDBid 1CUN.]    The protein spectrin, so called because it was discovered in erythrocyte  ghosts, accounts for —75% of the erythrocyte membrane skeleton. It is com¬  posed of two similar polypeptide chains, a 280-kD a subunit and a 246-kD  (3 subunit, which each consist of repeating 106-residue segments that fold into  triple-stranded a-helical coiled coils (Fig. 9-28). These large polypeptides are  loosely intertwined to form a flexible wormlike a(3 dimer that is — 1000 A  long. Two such heterodimers further associate in a head-to-head manner to  form an (a(3) 2 tetramer. Spectrin is a homolog of dystrophin, the muscle pro¬  tein that is defective in muscular dystrophy (Section 7-2A).   r There are —100,000 spectrin tetramers per cell, and they are cross-linked  at both ends by attachments to other cytoskeletal proteins. Together, these  proteins form a dense and irregular protein meshwork that underlies the ery¬  throcyte plasma membrane (Fig. 9-29). A defect or deficiency in spectrin syn¬  thesis causes hereditary spherocytosis, in which erythrocytes are spheroidal  and relatively fragile and inflexible. Individuals with the disease suffer from  anemia due to erythrocyte lysis and the removal of spherocytic cells by the  spleen (which normally functions to filter out aged and hence inflexible ery¬  throcytes from the blood at the end of their —120-day lifetimes).   Spectrin also associates with an 1880-residue protein known as ankyrin,  which binds to an integral membrane ion channel protein. This attachment  anchors the membrane skeleton to the membrane. Immunochemical studies  have revealed spectrin-like and ankyrin-like proteins in a variety of tissues, in  addition to erythrocytes. Ankyrin s N-terminal 798-residue segment consists  almost entirely of 24 tandem —33-residue repeats known as ankyrin repeats  (Fig. 9-30), which also occur in a variety of other proteins. Each ankyrin re¬  peat consists of two short (8- or 9-residue) antiparallel a helices followed by  a long loop. These structures are arranged in a right-handed helical stack. The  entire assembly forms an elongated concave surface that is postulated to bind  various integral proteins as well as spectrin.   The interaction of membrane components with the underlying skeleton  helps explain why integral membrane proteins exhibit different degrees of mo¬  bility within the membrane: Some integral proteins are firmly attached to el¬  ements of the cytoskeleton or are trapped within the spaces defined by those  “fences.” Other membrane proteins may be able to squeeze through gaps or  “gates” between cytoskeletal components, whereas still other proteins can dif¬  fuse freely without interacting with the cytoskeleton at all (Fig. 9-31). Support  for this gates and fences model comes from the finding that partial destruc¬  tion of the cytoskeleton results in freer protein diffusion.   (a) An electron micrograph of an erythrocyte membrane skeleton that  has been stretched to an area 9 to 10 times greater than that of the  native membrane. Stretching makes it possible to obtain clear images  of the membrane skeleton, which in its native state is densely packed  and irregularly flexed. Note the predominantly hexagonal network  composed of spectrin tetramers. [Courtesy of Daniel Branton, Harvard    University.] (b) A model of the erythrocyte membrane skeleton with an  inset showing its relationship to the intact erythrocyte. The junctions  between spectrin tetramers include actin and tropomyosin (Section  7-2) and band 4.1 protein (named after its position in an SDS-PAGE  electrophoretogram). [After Goodman, S.R., Krebs, K.E., Whitfield, C.F.,  Riederer, B.M., and Zagen, I.S., CRC Crit. Rev. Biochem. 23, 196  (1988).]   to 24. The polypeptide is shown in ribbon form colored in rainbow  order from its N-terminus (blue, repeat 13) to its C-terminus (red,  repeat 24). [Based on an X-ray structure by Peter Michaely, University  of Texas Southwestern Medical Center, Dallas, Texas. PDBid IN11.]   FIG. 9-31 Model rationalizing the various mobilities of  membrane proteins. Protein A, which interacts tightly with the  underlying cytoskeleton, is immobile. Protein B is free to rotate within  the confines of the cytoskeletal “fences.” Protein C diffuses by  traveling through “gates” in the cytoskeleton. The diffusion of some  membrane proteins is not affected by the cytoskeleton. [After Edidin,  M., Trends Cell Biol. 2, 378 (1992).]    FIG. 9-32 Asymmetric distribution of  membrane phospholipids in the human  erythrocyte membrane. The phospholipid  content is expressed as mol %. [After Rothman,  J.E. and Lenard, J., Science 194, 1744 (1977).]   FIG. 9-33 Electron micrograph of  endoplasmic reticulum. This membrane  network is contiguous with the nuclear  membrane. The so-called smooth ER is the  site of synthesis of membrane lipids, and the  rough ER, with its associated ribosomes, is the  site of synthesis of membrane and secretory  proteins. [© Omikron/Photo Researchers, Inc.]   The lipid and protein components of membranes do not occur in equal pro¬  portions on the two sides of biological membranes. For example, membrane  glycoproteins and glycolipids are invariably oriented with their carbohydrate moi¬  eties facing the cells exterior. The asymmetric distribution of certain membrane  lipids between the inner and outer leaflets of a membrane was first established  through the use of phospholipases (Section 9-1C). Phospholipases cannot pass  through membranes, so phospholipids on only the external surface of intact  cells are susceptible to hydrolysis by these enzymes. Such studies reveal that  lipids in biological membranes are asymmetrically distributed (e.g., Fig. 9-32).  How does this asymmetry arise?   In eukaryotes, the enzymes that synthesize membrane lipids are mostly in¬  tegral membrane proteins of the endoplasmic reticulum (ER; the intercon¬  nected membranous vesicles that occupy much of the cytosol; Fig. 9-33),  whereas in prokaryotes, lipids are synthesized by integral membrane proteins  in the plasma membrane. Hence, membrane lipids are fabricated on site.  Eugene Kennedy and James Rothman demonstrated this to be the case in bac¬  teria through the use of selective labeling. They gave growing bacteria a 1-  minute pulse of 32 PC >4 in order to radioactively label the phosphoryl groups  of only the newly synthesized phospholipids. Immediately afterward, they  added trinitrobenzenesulfonic acid (TNBS), a membrane-impermeable  reagent that combines with phosphatidylethanolamine (PE; Fig. 9-34). Analysis  of the resulting doubly labeled membranes showed that none of the TNBS-  labeled PE was radioactively labeled. This observation indicates that newly made  PE is synthesized on the cytoplasmic face of the membrane (Fig. 9-35, upper right).   However, if an interval of only 3 minutes was allowed to elapse between  the 32 PC >4 pulse and the TNBS addition, about half of the 32 P-labeled PE  was also TNBS labeled (Fig. 9-35, lower right)- This observation indicates that  the flip-flop rate of PE in the bacterial membrane is ~ 100,000-fold greater  than it is in bilayers consisting of only phospholipids (where the flip-flop rates  have half-times of many days).   How do phospholipids synthesized on one side of the membrane reach its  other side so quickly? Phospholipid flip-flops in bacteria as well as eukaryotes  appear to be facilitated in two ways:   1. Membrane proteins known as flippases catalyze the flip-flops of specific  phospholipids. These proteins tend to equilibrate the distribution of  their corresponding phospholipids across a bilayer; that is, the net trans¬  port of a phospholipid is from the side of the bilayer with the higher  concentration of the phospholipid to the opposite side. Such a process,  as we will see in Section 10-1, is a form of facilitated diffusion.   2. Membrane proteins known as phospholipid translocases transport  specific phospholipids across a bilayer in a process that is driven by    ATP hydrolysis. These proteins can transport certain phospholipids  from the side of a bilayer that has the lower concentration of the  phospholipid to the opposite side, thereby establishing a nonequilib¬  rium distribution of the phospholipid. Such a process, as we will see  in Section 10-3, is a form of active transport.   The observed distribution of phospholipids across membranes (e.g., Fig. 9-32)  therefore appears to arise from the membrane orientations of the enzymes that    FIG. 9-35 The location of lipid synthesis in a  bacterial membrane. Newly synthesized PE was labeled  by a 1-min pulse of 32 P0 3_ (yellow head groups), and the  PE on the cell surface was independently labeled by  treatment with the membrane-impermeable reagent TNBS.  When TNBS labeling (red circles) occurred  immediately after the 32 P pulse, none of the 32 P-labeled  PE was TNBS labeled (upper right), thereby indicating that  the PE is synthesized on the cytoplasmic face of the  membrane. If, however, there was even a few minutes’  delay between the two labeling procedures, much of the  TNBS-labeled PE in the external face of the membrane  was also 32 P labeled (lower right).   Chapter 9 Lipids and Biological Membranes synthesize phospholipids combined with the countervailing tendencies of ATP-   dependent phospholipid translocases that generate asymmetric phospholipid  distributions and flipases that equilibrate these distributions. The importance  of these lipid transport systems is demonstrated by the observation that the  presence of phosphatidylserine on the exteriors of many cells induces blood  clotting (i.e., it is an indication of tissue damage) and, in erythrocytes, marks  the cell for removal from the circulation.   In all cells, new membranes are generated by the expansion of existing mem¬  branes. In eukaryotic cells, lipids synthesized on the cytoplasmic face of the  ER are transported to other parts of the cell by membranous vesicles that bud  off from the ER and fuse with other cellular membranes (Section 9-4E). These  vesicles also carry membrane proteins.   Lipid Rafts Are Membrane Subdomains. Lipids and proteins in membranes  can also be laterally organized. Thus the plasma membranes of many eukary¬  otic cells have two or more distinct domains that have different functions. For  example, the plasma membranes of epithelial cells (the cells lining body cav¬  ities and free surfaces) have an apical domain, which faces the lumen (inte¬  rior) of the cavity and often has a specialized function (such as the absorption  of nutrients in intestinal brush border cells), and a basolateral domain, which  covers the remainder of the cell. These two domains, which do not intermix,  have different compositions of both lipids and proteins.   In addition, the hundreds of different lipids and proteins within a given  plasma membrane domain may not be uniformly mixed but instead often seg¬  regate to form microdomains that are enriched in certain lipids and proteins.  This may result from specific interactions between integral membrane proteins  and particular types of membrane lipids. Divalent metal ions, notably Ca 2+ ,  which bind to negatively charged lipid head groups such as those of phos¬  phatidylserine, may also cause clustering of these lipids.   One type of microdomain, termed a lipid raft, appears to consist of closely  packed glycosphingolipids (which occur only in the outer leaflet of the plasma  membrane) and cholesterol. By themselves, glycosphingolipids cannot form bilay¬  ers because their large head groups prevent the requisite close packing of their pre¬  dominantly saturated hydrophobic tails. Conversely, cholesterol by itself does not  form a bilayer due to its small head group. It is therefore likely that the glyco¬  sphingolipids in lipid rafts associate laterally via weak interactions between their car¬  bohydrate head groups , and the voids between their tails are filled in by cholesterol.   Owing to the close packing of their component lipids and the long, saturated  sphingolipid tails, sphingolipid—cholesterol rafts have a more ordered or crystalline  arrangement than other regions of the membrane and are more resistant to solu¬  bilization by detergents. The rafts may diffuse laterally within the membrane.  Certain proteins preferentially associate with the rafts, including many GPI-linked  proteins and some of the proteins that participate in transmembrane signaling  processes (Chapter 13). This suggests that lipid rafts, which are probably present  in all cell types, function as platforms for the assembly of complex intercellular  signaling systems. Several viruses, including influenza virus, measles virus, Ebola  virus, and HIV, localize to lipid rafts, which therefore appear to be the sites from  which these viruses enter uninfected cells and bud from infected cells. It should  be noted that lipid rafts are highly dynamic structures that rapidly exchange both  proteins and lipids with their surrounding membrane as a consequence of the weak  and transient interactions between membrane components.   In contrast to membrane lipids, membrane proteins do not change their  orientation in the membrane after they are synthesized. Membrane proteins,  like all proteins, are ribosomally synthesized under the direction of messen-   tide grows from its N-terminus to its C-terminus by the stepwise addition  of amino acid residues. Ribosomes may be free in the cytosol or bound to  the ER to form the rough endoplasmic reticulum (RER, so called because  of the knobby appearance its bound ribosomes give it; Fig. 9-33). Free ri¬  bosomes synthesize mostly soluble and mitochondrial proteins, whereas mem¬  brane-bound ribosomes manufacture transmembrane proteins and proteins  destined for secretion , operation within the ER, and incorporation into lyso-  somes (membranous vesicles containing a battery of hydrolytic enzymes that  degrade and recycle cell components). The latter proteins initially appear in  the ER.   How are RER-destined proteins differentiated from other proteins? And how  do these large, relatively polar molecules pass through the RER membrane?   These processes occur via the secretory pathway, which was first described by  Gunter Blobel, Cesar Milstein, and David Sabatini around 1975. Since —25%  of the various proteins synthesized by all types of cells are integral proteins  and many others are secreted, — 40% of the various types of proteins that a cell  synthesizes must be processed via the secretory pathway or some other protein tar¬  geting pathway. Here we outline the secretory pathway, which is diagrammed  in Fig. 9-36:   1. All secreted, ER-resident, and lysosomal proteins, as well as many TM pro¬  teins, are synthesized with leading (N-terminal) 13 - to 36-residue signal  peptides . These signal peptides consist of a 6- to 15-residue hydropho¬  bic core flanked by several relatively hydrophilic residues that usually    Growing protein  folds, is modified  by ER enzymes,  and is anchored  in membrane.    FIG. 9-36 The secretory pathway. This composite figure depicts of an integral protein. Details are given in the text. O See the  the ribosomal synthesis, membrane insertion, and initial glycosylation Animated Figures.    FIG. 9-37 The N-terminal sequences of some eukaryotic preproteins. The hy¬  drophobic cores (brown) of most signal peptides are preceded by basic residues (blue).  [After Watson, M.E.E., Nucleic Acids Res. 12, 5147-5156 (1984).]   include one or more basic residues near the N-terminus (Fig. 9-37).  Signal peptides otherwise have little sequence similarity. However, a va¬  riety of evidence indicates they form a helices in nonpolar environments.   2. When the signal peptide first protrudes beyond the ribosomal surface  (when the polypeptide is at least ~40 residues long), the signal recog¬  nition particle (SRP), a 325-kD complex of six different polypeptides  and a 300-nucleotide RNA molecule, binds to both the signal peptide  and the ribosome (Section 27-5B). At the same time, the SRPs bound  guanosine diphosphate (GDP; the guanine analog of ADP) is re¬  placed by guanosine triphosphate (GTP; the guanine analog of ATP).  The resulting conformational change in the SRP causes the ribosome  to arrest further polypeptide growth, thereby preventing the RER-  destined protein from being released into the cytosol.   3. The SRP—ribosome complex diffuses to the RER surface, where it  binds to the SRP receptor (SR; also called docking protein) in com¬  plex with the translocon, a protein pore in the ER membrane through  which the growing polypeptide will be extruded. In forming the  SR—translocon complex, the SRs bound GDP is replaced by GTP.   4. The SRP and SR stimulate each other to hydrolyze their bound GTP  to GDP (which is energetically equivalent to ATP hydrolysis), result¬  ing in conformational changes that cause them to dissociate from each  other and from the ribosome—translocon complex. This permits the  bound ribosome to resume polypeptide synthesis such that the grow¬  ing polypeptides N-terminus passes through the translocon into the lu¬  men of the ER. Most ribosomal processes, as we will see in Section 27-4,  are driven by GTP hydrolysis.   5. Shortly after the signal peptide enters the ER lumen, it is specifically  cleaved from the growing polypeptide by a membrane-bound signal  peptidase (polypeptide chains with their signal peptide still attached  are known as preproteins; signal peptides are alternatively called pre¬  sequences) .   6. The nascent (growing) polypeptide starts to fold to its native confor¬  mation, a process that is facilitated by its interaction with the ER-  resident chaperone protein Hsp70 (Section 6-5B). Enzymes in the ER  lumen then initiate posttranslational modification of the polypep¬  tide, such as the specific attachments of “core” carbohydrates to form  glycoproteins (Section 8-3C) and the formation of disulfide bonds as  facilitated by protein disulfide isomerase (Section 6-5A). Once the protein   has folded, it cannot be pulled back through the membrane. Secretory, Section 4 Membrane Structure and Assembly   membrane into the lumen. TM proteins, in contrast, contain one or   7. When the synthesis of the polypeptide is completed, it is released from  both the ribosome and the translocon. The ribosome detaches from  the RER, and its two subunits dissociate.   The secretory pathway also functions in prokaryotes for the insertion of cer¬  tain proteins into the cell membrane (whose exterior is equivalent to the ER  lumen). Indeed, all forms of life yet tested have homologous SRPs, SRs, and  translocons. Nevertheless, it should be noted that cells have several mecha¬  nisms for installing proteins in or transporting them through membranes. For  example, cytoplasmically synthesized proteins that reside in the mitochondrion  reach their destinations via mechanisms that are substantially different from  that of the secretory pathway.   The Translocon Is a Multifunctional Transmembrane Pore. In 1975, Blobel  postulated that protein transport through the RER membrane is mediated by  an aqueous TM channel. However, it was not until 1991 that he was able to  experimentally demonstrate its existence. These channels, now called translo¬  cons, enclose aqueous pores that completely span the ER membrane, as shown  by linking nascent polypeptide chains to fluorescent dyes whose fluorescence  is sensitive to the polarity of their environment. The channel-forming compo¬  nent of the translocon is a heterotrimeric protein named Sec61 in mammals  and SecY in prokaryotes. This protein is conserved throughout all kingdoms of  life and hence is likely to have a similar structure and function in all organisms.   The X-ray structure of the SecY complex from the archaeon Methanococcus  jannaschii reveals that the a, (3, and y subunits, respectively, have 10, 1, and  1 TM a helices (Fig. 9-38). The a subunits TM helices are wrapped around  an hourglass-shaped channel whose minimum diameter is ~3 A. The channel    FIG. 9-38 Structure and function of the M. jannaschii SecY  complex, (a) X-Ray structure of the complex, with shading indicating  the positions of membrane phospholipid head groups ( violet) and  hydrocarbon tails (pink). The a subunit of SecY is colored in rainbow  order from its N-terminus (dark blue ) to its C-terminus (red), the p  subunit is red, and the y subunit is purple, (b) View of SecY from the  cytosol. The translocon’s putative lateral gate is on the left between  helices 2 and 7. (c) Model for the insertion of a TM helix into a    membrane. The translocon (blue) is viewed as in Part b. A polypeptide  chain (yellow) is shown bound in the translocon’s pore during its  translocation through the membrane, and a signal-anchor sequence  (red) is shown passing through the translocon’s lateral gate and being  released into the membrane (arrow). [Parts a and b courtesy of  Stephen Harrison and Tom Rapoport, Harvard Medical School. PDBid  1RH5. Part c after a drawing by Dobberstein, B. and Sinning, L,  Science 303, 320 (2004).]   FIG. 9-39 Electron micrograph of the  Golgi apparatus. In this array of flattened  vesicles, newly synthesized transmembrane and  secretory proteins undergo processing and  sorting. [© Biophoto Associates/Photo  Researchers, Inc.]    is blocked at its extracellular end by a short relatively hydrophilic helix (blue  unnumbered helix in Figs. 9-38/2 and b). It is proposed that an incoming sig¬  nal peptide pushes this helix aside and hence that the helix functions as a plug  to prevent small molecules from leaking across the membrane in the absence  of a translocating polypeptide. The maximum diameter of an extended  polypeptide is ~12 A and that of an a helix is ~14 A. In order for SecY to  function as a channel for translocating polypeptides, the central pore must ex¬  pand, a structural change that would require relatively simple hingelike mo¬  tions involving conserved Gly residues.   In addition to forming a conduit for soluble proteins to pass through the  membrane, the translocon must mediate the insertion of an integral proteins TM  segments into the membrane. The X-ray structure of SecY suggests that this oc¬  curs by the opening of the C-shaped a subunit, as is diagramed in Fig. 9-38 c,  to permit the lateral installation of the TM segment into the membrane.   The signal peptides of many TM proteins are not cleaved by signal peptidase  but , instead^ are inserted into the membrane. Such so-called signal-anchor  sequences may be oriented with either their N- or C-termini in the cytosol. If the  N-terminus is installed in the cytosol, then the polypeptide must have looped  around before being inserted into the membrane. Moreover, for polytopic  (multispanning) TM proteins such as the SecY a subunit itself, this must oc¬  cur for each successive TM helix. Since it seems unlikely that the SecY chan¬  nel could expand to simultaneously accommodate numerous TM helices, these  helices are probably installed in the membrane one or two at a time. The  mechanisms by which the translocon recognizes TM segments are not well un¬  derstood. For example, the experimental deletion or insertion of a TM helix  in a polytopic protein does not necessarily change the membrane orientations  of the succeeding TM helices; when two successive TM helices have the same  preferred orientation, one of them may be forced out of the membrane.   Shortly after their synthesis in the rough ER, partially processed transmembrane,  secretory, and lysosomal proteins appear in the Golgi apparatus (Fig. 9-39). This  0.5- to 1.0-|Jim-diameter organelle consists of a stack of three to six or more  (depending on the species) flattened and functionally distinct membranous sacs  known as cisternae, where further posttranslational processing, mainly glyco-  sylation, occurs (Section 8-3C). The Golgi stack (Fig. 9-40) has two distinct  faces, each composed of a network of interconnected membranous tubules:  the cis Golgi network, which is opposite the ER and is the port through  which proteins enter the Golgi apparatus; and the trans Golgi network,  through which processed proteins exit to their final destinations. The inter¬  vening Golgi stack contains at least three different types of sacs, the cis, medial,  and trans cisternae, each of which contains different sets of glycoprotein¬  processing enzymes.   Proteins transit from one end of the Golgi stack to the other while being  modified in a stepwise manner. The proteins are transported via two mechanisms:   1. They are conveyed between successive Golgi compartments in the cis  to trans direction as cargo within membranous vesicles that bud off of  one compartment and fuse with a successive compartment, a process  known as forward or anterograde transport.   2. They are carried as passengers in Golgi compartments that transit the  Golgi stack; that is, the cis cisternae eventually become trans cisternae,  a process called cisternal progression or maturation. (Golgi-resident  proteins may migrate backward by retrograde transport from one  compartment to the preceding one via membranous vesicles.)   The cisternal progression mechanism has been clearly shown to occur, but the  significance of the anterograde transport mechanism is as yet unclear. In any   FIG. 9-40 The posttranslational  processing of proteins. Membrane, secretory,  and lysosomal proteins are synthesized by  RER-associated ribosomes (gray dots ; top). As  they are synthesized, the proteins (red dots ) are  either injected into the lumen of the ER or  inserted into its membrane. After initial  processing in the ER, the proteins are  encapsulated in vesicles that bud off from the  ER membrane and subsequently fuse with the  cis Golgi network (top). The proteins are  progressively processed in the cis, medial, and  trans cisternae of the Golgi. Finally, in the trans  Golgi network (bottom), the completed  glycoproteins are sorted for delivery to their final  destinations: the plasma membrane, secretory  vesicles, or lysosomes, to which they are  transported by yet other vesicles.    List, in order, all the cellular compart¬  ments involved in synthesizing a  secreted protein.    case, on reaching the trans Golgi network, the now mature proteins are sorted  and sent to their final cellular destinations.   Membrane, Secretory, and Lysosomal Proteins Are Transported in Coated  Vesicles. The vehicles in which proteins are transported between the RER,  the different compartments of the Golgi apparatus, and their final destinations  are known as coated vesicles (Fig. 9-41). This is because these 60- to 150-  nm-diameter membranous sacs are initially encased on their outer (cytosolic) faces  by specific proteins that act as flexible scaffolding in promoting vesicle forma¬  tion. A vesicle buds off from its membrane of origin and later fuses to its tar¬  get membrane. This process preserves the orientation of the transmembrane protein   FIG. 9-41 Electron micrographs of coated vesicles. vesicles, (c) COPII-coated vesicles. The inserts in Parts b and c show   (a) Clathrin-coated vesicles. Note their polyhedral character. [Courtesy the respective vesicles at higher magnification. [Courtesy of Lelio Orci,  of Barbara Pearse, Medical Research Council, U.K.] (b) COPI-coated University of Geneva, Switzerland.]    FIG. 9-42 Fusion of a vesicle with the  plasma membrane. The inside of the vesicle  and the exterior of the cell are topologically  equivalent Fusion of the vesicle with the plasma  membrane preserves the orientation of the  integral proteins embedded in the vesicle bilayer  because the same side of the protein is always  immersed in the cytosol. Note that soluble  proteins packaged inside a secretory vesicle that  fuses with the plasma membrane would be  released outside the cell.   (Fig. 9-42), because the lumens of the ER and the Golgi cisternae are topologi¬  cally equivalent to the outside of the cell. This explains why the carbohydrate  moieties of integral glycoproteins and the GPI anchors of GPI-linked proteins  occur only on the external surfaces of plasma membranes.   The three known types of coated vesicles are characterized by their pro¬  tein coats:   1. Clathrin (Fig. 9-41 a), a protein that forms a polyhedral framework  around vesicles that transport TM, GPI-linked, and secreted proteins  from the Golgi to the plasma membrane (see below).   2. COPI protein (Fig. 9-41 b; COP for coat protein), which forms what  appears to be a fuzzy rather than a polyhedral coating about vesicles  that carry out both the anterograde and retrograde transport of pro¬  teins between successive Golgi compartments. In addition, COPI-  coated vesicles return escaped ER-resident proteins from the Golgi back  to the ER (see below). The COPI protomer, which contains seven  different subunits, is named coatomer.   3. COPII protein (Fig. 9-41 c), which transports proteins from the ER to  the Golgi. The COPII vesicle components are then returned to the ER  by COPI-coated vesicles (the COPI vesicle components entering the  ER are presumably recycled by COPII-coated vesicles). The COPII  coat consists of two conserved protein heterodimers.   All of the above coated vesicles also bear receptors that bind the proteins be¬  ing transported, as well as proteins that mediate the fusion of these vesicles  with their target membranes (Section 9-4F).    Clathrin Forms Flexible Cages. Clathrin-coated vesicles are structurally bet¬  ter characterized than those coated with COPI or COPII. The clathrin net¬  work is built from proteins known as triskelions (Fig. 9-43), which consist  of three heavy chains (190 kD) that each bind one of two homologous light  chains (24—27 kD). The triskelions assemble to form polyhedral cages in which  each vertex is the center (hub) of a triskelion and the ~150-A-long edges are  formed by the overlapping legs of four triskelions (Fig. 9-44). The clathrin  light chains are not required for clathrin cage assembly. In fact, they inhibit  heavy chain polymerization in vitro , suggesting that they play a regulatory role  in clathrin cage formation in the cytosol.   A clathrin polyhedron, which has 12 pentagonal faces and a variable num¬  ber of hexagonal faces, is the most parsimonious way of enclosing a spheroidal  object in a polyhedral cage. The volume enclosed by a clathrin polyhedron in¬  creases with the number of hexagonal faces.   The triskelion legs, which have a total length of ~450 A, exhibit consid¬  erable flexibility (Fig. 9-43). This is a functional necessity for the formation  of different-sized vesicles as well as for the budding of a vesicle from a mem¬  brane surface, which requires a large change in its curvature. The clathrin  heavy chains appear to flex mainly along a portion of the knee (Fig. 9-44 A)  between the proximal and distal segments that is free of contacts with other  molecules in clathrin cages.   FIG. 9-43 An electron micrograph of  triskelions. The variable orientations of their  legs are indicative of their flexibility. [Courtesy of  Daniel Branton, Harvard University.]   FIG. 9-44 Anatomy of clathrin-coated vesicles, (a) A cryo-  EM-based image of a light chain-free clathrin cage from bovine brain  at 7.9 A resolution. The particle shown, a so-called hexagonal barrel,  which has D 6 symmetry (the symmetry of a hexagonal prism), consists  of 36 triskelions. Three of its interdigitated but symmetry unrelated  triskelions are drawn in red, yellow, and green, (b) A cryo-EM-based  image of a triskelion labeled with the names of its various segments.  The N-terminus of each heavy chain occupies the terminal domain  and its C-terminus is located in the vertex joining the three heavy  chains to form the triskelion. (c) Diagrams of the three polyhedral  structures that are formed when triskelions assemble into clathrin    cages in vitro. The minicoat has tetrahedral (7) symmetry, the  hexagonal barrel has D 6 symmetry, and the soccer ball has  icosahedral  symmetry (symmetry is discussed in Section 6-3).  These polyhedra consist of 28, 36, and 60 triskelions, respectively.  The arrangement of one triskelion within the “hexagonal barrel” is  indicated in blue. In vivo, clathrin forms membrane-enclosing  polyhedral cages with a large range of different sizes (number of  hexagons). The hexagonal barrel seen in Part a is only —700 A in  diameter, whereas clathrin-coated membranous vesicles are typically  — 1200 A in diameter or larger. [Courtesy of Stephen Harrison, Tomas  Kirchhausen, and Thomas Walz, Harvard Medical School.]   Chapter 9 Lipids and Biological Membranes In addition to transporting membrane and secretory proteins between the   Golgi apparatus and the plasma membrane, clathrin-coated vesicles partici¬  pate in endocytosis. In this process (discussed in Section 20-IB), a portion  of the plasma membrane invaginates to form a clathrin-coated vesicle that  engulfs specific proteins from the extracellular medium and then transports  them to intracellular destinations.   T Proteins Are Directed to the Lysosome by Carbohydrate Recognition  Markers. The signals that direct particular proteins to the various types of  coated vesicles for transport between cellular compartments are not entirely un¬  derstood. However, the trafficking of lysosomal proteins is known to depend  on their oligosaccharides. A clue to the nature of this process was provided by  the human hereditary defect known as I-cell disease (alternatively, mucolipi¬  dosis II) which, in homozygotes, is characterized by severe progressive psy¬  chomotor retardation, skeletal deformities, and death by age 10. The lysosomes  in the connective tissue of I-cell disease victims contain large inclusions (after  which the disease is named) of glycosaminoglycans and glycolipids as a result  of the absence of several lysosomal hydrolases. These enzymes are synthesized  on the RER with their correct amino acid sequences, but rather than being  dispatched to the lysosomes are secreted into the extracellular medium. This  misdirection results from the absence of a mannose-6-phosphate recognition  marker on the carbohydrate moieties of these hydrolases because of a defi¬  ciency of an enzyme required for mannose phosphorylation of the lysosomal  proteins. The mannose-6-phosphate residues are normally bound by a recep¬  tor in the coated vesicles that transports lysosomal hydrolases from the Golgi  apparatus to the lysosomes. No doubt, other glycoproteins are directed to their  intracellular destinations by similar carbohydrate markers.   ER-Resident Proteins Have the C-Terminal Sequence KDEL. Most soluble  ER-resident proteins in mammals have the C-terminal sequence KDEL  (HDEL in yeast), KKXX, or KXKXXX (where X represents any amino acid  residue), whose alteration results in the secretion of the resulting protein. By  what means are these proteins selectively retained in the ER? Since many ER-  resident proteins freely diffuse within the ER, it seems unlikely that they are  immobilized by membrane-bound receptors within the ER. Rather, ER-resident  proteins, like secretory and lysosomal proteins, readily leave the ER via COPII-  coated vesicles, but ER-resident proteins are promptly retrieved from the Golgi  and returned to the ER in COPI-coated vesicles. Indeed, coatomer binds the  Lys residues in the C-terminal KKXX motif of transmembrane proteins, which  presumably permits it to gather these proteins into COPI-coated vesicles.  Furthermore, genetically appending KDEL to the lysosomal protease  cathepsin D causes it to accumulate in the ER, but it nevertheless acquires an  7V-acetylglucosaminyl-1 -phosphate group, a modification that is made in an  early Golgi compartment. Presumably, a membrane-bound receptor in a post-  ER compartment binds the KDEL signal and the resulting complex is returned  to the ER in a COPI-coated vesicle. KDEL receptors have, in fact, been iden¬  tified in yeast and humans. However, the observation that former KDEL pro¬  teins whose KDEL sequences have been deleted are, nevertheless, secreted  relatively slowly suggests that there are mechanisms for retaining these pro¬  teins in the ER by actively withholding them from the bulk flow of proteins  through the secretory pathway.   In all cells, new membranes are generated by the expansion of existing membranes.  In eukaryotes, this process occurs mainly via vesicle trafficking in which a vesi¬  cle buds off from one membrane (e.g., that of the Golgi apparatus) and fuses  to a different membrane (e.g., the plasma membrane or that of the lysosome),    FIG. 9-45 Vesicle fusion at a synapse, (a) An electron micrograph of a frog neuro¬  muscular synapse. Synaptic vesicles are undergoing fusion ( arrows ) with the presynaptic  plasma membrane (top). [Courtesy of John Heuser, Washington University School of  Medicine, St. Louis, Missouri.] (b) This process discharges the neurotransmitter contents of  the synaptic vesicles into the synaptic cleft, the space between the neuron and the muscle  cell (the postsynaptic cell).    thereby transferring both lipids and proteins from the parent to the target  membrane.   On arriving at its target membrane, a vesicle fuses with it, thereby releas¬  ing its contents on the opposite side of the target membrane (Fig. 9-42). We  have already seen how proteins are transported from the ER through the Golgi  apparatus and secreted by this mechanism. Other substances are also secreted  in this way, such as neurotransmitters, the small molecules released by neu¬  rons (nerve cells; Fig. 9 -45). When a nerve impulse in a presynaptic cell  reaches a synapse (the junction between neurons or between neurons and  muscles), it triggers the fusion of neurotransmitter-containing synaptic vesi¬  cles with the presynaptic membrane (a specialized section of the neurons  plasma membrane). This releases the neurotransmitter into the ~200-A-wide  synaptic cleft, a process called exocytosis. In less than 0.1 ms, the neuro¬  transmitter diffuses across the synaptic cleft to the postsynaptic membrane,  where it binds to specific receptors that then trigger the continuation of the  nerve impulse or muscle contraction in the postsynaptic cell.   Biological membranes do not spontaneously fuse. Indeed, being negatively  charged, they strongly repel each other at short distances. This repulsive force  must be overcome if biological membranes are to fuse. How do vesicles fuse  and why do they fuse only with their target membranes?   Extensive investigations, in large part by James Rothman, have identified  numerous proteins that mediate vesicle fusion with their target membranes.  Among these are integral or lipid-linked proteins known as SNAREs.  R-SNAREs (which contain conserved Arg residues) usually associate with vesicle  membranes, and Q-SNAREs (which contain conserved Gin residues) usually as¬  sociate with target membranes. Interactions among these proteins, as we will see,  firmly anchor the vesicle to the target membrane, a process called docking.   The X-ray structure of a core SNARE complex is shown in Fig. 9-46.  Four parallel ~65-residue a helices wrap around each other with a gentle left-  handed twist. For the most part, the sequence of each helix has the expected  seven-residue repeat, (a-b-c-d-e-fg) „, with residues a and d hydrophobic  (Section 6-1C). However, the central layer of side chains along the length of  the four-helix bundle includes an Arg residue from the R-SNARE that is hy¬  drogen bonded to three Gin side chains, one from each of the Q-SNARE he¬  lices. These highly conserved polar residues are sealed off from the aqueous  environment so that their interactions serve to bring the four helices into    FIG. 9-46 X-Ray structure of a SNARE complex modeled between two mem¬  branes. The complex includes an R-SNARE (synaptobrevin, blue ) and two Q-SNARES  (syntaxin, red, and SNAP-25, green, a two-Q-domain-containing Q-SNARE) for a total of  four helices, which are shown here as ribbons. The transmembrane C-terminal extensions  of syntaxin and synaptobrevin are modeled as helices (yellow-green). The peptide segment  connecting the two Q-domain helices of SNAP-25 is speculatively represented as an un¬  structured loop (brown). The loop is anchored to the membrane via Cys-linked palmitoyl  groups (not shown). The cleavage sites for various clostridial neurotoxins (causing tetanus  and botulism; Box 9-3) are indicated by the arrows. [Courtesy of Axel Brunger, Stanford  University.]    The frequently fatal infectious diseases tetanus (which arises  from wound contamination) and botulism (a type of food poison¬  ing) are caused by certain anaerobic bacteria of the genus  Clostridium. These bacteria produce extremely potent protein neu¬  rotoxins that inhibit the release of neurotransmitters into synapses.  In fact, botulinum toxins are the most powerful known toxins; they  are —10 million times more toxic than cyanide.   There are seven serologically distinct types of botulinum neuro¬  toxins, designated BoNT/A through BoNT/G, and one type of  tetanus neurotoxin, TeTx. Each of these homologous proteins is  synthesized as a single — 150-kD polypeptide chain that is  cleaved by host proteases to yield an —50-kD light chain and an  — 100-kD heavy chain. The heavy chains bind to specific types of  neurons and facilitate the uptake of the light chain by endocytosis.  Each light chain is a protease that cleaves its target SNARE at a    specific site (see Fig. 9-46). SNARE cleavage prevents the forma¬  tion of the core complex and thereby halts the exocytosis of  synaptic vesicles. The heavy chain of TeTx specifically binds to  inhibitory neurons (which function to moderate excitatory nerve  impulses) and is thereby responsible for the spastic paralysis  characteristic of tetanus. The heavy chains of the BoNTs instead  bind to motor neurons (which innervate muscles) and thus cause  the flaccid paralysis characteristic of botulism.   The administration of carefully controlled quantities of botu¬  linum toxin (“Botox”) is medically useful in relieving the symptoms  of certain types of chronic muscle spasms. Moreover, this toxin is  being used cosmetically: Its injection into the skin relaxes the  small muscles causing wrinkles and hence these wrinkles  disappear for —3 months.    proper register. Since cells contain numerous different R-SNAREs and  Q-SNAREs, it seems likely that their interactions are at least partially respon¬  sible for the specificity that vesicles exhibit in fusing with their target mem¬  branes. Bacterial proteases that cleave SNAREs interfere with vesicle fusion,  with serious consequences (Box 9-3).   The association of an R-SNARE on a vesicle with Q-SNAREs on its target  membrane brings the two bilayers into close proximity. But what induces the fu¬  sion of the juxtaposed lipid bilayers? The answer, as is diagrammed in Fig. 9-47,  is that the mechanical forces arising from the formation of a ring of several  (estimated to be 5-10) SNARE complexes pulls together the apposing bilayers.  This expels the contacting lipids between them so as to join their outer leaflets,  a process known as hemifusion. Indeed, the pressure (force/area) within the  ring of trans-SNARE complexes is estimated to be 100 to 1000 atm. In the  resulting transient structure, no aqueous contact between the two membrane  systems has yet been established. However, as the fusion process proceeds (the  trans-SNARE complexes continue zipping up), the two inner leaflets of the  now partially joined membranes come together to form a new bilayer, whose  component lipids are subsequently similarly expelled to yield a fusion pore.  The fusion pore then rapidly expands, thereby fully joining the two mem¬  branes as well as their contents. Thus, vesicle fusion is driven by the protein  folding forming the trans-SNARE complexes. However, in vitro , this process  takes 30 to 40 minutes, whereas the fusion of a synaptic vesicle with the presy-  naptic membrane requires ~0.3 ms in vivo. This argues that other proteins  also participate in inducing bilayer fusion and, indeed, several other proteins  have been implicated.   Some Viruses Use Fusion Proteins to Infect Their Target Cells. Many  viruses, including those causing influenza and AIDS, arise by budding from  the plasma membrane of a virus-infected cell, much like the budding of a vesi¬  cle from a membrane. In order to infect a new cell, the membrane of the re¬  sulting membrane-enveloped virus must fuse with a membrane in its target  cell so as to deposit the virus’ cargo of nucleic acid in the cells cytoplasm.  Such membrane fusion events are mediated by protein systems that differ from  those responsible for the vesicle fusion events we discussed above. As an ex¬  ample, let us discuss how influenza virus mediates membrane fusion (other   Zipping: As the vesicle  approaches its target  membrane, the SNAREs begin  zipping together (docking)  from their N-termini, which    Hemifusion: As docking  proceeds, the increased  curvature and lateral tension  induce the approaching  bilayer leaflets to fuse,    The two bilayers leaflets that were  originally farthest apart are brought  together to form a new bi layer.    Fusion pore formation: The continuing  SNARE-induced lateral tension causes  membrane breakdown, resulting in the  formation of a fusion pore.    FIG. 9-47 A model for SNARE-mediated vesicle fusion. Here and red worms. [After a drawing by Chen, Y.A. and Scheller, R.H.,   R-SNAREs and Q-SNAREs are schematically represented by blue Nature Rev. Mol. Cell Biol. 2, 98 (2001).]    membrane-enveloped viruses use similar systems). Virus-mediated membrane  fusion occurs in three stages:   3. Fusion of the viral membrane with a host cell membrane so as to re¬  lease the viral genome into the host cell cytoplasm.   The major integral protein component of the membrane that envelops in¬  fluenza virus is named hemagglutinin (HA) because it causes erythrocytes to  agglutinate (clump together). HA, a so-called viral fusion protein, mediates  influenza host cell recognition by binding to specific glycoproteins that act as  cell-surface receptors. These glycoproteins (like glycophorin A in erythrocytes;  Section 9-3A) bear terminal 7V-acetylneuraminic acid (sialic acid; Fig. 8-6)  residues on their carbohydrate groups. HA is synthesized as a homotrimer of  550-residue subunits that are each anchored to the membrane by a single  transmembrane helix (residues 524-540). However, HAs Arg 329 is posttrans-  lationally excised by host cell proteases to yield two peptides, designated HA1  and HA2, that remain linked by a disulfide bond.   After budding from the original host cell, an influenza virus particle can in¬  fect a new host cell. The virus binds to the plasma membrane of the new cell  and is taken into the cell by the invagination of the membrane via a process  known as receptor-mediated endocytosis (Section 20- IB) that superficially    FIG. 9-48 X-Ray structure of influenza  hemagglutinin, (a) Ribbon diagram of the BHA  monomer. HA1 is green and HA2 is cyan.   (b) Ribbon diagram of the BHA trimer. Each  HA1 and HA2 chain is drawn in a different  color. The orientations of the green HA1 and the  cyan HA2 are the same as in Part a. [Based on  an X-ray structure by John Skehel, National  Institute for Medical Research, London, U.K., and  Don Wiley, Harvard University. PD Bid 4HMG.]   • Describe the structure of a lipid raft. Why  might a raft be associated with a particu¬  lar cellular activity?   • Summarize the steps of the secretory  pathway, including the function of the  translocon.   • Describe how a membrane protein, a  secreted protein, and a lysosomal protein  are transported from the RER to their final  destinations.   • Why does it make sense that carbohy¬  drate groups serve as intracellular ad¬  dressing signals?   • Describe how SNAREs and viral fusion  proteins such as hemagglutinin act to  bring specific membranes close together  and trigger their fusion.   resembles the reverse of the fusion of a vesicle with a membrane (Fig. 9-42).  The resulting intracellular vesicle with the virus bound to its inner surface then  fuses with a vesicle known as an endosome that has an internal pH of ~5.  The consequent drop in pH triggers a conformational change in which the  N-terminal end of HA2, a conserved, hydrophobic, ~24-residue segment  known as a fusion peptide, inserts into the host cells endosomal membrane,  thereby tightly linking it to the viral membrane preparatory to their fusion.   Our understanding of how HA mediates the fusion of viral and host cell  membranes is based largely on X-ray structural studies by Don Wiley and John  Skehel. HAs hydrophobic membrane anchor interferes with its crystallization.  However, the proteolytic removal of HAs transmembrane helix yields a crystal-  lizable protein named BHA, whose X-ray structure includes a globular region that  is perched on a long fibrous stalk projecting from the viral membrane surface  (Fig. 9-48/*). The globular region contains the sialic acid-binding pocket,  whereas the fibrous stalk includes a remarkable 76-A-long a helix (53 residues  in 14 turns). The dominant interaction stabilizing BHAs trimeric structure is a  triple-stranded coiled coil consisting of the long a helices from each of its pro-  tomers (Fig. 9-48 £). Curiously, BHAs fusion peptides are buried in its hydropho¬  bic interior, -TOO A from the sialic acid-binding sites at the “top” of the protein.   In the endosome, HA undergoes a dramatic conformational change  (Fig. 9-49), the nature of which was elucidated by the X-ray structure of  a portion of BHA named TBHA2 that consists of BHAs long helix and  some of its flanking regions. In this conformational change, segments A and  B at the N-terminus of TBHA2 (the red and orange segments in Fig. 9-49) un¬  dergo a jackknife-like movement of —100 A in a way that extends the top  of the long helix by -TO helical turns toward the endosomal membrane. This  translocates the fusion peptide (which is absent in TBHA2 but would extend  beyond its N-terminus at the top of segment A) by at least —100 A above its  position in BHA, thereby allowing it to insert into the host membrane. At the  same time, the long helix is shortened from the bottom by similar shifts of  segments D and E (green and blue in Fig. 9-49). These conformational  changes, which simultaneously occur in several closely spaced HA molecules in  the viral membrane, draw together the viral and host membranes so as to facil¬  itate their fusion in a manner similar to that postulated for SNARE complexes.   FIG. 9-49 Schematic drawing comparing  the structures of BHA and TBHA2. This  drawing indicates the positions and heights  above the viral membrane surface of TBHA2’s  various structural elements in the HA trimer (left)  and in its low-pH form (right). In the low-pH  form, the fusion peptide would protrude well  above the receptor-binding heads where it would  presumably insert itself into the endosomal  membrane.   • Lipids are a diverse group of molecules that are soluble in organic  solvents and, in contrast to other major types of biomolecules,  do not form polymers.   • Fatty acids are carboxylic acids whose chain lengths and degrees  of unsaturation vary.   • Adipocytes and other cells contain stores of triacylglycerols,  which consist of three fatty acids esterified to glycerol.   • Glycerophospholipids are amphiphilic molecules that contain  two fatty acid chains and a polar head group.   • Cholesterol, steroid hormones, and vitamin D are all based on a  four-ring structure.   • The arachidonic acid derivatives prostaglandins, prostacyclins,  thromboxanes, leukotrienes, and lipoxins are signaling molecules  that have diverse physiological roles.   • Glycerophospholipids and sphingolipids form bilayers in which  their nonpolar tails associate with each other and their polar head  groups are exposed to the aqueous solvent.   • Although the transverse diffusion of a lipid across a bilayer is  extremely slow, lipids rapidly diffuse in the plane of the bilayer.  Bilayer fluidity varies with temperature and with the chain lengths  and degree of saturation of its component fatty acid residues.   • The proteins of biological membranes include integral (intrinsic)  proteins that contain one or more transmembrane a helices or a  |3 barrel. In all cases, the membrane-exposed surface of the pro¬  tein is hydrophobic.    • Other membrane-associated proteins may be anchored to the  membrane via isoprenoid, fatty acid, or glycosylphosphatidylinositol  (GPI) groups. Peripheral (extrinsic) proteins are loosely associated  with the membrane surface.   • The fluid mosaic model of membrane structure accounts for the  lateral diffusion of membrane proteins and lipids.   • The arrangement of membrane proteins in the lipid bilayer may de¬  pend on their interactions with an underlying protein skeleton, as in  the erythrocyte.   • Membrane proteins and lipids are distributed asymmetrically in  the two leaflets of the bilayer and may form domains such as lipid  rafts.   • Transmembrane (TM), secretory, and lysosomal proteins are syn¬  thesized by means of the secretory pathway. A signal peptide di¬  rects a growing polypeptide chain through the RER membrane  via a protein pore called the translocon, which also functions to  laterally install TM proteins into the RER membrane.   • Coated vesicles transport membrane-embedded and luminal pro¬  teins from the ER to the Golgi apparatus for further processing,  and from there to other membranes. The proteins coating these  vesicles may consist largely of clathrin, which forms polyhedral  cages, or COPI or COPII, which form coats with an amorphous  appearance.   • The fusion of vesicles with membranes occurs via a complex  process that involves SNAREs. These form four-helix bundles  that bring two membranes into proximity, which in turn induces  membrane fusion.   • Viral fusion proteins mediate the fusion of membrane-enveloped  viruses such as influenza virus with their host cell membranes so  as to release the viral nucleic acids into the host cell cytoplasm.   1. Does trans -oleic acid have a higher or lower melting point than  cis -oleic acid? Explain.   2. Which triacylglycerol yields more energy on oxidation: one con¬  taining three residues of linolenic acid or three residues of stearic  acid?   3. Draw the structure of a glycerophospholipid that has a saturated  C 1( 5 fatty acyl group at position 1, a monounsaturated C 18 fatty  acyl group at position 2, and an ethanolamine head group.   H 3 C(CH 2 ) 5 CH O CH 2 — O—C—(CH 2 )i 2 CH 3   3. How many different types of triacylglycerols could incorporate  the fatty acids shown in Fig. 9-1?   (b) phospholipase A 2 ; (c) phospholipase C; (d) phospholipase D?   7. Which of the glycerophospholipid head groups listed in Table  9-2 can form hydrogen bonds?   8. Does the phosphatidylglycerol “head group” of cardiolipin (Table  9-2) project out of a lipid bilayer like other glycerophospholipid  head groups?   9. Sphingosine-1-phosphate can serve as a cell signaling molecule,  (a) Draw the structure of this compound, (b) What chemical re¬  actions must occur in converting a sphingomyelin molecule to  sphingosine-1 -phosphate?    10. Lipids known as sulfatides occur in cells of the central nervous  system. To which class of lipid does the sulfatide shown below  belong? How does it differ from a typical member of that class?   11. In some autoimmune diseases, an individual develops antibodies  that recognize cell constituents such as DNA and phospholipids.  Some of the antibodies react with both DNA and phospholipids.  What is the structural basis for this cross-reactivity?   12. Most hormones, such as peptide hormones, exert their effects by  binding to cell-surface receptors. However, steroid hormones do  so by binding to cytosolic receptors. How is this possible?   13. Animals cannot synthesize linoleic acid (a precursor of arachi-  donic acid) and therefore must obtain this essential fatty acid  from their diet. Explain why cultured animal cells can survive in  the absence of linoleic acid.   14. Many bacterial fatty acids contain branches and even rings. What  effect do these lipids have on membrane fluidity, compared to  their straight-chain counterparts?   17. When bacteria growing at 20°C are warmed to 30°C, are they  more likely to synthesize membrane lipids with (a) saturated or  unsaturated fatty acids, and (b) short-chain or long-chain fatty  acids? Explain.   18. Do membranes of the Antarctic icefish (see page 176) have longer  and/or more saturated fatty acids compared to tropical fish?  Explain.   19. (a) How many turns of an a helix are required to span a lipid  bilayer (~30 A across)? (b) What is the minimum number of  residues required? (c) Why do most transmembrane helices con¬  tain more than the minimum number of residues?   20. The distance between the C a atoms in a P sheet is ~3.5 A. Can  a single 9-residue segment with a P conformation serve as the  transmembrane portion of an integral membrane protein?   21. Are the following lipid samples likely to correspond to the inner  or outer leaflet of a eukaryotic plasma membrane? (a) 20% phos¬  phatidylcholine, 13% phosphatidylserine, 63% other lipids,  (b) 35% phosphatidylcholine, 15% gangliosides, 5% cholesterol,  45% other lipids.   22. Describe the labeling pattern of glycophorin A when a membrane-  impermeable protein-labeling reagent is added to (a) a prepara¬  tion of solubilized erythrocyte proteins; (b) intact erythrocyte   ghosts; and (c) erythrocyte ghosts that are initially leaky and then  immediately sealed and transferred to a solution that does not  contain the labeling reagent.   23. Predict the effect of a mutation in signal peptidase that narrows  its specificity so that it cleaves only between two Leu residues.   24. Explain why a drug that interferes with the disassembly of a  SNARE complex would block neurotransmission.    Some of the more unusual  membrane lipids occur in prokaryotes. For example, archaeal mem¬  branes are not built from glycerophospholipids. What lipids are used,  and how do their physical properties affect archaeal membranes?  How would you describe the structure of the lipopolysaccharides that  occur in gram-negative bacteria? Why are these molecules also  known as endotoxins? A certain group of bacteria synthesize unusual  lipids called ladderanes. How do these molecules help the bacteria  perform their specialized metabolic functions?   Edidin, M., Lipids on the frontier: a century of cell-membrane bilayers,  Nature Rev. Mol. Cell Biol. 4, 414-418 (2003). [A short account of  the history of the study of membranes.]   Engelman, D.M., Membranes are more mosaic than fluid, Nature 438,  578-580 (2005). [A brief review updating the classic model with  more proteins and variable bilayer thickness.]   Gurr, M.I., Harwood, J.L., and Frayn, K.N., Lipid Biochemistry: An  Introduction (5th ed.), Blackwell Science (2002).   van Meer, G., Voelker, D.R., and Feigenson, G.W., Membrane lipids:  where they are and how they behave, Nature Rev. Mol. Cell Biol. 9,  112-124 (2008).   Vance, D.E. and Vance, J. (Eds.), Biochemistry of Lipids, Lipoproteins, and  Membranes (5th ed.), Elsevier (2008).   Fujiyoshi, Y. and Unwin, N., Electron crystallography of proteins in  membranes, Curr. Opin. Struct. Biol. 18, 587-592 (2008).   Grum, V.L., Li, D., MacDonald, R.I., and Mondragon, A., Structures of  two repeats of spectrin suggest models of flexibility, Cell 98,  523-535 (1999).   Popot, J.-L. and Engelman, D.M., Helical membrane protein folding, sta¬  bility, and evolution, Annu. Rev. Biochem. 69, 881-922 (2000).  [Shows a number of protein structures and discusses many features  of transmembrane proteins.]   Sharom, F.J., and Lehto, M.T., Glycosylphosphatidylinositol-anchored  proteins: structure, function, and cleavage by phosphatidylinositol-  specific phospholipase C, Biochem. Cell Biol. 80, 535-549 (2002).  [Includes a discussion of the GPI anchor and its importance for pro¬  tein localization and function.]   Subramaniam, S., The structure of bacteriorhodopsin: an emerging con¬  sensus, Curr. Opin. Struct. Biol. 9, 462-468 (1999). [Compares the  six structures of bacteriorhodopsin that have been independently de¬  termined by electron or X-ray crystallography and finds them to be  remarkably similar.]   Wimley, W.C., The versatile P-barrel membrane protein, Curr. Opin.  Struct. Biol. 13, 404-411 (2003). [Reviews the basic principles of  construction for transmembrane P barrels.]    Zhang, F.L. and Casey, P.J., Protein prenylation: molecular mechanisms and  functional consequences, Annu. Rev. Biochem. 65, 241-269 (1996).   Alder, N.N. and Johnson, A.E., Cotranslational membrane protein bio¬  genesis at the endoplasmic reticulum, J. Biol. Chem. 279,  22787-22790 (2004).   Cross, B.C.S., Sinning, I., Luirink, J., and High, S., Delivering proteins  for export from the cytosol, Nature Rev. Mol. Cell Biol. 10, 255-264  (2009).   Keenan, R.J., Fraymann, D.M., Stroud, R.M., and Walter, P., The signal  recognition particle, Annu. Rev. Biochem. 70, 755-775 (2001).   van den Berg, B., Clemons, W.M., Jr., Collinson, I., Modis, Y., Hart¬  mann, E., Harrison, S.C., and Rapaport, T.A., X-Ray structure of a  protein-conducting channel, Nature 427, 36-44 (2004). [The X-ray  structure of SecY.]   Brodsky, F.M., Chen, C.-Y., Knuehl, C., Towler, M.C., and Wakeham,  D.E., Biological basket weaving: formation and function of clathrin-  coated vesicles, Annu. Rev. Cell Dev. Biol. 17, 515-568 (2001).   Skehel, J.J. and Wiley, D.C., Receptor binding and membrane fusion in  virus entry: The influenza hemagglutinin, Annu. Rev. Biochem. 69,  531-569 (2000).   Wickner, W. and Schekman, R. Membrane fusion, Nature Struct. Mol.  Biol. 15, 658-664 (2008).   Ungar, D. and Hughson, F.M., SNARE protein structure and function,  Annu. Rev. Cell Dev. Biol. 19, 493-517 (2003).   Ungermann, C. and Langosch, D., Functions of SNAREs in intracellu¬  lar membrane fusion and lipid bilayer mixing. J. Cell Sci. 118,  3819-3828 (2005).   A lonophores Carry Ions across Membranes  B Porins Contain (3 Barrels  C Ion Channels Are Highly Selective  D Aquaporins Mediate the Transmembrane  Movement of Water   A The (Na + -K + )-ATPase Transports Ions in  Opposite Directions   B The Ca 2+ -ATPase Pumps Ca 2+ Out of the  Cytosol    This scanning electron micrograph shows a kidney glomerulus, the structure that filters  small molecules from the blood. In order to eliminate the waste materials and retain the  nutrients present in the filtrate, kidney cells rely on a variety of membrane proteins to  transport water, ions, and other substances. [Susumu Nishinaga/Photo Researchers, Inc.]    Cells are separated from their environments by plasma membranes. Eukaryotic  cells, in addition, are compartmentalized by intracellular membranes that form  the boundaries and internal structures of their various organelles. Biological  membranes present formidable barriers to the passage of ionic and polar sub¬  stances, so these substances can traverse membranes only through the action of spe¬  cific transport proteins. Such proteins are therefore required to mediate the  transmembrane movements of ions, such as Na + , K + , Ca 2+ , and Cl' , as well  as metabolites such as pyruvate, amino acids, sugars, and nucleotides, and even  water. Transport proteins are also responsible for all biological electrochemical  phenomena such as neurotransmission. More complicated processes (e.g., en-  docytosis) are required to move larger substances such as proteins and macro-  molecular aggregates across membranes.   We begin our discussion of membrane transport by considering the ther¬  modynamics of this process. We will then examine the structures and mech¬  anisms of several different types of transport systems.    • The free energy change for moving a substance across a membrane depends on  the concentrations on each side of the membrane and, for ions, on the membrane  potential.   • For a substance that cannot diffuse directly across a membrane, transport may be  mediated by a protein and may require free energy input.    The diffusion of a substance between two sides of a membrane   thermodynamically resembles a chemical equilibration. We saw in Section  1-3D that the free energy of a solute, A, varies with its concentration:   where G A is the chemical potential (partial molar free energy) of A (the bar  indicates quantity per mole) and G A is the chemical potential of its standard  state. Thus, a difference in the concentrations of the substance on two sides  of a membrane generates a chemical potential difference:   A G a = G A (in) — G A (out) = ^Tln^-i!^ —^ [10-2]    Consequently,_if the concentration of A outside the membrane is greater than  that inside, A G A for the transfer of A from outside to inside will be negative  and the spontaneous net flow of A will be inward. If, however, [A] is greater  inside than outside, A G A is positive and an inward net flow of A can occur  only if an exergonic process, such as ATP hydrolysis, is coupled to it to make  the overall free energy change negative (see Sample Calculation 10-1).   The transmembrane movement of ions also results in charge differences  across the membrane, thereby generating an electrical potential difference,  A'T' = ty(in) — ^(out), where is termed the membrane potential.  Consequently, if A is ionic, Eq. 10-2 must be amended to include the electri¬  cal work required to transfer a mole of A across the membrane from outside  to inside:    where Z A is the ionic charge of A; 3% the Faraday constant, is the charge_of a  mole of electrons (96,485 C • mol 1 ; C is the symbol for coulomb); and G A is  now termed the electrochemical potential of A (see Sample Calculation 10-2).  The membrane potentials of living cells are commonly as large as —100 mV  (inside negative; note that 1 V = 1 J • C X ), which for a 50-A-thick mem¬  brane corresponds to a voltage gradient of 200,000 V • cm 1 . Hence, the last  term in Eq. 10-3 is often significant for ionic substances, particularly in mi¬  tochondria (Chapter 18) and in neurotransmission (Section 10-2C).    Show that A G < 0 when Ca  from the endoplasmic reticulum (where  [Ca 2+ ] = 1 mM) to the cytosol (where  [Ca 2+ ] = 0.1 |xM). Assume AT 1 = 0.    Transport May Be Mediated or Nonmediated. There are two types of trans¬  port processes: nonmediated transport and mediated transport. Nonmediated  transport occurs through simple diffusion. In contrast, mediated transport oc¬  curs through the action of specific carrier proteins. The driving force for the    Calculate the free energy required to move 1 mol of Na + ions from outside the  cell (where [Na + ] = 150 mM) to the inside (where [Na + ] = 5 mM) when the  membrane potential is -70 mV and the temperature is 37°C.   + (1)(96,485 J • V" 1 • mol _1 )(—0.070 V)   = -8763 J-mol -1 - 6754 J-mol -1  = —15,500 J • mol -1 = —15.5 kj • mol -1   • How can you predict whether it will be  thermodynamically favorable for an un¬  charged substance to move from one side  of a membrane to the other?   • If the substance is charged, what addi¬  tional information do you need in order to  predict whether its transmembrane move¬  ment will be spontaneous?   • Give an example of a substance that can  diffuse across a membrane. Give an exam¬  ple of a substance that must cross a  membrane by mediated transport.    nonmediated flow of a substance through a medium is its chemical potential  gradient. Thus, the substance diffuses in the direction that eliminates its concentra¬  tion gradient , at a rate proportional to the magnitude of the gradient. The rate of  diffusion of a substance also depends on its solubility in the membrane's nonpolar  core. Consequently, nonpolar molecules such as steroids and 0 2 readily diffuse  through biological membranes by nonmediated transport, according to their  concentration gradients across the membranes.   Mediated transport is classified into two categories depending on the ther¬  modynamics of the system:   1. Passive-mediated transport, or facilitated diffusion, in which a spe¬  cific molecule flows from high concentration to low concentration.   2. Active transport, in which a specific molecule is transported from low  concentration to high concentration, that is, against its concentration  gradient. Such an endergonic process must be coupled to a sufficiently  exergonic process to make it favorable (i.e., AG < 0).    • Passive-mediated transport is carried out by ionophores, porins, ion channels,  aquaporins, and transport proteins.   • The coordinated opening and closing of ion channels generates an action poten¬  tial in nerve cells.    Substances that are too large or too polar to diffuse across lipid bilayers on their  own may be conveyed across membranes via proteins or other molecules that  are variously called carriers, permeases, channels, and transporters. These  transporters operate under the same thermodynamic principles but vary widely  in structure and mechanism, particularly as it relates to their selectivity.   Ionophores are organic molecules of diverse types, usually of bacterial origin,  that increase the permeability of membranes to ions. These molecules often  exert an antibiotic effect by discharging the vital ion concentration gradients  that cells actively maintain.   1. Carrier ionophores, which increase the permeabilities of membranes to  their selected ion by binding it, diffusing through the membrane, and re¬  leasing the ion on the other side (Fig. 10-1 d). For net transport to occur,  the uncomplexed ionophore must then return to the original side of the  membrane ready to repeat the process. Carriers therefore share the com¬  mon property that their ionic complexes are soluble in nonpolar solvents.   2. Channel-forming ionophores , which form transmembrane channels or  pores through which their selected ions can diffuse (Fig. 10-1 b).   Both types of ionophores transport ions at a remarkable rate. For example, a  single molecule of the carrier ionophore valinomycin transports up to 1 0 4 K +  ions per second across a membrane. However, since ionophores passively permit  ions to diffuse across a membrane in either direction, their effect can only be to  equilibrate the concentrations of their selected ions across the membrane.   FIG. 1 0-1 Ionophore action, (a) Carrier  ionophores transport ions by diffusing through  the lipid bilayer, (b) Channel-forming ionophores  span the membrane with a channel through  which ions can diffuse.    Valinomycin, which is one of the best characterized ionophores, specifically  binds K + . It is a cyclic molecule containing D- and L-amino acid residues that  participate in ester linkages as well as peptide bonds (Fig. 10-2#). The X-ray  structure of valinomycin s K + complex (Fig. 10-2 b) indicates that the K + ion is  octahedrally coordinated by the carbonyl groups of its six Val residues, and the  cyclic valinomycin backbone surrounds the K coordination shell. The methyl  and isopropyl side chains project outward to provide the complex with a non¬  polar exterior that makes it soluble in the hydrophobic cores of lipid bilayers.   The K + ion (ionic radius, r = 1.33 A) fits snugly into valinomycins coor¬  dination site, but the site is too large for Na + (r = 0.95 A) or Li + (r = 0.60 A)  to coordinate with all six carbonyl oxygens. Valinomycin therefore has 10,000-fold  greater binding affinity for K + than for Na + . No other known substance dis¬  criminates better between Na + and K + .   The porins, introduced in Section 9-3A, are (3 barrel structures with a central  aqueous channel. In the E. coli OmpF porin (Fig. 9-23), the channel is con¬  stricted to form an elliptical pore with a minimum cross section of 7 X 11 A.  Consequently, solutes of more than ~600 D are too large to pass through the    FIG. 1 0-2 Valinomycin. (a) This cyclic ionophore contains ester and  amide bonds and d- as well as L-amino acids. ( b ) X-Ray structure of  valinomycin in complex with a K + ion colored according to atom type  (C green, H white, N blue, 0 red, and K + purple) and embedded in its  molecular surface (blue mesh). Note that the K + ion is octahedrally  coordinated by the carbonyl oxygen atoms of six Val residues and that the  surface of the complex is largely covered with methyl and isopropyl groups.  [Based on an X-ray structure by Max Dobler, ETH, Zurich, Switzerland.]   FIG. 1 0-3 Structure of a maltoporin  subunit in complex with a maltodextrin of six  glycosyl units. The polypeptide backbone of this  E. coli protein is represented by a multithreaded  cyan ribbon. Five glucose residues of  maltodextrin and the aromatic side chains lining  the transport channel are shown in space-filling  form with N blue, 0 red, protein C gold, and  glucosyl C green. The “greasy slide” consists of  the aromatic side chains of six residues. Tyr 118,  which projects into the channel, helps restrict  passage to glucosyl residues. [Based on an  X-ray structure by Tilman Schirmer, University of  Basel, Basel, Switzerland. PDBid 1MP0J    channel. OmpF is weakly cation selective; other porins are more selective for  anions. In general, the size of the channel and the residues that form its walls  determine what types of substances can pass through.   Solute selectivity is elegantly illustrated by maltoporin. This bacterial  outer membrane protein facilitates the diffusion of maltodextrins, which are  the a(l—>4)-linked glucose oligosaccharide degradation products of starch  (Section 8-2C). The X-ray structure of E. coli maltoporin reveals that the pro¬  tein is structurally similar to OmpF porin but is a homotrimer of 18-stranded  rather than 16-stranded antiparallel (3 barrels. Three long loops from the ex¬  tracellular face of each maltoporin subunit fold inward into the barrel, thereby  constricting the channel near the center of the membrane to a diameter of  ~5 A and giving the channel an hourglass-like cross section. The channel is  lined on one side with a series of six contiguous aromatic side chains arranged  in a left-handed helical path that matches the left-hand helical curvature of  a-amylose (Fig. 8-10). This so-called greasy slide extends from one end of the  channel, through its constriction, to the other end (Fig. 10-3).   How does the greasy slide work? The hydrophobic faces of the maltodex¬  trin glucose residues stack on aromatic side chains, as is often observed in  complexes of sugars with proteins. The glucose hydroxyl groups, which are  arranged in two strips along opposite edges of the maltodextrins, form numer¬  ous hydrogen bonds with polar and charged side chains that line the channel.  Tyr 118, which protrudes into the channel opposite the greasy slide, apparently  functions as a steric barrier that only permits the passage of near-planar groups  such as glucosyl residues. Thus, the hook-shaped sucrose (a glucose-fructose  disaccharide) passes only very slowly through the maltoporin channel.   At the start of the translocation process, the entering glucosyl residue in¬  teracts with the readily accessible end of the greasy slide in the extracellular  vestibule of the channel. Further translocation along the helical channel re¬  quires the maltodextrin to follow a screwlike path that maintains the helical  structure of the oligosaccharide, much like the movement of a bolt through a  nut, thereby excluding molecules of comparable size that have different shapes.  The translocation process is unlikely to encounter any large energy barrier due to  the smooth surface of the greasy slide and the multiple polar groups at the chan¬  nel constriction that would permit the essentially continuous exchange of hydrogen  bonds as a maltodextrin moves through the constriction.   All cells contain ion-specific channels that allow the rapid passage of ions such  as Na + , K + , and Cl . The movement of these ions through such channels, along  with their movement through active transporters (discussed in Section 10-3),  is essential for maintaining osmotic balance, for signal transduction (Section  13-4A), and for effecting changes in membrane potential that are responsible  for neurotransmission. Mammalian cells, for example, maintain a nonequilib¬  rium distribution of ions on either side of the plasma membrane: — 150 mM  Na + and ~4 mM K + in the extracellular fluid, and ~12 mM Na + and ~140  mM K + inside the cell.   The Structure of the KcsA K + Channel Explains Its Selectivity and  Speed. Potassium ions passively diffuse from the cytoplasm to the extracellular  space through transmembrane proteins known as K + channels. Although there  is a large diversity of K + channels, even within a single organism, all of them  have similar sequences, exhibit comparable permeability characteristics, and most  importantly, are at least 10,000-fold more permeable to K + than Na + . Since  this high selectivity (around the same as that of valinomycin; Section 10-2A)  implies energetically strong interactions between K + and the protein, how can  the K + channel maintain its observed nearly diffusion-limited throughput rate  of up to 10 8 ions per second (a 10 4 -fold greater rate than that of valinomycin)?   FIG. 1 0-4 X-Ray structure of the KcsA K + channel, (a) Ribbon diagram of the  tetramer as viewed from within the plane of the membrane, with the cytoplasm below and  the extracellular region above. The protein’s fourfold axis of rotation is vertical and each of  its identical subunits is differently colored. Each subunit has an inner helix that forms part  of the central pore, an outer helix that contacts the membrane interior, and a turret that  projects out into the extracellular space. The selectivity filter at the extracellular end of the  protein allows the passage of K + ions but not Na + ions, (b) Cutaway diagram viewed  similarly to Part a in which the K + channel is represented by its solvent-accessible surface.  The surface is colored according to its physical properties with negatively charged areas  red, uncharged areas white, positively charged areas blue, and hydrophobic areas of the  central pore yellow. K + ions are represented by green spheres. [Courtesy of Roderick  MacKinnon, Rockefeller University. PDBid 1BL8J    One of the best characterized ion channels is a K + channel from  Streptomyces lividans named KcsA. This 158-residue integral membrane pro¬  tein, like all known K + channels, functions as a homotetramer. The X-ray  structure of KcsAs N-terminal 125-residue segment, determined by Roderick  MacKinnon, reveals that each of its subunits contains two nearly parallel trans¬  membrane helices plus a shorter helix (Fig. 10-4^). Four such subunits asso¬  ciate to form a fourfold symmetric assembly surrounding a central pore. The  four inner helices, which largely form the pore, pack against each other near  the cytoplasmic side of the membrane much like the poles of an inverted  teepee. The four outer helices, which face the lipid bilayer, buttress the inner  helices. The central pore can accommodate several K + ions (Fig. 10-4£).   The 45-A-long central pore has variable width: It starts at its cytoplasmic  side as an ~6-A-diameter tunnel whose entrance is lined with four anionic  side chains (red area at the bottom of Fig. 10-4£) that presumably attract  cations and repel anions. The pore then widens to form an ~ 10-A-diameter  cavity. These regions of the central pore are wide enough so that a K + ion  could move through them in its hydrated state. However, the upper part of  the pore, called the selectivity filter, narrows to 3 A, thereby forcing a transit¬  ing K + ion to shed its waters of hydration. The walls of the pore and the cav¬  ity are lined with hydrophobic groups that interact minimally with diffusing  ions (yellow area of the pore in Fig. 10-4£). However, the selectivity filter (red   FIG. 1 0-5 Portions of the KcsA K +  channel responsible for its ion selectivity. The   protein is viewed similarly to Fig. 10-4 but with  the front and back subunits omitted for clarity.  The residues forming the cavity ( bottom ) and  selectivity filter (top) are shown with atoms  colored according to type (C yellow, N blue, 0  red, and K + ions represented by green spheres).  The water and protein 0 atoms that ligand the  K + ions, including those contributed by the front  and back subunits, are represented by red  spheres. The coordination polyhedra formed by  these 0 atoms are outlined by thin white lines.  [Based on an X-ray structure by Roderick  MacKinnon, Rockefeller University. PDBid  1 K4C.] Q See Interactive Exercise 5.   How does the K + channel discriminate so acutely between K + and  Na + ions? The main chain O atoms lining the selectivity filter form a  stack of rings (Fig. 10-5, top) that provide a series of closely spaced sites  of appropriate dimensions for coordinating dehydrated K + ions but not  the smaller Na + ions. The structure of the protein surrounding the se¬  lectivity filter suggests that the diameter of the pore is rigidly main¬  tained, thus making the energy of a dehydrated Na + in the selectivity  filter considerably higher than that of hydrated Na + and thereby ac¬  counting for the K + channels high selectivity for K + ions.   What is the function of the cavity? Energy calculations indicate that an  ion moving through a narrow transmembrane pore must surmount an en¬  ergy barrier that is maximal at the center of the membrane. The existence  of the cavity reduces this electrostatic destabilization by surrounding the  ion with polarizable water molecules (Fig. 10-5, bottom). The cavity holds  ~40 additional water molecules, which are disordered and therefore not  seen in the X-ray structure. Remarkably, the K + ion occupying the cav¬  ity is liganded by 8 ordered water molecules located at the corners of a  square antiprism (a cube with one face twisted by 45° with respect to the  opposite face). K + in aqueous solution is known to have such an inner  hydration shell but it had never before been visualized.   How does KcsA support such a high throughput of K + ions (up to  10 8 ions per second)? Figure 10-5 shows a string of regularly spaced K +  ions: four in the selectivity filter and two more just outside it on its ex¬  tracellular (top) side. Such closely spaced positive ions would strongly  repel one another and hence represent a high energy situation. However,  the X-ray structure is an average of many KcsA molecules, and a vari¬  ety of evidence indicates that within a single channel, the K + ions in the pore  actually alternate with water molecules. This arrangement means that each K +  ion is surrounded by O atoms (from H 2 0 or protein carbonyl groups) at each  position along the selectivity filter. As a K + ion moves into the selectivity fil¬  ter from the cavity, it exchanges some of its hydrating water molecules for pro¬  tein ligands, then does the reverse to restore its hydration shell when it exits  the selectivity filter to enter the extracellular solution. Within the selectivity  filter, the ligands are spaced and oriented such that there is little free energy  change (< 12 kj • mol X ) as a K + ion moves to successive positions. This level  free energy landscape allows the rapid movement of K + ions through the ion  channel. In addition, mutual electrostatic repulsions between successive K +  ions balance the attractive interactions holding these ions in the selectivity fil¬  ter and hence further facilitate their rapid transit.    Ion Channels Are Gated. The physiological functions of ion channels depend  not only on their exquisite ion specificity and speed of transport but also on  their ability to be selectively opened or closed. For example, the ion gradients  across cell membranes, which are generated by specific energy-driven pumps  (Section 10-3), are discharged through Na + and K + channels. However, the  pumps could not keep up with the massive fluxes of ions passing through the  open channels, so ion channels are normally shut and only open transiently to  perform some specific task for the cell. The opening and closing of ion channels,  a process known as gating, can occur in response to a variety of stimuli:    1. Mechanosensitive channels open in response to local deformations in  the lipid bilayer. Consequently, they respond to direct physical stimuli  such as touch, sound, and changes in osmotic pressure.   2. Ligand-gated channels open in response to an extracellular chemical  stimulus such as a neurotransmitter.   4. Voltage-gated channels open in response to a change in membrane  potential. Multicellular organisms contain numerous varieties of voltage¬  gated channels, including those responsible for generating nerve impulses.   Nerve Impulses Are Propagated by Action Potentials. As an example of the  functions of voltage-gated channels, let us consider electrical signaling events  in neurons (nerve cells). The stimulation of a neuron, a cell specialized for  electrical signaling, causes Na + channels to open so that Na + ions sponta¬  neously flow into the cell. The consequent local increase in membrane poten¬  tial induces neighboring voltage-gated Na + channels to open. The resulting  local depolarization of the membrane induces nearby voltage-gated K + chan¬  nels to open. This allows K + ions to spontaneously flow out of the cell in a  process called hyperpolarization (Fig. 10-6). However, well before the distri¬  bution of Na + and K + ions across the membrane equilibrates, the Na + and  K + channels spontaneously close. Yet, because depolarization of a membrane  segment induces the voltage-gated Na + channels in a neighboring membrane  segment to open, which induces their neighboring voltage-gated Na + chan¬  nels to open, etc., a wave of transient change in the membrane potential, called  an action potential, travels along the length of the nerve cell (which may be  over 1 m long). The action potential, which travels at ~10 m/s, propagates  in only one direction because, after the ion channels have spontaneously  closed, they resist reopening until the membrane potential has regained its rest¬  ing value, which takes a few milliseconds (Fig. 10-6).   As an action potential is propagated along the length of a nerve cell, it is  continuously renewed so that its signal strength remains constant (in contrast,  an electrical impulse traveling down a wire dissipates as a consequence of resis¬  tive and capacitive effects). Nevertheless, the relative ion imbalance responsible  for the resting membrane potential is small; only a tiny fraction of a nerve cells  Na + —K + gradient (which is generated by ion pumps; Section 10-3A) is discharged  by a single nerve impulse (only one K + ion per 3000-300,000 in the cytosol  is exchanged for extracellular Na + as indicated by measurements with radioac¬  tive Na + ). A nerve cell can therefore transmit a nerve impulse every few mil¬  liseconds without letup. This capacity to fire rapidly is an essential feature of  neuronal communications: Since action potentials all have the same amplitude,  the magnitude of a stimulus is conveyed by the rate at which a nerve fires.   FIG. 1 0-6 Time course of an action potential, (a) The neuron  membrane undergoes rapid depolarization, followed by a nearly as  rapid hyperpolarization and then a slow recovery to its resting  potential, (b) The depolarization is caused by a transient increase in   Na + permeability (conductance), whereas the hyperpolarization results  from a more prolonged increase in K + permeability that begins a  fraction of a millisecond later. [After Hodgkin, A.L and Huxley, A.F., J.  Physiol. 117, 530 (1952).]    Voltage Gating in Kv Channels Is Triggered by the Motion of a Positively  Charged Protein Helix. The subunits of all voltage-gated K + channels con¬  tain an ~220-residue N-terminal cytoplasmic domain, an ~250-residue trans¬  membrane domain consisting of six helices, SI to S6, and an ~ 150-residue  C-terminal cytoplasmic domain (Fig. 10-7). S5 and S6 are respectively  homologous to the outer and inner transmembrane helices of the KcsA chan¬  nel (Fig. 10-4), and their intervening P-loop, which forms the selectivity  filter, includes the same TVGYG signature sequence that occurs in KcsA. In  the voltage-gated K + channels known as Kv channels, a conserved, cytoplas¬  mic, ~ 100-residue so-called Tl domain precedes the transmembrane domain.   Kv channels closely resemble the KcsA channel in their tetrameric pore struc¬  ture, but what is the nature of the gating machinery in the voltage-gated ion chan¬  nels? The ~ 19-residue S4 helix, which contains around five positively charged side  chains spaced about every three residues on an otherwise hydrophobic polypep¬  tide, appears to act as a voltage sensor. Various experimental approaches indicate  that when the membrane potential increases (the inside becomes less negative), the  S4 helix is pulled toward the extracellular side of the membrane.   The X-ray structure of a Kv channel from rat brain named Kvl.2 (Fig. 10-8)  shows how gating might occur. As the membrane begins to depolarize, the four  conserved Arg residues on the S4 helix are drawn toward the extracellular sur¬  face (up in Fig. 10-9), pulling on the S4—S5 linker helix (not drawn in Fig.  10-7) and with it the S5 helix. This splays the ends of the S6 helices so as to  enlarge the intracellular entrance to the K + channel (Fig. 10-9 A). During re¬  polarization, as the cell interior becomes more negative, the positively charged  S4 helix drops back toward the cytoplasmic side of the membrane, pressing  down on the S4-S5 lever to close off the channel (Fig. 10-9c).    Ion Channels Have a Second Gate. Electrophysiological measurements in¬  dicate that Kv channels spontaneously close a few milliseconds after opening   FIG. 1 0-8 X-Ray structure of the Kvl.2 voltage-gated K +  channel, (a) View along the tetrameric protein’s fourfold axis from the  extracellular side of the membrane in which it is embedded. Each of  its four identical subunits is colored differently, and the T1 domain has  been omitted for clarity The fragmented appearance of the  polypeptide chains is due to the high mobilities of the missing  segments. The S5 and S6 helices with their intervening P loops form  the pore for K + ions (delineated by green spheres). Helices SI to S4  form a separate intramembrane voltage-sensing domain that    associates with the S5 and S6 helices of the clockwise adjacent  subunit, (b) View perpendicular to that in Part a with the extracellular  side of the membrane above. The pore and voltage sensing domains  span a distance of 30 A, the thickness of the membrane’s  hydrophobic core. The Tl domain, which occupies the cytoplasm,  forms the vestibule of the transmembrane K + channel. The four large  openings between the Tl domain and the K + channel are the portals  through which K + ions enter the K + channel. [Based on an X-ray  structure by Roderick MacKinnon, Rockefeller University. PDBid 2A79.]   FIG. 1 0-9 Operation of the transmembrane domain of the  Kvl.2 voltage-gated K + channel, (a) Side view of a single subunit  with its extracellular side at the top. The pore-forming helices (S5 and  S6) are gray, and the voltage-sensor domain (helices S1-S4) is blue.  The S4-S5 linker helix (not drawn in Fig. 10-7), which is parallel to  the membrane surface, connects the voltage-sensing domain to the  domain forming the K + pore (delineated by green spheres). Four Arg  residues on S4 are shown in ball-and-stick form with C atoms gold  and N atoms blue. Two of these residues interact with the protein,    and two project into the lipid bilayer. ( b ) Side view of the channel  tetramer in its open conformation. The S5 helices are gray, the S6  helices are blue, and the S4-S5 linker helix is red. (c) Hypothetical  model of the channel in its closed conformation, colored as in Part b.  The downward movement of the Arg-bearing S4 helix (not shown in  this drawing) pushes down on the S4-S5 linker helix to pinch off the  pore at its cytoplasmic end (bottom). [Courtesy of Roderick  MacKinnon, Rockefeller University. PDBid 2A79.]   and do not reopen until after the membrane has regained its resting mem¬  brane potential. Evidently, the Kv channel contains two voltage-sensitive gates ,  one to open the channel on an increase in membrane potential and one to inac¬  tivate it a short time later. This inactivation of the Kv channel is abolished by  proteolytically excising its N-terminal 20-residue segment, whose NMR struc¬  ture is a ball-like assembly. In the intact Kv channel, this “inactivation ball”  is tethered to the end of a flexible 65-residue peptide segment (Fig. 10-7), sug¬  gesting that channel inactivation normally occurs when the ball swings around  to bind in the mouth of the open K + pore, thereby blocking the passage of  K + ions. The mobility of this “ball and chain” is presumably why it is not  visible in the X-ray structure of Kvl.2 (Fig. 10-8).   The Tl tetramer does not have an axial channel, so the inactivation ball  must find its way to block the central pore through the side portals between  the Tl tetramer and the central pore (Fig. 10-8^). These portals, which are  15 to 20 A across, are lined with negatively charged groups that presumably  attract K + ions.   The cytoplasmic entrance to the K + channel pore is only 6 A in diame¬  ter, too narrow to admit the inactivation ball. Therefore, it appears that the ball  peptide must unfold in order to enter the pore. The first 10 residues of the un¬  folded ball peptide are predominantly hydrophobic and presumably make  contact with the hydrophobic residues lining the Kv channel pore. The next  10 residues, which are largely hydrophilic and contain several cationic groups,  bind to anionic groups lining the entrance to the side portals in Tl. Thus, the  inactivation peptide acts more like a snake than a ball and chain. A Kv chan¬  nel engineered so that only one subunit has an inactivation peptide still  becomes inactivated but at one-fourth the rate of normal Kv channels.  Apparently, any of the normal Kv channels four inactivation peptides can block  the channel and it is simply a matter of chance as to which one does so.   Other Voltage-Gated Cation Channels Contain a Central Pore. Voltage¬  gated Na + and Ca 2+ channels appear to resemble K + channels, although  rather than forming homotetramers, they are monomers of four consecutive  domains, each of which is homologous to a K + channel subunit, separated by  often large cytoplasmic loops. These domains presumably assume a pseudo-  tetrameric arrangement about a central pore resembling that of voltage-gated K +   Chapter 10 Membrane Transport channels. This structural homology suggests that voltage-gated ion channels   share a common architecture in which differences in ion selectivity arise from  precise stereochemical variations within the central pore. However, outside of  their conserved transmembrane core, voltage-gated ion channels with differ¬  ent ion selectivities are highly divergent. For example, the Tl domain of Kv  channels is absent in other types of voltage-gated ion channels.   Cl - Channels Differ from Cation Channels. Cl - channels, which occur in  all cell types, permit the transmembrane movement of chloride ions along their  concentration gradient. In mammals, the extracellular Cl concentration is  ~120 mM and the intracellular concentration is ~4 mM.   C1C Cl channels form a large family of anion channels that occur widely  in both prokaryotes and eukaryotes. The X-ray structures of C1C Cl chan¬  nels from two species of bacteria, determined by Raimund Dutzler and  MacKinnon, reveal, as biophysical measurements had previously suggested,  that C1C Cl channels are homodimers with each subunit forming an anion-  selective pore (Fig. 10-10). Each subunit consists mainly of 18 mostly trans¬  membrane a helices that are remarkably tilted with respect to the membrane  plane and have variable lengths compared to the transmembrane helices in  other integral proteins of known structures.   The specificity of the Cl channel results from an electrostatic field es¬  tablished by basic amino acids on the protein surface, which helps funnel an¬  ions toward the pore, and by a selectivity filter formed by the N-terminal  ends of several a helices. Because the polar groups of an a helix are all aligned  (Fig. 6-7), it forms a strong electrical dipole with its N-terminal end posi¬  tively charged. This feature of the selectivity filter helps attract Cl ions,  which are specifically coordinated by main chain amide nitrogens and side  chain hydroxyls from Ser and Tyr residues. A positively charged residue such  as Lys or Arg, if it were present in the selectivity filter, would probably bind  a Cl ion too tightly to facilitate its rapid transit through the channel.   Unlike the K + channel, which has a central aqueous cavity (Fig. 10-4£),  the Cl channel is hourglass-shaped, with its narrowest part in the center of  the membrane and flanked by wider aqueous “vestibules.” A conserved Glu  side chain projects into the pore. This group would repel other anions, sug¬  gesting that rapid Cl flux requires a protein conformational change in which  the Glu side chain moves aside. Another anion could push the Glu away,  which explains why some Cl channels appear to be activated by Cl ions;  that is, they open in response to a certain concentration of Cl in the extra¬  cellular fluid.   Salmonella typhimurium. Each subunit of the homodimer contains  18 a helices of variable lengths. The subunits are drawn in ribbon  form with one colored in rainbow order from its N-terminus (blue) to  its C-terminus (red) and the other pink. The two C\~ ions bound in  the selectivity filter of each subunit are represented by pale green   spheres, (a) View from within the membrane with the extracellular  surface above and the 2-fold axis relating the two subunits vertical.   (b) View from the extracellular side of the membrane along the  molecular 2-fold axis. [Based on an X-ray structure by Raimund  Dutzler and Roderick MacKinnon, Rockefeller University. PDBid 10TSJ   The observed rapid passage of water molecules across biological membranes had  long been assumed to occur via simple diffusion that was made possible by the  small size of water molecules and their high concentrations in biological systems.  However, certain cells, such as those in the kidney, can sustain particularly rapid  rates of water transport, which can be reversibly inhibited by mercuric ions. This  suggested the existence of previously unrecognized protein pores that conduct  water through biological membranes. The first of these elusive proteins was dis¬  covered in 1992 by Peter Agre, who named them aquaporins.   Aquaporins are widely distributed in nature; plants may have as many as  50 different aquaporins. The 13 known mammalian aquaporins are expressed  at high levels in tissues that rapidly transport water, including kidneys, sali¬  vary glands, and lacrimal glands (which produce tears). Aquaporins permit the  passage of water molecules at an extremely high rate (~3 X 10 9 per second)  but do not permit the transport of solutes (e.g., glycerol or urea) or ions, in¬  cluding, most surprisingly, protons (really hydronium ions; H 3 0 + ), whose free  passage would discharge the cells membrane potential.   The most extensively characterized member of the aquaporin family,  AQP1, is a homotetrameric glycoprotein. Its X-ray and electron crystallographic  structures reveal that each of its subunits consists mainly of six transmembrane  a helices plus two shorter helices that lie within the bilayer (Fig. 10-11). These  helices are arranged so as to form an elongated hourglass-shaped central pore   FIG. 10-11 X-Ray structure of the aquaporin AQP1 from  bovine erythrocytes, (a) Ribbon diagram of an aquaporin subunit  colored in rainbow order from its N-terminus (blue) to its C-terminus  (red). The view is from within the membrane with the extracellular  surface above. The four water molecules that occupy the central  portion of the water-transport channel are represented by red spheres.  (b) View of the aquaporin tetramer from the extracellular surface. The  subunit in the upper right is drawn in space-filling form with C green,    N blue, and 0 red; that in the upper left is drawn in ribbon form  colored in rainbow order from its N-terminus (blue) to its C-terminus  (red), that in the lower left is represented by its solvent-accessible  surface; and that in the lower right displays the side chains (green) at  the constriction site. Each subunit forms a water-transport channel,  which is most clearly visible in the subunit drawn in space-filling form.  [Based on an X-ray structure by Bing Jap, University of California at  Berkeley. PDBid 1J4N.]   FIG. 1 0-1 2 Schematic drawing of the water-conducting pore of aquaporin AQP1.   The pore is viewed from within the membrane with the extracellular surface above. The  positions of residues critical for preventing the passage of protons, other ions, and small  molecule solutes are indicated. [Courtesy of Peter Agre, The Johns Hopkins School of  Medicine.]    that, at its narrowest point, the so-called constriction region, is ~2.8 A wide,  which is the van der Waals diameter of a water molecule (Fig. 10-12). Much  of the pore is lined with hydrophobic groups whose lack of strong interactions  with water molecules hastens their passage through the pore. However, for a  water molecule to transit the constriction region, it must be separated from  other water molecules to which it is hydrogen-bonded. This is facilitated by  the side chains of highly conserved Arg and His residues as well as several back¬  bone carbonyl groups that form hydrogen bonds to a transiting water mole¬  cule and hence readily displace its associated water molecules, much as occurs  with K + in the selectivity filter of the KcsA channel (Section 10-2C).   If water were to pass through aquaporin as an uninterrupted chain of  hydrogen-bonded molecules, then protons would pass even more rapidly through  the channel via proton jumping (Fig. 2-15; in order for more than one such  series of proton jumps to occur, each water molecule in the chain must reori¬  ent such that one of its protons forms a hydrogen bond to the next water mol¬  ecule in the chain). However, aquaporin interrupts this process by forming  hydrogen bonds from the side chain NH 2 groups of two highly conserved Asn  residues to a water molecule that is centrally located in the pore (Fig. 10-12).  Consequently, although this central water molecule can readily donate hydro¬  gen bonds to its neighboring water molecules in the hydrogen-bonded chain,  it cannot accept one from them nor reorient, thereby severing the “proton¬  conducting wire.”   Up to this point, we have examined the structures and functions of membrane  proteins that form a physical passageway for small molecules, ions, or water.  Membrane proteins known as connexins also form such channels, in the form  of gap junctions between cells (see Box 10-1). However, not all membrane  transport proteins offer a discrete bilayer-spanning pore. Instead, some pro¬  teins undergo conformational changes to move substances from one side of  the membrane to the other. The erythrocyte glucose transporter (also known  as GLUTl) is such a protein.   Biochemical evidence indicates that GLUTl has glucose-binding sites on  both sides of the membrane. John Barnett showed that adding a propyl group  to glucose Cl prevents glucose binding to the outer surface of the membrane,  whereas adding a propyl group to C6 prevents binding to the inner surface.  He therefore proposed that this transmembrane protein has two alternate con¬  formations: one with the glucose site facing the external cell surface, requir¬  ing Ol contact and leaving 06 free, and the other with the glucose site facing  the internal cell surface, requiring 06 contact and leaving Ol free. Transport  apparently occurs as follows (Fig. 10-13):   1. Glucose binds to the protein on one face of the membrane.   2. A conformational change closes the first binding site and exposes the  binding site on the other side of the membrane (transport).   4. The transport cycle is completed by the reversion of GLUTl to its ini¬  tial conformation in the absence of bound glucose (recovery).   This transport cycle can occur in either direction, according to the relative  concentrations of intracellular and extracellular glucose. GLUTl provides a    Most eukaryotic cells are in metabolic as well as physical contact  with neighboring cells. This contact is brought about by tubular  structures, named gap junctions, that join discrete regions of  neighboring plasma membranes much like hollow rivets. Gap junc¬  tions consist of two apposed plasma membrane-embedded com¬  plexes. Small molecules and ions, but not macromolecules, can  pass between cells via the gap junction’s central channel.    These intercellular channels are so widespread that many whole  organs are continuous from within. Thus, gap junctions are  important intercellular communication channels. For example, the  synchronized contraction of heart muscle is brought about by  flows of ions through gap junctions, and gap junctions serve as  conduits for some of the substances that mediate embryonic  development.   Mammalian gap junction channels are 16 to 20 A in diameter,  which Werner Loewenstein established by microinjecting single  cells with fluorescent molecules of various sizes and observing  with a fluorescence microscope whether the fluorescent probe  passed into neighboring cells. The molecules and ions that can  pass freely between neighboring cells are limited in molecular  mass to a maximum of —1000 D; macromolecules such as pro¬  teins and nucleic acids cannot leave a cell via this route.   The diameter of a gap junction channel varies with Ca 2+ con¬  centration: The channels are fully open when the Ca 2+ level is  <10 -7 M and become narrower as the Ca 2+ concentration in¬  creases until, above 5 X 10 -5 M, they close. This shutter system  is thought to protect communities of interconnected cells from the  otherwise catastrophic damage that would result from the death  of even one of their members. Cells generally maintain very low  cytosolic Ca 2+ concentrations (<10 -7 M) by actively pumping  Ca 2+ out of the cell as well as into their mitochondria and endo¬  plasmic reticulum (Section 10-3B). Ca 2+ floods back into leaky or  metabolically depressed cells, thereby inducing closure of their  gap junctions and sealing them off from their neighbors.   Gap junctions are constructed from a single sort of protein  subunit known as a connexin. A single gap junction consists of    two hexagonal rings of connexins, called connexons, one from  each of the adjoining plasma membranes. A given animal  expresses numerous genetically distinct connexins (21 in humans),  with molecular masses ranging from 25 to 50 kD. At least some  connexons may be formed from two or more species of connexins,  and the gap junctions joining two cells may consist of two different  types of connexons. These various types of gap junctions presum¬  ably differ in their selectivities for the substances they transmit.   The X-ray structure of the gap junction formed by the 226-  residue human connexin 26 , determined by Tomitake Tsukihara,  reveals a symmetrical assembly with a height of 155 A and a  maximal diameter of 92 A that encloses a central channel.   Each connexin consists of an up-and-down bundle of four a he¬  lices. In this model, the protein is drawn as ribbons embedded in a  semitransparent molecular surface. Each connexin of the upper  connexon has a different color, whereas one connexin in the lower  connexon is colored in rainbow order from its N-terminus (blue) to  its C-terminus (red) with the remaining connexins gray.   The extracellular portion of each connexin extends from the cell  surface and interdigitates with the opposite connexon by 6 A to  span an intercellular gap of 40 A. The central channel has a diam¬  eter of —40 A at its cytosolic entrance that funnels down to 14 A  near the extracellular surface of the membrane and then widens to  25 A in the extracellular space. The entrance to the channel is  positively charged and would attract negatively charged molecules.  However, the region of maximal channel constriction is negatively  charged, which should also affect the channel’s charge selectivity.    [Based on an X-ray structure by Tomitake Tsukihara, University of  Osaka, Japan. PDBid 2ZW3.]    FIG. 1 0-1 3 Model for glucose transport. The transport protein  alternates between two mutually exclusive conformations. Glucose (green)  is not drawn to scale. [After Baldwin, S.A. and Lienhard, G.E., Trends  Biochem. Sci. 6, 210 (1981).] O See the Animated Figures.   Explain why ions cannot cross a membrane via the glucose  transporter.    means of equilibrating the glucose concentration across the erythrocyte mem¬  brane without any accompanying leakage of small molecules or ions (as might  occur through an always-open channel such as a porin).   Biochemical analyses indicate that GLUTl has 12 membrane-spanning  a helices and probably forms a tetramer in the membrane. This protein belongs  to a large family of transporters that move amino acids and other small mol¬  ecules across membranes. All known transport proteins appear to be asymmetri¬  cally situated transmembrane proteins that alternate between two conformational    Glucose and many other compounds can enter cells by a non¬  mediated pathway; that is, they slowly diffuse into cells at a rate  proportional to their membrane solubility and their concentrations  on either side of the membrane. This is a linear process: The flux  (rate of transport per unit area) of a substance across the mem¬  brane increases with the magnitude of its concentration gradient  (the difference between its internal and external concentrations). If  the same substance, say glucose, moves across a membrane by  means of a transport protein, its flux is no longer linear. This is  one of four characteristics that distinguish mediated from non¬  mediated transport:   1. Speed and specificity. The solubilities of the chemically  similar sugars D-glucose and D-mannitol in a synthetic lipid  bilayer are similar. However, the rate at which glucose moves  through the erythrocyte membrane is four orders of magnitude  faster than that of D-mannitol. The erythrocyte membrane must  therefore contain a system that transports glucose and that  can distinguish D-glucose from D-mannitol.   2 . Saturation. The rate of glucose transport into an erythrocyte  does not increase infinitely as the external glucose concentra¬  tion increases: The rate gradually approaches a maximum. Such  an observation is evidence that a specific number of sites on  the membrane are involved in the transport of glucose. At high  [glucose], the transporters become saturated, much like myoglo¬  bin becomes saturated with 0 2 at high p0 2 (Fig. 7-4). As ex¬  pected, the plot of glucose flux versus [glucose] is hyperbolic.  The nonmediated glucose flux increases linearly with [glucose]  but would not visibly depart from the baseline on the scale of  the graph.   3. Competition. The above curve is shifted to the right in the  presence of a substance that competes with glucose for bind¬  ing to the transporter; for example, 6-O-benzyl-D-galactose has  this effect. Competition is not a feature of nonmediated trans¬  port, since no transport protein is involved.   4. Inactivation. Reagents that chemically modify proteins and  hence may affect their functions may eliminate the rapid, satu-  ratable flux of glucose into the erythrocyte. The susceptibility of  the erythrocyte glucose transport system to protein-modifying  reagents is additional proof that it is a protein.   [Graph based on data from Stein, W.D., Movement of Molecules across   states in which the ligand binding sites are exposed, in turn, to opposite sides of  the membrane. Such a mechanism is analogous to the T —» R allosteric tran¬  sition of proteins such as hemoglobin (Section 7-IB). In fact, many of the fea¬  tures of ligand-binding proteins such as myoglobin and hemoglobin also apply  to transport proteins (Box 10-2).   Some transporters can transport more than one substance. For example,  the bacterial oxalate transporter transports oxalate into the cell and transports  formate out.   Some transport proteins move more than one substance at a time. Hence, it  is useful to categorize mediated transport according to the stoichiometry of  the transport process (Fig. 10-14; opposite ):   1. A uniport involves the movement of a single molecule at a time.  GLUT1 is a uniport system.   2. A symport simultaneously transports two different molecules in the  same direction.   3. An antiport simultaneously transports two different molecules in  opposite directions.   • What are the similarities and differences  among ionophores, porins, ion channels,  and passive-mediated transport proteins?  What determines the direction of solute  movement?   • Which transporters provide an open pas¬  sageway for the transmembrane move¬  ment of a solute? Which transporters  undergo a conformational change as part  of their transport mechanism?   • What structural features allow porins and  ion channels to discriminate among ions?   • How do transport proteins prevent the  transmembrane movement of water and  ions?   • How do aquaporins allow the passage of  H 2 0 but not H 3 0 + ?   • Use the terminology of allosteric proteins  to discuss the operation of proteins that  carry out uniport, symport, and antiport  transport processes.   FIG. 10-15 X-Ray structure of shark  (Na + -K + )-ATPase. The protein is drawn in  ribbon form viewed parallel to the plane of the  membrane (gray box) with the cytosol above. The  a subunit is colored in rainbow order from its  N-terminus (blue) to its C-terminus (red), the p  subunit is magenta, and the regulatory subunit,  called FXYD, is brown. Two bound K + ions in  the transmembrane (M domain) are drawn as  light blue spheres, and the MgF|“ ion, which  marks the protein’s ATPase active site between  the N and A domains, is shown with Mg pink  and F light green. [Based on an X-ray structure  by Chikashi Toyoshima, University of Tokyo,  Japan. PDBid 2ZXE]   • Pumps use the free energy of ATP to transport ions against their gradient.   • ABC transporters move amphipathic substances from one side of the membrane  to the other.   • Secondary active transporters use existing ion gradients to drive the unfavorable  transport of a second substance.   Passive-mediated transporters, including porins, ion channels, and proteins  such as GLUT 1, facilitate the transmembrane movement of substances accord¬  ing to the relative concentrations of the substance on either side of the mem¬  brane. For example, the glucose concentration in the blood plasma (~5 mM)  is generally higher than in cells, so GLUTl allows glucose to enter the  erythrocyte to be metabolized. Many substances, however, are available on one  side of a membrane in lower concentrations than are required on the other  side of the membrane. Such substances must be actively and selectively trans¬  ported across the membrane against their concentration gradients.   Active transport is an endergonic process that, in most cases, is coupled  to the hydrolysis of ATP. Several families of ATP-dependent transporters have  been identified:   1. P-type ATPases undergo phosphorylation as they transport cations  such as Na + , K + , and Ca 2+ across the membrane.   2. F-type ATPases are proton-transporting complexes located in mito¬  chondria and bacterial membranes. Instead of using the free energy of  ATP to pump protons against their gradient, these proteins usually  operate in reverse to synthesize ATP, as we will see in Section 18-3.   3. V-type ATPases resemble the F-type ATPases and occur in plant vac¬  uoles and acidic vesicles such as animal lysosomes.   5. ABC transporters are named for their ATP-binding cassette and trans¬  port a wide variety of substances, including ions, small metabolites,  and drug molecules.   In this section, we examine two P-type ATPases and an ABC transporter;  these proteins carry out primary active transport. In secondary active trans¬  port, the free energy of the electrochemical gradient generated by another  mechanism, such as an ion-pumping ATPase, is used to transport a molecule  against its concentration gradient.    A The (Na + -K + )-ATPase Transports Ions in Opposite Directions   One of the most thoroughly studied active transport systems is the  (Na + —K + )—ATPase in the plasma membranes of higher eukaryotes, which was  first characterized by Jens Skou. This transmembrane protein consists of two  types of subunits: The ~ 1000-residue a subunit contains the enzymes ATP  and ion binding sites, and the ~300-residue (3 subunit facilitates the correct  insertion of the a subunit into the plasma membrane. Chikashi Toyoshima  determined the X-ray structure of shark (Na + -K + )-ATPase in complex with  K + ions, an MgFf ion (a P z mimic), and a regulatory protein (Fig. 10-15).  The a subunit of this ~-T60-A-long protein consists of a transmembrane do¬  main (M) composed of 10 helices of varied lengths and, from top to bottom  in Fig. 10-15, three well-separated cytoplasmic domains: the nucleotide¬  binding domain (N), which binds ATP; the actuator domain (A), so named  because it participates in the transmission of major conformational changes  (see below); and the phosphorylation domain (P), which contains the proteins  phosphorylatable Asp residue.   The (Na + -K + )-ATPase is often called the (Na + -K + ) pump because it  pumps Na + out of and K + into the cell with the concomitant hydrolysis of  intracellular ATP. The overall stoichiometry of the reaction is   The (Na + —K + )—ATPase is an antiport that generates a charge separation  across the membrane, since three positive charges exit the cell for every two  that enter. This extrusion of Na + enables animal cells to control their water  content osmotically; without functioning (Net —K + )—A TPases to maintain a low  internal [Na + ], water would osmotically rush in to such an extent that animal  cells, which lack cell walls , would swell and burst. The electrochemical gradient  generated by the (Na + —K + )—ATPase is also responsible for the electrical ex¬  citability of nerve cells (Section 10-2C). In fact, all cells expend a large frac¬  tion of the ATP they produce (up to 70% in nerve cells) to maintain their  required cytosolic Na + and K + concentrations.   The key to the (Na + —K + )—ATPase is the phosphorylation of a specific  Asp residue of the transport protein. ATP phosphorylates the transporter only  in the presence of Na + , whereas the resulting aspartyl phosphate residue (at  right) is subject to hydrolysis only in the presence of K + . This suggests that  the (Na + -K + )-ATPase has two conformational states (called El and E2) with  different structures, different catalytic activities, and different ligand specifici¬  ties. The protein appears to operate in the following manner (Fig. 10-16):   1. El • ATP, which acquired its ATP inside the cell, binds three Na + ions  to yield the ternary complex El • ATP • 3Na + .    FIG. 1 0-1 6 Scheme for the active transport of Na + and K + by the (Na + -K + )-ATPase.   Draw simple shapes to represent the ATPase and its substrates and products at each step of the reaction cycle.    The cardiac glycosides are natural products that increase the  intensity of heart muscle contraction. Indeed, digitalis, an extract  of purple foxglove leaves, which contains a mixture of cardiac  glycosides including digitalin (see below), has been used to treat  congestive heart failure for centuries. The cardiac glycoside  ouabain (pronounced wabane), a product of the East African  ouabio tree, has been long used as an arrow poison.    [Na + ] stimulates the cardiac (Na + -Ca 2+ ) antiport system, which  pumps Na + out of and Ca 2+ into the cell, ultimately boosting the  [Ca 2+ ] in the sarcoplasmic reticulum. Thus, the release of Ca 2+ to  trigger muscle contraction (Section 7-2B) produces a larger than  normal increase in cytosolic [Ca 2+ ], thereby intensifying the force  of cardiac muscle contraction. Ouabain, which was once thought  to be produced only by plants, has recently been discovered to be    These two steroids, which are still among the most commonly  prescribed cardiac drugs, inhibit the (Na + -K + )-ATPase by binding  strongly to an externally exposed portion of the protein so as to  block Step 5 in Fig. 10-16. The resultant increase in intracellular    an animal hormone that is secreted by the adrenal cortex and  functions to regulate cellular [Na + ] and overall body salt and  water balance.    3. This “high-energy” intermediate relaxes to its “low-energy” conforma¬  tion, E2—P • 3Na + , and releases its bound Na + outside the cell; that  is, Na + is transported through the membrane.   4. E2—P binds two K + ions from outside the cell to form an E2—P • 2K +  complex. This corresponds to the structure shown in Fig. 10-15.   5. The phosphate group is hydrolyzed, yielding E2 • 2K + .   6 . E2 • 2K + changes conformation to El, binds ATP, and releases its two  K + ions inside the cell, thereby completing the transport cycle.   The enzyme appears to have only one set of cation binding sites (Fig. 10-15),  which apparently changes both its orientation and its specificity during the  course of the transport cycle.   Although each of the above reaction steps is individually reversible, the  cycle, as diagramed in Fig. 10-16, circulates only in the clockwise direction  under normal physiological conditions. This is because ATP hydrolysis and  ion transport are coupled vectorial (unidirectional) processes. The vectorial na¬  ture of the reaction cycle results from the alternation of some of the steps of  the exergonic ATP hydrolysis reaction (Step 2, Step 5, and ATP binding in  Step 6) with some of the steps of the endergonic ion transport process (Steps  3 + 4 and K + release in Step 6). Thus, neither reaction can go to completion  unless the other one also does. Study of the (Na + —K + )—ATPase has been greatly  facilitated by the use of glycosides that inhibit the transporter (Box 10-3).   B The Ca 2+ -ATPase Pumps Ca 2+ Out of the Cytosol   Transient increases in cytosolic [Ca 2+ ] trigger numerous cellular responses in¬  cluding muscle contraction (Section 7-2B), the release of neurotransmitters,    FIG. 10-17 Scheme for the active transport of Ca 2+ by the  Ca 2+ -ATPase. Here (in) refers to the cytosol and (out) refers to the  outside of the cell for plasma membrane Ca 2+ -ATPase or the lumen    of the endoplasmic reticulum (or sarcoplasmic reticulum) for the  Ca 2+ -ATPase of that membrane.    and glycogen breakdown (Section 16-3). Moreover, Ca 2+ is an important ac¬  tivator of oxidative metabolism (Section 18-4).   The [Ca 2+ ] in the cytosol (~0.1 (JiM) is four orders of magnitude less  than it is in the extracellular spaces [~1500 |JiM; intracellular Ca 2+ might  otherwise combine with phosphate to form Ca 3 (P 04 ) 2 , which has a maximum  solubility of only 65 |JiM]. This large concentration gradient is maintained by  the active transport of Ca 2+ across the plasma membrane and the endoplas¬  mic reticulum (the sarcoplasmic reticulum in muscle) by a Ca 2+ —ATPase. This  Ca 2+ pump actively pumps two Ca 2+ ions out of the cytosol at the expense  of ATP hydrolysis, while countertransporting two or three protons. The  mechanism of the Ca 2+ -ATPase (Fig. 10-17) resembles that of the  (Na + -K + )-ATPase (Fig. 10-16).   The superimposed X-ray structures of the Ca 2+ —ATPase from rabbit mus¬  cle sarcoplasmic reticulum in its El and E2 conformations, determined by  Chikashi Toyoshima, are shown in Fig. 10-18. Two Ca 2+ ions bind within a  bundle of 10 transmembrane helices. Three additional domains form a large    FIG. 1 0-1 8 X-Ray structures of the Ca 2+ -free and Ca 2+ -bound Ca 2+ -ATPase.   The Ca 2+ -free form, E2, is green with black helix numbers, and the Ca 2+ -bound form,  E1Ca 2+ , is violet with yellow helix numbers. These proteins, which are superimposed on  their transmembrane domains, are viewed from within the membrane with the cytosolic side  up. Ten transmembrane helices form the M (for membrane) domain, ATP binds to the N (for  nucleotide-binding) domain, the Asp residue that is phosphorylated during the reaction cycle  is located on the P (for phosphorylation) domain, and the A (for actuator) domain is so  named because it participates in the transmission of major conformational changes. Dashed  lines highlight the orientations of a helix in the N domain in the two conformations and the  horizontal lines delineate the membrane. [Courtesy of Chikashi Toyoshima, University of  Tokyo, Japan. PDBids 1EUL and 1WI0J   Chapter 10 Membrane Transport structure on the cytoplasmic side of the membrane. The differences between   the Ca 2+ -bound (El) and the Ca 2+ -free (E2) structures indicate that the  transporter undergoes extensive rearrangements, particularly in the positions  of the cytoplasmic domains but also in the Ca 2+ -transporting membrane do¬  main, during the reaction cycle. These changes apparently mediate communi¬  cation between the Ca 2+ -binding sites and the ~80-A-distant site where  bound ATP is hydrolyzed.   The inability of anticancer drugs to kill cancer cells is frequently traced to the  overexpression of a membrane protein known as P-glycoprotein. This mem¬  ber of the ABC family of transporters pumps a variety of amphiphilic  substances—including many drugs—out of the cell, so that it is also called a  multidrug resistance (MDR) transporter. Similar proteins in bacteria con¬  tribute to their antibiotic resistance.   The ABC transporters, which pump ions, sugars, amino acids, and other  polar and nonpolar substances, are built from four modules: two highly con¬  served cytoplasmic nucleotide-binding domains, and two transmembrane do¬  mains that typically contain six transmembrane helices each. In bacteria, the  four domains are contained on two or four separate polypeptides, and in eu¬  karyotes, a single polypeptide includes all four domains. Bacterial ABC trans¬  porters mediate the uptake as well as the efflux of a variety of compounds,  whereas their eukaryotic counterparts apparently operate only as exporters that  transport material out of the cell or into intracellular compartments such as  the endoplasmic reticulum.   The X-ray structure of mouse P-glycoprotein, determined by Geoffrey  Chang, shows a dimeric protein with pseudo-2-fold symmetry (Fig. 10-19).  Two bundles of six transmembrane a helices span the membrane and extend  into the cytoplasm, defining a large internal cavity that is open to both the  cytoplasm and the inner leaflet of the membrane. The two ATP-binding do¬  mains in the cytoplasm are separated by ~30 A. Conformational changes re¬  sulting from ATP binding and hydrolysis in this part of the protein appear to  be transmitted to the transmembrane domains in a way that allows the two  subunits of the transporter to move in concert.   A lipid-soluble molecule, such as a drug molecule, apparently enters  P-glycoprotein through a portal in the membrane-spanning domain and binds  in a pocket defined mostly by hydrophobic and aromatic residues. ATP bind¬  ing triggers a dramatic conformational change that includes dimerization of  the two cytoplasmic nucleotide-binding domains and a shift to an outward¬  facing conformation (Fig. 10-20). This structural change, which has been ob¬  served in the X-ray structures of similar transporters in their nucleotide-bound  state, closes off the original binding cavity and exposes the bound drug mol¬  ecule to the outer leaflet or the extracellular space. ATP hydrolysis presumably  restores the transporter to its inward-facing conformation, ready to bind an¬  other drug molecule. Because the transporter can bind a molecule from inside  the cell or from the inner leaflet and release it outside the cell or into the  outer leaflet, the transporter operates as a flippase (Section 9-4C) for lipid-  soluble substances, including membrane lipids.    FIG. 10-19 X-Ray structure of mouse P-glycoprotein. The peptide chain is drawn in  ribbon form colored in rainbow order from its N-terminus (blue) to its C-terminus (red). The  view is parallel to the membrane (delineated by the horizontal lines) with the cytosol below.  The two cytoplasmic nucleotide-binding domains (NBD1 and NBD2) are indicated. This  “inward-facing” conformation of P-glycoprotein contains no bound ligands or nucleotides.  [Based on an X-ray structure by Geoffrey Chang, The Scripps Research Institute, La Jolla.  PDBid 3G5U.]   FIG. 1 0-20 Model for P-glycoprotein  function. The gray shapes represent the surface  of the transporter in its inward-facing (left) and  outward-facing (right) conformations. A ligand  (magenta) may originate from outside the cell  but partitions into the inner leaflet of the plasma  membrane (delineated by the horizontal lines)  before entering the transporter’s binding pocket,  whose residues are represented by cyan  spheres. The binding of ATP (yellow) triggers  extensive conformational changes that bring the  nucleotide-binding domains together and open  the ligand-binding site to the extracellular space.  [Courtesy of Geoffrey Chang, The Scripps  Research Institute, La Jolla.]    T CFTR Is an ABC Transporter. Only one of the thousands of known ABC  transporters (48 in humans) functions as an ion channel rather than a  pump: the cystic fibrosis transmembrane conductance regulator (CFTR).  This 1480-residue protein, which is defective in individuals with the inherited  disease cystic fibrosis (Box 3-1), allows Cl ions to flow out of the cell, follow¬  ing their concentration gradient. ATP binding to CFTRs two nucleotide-bind¬  ing domains appears to open the Cl channel, and the hydrolysis of one ATP  closes it (the other ATP remains intact). However, the CFTR channel can open  only if its regulatory domain (which is unique among ABC transporters) has been  phosphorylated, thus regulating the flow of ions across the membrane. CFTR  does not exhibit high specificity for Cl ions, suggesting that the channel lacks  a selectivity filter analogous to that in the KcsA K channel (Section 10-2C).   The maintenance of electrical neutrality requires that the Cl ions trans¬  ported by the CFTR be accompanied by positively charged ions, mainly Na + .  The transported ions are osmotically accompanied by water, thus maintaining  the proper level of fluidity in secretions of the airways, intestinal tract, and  the ducts of the pancreas, testes, and sweat glands. Although over 1000 mu¬  tations of the CFTR have been described, in about 70% of cystic fibrosis cases,  Phe 508 of the CFTR has been deleted. Even though this mutant CFTR is  functional, it folds much more slowly than the wild-type protein and hence  is degraded before it can be installed in the plasma membrane.   Homozygotes for defective or absent CFTRs have problems in many of  the organs mentioned above but especially in their lungs (heterozygotes are  asymptomatic). This is because the reduced Cl export results in thickened  mucus that the lungs cannot easily clear. Since the flow of mucus is the ma¬  jor way in which the lungs eliminate foreign particles such as bacteria, indi¬  viduals with cystic fibrosis suffer from chronic lung infections leading to severe  progressive lung damage and early death.   Systems such as the (Na + -K + )-ATPase generate electrochemical gradients  across membranes. The free energy stored in an electrochemical gradient  (Eq. 10-3) can be harnessed to power various endergonic physiological  processes. For example, cells of the intestinal epithelium take up dietary glucose   The brushlike villi lining the small intestine greatly increase its surface  area (a), thereby facilitating the absorption of nutrients. The brush  border cells from which the villi are formed (b) concentrate glucose  from the intestinal lumen in symport with Na + (c), a process that is    driven by the (Na + -K + )-ATPase, which is located on the capillary  side of the cell and functions to maintain a low internal [Na + ]. The  glucose is exported to the bloodstream via a separate passive-  mediated uniport system similar to GLUT1.    by Na + -dependent symport (Fig. 10-21). The immediate energy source for this  “uphill” transport process is the Na + gradient. This process is an example of  secondary active transport because the Na + gradient in these cells is maintained  by the (Na + -K^)—ATPase. The Na + -glucose transport system concentrates  glucose inside the cell. Glucose is then transported into the capillaries through  a passive-mediated glucose uniport (which resembles GLUTl; Fig. 10-13).  Thus, since glucose enhances Na + resorption, which in turn enhances water  resorption, glucose (possibly as sucrose), in addition to salt and water,  should be fed to individuals suffering from severe salt and water losses due to  diarrhea (drinking only water or a salt solution is ineffective since they are rap¬  idly excreted from the gastrointestinal tract). It is estimated that the introduc¬  tion of this simple and inexpensive treatment, called oral rehydration therapy,  has decreased the annual number of human deaths from severe diarrhea,  mostly in children in less-developed countries, from 4.6 to 1.6 million.   Lactose Permease Requires a Proton Gradient. Gram-negative bacteria  such as E. coli contain several active transport systems for concentrating sug¬  ars. One extensively studied system, lactose permease (also known as galac-  toside permease), utilizes the proton gradient across the bacterial cell membrane  to cotransport H + and lactose. The proton gradient is metabolically generated  through oxidative metabolism in a manner similar to that in mitochondria  (Section 18-2). The electrochemical potential gradient created by both these  systems is used mainly to drive the synthesis of ATP.   Lactose permease is a 417-residue monomer that consists largely of 12  transmembrane helices with its N- and C-termini in the cytoplasm. A large  number of transport proteins, including the passive transporter GLUTl  (Section 10-2E), belong to the same family of distantly related proteins and  have similar structures and transport mechanisms. Like (Na + -K + )-ATPase,  lactose permease has two major conformational states (Fig. 10-22):   1. E-1, which has a low-affinity lactose-binding site facing the interior of  the cell.   2. E-2, which has a high-affinity lactose-binding site facing the exterior  of the cell.    FIG. 1 0-22 Scheme for the cotransport of H + and lactose by  lactose permease in E. coli. H + binds first to E-2 outside the cell,  followed by lactose. They are sequentially released from E-1 inside the cell.  E-2 must bind to both lactose and H + in order to change conformation    to E-1, thereby cotransporting these substances into the cell. E-1  changes conformation to E-2 when neither lactose nor H + is bound,  thus completing the transport cycle.    Ronald Kaback established that E-1 and E-2 can interconvert only when their  H + - and lactose-binding sites are either both filled or both empty. This pre¬  vents dissipation of the H + gradient without cotransport of lactose into the  cell. It also prevents transport of lactose out of the cell since this would re¬  quire co transport of H + against its concentration gradient.   The X-ray structure of lactose permease in complex with a tight-binding  lactose analog, determined by Kaback and So Iwata, reveals that this protein  consists of two structurally similar and twofold symmetrically positioned  domains containing six transmembrane helices each (Fig. 10-23^). A large   FIG. 1 0-23 X-Ray structure of lactose permease from E. coli.   (a) Ribbon diagram as viewed from the membrane with the cytoplasmic  side up. The protein’s 12 transmembrane helices are colored in rainbow  order from the N-terminus ( purple ) to the C-terminus (pink). The  bound lactose analog is represented by black spheres, (b) Surface    model viewed as in Part a but with the two helices closest to the  viewer in Part a removed to reveal the lactose-binding cavity. The  surface is colored according to its electrostatic potential with positively  charged areas blue, negatively charged areas red, and neutral areas  white. [Courtesy of H. Ronald Kaback, UCLA. PDBid 1 PV7.]   • Explain why the (Na + -K + )-ATPase and  the Ca 2+ -ATPase carry out transport in  one direction only.   • Explain how an ABC transporter would  operate as a flippase.   • What is the ultimate source of free energy  for secondary active transport?    internal hydrophilic cavity is open to the cytoplasmic side of the membrane  (Fig. 10-23^) so that the structure represents the E-l state of the protein.  The lactose analog is bound in the cavity at a position that is approximately  equidistant from both sides of the membrane, consistent with the model that  the lactose-binding site is alternately accessible from each side of the mem¬  brane (e.g., Fig. 10-13). Arg, His, and Glu residues that mutational studies  have implicated in proton translocation are located in the vicinity of the lac¬  tose-binding site.   • The mediated and nonmediated transport of a substance across a  membrane is driven by its chemical potential difference.   • Ionophores facilitate ion diffusion by binding an ion, diffusing  through the membrane, and then releasing the ion; or by form¬  ing a channel.   • Porins form (3 barrel structures about a central channel that is  selective for anions, cations, or certain small molecules.   • Ion channels mediate changes in membrane potential by allowing  the rapid and spontaneous transport of ions. Ion channels are  highly solute-selective and open and close (gate) in response to  various stimuli. Nerve impulses involve ion channels.   • Aquaporins contain channels that allow the rapid transmembrane  diffusion of water but not protons.    • Transport proteins such as GLUTl alternate between two confor¬  mational states that expose the ligand-binding site to opposite  sides of the membrane.   • Active transport, in most cases, is driven by ATP hydrolysis. In  the (Na + -K + )-ATPase and Ca 2+ -ATPase, ATP hydrolysis and  ion transport are coupled and vectorial.   • ABC transporters use ATP hydrolysis to trigger conformational  changes that move substances, including amphiphilic molecules,  from one side of the membrane to the other.   • In secondary active transport, an ion gradient maintained by an  ATPase or other free energy-capturing cellular process drives the  transport of another substance. For example, the transport of lac¬  tose into a cell by lactose permease is driven by the cotransport  of H + , whose gradient is maintained by oxidative metabolism.    1. Calculate the free energy change for glucose entry into cells when  the extracellular concentration is 5 mM and the intracellular con¬  centration is 3 mM.   2. (a) Calculate the chemical potential difference when intracellular  [Na + ] = 10 mM and extracellular [Na + ] = 130 mM at 37°C.  (b) What would the electrochemical potential be if the membrane  potential were —60 mV (inside negative)?   3. For the problem in Sample Calculation 10-1, calculate A Cat 37°C  when the membrane potential is (a) — 30 mV (cytosol negative)    and (b) +150 mV. In which case is Ca 2+ movement in the indi¬  cated direction thermodynamically favorable?   4. Calculate the free energy required to move 1 mol of K + ions  from the outside of the cell (where [K + ] = 4 mM) to the inside  (where [K + ] = 140 mM) when the membrane potential is —70  mV and the temperature is 37°C.   5. Indicate whether the following compounds are likely to cross a  membrane by nonmediated or mediated transport: (a) ethanol,  (b) glycine, (c) cholesterol, (d) ATP.    Rank the rate of transmembrane diffusion of the following com¬  pounds:   What happens to K + transport by valinomycin when the mem¬  brane is cooled below its transition temperature?   How long would it take 100 molecules of valinomycin to trans¬  port enough K + to change the concentration inside an erythro¬  cyte of volume 100 fxm 3 by 10 mM? (Assume that the  valinomycin does not also transport any K + out of the cell, which  it really does, and that the valinomycin molecules outside the cell  are always saturated with K + .)   The smallest (Tbarrel protein contains only eight (3 strands.  Explain why porins, which are also (3 barrels, usually contain at  least 16 or 18 strands.   Which amino acids would you expect to be particularly abun¬  dant at the entrance of a porin that is specific for phosphate ions?   The diameter of the KcsA K + channel is ~ 6 A. Why can’t H 2 0  (diameter 2.73 A) pass through this channel?   In addition to neurons, muscle cells undergo depolarization, al¬  though smaller and slower than in the neuron, as a result of the  activity of the acetylcholine receptor.   (a) The acetylcholine receptor is also a gated ion channel. What  triggers the gate to open?   (b) The acetylcholine receptor/ion channel is specific for Na +  ions. Would Na + ions flow in or out? Why?   (c) How would the Na + flow through the ion channel change  the membrane potential?   ch 3 - (CH 2 ) 15 - o -p - ch 2 - ch 2 -n + (ch 3 ) 3   (c) In what part of the cell would the drug tend to accumulate?  Explain.   (d) Miltefosine binds to a protein that also binds some sphin-  golipids and some glycerophospholipids. What feature com¬  mon to all these compounds is recognized by the protein?  The protein does not bind triacylglycerols.   In eukaryotes, ribosomes (approximate mass 4 X 10 6 D) are  assembled inside the nucleus, which is enclosed by a double  membrane. Protein synthesis occurs in the cytosol, (a) Could a  protein similar to a porin or the glucose transporter be responsi¬  ble for transporting ribosomes into the cytoplasm? Explain,  (b) Would free energy be required to move a ribosome from the  nucleus to the cytoplasm? Why or why not?    The rate of movement (flux) of a substance X into cells was  measured at different concentrations of X to construct the fol¬  lowing graph.   (a) Does this information suggest that the movement of X into  the cells is mediated by a protein transporter? Explain.   (b) What additional experiment could you perform to verify that  a transport protein is or is not involved?   16. Endothelial cells and pericytes in the retina of the eye have dif¬  ferent mechanisms for glucose uptake. The figure shows the rate  of glucose uptake for each type of cell in the presence of increas¬  ing amounts of sodium. What do these results reveal about the  glucose transporter in each cell type?   17. Explain why Na + and K + ions usually move more slowly  through pumps than through channels.   18. Why would overexpression of an MDR transporter in a cancer  cell make the cancer more difficult to treat?   19. Cells in the wall of the mammalian stomach secrete HCl at a con¬  centration of 0.15 M. The secreted protons, which are derived  from the intracellular hydration of C0 2 by carbonic anhydrase,  are pumped out by an (H + -K + )-ATPase antiport. A K + -Cl co¬  transporter is also required to complete the overall transport  process, (a) Calculate the pH of the secreted HCl. How does this  compare to the cytosolic pH (7.4)? (b) Write the reaction catalyzed  by carbonic anhydrase. (c) Draw a diagram to show how the ac¬  tion of both transport proteins results in the secretion of HCl.   20. Kidney cells contain a channel that allows intracellular ammonia  to exit the cells, (a) Why did researchers originally believe that cells  had no need for such a channel? (b) What is the free energy source  for ammonia transport via the channel? (c) The same kidney cells  also contain a proton pump that expels H from the cells. What  is the free energy source for this pump, and how does its action  prevent ammonia from moving back into the kidney cells?   21. If the ATP supply in the cell shown in Fig. 10-21 c suddenly van¬  ished, would the intracellular glucose concentration increase, de¬  crease, or remain the same?   22. The bacterial Na + -H + antiporter is a secondary active transport  protein that excretes excess Na + from the cell. Is the extracellu¬  lar pH higher or lower than the intracellular pH?   Case Study 14 Shavings from the Carpenter’s Bench:  The Biological Role of the Insulin C-peptide  Focus concept: Recent experiments indicate that the insulin C-peptide,  which is removed on conversion of proinsulin to insulin, may have  biological activity in its own right.   Case Study 17 A Possible Mechanism for Blindness  Associated with Diabetes: Na + -Dependent  Glucose Uptake by Retinal Cells   Focus concept: Glucose transport into cells can influence collagen  synthesis, which causes the basement membrane thickening asso¬  ciated with diabetic retinopathy.    The M2 protein encoded by  the influenza virus is essential for infection. What ion is transported  by the M2 protein, and what role does ion transport play in the flu  virus life cycle? What drugs target the M2 protein? Why is it nec¬  essary to develop new drugs to block M2 function?   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Jiang, Y., Lee, A., Chen, J., Ruta, V., Cadene, M., Chait, B.T., and Mac¬  Kinnon, R., X-Ray structure of a voltage-dependent K + channel,  Nature 423, 33-41 (2003).   Long, S.B., Campbell, E.B., and MacKinnon, R., Voltage sensor of  Kvl.2: Structural basis of electromechanical coupling, Science 30,  903-908 (2005).   Roux, B., Ion conduction and selectivity in K + channels, Annu. Rev. Bio-  phys. Biomol. Struct. 34, 153-171 (2005).   Zhou, Y., Morais-Cabral, J.H., Kaufman, A., and MacKinnon, R., Chem¬  istry of ion coordination and hydration revealed by a K +  channel-Fab complex at 2.0 A resolution, Nature 414, 43-48 (2001).   Fu, D. and Lu, M., The structural basis of water permeation and proton  exclusion in aquaporins, Mol. Membrane Biol. 24, 366-374 (2007).  [A review.]   King, L.S., Kozono, D., and Agre, P., From structure to disease: The  evolving tale of aquaporin biology, Nature Rev. Mol. Cell Biol. 5,  687-698 (2004).   Sui, H., Han, B.-G., Lee, J.K., and Jap, B.K., Structural basis of water-  specific transport through the AQP1 water channel, Nature 414,  872-878 (2001).   Abramson, J., Smirnova, I., Kasho, V., Verner, G., Kaback, H.R., and  Iwata, S., Structure and mechanism of the lactose permease of  Escherichia coli, Science 301, 610-615 (2003).   Aller, S.G., Yu, J., Ward, A., Weng, Y, Chittaboina, S., Zhuo, R., Har¬  rell, P.M., Trinh, Y.T., Zhang, Q., Urbatsch, I.L., and Chang, G.,  Structure of P-glycoprotein reveals a molecular basis for poly-specific  drug binding, Science 323, 1718-1722 (2009).   Bezanilla, F., How membrane proteins sense voltage, Nature Rev. Mol.  Cell Biol. 9, 323-332 (2008).   Gouaux, E. and MacKinnon, R., Principles of selective ion transport in  channels and pumps, Science 310, 1461-1465 (2005). [Compares  several transport proteins of known structure and discusses the se¬  lectivity of Na + , K + , Ca 2+ , and Cl transport.]   Hille, B., Ionic Channels of Excitable Membranes (3rd ed.), Sinauer Asso¬  ciates (2001).   Jones, P.M., O’Mara, M.L., and George, A.M., ABC transporters: a rid¬  dle wrapped in a mystery inside an enigma, Trends Biochem. Sci. 34,  520-531 (2009).   Kaplan, J.H., Biochemistry of Na,K-ATPase, Annu. Rev. Biochem. 71,  511-535 (2002).   Krishnamurthy, H., Piscitelli, C.L., and Gouaux, E., Unlocking the mo¬  lecular secrets of sodium-coupled transporters, Nature 459, 347-354  (2009).   Long, S.B., Tao, X., Campbell, E.B., and MacKinnon, R., Atomic struc¬  ture of a voltage-dependent K channel in a lipid membrane-like  environment, Nature 450, 376-382 (2007).   Olesen, C., Picard, M., Winther, A.-M.L., Gyrup, C., Morth, J.P., Oxvig,  C., Moller, J.V., and Nissen, P., The structural basis of calcium trans¬  port by the calcium pump, Nature 450, 1036-1042 (2007).   Rees, D.C., Johnson, E., and Lewinson, O., ABC transporters: the power  to change, Nature Rev. Mol. Cell Biol. 10, 218-227 (2009).   Shinoda, T., Ogawa, H., Cornelius, F., and Toyoshima, C., Crystal struc¬  ture of the sodium-potassium pump at 2.4 A resolution, Nature  459, 446-450 (2009). [The shark enzyme.]   Toyoshima, C. and Inesi, G., Structural basis of ion pumping by Ca 2+ -  ATPase of the sarcoplasmic reticulum, Annu. Rev. Biochem. 73,  269-292 (2004).   Wright, E.M., Hirayama, B.A., and Loo, D.F., Active sugar transport in  health and disease, J. Intern. Med. 261, 32-43 (2007).   Substances such as crude oil, a mixture of hydrocarbons, would persist unchanged for  many years because they are relatively unreactive in the aqueous environment.  Fortunately, microbial enzymes can catalyze the breakdown of these substances,  accelerating their decay by many orders of magnitude. [Michael B. Watkins/U.S.  Navy/Landov LLCJ   Living systems are shaped by an enormous variety of biochemical reactions,  nearly all of which are mediated by a series of remarkable biological catalysts  known as enzymes. Enzymology, the study of enzymes, has its roots in the  early days of biochemistry; both disciplines evolved together from nineteenth  century investigations of fermentation and digestion. Initially, the inability to  reproduce most biochemical reactions in the laboratory led Louis Pasteur and  others to assume that living systems were endowed with a “vital force” that  permitted them to evade the laws of nature governing inanimate matter. Some  investigators, however, notably Justus von Liebig, argued that biological  processes were caused by the action of chemical substances that were then  known as “ferments.” Indeed, the name “enzyme” (Greek: en, in + zyme, yeast)  was coined in 1878 in an effort to emphasize that there is something in yeast,  as opposed to the yeast itself, that catalyzes the reactions of fermentation.  Eventually, Eduard Buchner showed that a cell-free yeast extract could in fact  carry out the synthesis of ethanol from glucose (alcoholic fermentation;  Section 15-3B):   C 6 H 12 0 6 —> 2 CH 3 CH 2 OH + 2C0 2   A Enzymes Are Classified by the Type of Reaction  They Catalyze   B X-Ray Structures Provide Information about  Catalysis, Substrate Specificity, and Evolution   The chemical composition of enzymes was not firmly established until  1926, when James Sumner crystallized jack bean urease, which catalyzes the  hydrolysis of urea to NH 3 and C0 2 , and demonstrated that the crystals con¬  sist of protein. Enzymological experience since then has amply demonstrated  that nearly all biochemical reactions are catalyzed by proteins, although there  are some notable exceptions.   Certain species of RNA molecules known as ribozymes also have enzy¬  matic activity. These include ribosomal RNA, which catalyzes the formation  of peptide bonds between amino acids during protein synthesis. In fact, lab¬  oratory experiments have produced ribozymes that can catalyze reactions sim¬  ilar to those required for replicating DNA, transcribing it to RNA, and  attaching amino acids to transfer RNA. These findings are consistent with a  precellular world in which RNA molecules enjoyed a more exalted position as  the catalytic workhorses of biochemistry. The present-day RNA catalysts are  presumably vestiges of this earlier “RNA world.” Proteins have largely eclipsed   Chapter 11 Enzymatic Catalysis RNA as cellular catalysts, probably because of the greater chemical versatility   of proteins. Whereas nucleic acids are polymers of four types of chemically  similar monomeric units, proteins have at their disposal 20 types of amino  acids with a greater variety of functional groups.   This chapter is concerned with one of the central questions of biochem¬  istry: How do enzymes work? We will see that enzymes increase the rates of  chemical reactions by lowering the free energy barrier that separates the reac¬  tants and products. Enzymes accomplish this feat through various mechanisms  that depend on the arrangement of functional groups in the enzymes active  site, the region of the enzyme where catalysis occurs. In this chapter, we de¬  scribe these mechanisms, along with examples that illustrate how enzymes  combine several mechanisms to catalyze biological reactions. The following  chapter includes a discussion of enzyme kinetics, the study of the rates at  which such reactions occur.    Enzymes differ from ordinary chemical catalysts in reaction rate, reaction condi¬  tions, reaction specificity, and control.   The unique physical and chemical properties of the active site limit an enzyme’s  activity to specific substrates and reactions.    Biochemical research since Pasteurs era has shown that, although enzymes are  subject to the same laws of nature that govern the behavior of other substances,  enzymes differ from ordinary chemical catalysts in several important respects:   1. Higher reaction rates . The rates of enzymatically catalyzed reactions  are typically 10 6 to 10 12 times greater than those of the corresponding  uncatalyzed reactions (Table 11-1) and are at least several orders of  magnitude greater than those of the corresponding chemically catalyzed  reactions.   2. Milder reaction conditions . Enzymatically catalyzed reactions occur  under relatively mild conditions: temperatures below 100°C, atmos¬  pheric pressure, and nearly neutral pH. In contrast, efficient chemical  catalysis often requires elevated temperatures and pressures as well as  extremes of pH.   3. Greater reaction specificity . Enzymes have a vastly greater degree of  specificity with respect to the identities of both their substrates (reac¬  tants) and their products than do chemical catalysts; that is, enzymatic  reactions rarely have side products.    4. Capacity for regulation. The catalytic activities of many enzymes vary  in response to the concentrations of substances other than their sub¬  strates. The mechanisms of these regulatory processes include allosteric  control, covalent modification of enzymes, and variation of the  amounts of enzymes synthesized.    A Enzymes Are Classified by the Type of Reaction They Catalyze   Before delving further into the specific properties of enzymes, a word on nomen¬  clature is in order. Enzymes are commonly named by appending the suffix -ase  to the name of the enzymes substrate or to a phrase describing the enzyme’s cat¬  alytic action. Thus, urease catalyzes the hydrolysis of urea, and alcohol dehy¬  drogenase catalyzes the oxidation of primary and secondary alcohols to their  corresponding aldehydes and ketones by removing hydrogen. Since there were  at first no systematic rules for naming enzymes, this practice occasionally re¬  sulted in two different names being used for the same enzyme or, conversely, in  the same name being used for two different enzymes. Moreover, many enzymes,  such as catalase (which mediates the dismutation of H 2 0 2 to H 2 0 and 0 2 ),  were given names that provide no clue to their function. In an effort to elimi¬  nate this confusion and to provide rules for rationally naming the rapidly grow¬  ing number of newly discovered enzymes, the following scheme for the  systematic functional classification and nomenclature of enzymes was adopted  by the International Union of Biochemistry and Molecular Biology (IUBMB).   Enzymes are classified and named according to the nature of the chemical  reactions they catalyze. There are six major classes of enzymatic reactions (Table  11 - 2 ), as well as subclasses and sub-subclasses. Each enzyme is assigned two  names and a four-part classification number. Its accepted or recommended  name is convenient for everyday use and is often an enzymes previously used  name. Its systematic name is used when ambiguity must be minimized; it is  the name of its substrate(s) followed by a word ending in -ase specifying the  type of reaction the enzyme catalyzes according to its major group classification.  For example, the enzyme whose alternative name is aconitase (Section 17-3B)  has the systematic name aconitate hydratase and the classification number EC  4.2.1.3 (“EC” stands for Enzyme Commission, and the numbers represent the  class, subclass, sub-subclass, and an arbitrarily assigned serial number in the sub¬  subclass). For our purposes, the recommended name of an enzyme is usually ad¬  equate. However, EC classification numbers are increasingly used in various  Internet-accessible databases (Section 14-4C). Systematic names and EC classi¬  fication numbers can be obtained via the Internet (http://expasy.org/enzyme/).   The noncovalent forces through which substrates and other molecules bind to  enzymes are similar in character to the forces that dictate the conformations  of the proteins themselves (Section 6-4A). Both involve van der Waals, elec¬  trostatic, hydrogen bonding, and hydrophobic interactions. In general, a  substrate-binding site consists of an indentation or cleft on the surface of an  enzyme molecule that is complementary in shape to the substrate (geometric  complementarity). Moreover, the amino acid residues that form the binding  site are arranged to specifically attract the substrate (electronic complemen¬  tarity, Fig. 11-1). Molecules that differ in shape or functional group distribu¬  tion from the substrate cannot productively bind to the enzyme. X-Ray studies  indicate that the substrate-binding sites of most enzymes are largely preformed  but undergo some conformational change on substrate binding (a phenome¬  non called induced fit). The complementarity between enzymes and their  substrates is the basis of the “lock-and-key” model of enzyme function first  proposed by Emil Fischer in 1894. As we will see, such specific binding is  necessary but not sufficient for efficient catalysis.   The geometric and the electronic complementarity  between the enzyme and substrate depend on  noncovalent forces. Hydrophobic groups are  represented by an h in a brown circle, and  dashed lines represent hydrogen bonds.    FIG. 11-2 Stereospecificity in substrate  binding. The specific binding of a prochiral  molecule, such as citrate, in an enzyme active  site allows the enzyme to differentiate between  prochiral groups.    Enzymes Are Stereospecific. Enzymes are highly specific both in binding  chiral substrates and in catalyzing their reactions. This stereospecificity arises  because enzymes, by virtue of their inherent chirality (proteins consist of only  L-amino acids), form asymmetric active sites. For example, the enzyme aconi-  tase catalyzes the interconversion of citrate and isocitrate in the citric acid  cycle (Section 17-3B):    Citrate is a prochiral molecule; that is, it can become chiral through the sub¬  stitution of one of its two carboxymethyl (—CH 2 COO ) groups (chirality  is discussed in Section 4-2). These groups are chemically equivalent but oc¬  cupy different positions relative to the OH and COO groups (likewise, your  body is bilaterally symmetric but has distinguishable right and left sides).  Aconitase can therefore distinguish between them because citrate interacts  asymmetrically with the surface of the enzyme by making a three-point at¬  tachment (Fig. 11 -2). Because there is only one productive way for citrate to  bind to the enzyme, only one of its —CH 2 COO groups reacts to form iso¬  citrate. The stereospecificity of aconitase is by no means unusual. As we con¬  sider biochemical reactions, we will find that nearly all enzymes that participate  in chiral reactions are absolutely stereospecific.    Enzymes Vary in Geometric Specificity. The stereospecificity of enzymes is  not particularly surprising in light of the complementarity of an enzymes  binding site for its substrate. A substance of the wrong chirality will not fit  productively into an enzymatic binding site for much the same reason that  you cannot fit your right hand into your left glove. In addition to their stere¬  ospecificity, however, most enzymes are quite selective about the identities of  the chemical groups on their substrates. Indeed, such geometric specificity is  a more stringent requirement than is stereospecificity.   Enzymes vary considerably in their degree of geometric specificity. A few  enzymes are absolutely specific for only one compound. Most enzymes, how¬  ever, catalyze the reactions of a small range of related compounds although  with different efficiencies. For example, alcohol dehydrogenase catalyzes the  oxidation of ethanol (CH 3 CH 2 OH) to acetaldehyde (CH 3 CHO) faster than  it oxidizes methanol (CH 3 OH) to formaldehyde (H 2 CO) or isopropanol  [(CH 3 ) 2 CHOH] to acetone [(CH 3 ) 2 CO], even though methanol and iso¬  propanol differ from ethanol by only the deletion or addition of a CH 2 group.   Some enzymes, particularly digestive enzymes, are so permissive in their  ranges of acceptable substrates that their geometric specificities are more accu¬  rately described as preferences. Some enzymes are not even very specific in the  type of reaction they catalyze. For example, chymotrypsin, in addition to its abil¬  ity to mediate peptide bond hydrolysis, also catalyzes ester bond hydrolysis.   This property makes it convenient to measure chymotrypsin activity using Section 1 General Properties of Enzymes   small synthetic esters as substrates. Such permissiveness is much more the ex¬  ception than the rule. Indeed, most intracellular enzymes function in vivo to  catalyze a particular reaction on a particular substrate.    The functional groups of proteins, as we will see, can facilely participate in  acid—base reactions, form certain types of transient covalent bonds, and take part  in charge—charge interactions. They are, however, less suitable for catalyzing oxi¬  dation-reduction reactions and many types of group-transfer processes. Although  enzymes catalyze such reactions, they can do so only in association with small  cofactors, which essentially act as the enzymes “chemical teeth” (Fig. 11-3).   Cofactors may be metal ions, such as Cu 2+ , Fe 3+ , or Zn 2+ . The essen¬  tial nature of these cofactors explains why organisms require trace amounts of  certain elements in their diets. It also explains, in part, the toxic effects of cer¬  tain heavy metals. For example, Cd 2+ and Hg 2+ can replace Zn 2+ (all are in  the same group of the periodic table) in the active sites of certain enzymes,  including RNA polymerase, and thereby render these enzymes inactive.   Cofactors may also be organic molecules known as coenzymes. Some co¬  factors are only transiently associated with a given enzyme molecule, so that  they function as cosubstrates. Nicotinamide adenine dinucleotide (NAD + )  and nicotinamide adenine dinucleotide phosphate (NADP + ) are examples  of cosubstrates (Fig. 11 -4). For instance, NAD + is an obligatory oxidizing  agent in the alcohol dehydrogenase (ADH) reaction:   CH 3 CH 2 OH + NAD + s. CH 3 CH + NADH + H +   The product NADH dissociates from the enzyme for eventual reoxidation to  NAD + in an independent enzymatic reaction.    Nicotinamide adenine dinucleotide (NAD + )  Nicotinamide adenine dinucleotide phosphate (NADP +    FIG. 11-4 The structures and reaction of nicotinamide  adenine dinucleotide (NAD + ) and nicotinamide adenine  dinucleotide phosphate (NADP + ). Their reduced forms are  NADH and NADPH. These substances, which are collectively  referred to as the nicotinamide coenzymes or pyridine  nucleotides (nicotinamide is a pyridine derivative), function as  intracellular electron carriers. Reduction (addition of electrons)  formally involves the transfer of two hydrogen atoms (H*), or a  hydride ion and a proton (H:“ + H + ). Note that only the  nicotinamide ring is changed in the reaction.   • What is the relationship between cofactors,  coenzymes, cosubstrates, and prosthetic  groups?    Other cofactors, known as prosthetic groups, are permanently associated  with their protein, often by covalent bonds. For example, a heme prosthetic  group (Fig. 7-2) is tightly bound to proteins known as cytochromes (Fig.  6-32 and Box 18-1) through extensive hydrophobic and hydrogen-bonding in¬  teractions together with covalent bonds between the heme and specific protein  side chains.   A catalytically active enzyme-cofactor complex is called a holoenzyme.  The enzymatically inactive protein resulting from the removal of a holoen-  zyme’s cofactor is referred to as an apoenzyme; that is,   apoenzyme ( inactive ) + cofactor s holoenzyme ( active )    Coenzymes Must Be Regenerated. Coenzymes are chemically changed by  the enzymatic reactions in which they participate. In order to complete the cat¬  alytic cycle , the coenzyme must return to its original state. For a transiently bound  coenzyme (cosubstrate), the regeneration reaction may be catalyzed by a dif¬  ferent enzyme as we have seen to be the case for NAD + . However, for a pros¬  thetic group, regeneration occurs as part of the enzyme reaction sequence.    An enzyme provides a lower-energy pathway from substrate to product but does  not affect the overall free energy change for the reaction.    Much of our understanding of how enzymes catalyze chemical reactions comes  from transition state theory, which was developed in the 1930s, principally  by Henry Eyring. Consider a bimolecular reaction involving three atoms, such  as the reaction of a hydrogen atom with diatomic hydrogen (H 2 ) to yield a  new H 2 molecule and a different hydrogen atom:   In this reaction, H c must approach the diatomic molecule H A —H B so that, at  some point in the reaction, there exists a high-energy (unstable) complex rep¬  resented as H A ***H B ***H C . In this complex, the H A —H B covalent bond is in  the process of breaking while the H B —H c bond is in the process of forming.  The point of highest free energy is called the transition state of the system.   Reactants generally approach one another along the path of minimum free  energy, their so-called reaction coordinate. A plot of free energy versus the  reaction coordinate is called a transition state diagram or reaction coordi¬  nate diagram (Fig. 11-5). The reactants and products are states of minimum  free energy, and the transition state corresponds to the highest point of the  diagram. For the H + H 2 reaction, the reactants and products have the same  free energy (Fig. 11-5 a). If the atoms in the reacting system are of different  types, such as in the reaction   where A and B are the reactants, P and Q are the products, and X* represents  the transition state, the transition state diagram is no longer symmetrical  because there is a free energy difference between the reactants and products  (Fig. 11-5 b). In either case, AG*, the free energy of the transition state less  that of the reactants, is known as the free energy of activation.   Passage through the transition state requires only 10 13 to 10 14 s, so the  concentration of the transition state in a reacting system is small. Hence, the  decomposition of the transition state to products (or back to reactants) is    FIG. 11-5 Transition state diagrams, (a) The H + H 2 reaction.  The reactants and products correspond to low free energy structures.  The point of highest free energy is the transition state, in which the  reactants are partially converted to products. AG* is the free energy  of activation, the difference in free energy between the reactants and    the transition state, X*. (b) Transition state diagram for the reaction A  + B —» P + Q. This is a spontaneous reaction; that is, AG reac tion < 0  (the free energy of P + Q is less than the free energy of A + B).    postulated to be the rate-determining process of the overall reaction.  Thermodynamic arguments lead to the conclusion that the reaction rate  is proportional to e , where R is the gas constant and T is the absolute   temperature. Thus, the greater the value ofAG'\ the slower the reaction rate. This  is because the larger the AG^, the smaller the number of reactant molecules  that have sufficient thermal energy to achieve the transition state free energy.   Chemical reactions commonly consist of several steps. For a two-step  reaction such as   where I is an intermediate of the reaction, there are two transition states and  two activation energy barriers. The shape of the transition state diagram for  such a reaction reflects the relative rates of the two steps (Fig. 11-6). If the  activation energy of the first step is greater than that of the second step, then  the first step is slower than the second step, and conversely, if the activation   FIG. 11-6 Transition state diagram for a two-step reaction. The blue curve  represents a reaction (A —> I —» P) whose first step is rate determining, and the red curve  represents a reaction whose second step is rate determining.   FIG. 11-7 Effect of a catalyst on the transition state diagram of a reaction. Here  AAG* cat = AG*(uncat) - AG* (cat).    energy of the second step is greater. In a multistep reaction, the step with the  highest transition state free energy acts as a “bottleneck” and is therefore said   Catalysts Reduce AG*. Catalysts act by providing a reaction pathway with a  transition state whose free energy is lower than that in the uncatalyzed reaction  (Fig. 11 -7). The difference between the values of AG* for the uncatalyzed and  catalyzed reactions, AAG* at , indicates the efficiency of the catalyst. The rate  enhancement (ratio of the rates of the catalyzed and uncatalyzed reactions) is  given by e ^ G ^ /RT % Hence, at 25°C (298 K), a 10-fold rate enhancement re¬  quires a AAG* at of only 5.71 kj • mol X , which is less than half the free en¬  ergy of a typical hydrogen bond. Similarly, a millionfold rate acceleration  occurs when A A G* at ~ 34 kj • mol 1 , a small fraction of the free energy of  most covalent bonds (see Sample Calculation 11-1). Thus, from a theoretical  standpoint, tremendous catalytic efficiency seems within reach of the reactive  groups that occur in the active sites of enzymes. How these groups actually  function at the atomic level is the subject of much of this and other chapters.   Note that a catalyst lowers the free energy barrier by the same amount for  both the forward and reverse reactions (Fig. 11-7). Consequently, a catalyst  equally accelerates the forward and reverse reactions. Keep in mind also that  while a catalyst can accelerate the conversion of reactants to products (or prod¬  ucts back to reactants), the likelihood of the net reaction occurring in one di¬  rection or the other depends only on the free energy difference between the    By how much must an enzyme reduce the activation energy of a reaction at 37°C  in order for the reaction to occur 50 times faster than in the absence of the enzyme?   = (8.3145 J • K -1 • mol -1 )(37 + 273) Kin 50  = 10,000 J • mol 1 = 10 kj • mol 1    reactants and the products. If AG reaction < 0, the reaction proceeds sponta¬  neously from reactants toward products; if A G reaction > 0, the reverse reaction  proceeds spontaneously. An enzyme cannot alter AG reaction ; it can only decrease  AG* to allow the reaction to approach equilibrium (where the rates of the for¬  ward and reverse reactions are equal) more quickly than it would in the absence  of a catalyst. The actual velocity with which reactants are converted to prod¬  ucts is the subject of kinetics (Section 12-1).   • Sketch and label the various parts of  transition state diagrams for a reaction  with and without a catalyst.    • Amino acid side chains that can donate or accept protons can participate in  chemical reactions as acid or base catalysts.   • Nucleophilic groups can catalyze reactions through the transient formation of  covalent bonds with the substrate.   • In metal ion catalysis, the unique electronic properties of the metal ion facilitate  the reaction.   • Enzymes accelerate reactions by bringing reacting groups together and orienting  them for reaction.   • Transition state stabilization can significantly lower the activation energy for a  reaction.    Enzymes achieve their enormous rate accelerations via the same catalytic mech¬  anisms used by chemical catalysts. Enzymes have simply been better designed  through evolution. Enzymes, like other catalysts, reduce the free energy of the  transition state (AG*); that is, they stabilize the transition state of the catalyzed  reaction. What makes enzymes such effective catalysts is their specificity of sub¬  strate binding combined with their arrangement of catalytic groups. As we will  see, however, the distinction between substrate-binding groups and catalytic  groups is somewhat arbitrary.   The types of catalytic mechanisms that enzymes employ have been classi¬  fied as   In this section, we consider each of these types of mechanisms in turn. Much  can be learned about enzymatic reaction mechanisms by examining the corre¬  sponding nonenzymatic reactions of model compounds. Both types of reactions  can be described using the same conventions (Box 11-1).   General acid catalysis is a process in which proton transfer from an acid low¬  ers the free energy of a reactions transition state. For example, an uncatalyzed  keto-enol tautomerization reaction occurs quite slowly as a result of the high  free energy of its carbanion-like transition state (Fig. 11-8#; the transition  state is drawn in square brackets to indicate its instability). Proton donation  to the oxygen atom (Fig. 11 -8 b), however, reduces the carbanion character of  the transition state, thereby accelerating the reaction.   A reaction may also be stimulated by general base catalysis if its rate is in¬  creased by proton abstraction by a base (e.g., Fig. 11-8c). Some reactions may  be simultaneously subject to both processes; these are concerted acid-base  catalyzed reactions.    While it is sometimes sufficient to draw the structures of a reac¬  tion’s substrates and products, a full understanding of the reaction  mechanism requires knowing how electrons are reallocated as  covalent bonds are broken and re-formed. Biochemists use the  curved arrow convention to show how pairs of electrons are  rearranged during a reaction. Although single-electron reactions  also occur in biochemistry, we will focus on the more common  two-electron reactions.   The movement of an electron pair (which may be either a lone  pair or a pair forming a covalent bond) is symbolized by a curved  arrow emanating from the electron pair and pointing to the electron-  deficient center attracting the electron pair. For example, bond  breakage is shown as   An example of a more complicated reaction requiring several  curved arrows is the formation of an imine (Schiff base), a bio¬  chemically important reaction between an amine and an aldehyde  or ketone:    In the first reaction step, the amine’s unshared electron pair adds  to the electron-deficient carbonyl carbon while one electron pair  from its C=0 double bond transfers to the oxygen atom. Following  tautomerization (the shift of a hydrogen atom), the unshared  electron pair on the nitrogen atom of the carbinolamine intermedi¬  ate adds to the electron-deficient carbon atom with the elimination  of a hydroxide ion.   Familiarity with the relative electronegativities of atoms  0 > N » C ~ H   may be helpful in predicting the movement of electrons. In addi¬  tion, the rules of chemical reason apply to the system: For example,  there are never five bonds to a carbon atom or two bonds to a  hydrogen atom.    FIG. 11-8 Mechanisms of keto-enol  tautomerization. (a) Uncatalyzed, (b) General  acid catalyzed, (c) General base catalyzed.   The acid is represented as H—A and the  base as B.    Most enzymes are active within only a narrow pH range, typically  5 to 9. This is a result of the effects of pH on a combination of  factors: (1) the binding of substrate to enzyme, (2) the ionization  states of the amino acid residues involved in the catalytic activity of  the enzyme, (3) the ionization of the substrate, and (4) the variation  of protein structure (usually significant only at extremes of pH).   The rates of many enzymatic reactions exhibit bell-shaped  curves as a function of pH. For example, the pH dependence of  the rate of the reaction catalyzed by fumarase (Section 17-3G)  produces the following curve:    The observed pK's (the inflection points of the curve) often pro¬  vide valuable clues to the identities of the amino acid residues  essential for enzymatic activity. For example, an observed p K of  ~4 suggests that an Asp or Glu residue is essential to the en¬  zyme. Similarly, pK's of ~6 or —10 suggest the participation of a  His or a Lys residue, respectively. However, the pK of a given  acid-base group may vary by as much as several pH units from  its expected value, depending on its microenvironment (e.g., an  Asp residue in a nonpolar environment or in close proximity to  another Asp residue would attract protons more strongly than  otherwise and hence have a higher p K). Furthermore, pH effects  on an enzymatic rate may reflect denaturation of the enzyme  rather than protonation or deprotonation of specific catalytic  residues. The replacement of a particular residue by site-directed  mutagenesis or comparisons of enzyme variants generated by  evolution is a more reliable approach to identifying residues that  are required for substrate binding or catalysis.   In addition to pH, other environmental factors, such as temper¬  ature and salt concentration, affect enzyme catalysis, typically  producing bell-shaped activity curves as shown here for pH. The  optimal conditions (the peak of the activity curve) frequently corre¬  spond to the prevailing environmental conditions of the cell or  organism, indicating that evolution has fine-tuned enzymes for  maximum efficiency.    Such curves reflect the ionization of certain amino acid residues [Figure adapted from Tanford, C., Physical Chemistry of   that must be in a specific ionization state for enzymatic activity. Macromolecules, p. 647, Wiley (1961).]   Many types of biochemical reactions are susceptible to acid and/or base  catalysis. The side chains of the amino acid residues Asp, Glu, His, Cys, Tyr,  and Lys have p Ks in or near the physiological pH range (Table 4-1), which  permits them to act as acid and/or base catalysts. Indeed, the ability of enzymes  to arrange several catalytic groups around their substrates makes concerted  acid-base catalysis a common enzymatic mechanism. The catalytic activity of  these enzymes is sensitive to pH, since the pH influences the state of proton¬  ation of side chains at the active site (Box 11-2).    RNase A Is an Acid-Base Catalyst. Bovine pancreatic RNase A provides  an example of enzymatically mediated acid-base catalysis. This digestive en¬  zyme (Fig. 11-9) is secreted by the pancreas into the small intestine, where it  hydrolyzes RNA to its component nucleotides. The isolation of 2',3'-cyclic nu¬  cleotides from RNase A digests of RNA indicates that 2',3'-cyclic nucleotides    FIG. 11-9 X-Ray structure of bovine pancreatic RNase S. A nonhydrolyzable  substrate analog, the dinucleotide phosphonate UpcA (red), is bound in the active site.  RNase S is a catalytically active form of RNase A in which the peptide bond between  residues 20 and 21 has been hydrolyzed. [Illustration, Irving Geis/Geis Archives Trust. Copyright  Howard Hughes Medical Institute. Reproduced by permission.] Q See Interactive  Exercise 6.    are intermediates in the RNase A reaction (Fig. 11-10). The pH dependence  of the rate of the RNase A reaction suggests the involvement of two ionizable  residues with pK values of 5.4 and 6.4. This information, together with chem¬  ical derivatization and X-ray studies, indicates that RNase A has two essential His   His 12 acts as a  general base and  His 119 as a  general acid to  promote    FIG. 11-10 The RNase A mechanism. The bovine pancreatic RNase A-catalyzed hydrolysis  of RNA is a two-step process with the intermediate formation of a 2',3'-cyclic nucleotide.    residues, His 12 and His 119, that act in a concerted manner as general acid and  base catalysts. Evidently, RNase A catalyzes a two-step reaction (Fig. 11-10):   1. His 12, acting as a general base, abstracts a proton from an RNA 2'-OH  group, thereby promoting its nucleophilic attack on the adjacent phos¬  phorus atom. His 119, acting as a general acid, promotes bond scis¬  sion by protonating the leaving group.   2. After the leaving group departs, water enters the active site, and the  2',3'-cyclic intermediate is hydrolyzed through what is essentially the  reverse of the first step. Thus, His 12 now acts as a general acid and  His 119 as a general base to yield the hydrolyzed RNA and the en¬  zyme in its original state.   Covalent catalysis accelerates reaction rates through the transient formation of  a catalyst-substrate covalent bond. Usually, this covalent bond is formed by the  reaction of a nucleophilic group on the catalyst with an electrophilic group  on the substrate, and hence this form of catalysis is often also called nucle¬  ophilic catalysis. The decarboxylation of acetoacetate, as chemically catalyzed  by primary amines, is an example of such a process (Fig. 11-11). In the first  stage of this reaction, the amine, a nucleophile, attacks the carbonyl group  of acetoacetate to form a Schiff base (imine bond; the same reaction shown  in Box 11-1):   The protonated nitrogen atom of the covalent intermediate then acts as an  electron sink (Fig. 11-11, bottom) to reduce the high-energy enolate character  of the transition state. The formation and decomposition of the Schiff base  occurs quite rapidly so that it is not the rate-determining step of this reaction.   1. The nucleophilic reaction between the catalyst and the substrate to  form a covalent bond.    FIG. 11-11 The decarboxylation of  acetoacetate. The uncatalyzed reaction  mechanism is at the top, and the mechanism as  catalyzed by primary amines is at the bottom.    Enzymatic Catalysis 2. The withdrawal of electrons from the reaction center by the now elec¬   3. The elimination of the catalyst, a reaction that is essentially the reverse  of stage 1.   The nucleophilicity of a substance is closely related to its basicity. Indeed, the  mechanism of nucleophilic catalysis resembles that of base catalysis except that,  instead of abstracting a proton from the substrate, the catalyst nucleophilically  attacks the substrate to form a covalent bond. Biologically important nucle¬  ophiles are negatively charged or contain unshared electron pairs that easily  form covalent bonds with electron-deficient centers (Fig. 11-12#). Electrophiles,  in contrast, include groups that are positively charged, contain an unfilled  valence electron shell, or contain an electronegative atom (Fig. 11-12^).   An important aspect of covalent catalysis is that the more stable the cova¬  lent bond formed, the less easily it can decompose in the final steps of a reaction.  A good covalent catalyst must therefore combine the seemingly contradictory  properties of high nucleophilicity and the ability to form a good leaving group,  that is, to easily reverse the bond formation step. Groups with high polariz¬  ability (highly mobile electrons), such as imidazole and thiol groups, have these  properties and hence make good covalent catalysts. Functional groups in pro¬  teins that act in this way include the unprotonated amino group of Fys, the  imidazole group of His, the thiol group of Cys, the carboxyl group of Asp,  and the hydroxyl group of Ser. In addition, several coenzymes, notably thi¬  amine pyrophosphate (Section 15-3B) and pyridoxal phosphate (Section 21-2A),  function in association with their apoenzymes as covalent catalysts. The large  variety of covalently linked enzyme-substrate reaction intermediates that have  been isolated demonstrates that enzymes commonly employ covalent catalytic  mechanisms.   Nearly one-third of all known enzymes require metal ions for catalytic activ¬  ity. This group of enzymes includes the metalloenzymes, which contain  tightly bound metal ion cofactors, most commonly transition metal ions  such as Fe 2+ , Fe 3+ , Cu 2+ , Mn 2+ , or Co 2+ . These catalytically essential metal ions  are distinct from ions such as Na + , K + , or Ca 2+ , which often play a structural   (a) Nucleophilic groups such as hydroxyl, sulfhydryl, amino, and imidazole groups are nucle¬  ophiles in their basic forms. ( b ) Electrophilic groups contain an electron-deficient atom (red).    Identify the amino acid side chains that include the nucleophiles and  electrophiles shown here.    rather than a catalytic role in enzymes. Ions such as Mg 2+ and Zn 2+ may be  either structural or catalytic.   Metal ions participate in the catalytic process in three major ways:   1. By binding to substrates to orient them properly for reaction.   2. By mediating oxidation-reduction reactions through reversible changes  in the metal ions oxidation state.   In many metal ion—catalyzed reactions, the metal ion acts in much the  same way as a proton to neutralize negative charge. Yet metal ions are often  much more effective catalysts than protons because metal ions can be present  in high concentrations at neutral pH (at which [H + ] = 10 7 M) and may  have charges greater than +1.   A metal ions charge also makes its bound water molecules more acidic  than free H 2 0 and therefore a source of nucleophilic OH ions even below  neutral pH. An excellent example of this phenomenon occurs in the catalytic  mechanism of carbonic anhydrase (Box 2-2), a widely occurring enzyme that  catalyzes the reaction   Carbonic anhydrase contains an essential Zn 2+ ion that the enzymes  X-ray structure indicates lies at the bottom of a 15-A-deep active site cleft,  where it is tetrahedrally coordinated by three evolutionarily invariant His side  chains (Fig. 11 -13 a). The Zn 2+ ion polarizes a water molecule (not visual¬  ized in Fig. 11-13 a) so that it ionizes to form OH , which nucleophilically  attacks the substrate C0 2 to yield HC0 3 (Fig. 11-13^). The proton pro¬  duced in the reaction is shuttled to the enzymes surface through base cataly¬  sis that is facilitated by a fourth His residue (His 64). The enzymes catalytic  site is then regenerated by the binding of another H 2 0 to the Zn 2+ ion.    Although enzymes employ catalytic mechanisms that resemble those of organic  model reactions, they are far more catalytically efficient than the models. Such  efficiency must arise from the specific physical conditions at enzyme catalytic  sites that promote the corresponding chemical reactions. The most obvious ef¬  fects are proximity and orientation: Reactants must come together with the  proper spatial relationship for a reaction to occur. Consider the bimolecular  reaction of imidazole with />-nitrophenylacetate.   The progress of the reaction is conveniently monitored by the appearance of  the intensely yellow />-nitrophenolate ion. The related intramolecular reaction    FIG. 11-13 The role of Zn 2+ in carbonic  anhydrase. (a) The active site of the human  enzyme. The polypeptide is shown in ribbon form  (gold) with its side chains shown in stick form  colored according to atom type (C green, N blue,  and 0 red). The Zn 2+ ion (cyan sphere) is  liganded to three His side chains and the HCO3  ion, which is shown in ball-and-stick form. The  HCO3 ion also makes van der Waals contacts  (dot surface) and hydrogen bonds (dashed gray  lines) with a Thr and Glu residue. [Based on an  X-ray structure by K.K. Kannan, Bhabha Atomic  Research Center, Bombay, India. PDBid 1 HCBJ  O See Interactive Exercise 7. (b) The reaction  catalyzed by carbonic anhydrase. Im represents  the His imidazole group.   Chapter 11 Enzymatic Catalysis occurs about 24 times faster. Thus, when the imidazole is covalently attached   to the reactant, it is 24 times more effective than when it is free in solution.  This rate enhancement results from both proximity and orientation effects.   By simply binding their substrates, enzymes facilitate their catalyzed reac¬  tions in four ways:   1. Enzymes bring substrates into contact with their catalytic groups and, in  reactions with more than one substrate, with each other. However, cal¬  culations based on simple model systems suggest that such proximity ef¬  fects alone can enhance reaction rates by no more than a factor of —5.   2. Enzymes bind their substrates in the proper orientations for reaction.  Molecules are not equally reactive in all directions. Rather, they react most  readily if they have the proper relative orientation. For example, in an S N 2  (bimolecular nucleophilic substitution) reaction, the incoming nucleophile  optimally attacks its target along the direction opposite to that of the bond  to the leaving group (Fig. 11-14). Reacting atoms whose approaches de¬  viate by as little as 10° from this optimum direction are significantly less  reactive. It is estimated that properly orienting substrates can increase re¬  action rates by a factor of up to —100. Enzymes, as we will see, align their  substrates and catalytic groups so as to optimize reactivity.   3. Charged groups may help stabilize the transition state of the reaction,  a phenomenon termed electrostatic catalysis. The expulsion of water  from the active site may enhance this effect. The charge distribution  around the active sites of enzymes may also guide polar substrates to¬  ward their binding site.   4. Enzymes freeze out the relative translational and rotational motions of  their substrates and catalytic groups. This is an important aspect of  catalysis because, in the transition state, the reacting groups have little  relative motion. Indeed, experiments with model compounds suggest  that this ejfect can promote rate enhancements of up to — 10 7 \   FIG. 11-14 The geometry of an S N 2 reaction. (1) The   attacking nucleophile, Y, must approach the tetrahedrally coordinated  and hence sp 3 -hybridized C atom along the direction opposite that of  its bond to the leaving group, X. In the transition state of the reaction,  the C atom becomes trigonal bipyramidally coordinated and hence  sp 2 -p hybridized, with the p orbital (blue) forming partial bonds to X  and Y. The three sp 2 orbitals form bonds to the C atom’s three other    substituents (R, R', and R"), which have shifted their positions into the  plane perpendicular to the X—C—Y axis (curved arrows). Any  deviation from this optimal geometry would increase the free energy  of the transition state, AG*, and hence reduce the rate of the  reaction. ( 2 ) The transition state then decomposes to products in  which R, R', and R" have inverted their positions about the C atom,  which has rehybridized to sp 3 , and X - has been released.   Bringing substrates and catalytic groups together in a reactive orientation or¬  ders them and therefore has a substantial entropic penalty. The free energy re¬  quired to overcome this entropy loss is supplied by the binding energy of the  substrate(s) to the enzyme and contributes to the decreased AAG*. Keep in  mind, however, that enzymes are dynamic molecules that may adopt a variety  of different conformations before and after binding their substrates. Indeed,  such flexibility is essential in order for an enzyme to productively interact with  its substrates, transition state, and products during the course of a reaction.    The rate enhancements effected by enzymes are often greater than can be rea¬  sonably accounted for by the catalytic mechanisms discussed so far. However,  we have not yet considered one of the most important mechanisms of enzy¬  matic catalysis: An enzyme may bind the transition state of the reaction it cat¬  alyzes with greater affinity than its substrates or products. When taken together  with the previously described catalytic mechanisms, preferential transition state  binding explains the observed rates of enzyme-catalyzed reactions.   The original concept of transition state binding proposed that enzymes  mechanically strained their substrates toward the transition state geometry  through binding sites into which undistorted substrates did not properly fit. Such  strain promotes many organic reactions. For example, the rate of the reaction    is 315 times faster when R is CH 3 rather than H because of the greater steric  repulsion between the CH 3 groups and the reacting groups. The strained re¬  actant more closely resembles the transition state of the reaction than does the  corresponding unstrained reactant. Thus, as was first suggested by Linus  Pauling and further amplified by Richard Wolfenden and Gustav Lienhard,  enzymes that preferentially bind the transition state structure increase its concen¬  tration and therefore proportionally increase the reaction rate.   The more tightly an enzyme binds its reactions transition state relative to  the substrate, the greater is the rate of the catalyzed reaction relative to that  of the uncatalyzed reaction; that is, catalysis results from the preferential bind¬  ing and therefore the stabilization of the transition state relative to the sub¬  strate (Fig. 11-15). In other words, the free energy difference between an  enzyme-substrate complex (ES) and an enzyme-transition state complex (ES*) is  less than the free energy difference between S and S* in an uncatalyzed reaction.   As we saw in Section 11-2, the rate enhancement of a catalyzed reaction  is given by e AAG ^ tlRT where AAG* at is the difference in the values of AG* for  the uncatalyzed (AGfj) and the catalyzed (AG|) reactions. Thus, a rate en¬  hancement of 10 6 , which requires that an enzyme bind its transition state  complex with 10 6 -fold higher affinity than its substrate, corresponds to a  34.2 kj • mol 1 stabilization at 25°C, roughly the free energy of two hy¬  drogen bonds. Consequently, the enzymatic binding of a transition state by two  hydrogen bonds that cannot form when the substrate first binds to the enzyme  should result in a rate enhancement of ~10 6 based on this effect alone.   It is commonly observed that an enzyme binds poor substrates, which have  low reaction rates, as well as or even better than good ones, which have high  reaction rates. Thus, a good substrate does not necessarily bind to its enzyme  with high affinity, but it does so on activation to the transition state. Thus,   FIG. 11-15 Effect of preferential  transition state binding. The reaction  coordinate diagram for a hypothetical enzyme-  catalyzed reaction involving a single substrate is  blue, and the diagram for the corresponding  uncatalyzed reaction is red. AGn is the free  energy of activation for the nonenzymatic  reaction and AG| is the free energy of activation  for the enzyme-catalyzed reaction. The small dips  in the reaction coordinate diagram for the  enzyme-catalyzed reaction arise from the binding  of substrate and product to the enzyme. O See  the Animated Figures.   Chapter 11 Enzymatic Catalysis Fischers lock-and-key model for enzyme action applies more to transition state   For at least some enzymes, transition state stabilization makes only a mi¬  nor contribution to rate enhancement. In many such cases, as Thomas Bruice  has explained, the enzyme promotes the reaction by instead stabilizing the so-  called near attack conformation, a step along the reaction coordinate in which  the reactants are properly oriented and in van der Waals contact but have not  yet reached the transition state. These findings suggest that enzymes have  evolved to use a variety of strategies—including stabilizing the transition state  or something leading up to it—to accelerate chemical reactions.   Transition State Analogs Are Enzyme Inhibitors. If an enzyme preferentially  binds its transition state , then it can be expected that transition state analogs,  stable molecules that geometrically and electronically resemble the transition state ,  are potent inhibitors of the enzyme. For example, the reaction catalyzed by pro¬  line racemase from Clostridium sticklandii is thought to occur via a planar  transition state:   Proline racemase is inhibited by the planar analogs of proline, pyrrole-2-  carboxylate and A-1 -pyrroline-2-carboxylate,   • Describe how protein functional groups  can act as acid and base catalysts. How  is it possible for a single amino acid side  chain to function as both?   • Explain how nucleophiles function as  covalent catalysts. Which amino acids are  good at this?   • List the ways in which metal ions partici¬  pate in catalysis.   • What roles do proximity and orientation  play in enzymatic catalysis?   • Why is it unlikely that nonenzymatic cata¬  lysts operate by preferentially binding the  transition state?   both of which bind to the enzyme with 160-fold greater affinity than does  proline. These compounds are therefore thought to be analogs of the transi¬  tion state in the proline racemase reaction.   Hundreds of transition state analogs for various enzymes have been re¬  ported. Some are naturally occurring antibiotics. Others were designed to in¬  vestigate the mechanism of particular enzymes or to act as specific enzyme  inhibitors for therapeutic or agricultural use. Indeed, the theory that enzymes  bind transition states with higher affinity than substrates has led to a rational  basis for drug design based on the understanding of specific enzyme reaction mech¬  anisms (Section 12-4).    • Model building indicates that binding to lysozyme distorts the substrate sugar  residue.   • Lysozyme’s active site Asp and Glu residues promote substrate hydrolysis by  acid-base catalysis, covalent catalysis, and stabilization of an oxonium ion transi¬  tion state.    In the remainder of this chapter, we investigate the catalytic mechanisms of  some well-characterized enzymes. In doing so, we will see how enzymes apply  the catalytic principles described in the preceding section.   FIG. 11-16 The lysozyme cleavage site. The enzyme cleaves after a p(1—>4)  linkage in the alternating NAG—NAM polysaccharide component of bacterial cell walls.    Lysozyme is an enzyme that destroys bacterial cell walls. It does so by hy¬  drolyzing the (3(1—>4) glycosidic linkages from A^acetylmuramic acid (NAM  or MurNAc) to A^acetylglucosamine (NAG or GlcNAc) in cell wall pepti-  doglycans (Fig. 11-16 and Section 8-3B). It likewise hydrolyzes (3(1—>4)-  linked poly(NAG) (chitin; Section 8-2B), a cell wall constituent of most fungi  as well as the major component of the exoskeletons of insects and crustaceans.  Lysozyme occurs widely in the cells and secretions of vertebrates, where it  probably functions as a bactericidal agent or helps dispose of bacteria after  they have been killed by other means.   Hen egg white (HEW) lysozyme is the most widely studied species of  lysozyme and is one of the mechanistically best understood enzymes. It is a  rather small protein (14.3 kD) whose single polypeptide chain consists of 129  amino acid residues and is internally cross-linked by four disulfide bonds.  Lysozyme catalyzes the hydrolysis of its substrate at a rate that is ~10 8 -fold  greater than that of the uncatalyzed reaction.   The X-ray structure of HEW lysozyme (Fig. 11-17), which was elucidated by  David Phillips in 1965 (and hence was the first enzyme to have its structure   FIG. 11-17 X-Ray structure of HEW  lysozyme in complex with (NAG) 6 . The protein  is represented by its transparent molecular  surface with its polypeptide chain in worm form  colored in rainbow order from blue at its  N-terminus to red at its C-terminus. The (NAG) 6 ,  which is drawn in stick form with its sugar rings  designated A, at its nonreducing end, through F,  at its reducing end, binds in a deep cleft in the  enzyme surface. Rings A, B, and C (colored  according to atom type with C green, N blue,  and 0 red) are observed in the X-ray structure  of the complex of (NAG) 3 with lysozyme; the  positions of rings D, E, and F (C atoms cyan)  were inferred by model building. The side chains  of lysozyme’s active site residues, Glu 35 and  Asp 52, which are drawn in space-filling form  (C atoms yellow), catalyze the hydrolysis of the  glycosidic bond between rings D and E. [Based  on an X-ray structure by David Phillips, Oxford  University. PDBid 1HEW.] O See Interactive  Exercise 8 and Kinemage Exercise 9-1.    determined), shows that the protein molecule is roughly ellipsoidal in shape  with dimensions 30 X 30 X 45 A. Its most striking feature is a prominent cleft,  the substrate-binding site, that traverses one face of the molecule.   The elucidation of an enzymes mechanism of action requires a knowl¬  edge of the structure of its enzyme-substrate complex. This is because, even  if the active site residues have been identified through chemical and physical  means, their three-dimensional arrangement relative to the substrate as well  as to each other must be known in order to understand how the enzyme  works. However, an enzyme binds its good substrates only transiently before  it catalyzes a reaction and releases products. Consequently, much of our struc¬  tural knowledge of enzyme-substrate complexes comes from X-ray studies of  enzymes in their complexes with substrate analogs that bind but do not react  or react very slowly.   Phillips 5 X-ray structure of lysozyme includes the trisaccharide (NAG) 3 ,  which is only slowly hydrolyzed by lysozyme. However, the enzyme efficiently  catalyzes the hydrolysis of substrates containing at least six saccharide units,  so Phillips used model building to investigate how a larger substrate could  bind to the enzyme. Lysozymes active site cleft is long enough to accommo¬  date an oligosaccharide of six residues (designated A to F in Fig. 11-17).  However, the fourth residue (D) appeared unable to bind to the enzyme be¬  cause its C6 and 06 atoms too closely contacted protein side chains and  residue C. This steric interference could be relieved by distorting the glucose  ring from its normal chair conformation to that of a half-chair (Fig. 11-18).  This distortion moves the C6 group from its normal equatorial position to an  axial position, where it makes no close contacts and can hydrogen bond to  the backbone carbonyl group of Gin 57 and the backbone amido group of  Val 109. The other saccharide residues apparently bind to the enzyme without  distortion and with a number of favorable hydrogen-bonding and van der Waals  contacts. Some of these hydrogen bonds are diagrammed in Fig. 11-19.   In the enzymes natural substrate, every second residue is an NAM. Model  building, however, indicated that the enzyme could not accommodate a lactyl  side chain in subsites C or E. Hence, the NAM residues must bind at posi¬  tions B, D, and F, as drawn in Fig. 11-19. Moreover, since lysozyme hydrolyzes  (NAG)6 between residues D and E, it must cleave the glycosidic bond in its  natural substrate between the NAM residue in subsite D and the NAG residue  in subsite E.   The bond that lysozyme cleaves was identified by allowing lysozyme to  catalyze the hydrolysis of (NAG) 3 in H 2 18 0. The bond cleaved could either  be between Cl and the bridge oxygen Ol or between Ol and C4 of the next  sugar ring. The product of the hydrolysis reaction had 18 0 bonded to the Cl  atom of its newly liberated reducing terminus, thereby demonstrating that  bond cleavage occurs between Cl and Ol:   In addition, this reaction occurs with retention of configuration so that the  D-ring product remains the (3 anomer.    FIG. 11-18 Chair and half-chair conformations. Hexose rings normally assume the  chair conformation. However, binding to lysozyme distorts the D ring into the half-chair  conformation in which atoms Cl, C2, C5, and 05 are coplanar. © See the Animated  Figures.    FIG. 11-19 The interactions of lysozyme with its substrate. The view is into the binding cleft  with the heavier edges of the rings facing the outside of the enzyme and the lighter ones against the  bottom of the cleft [Modified from a drawing by Irving GeisJ Q See Kinemage Exercise 9-2.   The reaction catalyzed by lysozyme, the hydrolysis of a glycoside, is the con¬  version of an acetal to a hemiacetal. Nonenzymatic acetal hydrolysis is an acid-  catalyzed reaction that involves the protonation of a reactant oxygen atom  followed by cleavage of its C—O bond (Fig. 11-20). This results in the tran¬  sient formation of a resonance-stabilized carbocation that is called an oxonium  ion. To attain resonance stabilization, the oxonium ions R and R' groups must  be coplanar with its C, O, and H atoms. The oxonium ion then adds water  to yield the hemiacetal and regenerate the acid catalyst. An  enzyme that mediates acetal hydrolysis should therefore in¬  clude an acid catalyst and possibly a group that can stabilize  an oxonium ion transition state.   The only functional groups in the immediate vicinity of  lysozymes reactive center that have the required catalytic  properties are the side chains of Glu 35 and Asp 52. These  side chains, which are disposed to either side of the glyco-  sidic linkage to be cleaved (Fig. 11-17), have markedly dif¬  ferent environments. Asp 52 is surrounded by several  conserved polar residues with which it forms a complex hy¬  drogen-bonded network. Asp 52 is therefore predicted to  have a normal p K; that is, it should be unprotonated and  hence negatively charged throughout the 3 to 8 pH range  over which lysozyme is catalytically active. Thus, Asp 52 can  function to electrostatically stabilize an oxonium ion. In con¬  trast, the carboxyl group of Glu 35 is nestled in a predom¬  inantly nonpolar pocket where it is likely to remain   to form an alcohol (R'OH) and a resonance-stabilized carbocation (oxonium ion).   The addition of water to the oxonium ion forms the hemiacetal and regenerates the H +  catalyst. Note that the oxonium ion’s C, 0, H, R, and R' atoms all lie in the same plane.    FIG. 1 1-21 The lysozyme reaction mechanism. Glu 35 acts  as an acid catalyst, and Asp 52 acts as a covalent catalyst. Only the  substrate D and E rings are shown. R represents the A/-acetyl group  at C2, and R' represents the CH 3 CHCOO“ group at C3. The  resonance-stabilized oxonium ion transition state requires that Cl, C2,    C5, and 05 be coplanar (shading), creating a half-chair conformation.  Step 5 includes the participation of an oxonium ion transition state  that is not shown. Q See Kinemage Exercise 9.   protonated at unusually high pH values for carboxyl groups. This residue can  therefore act as an acid catalyst. Studies using protein-modifying reagents and  site-directed mutagenesis (e.g., changing Asp 52 to Asn and Glu 35 to Gin)  have verified that these residues are catalytically important.   1. The enzyme attaches to a bacterial cell wall by binding to a hexasac-  charide unit. This distorts the D residue toward the half-chair confor¬  mation.   2. Glu 35 transfers its proton to the Ol atom bridging the D and E rings,  thereby facilitating cleavage of the Cl—Ol bond (general acid catal¬  ysis). This step converts the D ring to a resonance-stabilized oxonium  ion transition state whose formation is facilitated by the strain distort¬  ing it to the half-chair conformation (catalysis by the preferential bind¬  ing of the transition state). The positively charged oxonium ion is  stabilized by the presence of the nearby negatively charged Asp 52 car-  boxylate group (electrostatic catalysis). The E-ring product is released.   3. The Asp 52 carboxylate group nucleophilically attacks the now electron-  poor Cl of the D ring to form a covalent glycosyl—enzyme intermedi¬  ate (covalent catalysis).   5. Hydrolysis of the covalent bond with the assistance of Glu 35 (general  base catalysis), which involves another oxonium ion transition state, re¬  generates the active site groups. The enzyme then releases the D-ring  product, completing the catalytic cycle.   The double-displacement mechanism diagrammed in Fig. 11-21 allows  the incoming water molecule to attach to the same face of the D residue as  the E residue it replaces. Consequently, the configuration of the D residue is  retained. A single-displacement reaction, in which water directly displaces the  leaving group, would invert the configuration at Cl of the D ring between  the substrate and product, a result that is not observed.   Experimental Evidence Supports the Role of Strain in the Lysozyme  Mechanism. Many of the structural and mechanistic investigations of  lysozyme have focused on the catalytic role of strain. For example, the X-ray  structure of lysozyme in complex with NAM—NAG—NAM shows that the  trisaccharide binds, as predicted, to the B, C, and D subsites of lysozyme, with  the NAM in the D subsite distorted to the half-chair conformation. This strained  conformation is stabilized by a strong hydrogen bond between the D ring 06  and the backbone NH of Val 109 (as predicted by model building; Fig. 11-19).  Indeed, the mutation of Val 109 to Pro, which lacks the NH group to make  such a hydrogen bond, inactivates the enzyme.   As we have discussed in Section 11-3E, an enzyme that catalyzes a reac¬  tion by the preferential binding of its transition state has a greater binding  affinity for an inhibitor that has the transition state geometry (a transition  state analog) than it does for its substrate. The 8-lactone analog of (NAGfy  (Fig. 11 -22), which binds tightly to lysozyme, is a transition state analog of  lysozyme since this compound's lactone ring has the half chair conformation that  geometrically resembles the proposed oxonium ion transition state of the substrates  D ring. X-Ray studies confirm that this inhibitor binds to lysozyme such that  the lactone ring occupies the D subsite in a half-chair-like conformation.    FIG. 1 1 -22 Transition state analog inhibition of lysozyme. The 8-lactone analog of  (NAG) 4 (top) resembles the transition state of the lysozyme reaction (bottom). Note that  atoms Cl, C2, C5, and 05 in each structure are coplanar (as indicated by shading), consis¬  tent with the half-chair conformation of the hexose ring.   FIG. 1 1 -23 Position of the D ring during  catalysis by lysozyme. The position of the  substrate’s C and D rings and the catalytic Asp  52 are shown in superimposed X-ray structures  of the covalent complex between E35Q lysozyme  and NAG2FGIcF (C green, N blue, 0 red, and F  magenta) and the noncovalent complex between  lysozyme and a NAM-NAG—NAM substrate (C  yellow, N blue, and 0 red). Note that the  covalent bond between Asp 52 and Cl of the D  ring forms when the D ring in the noncovalent  complex relaxes from its distorted half-chair  conformation to an undistorted chair  conformation and the side chain of Asp 52  undergoes a 45° rotation about its C a —Cp bond.  [Based on X-ray structures by David Vocadlo and  Stephen Withers, University of British Columbia,  Vancouver, Canada; and Michael James,   University of Alberta, Edmonton, Canada. PDBid  1 H6MJ O See Kinemage Exercise 9-2.   • Using lysozyme as an example, what can  an enzyme’s structure reveal about its  catalytic mechanism?   • Why was an oxonium ion expected to be  involved in the lysozyme reaction?   • Describe the experimental evidence that  supports lysozyme’s catalysis by  acid-base catalysis, covalent catalysis,  and transition state stabilization.    Mass Spectrometry and X-Ray Crystallography Provide Support for  Covalent Catalysis. The existence of a covalent reaction intermediate was dif¬  ficult to verify. The lifetime of a glucosyl oxonium ion in water is —^10 12 s.  In order for such a short-lived intermediate to be experimentally observed, its  rate of formation must be made significantly greater than its rate of break¬  down. To do this, Stephen Withers capitalized on three phenomena. First, if  the reaction goes through an oxonium ion transition state, its formation  should be slowed by the electron-withdrawing effects of substituting F (the  most electronegative element) at C2 of the D ring. Second, mutating Glu 35  to Gin (E35Q) removes the general acid—base catalyst, further slowing all steps  involving the oxonium ion transition state. Third, substituting an additional  F atom at Cl of the D ring accelerates the formation of the intermediate be¬  cause this F is a good leaving group without the need of general acid cataly¬  sis. Making all three of these changes should increase the rate of formation of  the proposed covalent intermediate relative to its breakdown and hence  should result in its accumulation. Withers therefore incubated E35Q HEW  lysozyme with NAG-|3(1—>4)-2-deoxy-2-fluoro-|3-D-glucopyranosyl fluoride  (NAG2FGlcF; at left). Electrospray ionization mass spectrometry (ESI-MS;  Section 5-3D) of this reaction mixture revealed a sharp peak at 14,683 D,  consistent with the formation of the proposed covalent intermediate [lysozyme  has a molecular mass of 14,314 D and that of the NAG-(3(1—>4)-2-deoxy-2-  fluoro-(3-D-glucopyranosyl group is 369 D].   The X-ray structure of this covalent complex reveals an ~ 1.4-A-long co¬  valent bond between Cl of the D ring and a side chain carboxyl O of Asp  52. This D ring adopts an undistorted chair conformation, thus indicating  that it is a reaction intermediate rather than an approximation of the transi¬  tion state. The superposition of this covalent complex with that of the above-  described complex of NAM—NAG—NAM with wild-type HEW  lysozyme reveals how this covalent bond forms (Fig. 11-23). The shortening  of the 3.2-A distance between the D ring Cl and the Asp 52 O in the  NAM—NAG—NAM complex to ~ 1.4 A in the covalent complex occurs as  the D ring relaxes from the half-chair to the chair conformation and the Asp  52 side chain rotates ~45° about its C a —Cp bond.    • The catalytically active Ser, His, and Asp residues of serine proteases were identi¬  fied by chemical labeling and structural analysis.   • A binding pocket determines the substrate specificity of the various serine proteases.   • Serine proteases catalyze peptide bond hydrolysis via proximity and orientation  effects, acid-base catalysis, covalent catalysis, electrostatic catalysis, and transi¬  tion state stabilization.    Our next example of enzymatic mechanisms is a diverse and widespread group  of proteolytic enzymes known as serine proteases, so named because they have  a common catalytic mechanism involving a peculiarly reactive Ser residue. The  serine proteases include digestive enzymes from prokaryotes and eukary¬  otes, as well as more specialized proteins that participate in development,  blood coagulation (clotting), inflammation, and numerous other processes. In  this section, we focus on some of the best studied serine proteases: chy-  motrypsin, trypsin, and elastase.   Chymotrypsin, trypsin, and elastase are digestive enzymes that are synthesized  by the pancreas and secreted into the duodenum (the small intestines upper  loop). All these enzymes catalyze the hydrolysis of peptide (amide) bonds but  with different specificities for the side chains flanking the scissile (to be  cleaved) peptide bond. Chymotrypsin is specific for a bulky hydrophobic  residue preceding the scissile bond, trypsin is specific for a positively charged  residue, and elastase is specific for a small neutral residue (Table 5-4). Together,  they form a potent digestive team.   Chymotrypsins catalytically important groups were identified by chemi¬  cal labeling studies. A diagnostic test for the presence of the active site Ser of  serine proteases is its reaction with diisopropylphosphofluoridate (DIPF),  which irreversibly inactivates the enzyme. Other Ser residues, including those  on the same protein, do not react with DIPF. DIPF reacts only with Ser 195 of  chymotrypsin , thereby demonstrating that this residue is the enzymes active site Ser.  This specificity makes DIPF and related compounds extremely toxic (Box 11-3).    The use of DIPF as an enzyme-inactivating agent came about  through the discovery that organophosphorus compounds such as  DIPF are potent nerve poisons. The neurotoxicity of DIPF arises  from its ability to inactivate acetylcholinesterase, an enzyme that  catalyzes the hydrolysis of acetylcholine:   The esterase activity of acetylcholinesterase, like that of  chymotrypsin (Section 1 1-1 B), requires a reactive Ser residue.   Acetylcholine is a neurotransmitter: It transmits nerve impulses  across certain types of synapses (junctions between nerve cells).  Acetylcholinesterase in the synapse normally degrades acetylcholine  so that the nerve impulse has a duration of only a millisecond or so.  The inactivation of acetylcholinesterase prevents hydrolysis of the  neurotransmitter. As a result, the acetylcholine receptor, which is a  Na + -K + channel, remains open for longer than normal, thereby  interfering with the regular sequence of nerve impulses. DIPF is so  toxic to humans (death occurs through the inability to breathe) that  it has been used militarily as a nerve gas. Related compounds, such  as parathion and malathion, are useful insecticides because they  are far more toxic to insects than to mammals.   Neurotoxins such as DIPF and sarin (which gained notoriety after  its release by terrorists in a Tokyo subway in 1995)   This enzyme occurs as two isoforms (one has Arg at position  192 and the other has Gin) with different activities, and individuals  express widely differing levels of the enzyme. These factors may  account for the large observed differences in individuals’ sensitivity  to nerve poisons.    A second catalytically important residue, His 57, was discovered through  affinity labeling. In this technique, a substrate analog bearing a reactive group  specifically binds at the enzymes active site, where it reacts to form a stable  covalent bond with a nearby susceptible group (these reactive substrate analogs  have been dubbed the “Trojan horses” of biochemistry). The affinity labeled  group(s) can subsequently be isolated and identified.   Chymotrypsin specifically binds tosyl-L-phenylalanine chloromethyl-  ketone (TPCK) because of its resemblance to a Phe residue (one of chymotrypsin s  preferred substrate residues). Active site-bound TPCK’s chloromethylketone  group is a strong alkylating agent; it reacts only with His 57 (Fig. 11-24), thereby  inactivating the enzyme. Trypsin, which prefers basic residues, is similarly inac¬  tivated by tosyl-L-lysine chloromethylketone (at left).    B X-Ray Structures Provide Information about Catalysis, Substrate  Specificity, and Evolution   Chymotrypsin, trypsin, and elastase are strikingly similar: The primary struc¬  tures of these ~240-residue enzymes are ~40% identical (for comparison, the  a and (3 chains of human hemoglobin have 44% sequence identity).  Furthermore, all these enzymes have a reactive Ser and a catalytically essential  His. It therefore came as no surprise when their X-ray structures all proved to  be closely related.    FIG. 1 1 -24 Reaction of TPCK with His 57 of chymotrypsin.    The structure of bovine chymotrypsin was elucidated in 1967 by David  Blow. This was followed by the determination of the structures of bovine  trypsin (Fig. 11 -25) by Robert Stroud and Richard Dickerson, and porcine  elastase by David Shotton and Herman Watson. Each of these proteins is  folded into two domains, both of which have extensive regions of antiparallel   FIG. 1 1 -25 X-Ray structure of bovine  trypsin in covalent complex with its inhibitor  leupeptin. The protein is represented by its  transparent molecular surface with its  polypeptide chain in worm form colored in  rainbow order from its N-terminus (blue) to its  C-terminus (red). The side chains of the catalytic  triad, Ser 195, His 57, and Asp 102, are drawn  in ball-and-stick form colored according to atom  type (C green, N blue, 0 red) with hydrogen  bonds represented by dashed black lines.  Leupeptin (acetyl-Leu-Leu-Arg in which the  terminal carboxyl group is replaced by —CHO) is  drawn in stick form (C cyan, N blue, 0 red) with  its Arg side chain occupying the enzyme’s  specificity pocket (magenta mesh). LI and L2  are surface loops that span the walls of the  specificity pocket. [Based on an X-ray structure  by Daniel Koshland, Jr., University of California at  Berkeley. PDBid 2AGI.] O See Kinemage  Exercise 10-1.   FIG. 1 1 -26 The active site residues of chymotrypsin. The view is in approximately  the same direction as in Fig. 11-25. Residues are drawn in ball-and-stick form with  C green, N blue, and 0 red. The catalytic triad consists of Ser 195, His 57, and Asp 102.  [Based on an X-ray structure by Daniel Koshland, Jr., University of California at Berkeley.  PDBid 2AGI.]   FIG. 1 1 -27 Specificity pockets of three  serine proteases. The side chains of key  residues that determine the size and nature of  the specificity pocket are shown along with a  representative substrate for each enzyme.  Chymotrypsin prefers to cleave peptide bonds  following large hydrophobic side chains; trypsin  prefers Lys or Arg; and elastase prefers Ala, Gly,  or Val. [After a drawing in Branden, C. and Tooze,  J., Introduction to Protein Structure (2nd ed.),  Garland Publishing, p. 213 (1999).]    (3 sheets in a barrel-like arrangement but contain little helix. For convenience  in comparing the structures of these three enzymes, we will assign them the same  residue numbering system—that of bovine chymotrypsinogen, the 245-residue  precursor of chymotrypsin (Section 11-5D).   In all three structures, the catalytically essential His 57 and Ser 195  residues are located in the enzymes substrate-binding site (center of Fig. 11-25).  The X-ray structures also show that Asp 102, which is present in all serine  proteases, is buried in a nearby solvent-inaccessible pocket. These three invari¬  ant residues form a hydrogen-bonded constellation referred to as the catalytic triad  (Figs. 11-25 and 11-26).   Substrate Specificities Are Only Partially Rationalized. The X-ray struc¬  tures of the above three enzymes suggest the basis for their differing substrate  specificities (Fig. 11-27):   1. In chymotrypsin, the bulky aromatic side chain of the preferred Phe,  Trp, or Tyr residue that contributes the carbonyl group of the scissile  peptide fits snugly into a slitlike hydrophobic pocket located near the  catalytic groups.   2. In trypsin, the residue corresponding to chymotrypsin Ser 189, which  lies at the bottom of the binding pocket, is the anionic residue Asp.  The cationic side chains of trypsins preferred residues, Arg and Lys,  can therefore form ion pairs with this Asp residue. The rest of chy-  motrypsins specificity pocket is preserved in trypsin so that it can ac¬  commodate the bulky side chains of Arg and Lys (Fig. 11-25).   3. Elastase is so named because it rapidly hydrolyzes the otherwise nearly  indigestible Ala, Gly, and Val-rich protein elastin (a major connective  tissue component). Elastases binding pocket is largely occluded by the  side chains of Val and Thr residues that replace the Gly residues lin¬  ing the specificity pockets in both chymotrypsin and trypsin.  Consequently elastase, whose substrate-binding site is better described  as merely a depression, specifically cleaves peptide bonds after small  neutral residues, particularly Ala. In contrast, chymotrypsin and trypsin  hydrolyze such peptide bonds extremely slowly because these small sub¬  strates cannot be sufficiently immobilized on the enzyme surface for  efficient catalysis to occur.   Despite the foregoing, changing trypsins Asp 189 to Ser by site-directed mu¬  tagenesis (Section 3-5D) does not switch its specificity to that of chymotrypsin   trypsins specificity pocket with those of chymotrypsin fails to significantly im¬  prove this catalytic activity. However, trypsin is converted to a reasonably ac¬  tive chymotrypsin-like enzyme when two surface loops that connect the walls  of the specificity pocket, LI (residues 185-188) and L2 (residues 221-225),  are also replaced by those of chymotrypsin. These loops, which are conserved  in each enzyme, are apparently necessary not for substrate binding per se but  for properly positioning the scissile bond. These results highlight an important  caveat for genetic engineers: Enzymes are so exquisitely tailored to their func¬  tions that they often respond to mutagenic tinkering in unexpected ways.   Serine Proteases Exhibit Divergent and Convergent Evolution. We have  seen that sequence and structural similarities among proteins reveal their evo¬  lutionary relationships (Sections 5-4 and 6-2D). The great similarities among  chymotrypsin ., trypsin, and elastase indicate that these proteins arose through du¬  plications of an ancestral serine protease gene followed by the divergent evolution  of the resulting enzymes. Indeed, the close structural resemblance of these pan¬  creatic enzymes to certain bacterial proteases indicates that the primordial  trypsin gene arose before the divergence of prokaryotes and eukaryotes.   There are several serine proteases whose primary and tertiary structures  bear no discernible relationship to each other or to chymotrypsin.   Nevertheless, these proteins also contain catalytic triads at their active sites  whose structures closely resemble that of chymotrypsin. These enzymes include  subtilisin, an endopeptidase that was originally isolated from Bacillus subtilis,  and wheat germ serine carboxypeptidase II, an exopeptidase. Since the or¬  ders of the corresponding active site residues in the amino acid sequences of  these serine proteases are quite different (Fig. 11-28), it seems highly improb¬  able that they could have evolved from a common ancestor protein. These   FIG. 1 1 -28 Diagram indicating the relative positions of the active site residues  of three unrelated serine proteases. The catalytic triads in subtilisin, chymotrypsin, and  serine carboxypeptidase II each consist of Ser, His, and Asp residues. The peptide  backbones of Ser 214, Trp 215, and Gly 216 in chymotrypsin, and their counterparts in  subtilisin, participate in substrate-binding interactions. [After Robertus, J.D., Alden, R.A., Birktoft,  J.J., Kraut, J., Powers, J.C., and Wilcox, P.E., Biochemistry 11, 2449 (1972).] Q See  Kinemage Exercise 10-2.    General acid catalysis  aids breakdown of the  2 1 tetrahedral intermediate  / to the acyl-enzyme  intermediate.    General base catalysis  and nucleophilic attack  to form tetrahedral (4  intermediate.   General acid  catalysis aids  breakdown of  tetrahedral  intermediate to  the carboxyl  product and the  active enzyme.    FIG. 1 1 -29 The catalytic mechanism of the serine  proteases.    Summarize the roles of Asp 102, His 57, and Ser 195 in this    enzymes apparently constitute a remarkable example of convergent evolution:  Nature seems to have independently discovered the same catalytic mechanism sev¬  eral times.   A catalytic mechanism based on considerable chemical and structural data has  been formulated and is given here in terms of chymotrypsin (Fig. 11-29),  although it applies to all serine proteases and certain other hydrolytic enzymes:   1. After chymotrypsin has bound a substrate, Ser 195 nucleophilically at¬  tacks the scissile peptides carbonyl group to form the tetrahedral  intermediate, which resembles the reactions transition state (covalent  catalysis). X-Ray studies indicate that Ser 195 is ideally positioned to  carry out this nucleophilic attack (proximity and orientation effects).  This nucleophilic attack involves transfer of a proton to the imidazole  ring of His 57, thereby forming an imidazolium ion (general base catal¬  ysis). This process is aided by the polarizing effect of the unsolvated  carboxylate ion of Asp 102, which is hydrogen bonded to His 57 (elec¬  trostatic catalysis). The tetrahedral intermediate has a well-defined, al¬  though transient, existence. We will see that much of chymotrypsins  catalytic power derives from its preferential binding of the transition state  leading to this intermediate (transition state binding catalysis).   2. The tetrahedral intermediate decomposes to the acyl-enzyme inter¬  mediate under the driving force of proton donation from N3 of His  57 (general acid catalysis) facilitated by the polarizing effect of Asp 102  on His 57 (electrostatic catalysis).   3. The amine leaving group (R'NH 2 , the new N-terminal portion of the  cleaved polypeptide chain) is released from the enzyme and replaced  by water from the solvent.   4. The acyl—enzyme intermediate, which is highly susceptible to hy¬  drolytic cleavage, adds water by the reversal of Step 2, yielding a sec¬  ond tetrahedral intermediate.   5. The reversal of Step 1 yields the carboxylate product (the new C-  terminal portion of the cleaved polypeptide chain), thereby regenerat¬  ing the active enzyme. In this process, water is the attacking  nucleophile and Ser 195 is the leaving group.    Serine Proteases Preferentially Bind the Transition State. Detailed com¬  parisons of the X-ray structures of several serine protease-inhibitor complexes  have revealed a further structural basis for catalysis in these enzymes (Fig. 11-30):   1. The conformational rearrangement that occurs with the formation of  the tetrahedral intermediate causes the now anionic carbonyl oxygen of  the scissile peptide to move deeper into the active site so as to occupy a  previously unoccupied position called the oxyanion hole.   2. There, it forms two hydrogen bonds with the enzyme that cannot form  when the carbonyl group is in its normal trigonal conformation. The  two enzymatic hydrogen bond donors were first noted by Joseph Kraut  to occupy corresponding positions in chymotrypsin and subtilisin. He  proposed the existence of the oxyanion hole on the basis of the prem¬  ise that convergent evolution had made the active sites of these unre¬  lated enzymes functionally identical.   3. The tetrahedral distortion, moreover, permits the formation of an oth¬  erwise unsatisfied hydrogen bond between the enzyme and the back¬  bone NH group of the residue preceding the scissile peptide bond.   This preferential binding of the transition state (or the tetrahedral intermediate)  over the enzyme—substrate complex or the acyl—enzyme intermediate is responsible   FIG. 1 1 -30 Transition state stabilization in the serine  proteases, (a) When the substrate binds to the enzyme, the trigonal  carbonyl carbon of the scissile peptide is conformationally constrained  from binding in the oxyanion hole (upper left), (b) In the tetrahedral  intermediate, the now charged carbonyl oxygen of the scissile peptide  (the oxyanion) enters the oxyanion hole and hydrogen bonds to the  backbone NH groups of Gly 193 and Ser 195. The consequent   conformational change permits the NH group of the residue  preceding the scissile peptide bond to form an otherwise unsatisfied  hydrogen bond to Gly 193. Serine proteases therefore preferentially  bind the tetrahedral intermediate. [After Robertus, J.D., Kraut, J., Alden,  R.A., and Birktoft, J.J., Biochemistry 11 , 4302 (1972).]    for much of the catalytic efficiency of serine proteases. Thus, mutating any or all  of the residues in chymotrypsins catalytic triad yields enzymes that still en¬  hance proteolysis by ~5 X 10 4 -fold over the uncatalyzed reaction (versus a  rate enhancement of ~10 10 for the native enzyme). Similarly, the reason that  DIPF is such an effective inhibitor of serine proteases is because its tetrahe¬  dral phosphate group makes this compound a transition state analog.   Low-Barrier Hydrogen Bonds May Stabilize the Transition State. The tran¬  sition state of the chymotrypsin reaction is stabilized not just through the for¬  mation of additional hydrogen bonds in the oxyanion hole but possibly also  by the formation of an unusually strong hydrogen bond. Proton transfers be¬  tween hydrogen bonded groups (D—H***A) occur at physiologically reason¬  able rates only when the p K of the proton donor is no more than 2 or 3 pH  units greater than that of the protonated form of the proton acceptor.  However, when the pK s of the hydrogen bonding donor (D) and acceptor  (A) groups are nearly equal, the distinction between them breaks down: The  hydrogen atom becomes more or less equally shared between them (D***H***A).  Such low-barrier hydrogen bonds (LBHBs) are unusually short and strong.  Their free energies of formation, as measured in the gas phase, are as high as  — 40 to — 80 kj • mol 1 (versus —12 to — 30 kj • mol 1 for normal hydro¬  gen bonds) and they exhibit a D***A bond length of <2.55 A for O—H***0  and <2.65 A for N—H---0 (versus 2.8 to 3.1 A for normal hydrogen bonds).   LBHBs are unlikely to exist in dilute aqueous solution because water mol¬  ecules, which are excellent hydrogen bond donors and acceptors, effectively  compete with D—H and A for hydrogen bonding sites. However, LBHBs may  exist in the nonaqueous active sites of enzymes. In fact, experimental evidence  indicates that in the serine protease catalytic triad, the pK’s of the protonated  His and Asp are nearly equal, and the hydrogen bond between His and Asp  has an unusually short N***0 distance of 2.62 A with the H atom nearly cen¬  tered between the N and O atoms. These findings are consistent with the for¬  mation of an LBHB in the transition state. This suggests that the enzyme uses  the “strategy” of converting a weak hydrogen bond in the initial enzyme-   substrate complex to a strong hydrogen bond in the transition state, thereby Section 5 Serine Proteases   facilitating proton transfer while applying the difference in the free energy be¬  tween the normal and low-barrier hydrogen bonds to preferentially binding  the transition state.   Although several studies have revealed the existence of unusually short hy¬  drogen bonds in enzyme active sites, it is far more difficult to demonstrate ex¬  perimentally that they are unusually strong, as LBHBs are predicted to be. In  fact, several studies of the strengths of unusually short hydrogen bonds in or¬  ganic model compounds in nonaqueous solutions suggest that these hydrogen  bonds are not unusually strong. Consequently, a lively debate has ensued as  to the catalytic significance of LBHBs. However, if enzymes do not form  LBHBs, it remains to be explained how the conjugate base of an acidic group  (e.g., Asp 102) partially abstracts a proton from a far more basic group (e.g.,   The Tetrahedral Intermediate Resembles the Complex of Trypsin with  Trypsin Inhibitor. Perhaps the most convincing structural evidence for the ex¬  istence of the tetrahedral intermediate was provided by Robert Huber in an  X-ray study of the complex between bovine pancreatic trypsin inhibitor   (BPTI) and trypsin. The 58-residue BPTI binds to the active site region of  trypsin to form a complex with a tightly packed interface and a network of  hydrogen-bonded cross-links. This interaction prevents any trypsin that is pre¬  maturely activated in the pancreas from digesting that organ (Section 11-5D).   The portion of BPTI in contact with the trypsin active site resembles  bound substrate. A specific Lys side chain of BPTI occupies the trypsin  specificity pocket (Fig. 11-31 a), and the inhibitor’s Lys-Ala peptide bond is  positioned as if it were the scissile peptide bond (Fig. 11-31 b). What is most  remarkable about the BPTI-trypsin complex is that its conformation is well  along the reaction coordinate toward the tetrahedral intermediate: The side  chain oxygen of trypsin Ser 195, the active Ser, is in closer-than-van der    FIG. 11-31 The trypsin-BPTI complex, (a) The X-ray structure is shown as a  cutaway model indicating how trypsin (red) binds BPTI (green). The green protrusion  extending into the red cavity near the center of the figure represents the inhibitor’s Lys 15  side chain occupying trypsin’s specificity pocket. Note the close complementary fit of the  two proteins. [Courtesy of Michael Connolly, New York University.] (b) Trypsin Ser 195 is in  closer-than-van der Waals contact with the carbonyl carbon of BPTI’s scissile peptide,  which is pyramidally distorted toward Ser 195. The normal proteolytic reaction is apparently  arrested somewhere along the reaction coordinate preceding the tetrahedral intermediate.  Q See Kinemage Exercise 10-1.   Chapter 11 Enzymatic Catalysis Waals contact (2.6 A) with the pyramidally distorted carbonyl carbon of   BPTI’s “scissile” peptide. However, the proteolytic reaction cannot proceed  past this point because of the rigidity of the complex and because it is so  tightly sealed that the leaving group cannot leave and water cannot enter the  reaction site.   Protease inhibitors are common in nature, where they have protective and  regulatory functions. For example, certain plants release protease inhibitors in  response to insect bites, thereby causing the offending insect to starve by in¬  activating its digestive enzymes. Protease inhibitors constitute ~10% of the  blood plasma proteins. For instance, a x -proteinase inhibitor, which is se¬  creted by the liver, inhibits leukocyte elastase (leukocytes are white blood  cells; the action of leukocyte elastase is thought to be part of the inflamma¬  tory process). Pathological variants of a r -proteinase inhibitor with reduced ac¬  tivity are associated with pulmonary emphysema, a degenerative disease of  the lungs resulting from the hydrolysis of its elastic fibers. Smokers also suf¬  fer from reduced activity of their a r -proteinase inhibitor because smoking ox¬  idizes a required Met residue.   The Tetrahedral Intermediate Has Been Directly Observed. Because the  tetrahedral intermediate resembles the transition state of the serine protease  reaction, it is thought to be short-lived and unstable. However, a series of X-  ray structures of porcine pancreatic elastase with a peptide substrate have  shown the progress of the reaction from the acyl-enzyme intermediate stage  to the release of product. This second phase of the proteolysis reaction in¬  cludes a tetrahedral intermediate (Fig. 11-29).   This acyl—enzyme complex, which is stable at pH 5.0, exhibits the ex¬  pected structure, with the substrates C-terminal lie residue covalently linked  via an ester bond to Ser 195 (Fig. 11-32^). In this first view of a serine pro-   FIG. 1 1 -32 Structure of the acyl-enzyme and tetrahedral  intermediates. Porcine pancreatic elastase was incubated with a  heptapeptide substrate (YPFVEPI, using the one-letter code). Only  residues 4-7 are visible. The protease residues are specified by the  three-letter code. Atoms are colored according to type with elastase C  green, substrate C cyan, N blue, 0 red, and S yellow, (a) At pH 5.0, a  covalent bond ( magenta ) links the Ser 195 0 atom to the C-terminal  (17) C atom of the substrate. A water molecule (orange sphere )  appears poised to nucleophilically attack the acyl-enzyme’s carbonyl C    atom. The dashed lines represent catalytically important hydrogen  bonds, and the dotted line indicates the trajectory that the water  molecule presumably follows in nucleophilically attacking the acyl  group’s carbonyl C atom, (b) When the complex is brought to pH 9.0  and then rapidly frozen, the water molecule becomes a hydroxyl  substituent ( orange ) to the carbonyl C atom, thereby yielding the  tetrahedral intermediate, which resembles the transition state. [Based  on X-ray structures by Christopher Schofield and Janos Hadju,  University of Oxford, U.K. PDBids (a) 1 HAX and (b) 1 HAZ]   tion toward a tetrahedral geometry A water molecule is located near the in¬  termediates ester bond, hydrogen-bonded to His 57, where it appears poised  to nucleophilically attack the ester linkage. At pH 5.0, His 57 is protonated  and acts as a hydrogen bond donor to water (it cannot function as a base cat¬  alyst at this pH).   To assess the next step of the reaction, the acyl-enzyme crystal was im¬  mersed in a solution of pH 9.0 (recall that protein crystals contain large solvent-  filled spaces, so protons and other small substances can diffuse into and out  of the crystallized enzyme’s active site; Section 6-2A). The resulting de¬  protonation of His 57 at pH 9.0 triggered the hydrolytic reaction (Step 4 of  Fig. 11-29). After a period of 1 or 2 minutes, the crystals were frozen in liq¬  uid nitrogen to halt the reaction so that the X-ray structure of the enzyme  complex could be determined. In this way, the tetrahedral intermediate was  trapped and observed (Fig. 11-32^).   During formation of the tetrahedral intermediate, the oxyanion hole does  not undergo any change in its structure, but the peptide substrate moves  within its binding pocket and becomes distorted toward a tetrahedral geome¬  try. The tetrahedral intermediate has the expected shape, similar to known  transition state analog inhibitors. However, it does not bind so tightly to the  amide group of the oxyanion hole (Fig. 11-30) that it would not be able to  subsequently dissociate.   Proteolytic enzymes are usually biosynthesized as somewhat larger inactive pre¬  cursors known as zymogens (enzyme precursors, in general, are known as  proenzymes). In the case of digestive enzymes, the reason for this is clear: If  these enzymes were synthesized in their active forms, they would digest the  tissues that synthesized them. Indeed, acute pancreatitis, a painful and some¬  times fatal condition that can be precipitated by pancreatic trauma, is charac¬  terized by the premature activation of the digestive enzymes synthesized by  that organ.   The activation of trypsinogen, the zymogen of trypsin, occurs when  trypsinogen enters the duodenum from the pancreas. Enteropeptidase, a  serine protease whose secretion from the duodenal mucosa is under hor¬  monal control, excises the N-terminal hexapeptide from trypsinogen by  specifically cleaving its Lys 15—lie 16 peptide bond (Fig. 11-33). Since this  activating cleavage occurs at a trypsin-sensitive site (recall that trypsin cleaves  after Arg and Lys residues), the small amount of trypsin produced by en¬  teropeptidase also catalyzes trypsinogen activation, generating even more  trypsin, etc. Thus, trypsinogen activation is said to be auto catalytic.   FIG. 1 1 -33 The activation of trypsinogen to trypsin. Proteolytic excision of the  N-terminal hexapeptide is catalyzed by either enteropeptidase or trypsin. The  chymotrypsinogen residue-numbering system is used here; that is, Val 10 is actually  trypsinogen’s N-terminus and lie 16 is trypsin’s N-terminus.    When a blood vessel is damaged, a clot forms as a result of the  aggregation of platelets (small enucleated blood cells) and the forma¬  tion of an insoluble fibrin network that traps additional blood cells.    Fibrin is produced from the soluble circulating protein fibrinogen  through the action of the serine protease thrombin. Thrombin is  the last in a series of coagulation enzymes that are sequentially  activated by proteolysis of their zymogen forms. The overall process  is known as the coagulation cascade (opposite), although experi¬  mental evidence shows that the pathway is not strictly linear, as  the waterfall analogy might suggest.   The various components of the coagulation cascade, which in¬  clude enzymes as well as nonenzymatic protein cofactors, are as¬  signed Roman numerals, largely for historical reasons that do not  reflect their order of action in vivo. The suffix a denotes an active  factor. The catalytic domains of the coagulation proteases resemble  trypsin in sequence and mechanism but are much more specific for  their substrates. Additional domains mediate interactions with cofac¬  tors and help anchor the proteins to the platelet membrane, which  serves as a stage for many of the coagulation reactions.   Coagulation is initiated when a membrane protein (tissue  factor) exposed to the bloodstream by tissue damage forms a    complex with circulating factor VII or Vila (factor Vila is gener¬  ated from factor VII by trace amounts of other coagulation pro¬  teases, including factor Vila itself). The tissue factor-Vlla complex  proteolytically converts the zymogen factor X to factor Xa. Factor  Xa then converts prothrombin to thrombin, which subsequently  cleaves fibrinogen to form fibrin. The tissue factor-dependent  steps of coagulation are known as the extrinsic pathway be¬  cause the source of tissue factor is extravascular. The extrinsic  pathway is quickly dampened through the action of a protein that  inhibits factor VII once factor Xa has been generated.   Sustained thrombin activation requires the activity of the intrin¬  sic pathway (so named because all its components are present in  the circulation). The intrinsic pathway is stimulated by the tissue  factor-Vlla complex, which converts factor IX to its active form,  factor IXa. The ensuing thrombin activates a number of compo¬  nents of the intrinsic pathway, including factor XI, a protease that  activates factor IX, to maintain coagulation in the absence of  tissue factor or factor Vila. Thrombin also activates factors V and  VIII, which are cofactors rather than proteases. Factor Va pro¬  motes prothrombin activation by factor Xa by as much as 20,000-  fold, and factor Villa promotes factor X activation by factor IXa by  a similar amount. Thus, thrombin promotes its own activation  through a feedback mechanism that amplifies the preceding steps  of the cascade. Factor XIII is also activated by thrombin. Factor  XII la, which is not a serine protease, chemically cross-links fibrin  molecules through formation of peptide bonds between glutamate  and lysine side chains, which forms a strong fibrin network.   The intrinsic pathway of coagulation can be triggered by expo¬  sure to negatively charged surfaces such as glass. Consequently,  blood clots when it is collected in a clean glass test tube. In the  absence of tissue factor, a fibrin clot may not appear for several  minutes, but when tissue factor is present, a clot forms within a  few seconds. This suggests that rapid blood clotting in vivo re¬  quires tissue factor as well as the proteins of the intrinsic path¬  way. Additional evidence for the importance of the extrinsic  pathway is that individuals who are deficient in factor VII tend to  bleed excessively. Abnormal bleeding also results from congenital    Chymotrypsinogen is then activated by trypsin-catalyzed cleavage of its Arg  15-Ile 16 peptide bond.   Proelastase, the zymogen of elastase, is activated by a single tryptic cleav¬  age that excises a short N-terminal peptide. Trypsin also activates pancreatic  procarboxypeptidases A and B and prophospholipase A 2 (Section 9-1C).  The autocatalytic nature of trypsinogen activation and the fact that trypsin ac¬  tivates other hydrolytic enzymes makes it essential that trypsinogen not be ac¬  tivated in the pancreas. We have seen that the all but irreversible binding of  trypsin inhibitors such as BPTI to trypsin is a defense against trypsinogen s  inappropriate activation.   Sequential proenzyme activation makes it possible to quickly generate  large quantities of active enzymes in response to diverse physiological signals.  For example, the serine proteases that lead to blood clotting are synthesized  as zymogens by the liver and circulate until they are activated by injury to a  blood vessel (Box 11-4).   The sequential activation of zymogens in the  coagulation cascade leads to a burst of thrombin  activity, since trace amounts of factors Vila, IXa,  and Xa can activate much larger amounts of  their respective substrates. The potential for  amplification in the coagulation cascade is re¬  flected in the plasma concentrations of the  coagulation proteins (see table).    ^Concentrations calculated from data in High, K.A.  and Roberts, H.R. (Eds.), Molecular Basis of  Thrombosis and Hemostasis, Marcel Dekker (1995).    Perhaps not surprisingly, thrombin eventually triggers mecha¬  nisms that shut down clot formation, thereby limiting the duration  of the clotting process and hence the extent of the clot. Such  control of clotting is of extreme physiological importance since the  formation of even one inappropriate blood clot within an individ¬  ual’s lifetime may have fatal consequences.    Zymogens Have Distorted Active Sites. Since the zymogens of trypsin, chy-  motrypsin, and elastase have all their catalytic residues, why aren’t they enzy¬  matically active? Comparisons of the X-ray structures of trypsinogen with that  of trypsin, and of chymotrypsinogen with that of chymotrypsin, show that on  activation, the newly liberated N-terminal lie 16 residue moves from the sur¬  face of the protein to an internal position, where its free cationic amino group  forms an ion pair with the invariant anionic Asp 194, which is close to the cat¬  alytic triad (Fig. 11-26). Without this conformational change, the enzyme can¬  not properly bind its substrate or stabilize the tetrahedral intermediate because  its specificity pocket and oxyanion hole are improperly formed. This provides  further structural evidence favoring the role of preferential transition state bind¬  ing in the catalytic mechanism of serine proteases. Nevertheless, because their  catalytic triads are structurally intact, the zymogens of serine proteases actually  have low levels of enzymatic activity, an observation that was made only after  the above structural comparisons suggested that this might be the case.   • What are some ways that active site  residues can be identified?   • Summarize the roles of the residues that  make up the catalytic triad of serine pro¬  teases.   • What role do low-barrier hydrogen bonds  play in serine protease catalysis?   • Enzymes, almost all of which are proteins, are grouped into six  mechanistic classes.   • Enzymes accelerate reactions by factors of up to at least 10 15 .   • The substrate specificity of an enzyme depends on the geometric  and electronic character of its active site.   • Some enzymes catalyze reactions with the assistance of metal ion  cofactors or organic coenzymes that function as reversibly bound  cosubstrates or as permanently associated prosthetic groups. Many  coenzymes are derived from vitamins.   • Enzymes catalyze reactions by facilitating a reaction pathway with  a lower activation free energy, A G*, which is the free energy re¬  quired to reach the transition state, the point of highest free en¬  ergy in the reaction.   • Enzymes use the same catalytic mechanisms employed by chemi¬  cal catalysts, including general acid and general base catalysis, co¬  valent catalysis, and metal ion catalysis.   • The arrangement of functional groups in an enzyme active site al¬  lows catalysis by proximity and orientation effects as well as elec¬  trostatic catalysis.    • A particularly important mechanism of enzyme-mediated cataly¬  sis is the preferential binding of the transition state of the cat¬  alyzed reaction.   • In the catalytic mechanism of lysozyme, Glu 35 in its protonated  form acts as an acid catalyst to cleave the polysaccharide substrate  between its D and E rings, and Asp 52 in its anionic state forms  a covalent bond to Cl of the D ring.   • The reaction is facilitated by the distortion of residue D to the  planar half-chair conformation, which resembles the reaction’s ox-  onium ion transition state.   • Serine proteases contain a Ser-His-Asp catalytic triad near a bind¬  ing pocket that helps determine the enzymes’ substrate specificity.   • Catalysis in the serine proteases occurs through acid-base cataly¬  sis, covalent catalysis, proximity and orientation effects, electro¬  static catalysis, and by preferential transition state binding in the  oxyanion hole.   • Synthesis of pancreatic proteases as inactive zymogens protects the  pancreas from self-digestion. Zymogens are activated by specific  proteolytic cleavages.   (b) It increases the rate at which a chemical reaction approaches  equilibrium relative to its uncatalyzed rate.   2. What is the relationship between the rate of an enzyme-catalyzed  reaction and the rate of the corresponding uncatalyzed reaction?  Do enzymes enhance the rates of slow uncatalyzed reactions as  much as they enhance the rates of fast uncatalyzed reactions?    3. Which type of enzyme (Table 11-2) catalyzes the following  reactions?   4. Which type of enzyme (Table 11-2) catalyzes the following  reactions?   5. On the free energy diagram shown, label the intermediate(s) and  transition state(s). Is the reaction thermodynamically favorable?   6. Draw a transition state diagram of (a) a nonenzymatic reaction  and the corresponding enzyme-catalyzed reaction in which (b) S  binds loosely to the enzyme and (c) S binds very tightly to the  enzyme. Compare AG^ for each case. Why is tight binding of S  not advantageous?   7. Approximately how much does staphylococcal nuclease (Table 11-1)  decrease the activation free energy (A G*) of its reaction (the hy¬  drolysis of a phosphodiester bond) at 25°C?   8. Calculate the rate enhancement that could be accomplished by  an enzyme forming one low-barrier hydrogen bond with its tran¬  sition state at 25°C.   9. Using the reaction shown in Box 11-1 (the attack of an amine  on the carbonyl group of a ketone) as a starting point, draw  curved arrows to represent the base-catalyzed reaction (when the  group —B: is present).   10. Using the reaction shown in Box 11-1 (the attack of an amine  on the carbonyl group of a ketone) as a starting point, draw  curved arrows to represent the acid-catalyzed reaction (when the  group —A—H is present).   11. Explain why enzyme activity varies with temperature, as shown  here.   12. The covalent catalytic mechanism of an enzyme depends on a  single active site Cys whose p K is 8. A mutation in a nearby  residue alters the microenvironment so that this p K increases  to 10. Would the mutation cause the reaction rate to increase or  decrease? Explain.   13. Studies at different pH’s show that an enzyme has two catalyti-  cally important residues whose p K’s are ~4 and — 10. Chemical  modification experiments indicate that a Glu and a Lys residue  are essential for activity. Match the residues to their piCs and  explain whether they are likely to act as acid or base catalysts.   14. Explain why RNase A cannot catalyze the hydrolysis of DNA.   13. Urease, the first enzyme to be crystallized, is inhibited in the pres¬  ence of Hg, Cd, or Co ions. What does this information suggest  about the catalytic mechanism of urease?   16. What feature of RNA would allow it to function as a ribozyme?  Why are there no naturally occurring DNA enzymes?   18. Suggest a transition state analog for proline racemase that differs  from those discussed in the text. Justify your suggestion.   19. Explain why lysozyme cleaves the artificial substrate (NAG )4  —4000 times more slowly than it cleaves (NAG) 6 .   20. Would you expect lysozyme to hydrolyze cellulose? Why or  why not?   21. Lysozyme residues Asp 101 and Arg 114 are required for efficient  catalysis, although they are located at some distance from the ac¬  tive site Glu 33 and Asp 52. Substituting Ala for either Asp 101  or Arg 114 does not significantly alter the enzyme’s tertiary struc¬  ture, but it significantly reduces its catalytic activity. Explain.   22. Predict the effect on lysozyme’s activity of mutating Glu 35 to  Asp and Asp 52 to Glu.   24. Predict the effect of mutating Asp 102 of trypsin to Asn (a) on  substrate binding and (b) on catalysis.   25. Diagram the hydrogen-bonding interactions of the catalytic triad  His-Lys-Ser during catalysis in a hypothetical hydrolytic enzyme.   26. The comparison of the active site geometries of chymotrypsin  and subtilisin under the assumption that their similarities have  catalytic significance has led to greater mechanistic understand¬  ing of both these enzymes. Discuss the validity of this strategy.   27. Under certain conditions, peptide bond formation rather than  peptide bond hydrolysis is thermodynamically favorable. Would  you expect chymotrypsin to catalyze peptide bond formation?   28. Why is the broad substrate specificity of chymotrypsin advanta¬  geous in vivo ? Why would this be a disadvantage for some other  proteases?   29. Tofu (bean curd), a high-protein soybean product, is prepared in  such a way as to remove the trypsin inhibitor present in soybeans.  Explain the reason(s) for this treatment.   30. Many of the cell’s hydrolytic enzymes are located in the lysosome,  where the pH is ~5. What would you expect to be the optimum  pH for these enzymes, and why would this protect the rest of the  cell from the destructive power of these enzymes upon the acci¬  dental rupture of a lysosome?   31. A genetic defect in coagulation factor IX causes hemophilia b, a  disease characterized by a tendency to bleed profusely after very  minor trauma. However, a genetic defect in coagulation factor XI  has only mild clinical symptoms. Explain this discrepancy in  terms of the mechanism for activation of coagulation proteases  shown in Box 11-4.   Case Study 11 Nonenzymatic Deamidation of  Asparagine and Glutamine Residues in Proteins  Focus concept: Factors influencing nonenzymatic hydrolytic deami¬  dation of Asn and Gin residues in proteins are examined and pos¬  sible mechanisms for the reactions are proposed.    Eukaryotic cells contain a class  of proteases known as caspases. Which catalytic residues give  these proteases their name? What role do these residues play in  catalysis, and how does the caspase mechanism compare to the  mechanism of serine proteases? How are the caspases activated  and what is the result of their activity in the cell?   Benkovic, S.J. and Hammes-Schiffer, S., A perspective on enzyme catal¬  ysis, Science 301, 1196-1202 (2003). [Includes a history of some of  the theories and experiments on catalysis.]   Bruice, T.C. and Benkovic, S.J., Chemical basis for enzyme catalysis, Bio¬  chemistry 39, 6267-6274 (2000).   Gerlt, J.A., Protein engineering to study enzyme catalytic mechanisms,  Curr. Opin. Struct. Biol. 4, 393-600 (1994). [Describes how infor¬  mation can be gained from mutagenesis and structural analysis of  enzymes.]   Hackney, D.D., Binding energy and catalysis, in Sigman, D.S. and Boyer,  P.D. (Eds.), The Enzymes (3rd ed.), Vol. 19, pp. 1-36, Academic  Press (1990).   Kraut, J., How do enzymes work? Science 242, 333-340 (1988). [A brief  and very readable review of transition state theory and applications.]   Schramm, V.L., Enzymatic transition states and transition state analogues,  Curr. Opin. Struct. Biol. 15, 604-613 (2005).   Tipton, K.E, The naming of parts, Trends Biochem. Sci. 18, 113-115  (1993). [A discussion of the advantages of a consistent naming  scheme for enzymes and the difficulties of formulating one.]   Kirby, A.J., The lysozyme mechanism sorted—after 50 years, Nature  Struct. Biol. 8, 737-739 (2001). [Briefly summarizes the theoretical  and experimental evidence for a covalent intermediate in the  lysozyme mechanism.]   McKenzie, H.A. and White, EH., Jr., Lysozyme and a-lactalbumin:  Structure, function and interrelationships, Adv. Protein Chem. 41,  173-315 (1991).   Strynadka, N.C.J. and James, M.N.G., Lysozyme revisited: crystallo¬  graphic evidence for distortion of an A^acetylmuramic acid residue  bound in site D, J. Mol. Biol. 220, 401-424 (1991).    Wolfenden, R., Benchmark reaction rates, the stability of biological  molecules in water, and the evolution of catalytic power in enzymes,  Annu. Rev. Biochem. 80, 645-667 (2011).   Cleland, W.W., Frey, P.A., and Gerlt, J.A., The low barrier hydrogen bond  in enzymatic catalysis, J. Biol. Chem. 273, 25529-25532 (1998).   Davie, E.W., Biochemical and molecular aspects of the coagulation cas¬  cade, Thromb. Haemost. 74, 1-6 (1995). [A brief review by one of  the pioneers of the cascade hypothesis.]   Fersht, A., Structure and Mechanism in Protein Science. Freeman (1999).  [Includes detailed reaction mechanisms for chymotrypsin and other  enzymes.]   Perona, J.J. and Craik, C.S., Evolutionary divergence of substrate speci¬  ficity within the chymotrypsin-like protease fold, J. Biol. Chem. 272,  29987-29990 (1997). [Summarizes research identifying the struc¬  tural basis of substrate specificity in chymotrypsin and related en¬  zymes.]   Radisky, E.S., Lee, J.M., Lu, C-J.K., and Koshland, D.E., Jr., Insights  into the serine protease mechanism from atomic resolution struc¬  tures of trypsin reaction intermediates, Proc. Natl. Acad. Sci. 103,  6835-6840 (2006). [Superpositions of X-ray structures representing  the enzyme-substrate complex, tetrahedral intermediate, and  acyl-enzyme intermediate illustrate the progress of the reaction.]   Wilmouth, R.C., Edman, K., Neutze, R., Wright, P.A., Clifton, I.J.,  Schneider, T.R., Schofield, C.J., and Hajdu, J., X-Ray snapshots of  serine protease catalysis reveal a tetrahedral intermediate, Nature  Struct. Biol. 8, 689-694 (2001). [Reports the first structural evidence  for a tetrahedral intermediate in the hydrolysis reaction catalyzed by  a serine protease.]    Atorvastatin (Lipitor®), one of the most commonly prescribed drugs, binds to and  inhibits the activity of the enzyme HMG-CoA reductase in a way that can be quanti¬  fied through enzyme kinetics. [Based on an X-ray structure by Johann Deisenhofer.  PDBid 1HWKJ    Early enzymologists, often working with crude preparations of yeast or liver  cells, could do little more than observe the conversion of substrates to prod¬  ucts catalyzed by as yet unpurified enzymes. Measuring the rates of such re¬  actions therefore came to be a powerful tool for characterizing enzyme activity.  The application of simple mathematical models to enzyme activity under vary¬  ing laboratory conditions, and in the presence of competing substrates or en¬  zyme inhibitors, made it possible to deduce the probable physiological  functions and regulatory mechanisms of various enzymes.   The study of enzymatic reaction rates, or enzyme kinetics, is no less im¬  portant now than it was early in the twentieth century. In many cases, the rate  of a reaction and how the rate changes in response to different conditions re¬  veal the path followed by the reactants and are therefore indicative of the re¬  action mechanism. Kinetic data, combined with detailed information about  an enzymes structure and its catalytic mechanisms, provide some of the most  powerful clues to the enzymes biological function and may suggest ways to  modify it for therapeutic purposes.   We begin our consideration of enzyme kinetics by reviewing chemical ki¬  netics. Following that, we derive the basic equations of enzyme kinetics and  describe the effects of inhibitors on enzymes. We also consider some examples  of enzyme control that highlight several aspects of enzyme function. Finally,  we describe some practical applications of enzyme inhibition, the development  of enzyme inhibitors as drugs.   B Enzyme Kinetics Often Follows the  Michaelis-Menten Equation  C Kinetic Data Can Provide Values of l/ max  and K m   A Competitive Inhibition Involves Inhibitor Binding  at an Enzyme’s Substrate Binding Site  B Uncompetitive Inhibition Involves Inhibitor  Binding to the Enzyme-Substrate Complex  C Mixed Inhibition Involves Inhibitor Binding to  Both the Free Enzyme and the  Enzyme-Substrate Complex   A Allosteric Control Involves Binding at a Site  Other than the Active Site  B Control by Covalent Modification Usually  Involves Protein Phosphorylation   A Drug Discovery Employs a Variety of Techniques  B A Drug’s Bioavailability Depends on How It Is  Absorbed and Transported in the Body  C Clinical Trials Test for Efficacy and Safety  D Cytochromes P450 Are Often Implicated in  Adverse Drug Reactions    Simple rate equations describe the progress of first-order and second-order reactions.  The Michaelis-Menten equation relates the initial velocity of a reaction to the maximal  reaction velocity and the Michaelis constant for a particular enzyme and substrate.   An enzyme’s overall catalytic efficiency is expressed as k cat /K M .   A Lineweaver-Burk plot can be used to present kinetic data and to calculate  values for K M and l/ max .   Bisubstrate reactions can occur by an Ordered or Random sequential mechanism  or by a Ping Pong mechanism.   Chapter 12 Enzyme Kinetics, Inhibition, and Control Kinetic measurements of enzymatically catalyzed reactions are among the most   powerful techniques for elucidating the catalytic mechanisms of enzymes.  Enzyme kinetics is a branch of chemical kinetics, so we begin this section by  reviewing the principles of chemical kinetics.   where A represents reactants and P represents products, may actually occur  through a sequence of elementary reactions (simple molecular processes) such  as   Here, I x and I 2 symbolize intermediates in the reaction. Each elementary re¬  action can be characterized with respect to the number of reacting species and  the rate at which they interact. Descriptions of each elementary reaction collec¬  tively constitute the mechanistic description of the overall reaction process. Even a  complicated enzyme-catalyzed reaction can be analyzed in terms of its com¬  ponent elementary reactions.    Reaction Order Indicates the Number of Molecules Participating in an  Elementary Reaction. At constant temperature, the rate of an elementary re¬  action is proportional to the frequency with which the reacting molecules come  together. The proportionality constant is known as a rate constant and is sym¬  bolized k. For the elementary reaction A—> P, the instantaneous rate of  appearance of product or disappearance of reactant, which is called the velocity  ( v) of the reaction, is    Determine the velocity of the elementary  reaction X + Y —> Z when the sample con¬  tains 3 |xM X and 3 |xM Y and k for the  reaction is 400 M 1 • s l .    Use Equation 12-3 and make sure that all  units are consistent:   = (400 M -1 • S -1 )(3 X 10 -6 M)   = 6 X 10 -9 M • s -1  = 6 nM • s 1    In other words, the reaction velocity at any time point is proportional to the  concentration of the reactant A. This is an example of a first-order reaction.  Since the velocity has units of molar per second (M • s x ), the first-order rate  constant must have units of reciprocal seconds (s l ). The reaction order of  an elementary reaction corresponds to the molecularity of the reaction ., which is  the number of molecules that must simultaneously collide to generate a product.  Thus, a first-order elementary reaction is a unimolecular reaction.   Consider the elementary reaction 2A —> P. This bimolecular reaction is a  second-order reaction, and its instantaneous velocity is described by    In this case, the reaction velocity is proportional to the square of the concen¬  tration of A, and the second-order rate constant k has units of M 1 • s 1 .   The bimolecular reaction A + B —» P is also a second-order reaction with  an instantaneous velocity described by   Here, the reaction is said to be first order in [A] and first order in [B] (see  Sample Calculation 12-1). Unimolecular and bimolecular reactions are  common. Termolecular reactions are unusual because the simultaneous col¬  lision of three molecules is a rare event. Fourth- and higher-order reactions  are unknown.    A Rate Equation Indicates the Progress of a Reaction as a Function of  Time. A rate equation can be derived from the equations that describe the  instantaneous reaction velocity. Thus, a first-order rate equation is obtained by  rearranging Eq. 12-1   Equation 12-6 is a linear equation of the form y — mx + b and can be plot¬  ted as in Fig. 12-1. Therefore, if a reaction is first order, a plot of In [A] ver¬  sus t will yield a straight line whose slope is — k (the negative of the first-order  rate constant) and whose intercept on the In [A] axis is ln[A] 0 .   One of the hallmarks of a first-order reaction is that the time for half of  the reactant initially present to decompose, its half-time or half-life , *i/ 2 > I s a  constant and hence independent of the initial concentration of the reactant. This  is easily demonstrated by substituting the relationship [A] = [A] 0 /2 when  t ~ h /2 i nto Eq. 12-6 and rearranging:    FIG. 12-1 A plot of a first-order rate  equation. The slope of the line obtained when  In [A] is plotted against time gives the rate  constant k.    Substances that are inherently unstable, such as radioactive nuclei, decompose  through first-order reactions (see Sample Calculation 12-2).   In a second-order reaction with one type of reactant, 2A —> P, the varia¬  tion of [A] with time is quite different from that in a first-order reaction.  Rearranging Eq. 12-2 and integrating it over the same limits used for the first-  order reaction yields    Equation 12-11 is a linear equation in terms of the variables 1 / [A] and t. The  halftime for a second-order reaction is expressed t x / 2 — \/ k[A\ Q and therefore,  in contrast to a first-order reaction, depends on the initial reactant concentration.  Equations 12-6 and 12-11 may be used to distinguish a first-order from a  second-order reaction by plotting In [A] versus t and l/[A] versus t and  observing which, if any, of these plots is linear.    The decay of a hypothetical radioisotope  has a rate constant of 0.01 s \ How much  time is required for half of a 1-g sample of  the isotope to decay?    The units of the rate constant indicate a  first-order process. Thus, the half-life is in¬  dependent of concentration. The half-life  of the isotope (the half-time for its decay)  is given by Eq. 12-9:   Chapter 12 Enzyme Kinetics, Inhibition, and Control To experimentally determine the rate constant for the second-order reac¬   tion A + B —» P, it is often convenient to increase the concentration of one  reactant relative to the other, for example, [B] » [A]. Under these condi¬  tions, [B] does not change significantly over the course of the reaction. The  reaction rate therefore depends only on [A], the concentration of the reactant  that is present in limited amounts. Hence, the reaction appears to be first or¬  der with respect to A and is therefore said to be a pseudo-first-order reac¬  tion. The reaction is first order with respect to B when [A] [B].   Enzymes catalyze a tremendous variety of reactions using different combina¬  tions of five basic catalytic mechanisms (Section 11-3). Some enzymes act on  only a single substrate molecule; others act on two or more different substrate  molecules whose order of binding may or may not be obligatory. Some en¬  zymes form covalently bound intermediate complexes with their substrates;  others do not. Yet all enzymes can be analyzed such that their reaction rates as  well as their overall efficiency can be quantified.   The study of enzyme kinetics began in 1902 when Adrian Brown investi¬  gated the rate of hydrolysis of sucrose by the yeast enzyme (i-fructofuranosidase:   Brown found that when the sucrose concentration is much higher than that  of the enzyme, the reaction rate becomes independent of the sucrose concen¬  tration; that is, the rate is zeroth order with respect to sucrose. He therefore  proposed that the overall reaction is composed of two elementary reactions in  which the substrate forms a complex with the enzyme that subsequently de¬  composes to products, regenerating enzyme:   E + S ES -> P + E [ 12 - 12 ]   Here E, S, ES, and P symbolize the enzyme, substrate, enzyme-substrate  complex, and products, respectively. According to this model, when the sub¬  strate concentration becomes high enough to entirely convert the enzyme to  the ES form, the second step of the reaction becomes rate limiting and the  overall reaction rate becomes insensitive to further increases in substrate con¬  centration.   Each of the elementary reactions that make up the above enzymatic reac¬  tion is characterized by a rate constant: k x and k- X are the forward and re¬  verse rate constants for formation of the ES complex (the first reaction), and  k 2 is the rate constant for the decomposition of ES to P (the second reaction).  Here we assume, for the sake of mathematical simplicity, that the second  reaction is irreversible; that is, no P is converted back to S.   The Michaelis-Menten Equation Assumes that ES Maintains a Steady  State. The Michaelis-Menten equation describes the rate of the enzymatic re¬  action represented by Eq. 12-12 as a function of substrate concentration. In  this kinetic scheme, the formation of product from ES is a first-order process.  Thus, the rate of formation of product can be expressed as the product of the  rate constant of the reaction yielding product and the concentration of its im¬  mediately preceding intermediate. The general expression for the velocity (rate)  of Reaction 12-12 is therefore   The overall rate of production of ES is the difference between the rates of   the elementary reactions leading to its appearance and those resulting in its  disappearance:    This equation cannot be explicitly integrated, however, without simplifying as¬  sumptions. Two possibilities are   1. Assumption of equilibrium. In 1913, Leonor Michaelis and Maud  Menten, building on the work of Victor Henri, assumed that  k-i k 2 , so that the first step of the reaction reaches equilibrium:    Here K s is the dissociation constant of the first step in the enzymatic  reaction. With this assumption, Eq. 12-14 can be integrated. Although  this assumption is often not correct, in recognition of the importance  of this pioneering work, the enzyme—substrate complex, ES, is known   2. Assumption of steady state. Figure 12-2 illustrates the progress curves  of the various participants in Reaction 12-12 under the physiologically  common condition that substrate is in great excess over enzyme  ([S] [E]). With the exception of the initial stage of the reaction,   which is usually over within milliseconds of mixing E and S, [ES]  remains approximately constant until the substrate is nearly exhausted.  Hence, the rate of synthesis of ES must equal its rate of consumption  over most of the course of the reaction. In other words, ES maintains  a steady state and [ES] can be treated as having a constant value:   This so-called steady state assumption, a more general condition than  that of equilibrium, was first proposed in 1925 by George E. Briggs  and John B.S. Haldane (Box 12-1).   FIG. 1 2-2 The progress curves for a  simple enzyme-catalyzed reaction. With the  exception of the initial phase of the reaction  (before the darker block), the slopes of the  progress curves for [E] and [ES] are essentially  zero as long as [S] » [E] (within the darker  block). [After Segel, I.H., Enzyme Kinetics, p. 27,  Wiley (1993).] O See the Animated Figures.   J.B.S. Haldane (1892-1964) John  Burdon Sanderson Haldane, the son of a  prominent physiologist, was a gifted scientist  and writer whose major contributions include  the application of mathematics to areas of  biology such as genetics and enzyme  kinetics. As a scientist as well as a  philosopher, he was aware of developments  in relativity theory and quantum mechanics and was influenced by  a practical philosophy that the natural world obeyed the laws of  logic and arithmetic.   When Haldane published his book Enzymes in 1930, the idea  that enzymes are proteins, rather than small catalysts surrounded  by an amorphous protein “colloid,” was still controversial. However,  even in the absence of structural information, scientists such as  Leonor Michaelis and Maud Menten had already applied the  principles of thermodynamics to derive some basic equations  related to enzyme kinetics. Michaelis and Menten proposed in  1913 that during a reaction, an enzyme and its substrate are in  equilibrium with a complex of enzyme and substrate. In 1925,  Haldane argued that this was not strictly true, since some  enzyme-substrate complex does not dissociate to free enzyme  and substrate but instead goes on to form product. When the  enzyme and substrate are first mixed together, the concentration  of the enzyme-substrate complex increases, but after a time the  concentration of the complex levels off because the complex is  constantly forming and breaking down to generate product. This    principle, the so-called steady state assumption, underlies modern  theories of enzyme activity.   Even without knowing what enzymes were made of, Haldane  showed great insight in proposing that an enzyme could catalyze a  reaction by bringing its substrates into a strained or out-of-  equilibrium arrangement. This idea refined Emil Fischer’s earlier lock-  and-key simile (and was later elaborated further by Linus Pauling).  Haldane’s idea of strain was not fully appreciated until around 1970,  after the X-ray structures of several enzymes, including lysozyme  and chymotrypsin (Chapter 11), had been examined.   In addition to his work in enzymology, Haldane articulated the  role of genes in heredity and formulated mathematical estimates  of mutation rates-many years before the nature of genes or the  structure of DNA were known. However, in addition to being a  theorist, Haldane was an experimentalist who frequently performed  unpleasant or dangerous experiments on himself. For example, he  ingested sodium bicarbonate and ammonium chloride in order to  investigate their effect on breathing rate. Beyond the laboratory,  Haldane was well known for his efforts to popularize science. In  Daedalus, or, Science and the Future, Haldane commented on the  status of various branches of science circa 1924 and speculated  about future developments. His thoughts and his persona are  believed to have inspired various plots and characters in other  writers’ works of science fiction.   Briggs, G.E. and Haldane, J.B.S., A note on the kinetics of enzyme action, Biochem.   In order to be useful, kinetic expressions for overall reactions must be  formulated in terms of experimentally measurable quantities. The quantities  [ES] and [E] are not, in general, directly measurable, but the total enzyme  concentration   is usually readily determined. The rate equation for the overall enzymatic re¬  action as a function of [S] and [E] can then be derived. First, Eq. 12-14 is  combined with the steady state assumption (Eq. 12-16) to give    £ X [E][S] = *_![ES] + k 2 [ES]  Letting [E] = [E] T — [ES] and rearranging yields    The expression for the initial velocity ( v Q ) of the reaction, the velocity  (Eq. 12-13) at t = 0, thereby becomes    Both [E]t and [S] are experimentally measurable quantities. In order to meet  the conditions of the steady state assumption, the concentration of the sub¬  strate must be much greater than the concentration of the enzyme, which al¬  lows each enzyme molecule to repeatedly bind a molecule of substrate and  convert it to product, so that [ES] is constant. The use of the initial veloc¬  ity (operationally taken as the velocity measured before more than ~10% of  the substrate has been converted to product)—rather than just the velocity—  minimizes such complicating factors as the effects of reversible reactions,  inhibition of the enzyme by its product(s), and progressive inactivation of the  enzyme. (This is also why the rate of the reverse reaction in Eq. 12-12 can  be assumed to be zero.)   The maximal velocity of a reaction, V mstK > occurs at high substrate con¬  centrations when the enzyme is saturated, that is, when it is entirely in the  ES form:    This expression, the Michaelis-Menten equation , is the basic equation of en¬  zyme kinetics. It describes a rectangular hyperbola such as that plotted in  Fig. 12-3. The saturation function for oxygen binding to myoglobin (Eq. 7-6)  has the same algebraic form.    The Michaelis Constant Has a Simple Operational Definition. At the sub¬  strate concentration at which [S] = K M , Eq. 12-25 yields v 0 = V max /2 so that  K m is the substrate concentration at which the reaction velocity is half-maximal.   FIG. 1 2-3 A plot of the initial velocity v Q of a simple enzymatic reaction versus  the substrate concentration [S]. Points are plotted in 0.5 K M intervals of substrate  concentration between 0.5 K M and 5 K M . © See the Animated Figures.   Compare the curve shown here to the oxygen saturation curve for myoglobin  (Fig. 7-4).   TABLE 12-1 The Values of K Ml /c cat , and k cat /K M for Some Enzymes and Substrates    Therefore, if an enzyme has a small value of K M , it achieves maximal catalytic  efficiency at low substrate concentrations.   The K m is unique for each enzyme-substrate pair: Different substrates that  react with a given enzyme do so with different K M values. Likewise, different  enzymes that act on the same substrate have different K M values. The magni¬  tude of K m varies widely with the identity of the enzyme and the nature of  the substrate (Table 12-1J. It is also a function of temperature and pH. The  Michaelis constant (Eq. 12-20) can be expressed as   K M = — L + f=K s + f [12-26]   Since K s is the dissociation constant of the Michaelis complex (Eq. 12-15), as  K s decreases, the enzymes affinity for substrate increases. K M is therefore also  a measure of the affinity of the enzyme for its substrate, provided k 2 /k j is  small compared to K s , that is, k 2 < k-\ so that the ES —>P reaction proceeds  more slowly than ES reverts to E + S.   k cat /K m Is a Measure of Catalytic Efficiency. We can define the catalytic  constant, & cat , of an enzyme as   This quantity is also known as the turnover number of an enzyme because  it is the number of reaction processes (turnovers) that each active site catalyzes  per unit time. The turnover numbers for a selection of enzymes are given  in Table 12-1. Note that these quantities vary by over eight orders of mag¬  nitude. Equation 12-24 indicates that for a simple system, such as the  Michaelis-Menten model reaction (Eq. 12-12), k CSLt = k 2 . For enzymes with  more complicated mechanisms (e.g., multiple substrates or multiple reaction  intermediates), k cgit may be a function of several rate constants. Note that  whereas k cgLt is a constant, V maxi depends on the concentration of the enzyme  present in the experimental system. V mgtx increases as [E]t increases.   When [S] « K m , very little ES is formed. Consequently, [E] ~ [E] T , so  Eq. 12-23 reduces to a second-order rate equation:   Here, k C2it /K M is the apparent second-order rate constant of the enzymatic    How is the rate of a reaction related to its activation energy  (Section 11-2)? Consider a bimolecular reaction that proceeds  along the following pathway:   where X* represents the transition state. The rate of the reaction  can be expressed as    where k is the ordinary rate constant of the elementary reaction and  k' is the rate constant for the decomposition of X* to products.   Although X* is unstable, it is assumed to be in rapid equilib¬  rium with the reactants; that is,    where K 4 is an equilibrium constant. This central assumption of  transition state theory permits the powerful formalism of thermo¬  dynamics to be applied to the theory of reaction rates.   Since K* is an equilibrium constant, it can be expressed as    where T is the absolute temperature and R (8.3145 J • K -1 • mol -1 )  is the gas constant (this relationship between equilibrium constants  and free energy is derived in Section 1-3D). Combining the three  preceding equations yields    This equation indicates that the rate of a reaction not only de¬  pends on the concentrations of its reactants, but also decreases  exponentially with AG*. Thus, the larger the difference between the  free energy of the transition state and that of the reactants (the free  energy of activation), that is, the less stable the transition state, the  slower the reaction proceeds.    We must now evaluate k\ the rate at which X* decomposes.  The transition state structure is held together by a bond that is  assumed to be so weak that it flies apart during its first vibrational  excursion. Therefore, k' is expressed   where v is the vibrational frequency of the bond that breaks as  X* decomposes to products, and k, the transmission coefficient,  is the probability that the breakdown of X* will be in the direction  of product formation rather than back to reactants. For most spon¬  taneous reactions, k is assumed to be 1.0 (although this number,  which must be between 0 and 1, can rarely be calculated with  confidence).   where, in this case, e is the average energy of the vibration that  leads to the decomposition of X*, and h (6.6261 X 10 -34 J • s)  is Planck’s constant. Statistical mechanics tells us that at a tem¬  perature T, the classical energy of an oscillator is   where k B (1.3807 X 10 -23 J • K -1 ) is the Boltzmann constant  and k B T is essentially the available thermal energy. Combining Eqs.   Thus, combining Eqs. 12-A, 12-D, and 12-H yields the expression  for the rate constant of the elementary reaction:   This equation indicates that as the temperature rises, so that there  is increased thermal energy available to drive the reacting complex  over the activation barrier (AG*), the reaction speeds up.   reaction; the rate of the reaction varies directly with how often enzyme and  substrate encounter one another in solution. The quantity k CdX /K M is therefore  a measure of an enzyme's catalytic efficiency   There is an upper limit to the value of k C2it /K M : It can be no greater than  ky that is, the decomposition of ES to E + P can occur no more frequently  than E and S come together to form ES. The most efficient enzymes have  kca.JK m v alues near the diffusion-controlled limit of 10 8 to 10 9 M -1 • s _1 .  These enzymes catalyze a reaction almost every time they encounter a sub¬  strate molecule and hence have achieved a state of virtual catalytic perfection.  The relationship between the catalytic rate and the thermodynamics of the  transition state can now be appreciated (Box 12-2).   C Kinetic Data Can Provide Values of K max and K M   There are several methods for determining the values of the parameters of the  Michaelis-Menten equation (i.e., V max and Kf). At very high values of [S], the    An enzyme-catalyzed reaction has a K M of  1 mM and a V m2tx of 5 nM • s _1 . What is  the reaction velocity when the substrate  concentration is (a) 0.25 mM, (b) 1.5 mM,  or (c) 10 mM?    (5 nM • s 1 )(1.5 mM)  (1 mM) + (1.5 mM)    (5 nM • s _1 )(10 mM)  (1 mM) + (10 mM)   Note: When units in the numerator and  denominator cancel, it is unnecessary to  convert them to standard units before per¬  forming the calculation.    initial velocity, v Q , asymptotically approaches V max (see Sample Calculation 12-3).  In practice, however, it is very difficult to assess V max accurately from direct  plots of v Q versus [S] such as Fig. 12-3, because, even at substrate concentra¬  tions as high as [S] = 10 K M , Eq. 12-25 indicates that v Q is only 91% of  V max , so that the value of V max will almost certainly be underestimated.   A better method for determining the values of V max and K M , which was  formulated by Hans Lineweaver and Dean Burk, uses the reciprocal of the  Michaelis-Menten equation (Eq. 12-25):    This is a linear equation in l/v Q and 1 / [S]. If these quantities are plotted to  obtain the so-called Lineweaver-Burk or double-reciprocal plot, the slope  of the line is K M / V max , the l/v Q intercept is 1 / V max , and the extrapolated  1 /[S] intercept is -1 /K M (Fig. 12-4 and Sample Calculation 12-4).   As can be seen in Fig. 12-3, the best estimates of kinetic parameters are  obtained by collecting data over a range of [S] from ~0.5 K M to ~5 K M .  Thus, a disadvantage of the Lineweaver-Burk plots is that most experimental    Determine K M and V mgx for an enzyme from the following data using Eq. 12-29:    First, convert the data to reciprocal form (1 /[S] in units of mM -1 , and l/v Q in  units of mM -1 • s). Next, make a plot of l/v Q versus 1 /[S]. The x- and /-inter¬  cepts can be estimated by extrapolation of the straight line or can be calculated by  linear regression. According to Eq. 12-29 and Fig. 12-4, the /-intercept, which has  a value of ~0.1 |jlM 1 • s, is equivalent to l/V mzx , so V^nax (the reciprocal of the  /-intercept) is 10 jjlM • s 1 . The x-intercept, —0.33 mM \ is equivalent to  -1/ so K m (the negative reciprocal of the ^-intercept) is equal to 3.0 mM.    FIG. 1 2-4 A double-reciprocal  (Lineweaver-Burk) plot. The error bars  represent ±0.051/ max - The indicated points are  the same as those in Fig. 12-3. Note the large  effect of small errors at small [S] (large 1 /[S])  and the crowding together of points at large [S].  C' See the Animated Figures.   measurements of [S] are crowded onto the left side of the graph (Fig. 12-4). Section 1 Reaction Kinetics   Moreover, for small values of [S], small errors in v Q lead to large errors in 1 / v Q  and hence to large errors in K M and V^nax.   Several other types of plots, each with its advantages and disadvantages,  can also be used to determine K M and V max from kinetic data. However, ki¬  netic data are now commonly analyzed by computer using mathematically so¬  phisticated statistical treatments. Nevertheless, Lineweaver-Burk plots are still  valuable for the visual presentation of kinetic data.   Steady State Kinetics Cannot Unambiguously Establish a Reaction  Mechanism. Although steady state kinetics provides valuable information  about the rates of buildup and breakdown of ES, it provides little insight as  to the nature of ES. Thus, an enzymatic reaction may, in reality, pass through  several more or less stable intermediate states such as    Unfortunately, steady state kinetic measurements are incapable of revealing the  number of intermediates in an enzyme-catalyzed reaction. Thus, such mea¬  surements of a multistep reaction can be likened to a “black box” containing  a system of water pipes with one inlet and one drain:   At steady state, that is, after the pipes have filled with water, the relationship  between input pressure and output flow can be measured. However, such  measurements yield no information concerning the detailed construction of  the plumbing connecting the inlet to the drain. This would require additional  information, such as opening the box and tracing the pipes. Likewise, steady  state kinetic measurements can provide a phenomenological description of en¬  zymatic behavior, but the nature of the intermediates remains indeterminate.  The existence of intermediates must be verified independently, for example,  by identifying them through the use of spectroscopic techniques.   The foregoing highlights a central principle of enzymology: The steady  state kinetic analysis of a reaction cannot unambiguously establish its mechanism.  This is because no matter how simple, elegant, or rational a postulated mech¬  anism, there are an infinite number of alternative mechanisms that can also  account for the kinetic data. Usually, it is the simpler mechanism that turns  out to be correct, but this is not always the case. However, if kinetic data are  not compatible with a given mechanism, then that mechanism must be rejected.   Chapter 12 Enzyme Kinetics, Inhibition, and Control Therefore, although kinetics cannot be used to establish a mechanism unam¬  biguously without confirming data, such as the physical demonstration of an  intermediate’s existence, the steady state kinetic analysis of a reaction is of great  value because it can be used to eliminate proposed mechanisms.   We have heretofore been concerned with simple, single-substrate reactions that  obey the Michaelis—Menten model (Reaction 12-12). Yet, enzymatic reactions  requiring multiple substrates and yielding multiple products are far more com¬  mon. Indeed, those involving two substrates and yielding two products   account for —60% of known biochemical reactions. Almost all of these so-  called bisubstrate reactions are either transfer reactions in which the enzyme  catalyzes the transfer of a specific functional group, X, from one of the sub¬  strates to the other:   or oxidation—reduction reactions in which reducing equivalents are transferred  between the two substrates. For example, the hydrolysis of a peptide bond by  trypsin (Section 11-5) is the transfer of the peptide carbonyl group from the  peptide nitrogen atom to water (Fig. 12-5 a), whereas in the alcohol dehydro¬  genase reaction (Section 11-1B), a hydride ion is formally transferred from  ethanol to NAD + (Fig. 12-5 b). Although bisubstrate reactions could, in  principle, occur through a vast variety of mechanisms, only a few types are  commonly observed.   Sequential Reactions Occur via Single Displacements. Reactions in which  all substrates must combine with the enzyme before a reaction can occur and prod¬  ucts be released are known as sequential reactions . In such reactions, the group  being transferred, X, is directly passed from A (= P—X) to B, yielding P  and Q (= B—X). Hence, such reactions are also called single-displacement  reactions.   Sequential reactions can be subclassified into those with a compulsory or¬  der of substrate addition to the enzyme, which are said to have an Ordered  mechanism, and those with no preference for the order of substrate addition,  which are described as having a Random mechanism. In the Ordered mech¬  anism, the binding of the first substrate is apparently required for the enzyme  to form the binding site for the second substrate, whereas in the Random  mechanism, both binding sites are present on the free enzyme.    FIG. 1 2-5 Some bisubstrate reactions, (a) In the peptide hydrolysis reaction  catalyzed by trypsin, the peptide carbonyl group, with its pendent polypeptide chain, R 1( is  transferred from the peptide nitrogen atom to a water molecule, (b) In the alcohol  dehydrogenase reaction, a hydride ion is formally transferred from ethanol to NAD + .    In a notation developed by W.W. Cleland, substrates are designated by the  letters A and B in the order that they add to the enzyme, products are desig¬  nated by P and Q in the order that they leave the enzyme, the enzyme is rep¬  resented by a horizontal line, and successive additions of substrates and releases  of products are denoted by vertical arrows. An Ordered bisubstrate reaction is  thereby diagrammed:    where A and B are said to be the leading and following substrates, respec¬  tively. Many NAD + - and NADP + -requiring dehydrogenases follow an Ordered  bisubstrate mechanism in which the coenzyme is the leading substrate.   Some dehydrogenases and kinases operate through Random bisubstrate mech¬  anisms (kinases are enzymes that transfer phosphoryl groups from ATP to  other compounds or vice versa).    Ping Pong Reactions Occur via Double Displacements. Group-transfer re¬  actions in which one or more products are released before all substrates have been  added are known as Ping Pong reactions . The Ping Pong bisubstrate reaction  is represented by    Here, a functional group X of the first substrate A (= P—X) is displaced from  the substrate by the enzyme E to yield the first product P and a stable en¬  zyme form F (= E—X) in which X is tightly (often covalently) bound to the  enzyme (Ping). In the second stage of the reaction, X is displaced from the  enzyme by the second substrate B to yield the second product Q (— B—X),  thereby regenerating the original form of the enzyme, E (Pong). Such reac¬  tions are therefore also known as double-displacement reactions. Note that  in Ping Pong reactions , the substrates A and B do not encounter one another on  the surface of the enzyme. Many enzymes, including trypsin (in which F is the  acyl-enzyme intermediate; Section 11-5), transaminases, and some flavo-  enzymes, react with Ping Pong mechanisms.   The rate equations that describe the foregoing bisubstrate mechanisms are con¬  siderably more complicated than the equation for a single-substrate reaction.  In fact, the equations for bisubstrate mechanisms (which are beyond the scope  of this text) contain as many as four kinetic constants versus two ( V msx and Kjf)  for the Michaelis-Menten equation. Nevertheless, steady state kinetic measure¬  ments can be used to distinguish among the various bisubstrate mechanisms.   • Write the rate equations for a first-order  and a second-order reaction.   • If you know a reaction’s half-life, can you  determine its rate constant? What other  information do you need?   • What are the differences between instan¬  taneous velocity, initial velocity, and maxi¬  mal velocity for an enzymatic reaction?   • What do the values of K M and k cai /K M  reveal about an enzyme?   • Write the Lineweaver-Burk (double¬  reciprocal) equation and describe the  features of a Lineweaver-Burk plot.   • Why can’t enzyme kinetics prove that a  particular enzyme mechanism is correct?   • Use Cleland notation to describe Ordered  and Random sequential reactions and a  Ping Pong reaction.   • Enzyme inhibitors interact reversibly or irreversibly with an enzyme to alter its K M  and/or l/ max values.   • A competitive inhibitor binds to the enzyme’s active site and increases the appar¬  ent K m for the reaction.   • An uncompetitive enzyme inhibitor affects catalytic activity such that both the  apparent K M and the apparent l/ max decrease.   • A mixed enzyme inhibitor alters both catalytic activity and substrate binding such  that the apparent l/ max decreases and the apparent K M may increase or decrease.    Many substances alter the activity of an enzyme by combining with it in a  way that influences the binding of substrate and/or its turnover number.  Substances that reduce an enzyme's activity in this way are known as inhibitors .  A large part of the modern pharmaceutical arsenal consists of enzyme in¬  hibitors. For example, AIDS is treated almost exclusively with drugs that in¬  hibit the activities of certain viral enzymes.   Inhibitors act through a variety of mechanisms. Irreversible enzyme in¬  hibitors, or inactivators, bind to the enzyme so tightly that they permanently  block the enzymes activity. Reagents that chemically modify specific amino  acid residues can act as inactivators. For example, the compounds used to iden¬  tify the catalytic Ser and His residues of serine proteases (Section 11-5A) are  inactivators of these enzymes.   Reversible enzyme inhibitors diminish an enzyme s activity by interacting re¬  versibly with it. Some enzyme inhibitors are substances that structurally resem¬  ble their enzymes substrates but either do not react or react very slowly. These  substances are commonly used to probe the chemical and conformational na¬  ture of an enzymes active site in an effort to elucidate the enzymes catalytic  mechanism. Other inhibitors affect catalytic activity without interfering with  substrate binding. Many do both. In this section, we discuss several of the sim¬  plest mechanisms for reversible inhibition and their effects on the kinetic behav¬  ior of enzymes that follow the Michaelis—Menten model. As in the preceding  discussion, we will base our analysis on a simple one-substrate reaction model.    A Competitive Inhibition Involves Inhibitor Binding at an Enzyme’s  Substrate Binding Site   A substance that competes directly with a normal substrate for an enzymes  substrate-binding site is known as a competitive inhibitor. Such an inhibitor  usually resembles the substrate so that it specifically binds to the active site  but differs from the substrate so that it cannot react as the substrate does. For  example, succinate dehydrogenase, a citric acid cycle enzyme that converts  succinate to fumarate (Section 17-3F), is competitively inhibited by mal-  onate, which structurally resembles succinate but cannot be dehydrogenated:   (b) X-Ray structure of influenza virus neuraminidase in complex with its  competitive inhibitor, oseltamivir carboxylate. The active site region of  the H5N1 avian flu neuraminidase is represented by its semitranspar¬  ent surface diagram. The oseltamivir carboxylate is shown in space¬  filling form with C green, 0 red, and N blue. [Based on an X-ray  structure by John Skehel, MRC National Institute for Medical Research,  London, U.K. PDBid 2HU4.]    The effectiveness of malonate as a competitive inhibitor of suc¬  cinate dehydrogenase strongly suggests that the enzymes sub¬  strate-binding site is designed to bind both of the substrates  carboxylate groups, presumably through the influence of appro¬  priately placed positively charged residues.   Similar principles are responsible for product inhibition. In  this phenomenon, a product of the reaction, which necessarily  is able to bind to the enzymes active site, may accumulate and  compete with substrate for binding to the enzyme in subsequent catalytic  cycles. Product inhibition is one way in which the cell controls the activities  of its enzymes (Section 12-3).   Computer-assisted design of compounds that fit snugly into an enzymes  active site is an essential part of modern drug development (discussed fur¬  ther in Section 12-4). For example, the anti-influenza drug oseltamivir  (Tamiflu) is hydrolyzed in the liver (Fig. 12-6^) to yield oseltamivir carboxy¬  late, a competitive inhibitor of influenza neuraminidase that binds to this en¬  zyme s active site (Fig. \2-6b). Neuraminidase catalyzes the hydrolysis of  neuraminic acid (sialic acid; Section 8-1C) to help the viral particles escape  from the host cell surface, where membrane glycoproteins contain sialic acid  residues. Oseltamivir carboxylate, which can significantly limit the production  of the flu virus, has an inhibition constant (AJ, the dissociation constant for  enzyme—inhibitor binding, see below) of less than 1 nM with neuraminidase  (the enzyme’s K M is in the micromolar range).   Transition state analogs are particularly effective inhibitors. This is be¬  cause effective catalysis often depends on an enzymes ability to bind to and  stabilize its reactions transition state (Section 11-3E). A compound that mim¬  ics a transition state exploits these binding interactions in ways that a substrate  analog cannot. For example, adenosine deaminase converts the nucleoside  adenosine to inosine as follows:    The K m of the enzyme for the substrate adenosine is 3X10 5 M. The prod¬  uct inosine acts as an inhibitor of the reaction, with a AJ value of 3 X 10“ 4 M.  However, a transition state analog, 1,6-dihydroinosine (at left), inhibits the  reaction with a of 1.5 X 10 13 M. Several of the drugs used to block  HIV protease, an essential enzyme for production of the human immuno¬  deficiency virus, are compounds that were designed to mimic the enzymes  transition state and bind to the enzyme with high affinity (Box 12-3).    The human immunodeficiency virus (HIV) causes acquired im¬  munodeficiency syndrome (AIDS) by infecting and destroying the  host’s immune system. In the first steps of infection, HIV attaches  to a target cell and injects its genetic material (which is RNA  rather than DNA) into the host cell. The viral RNA is transcribed  into DNA by a viral enzyme called reverse transcriptase  (Box 25-2). After this DNA is integrated into the host’s genome,  the cell can produce more viral RNA and proteins for packaging it  into new viral particles that bud off from the host cell membrane.   Most of the viral proteins are synthesized as parts of larger  polypeptide precursors known as polyproteins. Consequently,  proteolytic processing by the virally encoded HIV protease to    release these viral proteins is necessary for viral reproduction. In  the absence of an effective vaccine for HIV, efforts to prevent and  treat AIDS have led to the development of compounds that inhibit  HIV reverse transcriptase and HIV protease.   Several inhibitors of reverse transcriptase have been developed.  The archetype is AZT (3'-azido-3'-deoxythymidine; Zidovudine),  which is taken up by cells, phosphorylated, and incorporated into  the DNA chains synthesized from the HIV template by reverse  transcriptase. Because AZT lacks a 3'-OH group, it cannot sup¬  port further polynucleotide chain elongation and therefore termi¬  nates DNA synthesis (Section 3-4C). Most cellular DNA  polymerases have a low affinity for phosphorylated AZT, but re¬  verse transcriptase has a high affinity for this drug, which makes  AZT effective against viral replication. Other nucleoside analogs  that are used to treat HIV infection are 2',3'-dideoxycytidine  (ddC, Zalcitabine) and 2',3'-dideoxyinosine (ddl, Didanosine;  inosine nucleotides are metabolically converted to adenosine and  guanosine nucleotides), which also act as chain terminators.  Nonnucleoside compounds such as nevirapine do not bind to the  reverse transcriptase active site but instead bind to a hydrophobic  pocket elsewhere on the enzyme.    HIV protease is a homodimer of 99-residue subunits. Mecha¬  nistically, it is a so-called aspartic protease, a family of proteases  that includes pepsin (the gastric protease that operates at low    The Degree of Competitive Inhibition Varies with the Fraction of Enzyme  That Has Bound Inhibitor. The general model for competitive inhibition is  given by the following reaction scheme:    pH). Comparisons among the active sites of the aspartic  proteases were instrumental in designing HIV protease inhibitors.  HIV protease cleaves a number of specific peptide bonds, including  Phe-Pro and Tyr-Pro peptide bonds in its physiological  substrates, the HIV polyproteins. Inhibitors based on these se¬  quences should therefore selectively inhibit the viral protease. The  peptidomimetic (peptide-imitating) drugs ritonavir and saquinavir    contain phenyl and other bulky groups that  bind in the HIV protease active site. Of  perhaps even greater importance, these  drugs have the geometry of the catalyzed  reaction’s tetrahedral transition state (red). The  enzyme’s peptide substrate is shown for com¬  parison. O See Interactive Exercise 9.   The efficacy of anti-HIV agents, like that  of many drugs, is limited by their side ef¬  fects. Despite their preference for viral en¬  zymes, anti-HIV drugs also interfere with  normal cellular processes. For example, the  inhibition of DNA synthesis by reverse tran¬  scriptase inhibitors in rapidly dividing cells,  such as the bone marrow cells that give rise  to erythrocytes, can lead to severe anemia.  Other side effects include nausea, kidney  stones, and rashes. Side effects are particu¬  larly problematic in HIV infection, because  drugs must be taken several times daily for  many years, if not for a lifetime.   Acquired resistance also limits the effec¬  tiveness of antiviral drugs. This is a signifi¬  cant problem in HIV infection because the  error-prone reverse transcriptase allows HIV  to mutate rapidly. Numerous mutations in  HIV are known to be associated with drug  resistance.   In the case of HIV infection, it seems  unlikely that a single drug will prove to be a  “magic bullet,” in part because HIV infects  many cell types, but mostly because of its  ability to rapidly evolve resistance against  any one drug. The outstanding success of  anti-HIV therapy rests on combination ther¬  apy, in which several different drugs are  administered simultaneously. This success¬  fully keeps AIDS at bay by reducing levels  of HIV, in some cases to undetectable levels, thereby reducing  the probability that HIV will evolve a drug-resistant variant. The  advantages of using an inhibitor “cocktail” containing inhibitors of  reverse transcriptase and HIV protease include (1) decreasing  the likelihood that a viral strain will simultaneously develop resist¬  ance to every compound in the mix and (2) decreasing the doses  and hence the side effects of the individual compounds.    Here, I is the inhibitor, El is the catalytically inactive enzyme—inhibitor com¬  plex, and it is assumed that the inhibitor binds reversibly to the enzyme and  is in rapid equilibrium with it so that    A competitive inhibitor therefore reduces the concentration of free enzyme avail¬  able for substrate binding.   The Michaelis—Menten equation for a competitively inhibited reaction is  derived as before (Section 12-IB), but with an additional term to account for  the fraction of [E] T that binds to I to form El ([E] T = [E] + [ES] + [El]).  The resulting equation,    is the Michaelis-Menten equation that has been modified by a factor, a, which  is defined as   Note that a, a function of the inhibitor s concentration and its affinity for the  enzyme, cannot be less than 1. Comparison of Eqs. 12-25 and 12-31 indi¬  cates that Kff = olK m , where is the apparent K M , that is, the K M value   that would be measured in the absence of the knowledge that inhibitor is pres¬  ent. Figure 12-7 shows the hyperbolic plots of Eq. 12-31 for increasing val¬  ues of a. The presence of I makes [S] appear to be less than it really is (makes  K m appear to be larger than it really is), a consequence of the binding of  I and S to E being mutually exclusive. However, increasing [S] can overwhelm  a competitive inhibitor. In fact, a is the factor by which [S] must be increased  in order to overcome the effect of the presence of inhibitor. As [S] approaches in¬  finity, v Q approaches V max for any value of a (that is, for any concentration of  inhibitor). Thus, the inhibitor does not affect the enzymes turnover number.  Competitive inhibition is the principle behind the use of ethanol to treat  methanol poisoning. Methanol itself is only mildly toxic. However, the  liver enzyme alcohol dehydrogenase converts methanol to the highly toxic  formaldehyde (at left), only small amounts of which cause blindness and death.  Ethanol competes with methanol for binding to the active site of liver alco¬  hol dehydrogenase, thereby slowing the production of formaldehyde from  methanol (the ethanol is converted to the readily metabolized acetaldehyde;  at left). Thus, through the administration of ethanol, a large portion of the  methanol will be harmlessly excreted from the body in the urine before it can  be converted to formaldehyde. The same principle underlies the use of ethanol  to treat antifreeze (ethylene glycol, HOCH 2 CH 2 OH) poison¬  ing, which, due to its sweet taste, often afflicts cats and dogs.    A plot of this equation is linear and has a slope of a    FIG. 1 2-7 A plot of v Q versus [S] for a Michaelis-Menten reaction  in the presence of different concentrations of a competitive inhibitor.    FIG. 1 2-8 A Lineweaver-Burk plot of the competitively inhibited  Michaelis-Menten enzyme described by Fig. 12-7. Note that all lines intersect on the  1 /v Q axis at 1 /l/ max . The varying slopes indicate the effect of the inhibitor on aK M =    1 /[S] intercept of — l/a K M , and a l/// Q intercept of l/V max (Fig. 12-8). The  double-reciprocal plots for a competitive inhibitor at various concentrations of I in¬  tersect at 1 / V max on the l/v Q axis, a property that is diagnostic of competitive in¬  hibition.   The value of K± for a competitive inhibitor can be determined from the  plot of KZZ = (1 + [ I \/Ki)K m versus [I]; its intercept on the [I] axis is — AJ.  AJ can also be calculated from Eq. 12-32 if KZZ — aK M is determined at a  known [I] for an enzyme of known K M (see Sample Calculation 12-5).  Comparing the K\ values of competitive inhibitors with different structures  can provide information about the binding properties of an enzyme’s active  site and hence its catalytic mechanism. For example, to ascertain the  importance of the various segments of an ATP molecule   for binding to the active site of an ATP-requiring enzyme, one might deter¬  mine the Ki, say, for ADP, AMP, ribose, triphosphate, etc. Since many of these  ATP components are unreactive, inhibition studies are the most convenient  method of monitoring their binding to the enzyme.    An enzyme has a K M of 8 jjlM in the ab¬  sence of a competitive inhibitor and a K  of 12 |xM in the presence of 3 |xM of  the inhibitor. Calculate AJ.    First calculate the value of a when  K m = 8 |xM and = 12 |jlM:   chapter 12 Enzyme Kinetics, inhibition, and Control g Uncompetitive Inhibition Involves Inhibitor Binding to the   In uncompetitive inhibition, the inhibitor binds directly to the enzyme-  substrate complex but not to the free enzyme:    In this case, the inhibitor binding step has the dissociation constant    The binding of uncompetitive inhibitor, which need not resemble substrate,  presumably distorts the active site , thereby rendering the enzyme catalytically  inactive.   The Michaelis-Menten equation for uncompetitive inhibition and the  equation for its double-reciprocal plot are given in Table 12-2. The double¬  reciprocal plot consists of a family of parallel lines (Fig. 12-9) with  slope K m / V max , 1 / V 0 intercepts of a'/ V m2iX , and 1 /[S] intercepts of —a ’/K M .  Note that in uncompetitive inhibition, both = K m /ol' and V^ 9 X = V mSLX /a'  are decreased, but that = K M /V mSiX . In contrast to the case for   competitive inhibition, adding substrate does not reverse the effect of an un¬  competitive inhibitor because the binding of substrate does not interfere with  the binding of uncompetitive inhibitor.   Uncompetitive inhibition requires that the inhibitor affect the catalytic  function of the enzyme but not its substrate binding. This is difficult to en¬  vision for single-substrate enzymes. In actuality, uncompetitive inhibition is  significant only for multisubstrate enzymes.   FIG. 1 2-9 A Lineweaver-Burk plot of a Michaelis-Menten enzyme in the  presence of an uncompetitive inhibitor. Note that all lines have identical slopes of K M /V max .    C Mixed Inhibition Involves Inhibitor Binding to Both the Free  Enzyme and the Enzyme-Substrate Complex   Many reversible inhibitors interact with the enzyme in a way that affects sub¬  strate binding as well as catalytic activity. In other words, both the enzyme  and the enzyme—substrate complex bind inhibitor, resulting in the following  model:    This phenomenon is known as mixed inhibition (alternatively, noncompet¬  itive inhibition). Presumably, a mixed inhibitor binds to enzyme sites that par¬  ticipate in both substrate binding and catalysis. For example, metal ions, which  do not compete directly with substrates for binding to an enzyme active site,  as do competitive inhibitors, may act as mixed inhibitors. The two dissocia¬  tion constants for inhibitor binding   The Michaelis-Menten equation and the corresponding double-reciprocal  equation for mixed inhibition are given in Table 12-2. As in uncompetitive  inhibition, the apparent values of K M and V^ ax are modulated by the pres¬  ence of inhibitor. The name mixed inhibition arises from the fact that the de¬  nominator of the Michaelis-Menten equation has the factor a multiplying K M  as in competitive inhibition and the factor a' multiplying [S] as in uncom¬  petitive inhibition. Thus K = (a/a')A}^and = V msx ./a'.   • Why might an enzyme’s substrate, transi¬  tion state, and product all serve as start¬  ing points for the design of a competitive  inhibitor?   • Describe the effects of competitive, un¬  competitive, and mixed inhibitors on K M  and I/max 1   • How does pure noncompetitive inhibition  differ from other forms of inhibition?    Depending on the relative values of a and a' (and hence K\ and K\),  may increase (as in competitive inhibition) or decrease. If the enzyme and  enzyme-substrate complex bind I with equal affinity, then a = a' and the K$ p  value is unchanged from the K M for the reaction in the absence of inhibitor.  In this case, only V^n ax is affected, a phenomenon that is named pure non¬  competitive inhibition.   Double-reciprocal plots for mixed inhibition consist of lines that have the  slope olK m / V^n ax , a \/v Q intercept of a'/ V max , and a 1 /[S] intercept of  —ol'/olK m (Fig. 12-10). The lines for increasing values of [I] (representing in¬  creasing saturation of E and ES with I) intersect to the left of the \/v Q axis.  For pure noncompetitive inhibition, the lines intersect on the 1 / [S] axis at  —1 /K m . As with uncompetitive inhibition, substrate binding does not reverse  the effects of mixed inhibition.   The kinetics of an enzyme inactivator (an irreversible inhibitor) resembles  that of a pure noncompetitive inhibitor because the inactivator reduces the  concentration of functional enzyme at all substrate concentrations.  Consequently, V^ax decreases and K M is unchanged. The double-reciprocal  plots for irreversible inactivation therefore resemble those for pure noncom¬  petitive inhibition (the lines intersect on the 1 /[S] axis).    • Allosteric effectors bind to multisubunit enzymes such as aspartate transcarbamoyl-  ase, thereby inducing cooperative conformational changes that alter the enzyme’s  catalytic activity.    • Phosphorylation and dephosphorylation of an enzyme such as glycogen phospho-  rylase can control its activity by shifting the equilibrium between more active and  less active conformations.    An organism must be able to control the catalytic activities of its component  enzymes so that it can coordinate its numerous metabolic processes, respond    to changes in its environment, and grow and differentiate, all in an orderly  manner. There are two ways that this may occur:   1. Control of enzyme availability . The amount of a given enzyme in a  cell depends on both its rate of synthesis and its rate of degradation.  Each of these rates is directly controlled by the cell and is subject to  dramatic changes over time spans of minutes (in bacteria) to hours (in  higher organisms).   2. Control of enzyme activity. As we have already seen, an enzymes activ¬  ity can be inhibited by the accumulation of product and by the presence  of other types of inhibitors, In fact, an enzymes catalytic activity can be  modulated—either negatively or positively—through structural alter¬  ations that influence the enzymes substrate-binding affinity or turnover  number. Just as hemoglobins oxygen affinity is allosterically regulated by  the binding of ligands such as 0 2 , C0 2 , H + , and BPG (Section 7-ID),  an enzymes substrate-binding affinity may likewise vary with the bind¬  ing of small molecules, called allosteric effectors. Allosteric mechanisms  can cause large changes in enzymatic activity. The activities of many en¬  zymes are similarly controlled by covalent modification, usually phos¬  phorylation and dephosphorylation of specific Ser, Thr, or Tyr residues.   In the following sections we discuss the control of enzyme activity by allosteric  interactions and covalent modification.    A Allosteric Control Involves Binding at a Site Other  than the Active Site   In this section we examine allosteric control of enzymatic activity by consid¬  ering one example—aspartate transcarbamoylase (ATCase) from E. coli.  Other allosteric enzymes are discussed in later chapters.   Aspartate transcarbamoylase catalyzes the formation of Af-carbamoyl aspar¬  tate from carbamoyl phosphate and aspartate:    This reaction is the first step unique to the biosynthesis of pyrimidines  (Section 23-2A). The allosteric behavior of E. coli ATCase has been investi¬  gated by John Gerhart and Howard Schachman, who demonstrated that both  of its substrates bind cooperatively to the enzyme. Moreover, ATCase  is allosterically inhibited by cytidine triphosphate (CTP), a pyrim¬  idine nucleotide, and is allosterically activated by adenosine triphos¬  phate (ATP), a purine nucleotide.   The v Q versus [S] curve for ATCase (Fig. 12-11) is sigmoidal,  rather than hyperbolic as it is in enzymes that follow the  Michaelis—Menten model. This is consistent with cooperative sub-   Reaction velocity was measured in the absence of allosteric effectors, in the  presence of 0.4 mM CTP (an inhibitor), and in the presence of 2.0 mM ATP (an  activator). [After Kantrowitz, E.R., Pastra-Landis, S.C., and Lipscomb, W.N., Trends  Biochem. Sci. 5, 125 (1980).] O See the Animated Figures.   CTP, the end product of the pathway, inhibits ATCase, which catalyzes the pathway’s first  step.    strate binding (recall that hemoglobins 0 2 -binding curve is also sigmoidal;  Fig. 7-6). ATCase s allosteric effectors shift the entire curve to the right or the  left: At a given substrate concentration, CTP decreases the enzymes catalytic  rate, whereas ATP increases it.   CTP, which is a product of the pyrimidine biosynthetic pathway, is an ex¬  ample of a feedback inhibitor, since it inhibits an earlier step in its own biosyn¬  thesis (Fig. 12-12). Thus, when CTP levels are high, CTP binds to ATCase,  thereby reducing the rate of CTP synthesis. Conversely, when cellular [CTP]  decreases, CTP dissociates from ATCase and CTP synthesis accelerates.   The metabolic significance of the ATP activation of ATCase is that it tends  to coordinate the rates of synthesis of purine and pyrimidine nucleotides,  which are required in roughly equal amounts in nucleic acid biosynthesis. For  instance, if the ATP concentration is much greater than that of CTP, ATCase  is activated to synthesize pyrimidine nucleotides until the concentrations of  ATP and CTP become balanced. Conversely, if the CTP concentration is  greater than that of ATP, CTP inhibition of ATCase permits purine nucleotide  biosynthesis to balance the ATP and CTP concentrations.   Allosteric Changes Alter ATCase’s Substrate-Binding Sites. E. ca/z ATCase  (300 kD) has the subunit composition c 6 r 6 , where c and r represent its cat¬  alytic and regulatory subunits. The X-ray structure of ATCase (Fig. 12-13),  determined by William Lipscomb, reveals that the catalytic subunits are  arranged as two sets of trimers (c 3 ) in complex with three sets of regulatory  dimers (r 2 ). Each regulatory dimer joins two catalytic subunits in different  c 3 trimers.   The isolated catalytic trimers are catalytically active, have a maximum cat¬  alytic rate greater than that of intact ATCase, exhibit a noncooperative (hy¬  perbolic) substrate saturation curve, and are unaffected by the presence of ATP  or CTP The isolated regulatory dimers bind the allosteric effectors but are de¬  void of enzymatic activity. Evidently, the regulatory subunits allosterically reduce  the activity of the catalytic subunits in the intact enzyme.   T-state enzyme in complex with CTP is viewed (a) along the protein’s  molecular threefold axis of symmetry and (b) along a molecular  twofold axis of symmetry perpendicular to the view in Part a. The  polypeptide chains are drawn in worm form embedded in their  transparent molecular surface. The regulatory dimers ( yellow ) join the  upper catalytic trimer ( red) to the lower catalytic trimer (blue). CTP is  drawn in space-filling form colored according to atom type (C green,   0 red, N blue, and P orange), (c) The R-state enzyme in complex with  PALA viewed as in Part b. PALA (which is bound to the c subunits  but largely obscured here) is drawn in space-filling form. Note how  the rotation of the regulatory dimers in the T —> R transition causes  the catalytic trimers to move apart along the threefold axis. [Based on  X-ray structures by William Lipscomb, Harvard University. PD Bids  5AT1 and 8ATC.] O See Kinemage Exercise 11-1.    As allosteric theory predicts (Section 7-ID), the activator ATP preferen¬  tially binds to ATCase s active (R or high substrate affinity) state, whereas the  inhibitor CTP preferentially binds to the enzyme’s inactive (T or low substrate  affinity) state. Similarly, the unreactive bisubstrate analog Af-(phosphon-  acetyl)-L-aspartate (PALA) binds tightly to R-state but not to T-state  ATCase.   X-Ray structures have been determined for the T-state ATCase-CTP com-  : and the R-state ATCase-PALA complex (as a rule, unreactive substrate    analogs, such as PALA, form complexes with an enzyme that are more  amenable to structural analysis than complexes of the enzyme with rapidly re¬  acting substrates). Structural studies reveal that in the T —> R transition, the   Chapter 12 Enzyme Kinetics, Inhibition, and Control enzymes catalytic trimers separate along the molecular threefold axis by   ~11 A and reorient around this axis relative to each other by 12° (Fig. 12-13 b,c).  In addition, the regulatory dimers rotate clockwise by 15° around their twofold  axes and separate by ~4 A along the threefold axis. Such large quaternary  shifts are reminiscent of those in hemoglobin (Section 7-ID).   Each catalytic subunit of ATCase consists of a carbamoyl phosphate-binding  domain and an aspartate-binding domain. The binding of PALA to the en¬  zyme, which presumably mimics the binding of both substrates, induces a con¬  formational change that swings the two domains together such that their two  bound substrates can react to form product (Fig. 12-14). The conformational  changes—movements of up to 8 A for some residues—in a single catalytic sub¬  unit trigger ATCase s T —> R quaternary shift. ATCase s tertiary and quater¬  nary shifts are tightly coupled (i.e., ATCase closely follows the symmetry  model of allosterism; Section 7-ID). The binding of substrate to one catalytic  subunit therefore increases the substrate-binding affinity and catalytic activity of  the other five catalytic subunits and hence accounts for the enzymes coopera¬  tive substrate binding.   The structural basis for the effects of CTP and ATP on ATCase activity has  been partially unveiled. Both the inhibitor CTP and the activator ATP bind  to the same site on the outer edge of the regulatory subunit, about 60 A away    arrows ) are prevented by steric interference (purple bar) between the  contacting aspartic acid-binding domains, (b) The binding of  carbamoyl phosphate (CP) followed by aspartic acid (Asp) to their  respective binding sites causes the subunits to move apart and rotate    promote the reaction of their bound substrates to form products.  [Illustration, Irving Geis/Geis Archives Trust. Copyright Howard Hughes  Medical Institute. Reproduced with permission.] O See Kinemage  Exercises 11-1 and 11-2.   from the nearest catalytic site. CTP binds preferentially to the T state, increas- Section 3 Control of Enzyme Activity   ing its stability, whereas ATP binds preferentially to the R state, increasing its   The binding of CTP and ATP to their less favored enzyme states also has  structural consequences. When CTP binds to R-state ATCase, it induces a  contraction in the regulatory dimer that causes the catalytic trimers to come  together by 0.5 A (become more T-like, that is, less active). This, in turn, re¬  orients key residues in the enzymes active sites, thereby decreasing the en¬  zymes catalytic activity. ATP has essentially opposite effects when binding to  the T-state enzyme: It causes the catalytic trimers to move apart by 0.4 A (be¬  come more R-like, that is, more active), thereby reorienting key residues in  the enzyme’s active sites so as to increase the enzymes catalytic activity.   Allosteric Transitions in Other Enzymes Often Resemble Those of  Hemoglobin and ATCase. Allosteric enzymes are widely distributed in nature  and tend to occupy key regulatory positions in metabolic pathways. Such en¬  zymes are almost always symmetrical proteins containing at least two subunits,  in which quaternary structural changes communicate binding and catalytic ef¬  fects among all active sites in the enzyme. The quaternary shifts are primarily  rotations of subunits relative to one another. Secondary structures are largely  preserved in T —> R transitions, which is probably important for mechanically  transmitting allosteric effects over distances of tens of angstroms.   In addition to allosteric interactions, many enzymes may be subject to con¬  trol by covalent modification. In eukaryotes, by far the most common such  modification is phosphorylation and dephosphorylation (the attachment and  removal of a phosphoryl group) of the hydroxyl group of a Ser, Thr, or Tyr  residue.   Such enzymatic modification/demodification processes, which are catalyzed by  enzymes known as protein kinases and protein phosphatases, alter the ac¬  tivities of the modified proteins. Indeed, ~30% of human proteins, which  collectively participate in nearly all biological processes, are subject to control  by reversible phosphorylation.   As an example of an enzyme whose activity is controlled by covalent mod¬  ification, let us consider glycogen phosphorylase (or simply phosphorylase),  which catalyzes the phosphorolysis (bond cleavage by the substitution of a  phosphate group) of glycogen [the starchlike polysaccharide that consists   Chapter 5 Proteins: Primary Structure mainly of a(l—>4)-linked glucose residues; Section 8-2C] to yield glucose-1-   This is the rate-controlling step in the metabolic pathway of glycogen break¬  down, an important supplier of fuel for metabolic activities (Section 16-1).    IFIG. 12-15 X-Ray structure of rabbit muscle glycogen phosphorylase. Ribbon  diagram of the phosphorylase a homodimer viewed along its molecular two-fold axis of  symmetry. Each subunit consists of an N-terminal domain, which is subdivided into an  interface subdomain (residues 1-315) and a glycogen-binding subdomain (residues  316-484), and a C-terminal domain (residues 485-842). The enzyme’s several ligands are  drawn in stick form colored according to type with N blue, 0 red, P orange, and C atoms  as follows. The active site is marked by a bound glucose molecule (C yellow). Pyridoxal  phosphate (PLP) is covalently linked to the side chain of Lys 678 in the C-terminal domain  (C magenta). In addition, the enzyme binds its allosteric effector AMP (C cyan) and  maltoheptaose (C green), an a(1—»4)-l inked glucose heptamer, which is bound in the  enzyme’s glycogen storage site. Ser 14-P, the phosphoryl group on Ser 14, is drawn in  space-filling form. [X-Ray structure coordinates courtesy of Stephen Sprang, University of  Texas Southwest Medical Center.] O See Kinemage Exercise 14-1.   genetically distinct enzymes from the same organism; also called isoforms) of  glycogen phosphorylase, those from muscle, brain, and liver. Muscle glycogen  phosphorylase, which we discuss here, is a dimer of identical 842-residue sub¬  units. It is regulated both by allosteric interactions and by phosphorylation/  dephosphorylation. The phosphorylated form of the enzyme, phosphorylase ci,  has a phosphoryl group esterified to its Ser 14. The dephospho form is called  phosphorylase b.   The X-ray structures of phosphorylase a and phosphorylase b, which were  respectively determined by Robert Fletterick and Louise Johnson, are similar.   Both have a large N-terminal domain (484 residues; the largest known do¬  main) and a smaller C-terminal domain (Fig. 12-15). The N-terminal do¬  main, which is subdivided into a glycogen-binding subdomain and an interface  subdomain, contains the phosphorylation site (Ser 14), an allosteric effector  site, a glycogen-binding site (called the glycogen storage site), and all the in¬  tersubunit contacts in the dimer. The enzyme’s active site is located at the cen¬  ter of the subunit.   Phosphorylation and Dephosphorylation Can Alter Enzymatic Activity in a  Manner That Resembles Allosteric Control. Glycogen phosphorylase has  two conformational states, the enzymatically active R state and the enzymat¬  ically inactive T state (Fig. 12-16). The T-state enzyme is inactive because it  has a malformed active site and a surface loop (residues 282-284) that blocks  substrate access to its binding site. In contrast, in the R-state enzyme, the side  chain of Arg 569 has reoriented so as to bind the substrate phosphate ion and   FIG. 12-16 Conformational changes in glycogen  phosphorylase. Ribbon diagrams of one subunit of the dimeric  enzyme glycogen phosphorylase b (a) in the T state in the absence of  allosteric effectors and (b) in the R state with bound AMR The view  is of the lower ( orange ) subunit in Fig. 12-15 rotated by —45° about  a horizontal axis. The tower helix is blue; the N-terminal helix is cyan;  the N-terminal residues that change conformation on AMP binding are  red; and residues 282 to 286, the 280s loop, which in the R state  are disordered and hence not seen, are green. Of the groups that  are shown in space-filling representation, the side chain of Ser 14,  the phosphorylation site, and AMP, which binds only to the R state  enzyme, have C green, N blue, 0 red, and P orange; the side chain of    Arg 569, which reorients in the T —» R transition so as to interact with  the substrate phosphate, has C cyan and N blue; and sulfate ions  bound near Ser 14 and at the active site of the R state enzyme,  which mimic the sterically similar phosphate ions, have 0 red and S  orange. The PLP at the active site is drawn in stick form with C  magenta, N blue, 0 red, and P orange. [Based on X-ray structures by  Louise Johnson, Oxford University, U.K. PDBids 8GPB and  7GPB.] Q See Kinemage Exercises 14-2 and 14-3.    Consider the properties of a phosphate group to explain  why protein phosphorylation can trigger dramatic conforma¬  tional changes.    FIG. 12-17 The control of glycogen phosphorylase activity. The enzyme may  assume the enzymatically inactive T conformation {above) or the catalytically active R form  {below). The conformation of phosphorylase b is allosterically controlled by the effectors  AMP, ATP, and G6P and is mostly in the T state under physiological conditions. In contrast,  the phosphorylated form of the enzyme, phosphorylase a, is unresponsive to these effectors  and is mostly in the R state unless there is a high level of glucose. Thus, under usual  physiological conditions, the enzymatic activity of glycogen phosphorylase is largely  determined by its rates of phosphorylation and dephosphorylation.    the 282—284 loop no longer blocks the active site, thereby permitting the en¬  zyme to bind substrate and efficiently catalyze the phosphorolysis of glycogen.   The phosphorylation ofSer 14 promotes phosphorylases T (inactive) —» R (ac¬  tive) conformational change. Moreover, these different enzymatic forms respond  to different allosteric effectors (Fig. 12-17). Thus ATP and glucose-6-phos-  phate [G6P; to which the G1P product of the glycogen phosphorylase reac¬  tion is converted (Section 16-1C)] preferentially bind to the T state of  phosphorylase b and, in doing so, inactivate the enzyme, whereas AMP pref¬  erentially binds to the R state of phosphorylase b and hence activates it. In  contrast, phosphorylase as only allosteric effector is glucose, which binds to  the enzyme’s T state and inactivates the enzyme. Note that ATP, G6P, and glu¬  cose are present in relatively high concentrations in muscle under conditions  of low exertion, a state when glycogen breakdown would be superfluous,  whereas AMP is present in relatively high concentration in muscles under con¬  ditions of high exertion, a state when the G1P product of the phosphorylase  reaction helps fuel muscle contraction.   The phosphate group, with its double negative charge (a property not  shared with naturally occurring amino acid residues) and its covalent attach¬  ment to a protein, can induce dramatic conformational changes. In phospho¬  rylase a , the Ser 14 phosphate group forms ion pairs with two cationic Arg side  chains, thereby linking the active site, the subunit interface, and the N-  terminal region, the latter having undergone a large conformational shift from  its position in the T-state enzyme (Fig. 12-1 6a). These binding interactions  cause glycogen phosphorylases tower helices (Figs. 12-15 and 12-16) to tilt and  pull apart so as to pack more favorably, which in turn triggers a quaternary T  —» R transition, which largely consists of an ~ 10° relative rotation of the two  subunits. Thus the Ser 14-phosphate group functions as a sort of internal allosteric  effector that shifts the enzymes T R equilibrium in favor of the R state.    Cascades of Protein Kinases Enable Sensitive Responses to Metabolic  Needs. The enzymes that catalyze the phosphorylation and dephosphoryla¬  tion of glycogen phosphorylase are named phosphorylase kinase and phos-  phoprotein phosphatase (Fig. 12-17). The activities of these latter enzymes  are themselves controlled, both allosterically and by phosphorylation/dephos-  phorylation. As we will see in Section 13-2, protein kinases and protein phos¬  phatases are often linked in series (i.e., protein kinase X phosphorylates protein  kinase Y, which phosphorylates protein kinase Z). Such cascade arrangements  have far greater capacities for signal amplification than simple allosteric en¬  zymes, because in a cascade, a small fractional change in effector concentra¬  tion can cause a larger fractional change in enzyme activity. In addition, the  cascade system offers greater flexibility in responding to a large number of  allosteric effectors because the activity of each protein in the cascade may be  influenced by different sets of effectors. Indeed, just such cascades permit  glycogen phosphorylase to respond with great sensitivity to the metabolic  needs of the organism.   • Compare and contrast the actions of an  allosteric effector, a competitive enzyme  inhibitor, and a noncompetitive inhibitor.   • Explain the structural basis for cooperative  substrate binding and allosteric control in  ATCase.   • Why are such allosteric enzymes com¬  posed of more than one catalytic subunit?   • Describe how phosphorylation and de¬  phosphorylation control the activity of  glycogen phosphorylase.   • List some advantages of phosphorylation/  dephosphorylation cascade systems over  simple allosteric regulation.    • Structure-based design and combinatorial chemistry approaches are used to dis¬  cover new drugs.   • A drug’s effectiveness and safety are tested in clinical trials.   • Cytochrome P450 may modify a drug to alter its bioavailability or toxicity.    T The use of drugs to treat various maladies has a long history, but the mod¬  ern pharmaceutical industry, which is based on science (rather than tradi¬  tion or superstition), is a product of the twentieth century. For example, at  the beginning of the twentieth century, only a handful of drugs, apart from  folk medicines, were known, including digitalis, a heart stimulant from the  foxglove plant (Box 10-3); quinine (Fig. 12-18), obtained from the bark and  roots of the Cinchona tree, which was used to treat malaria; and mercury,  which was used to treat syphilis—a cure that was often worse than the dis¬  ease. Almost all drugs in use today were discovered and developed in the past  four decades. The majority of drugs act by modifying the activity of a recep¬  tor protein, with enzyme inhibitors constituting the second largest class of  drugs. Indeed, the techniques of enzyme kinetics have proved to be invaluable  for evaluating drug candidates.    FIG. 1 2-1 8 Quinine and chloroquine. These two compounds, which share a quino¬  line ring system, are effective antimalarial agents. The Plasmodium parasite multiplies within  red blood cells, where it proteolyzes hemoglobin to meet its nutritional needs. This process  releases heme, which in its soluble form is toxic to the parasite. The parasite sequesters  the heme in a crystalline (nontoxic) form. Quinine and chloroquine pass through cell mem¬  branes and inhibit heme crystallization in the parasite.   How are new drugs discovered? Nearly all drugs that have been in use for over  15 years were discovered by screening large numbers of compounds—either  synthetic compounds or those derived from natural products (often plants used  in folk remedies). Initial screening involves in vitro assessment of, for exam¬  ple, the degree of binding of a drug candidate to an enzyme that is implicated  in a disease of interest, that is, determining its value. Later, as the number  of drug candidates is winnowed down, more sensitive screens such as testing  in animals are employed.   A drug candidate that exhibits a desired effect is called a lead compound  (pronounced leed). A good lead compound binds to its target protein with a  dissociation constant (for an enzyme, an inhibition constant) of less than 1 |ulM.  Such a high affinity is necessary to minimize a drugs less specific binding  to other macromolecules in the body and to ensure that only low doses of  the drug need be taken.   A lead compound is used as a point of departure to design more efficacious  compounds. Even minor modifications to a drug candidate can result in ma¬  jor changes in its pharmacological properties. Thus, substitution of methyl,  chloro, hydroxyl, or benzyl groups at various places on a lead compound may  improve its action. For most drugs in use today, 5000 to 10,000 related com¬  pounds were typically synthesized and tested. Drug development is a system¬  atic and iterative process in which the most promising derivatives of the lead  compound serve as the starting points for the next round of derivatization and  testing.   Structure-Based Drug Design Accelerates Drug Discovery. Since the mid  1980s, dramatic advances in the speed and precision with which a macromol-  ecular structure can be determined by X-ray crystallography and NMR  (Section 6-2A) have enabled structure-based drug design, a process that  greatly reduces the number of compounds that need be synthesized in a drug  discovery program. As its name implies, structure-based drug design (also  called rational drug design) uses the structure of a receptor or enzyme in  complex with a drug candidate to guide the development of more efficacious  compounds. Such a structure will reveal, for example, the positions of the hy¬  drogen bond donors and acceptors in the binding site as well as cavities in the  binding site into which substituents might be placed on a drug candidate to in¬  crease its binding affinity. These direct visualization techniques are usually sup¬  plemented with molecular modeling tools such as the computation of the  minimum energy conformation of a proposed derivative, quantum mechanical  calculations that determine its charge distribution and hence how it would in¬  teract electrostatically with the protein, and docking simulations in which an in¬  hibitor candidate is computationally modeled into the binding site on the receptor  to assess potential interactions (see Bioinformatics Project 6). A structure-based  approach was used to develop the analgesics (pain relievers) Celebrex and Vioxx  (Section 20-6C) and to develop drugs to treat HIV infection (Box 12-3).   Combinatorial Chemistry and High-Throughput Screening Are Useful Drug  Discovery Tools. As structure-based methods were developed, it appeared that  they would become the dominant mode of drug discovery. However, the re¬  cent advent of combinatorial chemistry techniques to rapidly and inexpen¬  sively synthesize large numbers of related compounds combined with the  development of robotic high-throughput screening techniques has caused the  drug discovery “pendulum” to again swing toward the “make-many-com-  pounds-and-see-what-they-do” approach. If a lead compound can be synthe¬  sized in a stepwise manner from several smaller modules, then the substituents  on each of these modules can be varied in parallel to produce a library of re¬  lated compounds (Fig. 12-19).   FIG. 1 2-19 The combinatorial synthesis of arylidene diamides. If ten different  variants of each R group are used in the synthesis, then 1000 different derivatives  will be synthesized.    A variety of synthetic techniques have been developed that permit the  combinatorial synthesis of thousands of related compounds in a single proce¬  dure. Thus, whereas investigations into the importance of a hydrophobic group  at a particular position in a lead compound might previously have prompted  the individual syntheses of only the ethyl, propyl, and benzyl derivatives of  the compound, the use of combinatorial synthesis would permit the genera¬  tion of perhaps 100 different groups at that position. This would far more  effectively map out the potential range of the substituent and possibly identify  an unexpectedly active analog.   B A Drug's Bioavailability Depends on How It Is Absorbed and  Transported in the Body   The in vitro development of an effective drug candidate is only the first step  in the drug development process. Besides causing the desired response in its  isolated target protein , a useful drug must be delivered in sufficiently high  concentration to this protein where it resides in the human body. For example, a  drug that is administered orally (the most convenient route) must surmount  a series of formidable barriers: (1) The drug must be chemically stable in the  highly acidic environment of the stomach and must not be degraded by di¬  gestive enzymes; (2) it must be absorbed from the gastrointestinal tract into  the bloodstream, that is, it must pass through several cell membranes; (3) it  must not bind too tightly to other substances in the body (e.g., lipophilic sub¬  stances tend to be absorbed by certain plasma proteins and by fat tissue);   (4) it must survive derivatization by the battery of enzymes, mainly in the  liver, that detoxify xenobiotics (foreign compounds; note that the intestinal  blood flow drains directly into the liver via the portal vein so that the liver  processes all orally ingested substances before they reach the rest of the body);   (5) it must avoid rapid excretion by the kidneys; (6) it must pass from the  capillaries to its target tissue; (7) if it is targeted to the brain, it must cross  the blood-brain barrier, which blocks the passage of most polar substances;  and (8) if it is targeted to an intracellular protein, it must pass through the  plasma membrane and possibly one or more intracellular membranes.   The ways in which a drug interacts with the barriers listed above are known  as its pharmacokinetics. Thus, the bioavailability of a drug (the extent to  which it reaches its site of action, which is usually taken to be the systemic  circulation) depends on both the dose given and its pharmacokinetics. The  most effective drugs are usually a compromise; they are neither too lipophilic nor  too hydrophilic. In addition, their pA" values are usually in the range 6 to 8 so  that they can readily assume both their ionized and unionized forms at phys¬  iological pH’s. This permits them to cross cell membranes in their unionized  form and to bind to their target protein in their ionized form.   Above all else, a successful drug candidate must be safe and efficacious in hu¬  mans. Tests for these properties are initially carried out in animals, but since    humans and animals often react quite differently to a drug, it must ultimately  be tested in humans through clinical trials. In the United States, clinical tri¬  als are monitored by the Food and Drug Administration (FDA) and have three  increasingly detailed (and expensive) phases:   This phase is primarily designed to test the safety of a drug  candidate but is also used to determine its dosage range and  the optimal dosage method (e.g., oral versus injected) and fre¬  quency. It is usually carried out on a small number (20-100)  of normal, healthy volunteers, but in the case of a drug can¬  didate known to be highly toxic (e.g., a cancer chemothera¬  peutic agent), it is carried out on volunteer patients with the  target disease.   This phase mainly tests the efficacy of the drug against the  target disease in 100 to 500 volunteer patients but also re¬  fines the dosage range and checks for side effects. The effects  of the drug candidate are usually assessed via single blind  tests, procedures in which the patient is unaware of whether  he or she has received the drug or a control substance. Usually  the control substance is a placebo (an inert substance with  the same physical appearance, taste, etc., as the drug being  tested) but, in the case of a life-threatening disease, it is an  ethical necessity that the control substance be the best avail¬  able treatment against the disease.   This phase monitors adverse reactions from long-term use as  well as confirming efficacy in 1000 to 5000 patients. It pits  the drug candidate against control substances through the  statistical analysis of carefully designed double blind tests,  procedures in which neither the patients nor the clinical in¬  vestigators know whether a given patient has received the  drug or a control substance. This is done to minimize bias in  the subjective judgments the investigators must make.   Currently, only about 5 drug candidates in 5000 that enter preclinical  trials reach clinical trials. Of these, only one, on average, is ultimately approved  for clinical use, with the majority failing in Phase I trials. In recent years, the  preclinical portion of a drug discovery process has averaged ~3 years to com¬  plete, whereas successful clinical trials have usually required an additional 7 to  10 years. These successive stages of the drug discovery process are increasingly  expensive so that successfully bringing a drug to market costs, on average,  around $300 million.   The most time-consuming and expensive aspect of a drug development  program is identifying a drug candidates rare adverse reactions. Nevertheless,  it is not an uncommon experience for a drug to be brought to market only  to be withdrawn some months or years later when it is found to have caused  unanticipated life-threatening side effects in as few as 1 in 10,000 individuals  (the post-marketing surveillance of a drug is known as its Phase IV clinical  trial). For example, in 1997, the FDA withdrew its approval of the drug  fenfluramine (fen; at left ), which it had approved in 1973 for use as an ap¬  petite suppressant in short-term (a few weeks) weight-loss programs.  Fenfluramine had become widely prescribed, often for extended periods, to¬  gether with another appetite suppressant, phentermine (phen; approved in  1959; at left), a combination known as fen-phen (although the FDA had not  approved of the use of the two drugs in combination, once it approves a drug  for some purpose, a physician may prescribe it for any other purpose). The  withdrawal of fenfluramine was prompted by over 100 reports of heart valve  damage in individuals (mostly women) who had taken fen-phen for an average  of 12 months (phentermine was not withdrawn because the evidence indicated   that fenfluramine was the responsible agent). This rare side effect had not been  observed in the clinical trials of fenfluramine, in part because it is such an un¬  usual type of drug reaction that it had not been screened for.   More recently (2004), the widely prescribed analgesic Vioxx was with¬  drawn from use due to its previously undetected cardiac side effects, although  the closely related analgesic Celebrex remains available (Section 20-6C). The  diabetes drug rosiglitazone (Avandia), which is associated with an increased  risk of heart attack and stroke, likewise illustrates the failure of clinical trials  to detect small increases in relatively common diseases, particularly when pa¬  tients (e.g., diabetics) are already ill.   Why is it that a drug that is well tolerated by the majority of patients can  pose a danger to others? Differences in reactions to drugs arise from genetic dif¬  ferences among individuals as well as differences in their disease states, other drugs  they are taking, age, sex, and environmental factors. The cytochromes P450,  which function in large part to detoxify xenobiotics and which participate in  the metabolic clearance of the majority of drugs in use, provide instructive ex¬  amples of these phenomena.   The cytochromes P450 constitute a superfamily of heme-containing en¬  zymes that occur in nearly all living organisms, from bacteria to mammals  [their name arises from the characteristic 450-nm peak in their absorption  spectra when reacted in their Fe(II) state with CO]. The human genome en¬  codes 57 isozymes of cytochrome P450, around one-third of which occur in  the liver (P450 isozymes are named by the letters “CYP” followed by a num¬  ber designating its family, an uppercase letter designating its subfamily, and  often another number; e.g., CYP2D6). These mono oxygenases (Fig. 12-20),  which in animals are embedded in the endoplasmic reticulum membrane, cat¬  alyze reactions of the sort   RH + 0 2 + 2H + + 2e~ ROH + H 2 0   The electrons (e ) are supplied by NADPH, which passes them to cy¬  tochrome P450s heme prosthetic group via the intermediacy of the enzyme  NADPH-P450 reductase. Here RH represents a wide variety of usually  lipophilic compounds for which the different cytochromes P450 are specific.  They include polycyclic aromatic hydrocarbons [PAHs; frequently carcino¬  genic (cancer-causing) compounds that are present in tobacco smoke, broiled  meats, and other pyrolysis products], polycyclic biphenyls (PCBs; which were  widely used in electrical insulators and as plasticizers and are also carcino¬  genic), steroids (in whose syntheses cytochromes P450 participate), and many  different types of drugs. The xenobiotics are thereby converted to a more wa¬  ter-soluble form, which aids in their excretion by the kidneys. Moreover, the  newly generated hydroxyl groups are often enzymatically conjugated (cova¬  lently linked) to polar substances such as glucuronic acid (Section 8-1C),  glycine, sulfate, and acetate, which further enhances aqueous solubility. The  many types of cytochromes P450 in animals, which have different substrate  specificities (although these specificities tend to be broad and hence often  overlap), are thought to have arisen in response to the numerous toxins that  plants produce, presumably to discourage animals from eating them (other  P450s function to catalyze specific biosynthetic reactions).   Drug—drug interactions are often mediated by cytochromes P450. For ex¬  ample, if drug A is metabolized by or otherwise inhibits a cytochrome P450  isozyme that metabolizes drug B, then coadministering drugs A and B will  cause the bioavailability of drug B to increase above the value it would have  had if it alone had been administered. This phenomenon is of particular   FIG. 1 2-20 X-Ray structure of the human  cytochrome P450 CYP2C9 in complex with  the blood clotting inhibitor warfarin  (Coumadin). A cutaway diagram of the  enzyme’s active site region drawn with its  surface purple and with its polypeptide backbone  in ribbon form colored in rainbow order from  N-terminus (blue) to C-terminus (red). The heme  (seen edgewise), the Cys side chain that axially  ligands the heme Fe atom, and the warfarin are  shown in ball-and-stick form with C gray, N blue,  0 red, S yellow, and Fe orange. [Image courtesy  of Astex Therapeutics Limited. PDBid 10G5.]   Chapter 12 Enzyme Kinetics, Inhibition, and Control concern if drug B has a low therapeutic index (the ratio of the dose of the   drug that produces toxicity to that which produces the desired effect).  Conversely, if, as is often the case, drug A induces the increased expression of  the cytochrome P450 isozyme that metabolizes it and drug B, then coadminis¬  tering drugs A and B will reduce drug B s bioavailability, a phenomenon that  was first noted when certain antibiotics caused oral contraceptives to lose their  efficacy. Moreover, if drug B is metabolized to a toxic product, its increased  rate of reaction may result in an adverse reaction. Environmental pollutants  such as PAHs and PCBs are also known to induce the expression of specific  cytochrome P450 isozymes and thereby alter the rates at which certain drugs  are metabolized. Finally, some of these same effects may occur in patients with  liver disease, as well as arising from age-based, gender-based, and individual  differences in liver physiology.   Although many cytochromes P450 presumably evolved to detoxify and/or  help eliminate harmful substances, in several cases they have been shown to  participate in converting relatively innocuous compounds to toxic agents. For  example, acetaminophen, a widely used analgesic and antipyretic (fever re¬  ducer) is quite safe when taken in therapeutic doses (1.2 g/day for an adult)  but in large doses (>10 g) is highly toxic. This is because, in therapeutic  amounts, 95% of the acetaminophen present is enzymatically glucuronidated  or sulfated at its —OH group to the corresponding conjugates, which are read¬  ily excreted. The remaining 5% is converted, through the action of a  cytochrome P450 (CYP2E1), to acetimidoquinone (Fig. 12-21), which is then    FIG. 1 2-21 The metabolic reactions of acetaminophen. An overdose of  acetaminophen leads to the buildup of acetimidoquinone, which is toxic.    conjugated with glutathione (Section 4-3B). However, when acetaminophen is  taken in large amounts, the glucuronidation and sulfation pathways become  saturated and hence the cytochrome P450-mediated pathway becomes increas¬  ingly important. If hepatic (liver) glutathione is depleted faster than it can be  replaced, acetimidoquinone, a reactive compound, instead conjugates with the  sulfhydryl groups of cellular proteins, resulting in often fatal hepatotoxicity.   Many of the cytochromes P450 in humans are unusually polymorphic,  that is, there are several common alleles (variants) of the genes encoding each  of these enzymes in the human population. Alleles that cause diminished, en¬  hanced, and qualitatively altered rates of drug metabolism have been charac¬  terized for many of the cytochromes P450. The distributions of these various  alleles differs markedly among ethnic groups and hence probably arose to per¬  mit each group to cope with the toxins in its particular diet.   Polymorphism in a given cytochrome P450 results in differences between  individuals in the rates at which they metabolize certain drugs. For instance, in  cases in which a cytochrome P450 variant has absent or diminished activity, oth¬  erwise standard doses of a drug that the enzyme normally metabolizes may cause  the bioavailability of the drug to reach toxic levels. Conversely, if a particular  P450 enzyme has enhanced activity (usually because the gene encoding it has  been duplicated one or more times), higher than normal doses of a drug that  the enzyme metabolizes would have to be administered to obtain the required  therapeutic effect. However, if the drug is metabolized to a toxic product, this  may result in an adverse reaction. Several known P450 variants have altered sub¬  strate specificities and hence produce unusual metabolites, which also may cause  harmful side effects.   Experience has amply demonstrated that there is no such thing as a drug that  is entirely free of adverse reactions. However, as the enzymes and their variants  that participate in drug metabolism are characterized and as rapid and inexpen¬  sive genotyping methods are developed, it is becoming possible to tailor drug  treatment to an individuals genetic makeup rather than to the population as a  whole. This rapidly developing area of study is called pharmacogenomics.   • Compare structure-based drug design and  combinatorial chemistry as tools for devel¬  oping drug candidates.   • Summarize the purpose of phases I  through III of a clinical trial.   • Indicate how drugs that are well tolerated  by the majority of the population cause  adverse reactions in certain individuals.   • Elementary chemical reactions may be first order, second order,  or, rarely, third order. In each case, a rate equation describes the  progress of the reaction as a function of time.   • The Michaelis-Menten equation describes the relationship be¬  tween initial reaction velocity and substrate concentration under  steady state conditions.   • K m is the substrate concentration at which the reaction velocity is  half-maximal. The value of k C2X /K M indicates an enzyme’s catalytic  efficiency.   • Kinetic data can be plotted in double-reciprocal form to deter¬  mine K m and Enax-   • Bisubstrate reactions are classified as sequential (single displace¬  ment) or Ping Pong (double displacement). A sequential reaction  may proceed by an Ordered or Random mechanism.   • Reversible inhibitors reduce an enzyme’s activity by binding to  the substrate-binding site (competitive inhibition), to the    enzyme-substrate complex (uncompetitive inhibition), or to both  the enzyme and the enzyme-substrate complex (mixed inhibition).   • The activity of ATCase is increased by ATP and decreased by CTP,  which alter the conformation of the catalytic sites by stabilizing  the R and the T states of the enzyme, respectively.   • The enzymatic activity of glycogen phosphorylase is controlled by  its phosphorylation/dephosphorylation as well as by the influence  of allosteric effectors.   • An enzyme inhibitor can be developed for use as a drug through  structure-based and/or combinatorial methods. It must then be  tested for safety and efficacy in clinical trials.   • Adverse reactions to drugs and drug-drug interactions are often  mediated by a cytochrome P450.    Ping Pong reaction 367  double-displacement  reaction 367  inhibitor 368  inactivator 368  competitive inhibition 368  product inhibition 369  transition state analog 369  Ki 369  Kffi 372    1. Consider the nonenzymatic elementary reaction A —> B. When  the concentration of A is 20 mM, the reaction velocity is mea¬  sured as 5 |JiM B produced per minute, (a) Calculate the rate  constant for this reaction, (b) What is the molecularity of the  reaction?   2. The hypothetical elementary reaction 2A —» B + C has a rate  constant of 10 -6 M 1 • s \ What is the reaction velocity when  the concentration of A is 10 mM?   3. If there is 10 fxmol of the radioactive isotope 32 P (half-life 14  days) at t = 0, how much 32 P will remain at (a) 7 days, (b) 14  days, (c) 21 days, and (d) 70 days?   4. Calculate the half-life, in years, for the reaction 2 X —> Y when  the starting concentration of X is 6 |JiM and the rate constant is  3.6 X 1(T 3 M -1 • s -1 .   3. From the reaction data below, determine whether the reaction is  first order or second order and calculate the rate constant.    6. From the reaction data below, determine whether the reaction is  first order or second order and calculate the rate constant.    7. For an enzymatic reaction, draw curves that show the appropri¬  ate relationships between the variables in each plot below.    8. Explain why it is usually easier to calculate an enzyme’s reaction  velocity from the rate of appearance of product rather than the  rate of disappearance of a substrate.   10. Identify the enzymes in Table 12-1 whose catalytic efficiencies are  near the diffusion-controlled limit.   11. Calculate K M and V max from the following data:    12. Explain why each of the following data sets from a  Lineweaver-Burk plot are not individually ideal for determining  K m for an enzyme-catalyzed reaction that follows Michaelis-  Menten kinetics.    13. Is it necessary for measurements of reaction velocity to be ex¬  pressed in units of concentration per time (M • s \ for example)  in order to calculate an enzyme’s K M }   14. Is it necessary to know [E] T in order to determine (a) K Mi (b)  Knax> or (c) ^car   15. You are trying to determine the K M for an enzyme. Due to a lab  mishap, you have only two usable data points:    Use these data to calculate an approximate value for K M . Is this  value likely to be an overestimate or an underestimate of the true  value? Explain.   16. You are attempting to determine K M by measuring the reaction  velocity at different substrate concentrations, but you do not re¬  alize that the substrate tends to precipitate under the experimen¬  tal conditions you have chosen. How would this affect your  measurement of K M ?    17. You are constructing a velocity versus [substrate] curve for an en¬  zyme whose K m is believed to be about 2 |jlM. The enzyme con¬  centration is 200 nM and the substrate concentrations range  from 0.1 |JiM to 10 |JiM. What is wrong with this experimental  setup and how could you fix it?   18. Enzyme X and enzyme Y catalyze the same reaction and exhibit  the v 0 versus [S] curves shown below. Which enzyme is more  efficient at low [S]? Which is more efficient at high [S] ?   19. The K m for the reaction of chymotrypsin with A^-acetylvaline  ethyl ester is 8.8 X 10 2 M, and the K M for the reaction of chy¬  motrypsin with A^-acetyltyrosine ethyl ester is 6.6 X 10 4 M.  (a) Which substrate has the higher apparent affinity for the en¬  zyme? (b) Which substrate is likely to give a higher value for V ma J   20. Enzyme A catalyzes the reaction S —> P and has a K M of 50 |JiM  and a V max of 100 nM • s \ Enzyme B catalyzes the reaction  S —^ Q_ and has a K M of 5 mM and a V max of 120 nM • s \  When 100 |JiM of S is added to a mixture containing equivalent  amounts of enzymes A and B, after 1 minute which reaction  product will be more abundant: P or Q?   21. In a bisubstrate reaction, a small amount of the first product P  is isotopically labeled (P*) and added to the enzyme and the first  substrate A. No B or Q is present. Will A (= P—X) become  isotopically labeled (A*) if the reaction follows a Ping Pong  mechanism?    22. In a bisubstrate reaction, a small amount of the first product P  is isotopically labeled (P*) and added to the enzyme and the first  substrate A. No B or Q is present. Will A (= P—X) become  isotopically labeled (A*) if the reaction follows a Sequential  mechanism?   23. How would diisopropylphosphofluoridate (DIPF; Section 11-5A)  affect the apparent K M and V max of a sample of chymotrypsin?   24. Molecule A is the substrate for enzyme X. Which is more likely  to be a competitive inhibitor of enzyme X: molecule B or mol¬  ecule C? Explain.    25. Based on some preliminary measurements, you suspect that a sam¬  ple of enzyme contains an irreversible enzyme inhibitor. You decide  to dilute the sample 100-fold and remeasure the enzyme’s activity.  What would your results show if an irreversible inhibitor is present?   26. For the same enzyme sample described in Problem 25, what  would your results show if a reversible inhibitor is present?   27. Determine the type of inhibition of an enzymatic reaction from  the following data collected in the presence and absence of the  inhibitor.    28. Estimate K\ for a competitive inhibitor when [I] = 5 mM gives  an apparent value of K M that is three times the K M for the un¬  inhibited reaction.   29. For an enzyme-catalyzed reaction, the presence of 5 nM of a re¬  versible inhibitor yields a V max value that is 80% of the value in  the absence of the inhibitor. The K M value is unchanged, (a) What  type of inhibition is likely occurring? (b) What proportion of the  enzyme molecules have bound inhibitor? (c) Calculate the inhibi¬  tion constant.   30. Sphingosine-1-phosphate (SPP) is important for cell survival. The  synthesis of SPP from sphingosine and ATP is catalyzed by the  enzyme sphingosine kinase. An understanding of the kinetics of  the sphingosine kinase reaction may be important in the devel¬  opment of drugs to treat cancer. The velocity of the sphingosine  kinase reaction was measured in the presence and absence of  ^r^-sphingosine, a stereoisomer of sphingosine that inhibits the  enzyme. The results are shown below.   (a) What are the apparent K M and V max values in the presence  and absence of the inhibitor?   1. Dihydrofolate Reductase. Examine the structure of an enzyme  with an inhibitor bound to it.    2. HIV Protease. Compare the structures of complexes containing  HIV protease and an inhibitor.   Project 7 Enzyme Commission Classes and  Catalytic Site Alignments with PyMOL   1. The Enzyme Commission and IUPAC classification of  enzymes. Discover the systematic nature of enzyme classifica¬  tion, brought to you by the same organization (IUPAC) that de¬  termines systematic nomenclature for organic molecules.   2. Exploring the Catalytic Site Atlas. The Catalytic Site Atlas  is an annotated database of enzyme active sites. You’ll learn  to find the active sites of enzymes that you learn about in  class.   The combination of the PyMOL molecular graphics environment  and the ProMOL plug-in for PyMOL will enable you to study  and align the catalytic sites of many different enzymes.   Case 7 A Storage Protein from Seeds of Brassica  nigra Is a Serine Protease Inhibitor  Focus concept: Purification of a novel seed storage protein allows  sequence analysis and determination of the protein’s secondary  and tertiary structure.   Case 12 Production of Methanol in Ripening Fruit  Focus concept: The link between the production of methanol in  ripening fruit and the activity of pectin methylesterase, the enzyme  responsible for methanol production, is examined in wild-type and  transgenic tomato fruit.   Case 13 Inhibition of Alcohol Dehydrogenase  Focus concept: The inhibition of alcohol dehydrogenase by a for-  mamide compound is examined.   Focus concept: Site-directed mutagenesis is used to create mutant  creatine kinase enzymes so that the role of a single reactive cysteine  in binding and catalysis can be assessed.   Focus concept: The purification and kinetic analysis of an enzyme  that produces a product important in cell survival is the focus of  this study.    What kind of inhibitor is warfarin  (shown in Fig. 1 2-20) and what enzyme does it inhibit? What is the  physiological goal of warfarin therapy? Why do individuals vary in  their response to a standard dose of warfarin? Why can a patient’s  genotypic information help a physician choose an effective dose?   Biswanger, H., Enzyme Kinetics: Principles and Methods (2nd ed.),  Wiley-VCH (2008).   Cornish-Bowden, A., Fundamentals of Enzyme Kinetics (revised ed.), Port¬  land Press (1995). [A lucid and detailed account of enzyme kinetics.]   Fersht, A., Structure and Mechanism in Protein Science: A Guide to Enzyme  Catalysis and Protein Folding, W.H. Freeman (1999).   Segel, I.H., Enzyme Kinetics, Wiley-Interscience (1993). [A detailed and  understandable treatise providing full explanations of many aspects  of enzyme kinetics.]   Jin, L., Stec, B., Lipscomb, W.N., and Kantrowitz, E.R., Insights into the  mechanisms of catalysis and heterotropic regulation of Escherichia  coli aspartate transcarbamoylase based upon a structure of the en¬  zyme complexed with the bisubstrate analogue A-phosphonacetyl-  L-aspartate at 2.1 A, Proteins 37, 729-742 (1999).   Johnson, L.N. and Lewis, R.J., Structural basis for control by phospho¬  rylation, Chem. Rev. 101, 2209-2242 (2001).   Lipscomb, W.N., Structure and function of allosteric enzymes, Chemtracts —  Biochem. Mol. Biol. 2, 1-15 (1991).   Perutz, M., Mechanisms of Cooperativity and Allosteric Regulation in  Proteins, Cambridge University Press (1990).   Corey, E.J., Czako, B., and Kurd, L., Molecules and Medicine, Wiley  (2007). [Discusses the discovery, application, and mode of action of  numerous drug molecules.]    Corson, T.W. and Crews, C.M., Molecular understanding and modern  application of traditional medicines: triumphs and trials, Cell 130,  769-774 (2007).   Furge, L.L. and Guengerich, F.P., Cytochrome P450 enzymes in drug me¬  tabolism and chemical toxicology, Biochem. Mol. Biol. Educ. 34,  66-74 (2006).   Jorgenson, W.L., The many roles of computation in drug discovery, Sci¬  ence 303, 1813-1818 (2004).   Katzung, B.G. (Ed.), Basic & Clinical Pharmacology (10th ed.), McGraw-  Hill (2007).   Ohlstein, E.H., Ruffolo, R.R., Jr., and Elliott, J.D., Drug discovery in  the next millennium, Annu. Rev. Pharmacol. Toxicol. 40, 177-191  ( 2000 ).   White, R.E., High-throughput screening in drug metabolism and  pharmokinetic support of drug discovery, Annu. Rev. Pharmacol.  Toxicol. 40, 133-157 (2000).   Williams, P.A., Cosme, J., Ward, A., Angove, H.C., Vinkovic, D.M., and  Jhoti, H., Crystal structure of human cytochrome P450 2C9 with  bound warfarin, Nature 424, 464-468 (2003).   Wlodawer, A. and Vondrasek, J., Inhibitors of HIV-1 protease: A major  success of structure-assisted drug design, Annu. Rev. Biophys. Biomol.  Struct. 27 ', 249-284 (1998). [Reviews the development of some  HIV-1 protease inhibitors.]   C Steroid Hormones Regulate a Wide Variety of  Metabolic and Sexual Processes  D Growth Hormone Binds to Receptors in Muscle,  Bone, and Cartilage   A Receptor Tyrosine Kinases Transmit Signals  across the Cell Membrane  B Kinase Cascades Relay Signals to the Nucleus  C Some Receptors Are Associated with  Nonreceptor Tyrosine Kinases  D Protein Phosphatases Are Signaling Proteins in  Their Own Right   A G Protein-Coupled Receptors Contain Seven  Transmembrane Helices  B Heterotrimeric G Proteins Dissociate on  Activation   C Adenylate Cyclase Synthesizes cAMP to  Activate Protein Kinase A  D Phosphodiesterases Limit Second Messenger  Activity   A Ligand Binding Results in the Cytoplasmic  Release of the Second Messengers IP 3  and Ca 2+   B Calmodulin Is a Ca 2+ -Activated Switch  C DAG Is a Lipid-Soluble Second Messenger That  Activates Protein Kinase C  D Epilog: Complex Systems Have Emergent  Properties    Vanillin, an odorant extracted from vanilla beans and other sources, binds to receptors on  olfactory neurons and triggers a series of intracellular events, including heterotrimeric G  protein activation and second messenger production, that ultimately generate an  electronic signal to the brain. [© Martin Harvey/Peter Arnold, Inc.]    Living things coordinate their activities at every level of their organization  through complex biochemical signaling systems. Intercellular signals are me¬  diated by chemical messengers known as hormones and, in higher animals,  by neuronally transmitted electrochemical impulses. Intracellular communica¬  tions are maintained by the synthesis or alteration of a great variety of differ¬  ent substances that are often integral components of the processes they control.  For example, metabolic pathways, as we have seen (Section 12-3), are regu¬  lated by the feedback control of allosteric enzymes by metabolites in those  pathways or by the covalent modification of the enzymes. In this chapter we  consider the nature of chemical signals and how the signals are transmitted.   In general, every signaling pathway consists of a receptor protein that specif¬  ically binds a hormone or other ligand, a mechanism for transmitting the ligand¬  binding event to the cell interior, and a series of intracellular responses that may  involve the synthesis of a second messenger and/or chemical changes catalyzed by  kinases and phosphatases. These pathways often involve enzyme cascades , in  which a succession of events amplifies the signal.   We begin by discussing the functions of some representative human hor- Section 1 Hormones   mone systems. We then discuss the three major pathways whereby intercellu¬  lar signals are converted (transduced) to intracellular signals: those that  (1) involve receptor tyrosine kinases, (2) utilize heterotrimeric G proteins, and  (3) employ phosphoinositide cascades. Neurotransmission is discussed in  Section 10-2C.   • The pancreatic hormones insulin and glucagon help control fuel metabolism.   • Catecholamines produced by the adrenal medulla bind to a- and p-adrenergic  receptors on target cells.   • Steroid hormones regulate fuel metabolism, salt and water balance, and sexual  differentiation and function.   • Growth hormone exerts its effects by inducing dimerization of its receptor.   In higher animals, specialized ducdess endocrine glands (Fig. 13-1) synthe¬  size endocrine hormones that are released into the bloodstream in response  to external stimuli. These hormones are thereby carried to their target cells  (Fig. 13-2) in which they elicit a response. The human endocrine system se¬  cretes a wide variety of hormones that enable the body to:   1. Maintain homeostasis (a steady state; e.g., insulin and glucagon main¬  tain the blood glucose level within rigid limits during feast or famine).   2. Respond to a wide variety of external stimuli (such as the preparation  for “fight or flight” by epinephrine and norepinephrine).   3. Follow various cyclic and developmental programs (for instance, sex  hormones regulate sexual differentiation, maturation, the menstrual  cycle, and pregnancy).   Most hormones are either polypeptides, amino acid derivatives, or steroids, al¬  though there are important exceptions to this generalization. In any case, only  those cells with a specific receptor for a given hormone will respond to its presence  even though nearly all cells in the body may be exposed to the hormone. Hormonal  messages are therefore quite specifically addressed.   Although we discuss specific hormones and their function in many other  chapters, in this section we outline the activities of hormones produced by  some representative endocrine glands. These glands are not just a collection of  independent secretory organs but form a complex and highly interdependent    FIG. 1 3-1 The major glands of the  human endocrine system. Other tissues, such  as the intestines, also secrete endocrine  hormones.   FIG. 1 3-2 Endocrine signaling. Hormones  produced by endocrine cells reach their target  cells via the bloodstream. Only cells that display  the appropriate receptors can respond to the  hormones.   Chapter 13 Biochemical Signaling control network. Indeed, the secretion of many hormones is under feedback   control through the secretion of other hormones to which the original  hormone-secreting gland responds. The concentrations of circulating  hormones are typically measured using the radioimmunoassay developed by  Rosalyn Yalow (Box 13-1).   The pancreas is a large glandular organ, the bulk of which is an exocrine  gland dedicated to producing digestive enzymes—such as trypsin, chy-  motrypsin, RNase A, a-amylase, and phospholipase A 2 —that are secreted via  the pancreatic duct into the small intestine. However, ~1 to 2% of pancre¬  atic tissue consists of scattered clumps of cells known as islets of Langerhans,  which comprise an endocrine gland that functions to maintain energy home¬  ostasis. Pancreatic islets contain three types of cells, each of which secretes a  characteristic polypeptide hormone:   Insulin, which is secreted in response to high blood glucose levels, primarily  functions to stimulate muscle, liver, and adipose cells to store glucose for later  use by synthesizing glycogen, protein, and fat (Section 22-2). Glucagon, which  is secreted in response to low blood glucose, has essentially the opposite  effects: It stimulates the liver to release glucose through the breakdown of  glycogen (glycogenolysis; Section 16-1) and the synthesis of glucose from  noncarbohydrate precursors (gluconeogenesis, Section 16-4). It also stimu¬  lates adipose tissue to release fatty acids through lipolysis. Somatostatin, which  is also secreted by the hypothalamus, inhibits the release of insulin and  glucagon from their islet cells.   Polypeptide hormones, like other proteins destined for secretion, are ribo-  somally synthesized as prohormones, processed in the rough endoplasmic retic¬  ulum and Golgi apparatus to form the mature hormones, and then packaged  in secretory granules to await the signal for their release by exocytosis  (Sections 9-4D-F). The most potent physiological stimuli for the release of  insulin and glucagon are, respectively, high and low blood glucose concen¬  trations so that islet cells act as the body’s primary glucose sensors. However,  the release of the hormones is also influenced by the autonomic (involuntary)  nervous system and by hormones secreted by the gastrointestinal tract.    R = H Norepinephrine (noradrenalin)  R = CH 3 Epinephrine (adrenalin)   The adrenal glands consist of two distinct types of tissue: the medulla (core),  which is really an extension of the sympathetic nervous system (a part of the  autonomic nervous system), and the more typically glandular cortex (outer  layer). Here we consider the hormones of the adrenal medulla; those of the  cortex are discussed in the following section.   The adrenal medulla synthesizes two hormonally active catecholamines  (amine-containing derivatives of catechol, 1,2-dihydroxy benzene): norepineph¬  rine (noradrenalin) and its methyl derivative epinephrine (adrenalin; at left).  These hormones are synthesized from tyrosine as is described in Section 21-  6B and are stored in granules to await their exocytotic release under the con¬  trol of the sympathetic nervous system.   The biological effects of catecholamines are mediated by two classes of  plasma membrane receptors, the o i- and (3-adrenergic receptors (also known  as adrenoreceptors). These transmembrane glycoproteins were originally iden¬  tified on the basis of their varying responses to certain agonists (substances  that bind to a receptor so as to evoke a response) and antagonists (substances   Sussman Yalow was born on July 19, 1921,  in New York City. Neither of her parents had  the advantage of a high school education but  there was never a doubt that their two  children would make it through college. While  Rosalyn was at Hunter College, the college  for women in New York City’s college system  (now the City University of New York), Eve Curie had just  published the biography of her mother, Madame Marie Curie, a  “must-read” for every aspiring female scientist. Rosalyn’s goal at  that time was achieving a career in physics. Despite suggestions  from her family that becoming an elementary school teacher  would be more practical, she persisted.   In 1941, after graduating from college, Yalow received an offer  of a teaching assistantship in physics at the University of Illinois in  Champaign-Urbana. At the first meeting of the faculty of the  College of Engineering, she discovered that she was the only  woman among its 400 members. The dean of the faculty  congratulated her on her achievement and told her she was the  first woman there since 1917. The draft of young men into the  armed forces, even before America entered World War II, had  made possible her entrance into graduate school. On the first day  of graduate school she met Aaron Yalow, who was also beginning  graduate study in physics at Illinois and who in 1943 was to  become her husband.   In January 1945, Yalow received a Ph.D. in nuclear physics  and returned to New York as assistant engineer at the Federal  Telecommunications Laboratory—its only woman engineer. In 1946,  she returned to Hunter College to teach physics, not to women  but to returning veterans in a preengineering program. During  that time she became interested in the medical aspects of  radioisotopes. She joined the Bronx Veterans Administration (VA)  as a part-time consultant in December 1947. Even while teaching  full-time at Hunter, she equipped and developed the Radioisotope  Service at the VA hospital and initiated a variety of research    projects with several physicians. In January 1950, Yalow chose to  leave teaching and join the VA full time. That spring, Dr. Solomon  A. Berson joined the Service and began a 22-year partnership  that lasted until his death in 1972.   In their joint studies, Yalow and Berson concentrated on the  application of isotopes to clinical problems such as hormone  analysis. At the time, insulin was the hormone most readily  available in a highly purified form. In studying the reaction of  insulin with antibodies, Yalow and Berson recognized that they had  a tool with the potential for measuring the concentration of insulin  in a complex mixture such as blood. By 1959, they had developed  a practical method for quantifying insulin in human plasma, the  radioimmunoassay (RIA). RIA is now used to measure hundreds  of substances of biological interest.   The serum concentrations of insulin and other hormones are  extremely low, generally between 10“ 12 and 10 -7 M, so they  usually must be measured by indirect means. In RIAs, the  unknown concentration of a hormone, H, is determined by  measuring how much of a known amount of the radioactively  labeled hormone, H*, binds to a fixed quantity of anti-H antibody  in the presence of H. This competition reaction is easily calibrated  by constructing a standard curve indicating how much H* binds to  the antibody as a function of [Hj. The high binding affinity and  specificity of antibodies for their ligands gives RIAs the  advantages of great sensitivity and specificity.   By 1977, Yalow’s hospital was affiliated with the Mount Sinai  School of Medicine, and Yalow held the title of Distinguished  Service Professor. She was a member of the National Academy of  Sciences and was a recipient of numerous awards and honors,  including the 1977 Nobel Prize in Physiology or Medicine (Berson  died before this Nobel prize was awarded and so could not share  the prize) and the 1988 National Medal of Science. “The  excitement of learning separates youth from old age,” Rosalyn  Yalow has said. “As long as you’re learning, you’re not old.”*   Mostly abridged from Rosalyn Yalow’s autobiography, Les Prix Nobel. The Nobel Prizes  1977, Wilhelm Odelberg (Ed.), Nobel Foundation, 1978.   that bind to a receptor but fail to elicit a response, thereby blocking agonist  action). The (3- but not the a-adrenergic receptors, for example, are stimu¬  lated by isoproterenol but blocked by propranolol, whereas a- but not  (3-adrenergic receptors are blocked by phentolamine.    The a- and (3-adrenergic receptors, which occur on separate tissues in  mammals, generally respond differently and often oppositely to cate¬  cholamines. For instance, (3-adrenergic receptors stimulate glycogenolysis and  gluconeogenesis in liver (Sections 16-1 and 16-4), glycogenolysis in skeletal  muscle, lipolysis in adipose tissue, the relaxation of smooth (involuntary)   muscle in the bronchi and in the blood vessels supplying the skeletal (voluntary)  muscles, and increased heart action. In contrast, a-adrenergic receptors stim¬  ulate smooth muscle contraction in blood vessels supplying peripheral organs  such as skin and kidney, smooth muscle relaxation in the gastrointestinal tract,  and blood platelet aggregation. Most of these diverse effects are directed toward  a common end: the mobilization of energy resources and their shunting to where  they are most needed to prepare the body for action.   The tissue distributions of the a- and (3-adrenergic receptors and their vary¬  ing responses to different agonists and antagonists have important thera¬  peutic consequences. For example, propranolol is used to treat high blood  pressure and protects against heart attacks, whereas epinephrines bronchodila-  tor effects make it clinically useful in the treatment of asthma, a breathing  disorder caused by the inappropriate contraction of bronchial smooth muscle.   C Steroid Hormones Regulate a Wide Variety of Metabolic  and Sexual Processes   The adrenal cortex produces at least 50 different adrenocortical steroids . These  have been classified according to the physiological responses they evoke  (Section 9-IE):   1. The glucocorticoids affect carbohydrate, protein, and lipid metabo¬  lism in a manner nearly opposite to that of insulin and influence a  wide variety of other vital functions, including inflammatory reactions  and the capacity to cope with stress.   2. The mineralocorticoids largely function to regulate the excretion of  salt and water by the kidneys.   3. The androgens and estrogens affect sexual development and function.  They are made in larger quantities by the gonads.   Glucocorticoids, the most common of which is cortisol (also known as  hydrocortisone), and the mineralocorticoids, the most common of which is  aldosterone, are all C 2 i compounds (Fig. 9-11).   Steroids, being water insoluble, are transported in the blood in complex  with the glycoprotein transcortin and, to a lesser extent, with albumin. The  steroids spontaneously pass through the membranes of their target cells to the  cytosol, where they bind to their cognate steroid receptors. The steroid-  receptor complexes then migrate to the cell nucleus, where they function as  transcription factors to induce, or in some cases repress, the transcription of  specific genes (Section 28-3B). In this way, the glucocorticoids and the min¬  eralocorticoids influence the expression of numerous metabolic enzymes in  their respective target tissues. Thyroid hormones, which are also nonpolar,  function similarly. However, as we will see in the following sections, other hor¬  mones that are not lipid-soluble must bind to receptors on the cell surface in  order to trigger complex cascades of events within cells that ultimately influ¬  ence transcription as well as other cellular processes.   (testes in males , ovaries in females), in addition to producing sperm or ova, secrete  steroid hormones (androgens and estrogens) that regulate sexual differentiation , the  expression of secondary sex characteristics , and sexual behavior patterns. Although  testes and ovaries both synthesize androgens and estrogens, the testes predom¬  inantly secrete androgens, which are therefore known as male sex hormones,  whereas ovaries produce mostly estrogens, which are consequently termed  female sex hormones.   Androgens, of which testosterone (Fig. 9-11) is prototypic, lack the C 2  substituent at Cl7 that occurs in glucocorticoids and are therefore C 19 com¬  pounds. Estrogens, such as (5-estradiol (Fig. 9-11), resemble androgens but    are C 18 compounds. Interestingly, testosterone is an intermediate in estrogen  biosynthesis. Another class of ovarian steroids, C 2 i compounds called pro-  gestins, help mediate the menstrual cycle and pregnancy.   Androgens play a key role in sexual differentiation. If the gonads of an  embryonic male mammal are surgically removed, that individual will become  a phenotypic female. Evidently, mammals are programmed to develop as females  unless embryonically subjected to the influence of testicular hormones. Indeed, ge¬  netic males with absent or nonfunctional cytosolic androgen receptors are phe¬  notypic females, a condition named testicular feminization. Curiously,  estrogens appear to play no part in embryonic female sexual development,  although they are essential for female sexual maturation and function.   r Androgens that promote muscle growth are known as anabolic steroids.   Many individuals have taken anabolic steroids, both natural and synthetic,  in an effort to enhance their athletic performance or for cosmetic reasons.  However, because these substances and their metabolic products interact with  the various steroid receptors, their use may cause adverse side effects includ¬  ing cardiovascular disease, the development of breast tissue in males, mas-  culinization in females, temporary infertility in both sexes, and, in  adolescents, stunted growth due to accelerated bone maturation and preco¬  cious and/or exaggerated sexual development. Consequently, anabolic steroids  have been classified as controlled substances. Their use by competitive ath¬  letes has been banned to prevent an unfair advantage over athletes not tak¬  ing steroids.   Growth hormone (GH), a 19-residue polypeptide, is produced by the ante¬  rior lobe of the pituitary gland. Its binding to receptors directly stimulates  growth and metabolism in muscle, bone, and cartilage cells. GH also acts in¬  directly by stimulating the liver to produce additional growth factors.   Overproduction of GH, usually a consequence of a pituitary tumor, re¬  sults in excessive growth. If this condition commences while the skeleton is  still growing, that is, before its growth plates have ossified, then this excessive  growth is of normal proportions over the entire body, resulting in gigantism.  Moreover, since excessive GH inhibits the testosterone production necessary  for growth plate ossification, such “giants” continue growing throughout their  abnormally short lives. If, however, the skeleton has already matured, GH  stimulates only the growth of soft tissues, resulting in enlarged hands and feet  and thickened facial features, a condition named acromegaly. The opposite  problem, GH deficiency, which results in insufficient growth (dwarfism), can  be treated before skeletal maturity by regular injections of human GH (hGH;  animal GH is ineffective in humans).   Since hGH was, at first, available only from the pituitaries of cadavers, it  was in very short supply. Now, however, hGH can be synthesized in virtually  unlimited amounts via recombinant DNA techniques (Section 3-5D). Indeed,  hGH has been taken by individuals to increase their athletic prowess, although  there is no clear evidence that it does so. However, because of its adverse side  effects, which include high blood pressure, joint and muscle pain, and  acromegaly, as well as to eliminate any unfair competitive advantages in ath¬  letes, its nonmedical use is prohibited.   The GH Receptor Dimerizes on Hormone Binding. The 620-residue GH  receptor is a member of a large family of structurally related proteins. These  receptors consist of an N-terminal extracellular ligand-binding domain, a sin¬  gle transmembrane segment that is almost certainly helical, and a C-terminal  cytoplasmic domain that is not homologous within the superfamily but in  many cases functions as a tyrosine kinase (Section 13-2A).    FIG. 1 3-3 X-Ray structure of human  growth hormone (hGH) in complex with two  molecules of its receptor’s extracellular  domain (hGHbp). The proteins are shown in  ribbon form, with the hGH magenta and the two  hGHbp molecules, which together bind one  molecule of hGH, each colored in rainbow order  from its N-terminus (blue) to its C-terminus (red).  The dotted red lines indicate how the hGHbp  chains likely penetrate the membrane (light blue).  [Based on an X-ray structure by Abraham de  Vos and Anthony Kossiakoff, Genentech Inc.,  South San Francisco, California. PDBid  3HHR.] Q See Interactive Exercises 10.   • Explain why only certain cells respond to  hormones even though all cells in the  body are exposed to the hormone.   • List some hormones produced by the  pancreas, adrenal medulla, and adrenal  cortex. What types of molecules are these  hormones?   • Summarize the biological effects of insulin,  glucagon, norepinephrine, androgens,  estrogens, and growth hormone.   • What are some dangers of nonmedical  use of steroids and growth hormone?   • What is the significance of dimerization of  the GH receptor?   The X-ray structure of hGH in complex with the extracellular domain of  its binding protein (hGHbp) reveals that the complex consists of two mole¬  cules of hGHbp bound to a single hGH molecule (Fig. 13-3). hGH, like  many other protein growth factors, consists largely of a four-helix bundle. Each  hGHbp molecule consists of two structurally homologous domains, each of  which forms a topologically identical sandwich of a three- and a four-stranded  antiparallel (3 sheet that resembles the immunoglobulin fold (Section 7-3B).   The two hGHbp molecules bind to hGH with near twofold symmetry  about an axis that is roughly perpendicular to the helical axes of the hGH  four-helix bundle and, presumably, to the plane of the cell membrane to which  the intact hGH receptor is anchored (Fig. 13-3). The C-terminal domains of  the two hGHbp molecules are almost parallel and in contact with one an¬  other. Intriguingly, the two hGHbp molecules use essentially the same residues  to bind to sites that are on opposite sides of hGH s four-helix bundle and  which have no structural similarity.   The ligand-induced dimerization of hGHbp has important implications for  the mechanism of signal transduction. The dimerization, which does not oc¬  cur in the absence of hGH, apparently brings together the intact receptors in¬  tracellular domains in a way that activates intracellular effector proteins.  Indeed, hGH mutants that cannot induce receptor dimerization are biologi¬  cally inactive. Numerous other protein growth factors also induce the dimer¬  ization of their receptors.    • Dimerization and autophosphorylation allow receptor tyrosine kinases to become  active as protein tyrosine kinases.   • Adaptor proteins containing SH2 and SH3 domains can link a receptor tyrosine  kinase with G proteins and additional kinases that operate as a cascade.   • Protein phosphatases participate in signaling pathways by removing phosphoryl  groups from receptors and target proteins.    We have seen (Section 12-3B) that the activities of many enzymes are con¬  trolled by their covalent modification, mainly the phosphorylation of Ser and  Thr residues. A similar process forms the basis of one of the major intracel-    Phospho-Tyr residues mediate protein-protein interactions that are involved  in numerous cell functions. Consequently, the PTKs play a central role in sig¬  nal transduction, regulation of central metabolic pathways, cell cycle control,  and cell growth and differentiation. In this section, we discuss the proteins  that participate in this signaling system and how their activities are orches¬  trated to transmit signals within the cell.   The first step in all biochemical signaling pathways is the binding of a ligand to its  receptor protein. Box 13-2 discusses how receptor—ligand interactions are quan¬  titated. Insulin and many other polypeptide growth factors bind to receptors  whose C-terminal domains have tyrosine kinase activity. Such receptor tyrosine  kinases (RTKs) typically contain only a single transmembrane segment and are  monomers in the unliganded state. These structural features make it unlikely  that ligand binding to an extracellular domain manifests itself as a conforma¬  tional change in an intracellular domain (such a conformational shift seems more  likely to occur in receptors with multiple transmembrane segments). Indeed, the  most common mechanism for activating RTKs appears to be ligand-induced  dimerization of receptor proteins as is the case with the growth hormone recep¬  tor (Fig. 13-3; although it lacks tyrosine kinase activity). The insulin receptor is  unusual in that it is a dimer in the unliganded state. In this case, ligand bind¬  ing apparently induces a conformational change in the receptor.    Autophosphorylation Activates Receptor Tyrosine Kinases. When an RTK  dimerizes (or its conformation changes on ligand binding, in the case of the  insulin receptor), its cytoplasmic protein tyrosine kinase (PTK) domains are  brought close together so that they cross-phosphorylate each other on specific  Tyr residues. This autophosphorylation activates the PTK so that it can  phosphorylate other protein substrates.   How does autophosphorylation activate the PTK activity of the in¬  sulin receptor? The human protein is synthesized as a single 1382-  residue precursor peptide that is proteolytically processed to yield the  disulfide-linked a and (3 subunits of the mature receptor (Fig. 13-4).   Insulin binds to the receptors a subunits, which are entirely extracel¬  lular, whereas the (3 subunit contains a PTK domain on its intracellu¬  lar side.   subunits of the a 2 p 2 heterotetramer are linked by disulfide bonds  (short horizontal bars). The extracellular 731-residue a subunits form  the insulin-binding site. The cytoplasmic portions of the 620-residue p  subunits are tyrosine kinases that are activated by insulin binding.    Receptors, like other proteins, bind their corresponding ligands  according to the laws of mass action:    Here R and L represent receptor and ligand, and the reaction’s  dissociation constant is expressed:   where the total receptor concentration, [R] T , is equal to [R] + [R • L].  Equation 13-1 may be rearranged to a form analogous to the  Michaelis-Menten equation of enzyme kinetics (Section 12-1B):    where Y is the fractional occupation of the ligand-binding sites.  Equation 13-2 represents a hyperbolic curve (Fig. la) in which K L  may be operationally defined as the ligand concentration at which  the receptor is half-maximally occupied by ligand.   Although K L and [R] T may, in principle, be determined from an  analysis of a hyperbolic plot such as Fig. la, the analysis of a  linear form of the equation is a more common procedure.    Now, in keeping with customary receptor-binding nomenclature, let  us redefine [R • L] as B (for bound ligand), [L] as F (for free  ligand), and [R] T as B max . Then Eq. 13-3 becomes    A plot of B/F versus B, which is known as a Scatchard plot  (after George Scatchard, its originator), therefore yields a straight  line of slope ~\/K^ whose intercept on the B axis is B max    Figure 1 The binding of ligand to receptor, (a) A hyperbolic plot.  (b) A Scatchard plot. Here, B = [R • L], F = [L], and B max = [R] T .    (Fig. lb). Here, both B and F may be determined by filter-binding  assays as follows. Most receptors are insoluble membrane-bound  proteins and may therefore be separated from soluble free ligand  by filtration (receptors that have been solubilized may be sepa¬  rated from free ligand by filtration, for example, through nitrocellu¬  lose since proteins nonspecifically bind to nitrocellulose). Hence,  by using radioactively labeled ligand, the values of B and F ([R- L]  and [L]) may be determined, respectively, from the radioactivity on  the filter and that remaining in solution. The rate of R • L dissocia¬  tion is generally so slow (half-times of minutes to hours) as to  cause insignificant errors when the filter is washed to remove  residual free ligand.   Once the receptor-binding parameters for one ligand have  been determined, the dissociation constant of other ligands for the  same ligand-binding site may be determined through competitive  binding studies. The model describing this competitive binding is  analogous to the competitive inhibition of a Michaelis-Menten  enzyme (Section 12-2A):    where I is the competing ligand whose dissociation constant with  the receptor is expressed:    Thus, in direct analogy with the derivation of the equation describ¬  ing competitive inhibition:    The relative affinities of a ligand and an inhibitor may therefore be  determined by dividing Eq. 13-6 in the presence of inhibitor with  that in the absence of inhibitor:    When this ratio is 0.5 (50% inhibition), the competitor concentra¬  tion is referred to as [l 50 ]. Thus, solving Eq. 13-7 for K\ at 50%  inhibition:    The X-ray structure of the (3 subunit’s 306-residue PTK domain  (Fig. 13-5^) reveals a deeply clefted bilobal protein whose N-terminal do¬  main consists of a five-stranded (3 sheet and an a helix and whose larger  C-terminal domain is mainly a-helical. This structure, as we will repeatedly  see, is typical of the large family of protein kinases, enzymes that phosphory-   FIG. 1 3-5 X-Ray structure of the tyrosine kinase domain of  the insulin receptor, (a) The tyrosine kinase domain is shown in  the “standard” protein kinase orientation with its N-terminal domain  lavender, its C-terminal domain cyan, and its activation loop light blue.  Three phosphorylated Tyr side chains are shown in ball-and-stick form  with C green, N blue, 0 red, and P yellow. The ATP analog AMPPNP  is shown in space-filling form. Six residues of the substrate  polypeptide are shown in orange, and its phosphorylatable Tyr residue  is shown in magenta, (b) The polypeptide backbones of the   phosphorylated and unphosphorylated forms of the insulin receptor  tyrosine kinase domain are shown superimposed on their C-terminal  lobes. The phosphorylated protein is green with its activation loop  blue, and the unphosphorylated protein is yellow with its activation  loop red. The cyan arrow and black axis indicate the rotation required  to align the two N-terminal lobes. [Part a based on an X-ray structure  by and Part b courtesy of Stevan Hubbard, New York University  Medical School. PDBids 1IR3 and 1IRKJ ©See Interactive  Exercise 11.    late the OH groups of Tyr residues and/or Ser and Thr residues. Indeed, the  human genome encodes 90 PTKs and 388 protein Ser/Thr kinases (representing  >2% of human genes) that collectively phosphorylate an estimated one-third  of the proteins in human cells. In doing so, they play key roles in the signal¬  ing pathways by which many hormones, growth factors, neurotransmitters,  and toxins affect the functions of their target cells.   In the X-ray structure of the insulin receptor PTK domain (Fig. 13-5^),  the nonhydrolyzable ATP analog adenosine-5'-(P,7-imido)triphosphate  (AMPPNP; at right; alternatively ADPNP), is bound in the cleft between the  protein domains. There, its y-phosphate group is in close juxtaposition to the OH  group of the target Tyr residue in a substrate peptide that is also bound to the  protein. Three of the PTKs Tyr residues, which are located on the C-terminal do¬  main in its so-called activation loop, are phosphorylated. In its unphosphorylated  state, the 18-residue loop threads through the PTK active site so as to prevent the  binding of both ATP and protein substrates. When the three Tyr residues are phos¬  phorylated, the activation loop changes its conformation such that it does not oc¬  clude the active site (Fig. 13-5#) but instead forms part of the substrate recognition  site. In fact, the PTK activity of the insulin receptor increases with the degree of  phosphorylation of its three autophosphorylatable Tyr residues.   The conformational changes in PTK induced by phosphorylation and lig¬  and binding are depicted in Fig. 13-5 h. The PTKs N-terminal lobe under¬  goes a nearly rigid 21° rotation relative to the C-terminal lobe, a dramatic   conformational change that presumably positions critical residues for substrate  binding and catalysis.   Nearly all known PTKs have between one and three autophosphorylatable  Tyr residues in their activation loops, which assume similar conformations in  all phosphorylated PTKs of known structure. Moreover, many activated PTKs  also phosphorylate the opposing RTK at cytoplasmic Tyr residues outside of  the PTK domain. The specificity of PTKs for phosphorylating Tyr rather than  Ser or Thr is explained by the observation that the side chain of Tyr, but not  those of Ser or Thr, is long enough to reach the active site.   Although certain autophosphorylated RTKs directly phosphorylate their ultimate  target proteins, many do not do so. How, then, are these target proteins activated?  The answer, as we will see, is through a highly diverse and complicated set of in¬  terconnected signaling pathways involving cascades of associating proteins.   SH2 Domains Bind Phospho-Tyr Residues. The main substrates of the in¬  sulin receptor tyrosine kinase are known as insulin receptor substrates 1 and  2 (IRS-1 and IRS-2). When phosphorylated, these proteins can interact with  yet another set of proteins that contain one or two conserved ~ 100-residue  modules known as Src homology 2 (SH2) domains [because they are simi¬  lar to the sequence of a domain in the protein named Src (pronounced  “sarc”)]. SH2 domains bind phospho-Tyr residues with high affinity but do  not bind the far more abundant phospho-Ser and phospho-Thr residues. This  specificity has a simple explanation. X-Ray structural studies reveal that  phospho-Tyr interacts with an Arg at the bottom of a deep pocket (Fig. 13-6).  The side chains of Ser and Thr are too short to interact with this residue.   The SH2-containing proteins that interact with the IRSs and other phos¬  phorylated proteins have varied functions: Some are kinases, some are phos¬  phatases, and some are GTP-binding proteins known as G proteins (G  proteins also play a key role in non-RTK signaling pathways, as described in  Section 13-3). Consequently, hormone binding to its RTK can elicit a variety  of intracellular responses.   RTKs Indirectly Activate the G Protein Ras. Molecular genetic analysis of sig¬  naling in a variety of distantly related organisms revealed a remarkably conserved  pathway that regulates such essential functions as cell growth and  differentiation. Briefly, growth factor binding to its cognate RTK  activates a monomeric G protein named Ras that is anchored to  the inner surface of the plasma membrane by prenylation (Section  9-3B). Activated Ras then activates a kinase cascade that relays  the signal to the transcriptional apparatus in the nucleus.   The binding of a growth factor to its RTK leads to autophos¬  phorylation of the RTK, which then interacts with an SH2-con-  taining protein (Fig. 13-7, top left). Many proteins that contain  SH2 domains also have one or more unrelated 50- to 75-residue    FIG. 1 3-6 X-Ray structure of the Src SH2 domain in complex  with a target peptide. The protein is represented by its ribbon diagram  colored in rainbow order from its N-terminus (blue) to its C-terminus (red)  and embedded in its semitransparent surface diagram. An 11-residue  polypeptide containing the protein’s phospho-Tyr-Glu-Glu-lle (pYEEl) target  tetrapeptide is drawn in space-filling form with backbone C cyan, side  chain C green, N blue, 0 red, and P orange. The side chain of SH2 Arg  32, which interacts with the phosphoryl group of the peptide phospho-Tyr  (pTyr) residue, is drawn in stick form. [Based on an X-ray structure by John  Kuriyan, The Rockefeller University. PDBid ISPS.]   FIG. 1 3-7 The Ras signaling cascade. RTK binding to its  cognate growth factor (1) induces the autophosphorylation of the  RTK’s cytosolic domain (2). Grb2/Sem-5 binds to the resulting  phospho-Tyr-containing peptide segment via its SH2 domain (3) and  simultaneously binds to Pro-rich segments on Sos via its two SH3  domains (4). This activates Sos to exchange Ras’s bound GDP for  GTP (5), which activates Ras to bind to Raf (6). Then, in a so-called  kinase cascade, Raf, a Ser/Thr kinase, phosphorylates MEK (7), which  in turn phosphorylates MAPK (8), which then migrates to the nucleus    (9), where it phosphorylates transcription factors such as Fos, Jun,  and Myc (10), thereby modulating gene expression. Ras is eventually  inactivated by GTP hydrolysis (11), a process that is accelerated by  GTPase-activating proteins (GAPs). The kinase cascade eventually  returns to its resting state through the action of protein phosphatases  (Section 13-2D). [After Egan, S.E. and Weinberg, R.A., Nature 365,  782 (1993).] O See the Animated Figures.   FIG. 1 3-8 X-Ray structure of the SH3 domain from Abl protein in complex with  its 10-residue target Pro-rich polypeptide (APTMPPPLPP). The protein is represented  by its surface diagram (cyan) and the peptide is drawn in stick form with Pro C magenta,  other C green, N blue, 0 red, and S yellow. [Based on an X-ray structure by Andrea  Musacchio, European Molecular Biology Laboratory, Heidelberg, Germany. PDBid 1ABOJ    SH3 domains. SH3 domains, which bind Pro-rich sequences of 9 or 10  residues (Fig. 13-8), are also present in some proteins that lack SH2 domains.  In the signaling cascade shown in Fig. 13-7, the mammalian protein known  as Grb2 (Sem-5 in the nematode worm Caenorhabditis elegans) consists almost  entirely of an SH 2 domain flanked by two SH3 domains (Fig. 13-9).  Grb2/Sem-5 links the autophosphorylated RTK (via its SH 2 domain) to the  Pro-rich Sos protein (via its SH3 domains).   Inactive Ras has GDP bound in its nucleotide-binding site. The Grb 2 -Sos  complex activates Ras by inducing it to release its bound GDP and replace it  with GTP. Only the Ras • GTP complex is capable of further relaying the growth -  promoting signal from the RTK. Ras tightly binds both GTP and GDP and  hence must interact with Sos to exchange these nucleotides. Sos is therefore  known as a guanine nucleotide exchange factor (GEF). Most G proteins, as  we will see, have a corresponding GEE   The X-ray structure of Grb 2 (Fig. 13-9) suggests that its SH 2 domain is  flexibly linked to its two SH3 domains. How does the binding of such a pli¬  able adaptor (a linker that lacks enzymatic activity) to a phosphorylated RTK  cause Sos to activate Ras? Grb 2 and Sos bind one another so tightly that they  are essentially permanently associated in the cell. Hence, when Grb 2 binds to  a phosphorylated RTK, it recruits Sos to the inner surface of the plasma mem¬  brane, where the increased local concentration of Sos causes it to more read¬  ily bind to the membrane-anchored Ras and thus act as a GEF.    GAPs Accelerate the GTPase Activities of G Proteins. Ras is an enzyme (a  GTPase) that catalyzes the hydrolysis of its bound GTP to GDP + P z , thereby  limiting the magnitude of the cell’s response to the growth factor. Yet, Ras by  itself hydrolyzes only two to three GTPs per minute, too slowly for effective  signal transduction. Furthermore, for a signal to be more than a one-time  switch, there must be a mechanism for turning it off as well as on. These con¬  siderations led to the discovery of a 120-kD GTPase-activating protein  (GAP) named RasGAP that, on binding Ras • GTP, accelerates the rate of  GTP hydrolysis by a factor of 10 5 . Most G proteins, like Ras, also have a cor¬  responding GAP.   The mechanism whereby RasGAP activates the GTPase activity of Ras  was revealed by the X-ray structure of the 334-residue GTPase-activating do¬  main of RasGAP (GAP334) bound to Ras in its complex with GDP and A1F 3  (Fig. 13-10). GAP334 interacts with Ras over an extensive surface. The A1F 3 ,  which has trigonal planar geometry, binds to Ras at the expected position of  GTPs 7 phosphate group, with the A1 atom opposite a bound water mole¬  cule that presumably would be the attacking nucleophile in the GTPase reac¬  tion. Since A1—F and P—O bonds have similar lengths, the GDP-A1F 3 -H 2 0  assembly resembles the GTPase reactions expected transition state with the  A1F 3 mimicking the planar P0 3 group.   GAP334 binds to Ras with GAP334 s so-called finger loop inserted into  the Ras active site such that an Arg residue in the loop interacts with both    FIG. 1 3-9 X-Ray structure of Grb2. Its SH2 domain (green) is linked to its flanking  SH3 domains (cyan and orange) via apparently unstructured and hence flexible four-residue  linkers. [Based on an X-ray structure by Arnaud Ducruix, Universite de Paris-Sud, Gif sur  Yvette Cedex, France. PDBid 1GRI.]    FIG. 13-10 X-Ray structure of the  GAP334 • Ras • GDP • AIF 3 complex. The   active site regions of the proteins are shown as  ribbons with Ras cyan, its Gly 12 magenta,  GAP334 yellow, and its finger loop red. The  GDP, AIF 3 , and the side chains of Ras Gin 61  and GAP334 Arg 789 are drawn in stick form  with C green, N blue, 0 red, F yellow-green, P  orange, and Al pink; the water molecule is  represented by a red sphere; and hydrogen  bonds are shown as dashed lines. [Based on an  X-ray structure by Alfred Wittinghofer,  Max-Planck-lnstitut fur Molekulare Physiologie,  Dortmund, Germany. PDBid 1WQ1.]    the GDP’s (3 phosphate and the A1F 3 . In the Ras • GTP complex, this Arg  side chain would be in an excellent position to stabilize the developing nega¬  tive charge in the GTPase reactions transition state. Indeed, catalytically more  efficient G proteins contain an Arg residue that occupies a nearly identical  position.   A Kinase Cascade Completes the Signaling Pathway. The signaling path¬  way downstream of Ras consists of a linear cascade of protein kinases  (Fig. 13-7, righi). The Ser/Thr kinase Raf, which is activated by direct inter¬  action with Ras • GTP, phosphorylates a protein alternatively known as MEK  or MAP kinase kinase, thereby activating it as a kinase. Activated MEK phos¬  phorylates a family of proteins variously termed mitogen-activated protein  kinases (MAPKs) or extracellular-signal-regulated kinases (ERKs). A  MAPK must be phosphorylated at both its Thr and Tyr residues in the se¬  quence Thr-Glu-Tyr for full activity. MEK (which stands for MAP kinase/ARK  kinase-activating kinase) catalyzes both phosphorylations; it is therefore a pro¬  tein Ser/Thr kinase as well as a protein Tyr kinase.   The activated MAP kinases migrate from the cytosol to the nucleus, where  they phosphorylate a variety of proteins, including Fos, Jun, and Myc. These  proteins are transcription factors (proteins that induce the transcription of  their target genes; Section 28-3B): In their activated forms they stimulate var¬  ious genes to produce the effects commissioned by the extracellular presence  of the growth factor that initiated the signaling cascade. When insulin acti¬  vates the Ras signaling pathway, the result is an increase in protein synthesis  that supports cell growth and differentiation, a response consistent with in¬  sulins function as a signal of fuel abundance. Variant proteins encoded by  oncogenes subvert such signaling pathways so as to induce uncontrolled cell  growth (Box 13-3).   The advantage of a kinase cascade is that a small signal can be amplified  manyfold inside the cell. In addition, phosphorylation of more than one target  protein can lead to the simultaneous activation of several intracellular  processes. Thus as we will see (Section 22-2), insulin signaling mediates  changes in vesicle trafficking, enzyme activation, and gene expression.   Scaffold Proteins Organize and Position Protein Kinases. Eukaryotic cells  contain numerous different MAPK signaling cascades, each with a character-    The growth and differentiation of cells in the body are normally  strictly controlled. Thus, with few exceptions (e.g., blood-forming  cells and hair follicles), cells in the adult body are largely quies¬  cent. However, for a variety of reasons, a cell may be made to  proliferate uncontrollably to form a tumor.   Malignant tumors (cancers) grow in an invasive manner and  are almost invariably life threatening. They are responsible for  20% of the mortalities in the United States.   Among the many causes of cancer are viruses that carry  oncogenes (Greek: onkos, mass or tumor). For example, the  Rous sarcoma virus (RSV), which induces the formation of  sarcomas (cancers arising from connective tissues) in chickens,  contains four genes. Three of the genes are essential for viral  replication, whereas the fourth, v-src (v for viral, src for sarcoma),  an oncogene, induces tumor formation. What is the origin of v-src,  and how does it function? Hybridization studies by Michael Bishop  and Harold Varmus in 1976 led to the remarkable discovery that  uninfected chicken cells contain a gene, c-src (c for cellular), that  is homologous to v-src and that is highly conserved in a wide  variety of eukaryotes, suggesting that it is an essential cellular  gene. Apparently, v-src was originally acquired from a cellular  source by a non-tumor-forming ancestor of RSV. Both v-src and   An X-ray-based false-color image showing an axial section through a  human abdomen that has cancer of the liver. The liver is the large red  mass occupying much of the abdomen; the light patches on the liver  are cancerous tumors. A vertebra (dark green ) can be seen at the  lower center of the image. [Salisbury/Photo Researchers.]    c-src encode a 60-kD tyrosine kinase. However, whereas the  activity of c-src is strictly regulated, that of v-src is under no such  control and hence its presence maintains the host cell in a prolif¬  erative state. Since cells are not killed by an RSV infection, this  presumably enhances the viral replication rate.   Other oncogenes have been similarly linked to processes that  regulate cell growth. For example, the v-erbB oncogene specifies  a truncated version of the epidermal growth factor (EGF)  receptor, which lacks the EGF-binding domain but retains its  transmembrane segment and its tyrosine kinase domain. This  kinase phosphorylates its target proteins in the absence of an  extracellular signal, thereby driving uncontrolled cell proliferation.   The v-ras oncogene encodes a 21-kD protein, v-Ras, that  resembles cellular Ras but hydrolyzes GTP much more slowly. The  reduced braking effect of GTP hydrolysis on the rate of protein  phosphorylation leads to increased activation of the kinases down¬  stream of Ras (Fig. 13-7).   The transcription factors that respond to Ras-mediated signal¬  ing (e.g., Fos and Jun) are also encoded by proto-oncogenes,  the normal cellular analogs of oncogenes. The viral genes v-fos  and M-jun encode proteins that are nearly identical to their cellular  counterparts and mimic their effects on host cells but in an  uncontrolled fashion.   Oncogenes are not necessarily of viral origin. Indeed, few hu¬  man cancers are caused by viruses. Rather, they are caused by  proto-oncogenes that have mutated to form oncogenes. For exam¬  ple, a mutation in the c-ras gene, which converts Gly 12 of Ras  to Val, reduces Ras’s GTPase activity without affecting its ability to  stimulate protein phosphorylation. This prolongs the time that Ras  is in the “on” state, thereby inducing uncontrolled cell proliferation.  In fact, oncogenic versions of c-ras are among the most com¬  monly implicated oncogenes in human cancers.   To date, over 350 viral and cellular oncogenes have been  identified. The subversive effects of oncogene products arise  through their differences from the corresponding normal cellular  proteins: They may have different rates of synthesis and/or degra¬  dation; they may have altered cellular functions; or they may resist  control by cellular regulatory mechanisms. However, in order for a  normal cell to undergo a malignant transformation (become a  cancer cell), it must undergo several (an average of five) inde¬  pendent oncogenic events. This is a reflection of the complexity of  cellular signaling networks (cells respond to a variety of hormones,  growth factors, and transcription factors in partially overlapping  ways) and explains why the incidence of cancer increases with  age. Nevertheless, at the cellular level, a malignant transformation  is an extremely rare event because oncogenic mutations are infre¬  quent and because cells have highly effective defense mecha¬  nisms that guard against cancer.    istic set of component kinases, which in mammals comprise at least 12 MAP  kinases, 7 MAP kinase kinases (MKKs; e.g., MEK), and 14 MAP kinase  kinase kinases (MKKKs; e.g., Raf; Fig. 13-11). Although each MAPK is ac¬  tivated by a specific MKK, a given MKK can be activated by more than one  MKKK. Moreover, several pathways may be activated by a single type of re¬  ceptor. How then does a cell prevent inappropriate cross talk between closely  related signaling pathways? One way that this occurs is through the use of    p90 rsk , S6 kinase, Sos,  phospholipase A 2 ,  EGF receptor, Elk-1, Etsl,  Sapla, c-Myc, Tal, STATS    FIG. 1 3-1 1 MAP kinase cascades in mammalian cells. Each MAP kinase cascade  consists of an MKKK, an MKK, and a MAPK. Various external stimuli may each activate  one or more MKKKs, which in turn may activate one or more MKKs. However, the MKKs  are relatively specific for their target MAPKs. The activated MAPKs phosphorylate specific  transcription factors, which are then translocated to the nucleus, as well as specific kinases.  The resulting activated transcription factors and kinases then induce cellular responses  such as growth, differentiation, and apoptosis (programmed cell death; Section 28-4C).  [After Garrington, T.P. and Johnson, G.L, Curr. Opin. Cell Biol. 11, 212 (1999).]    scaffold proteins that bind some or all of the component protein kinases of  a particular signaling cascade so as to ensure that the protein kinases of a given  pathway interact only with one another. In addition, a scaffold protein can  control the subcellular location of its associated kinases.   The first known scaffold protein was discovered through the genetic analy¬  sis of a MAP kinase cascade in yeast, which demonstrated that this protein,  Ste5p, binds the MKKK, MKK, and MAPK components of the pathway and  that, in vivo, the scaffolds absence inactivates the pathway. Evidently, the in¬  teractions between successive kinase components of this MAP kinase cascade  are, by themselves, insufficient for signal transmission.   Many cell-surface receptors for growth factors and related molecules do not re¬  spond to ligand binding by autophosphorylation. These include the receptors  for growth hormone (Fig. 13-3), the cytokines (protein growth factors that reg¬  ulate the differentiation, proliferation, and activities of numerous types of cells,  most conspicuously blood cells), the interferons (protein growth factors that  stimulate antiviral defenses), and T cell receptors [which control the prolifer¬  ation of immune system cells known as T lymphocytes (T cells); Section 7-3].  Ligand binding induces these tyrosine kinase—associated receptors to dimerize  (and, in some cases, to trimerize), often with different types of subunits, in a way  that activates associated nonreceptor tyrosine kinases (NRTKs).   Chapter 13 Biochemical Signaling The Structure of Src Reveals Its Autoinhibitory Mechanism. Many of the   NRTKs belong to the Src family, which contains at least nine members, in¬  cluding Src, Fyn, and Lck. Most of these ~530-residue membrane-anchored  (by myristoylation) proteins have both an SH2 and an SH3 domain and all  have a PTK domain. Hence, a Src-related kinase may also be activated by an  autophosphorylated RTK. Although Src-related kinases are each associated  with different receptors, they phosphorylate overlapping sets of target proteins.  This complex web of interactions explains why different ligands often activate  some of the same signaling pathways.   Src consists of, from N- to C-terminus, a myristoylated N-terminal  “unique” domain that differs among Src family members, an SH3 domain, an  SH2 domain, a PTK domain, and a short C-terminal tail. Phosphorylation of  Tyr 416 in the PTK’s activation loop activates Src, whereas phosphorylation  of Tyr 527 in its C-terminal tail deactivates it. In vivo , Src is phosphorylated  at either Tyr 416 or Tyr 527, but not at both. The dephosphorylation of Tyr  527 or the binding of external ligands to the SH2 or the SH3 domain acti¬  vates Src, a state that is then maintained by the autophosphorylation of Tyr  416. When Tyr 527 is phosphorylated and no activating phosphopeptides are  available, Src s SH2 and SH3 domains function to deactivate its PTK domain;  that is, Src is then autoinhibited.   The X-ray structure of Src*AMPPNP lacking its N-terminal domain and  with Tyr 527 phosphorylated reveals the structural basis of Src autoinhibition  (Fig. 13-12). As biochemical studies had previously shown, the SH2 domain  binds phospho-Tyr 527, which occurs in the sequence pYNPG rather than the  pYEEI sequence characteristic of high-affinity Src SH2 target peptides.  Although the pYNP segment binds to SH2 as does the pYEE segment in  Fig. 13-6, the succeeding residues are poorly ordered in the X-ray structure and,  moreover, the SH2 pocket in which the lie side chain of pYEEI binds is un¬  occupied. Apparently, the phospho-Tyr 527-containing peptide segment binds  to the Src SH2 domain with reduced affinity relative to its target peptides.   The Src SH3 domain binds to the linker connecting the SH2 domain to  the N-terminal lobe of the PTK domain. Residues 249 to 253 of the linker  bind to the SH3 domain in much the same way as do SH3 s Pro-rich target  peptides (Fig. 13-8). However, the only Pro in this segment is residue 250.  The polar side chain of Gin 253, which occupies the position of the second  Pro in SH3 s normal Pro-X-X-Pro target sequence, does not enter the hy¬  drophobic binding pocket that this second Pro would occupy (Fig. 13-8) and  hence the path of the peptide deviates from that of Pro-rich target peptides at  this point. Apparently, this interaction is also weaker than those with Src s SH3  target peptides.   Since Src’s SH2 and SH3 domains bind the PTK domain on  the side opposite its active site, how is the PTK activity inhibited?  The two lobes of Src’s PTK domain are, for the most part, closely  superimposable on their counterparts in the PTK domains of phos¬  phorylated and hence activated protein kinases (e.g., Fig. 13-5^).   FIG. 1 3-1 2 X-Ray structure of Src • AMPPNP with Tyr 527  phosphorylated. The protein, which lacks its membrane-anchored N-terminal  domain, is oriented such that its PTK domain is seen in “standard” view  (compare it with Fig. 13-5a). The SH3 domain is orange, the SH2 domain is  magenta, the linker joining the SH2 domain to the PTK domain is green with  residues 249 to 253, which interact with the SH3 domain, yellow, the  N-terminal lobe of the PTK domain is lavender, its C-terminal lobe is cyan with  its activation loop light blue, and its C-terminal tail is red. The AMPPNP is  shown in space-filling form and Y416 (unphosphorylated) and pY527  (phosphorylated) are shown in ball-and-stick form, all with C green, N blue,   0 red, and P yellow. [Based on an X-ray structure by Stephen Harrison and  Michael Eck, Harvard Medical School. PDBid 2SRC.]    However, Src helix C (the only helix in the PTK’s N-terminal lobe) is  displaced from the interface between the N- and C-terminal lobes, a po¬  sition it occupies in other activated protein kinases (e.g., Fig. 13-5 a). Helix  C contains the conserved residue Glu 310 (using Src numbering), which in  other activated protein kinases projects into the catalytic cleft where it forms  a salt bridge with Lys 295, an important ligand of the substrate ATP s a and  (3 phosphates. In inactive Src, Glu 310 forms an alternative salt bridge with  Arg 409, and Lys 295 instead interacts with Asp 404. In activated Lck, Arg  409 forms a salt bridge with phospho-Tyr 416.   The foregoing structural observations suggest the following scenario for  Src activation (Fig 13-13):   1 . The dephosphorylation of Tyr 527 and/or the binding of the SH 2  and/or SH3 domains to their target peptides (for which SH 2 and SH3  have greater affinity than their internal Src-binding sites) releases these  domains from their PTK-bound positions shown in Fig. 13-12, thus  relaxing conformational constraints on the PTK domain. This allows  the PTK’s active site cleft to open, thereby disrupting the structure of  its partially helical activation loop (which occupies a blocking position  in the active site cleft; Fig. 13-12) so as to expose Tyr 416 to autophos¬  phorylation.   2 . The resulting phospho-Tyr 416 forms a salt bridge with Arg 409, which  sterically requires the structural reorganization of the activation loop to  its active, nonblocking conformation. The consequent rupture of the  Glu 310—Arg 409 salt bridge frees helix C to assume its active orien¬  tation which, in turn, allows Glu 310 to form its catalytically impor¬  tant salt bridge to Lys 295, thereby activating the Src PTK activity.   autoinhibited form (left), the SH2 domain ( magenta) binds to  phospho-Tyr 527, and the SH3 domain ( orange) binds to an internal  Pro-containing segment (yellow). Glu 310 forms a salt bridge to Arg  409, the partially helical activation loop (blue) blocks the active site,  and Tyr 416 is buried. In the active form (right), the SH2 and SH3  domains bind to a Src activator, Tyr 527 is dephosphorylated, the   activation loop has undergone a conformational change to expose Tyr  416 to phosphorylation, Glu 310 forms a salt bridge with Lys 295,  and phospho-Tyr 416 forms a salt bridge with Arg 409. The coloring  scheme and viewpoint largely match those in Fig. 13-12. [After Young,  M.A., Gonfloni, F., Superti-Furga, G., Roux, B., and Kuriyan, J., Cell 105,  115 (2001).]   FIG. 1 3-1 4 X-Ray structure of the Abl  PTK domain in complex with a truncated  derivative of Gleevec. The protein is viewed  from the right of the “standard” view of protein  kinases (e.g., Figs. 13-5a and 13-12), with its  N-terminal lobe lavender, its C-terminal lobe  cyan, and its activation loop light blue. The  truncated Gleevec, which occupies the PTK’s  ATP-binding site, is shown in space-filling form  with C green, N blue, and 0 red. [Based on an  X-ray structure by John Kuriyan, The Rockefeller  University. PDBid 1 FPU.]    T PTKs Are Targets of Anticancer Drugs. The hallmark of chronic mye¬  logenous leukemia (CML) is a specific chromosomal translocation form¬  ing the so-called Philadelphia chromosome in which the Abl gene (which  encodes the NRTK Abl) is fused with the Bcr gene (which encodes the pro¬  tein Ser/Thr kinase Bcr). The Abl portion of the resulting Bcr—Abl fusion pro¬  tein is constitutively activated (that is, continuously, without regulation),  probably because its Bcr portion oligomerizes. Hematopoietic stem cells (from  which all blood cells are descended) bearing the Philadelphia chromosome are  therefore primed to develop CML (malignancy requires several independent  genetic alterations; Box 13-3). Without a bone marrow transplant (a high-risk  procedure that is unavailable to most individuals due to the lack of a suitable  donor), CML is invariably fatal with an average survival time of ~6 years.   An inhibitor of Abl would be expected to prevent the proliferation of, and  even kill, CML cells. However, to be an effective anti-CML agent, such a sub¬  stance must not inhibit other protein kinases because this would almost cer¬  tainly cause serious side effects. Derivatives of 2-phenylaminopyrimidine bind  to Abl with exceptionally high affinity and specificity. One such derivative,  imatinib (trade name Gleevec),   which was developed by Brian Druker and Nicholas Lydon, has caused the re¬  mission of symptoms in >90% of CML patients with almost no serious side  effects. This unprecedented performance occurs, in part, because Gleevec does  not bind to other protein kinases.   Abl resembles Src but lacks Srcs C-terminal regulatory phosphorylation  site (Tyr 527; Figs. 13-12 and 13-13). The X-ray structure of Abls PTK do¬  main in complex with a truncated form of Gleevec, determined by John  Kuriyan (Fig. 13-14), reveals, as expected, that the drug binds in Abls  ATP-binding site. Abl thereby adopts an inactive conformation in which its  activation loop, which is not phosphorylated, assumes an autoinhibitory  conformation.   Gleevec was the first of several 2-phenylaminopyrimidine derivatives,  which inhibit specific protein kinases, to be approved by the FDA for clinical  use against certain cancers. In addition, several monoclonal antibodies (Box 7-5)  that bind to specific PTKs or their ligands are in clinical use as anticancer  agents [e.g., trastuzumab (trade name Herceptin), which is effective against  breast cancers that overexpress the RTK named HER2] . Such receptor-targeted  therapies hold enormous promise for controlling, if not curing, cancers by  specifically targeting the aberrant proteins that cause the cancers. In contrast,  most chemotherapeutic agents that are presently in use indiscriminately kill  fast-growing cells and hence tend to have debilitating side effects.   Intracellular signals must be “turned off” after the system has delivered its mes¬  sage so that the system can transmit future messages. In the case of protein  kinases, their activities are balanced by the activities of protein phosphatases  that hydrolyze the phosphoryl groups attached to Ser, Thr, or Tyr side chains  and thereby limit the effects of the signal that activated the kinase. Although   protein kinases have traditionally garnered more attention, mammalian cells Section 2 Receptor Tyrosine Kinases   express ~500 protein phosphatases (about the same number as protein ki¬  nases) with substrate specificities comparable to those of kinases.   Protein Tyrosine Phosphatases Are Multidomain Proteins. The enzymes  that dephosphorylate Tyr residues, the protein tyrosine phosphatases (PTPs),  are not just simple housekeeping enzymes but are important signal transduc¬  ers. These enzymes, 107 of which are encoded by the human genome, are  members of four families. Each tyrosine phosphatase contains at least one con¬  served ~240-residue phosphatase domain that has the 11-residue signature se¬  quence [(I/V)HCXAGXGR(S/T)G], the so-called CX 5 R motif, which  contains the enzymes catalytically essential Cys and Arg residues. During the  hydrolysis reaction, the phosphoryl group is transferred from the tyrosyl  residue of the substrate protein to the essential Cys on the enzyme, forming  a covalent Cys-phosphate intermediate that is subsequently hydrolyzed.   Some tyrosine phosphatases are constructed much like the receptor ty¬  rosine kinases; that is, they have an extracellular domain, a single transmem¬  brane helix, and a cytoplasmic domain consisting of a catalytically active  PTP domain that, in most cases, is followed by a second PTP domain with  little or no catalytic activity. These inactive PTP domains are, nevertheless,  highly conserved, which suggests that they have an important although as  yet unknown function. Biochemical and structural analyses indicate that  ligand-induced dimerization of a receptor-like PTP reduces its catalytic  activity, probably by blocking its active sites.   A second group of PTPs, intracellular PTPs, contain only one tyrosine  phosphatase domain, which is flanked by regions containing motifs, such as  SH2 domains, that participate in protein-protein interactions. The PTP  known as SHP-2, which is expressed in all mammalian cells, binds to a vari¬  ety of phosphorylated (that is, ligand-activated) RTKs. The X-ray structure of  SHP-2 lacking its C-terminal tail reveals two SH2 domains, followed by a  tyrosine phosphatase domain (Fig. 13-15). The N-terminal SH2 domain  (N-SH2) functions as an autoinhibitor by inserting a protein loop (labeled  D'E in Fig. 13-15) into the PTPs 9-A-deep catalytic cleft. When N-SH2 rec¬  ognizes and binds a phospho-Tyr group on a substrate protein, its conforma¬  tion changes, unmasking the PTP catalytic site so that the phosphatase can  hydrolyze another phospho-Tyr group on the target protein (activated RTKs  typically bear multiple phosphorylated Tyr residues).   The active site cleft of intracellular tyrosine phosphatases such as SHP-2 is  too deep to cleave phospho-Ser/Thr side chains. However, the active site pockets  of a third group of PTPs, the so-called dual-specificity tyrosine phosphatases,  are sufficiently shallow to bind both phospho-Tyr and phospho-Ser/Thr residues.   FIG. 1 3-1 5 X-Ray structure of the protein  tyrosine phosphatase SHP-2. Its N-SH2  domain is gold with its D'E loop red, its C-SH2  domain is green, and its PTP domain is cyan  with its 11-residue CX 5 R motif blue. The side  chain of the catalytically essential Cys residue is  shown in ball-and-stick form with C green and  S yellow. [Based on an X-ray structure by  Michael Eck and Steven Shoelson, Harvard  Medical School. PDBid 2SHP.]    T Bubonic Plague Virulence Requires a PTP. Bacteria lack PTKs and hence  do not synthesize phospho-Tyr residues. Nevertheless, PTPs are expressed  by bacteria of the genus Yersinia, most notably Yersinia pestis, the pathogen  that causes bubonic plague (the flea-transmitted “Black Death,” which, since  the sixth century, has been responsible for an estimated 200 million human  deaths including about one-third of the European population in the years  1347-1350). The Y pestis PTP, YopH, which is required for bacterial viru¬  lence, is far more catalytically active than other known PTPs. Hence, when  Yersinia injects YopH into a cell, the cells phospho-Tyr-containing proteins  are catastrophically dephosphorylated. Although YopH and mammalian PTPs  are only ~15% identical in sequence, they share a set of invariant residues  and have similar X-ray structures. This suggests that an ancestral Yersinia ac¬  quired a PTP gene from a eukaryote.    Protein Ser/Thr Phosphatases Participate in Numerous Regulatory  Processes. The protein Ser/Thr phosphatases in mammalian cells belong  to two protein families: the PPP family and the PPM family. The PPP and  PPM families are unrelated to each other or to the PTPs. X-Ray structures  have shown that PPP catalytic centers each contain an Fe 2+ (or possibly an  Fe 3+ ) ion and a Zn 2+ (or possibly an Mn 2+ ) ion, whereas PPM catalytic  centers each contain two Mn 2+ ions. These binuclear metal ion centers  nucleophilically activate water molecules to dephosphorylate substrates in a  single reaction step.   The PPP family member named phosphoprotein phosphatase-1  (PP1), as we will see, plays an important role in regulating glycogen   (a) The structure of an isolated scaffold (A) subunit HEAT repeats,  which are drawn here in different colors, each consist of two  antiparallel helices joined by a short linker. These stack on one  another with their corresponding helices nearly parallel to form an  ~100-A-long right-handed superhelix (helix of helices) with a hooklike  shape. [Courtesy of Bostjan Kobe, St. Vincent’s Institute of Medical  Research, Fitzroy, Victoria, Australia. X-Ray structure by David Barford,  University of Oxford, U.K. PDBid 1 B3U.] (b) The structure of a PP2A    heterotrimer viewed with the scaffold subunit oriented approximately  as in Part a. Here the scaffold (A; 589 residues) and regulatory  (B; 449 residues) subunits are drawn in worm form, each colored in  rainbow order from its N-terminus (blue) to its C-terminus (red). In  addition, the A subunit is embedded in its transparent molecular  surface. The catalytic (C; 309 residues) subunit (magenta) is drawn in  ribbon form. Note the close structural resemblance of the A and B  subunits. [Based on an X-ray structure by Yigong Shi, Princeton  University. PDBid 2NPP.]    metabolism (Section 16-3B). The PPP member known as PP2A participates  in a wide variety of regulatory processes including those governing metabo¬  lism, DNA replication, transcription, and development. PP2A is a het-  erotrimer that consists of a scaffold (A) subunit that binds both a catalytic (C)  subunit and a regulatory (B) subunit. The A subunit, which consists of 15 im¬  perfect tandem repeats of a 39-residue sequence termed HEAT (because it oc¬  curs in proteins named //untingtin, TT3, A subunit of PP2A, and TORI),  has a remarkable structure in which the HEAT repeats are joined in a horse-  shoe-shaped solenoidal arrangement (Fig. 13-16/z; opposite).   The X-ray structure of a PP2A holoenzyme (complete enzyme; Fig. 13-16^)  reveals, unexpectedly, that its regulatory subunit consists of 8 tandem HEAT-  like repeats arranged like those of the A subunit, despite their lack of sequence  similarity. The C subunit binds to the A subunits concave surface along a  ridge of conserved hydrophobic side chains spanning HEAT repeats 11 to 15.  The regulatory subunit similarly interacts with the A subunit s HEAT repeats  2 to 8 and also binds to the C subunit via a ridge spanning its own HEAT-  like repeats 6 to 8. The highly acidic, convex side of the regulatory subunit  (lower part of Fig. 13-16£) is thereby left unoccupied, which suggests that it  interacts with substrate proteins.   PP2As catalytic and scaffold subunits both have two isoforms, and there  are 16 isoforms of the regulatory subunit. This results in an enormous panoply  of enzymes that are targeted to different phosphoproteins in distinct subcel-  lular sites during different developmental stages. This complexity is a major  cause of our limited understanding of how PP2A carries out its diverse cellu¬  lar functions, even though it comprises between 0.3 and 1% of cellular  proteins.   r The PPP family also includes calcineurin (also called PP2B), a Ser/Thr  phosphatase that is activated by Ca 2+ . Calcineurin plays an essential role  in T cell proliferation. It is inhibited by the action of drugs such as  cyclosporin A, which is used clinically to suppress immune system function  following organ transplantation.   • Explain how the activation loop governs  substrate access to the active site of a  protein tyrosine kinase.   • Summarize the roles of SH 2 and SH3  domains, Ras, GTP, and protein kinases in  transmitting a signal from an RTK to a  transcription factor.   • What is the function of a GEF and a GAP  in signal transduction?   • Explain how a kinase cascade works. Why  is this advantageous in the context of  hormone signaling?   • Describe how SH 2 and SH3 domains and  Tyr phosphorylation influence PTK activity.   • Explain why cells contain an array of  protein phosphatases as well as protein  kinases.    • G protein-coupled receptors contain seven membrane-spanning helices and un¬  dergo conformational changes when a hormone binds.   • Ligand binding to a G protein-coupled receptor induces the a subunit of the  associated G protein to exchange GDP for GTP and dissociate from the p and 7  subunits.   • Adenylate cyclase is activated to produce cAMP, which in turn activates protein  kinase A.   • Signaling activity is limited through the action of phosphodiesterases that act on  cAMP and cGMP    The second major class of signal transduction pathways that we will discuss  involves heterotrimeric G proteins. These proteins are members of the su¬  perfamily of regulatory GTPases that are collectively known as G proteins,  which, as we have seen, are named for their ability to bind the guanine nu¬  cleotides GTP and GDP and hydrolyze GTP to GDP and P r The monomeric  G proteins are essential for a wide variety of processes, including signal trans¬  duction (e.g., Ras; Section 13-2B), vesicle trafficking (Section 9-4E), the  growth of actin micro filaments (Section 7-2C), translation (as ribosomal ac¬  cessory factors; Section 27-4), and protein targeting [as components of the sig¬  nal recognition particle (SRP) and the SRP receptor; Section 9-4D]. The many  G proteins share common structural motifs that bind guanine nucleotides and  catalyze the hydrolysis of GTP.    Many heterotrimeric G proteins participate in signal transduction systems  that consist of three major components (Fig. 13-17):   1. G protein-coupled receptors (GPCRs), transmembrane proteins that  bind their corresponding ligand (e.g., a hormone) on their extracellular  side, which induces a conformational change on their cytoplasmic side.   2. Heterotrimeric G proteins, which are anchored to the cytoplasmic side  of the plasma membrane and which are activated by a GPCR when it  binds its corresponding ligand.   3. Adenylate cyclase (AC), a transmembrane enzyme that is activated (or  in some cases inhibited) by activated heterotrimeric G proteins.    Activated AC catalyzes the synthesis of adenosine-3',5'-cyclic monophos¬  phate (3',5'-cyclic AMP or cAMP) from ATP.    The cAMP, in turn, binds to a variety of proteins so as to activate numerous cel¬  lular processes. Thus, as Earl Sutherland first showed, cAMP is a second messen¬  ger, that is, it intracellularly transmits the signal originated by the extracellular ligand.   What are the mechanisms through which the binding of ligand to an ex¬  tracellular receptor induces AC to synthesize cAMP in the cytosol? In answer¬  ing this question we will see that the signaling system outlined above has a  surprising complexity that endows it with immense capacity for both signal  amplification and regulatory flexibility.   The GPCRs form one of the largest known protein families (>800 species in  humans, which comprises 3.5% of the ~23,000 genes in the human genome).  This family includes receptors for nucleosides, nucleotides, Ca 2+ , catecholamines  (Section 13-IB) and other biogenic amines (e.g., histamine and serotonin;  Section 21-6B), the various eicosanoids (Section 9-IF), and a variety of  peptide and protein hormones. In addition, GPCRs have essential sensory  functions: They constitute the olfactory (odorant) and gustatory (taste) recep¬  tors (of which there are estimated to be 460 different types in humans), as  well as the several light-sensing proteins in the retina, which are known as  rhodopsins. The importance of these receptors is also evident from the fact  that over half of the therapeutic drugs presently in use target specific GPCRs.   The GPCRs are all integral membrane proteins with seven transmembrane  a helices of generally uniform size: 20 to 27 residues, which is sufficient to span  a lipid bilayer. However, their N- and C-terminal segments and the loops con¬  necting their transmembrane helices vary widely in length. These are the por¬  tions of the protein that participate in binding ligands (on their extracellular  side) and heterotrimeric G proteins (on their cytoplasmic side). Many GPCRs  are posttranslationally modified by A^glycosylation and/or by the palmitoylation  of a Cys residue, so they are also lipid-linked glycoproteins (Section 9-3B).   The structure of the human (3 2 -adrenergic receptor with bound carazolol  (at left), an antagonist, is shown in Figure 13-18. The ligand-binding site is   1) heterotrimeric G  protein to  exchange its  bound GDP for  GTP    The G protein*GTP complex  dissociates from the receptor  and activates adenylate  cyclase to produce the  second messenger cAMP    FIG. 1 3-1 7 Overview of Heterotrimeric G protein-dependent bound GDP for GTP, triggering a process that activates adenylate   signaling. In this widely used pathway, hormone binding to its recep- cyclase to produce the second messenger cAMP.    defined by a portion of the helical core of the protein as well as its extracel¬  lular loops. Aside from this general location, there are few similarities in the    structures of the binding sites in different GPCRs, which is  consistent with the observation that each receptor is specific  for only one or a few ligands.   GPCRs function much like allosteric proteins such as he¬  moglobin (Section 7-1). By alternating between two discrete  conformations , one with ligand bound and one without , the re¬  ceptor can transmit an extracellular signal to the cell interior. In  the case of GPCRs, this conformational change is propagated    FIG. 1 3-1 8 X-Ray structure of the human p 2 -adrenergic receptor.   The structure is viewed parallel to the plane of the membrane with its  approximate position therein indicated. The protein is represented by its  transparent molecular surface with its polypeptide chain in ribbon form  colored in rainbow order from its N-terminus (blue) to its C-terminus (red).  Note its bundle of seven nearly parallel transmembrane helices. The carazolol  ligand and the Cys-linked palmitoyl group are shown in space-filling form  with carazolol C magenta, palmitoyl and Cys side chain C green, N blue,   0 red, and S yellow. [Based on an X-ray structure by Brian Kobilka and  Raymond Stevens, Scripps Research Institute. PDBid 2RH1.]   Chapter 13 Biochemical Signaling through the receptors transmembrane helices to its cytoplasmic face so as to   permit the binding of the corresponding heterotrimeric G protein. This model  of receptor action is similar to the operation of membrane transport proteins  (e.g., Fig. 10-13); in fact, some membrane-bound receptors are ion channels  that switch between the open and closed conformations in response to ligand  binding.   Receptors Are Subject to Desensitization. A hallmark of biological signal¬  ing systems is that they adapt to long-term stimuli by reducing their response  to them, a process named desensitization. These signaling systems therefore re¬  spond to changes in stimulation levels rather than to their absolute values. In the  case of the (3-adrenergic receptor, continuous exposure to epinephrine leads to  the phosphorylation of one or more of the receptors Ser residues. This phos¬  phorylation, which is catalyzed by a specific kinase that acts on the  hormone-receptor complex but not on the receptor alone, reduces the recep¬  tors affinity for epinephrine. If the epinephrine level is reduced, the receptor  is slowly dephosphorylated, eventually restoring the cells initial epinephrine  sensitivity. Desensitization may also involve endocytosis of the receptor, which  decreases its availability on the cell surface.   Heterotrimeric G proteins, as their name implies, are G proteins that consist  of an a, (3, and y subunit (45, 37, and 9 kD, respectively). The X-ray struc¬  tures of entire heterotrimeric G proteins were independently determined by  Alfred Gilman and Stephan Sprang (Fig. 13-19) and by Heidi Hamm and Paul  Sigler. The large a subunit, designated G a , consists of two domains connected  by two polypeptide linkers (Fig. 13-19 a): (1) a highly conserved GTPase do¬  main that is structurally similar to those in monomeric G proteins such as Ras   (a) The G a subunit is pink with the segments known as Switch I, II,  and III green, blue, and red, respectively. A bound GDP is shown in  space-filling form with C green, N blue, 0 red, and P yellow. The Gp  subunit’s N-terminal segment is light blue and each blade of its  (3-propeller has a different color. The G 7 subunit is gold. The plasma  membrane is probably at the top of the drawing, as inferred from the  positions of the N terminus of G a and the neighboring C terminus of   G t which are lipid-linked in vivo, although the orientation of the  protein relative to the membrane is unknown, (b) View related to that  in Part a by a 90° rotation about the horizontal axis, that is, looking  from the general direction of the plasma membrane. The protein is  colored as in Part a except that the G a subunit is mainly gray. [Based  on an X-ray structure by Alfred Gilman and Stephan Sprang,   University of Texas Southwestern Medical Center. PDBid 1GP2J  O See Interactive Exercise 12.   and hence is known as a Ras-like domain, and (2) a helical domain that is Section 3 Heterotrimeric G Proteins   unique to heterotrimeric G proteins. The Ras-like domain contains the gua¬  nine nucleotide-binding site in a deep cleft and is anchored to the membrane  by a myristoyl or palmitoyl group, or both, covalently attached near the pro¬  teins N-terminus. The Gp subunit, which is anchored to the membrane via  prenylation of its C-terminus, consists of an N-terminal helical domain fol¬  lowed by a C-terminal domain comprising seven 4-stranded antiparallel (3 sheets  arranged like the blades of a propeller—a so-called (3 propeller (Fig. 13-19^).   The G 7 subunit consists mainly of two helical segments joined by a polypeptide  link (Fig. 13-19£). It is closely associated with Gp along its entire extended length  through mainly hydrophobic interactions. G 7 binds to Gp with such high affin¬  ity that they dissociate only under denaturing conditions. Consequently, we will  henceforth refer to their complex as Gp 7 .   In its unactivated state, a heterotrimeric G protein maintains its het¬  erotrimeric state and its G a subunit binds GDP. However, the binding of such  a G a • GDP—G^ complex to its cognate GPCR in complex with a ligand induces  the G a subunit to exchange its bound GDP for GTP. Thus, the ligand-GPCR  complex functions as the G a subunits guanine nucleotide exchange factor  (GEF).   When GTP is bound to G a , its y phosphoryl group promotes conforma¬  tional changes in three of G a ’s so-called switch regions (Fig. 13-19 a), caus-  ing G a to dissociate from Gp 7 . This occurs because the binding of GTP’s y  phosphoryl group and the binding of Gp 7 to G a are mutually exclusive; the  y phosphoryl group hydrogen bonds with side chains in Switches I and II so  as to prevent the segments from interacting with the loops and turns at the  bottom of Gp s (3-propeller. Switches I and II have counterparts in other  G proteins of known structure. Comparison of the X-ray structures of the  G a • GDP-Gp 7 complex and Gp 7 alone indicates that the structure of Gp 7  is unchanged by its association with G a • GDP. Nevertheless, both G a and  Gp 7 are active in signal transduction; they interact with additional cellular  components, as we discuss below.   The effect of G protein activation is short-lived, because G a is also a  GTPase that catalyzes the hydrolysis of its bound GTP to GDP + P z , although  at the relatively sluggish rate of 2 to 3 min \ GTP hydrolysis causes the het¬  erotrimeric G protein to reassemble as the inactive G a • GDP-Gp 7 complex.   This prevents a runaway response to ligand binding to a GPCR.   Heterotrimeric G Proteins Activate Other Proteins. A human cell can con¬  tain numerous different kinds of heterotrimeric G proteins, since there are 21  different a subunits, 6 different (3 subunits, and 12 different y subunits. This  heterozygosity presumably permits various cell types to respond in different  ways to a variety of stimuli.   One of the major targets of the heterotrimeric G protein system is the en¬  zyme adenylate cyclase (described more fully in the next section). For exam¬  ple, when a G a • GTP complex dissociates from Gp 7 , it may bind with high  affinity to AC, thereby activating the enzyme. Such a G a protein is known as  a stimulatory G protein, G sot . Other G a proteins, known as inhibitory G pro¬  teins, G i( *, inhibit AC activity. The heterotrimeric G s and G t proteins, which  differ in their a subunits, may actually contain the same (3 and y subunits.   Other types of heterotrimeric G proteins—acting through their G a or Gp 7  units—stimulate the opening of ion channels, participate in the phosphoinosi-  tide signaling system (Section 13-4), activate phosphodiesterases, and activate  protein kinases.   Because a single ligand-receptor interaction can activate more than one G  protein, this step of the signal transduction pathway serves to amplify the orig¬  inal extracellular signal. In addition, several types of ligand-receptor complexes  may activate the same G protein so that different extracellular signals elicit the  same cellular response.   chapter 13 Biochemical signaling c Adenylate Cyclase Synthesizes cAMP to Activate Protein Kinase A    Mammals have nine different isoforms of adenylate cyclase, which are each ex¬  pressed in a tissue-specific manner and differ in their regulatory properties.  These ~120-kD transmembrane glycoproteins each consist of a small N-ter-  minal domain (N), followed by two repeats of a unit consisting of a transmem¬  brane domain (M) followed by two consecutive cytoplasmic domains (C), thus  forming the sequence NM 1 C la C 1 bM 2 C2aC 2 b (Fig* 13-20). The 40% identical  C la and C 2a domains associate to form the enzymes catalytic core, whereas Cu,,  as well as C la and C 2a , bind regulatory molecules. For example, G sa binds to  C 2a to activate AC, and G ia binds to C la to inhibit the enzyme. Other regu¬  lators of AC activity include Ca 2+ and certain Ser/Thr protein kinases. Clearly,  cells can adjust their cAMP levels in response to a great variety of stimuli.   The structure of intact adenylate cyclase is not known, but X-ray struc¬  tural studies of the catalytic domains indicate that G sa • GTP binds to the  C ia • C 2a complex via its Switch II region. This binding alters the orientation  of the Ci a and C 2a domains so as to position their catalytic residues for the  efficient conversion of ATP to cAMP. When G sa hydrolyzes its bound GTP,  its Switch II region reorients so that it can no longer bind to C 2a , and the  adenylate cyclase reverts to its inactive conformation.    Protein Kinase A Is Activated by Binding Four cAMP. cAMP is a polar,  freely diffusing second messenger. In eukaryotic cells, its main target is pro¬  tein kinase A (PKA; also known as cAMP-dependent protein kinase or  cAPK), an enzyme that phosphorylates specific Ser or Thr residues of numer¬  ous cellular proteins. These proteins all contain a consensus kinase-recognition  sequence, Arg-Arg-X-Ser/Thr-Y, where Ser/Thr is the phosphorylation site, X  is any small residue, and Y is a large hydrophobic residue.   In the absence of cAMP, PKA is an inactive heterotetramer of two regu¬  latory and two catalytic subunits, R 2 C 2 . The cAMP binds to the regulatory  subunits to cause the dissociation of active catalytic monomers:   R 2 C 2 + 4 cAMP 2C + R 2 (cAMP) 4   The intracellular concentration of cAMP therefore determines the fraction of PKA  in its active form and thus the rate at which it phosphorylates its substrates.    FIG. 1 3-20 Schematic diagram of a  typical mammalian adenylate cyclase. The M 1   and M 2 domains are each predicted to contain  six transmembrane helices. C 1a and C 2a form the  enzyme’s pseudosymmetric catalytic core. The  domains with which various regulatory proteins  are known to interact are indicated. [After  Tesmer, J.J.G. and Sprang, S.R., Curr. Opin.   FIG. 1 3-22 X-Ray structure of the regulatory (R) subunit of bovine protein kinase A  (PKA) in complex with cAMP. Domain A is cyan and domain B is orange. The cAMP molecules  are shown in space-filling form colored according to atom type (C green, N blue, 0 red, and P yellow).   The region containing the autoinhibitor segment is magenta. The N-terminal 91 residues of the R subunit,  which mediate its dimerization, are absent from this polypeptide. [Based on an X-ray structure by Susan Taylor,  University of California at San Diego. PDBid 1 RGSJ    FIG. 1 3-21 X-Ray structure of the catalytic (C) subunit of mouse protein kinase A (PKA).   The protein, which is shown in its “standard” view, is in complex with ATP and a 20-residue peptide  segment of a naturally occurring protein kinase inhibitor. The N-terminal domain is pink, the  C-terminal domain is cyan, and the activation loop containing Thr 197 is light blue. The  polypeptide inhibitor is orange and its pseudotarget sequence, Arg-Arg-Asn-Ala-lle, is magenta  (the Ala, which replaces the Ser or Thr of a true substrate, is white). The substrate ATP and the  phosphoryl group of phospho-Thr 197 are shown in space-filling form and the side chains of  the catalytically essential Arg 165, Asp 166, and Thr 197 are shown in stick form, all colored  according to atom type (C green, N blue, 0 red, and P yellow). Note that the inhibitor’s  pseudotarget sequence is close to ATP’s 7 phosphate group, the group that the enzyme  transfers to the Ser or Thr of the target sequence. [Based on an X-ray structure by Susan  Taylor and Janusz Sowadski, University of California at San Diego. PDBid 1ATPJ © See  Interactive Exercise 13 and Kinemage Exercise 15.    The X-ray structure of the 350-residue C subunit of mouse PKA in com¬  plex with ATP and a 20-residue inhibitor peptide, which was determined by  Susan Taylor and Janusz Sowadski, is shown in Fig. 13-21. The C subunit  closely resembles other protein kinases of known structure (e.g., Figs. 13-5*2  and 13-12). In the PKA structure, the deep cleft between the lobes is occu¬  pied by ATP and a segment of the inhibitor peptide that resembles the   5- residue consensus sequence for phosphorylation except that the phosphory-  lated Ser/Thr is replaced by Ala. Thr 197, which is part of the activation loop,  must be phosphorylated for maximal activity. The phosphoryl group at Thr  197 interacts with Arg 165, a conserved catalytic residue that is adjacent to  Asp 166, the catalytic base that activates the substrate proteins target Ser/Thr  hydroxyl group for phosphorylation. Thus, the phosphoryl group at PKAs Thr  197 functions to properly orient its active site residues.   The R subunit of protein kinase A competitively inhibits its C subunit. The R  subunit contains two homologous cAMP-binding domains, A and B, and a so-  called autoinhibitor segment (Fig. 13-22). In the inactive R 2 C 2 complex, the  autoinhibitor segment, which resembles the C subunits substrate, binds in the C  subunits active site (as does the inhibitory peptide in Fig. 13-21) so as to block  substrate binding. Each R subunit cooperatively binds two cAMPs. When the B  domain lacks bound cAMP, it masks the A domain so as to prevent it from  binding cAMP. However, the binding of cAMP to the B domain triggers a  massive conformational change that permits the A domain to bind cAMP,  which in turn releases the now-active C subunits from the complex.   The targets of PKA include enzymes involved in glycogen metabolism. For  example, when epinephrine binds to the (3-adrenergic receptor of a muscle cell,  the sequential activation of a heterotrimeric G protein, adenylate cyclase, and  PKA leads to the activation of glycogen phosphorylase, thereby making glucose-   Each step of a signal transduction pathway can potentially be regulated, so  the nature and magnitude of the cellular response ultimately reflect the presence and  degree of activation or inhibition of all the preceding components of the pathway.  For example, the adenylate cyclase signaling pathway can be limited or reversed  through ligand activation of a receptor coupled to an inhibitory G protein. The  activity of the cAMP second messenger can be attenuated by the action of phos¬  phodiesterases that hydrolyze cAMP to AMP (see below). In addition, reactions  catalyzed by PKA are reversed by protein Ser/Thr phosphatases (Section 13-  2D). Some of these features of the adenylate cyclase signaling pathway are   binding of hormone to a stimulatory receptor R s (1) induces it to bind  the heterotrim eric G protein G s , which in turn stimulates the G sa subunit  (2) to exchange its bound GDP for GTP. The G sa • GTP complex then  dissociates from Gp 7 (3) and (4) stimulates adenylate cyclase (AC) to  convert ATP to cAMP (5). This stimulation stops when G sa catalyzes the  hydrolysis of its bound GTP to GDP ( 6 ). The binding of hormone to the  inhibitory receptor R, (7) triggers an almost identical chain of events  (8-11) except that the presence of the G ia • GTP complex inhibits  adenylate cyclase (10). cAMP activates protein kinase A (PKA; R 2 C 2 )    by binding to the regulatory dimer as R 2 • cAMP 4 , causing the catalytic  subunit C to dissociate (12), and activates various cellular proteins (13)  by catalyzing their phosphorylation. The sites of action of certain toxins  are indicated. Signaling is limited by the action of phosphatases (14)  and cAMP phosphodiesterase (15). Toxins such as cholera toxin and  pertussis toxin act by blocking the hydrolysis of GTP from either G sa •  GTP (6) or G ia • GTP (8), enhancing their activities.   Compare the effects of cholera toxin and pertussis toxin on  the cellular response.    illustrated in Fig. 13-23. Many drugs and toxins exert their effects by modify¬  ing components of the adenylate cyclase system (Box 13-4).   In any chemically based signaling system , the signal molecule must eventually be  eliminated in order to control the amplitude and duration of the signal and to  prevent interference with the reception of subsequent signals. In the case of cAMP,    Complex processes such as the adenylate cyclase signaling  system can be sabotaged by a variety of agents. For example, the  methylated purine derivatives caffeine (an ingredient of coffee  and tea), theophylline (an asthma treatment), and theobromine  (found in chocolate)    are stimulants because they antagonize adenosine receptors that  act through inhibitory G proteins. This antagonism results in an  increase in cAMP concentration.   Deadlier effects result from certain bacterial toxins that inter¬  fere with heterotrimeric G protein function. The toxin released by  Vibrio cholerae (the bacterium causing cholera) triggers massive  fluid loss of over a liter per hour from diarrhea. Victims die from  dehydration unless their lost water and salts are replaced.    Cholera toxin, an 87-kD protein of subunit composition AB 5 ,  binds to ganglioside G M1 (Fig. 9-9) on the surface of intestinal  cells via its B subunits. This permits the toxin to enter the cell,  probably via receptor-mediated endocytosis, where an —195-  residue proteolytic fragment of its A subunit is released. This  fragment catalyzes the transfer of the ADP-ribose unit from  NAD + to a specific Arg side chain of G sa .   ADP-ribosylated G sa • GTP can activate adenylate cyclase but  cannot hydrolyze its bound GTP (Fig. 13-23). As a consequence,  the adenylate cyclase is locked in its active state and cellular  cAMP levels increase —100-fold. Intestinal cells, which normally  respond to small increases in cAMP by secreting digestive fluid  (an HCO 3 -rich salt solution), pour out enormous quantities of this  fluid in response to the elevated cAMP concentrations.   Other bacterial toxins act similarly. Certain strains of E. coli cause  a diarrheal disease similar to but less serious than cholera through  their production of heat-labile enterotoxin, a protein that is closely  similar to cholera toxin (their A and B subunits are >80% identical)  and has the same mechanism of action. Pertussis toxin [secreted  by Bordetella pertussis, the bacterium that causes pertussis  (whooping cough), which is responsible for —400,000 infant deaths  per year worldwide] is an AB 5 protein homologous to cholera toxin  that ADP-ribosylates a specific Cys residue of G ia . The modified G ia  cannot exchange its bound GDP for GTP and therefore cannot  inhibit adenylate cyclase (Fig. 13-23).    this second messenger is hydrolyzed to AMP by enzymes known as cAMP-  phosphodiesterases (cAMP-PDEs).   The PDE superfamily, which includes both cAMP-PDEs and cGMP-PDEs  (cGMP is the guanine analog of cAMP), is encoded in mammals by at least 20  different genes grouped into 12 families (PDE1 through PDE 12). Moreover,  many of the mRNAs transcribed from these genes have alternative initiation sites  and alternative splice sites (Section 26-3B), so that mammals express ~50 PDE  isoforms. These are functionally distinguished by their substrate specificities (for  cAMP, cGMP, or both) and kinetic properties, their responses (or lack of them)   Chapter 13 Biochemical Signaling to various activators and inhibitors (see below), and their tissue, cellular, and   subcellular distributions. The PDEs have characteristic modular architectures  with a conserved ~270-residue catalytic domain near their C-termini and widely  divergent regulatory domains or motifs, usually in their N-terminal portions.  Some PDEs are membrane-anchored, whereas others are cytosolic.   PDE activity, as might be expected, is elaborately controlled. Depending  on its isoform, a PDE may be activated by one or more of a variety of agents,  including Ca 2+ ion and phosphorylation by PKA and insulin-stimulated  protein kinase. Phosphorylated PDEs are dephosphorylated by a variety of  protein phosphatases. Thus, the PDEs provide a means for cross talk between  cAMP-based signaling systems and those using other types of signals.   T PDEs are inhibited by a variety of drugs that influence such widely diver¬  gent disorders as asthma, congestive heart failure, depression, erectile dysfunc¬  tion, inflammation, and retinal degeneration. Sildenafil (trade name Viagra),   • Summarize the steps of signal transduc¬  tion from a GPCR to phosphorylation of  target proteins by PKA.   • Explain why a GPCR can be considered  to be an allosteric protein.   • Describe how G proteins are activated  and inactivated. What protein functions as  a GEF?   • What is the function of a second messen¬  ger such as cAMP?    a compound used to treat erectile dysfunction, specifically inhibits PDE5,  which hydrolyzes only cGMP. Sexual stimulation in males causes penile nerves  to release nitric oxide (NO), which activates guanylate cyclase to produce  cGMP from GTP. The cGMP induces vascular smooth muscle relaxation in  the penis, thereby increasing the inflow of blood, which results in an erection.  This cGMP is eventually hydrolyzed by PDE5. Sildenafil is therefore an ef¬  fective treatment in men who produce insufficient NO and hence cGMP to  otherwise generate a satisfactory erection.    Signal transduction via the phosphoinositide pathway generates the second  messenger inositol trisphosphate, which triggers Ca 2+ release, and diacylglycerol,  which activates protein kinase C.   In the presence of Ca 2+ , calmodulin binds and activates its target proteins.   A hormone can activate multiple signal transduction pathways to elicit a variety of  intracellular responses.    A discussion of signal transduction pathways would not be complete without a  consideration of the phosphoinositide pathway, which mediates the effects of  a variety of hormones. This signaling pathway requires a receptor with seven  transmembrane segments, a heterotrimeric G protein, a specific kinase, and a  phosphorylated glycerophospholipid that is a minor component of the plasma  membranes inner leaflet. It involves the production of three second messengers:  inositol-1,4,5-trisphosphate (IP3) , Ca 2+ , and 1,2 -diacylglycerol (DAG).   A Ligand Binding Results in the Cytoplasmic Release of the Second Section 4 The Phosphoinositide Pathway   Ligand binding to its receptor, such as epinephrine binding to the a x -adrenergic  receptor, activates a heterotrimeric G protein, G q , whose membrane-  anchored a subunit in complex with GTP diffuses laterally along the plasma  membrane to activate the membrane-bound enzyme phospholipase C (PLC;   Fig. 13-24, upper left). Activated PLC catalyzes the hydrolysis of phos-  phatidylinositol-4,5-bisphosphate (PIP 2 ) at its glycero-phospho bond    FIG. 1 3-24 The phosphoinositide signaling system. Ligand  binding to a cell-surface receptor R (1) activates phospholipase C  (PLC) through the heterotrimeric G protein G q ( 2 ). Phospholipase C  catalyzes the hydrolysis of PIP 2 to IP 3 and DAG (3). The water-soluble  IP 3 stimulates the release of Ca 2+ sequestered in the endoplasmic  reticulum (4), which in turn activates numerous cellular processes  through the intermediacy of calmodulin (CaM; 5). The nonpolar DAG  remains associated with the membrane, where it activates protein  kinase C (PKC; 6 ) to phosphorylate and thereby modulate the    activities of a number of cellular proteins (7). PKC activation also  requires the presence of the membrane lipid phosphatidylserine (PS)  and Ca 2+ . Phosphoinositide signaling is limited by GTP hydrolysis  on qa (8) and by inositol polyphosphate 5-phosphatase, which acts   Which components of the signaling system are membrane-  bound and which are soluble?    FIG. 1 3-25 The phospholipase C  reaction. Phospholipase C cleaves PIP 2 to  produce diacylglycerol (DAG) and inositol-1,4,5-  trisphosphate (IP 3 ), both of which are second  messengers. (The bis and tris prefixes denote,  respectively, two and three phosphoryl groups  that are linked separately to the inositol; in  di- and triphosphates, the phosphoryl groups are  linked sequentially.)   FIG. 1 3-26 X-Ray structure of rat testis  calmodulin. This monomeric 148-residue  protein, which is colored in rainbow order from  its N-terminus (blue) to its C-terminus (red),  contains two remarkably similar globular domains  separated by a seven-turn a helix. The two Ca 2+  ions bound to each domain are represented by  cyan spheres. The side chains liganding the  Ca 2+ ions are drawn in stick form colored  according to atom type (C green, N blue, and  0 red). [Based on an X-ray structure by Charles  Bugg, University of Alabama at Birmingham.  PDBid 3CLN.] Q See Kinemage Exercise  16-1.    (Section 9-1C), yielding inositol-1,4,5-trisphosphate (IP 3 ) and 1,2-diacylglycerol  (DAG; Fig. 13-25). PLC, which in mammals is actually a set of 13 isozymes,  several of which have splice variants, requires the presence of Ca 2+ for enzy¬  matic activity. It has a hydrophobic ridge consisting of three protein loops that  is postulated to penetrate into the membranes nonpolar region during  catalysis. This would explain how the enzyme can catalyze hydrolysis of the  membrane-bound PIP 2 , leaving the DAG product associated with the  membrane.   The Charged IP 3 Molecule Is a Water-Soluble Second Messenger. The   hydrolysis of PIP 2 sets in motion both cytoplasmic and membrane-bound  events. While DAG acts as a second messenger in the membrane (Section 13-4C),  IP 3 diffuses through the cytoplasm to the endoplasmic reticulum (ER). There,  it binds to and induces the opening of a Ca 2+ transport channel (an example  of a receptor that is also an ion channel), thereby allowing the efflux of  Ca 2+ from the ER. This causes the cytosolic [Ca 2+ ] to increase from ~0.1 piM  to as much as 10 mM, which triggers such diverse cellular processes as glu¬  cose mobilization and muscle contraction through the intermediacy of the  Ca 2+ -binding protein calmodulin (see below) and its homologs. The ER con¬  tains embedded Ca 2+ -ATPases that actively pump Ca 2+ from the cytosol back  into the ER (Section 10-3B) so that in the absence of IP 3 , the cytosolic [Ca 2+ ]  rapidly returns to its resting level.   Calmodulin (CaM) is a ubiquitous, eukaryotic Ca 2+ -binding protein that par¬  ticipates in numerous cellular regulatory processes. In some of these, CaM  functions as a free-floating monomeric protein, whereas in others it is a sub¬  unit of a larger protein. The X-ray structure of this highly conserved 148-  residue protein has a curious dumbbell-like shape in which two structurally  similar globular domains are connected by a seven-turn a helix (Fig. 13-26).  Note the close structural resemblance between CaM and the Ca 2+ -binding TnC  subunit of the muscle protein troponin (Fig. 7-31).   CaM’s two globular domains each contain two high-affinity Ca 2+ -binding  sites. The Ca 2+ ion in each of these sites is octahedrally coordinated by oxy¬  gen atoms from the backbone and side chains as well as from a protein-asso¬  ciated water molecule. Each of the Ca 2+ -binding sites is formed by nearly  superimposable helix-loop-helix motifs known as EF hands (Fig. 13-27) that  form the Ca 2+ -binding sites in numerous other Ca 2+ -binding proteins of  known structure.    FIG. 1 3-27 The EF hand. The Ca 2+ -binding sites in many proteins that sense the  level of Ca 2+ are formed by helix-loop-helix motifs named EF hands. [After Kretsinger,  R.H., Annu. Rev. Biochem. 45, 241 (1976).] O See Kinemage Exercise 16-1.    Ca 2+ -CaM Activates Its Target Proteins via an Intrasteric Mechanism.   The binding of Ca 2+ to either domain of CaM induces a conformational  change in that domain, which exposes an otherwise buried Met-rich hy¬  drophobic patch. This patch, in turn, binds with high affinity to the CaM-  binding domains of numerous Ca 2+ -regulated protein kinases. These  CaM-binding domains have little mutual sequence homology but are all ba¬  sic amphiphilic a helices.   Despite uncomplexed CaM’s extended appearance (Fig. 13-26), a variety  of studies indicate that both of its globular domains bind to a single target  helix. This was confirmed by the NMR structure (Fig. 13-28) of  (Ca 2+ ) 4 —CaM in complex with a target polypeptide, a segment of skeletal  muscle myosin light chain kinase (MLCK). This enzyme, a homolog of the  PKA C subunit, phosphorylates and thereby activates the light chains of the  muscle protein myosin (Section 7-2A). Thus, CaM’s central a helix serves as  a flexible tether rather than a rigid spacer, a property that probably allows  CaM to bind to a wide range of target polypeptides. Experiments show that  both of CaM’s globular domains are required for CaM to activate its targets:  CaM domains that have been separated by proteolytic cleavage bind to their  target peptides but do not cause enzyme activation.   How does Ca 2+ —CaM activate its target protein kinases? MLCK contains  a C-terminal segment whose sequence resembles that of MLCK’s target  polypeptide on the light chain of myosin but lacks a phosphorylation site. A  model of MLCK, based on the X-ray structure of the 30% identical C sub¬  unit of PKA, strongly suggests that this segment of MLCK acts as an autoin¬  hibitor by binding in the kinase’s active site. Indeed, the excision of MLCK’s  autoinhibitor peptide by limited proteolysis permanently activates the enzyme.   FIG. 1 3-28 NMR structure of calmodulin in complex with a  target polypeptide. The N-terminal domain of CaM (from the fruit fly  Drosophila melanogaster) is blue, its C-terminal domain is red, the  26-residue target polypeptide, which is from rabbit skeletal muscle  myosin light chain kinase (MLCK), is green, and the Ca 2+ ions are  represented by cyan spheres, (a) A view of the complex in which the  N-terminus of the target polypeptide is on the right. ( b ) The  perpendicular view as seen from the right side of the structure shown    in Part a. In both views, the pseudo-2-fold axis relating the N- and  C-terminal domains of CaM is approximately vertical. Note how the  segment that joins the two domains is unwound and bent (bottom  loop in Part b) so that CaM forms a globular protein that largely  encloses the helical target polypeptide within a hydrophobic tunnel in  a manner resembling two hands holding a rope. [Based on an NMR  structure by Marius Clore, Angela Gronenborn, and Ad Bax, NIH.  PDBid 2BBM.] © See Kinemage Exercise 16-2.    The exposed active site of  the protein kinase binds a (2  substrate protein.   Ca 2+ - CaM binds to its binding site on a   FIG. 1 3-29 A schematic diagram of the Ca 2+ -CaM-dependent  activation of protein kinases. Autoinhibited kinases have an N- or  C-terminal “pseudosubstrate” sequence (pink) that binds at or near the  enzyme’s active site (orange) so as to inhibit its function. The autoin-  hibitory segment is in close proximity with or overlaps a Ca 2+ -CaM    binding sequence. Consequently, Ca 2+ -CaM (green) binds to the  sequence so as to extract it from the enzyme’s active site, thereby  activating the enzyme to bind and phosphorylate other proteins  (purple), eliciting a cellular response. [After Crivici, A. and Ikura, M.,  Annu. Rev. Biophys. Biomol. Struct. 24, 88 (1995).]    MLCK’s CaM-binding segment overlaps the autoinhibitor peptide. Thus, the  binding of Ca 2+ -CaM to this peptide segment extracts the autoinhibitor from  MLCKs active site , thereby activating the enzyme (Fig. 13-29).   Ca 2+ -CaMs other target proteins are presumably activated in the same  way. In fact, the X-ray structures of several homologous protein kinases sup¬  port this so-called intrasteric mechanism. While the details of binding of the  autoinhibitory sequence differ for each of the protein kinases, the general  mode of autoinhibition and activation by Ca 2+ -CaM is the same.   PKA’s R subunit, as we have seen (Section 13-3C), contains a similar au¬  toinhibitory sequence adjacent to its two tandem cAMP-binding domains. In  this case, however, the autoinhibitory peptide is allosterically ejected from the  C subunit’s active site by the binding of cAMP to the R subunit (which lacks  a Ca 2+ -CaM-binding site).   C DAG Is a Lipid-Soluble Second Messenger That Activates  Protein Kinase C   The second product of the phospholipase C reaction, diacylglycerol (DAG),  is a lipid-soluble second messenger. It therefore remains embedded in the  plasma membrane, where it activates the membrane-bound protein kinase  C (PKC) to phosphorylate and thereby modulate the activities of several    different cellular proteins (Fig. 13-24, righi). Multiple PKC enzymes are  known; they differ in tissue expression, intracellular location, and their re¬  quirement for the DAG that activates them. PKC is a phosphorylated, cy¬  tosolic protein in its resting state. DAG increases the membrane affinity of  PKC and also helps stabilize its active conformation. The catalytic activities  of PKC and PKA are similar: Both kinases phosphorylate Ser and Thr  residues.   The X-ray structure of a DAG-bound segment of PKC shows that the  50-residue motif is largely knit together by two Zn 2+ ions, each of which is  tetrahedrally liganded by one His and three Cys side chains (Fig. 13-30). A  DAG analog, phorbol-13-acetate,   binds in a narrow groove between two long nonpolar loops. Very few soluble  proteins have such a large continuous nonpolar region, suggesting that this  portion of PKC inserts into the membrane. Full activation of PKC requires  phosphatidylserine (which is present only in the cytoplasmic leaflet of the  plasma membrane; Fig. 9-32) and, in some cases, a Ca 2+ ion (presumably  made available through the action of the IP 3 second messenger). Like other  signaling systems, the phosphoinositide system is limited by the destruction  of its second messengers, for example, through the action of inositol  polyphosphate 5-phosphatase (Fig. 13-24, lower left).   PLC Acts on Several Phospholipids to Release Different Second  Messengers. Choline-containing phospholipids hydrolyzed by phospholipase  C yield DAGs that differ from those released from PIP 2 and exert different  effects on PKC. Another lipid second messenger, sphingosine released from  sphingolipids, inhibits PKC.   The phosphoinositide signaling pathway in some cells yields a DAG that  is predominantly l-stearoyl-2-arachidonoyl-glycerol. This molecule is further  degraded to yield arachidonate, the precursor of the bioactive eicosanoids  (prostaglandins and thromboxanes; Fig. 9-12), and hence the phosphoinosi¬  tide pathway yields up to four different second messengers. In other cells, IP 3  and diacylglycerol are rapidly recycled to re-form PIP 2 in the inner leaflet of  the membrane. Some receptor tyrosine kinases activate an isoform of phos¬  pholipase C that contains two SH2 domains. This is another example of cross  talk, the interactions of different signal transduction pathways.   FIG. 1 3-30 X-Ray structure of a portion  of protein kinase C in complex with  phorbol-13-acetate. The protein tetrahedrally  ligands two Zn 2+ ions (cyan spheres ), each via  His and Cys side chains (shown in ball-and-stick  form). Phorbol-13-acetate (top), which mimics  the natural diacylglycerol ligand, binds between  two nonpolar protein loops. Atoms are colored  according to type with C green, N blue, 0 red,  and S yellow. [Based on an X-ray structure by  James Hurley, NIH. PDBid 1 PTRJ   Complex systems are, by definition, difficult to understand and substanti¬  ate. Familiar examples include the earths weather system, the economies of  large countries, the ecologies of even small areas, and the human brain.  Biological signal transduction systems, as is amply evident from a reading  of this chapter, are complex systems. Thus, a hormonal signal is typically  transduced through several intracellular signaling pathways, each of which  consists of numerous components, many of which interact with compo¬  nents of other signaling pathways. For example, the insulin signaling    system (Fig. 13-31), although not yet fully elucidated, is clearly highly  complex. Upon binding insulin, the insulin receptor autophosphorylates it¬  self at several Tyr residues (Section 13-2A) and then Tyr-phosphorylates its  target proteins, thereby activating several signaling pathways that control a  diverse array of effects:   1. Phosphorylation of the adaptor protein She, which generates a binding  site for Grb2 s SH2 domain, results in stimulation of a MAP kinase cas¬  cade (Section 13-2B), ultimately affecting growth and differentiation.   2. Phosphorylation of Gab-1 (Grb2-associated binder-1) similarly acti¬  vates the MAP kinase cascade.   FIG. 13-31 Insulin signal transduction. The binding of insulin  to the insulin receptor (IR) induces tyrosine phosphorylations (pY)  that lead to the activation of MAPK via activation of She (1) and  Gab-1 (2). The MAPK cascade regulates the expression of genes  involved in cellular growth and differentiation. Phosphorylation of IRS  proteins (3) activates the PI3K cascade, which leads to changes in  the phosphorylation states of several enzymes, so as to stimulate  glycogen synthesis as well as other metabolic pathways. The PI3K  cascade also participates in the control of vesicle trafficking, leading  to the translocation of the GLUT4 glucose transporter to the cell  surface and thus increasing the rate of glucose transport into the cell.  Glucose transport control is also exerted by the APS/Cbl system  (4) in a PI3K-independent manner involving lipid rafts (Section 9-4C).   Other symbols: Myc, Fos, and Jun (transcription factors), SHP-2 (an  SH2-containing protein tyrosine phosphatase), CAP (Cbl-associated  protein), C3G [a guanine nucleotide exchange factor (GEF)], Crkll  (an SH2/SH3-containing adaptor protein), PDK1 (phosphoinositide-  dependent protein kinase-1), PKB (protein kinase B, also named  Akt), GSK3P (glycogen synthase-3p, which is inhibited by  phosphorylation by PKB), mTOR (for mammalian target of rapamycin,  a PI3K-related protein kinase; rapamycin is an immunosuppressant),  S6 (a protein subunit of the eukaryotic ribosome’s small subunit  whose phosphorylation stimulates translation), and PKC£ and PKCA  (atypical isoforms of protein kinase C). [After Zick, Y., Trends Cell Biol.  11 , 437 (2001).]    3. Phosphorylation of insulin receptor substrate (IRS) proteins (Section  13-2A) activates enzymes known as phosphoinositide 3-kinases  (PI3Ks). These enzymes add a phosphoryl group to the 3'-OH group  of a phosphatidylinositol, often the 4,5-bisphosphate shown in Fig.  13-25. The 3-phosphorylated lipid activates phosphoinositide-  dependent protein kinase-1 (PDK1) that in turn initiates cascades  leading to glycogen synthesis (Section 16-3C) and the translocation of  the glucose transporter GLUT4 to the surface of insulin-responsive  cells (Section 22-2), as well as affecting cell growth and differentiation.   4. Phosphorylation of the APS/Cbl complex (APS for adaptor protein  containing yJekstrin homology and Src homology-2 domains; Cbl is  an SH2/SH3-binding docking protein that is a proto-oncogene prod¬  uct) leads to the stimulation of TC10 (a monomeric G protein) and  to the PI3K-independent regulation of glucose transport involving the  participation of lipid rafts (Section 9-4C).   Thus, by activating multiple pathways, a hormone such as insulin can trig¬  ger a variety of physiological effects that would not be possible in a one  hormone-one target regulatory system.   The predominant approach in science is reductionist: the effort to understand  a system in terms of its component parts. Thus chemists and biochemists ex¬  plain the properties of molecules in terms of the properties of their compo¬  nent atoms, cell biologists explain the nature of cells in terms of the properties  of their component macromolecules, and biologists explain the characteristics  of multicellular organisms in terms of the properties of their component cells.  However, complex systems have emergent properties that are not readily pre¬  dicted from an understanding of their component parts (i.e., the whole is  greater than the sum of its parts). Indeed, life itself is an emergent property  that arises from the numerous chemical reactions that occur in a cell.   In order to elucidate the emergent properties of a complex system, an in¬  tegrative approach is required. For signal transduction systems, such an ap¬  proach would entail determining how each of the components of each  signaling pathway in a cell interacts with all of the other such components  under the conditions that each of these components experiences within its lo¬  cal environment. Yet, existing techniques for doing so are crude at best.  Moreover, these systems are by no means static but vary, over multiple time  scales, in response to cellular and organismal programs. Consequently, the  means for understanding the holistic performance of cellular signal transduc¬  tion systems are only in their earliest stages of development. Such an under¬  standing is likely to have important biomedical consequences since many  diseases, including cancer, diabetes, and a variety of neurological disorders, are  caused by malfunctions of signal transduction systems.   • Describe how ligand binding to a receptor  leads to the production of IP 3 and DAG  and the release of Ca 2+ .   • Which components of the phosphoinosi¬  tide signaling system are soluble and  which are associated with the membrane?   • Describe the mechanisms that limit signal¬  ing by the phosphoinositide pathway.   • Compare Figures 13-7, 13-23, and 13-24.  What do all these pathways have in com¬  mon? How do they differ?   • How do biological signaling systems sup¬  port the idea that the whole is greater  than the sum of its parts?   • Hormones produced by endocrine glands and other tissues regu¬  late diverse physiological processes. The polypeptide hormones in¬  sulin and glucagon control fuel metabolism; fight-or-flight  responses are governed by epinephrine and norepinephrine bind¬  ing to a- and (3-adrenergic receptors; steroid hormones regulate  sexual development and function; and growth hormone stimulates  growth directly and indirectly.   • Hormone signals interact with target tissues by binding to recep¬  tors that transduce the signal to the interior of the cell.   • On ligand binding, receptor tyrosine kinases such as the insulin  receptor undergo autophosphorylation. This activates them to  phosphorylate their target proteins, in some cases triggering a ki¬  nase cascade.   • One growth-promoting pathway involves the monomeric G pro¬  tein Ras and leads to altered gene expression.   • Protein-protein interactions in signaling pathways may require  SH2 and SH3 domains.   • Protein kinases share a common structure that often includes  activation by displacement of an autoinhibitory segment.   • The effects of protein kinases are reversed by the activity of pro¬  tein phosphatases.   • The G protein-coupled receptors (GPCRs) have seven transmem¬  brane helices. On ligand binding, these receptors undergo a con¬  formational change that activates an associated heterotrimeric  G protein.   • The G protein exchanges GDP for GTP and dissociates. G a and  Gp 7 units may activate or inhibit targets such as adenylate cyclase,  which produces the cAMP activator of protein kinase A (PKA).   • In the phosphoinositide pathway, the hormone receptor is associated  with a G protein whose activation in turn activates phospholipase C.  This enzyme catalyzes the hydrolysis of phosphatidylinositol-4,  5 -bisphosphate to yield two second messengers.   • The soluble inositol- 1 ,4, 5 -trisphosphate (IP 3 ) second messenger  opens Ca 2+ channels, causing a rise in intracellular Ca 2+ that  activates protein kinases via the binding of the Ca 2+ -calmodulin  complex. The lipid second messenger diacylglycerol (DAG) acti¬  vates protein kinase C.   • Signaling pathways are complex systems in which a single extra¬  cellular signal can elicit multiple intracellular events, some of  which may also be triggered by other signaling pathways.   1 . Would the pancreatic hormone somatostatin require a receptor  on the surface of or in the cytosol of a target cell?   2 . Retinoic acid (a derivative of vitamin A; Section 9-IF) is a hor¬  mone that mediates immune system function. Would retinoic  acid require a receptor on the surface of a target cell?   3. What biochemical changes are required to convert tyrosine to  (a) norepinephrine and (b) epinephrine?   4. The steroid methandrostenolone is shown here, (a) How does it  differ structurally from testosterone? (b) Why might such drugs  be administered to burn victims?   3. Estimate the binding affinity of a ligand for its receptor from the  following data:    6 . Calculate the binding affinity of a ligand for its receptor from  the following data:    7. Hormone A binds to its receptor with a Af of 5 |JiM. In the pres¬  ence of 2.5 |JtM compound B and 2.0 pM hormone A, 1.0 pM  of A remains unbound. Calculate the binding affinity of B for  the hormone receptor.   8 . Propranolol binds to (3-adrenergic receptors with a ^ of  8.9 X 10 9 M. What concentration of propranolol would be re¬  quired to achieve a 50% reduction in the binding of the recep¬  tor agonist isoproterenol if the agonist concentration is 10 nM  and its dissociation constant for the receptor is 4.8 X 10 8 M?   9. Some bacterial signaling systems involve kinases that transfer a  phosphoryl group to a His side chain. Draw the phospho-His  side chain.   10. Why might a compound resembling ADP function as an in¬  hibitor of a protein kinase?   11 . An insulin receptor substrate (IRS) contains a so-called plekstrin  homology (PH) domain that binds to the inositol head group of  membrane lipids, a phosphotyrosine-binding (PTB) domain that    differs from an SH2 domain, and six to eight Tyr residues that  may be phosphorylated. Explain how each of these features con¬  tributes to signal transduction.   12 . Predict the effect on cell growth of an Sos mutation that de¬  creased its affinity for Ras.   13. Would the following alterations to Src be oncogenic? Explain,  (a) The deletion or inactivation of the SH3 domain, (b) The  mutation of Tyr 416 to Phe.   14. Would the following alterations to Src be oncogenic? Explain,  (a) The mutation of Tyr 327 to Phe. (b) The replacement of Src  residues 249 to 233 with the sequence APTMP.   15. Explain why a protein tyrosine phosphatase would include an  SH2 domain in addition to its phosphatase domain.   16. A growth factor that acts through a receptor tyrosine kinase stim¬  ulates cell division. Predict the effect of a viral protein that in¬  hibits the corresponding protein tyrosine phosphatase.   17. Retroviruses bearing oncogenes will infect cells from their corre¬  sponding host animal but will usually not transform them to cancer  cells. Yet, these retroviruses will readily transform immortalized  cells derived from the same organism. Explain.   18. Trastuzumab (Herceptin; Section 7-3) is an antibody that binds  to the extracellular domain of the growth factor receptor HER2.  Explain why trastuzumab would be an effective treatment for  cancer in which cells overexpress HER2.   19. How does the presence of the poorly hydrolyzable GTP analog  GTPyS (in which an O atom on the terminal phosphate is re¬  placed by an S atom) affect cAMP production by adenylate  cyclase?   20. Mycobacterium tuberculosis is an intracellular bacterium that  causes tuberculosis. The M. tuberculosis genome includes 17 genes  encoding adenylate cyclase, whereas free-living bacteria typically  have just one AC gene. Why might it be an advantage for M.  tuberculosis to generate high adenylate cyclase activity?   21 . One of the toxins produced by Bacillus anthracis (the cause of  anthrax) is known as EF, or edema factor (edema is the abnor¬  mal buildup of extracellular fluid). EF, which enters mammalian   host cells, is a calmodulin-activated adenylate cyclase. Explain  how this toxin causes edema.   22. Another B. anthracis toxin is lethal factor, LF, a protease that  cleaves members of the MAPK kinase family so that they cannot  bind to their downstream MAPK targets in white blood cells.  Explain how LF inhibits activation of the immune system dur¬  ing B. anthracis infection.   23. Explain why mutations of the Arg residue in G sa that is ADP-  ribosylated by cholera toxin are oncogenic mutations.   25. Phosphatidylethanolamine and PIP 2 containing identical fatty  acyl residues can be hydrolyzed with the same efficiency by a cer¬  tain phospholipase C. Will the hydrolysis products of the two  lipids have the same effect on protein kinase C? Explain.   27. Diacylglycerol is a substrate for the enzyme diacylglycerol kinase.  What is the product of this reaction?   28. Explain why activation of diacyglycerol kinase would limit sig¬  naling by the phosphoinositide pathway.   29. Lithium ion, which is used to treat bipolar disorder, interferes  with the phosphoinositide signaling pathway by inhibiting en¬  zymes such as inositol monophosphatase and inositol polyphos¬  phate 1-phosphatase. Predict the effect of Li + on the supply of  cellular inositol, a precursor of phosphatidylinositol and PIP 2 .   30. The SHP-2 protein participates in insulin signaling through its  indirect activation by the insulin receptor (Fig. 13-31), a recep¬  tor tyrosine kinase. Yet SHP-2 is a protein tyrosine phosphatase.  Is this a paradox? Explain.    The sense of smell, that is, the  detection of odorant molecules, is mediated by G protein-coupled  receptors. Are the number of such receptors in the human nose  enough to detect all possible odors? Following ligand binding to  one of these receptors, what happens in the cell? 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A Nutrition Involves Food Intake and Use  B Vitamins and Minerals Assist Metabolic  Reactions   D Thermodynamics Dictates the Direction and  Regulatory Capacity of Metabolic Pathways  E Metabolic Flux Must Be Controlled   B Coupled Reactions Drive Endergonic Processes  C Some Other Phosphorylated Compounds Have  High Phosphoryl Group-Transfer Potentials  D Thioesters Are Energy-Rich Compounds   A NAD + and FAD Are Electron Carriers  B The Nernst Equation Describes  Oxidation-Reduction Reactions  C Spontaneity Can Be Determined by Measuring  Reduction Potential Differences   A Labeled Metabolites Can Be Traced  B Studying Metabolic Pathways Often Involves  Perturbing the System  C Systems Biology Has Entered the Study  of Metabolism   The fungus Neurospora crassa (a common bread mold), with a modest 10,000 genes  and a relatively simple life cycle, is famous for its role in metabolic studies.  Experiments with mutant Neurospora led to the “one gene-one enzyme” hypothesis  in 1941. Species such as Neurospora make good model organisms because the  central metabolic pathways of all organisms are fundamentally the same.    Understanding the chemical compositions and three-dimensional structures of  biological molecules is not sufficient to understand how they are assembled into  organisms or how they function to sustain life. We must therefore examine the  reactions in which biological molecules are built and broken down. We must  also consider how free energy is consumed in building cellular materials and  carrying out cellular work and how free energy is generated from organic or  other sources. Metabolism, the overall process through which living systems  acquire and use free energy to carry out their various functions, is traditionally  divided into two parts:   1. Catabolism, or degradation, in which nutrients and cell constituents are  broken down to salvage their components and/or to generate energy.   2. Anabolism, or biosynthesis, in which biomolecules are synthesized  from simpler components.   In general, catabolic reactions carry out the exergonic oxidation of nutri¬  ent molecules. The free energy thereby released is used to drive such ender¬  gonic processes as anabolic reactions, the performance of mechanical work,  and the active transport of molecules against concentration gradients.  Exergonic and endergonic processes are often coupled through the intermediate  synthesis of a “high-energy” compound such as ATP. This simple principle un¬  derlies many of the chemical reactions presented in the following chapters. In  this chapter, we introduce the general features of metabolic reactions and the  roles of ATP and other compounds as energy carriers. Because many meta¬  bolic reactions are also oxidation-reduction reactions, we review the thermo¬  dynamics of these processes. Finally, we examine some approaches to studying  metabolic reactions.   • Different organisms use different strategies for capturing free energy from their  environment and can be classified by their requirement for oxygen.   • Mammalian nutrition involves the intake of macronutrients (proteins, carbohydrates,  and lipids) and micronutrients (vitamins and minerals).   • A metabolic pathway is a series of enzyme-catalyzed reactions, often located in a  specific part of the cell.   • The flux of material through a metabolic pathway varies with the activities of the  enzymes that catalyze irreversible reactions.   • These flux-controlling enzymes are regulated by allosteric mechanisms, covalent  modification, substrate cycling, and changes in gene expression.   A bewildering array of chemical reactions occur in any living cell. Yet the prin¬  ciples that govern metabolism are the same in all organisms, a result of their  common evolutionary origin and the constraints of the laws of thermodynam¬  ics. In fact, many of the specific reactions of metabolism are common to all  organisms, with variations due primarily to differences in the source of the  free energy that supports them.   Nutrition, the intake and utilization of food, affects health, development, and  performance. Food supplies the energy that powers life processes and provides  the raw materials to build and repair body tissues. The nutritional require¬  ments of an organism reflect its source of metabolic energy. For example, some  prokaryotes are autotrophs (Greek: autos , self + trophos , feeder), which can  synthesize all their cellular constituents from simple molecules such as H 2 0,  C0 2 , NH 3 , and H 2 S. There are two possible free energy sources for this  process. Chemolithotrophs (Greek: lithos, stone) obtain their energy through  the oxidation of inorganic compounds such as NH 3 , H 2 S, or even Fe 2+ :   2 NH 3 + 4 0 2 -^2 HN0 3 + 2 H 2 0  H 2 S + 2 0 2 H 2 S0 4   4 FeC0 3 + 0 2 + 6 H 2 0 ->4 Fe(OH) 3 + 4 C0 2   Photoautotrophs do so via photosynthesis, a process in which light energy  powers the transfer of electrons from inorganic donors to C0 2 to produce car¬  bohydrates, (CH 2 0)„, which are later oxidized to release free energy.  Heterotrophs (Greek: hetero, other) obtain free energy through the oxidation  of organic compounds (carbohydrates, lipids, and proteins) and hence ulti¬  mately depend on autotrophs and/or phototrophs for those substances.   Organisms can be further classified by the identity of the oxidizing agent  for nutrient breakdown. Obligate aerobes (which include animals) must use  0 2 , whereas anaerobes employ oxidizing agents such as sulfate or nitrate.  Facultative anaerobes, such as E. coli, can grow in either the presence or the  absence of 0 2 . Obligate anaerobes, in contrast, are poisoned by the presence  of 0 2 . Their metabolisms are thought to resemble those of the earliest life-  forms, which arose over 3.5 billion years ago when the earths atmosphere lacked  0 2 . Most of our discussion of metabolism will focus on aerobic processes.   Animals are obligate aerobic heterotrophs, whose nutrition depends on a  balanced intake of the macronutrients proteins, carbohydrates, and lipids.  These are broken down by the digestive system to their component amino  acids, monosaccharides, fatty acids, and glycerol, the major nutrients involved  in cellular metabolism, which are then transported by the circulatory system  to the tissues. The metabolic utilization of the latter substances also requires  the intake of 0 2 and water, as well as micronutrients composed of vitamins  and minerals.    Vitamins are organic molecules that an animal is unable to synthesize and  must therefore obtain from its diet. Vitamins can be divided into two groups,  water-soluble vitamins and fat-soluble vitamins. Table 14-1 lists many com¬  mon vitamins and the types of reactions or processes in which they partici¬  pate (we will consider the structures of these substances and their reaction  mechanisms in the appropriate sections of the text).   Table 14-2 lists the essential minerals and trace elements necessary for  metabolism. They participate in metabolic processes in many ways. Mg 2+ , for  example, is involved in nearly all reactions that involve ATP and other nu¬  cleotides, including the synthesis of DNA, RNA, and proteins. Zn 2+ is a co¬  factor in a variety of enzymes, including carbonic anhydrase (Section 11-3C).  Ca 2+ , in addition to being the major mineral component of bones and teeth,  is a vital participant in signal transduction processes (Section 13-4).   Most Water-Soluble Vitamins Are Converted to Coenzymes. Many coen¬  zymes (Section 11-1C) were discovered as growth factors for microorganisms  or as substances that cure nutritional deficiency diseases in humans and/or an¬  imals. For example, the NAD + component nicotinamide, or its carboxylic  acid analog nicotinic acid (niacin; Fig. 14-1), relieves the ultimately fatal    FIG. 1 4-1 The structures of nicotinamide  and nicotinic acid. These vitamins form the  redox-active components of the nicotinamide  coenzymes NAD + and NADP + (compare with  Fig. 11-4).   dietary deficiency disease in humans known as pellagra. The symptoms of Section 1 Overview of Metabolism   The water-soluble vitamins in the human diet are all coenzyme precur¬  sors. In contrast, the fat-soluble vitamins, with the exception of vitamin K  (Section 9-IF), are not components of coenzymes, although they are also re¬  quired in small amounts in the diets of many higher animals. The distant an¬  cestors of humans probably had the ability to synthesize the various vitamins,  as do many modern plants and microorganisms. Yet since vitamins are nor¬  mally available in the diets of animals, which all eat other organisms, or are  synthesized by the bacteria that normally inhabit their digestive systems, it  seems likely that the superfluous cellular machinery to synthesize them was  lost through evolution. For example, vitamin C (ascorbic acid) is required in  the diets of only humans, apes, and guinea pigs (Section 6-1C and Box 6-2)  because, in what is apparently a recent evolutionary loss, they lack a key en¬  zyme for ascorbic acid biosynthesis.   Metabolic pathways are series of connected enzymatic reactions that produce specific  products. Their reactants, intermediates, and products are referred to as metabo¬  lites. There are around 4000 known metabolic reactions, each catalyzed by a  distinct enzyme. The types of enzymes and metabolites in a given cell vary with  the identity of the organism, the cell type, its nutritional status, and its devel¬  opmental stage. Many metabolic pathways are branched and interconnected, so  delineating a pathway from a network of thousands of reactions is somewhat  arbitrary and is driven by tradition as much as by chemical logic.   In general, degradative and biosynthetic pathways are related as follows  (Fig. 14 -2): In degradative pathways, the major nutrients, referred to as com¬  plex metabolites, are exergonically broken down into simpler products. The  free energy released in the degradative process is conserved by the synthesis of  ATP from ADP + P z or by the reduction of the coenzyme NADP + (Fig. 11-4)  to NADPH. ATP and NADPH are the major free energy sources for biosyn¬  thetic reactions. We will consider the thermodynamic properties of ATP and  NADPH later in this chapter.   A striking characteristic of degradative metabolism is that the pathways for  the catabolism of a large number of diverse substances (carbohydrates, lipids, and  proteins) converge on a few common intermediates , in many cases, a two-carbon   FIG. 1 4-2 Roles of ATP and NADP + in metabolism. ATP and NADPH, generated  through the degradation of complex metabolites such as carbohydrates, lipids, and proteins,  are sources of free energy for biosynthetic and other reactions.   FIG. 1 4-3 Overview of catabolism. Complex metabolites such as carbohydrates,  proteins, and lipids are degraded first to their monomeric units, chiefly glucose, amino acids,  fatty acids, and glycerol, and then to the common intermediate, acetyl-CoA. The acetyl  group is oxidized to C0 2 via the citric acid cycle with concomitant reduction of NAD + and  FAD to NADH and FADH 2 . Reoxidation of NADH and FADH 2 by 0 2 during electron  transport and oxidative phosphorylation yields H 2 0 and ATR    acetyl unit linked to coenzyme A to form acetyl-coenzyme A (acetyl-CoA;  Section 14-2D). These intermediates are then further metabolized in a central  oxidative pathway. Figure 14-3 outlines the breakdown of various foodstuffs  to their monomeric units and then to acetyl-CoA. This is followed by the ox¬  idation of the acetyl carbons to C0 2 by the citric acid cycle (Chapter 17).  When one substance is oxidized (loses electrons), another must be reduced  (gain electrons; Box 14-1). The citric acid cycle thus produces the reduced  coenzymes NADH and FADH 2 (Section 14-3A), which then pass their elec¬  trons to 0 2 to produce H 2 0 in the processes of electron transport and  oxidative phosphorylation (Chapter 18).   Biosynthetic pathways carry out the opposite process. Relatively few  metabolites serve as starting materials for a host of varied products. In the next  several chapters, we discuss many catabolic and anabolic pathways in detail.   Enzymes Catalyze the Reactions of Metabolic Pathways. With a few ex¬  ceptions, the interconversions of metabolites in degradative and biosynthetic  pathways are catalyzed by enzymes. In the absence of enzymes, the reactions  would occur far too slowly to support life. In addition, the specificity of en¬  zymes guarantees the efficiency of metabolic reactions by preventing the for-   The carbon atoms in biological molecules can assume different  oxidation states depending on the atom to which they are bonded.  For example, a carbon atom bonded to less electronegative hydro¬  gen atoms is more reduced than a carbon atom bonded to highly  electronegative oxygen atoms.   The simplest way to determine the oxidation number (and  hence the oxidation state) of a particular carbon atom is to exam¬  ine each of its bonds and assign the electrons to the more elec¬  tronegative atom. In a C—0 bond, both electrons “belong” to 0; in  a C—H bond, both electrons “belong” to C; and in a C—C bond,  each carbon “owns” one electron. An atom’s oxidation number is  the number of valence electrons on the free atom (4 for carbon)  minus the number of its lone pair and assigned electrons. For ex¬  ample, the oxidation number of carbon in C0 2 is 4 - (0 + 0) =  +4, and the oxidation number of carbon in CH 4 is 4 - (0 + 8) =  -4. Keep in mind, however, that oxidation numbers are only  accounting devices; actual atomic charges are much closer to  neutrality.   The following compounds are listed according to the oxidation  state of the highlighted carbon atom. In general, the more oxidized  compounds have fewer electrons per C atom and are richer in  oxygen, and the more reduced compounds have more electrons  per C atom and are richer in hydrogen. But note that not all  reduction events (gain of electrons) or oxidation events (loss of  electrons) are associated with bonding to oxygen. For example,  when an alkane is converted to an alkene, the formation of a  carbon-carbon double bond involves the loss of electrons and  therefore is an oxidation reaction although no oxygen is involved.  Knowing the oxidation number of a carbon atom is seldom  required. However, it is useful to be able to determine whether the  oxidation state of a given atom increases or decreases during a  chemical reaction.   mation of useless or toxic by-products. Most importantly, enzymes provide a  mechanism for coupling an endergonic chemical reaction (which would not  occur on its own) with an energetically favorable reaction, as discussed below.   We will see examples of reactions catalyzed by all six classes of enzymes  introduced in Section 11-1 A. These reactions fall into four major types: oxi¬  dations and reductions (catalyzed by oxidoreductases), group-transfer reac¬  tions (catalyzed by transferases and hydrolases), eliminations, isomerizations,  and rearrangements (catalyzed by isomerases and mutases), and reactions  that make or break carbon-carbon bonds (catalyzed by hydrolases, lyases,  and ligases).   Metabolic Pathways Occur in Specific Cellular Locations. The compart-  mentation of the eukaryotic cytoplasm allows different metabolic pathways to  operate in different locations. For example, electron transport and oxidative  phosphorylation occur in the mitochondria, whereas glycolysis (a carbohy¬  drate degradation pathway) and fatty acid biosynthesis occur in the cytosol.   Posttranslational processing of  membrane & secretory proteins;  formation of plasma membrane  and secretory vesicles   Glycolysis, pentose phosphate  pathway, fatty acid biosynthesis,  many reactions of gluconeogenesis   Oxidative reactions catalyzed by  amino acid oxidases and catalase;  glyoxylate cycle reactions in plants   Citric acid cycle, electron transport  and oxidative phosphorylation, fatty  acid oxidation, amino acid breakdown    FIG. 1 4-4 Metabolic functions of eukaryotic organelles. Degradative and biosyn¬  thetic processes may occur in specialized compartments in the cell, or may involve several  compartments.    Without looking at the figure, summarize the major function of each cellular  compartment. Identify which compartments carry out degradative versus  synthetic processes.    Figure 14-4 shows the major metabolic functions of eukaryotic organelles.  Metabolic processes in prokaryotes, which lack organelles, may be localized to  particular areas of the cytosol.   The synthesis of metabolites in specific membrane-bounded compartments in  eukaryotic cells requires mechanisms to transport these substances between compart¬  ments. Accordingly, transport proteins (Chapter 10) are essential components  of many metabolic processes. For example, a transport protein is required to  move ATP, which is generated in the mitochondria, to the cytosol, where most  of it is consumed (Section 18-IB).   In multicellular organisms, compartmentation is carried a step further to  the level of tissues and organs. The mammalian liver, for example, is largely  responsible for the synthesis of glucose from noncarbohydrate precursors (glu¬  coneogenesis; Section 16-4) so as to maintain a relatively constant level of  glucose in the circulation, whereas adipose tissue is specialized for storage of  triacylglycerols. The interdependence of the metabolic functions of the vari¬  ous organs is the subject of Chapter 22.   An intriguing manifestation of specialization of tissues and subcellular  compartments is the existence of isozymes, enzymes that catalyze the same re¬  action but are encoded by different genes and have different kinetic or regu¬  latory properties. For example, we have seen that mammals have three  isozymes of glycogen phosphorylase, those expressed in muscle, brain, and liver  (Section 12-3B). Similarly, vertebrates possess two homologs of the enzyme  lactate dehydrogenase: the M type, which predominates in tissues subject to  anaerobic conditions such as skeletal muscle and liver, and the H type, which  predominates in aerobic tissues such as heart muscle. Lactate dehydrogenase    catalyzes the interconversion of pyruvate, a product of glycolysis, and lactate  (Section 15-3A). The M-type isozyme appears mainly to function in the re¬  duction by NADH of pyruvate to lactate, whereas the H-type enzyme appears  to be better adapted to catalyze the reverse reaction.   r The existence of isozymes allows for the testing of various illnesses. For ex¬  ample, heart attacks cause the death of heart muscle cells, which consequently  rupture and release H-type LDH into the blood. A blood test indicating the  presence of H-type LDH is therefore diagnostic of a heart attack.   D Thermodynamics Dictates the Direction and Regulatory Capacity  of Metabolic Pathways   Knowing the location of a metabolic pathway and enumerating its substrates  and products does not necessarily reveal how that pathway functions as part  of a larger network of interrelated biochemical processes. It is also necessary  to appreciate how fast end product can be generated by the pathway as well  as how pathway activity is regulated as the cells needs change. Conclusions  about a pathways output and its potential for regulation can be gleaned from  information about the thermodynamics of each enzyme-catalyzed step.   Recall from Section 1-3D that the free energy change AG of a biochem¬  ical process, such as the reaction   is related to the standard free energy change (A G of ) and the concentrations of  the reactants and products (Eq. 1-15):   Thus, the value of AG°' can be calculated from the equilibrium constant and  vice versa (see Sample Calculation 14-1).   When the reactants are present at values close to their equilibrium values,  [C] eq [D] eq /[A] eq [B] eq ~ fQ q , and AG~ 0. This is the case for many meta¬  bolic reactions, which are said to be near-equilibrium reactions. Because their  AG values are close to zero, they can be relatively easily reversed by changing  the ratio of products to reactants. When the reactants are in excess of their  equilibrium concentrations, the net reaction proceeds in the forward direction  until the excess reactants have been converted to products and equilibrium is  attained. Conversely, when products are in excess, the net reaction proceeds  in the reverse direction so as to convert products to reactants until the    Calculate the equilibrium constant for the hydrolysis of glucose-1-phosphate at  37°C, using the information in Table 14-3 (see p.448).   £ _ ^-(-20,900 J * mol -1 )/(8.3145 J * K _1 • mol - 9(310 K)   near-equilibrium reactions tend to act quickly to restore equilibrium concentra¬  tions y and the net rates of such reactions are effectively controlled by the relative  concentrations of substrates and products.   Other metabolic reactions function far from equilibrium; that is, they are  irreversible. This is because an enzyme catalyzing such a reaction has insufficient  catalytic activity (the rate of the reaction it catalyzes is too slow) to allow the  reaction to come to equilibrium under physiological conditions. Reactants there¬  fore accumulate in large excess of their equilibrium amounts, making AG « 0.  Changes in substrate concentrations therefore have relatively little effect on the  rate of an irreversible reaction; the enzyme is essentially saturated. Only changes  in the activity of the enzyme, through allosteric interactions, for example, can  significantly alter the rate. The enzyme is therefore analogous to a dam on a  river: It controls the flow of substrate through the reaction by varying its activityy  much as a dam controls the flow of a river by varying the opening of its floodgates.   Understanding the flux (rate of flow) of metabolites through a metabolic  pathway requires knowledge of which reactions are functioning near equilibrium  and which are far from it. Most enzymes in a metabolic pathway operate near  equilibrium and therefore have net rates that vary with their substrate concen¬  trations. However, certain enzymes that operate far from equilibrium are strate¬  gically located in metabolic pathways. This has several important implications:   1. Metabolic pathways are irreversible. A highly exergonic reaction (one  with AG « 0) is irreversible; that is, it goes to completion. If such a  reaction is part of a multistep pathway, it confers directionality on the  pathway; that is, it makes the entire pathway irreversible.   2. Every metabolic pathway has a first committed step. Although most  reactions in a metabolic pathway function close to equilibrium, there  is generally an irreversible (exergonic) reaction early in the pathway that  “commits” its product to continue down the pathway (likewise, water  that has gone over a dam cannot spontaneously return).   3. Catabolic and anabolic pathways differ. If a metabolite is converted  to another metabolite by an exergonic process, free energy must be sup¬  plied to convert the second metabolite back to the first. This energet¬  ically “uphill” process requires a different pathway for at least one of  the reaction steps.   The existence of independent interconversion routes, as we will see, is  an important property of metabolic pathways because it allows inde¬  pendent control of the two processes. If metabolite 2 is required by the  cell, it is necessary to “turn off” the pathway from 2 to 1 while “turn¬  ing on” the pathway from 1 to 2. Such independent control would be  impossible without different pathways.   Living organisms are thermodynamically open systems that tend to maintain a  steady state rather than reaching equilibrium (Section 1-3E). This is strikingly  demonstrated by the observation that, over a 40-year time span, a normal hu¬  man adult consumes literally tons of nutrients and imbibes over 20,000 L of  water but does so without major weight change. The flux of intermediates  through a metabolic pathway in a steady state is more or less constant; that is, the  rates of synthesis and breakdown of each pathway intermediate maintain it at a  constant concentration. A steady state far from equilibrium is thermodynamically  efficient, because only a nonequilibrium process (AG A 0) can perform useful  work. Indeed, living systems that have reached equilibrium are dead.   Since a metabolic pathway is a series of enzyme-catalyzed reactions, it is Section 1 Overview of Metabolism   easiest to describe the flux of metabolites through the pathway by considering  its reaction steps individually. The flux of metabolites, /, through each reaction  step is the rate of the forward reaction, zy, less that of the reverse reaction, v r \   At equilibrium, by definition, there is no flux (/ = 0), although zy and v r may  be quite large. In reactions that are far from equilibrium, zy » zy, the flux  is essentially equal to the rate of the forward reaction (/ — zy).   For the pathway as a whole, flux is set by the rate-determining step of the  pathway. By definition, this step is the pathways slowest step, which is often the  first committed step of the pathway. In some pathways, flux control is distrib¬  uted over several enzymes, all of which help determine the overall rate of flow  of metabolites through the pathway. Because a rate-determining step is slow rel¬  ative to other steps in the pathway, its product is removed by succeeding steps  in the pathway before it can equilibrate with reactant. Thus, the rate-determining  step functions far from equilibrium and has a large negative free energy change.   In an analogous manner, a dam creates a difference in water levels between its  upstream and downstream sides, and a large negative free energy change results  from the hydrostatic pressure difference. The dam can release water to generate  electricity, varying the water flow according to the need for electrical power.   Reactions that function near equilibrium respond rapidly to changes in  substrate concentration. For example, upon a sudden increase in the concen¬  tration of a reactant for a near-equilibrium reaction, the enzyme catalyzing it  would increase the net reaction rate so as to rapidly achieve the new equilib¬  rium level. Thus, a series of near-equilibrium reactions downstream from the  rate-determining step all have the same flux. Likewise, the flux of water in a  river is the same at all points downstream from a dam.   In practice, it is often possible to identify flux control points for a pathway  by identifying reactions that have large negative free energy changes. The rela¬  tive insensitivity of the rates of these nonequilibrium reactions to variations in  the concentrations of their substrates permits the establishment of a steady state  flux of metabolites through the pathway. Of course, flux through a pathway  must vary in response to the organisms requirements so as to reach a new steady  state. Altering the rates of the rate-determining steps can alter the flux of mate¬  rial through the entire pathway, often by an order of magnitude or more.   Cells use several mechanisms to control flux through the rate-determining  steps of metabolic pathways:   1. Allosteric control. Many enzymes are allosterically regulated (Section  12-3A) by effectors that are often substrates, products, or coenzymes  of the pathway but not necessarily of the enzyme in question. For ex¬  ample, in negative feedback regulation, the product of a pathway in¬  hibits an earlier step in the pathway:   Thus, as we have seen, CTP, a product of pyrimidine biosynthesis, in¬  hibits ATCase, which catalyzes the rate-determining step in the path¬  way (Fig. 12-11).   2. Covalent modification. Many enzymes that control pathway fluxes have  specific sites that may be enzymatically phosphorylated and dephospho-  rylated (Section 12-3B) or covalently modified in some other way. Such  enzymatic modification processes, which are themselves subject to con¬  trol by external signals such as hormones (Section 13-1), greatly alter the  activities of the modified enzymes. The signaling methods involved in  such flux control mechanisms are discussed in Chapter 13.   • Use the words obligate, facultative,  aerobic, anaerobic, autotroph, and  heterotroph to describe the metabolism  of a human, oak tree, E. coli, and  Methanococcus jannaschii (an organism  that lives in deepwater anoxic sediments).   • List the categories of macronutrients and  micronutrients required for mammalian  metabolism and provide examples of each.   • Explain the roles of ATP and NADPH in  catabolic and anabolic reactions.   • Give some reasons why enzymes are  essential for the operation of metabolic  pathways.   • Explain the metabolic significance of  reactions that function near equilibrium and  reactions that function far from equilibrium.   • Discuss the mechanisms by which the flux  through a metabolic pathway can be  controlled. Which mechanisms can rapidly  alter flux?    3. Substrate cycles. If zyand v r represent the rates of two opposing non¬  equilibrium reactions that are catalyzed by different enzymes, zy and  v r may be independently varied.    For example, flux (zy — v r ) can be increased not just by accelerating  the forward reaction but by slowing the reverse reaction. The flux  through such a substrate cycle, as we will see in Section 15-4, is more  sensitive to the concentrations of allosteric effectors than is the flux  through a single unopposed nonequilibrium reaction.   4. Genetic control. Enzyme concentrations, and hence enzyme activities, may  be altered by protein synthesis in response to metabolic needs. With the  sequencing of entire genomes, the genetic response of an organism to en¬  vironmental changes has become a major field of study. Transcriptomics  (the study of the entire collection of RNA transcribed by a cell) and pro-  teomics (the study of the complete set of proteins synthesized by a cell in  response to changing conditions) are part of the emerging field of systems  biology (Section 14-4C). Mechanisms of genetic control of enzyme con¬  centrations are a major concern of Part V of this text.   Mechanisms 1 to 3 can respond rapidly (within seconds or minutes) to exter¬  nal stimuli and are therefore classified as “short-term” control mechanisms.  Mechanism 4 responds more slowly to changing conditions (within hours or  days in higher organisms) and is therefore regarded as a “long-term” control  mechanism.   Control of most metabolic pathways involves several nonequilibrium steps.  Hence, the flux of material through a pathway that supplies intermediates for  use by an organism may depend on multiple effectors whose relative impor¬  tance reflects the overall metabolic demands of the organism at a given time.  Thus, a metabolic pathway is part of a supply-demand process.    • Organisms capture the free energy released on degradation of nutrients as  “high-energy" compounds such as ATP, whose subsequent breakdown is used to  power otherwise endergonic reactions.   • The “high energy" of ATP is related to the large negative free energy change for  hydrolysis of its phosphoanhydride bonds.   • ATP hydrolysis can be coupled to an endergonic reaction such that the net  reaction is favorable.   • Phosphoryl groups are transferred from compounds with high phosphoryl group-  transfer potentials to those with low phosphoryl group-transfer potentials.    C 6 H 12 0 6 + 6 0 2 —> 6 C0 2 + 6 H 2 0   C 16 H 32 0 2 + 23 0 2 —> 16 C0 2 +16 H 2 0   is even more exergonic (AG°' = —9781 kj • mol x ). Oxidative metabolism  proceeds in a stepwise fashion, so the released free energy can be recovered in  a manageable form at each exergonic step of the overall process. These 'packets”  of energy are conserved by the synthesis of a few types of “high-energy” interme¬  diates whose subsequent exergonic breakdown drives endergonic processes. These  intermediates therefore form a sort of free energy “currency” through which  free energy-producing reactions such as glucose oxidation or fatty acid oxi¬  dation “pay for” the free energy-consuming processes in biological systems  (Box 14-2).   The cell uses several forms of energy currency, including phosphorylated  compounds such as the nucleotide ATP (the cells primary energy currency),  compounds that contain thioester bonds, and reduced coenzymes such as  NADH. Each of these represents a source of free energy that the cell can use  in various ways, including the synthesis of ATP. We will first examine ATP  and then discuss the properties of other forms of energy currency.   Among the many scientists who fled Europe  for the United States in the 1930s was Fritz  i mi, if Lipmann, a German-born physician-turned-   ■ twentieth century, scientists were primarily  interested in the structures and compositions  of biological molecules, and not much was  known about their biosynthesis. Lipmann’s  contribution to this field centers on his understanding of  “energy-rich” phosphates and other “active” compounds.   Lipmann began his research career by studying creatine  phosphate, a compound that could provide energy for muscle  contraction. He, like many of his contemporaries, was puzzled by  the absence of an obvious link between this phosphorylated  compound and the known metabolic activity of a contracting  muscle, namely, converting glucose to lactate. One link was  discovered by Otto Warburg (Box 15-1), who showed that one  of the steps of glycolysis was accompanied by the incorporation  of inorganic phosphate. The resulting acyl phosphate  (1,3-bisphosphoglycerate) could then react with ADP to form ATP.   Lipmann wondered whether other phosphorylated compounds  might behave in a similar manner. Because the purification of  such labile (prone to degradation) compounds from whole cells  was impractical, Lipmann synthesized them himself. He was able  to show that cell extracts used synthetic acetyl phosphate to  produce ATP. Lipmann went on to propose that cells contain two  classes of phosphorylated compounds, which he termed “energy-  poor” and “energy-rich,” by which he meant compounds with low  and high negative free energies of hydrolysis (the “squiggle,” ~,  which is still used, was his symbol for an “energy-rich” bond).  Lipmann described a sort of “phosphate current” in which   The ATP, in turn, can power mechanical work such as muscle  contraction or drive biosynthetic reactions.   Until this point (1941), biochemists studying biosynthetic  processes were largely limited to working with whole animals or  relatively intact tissue slices. Lipmann’s insight regarding the role  of ATP freed researchers from their cumbersome and poorly  reproducible experimental systems. Biochemists could simply add  ATP to their cell-free preparations to reconstitute the biosynthetic  process.   Lipmann was intrigued by the discovery that a two-carbon  group, an “active acetate,” served as a precursor for the synthesis  of fatty acids and steroids. Was acetyl phosphate also the “active  acetate”? This proved not to be the case, although Lipmann was  able to show that the addition of a two-carbon unit to another  molecule (acetylation) required acetate, ATP, and a heat-stable  factor present in pigeon liver extracts. He isolated and determined  the structure of this factor, which he named coenzyme A. For this  seminal discovery, Lipmann was awarded the 1953 Nobel Prize in  Physiology or Medicine.   Kleinkauf, H., von Dohren, H., and Jaenicke, L (Eds.), The Roots of Modern  Biochemistry. Fritz Lipmann’s Squiggle and Its Consequences, Walter de Gruyter  (1988).   FIG. 1 4-5 The structure of ATP indicating  its relationship to ADP, AMP, and adenosine.   The phosphoryl groups, starting from AMP, are  referred to as the a-, p-, and 7 -phosphates.    TABLE 14-3 Standard Free Energies  of Phosphate Hydrolysis  of Some Compounds of  Biological Interest    Source: Mostly from Jencks, W.P., in Fasman, G.D.  (Ed.), Handbook of Biochemistry and Molecular  Biology (3rd ed.), Physical and Chemical Data,   The “high-energy” intermediate adenosine triphosphate (ATP; Fig. 14-5) oc¬  curs in all known life-forms. ATP consists of an adenosine moiety (adenine +  ribose) to which three phosphoryl (—PO 3 ) groups are sequentially linked via  a phosphoester bond followed by two phosphoanhydride bonds.   The biological importance of ATP rests in the large free energy change  that accompanies cleavage of its phosphoanhydride bonds. This occurs when  either a phosphoryl group is transferred to another compound, leaving ADP,  or a nucleotidyl (AMP) group is transferred, leaving pyrophosphate (P 2 0 2 ;  PP/). When the acceptor is water, the process is known as hydrolysis:   Most biological group-transfer reactions involve acceptors other than water.  However, knowing the free energy of hydrolysis of various phosphoryl com¬  pounds allows us to calculate the free energy of transfer of phosphoryl groups  to other acceptors by determining the difference in free energy of hydrolysis  of the phosphoryl donor and acceptor.   The A G of values for hydrolysis of several phosphorylated compounds of  biochemical importance are tabulated in Table 14-3. The negatives of these  values are often referred to as phosphoryl group-transfer potentials; they are  a measure of the tendency of phosphorylated compounds to transfer their  phosphoryl groups to water. Note that ATP has an intermediate phosphate  group-transfer potential. Under standard conditions, the compounds above  ATP in Table 14-3 can spontaneously transfer a phosphoryl group to ADP to  form ATP, which can, in turn, spontaneously transfer a phosphoryl group to  the appropriate groups to form the compounds listed below it. Note that a  favorable free energy change for a reaction does not indicate how quickly the  reaction occurs. Despite their high group-transfer potentials, ATP and related  phosphoryl compounds are kinetically stable and do not react at a signifi¬  cant rate unless acted upon by an appropriate enzyme.   What Is the Nature of the “Energy” in “High-Energy” Compounds? Bonds  whose hydrolysis proceeds with large negative values of A G of (customarily  more than —25 kj • mol l ) are often referred to as “high-energy” bonds or  “energy-rich” bonds and are frequently symbolized by the squiggle . Thus,  ATP can be represented as AR—P~P~P, where A, R, and P symbolize adenyl,  ribosyl, and phosphoryl groups, respectively. Yet the phosphoester bond join¬  ing the adenosyl group of ATP to its a-phosphoryl group appears to be not  greatly different in electronic character from the “high-energy” bonds bridg¬  ing its a- and (3- and its (3- and y-phosphoryl groups. In fact, none of these  bonds has any unusual properties, so the term “high-energy” bond is some¬  what of a misnomer (in any case, it should not be confused with the term  “bond energy,” which is defined as the energy required to break, not hydrolyze,  a covalent bond). Why, then, are the phosphoryl group-transfer reactions of  ATP so exergonic? Several factors appear to be responsible for the “high-  energy” character of phosphoanhydride bonds such as those in ATP (Fig. 14-6):   1 . The resonance stabilization of a phosphoanhydride bond is less than  that of its hydrolysis products. This is because a phosphoanhydride’s  two strongly electron-withdrawing groups must compete for the lone  pairs of electrons of its bridging oxygen atom, whereas this competi¬  tion is absent in the hydrolysis products. In other words, the electronic  requirements of the phosphoryl groups are less satisfied in a phospho¬  anhydride than in its hydrolysis products.   2 . Of perhaps greater importance is the destabilizing effect of the electro¬  static repulsions between the charged groups of a phosphoanhydride  compared to those of its hydrolysis products. In the physiological pH    range, ATP has three to four negative charges whose mutual electro¬  static repulsions are partially relieved by ATP hydrolysis.   3. Another destabilizing influence, which is difficult to assess, is the  smaller solvation energy of a phosphoanhydride compared to that of  its hydrolysis products. Some estimates suggest that this factor provides  the dominant thermodynamic driving force for the hydrolysis of phos-  phoanhydrides.   Of course, the free energy change for any reaction, including phosphoryl  group transfer from a “high-energy” compound, depends in part on the con¬  centrations of the reactants and products (Eq. 14-1). Furthermore, because  ATP and its hydrolysis products are ions, AG also depends on pH and ionic  strength (Box 14-3).   The hydrolysis of a “high-energy” compound, while releasing considerable free  energy, is not in itself a useful reaction. However, the exergonic reactions of  “high-energy” compounds can be coupled to endergonic processes to drive  them to completion. The thermodynamic explanation for the coupling of an  exergonic and an endergonic process is based on the additivity of free energy.  Consider the following two-step reaction pathway:    FIG. 1 4-6 Resonance and electrostatic  stabilization in a phosphoanhydride and its  hydrolytic products. The competing resonances  (curved arrows from the central 0) and  charge-charge repulsions (zigzag lines ) between  phosphoryl groups decrease the stability of a  phosphoanhydride relative to its hydrolysis  products.    If AG X > 0, Reaction 1 will not occur spontaneously. However, if AG 2 is suf¬  ficiently exergonic so that A G x + AG 2 < 0, then although the equilibrium  concentration of D in Reaction 1 will be relatively small, it will be larger than  that in Reaction 2. As Reaction 2 converts D to products, Reaction 1 will op¬  erate in the forward direction to replenish the equilibrium concentration of  D. The highly exergonic Reaction 2 therefore “drives” or “pulls” the ender¬  gonic Reaction 1, and the two reactions are said to be coupled through their  common intermediate, D. That these coupled reactions proceed spontaneously  can also be seen by summing Reactions 1 and 2 to yield the overall reaction    (1+2) A + B + E C + F + G AG 3    The standard conditions reflected in AG°' values never occur in  living organisms. Furthermore, other compounds that are present  at high concentrations and that can potentially interact with the  substrates and products of a metabolic reaction may dramatically  affect AG values. For example, Mg 2+ ions in cells partially neutral¬  ize the negative charges on the phosphate groups in ATP and its  hydrolysis products, thereby diminishing the electrostatic repulsions  that make ATP hydrolysis so exergonic. Similarly, changes in pH  alter the ionic character of phosphorylated compounds and there¬  fore alter their free energies.   In a given cell, the concentrations of many ions, coenzymes, and  metabolites vary with both location and time, often by several or¬  ders of magnitude. Intracellular ATP concentrations are maintained  within a relatively narrow range, usually 2-10 mM, but the concen¬  trations of ADP and P/ are more variable. Consider a typical cell  with [ATP] = 3.0 mM, [ADP] = 0.8 mM, and [Pj = 4.0 mM. Using  Eq. 14-1, the actual free energy of ATP hydrolysis at 37°C is  calculated as follows:   = -30.5 kJ • mol -1 + (8.3145 J • K _1  ((0.Q X 10~ 3 M)(4.0 X 10~ 3 M)  A (3.0 X 10~ 3 M)    This value is even greater than the standard free energy of ATP  hydrolysis. However, because of the difficulty in accurately mea¬  suring the concentrations of particular chemical species in a cell  or organelle, the AG’s for most in vivo reactions are little more  than estimates. For the sake of consistency, we will, for the most  part, use AG°' values in this textbook.   Chapter 14 Introduction to Metabolism where A G 3 = AG^ + AG 2 < 0. As long as the overall pathway is exergonic, it   To illustrate this concept, let us consider two examples of phosphoryl  group-transfer reactions. The initial step in the metabolism of glucose is its  conversion to glucose-6-phosphate (Section 15-2A). Yet the direct reaction  of glucose and P z is thermodynamically unfavorable (AG 0 ' = +13.8  kj • mol T ; Fig. 14 -7 a). In cells, however, this reaction is coupled to the ex¬  ergonic cleavage of ATP (for ATP hydrolysis, AG°' = —30.5 kj • mol X ), so  the overall reaction is thermodynamically favorable (AG°' = +13.8 — 30.5  = —16.7 kj • mol X ). ATP can be similarly regenerated (AG°' = +30.5 kj  • mol X ) by coupling its synthesis from ADP and P z to the even more exergonic  cleavage of phosphoenolpyruvate (A G°' = —61.9 kj • mol x ; Fig. 14-7^  and Section 15-2J).   Note that the half-reactions shown in Fig. 14-7 do not actually occur as  written in an enzyme active site. Hexokinase, the enzyme that catalyzes the  formation of glucose- 6 -phosphate (Fig. 14-7^), does not catalyze ATP hydrol¬  ysis but instead catalyzes the transfer of a phosphoryl group from ATP directly  to glucose. Likewise, pyruvate kinase, the enzyme that catalyzes the reaction  shown in Fig. \A-7 b, does not add a free phosphoryl group to ADP but trans¬  fers a phosphoryl group from phosphoenolpyruvate to ADP to form ATP.   free energy of the phosphoanhydride bonds of “high-energy” compounds such  as ATP can be used to drive reactions even when the phosphoryl groups are  not transferred to another organic compound. For example, ATP hydrolysis  (i.e., phosphoryl group transfer directly to H 2 0) provides the free energy for  the operation of molecular chaperones (Section 6-5B), muscle contraction  (Section 7 - 2 B), and transmembrane active transport (Section 10-3). In these   half-reaction 1 P z + glucose v glucose-6-P + H 2 0 +13.8   phosphorylation of glucose to form glucose-6-phosphate and ADR  (b) The phosphorylation of ADP by phosphoenolpyruvate to form ATP  and pyruvate. Each reaction has been conceptually decomposed into a    direct phosphorylation step (half-reaction 1) and a step in which ATP  is hydrolyzed (half-reaction 2). Both half-reactions proceed in the  direction that makes the overall reaction exergonic (AG < 0).   ATP. The exergonic hydrolysis of ATP and release of ADP and Pi renders these  changes irreversible and thereby drives the processes forward. GTP hydrolysis func¬  tions similarly to drive some of the reactions of signal transduction (Section  13-3B) and protein synthesis (Section 27-4).   In the absence of an appropriate enzyme, phosphoanhydride bonds are  stable; that is, they hydrolyze quite slowly, despite the large amount of free  energy released by these reactions. This is because these hydrolysis reactions have  unusually high free energies of activation (AG*; Section 11-2). Consequently,   ATP hydrolysis is thermodynamically favored but kinetically disfavored. For  example, consider the reaction of glucose with ATP that yields glucose-  6 -phosphate (Fig. \4-7d). AG* for the nonenzymatic transfer of a phospho-  ryl group from ATP to glucose is greater than that for ATP hydrolysis, so  the hydrolysis reaction predominates (although neither reaction occurs at a  biologically significant rate). However, in the presence of the appropriate  enzyme, hexokinase (Section 15-2A), glucose-6-phosphate is formed far more  rapidly than ATP is hydrolyzed. This is because the catalytic influence of the  enzyme reduces the activation energy for phosphoryl group transfer from ATP  to glucose to less than the activation energy for ATP hydrolysis. This ex¬  ample underscores the point that even a thermodynamically favored reaction  (AG < 0) may not occur in a living system in the absence of a specific enzyme  that catalyzes the reaction (i.e., lowers AG* to increase the rate of product for¬  mation; Box 12-2).   Inorganic Pyrophosphatase Catalyzes Additional Phosphoanhydride  Bond Cleavage. Although many reactions involving ATP yield ADP and P z   (orthophosphate cleavage), others yield AMP and PP Z (pyrophosphate  cleavage). In these latter cases, the PP Z is rapidly hydrolyzed to 2 P z by inor¬  ganic pyrophosphatase (A G°' = —19.2 kj • mol l ) so that the pyrophosphate  cleavage of ATP ultimately consumes two “high-energy” phosphoanhydride bonds.   The attachment of amino acids to tRNA molecules for protein synthesis is an  example of this phenomenon (Fig. 14-8 and Section 27-2B). The two steps  of the reaction are readily reversible because the free energies of hydrolysis of the  bonds formed are comparable to that of ATP hydrolysis. The overall reaction is  driven to completion by the irreversible hydrolysis of PP Z . Nucleic acid biosyn¬  thesis from nucleoside triphosphates also releases PP Z (Sections 25-1 and 26-1).    FIG. 14-8 Pyrophosphate cleavage in the synthesis of an aminoacyl-tRNA. (1) In   the first reaction step, the amino acid is adenylylated by ATP. (2) In the second step, a  tRNA molecule displaces the AMP moiety to form an aminoacyl-tRNA. (3) The exergonic  hydrolysis of pyrophosphate (AG°' = -19.2 kJ • mol -1 ) drives the reaction forward.   Chapter 14 Introduction to Metabolism The standard free energy changes of these reactions are around 0, so the subse¬   quent hydrolysis of PP, is also essential for the synthesis of nucleic acids.    FIG. 1 4-9 Position of ATP relative to  “high-energy” and “low-energy” phosphate  compounds. Phosphoryl groups flow from the  “high-energy” donors, via the ATP-ADP system,  to “low-energy” acceptors.    “High-energy” compounds other than ATP are essential for energy metabolism,  in part because they help maintain a relatively constant level of cellular ATP. ATP  is continually being hydrolyzed and regenerated. Indeed, experimental evidence in¬  dicates that the metabolic half-life of an ATP molecule varies from seconds to  minutes depending on the cell type and its metabolic activity. For instance, brain  cells have only a few seconds supply of ATP (which partly accounts for the rapid  deterioration of brain tissue by oxygen deprivation). An average person at rest  consumes and regenerates ATP at a rate of ~3 mol (1.5 kg) per hour and as  much as an order of magnitude faster during strenuous activity.   Just as ATP drives endergonic reactions through the exergonic process of  phosphoryl group transfer and phosphoanhydride hydrolysis, ATP itself can be  regenerated by coupling its formation to a more highly exergonic metabolic process.  As Table 14-3 indicates, in the thermodynamic hierarchy of phosphoryl-transfer  agents, ATP occupies the middle rank. ATP can therefore be formed from  ADP by direct transfer of a phosphoryl group from a “high-energy” compound  (e.g., phosphoenolpyruvate; Fig. 14-7 b and Section 15-2J). Such a reaction is  referred to as a substrate-level phosphorylation. Other mechanisms generate  ATP indirectly, using the energy supplied by transmembrane proton concen¬  tration gradients. In oxidative metabolism, this process is called oxidative  phosphorylation (Section 18-3), whereas in photosynthesis, it is termed  photophosphorylation (Section 19-2D).   The flow of energy from “high-energy” phosphate compounds to ATP and  from ATP to “low-energy” phosphate compounds is diagrammed in Fig. 14-9.  These reactions are catalyzed by enzymes known as kinases, which transfer phos¬  phoryl groups from ATP to other compounds or from phosphorylated com¬  pounds to ADP. We will revisit these processes in our discussions of carbohydrate  metabolism in Chapters 15 and 16.   The compounds whose phosphoryl group-transfer potentials are greater  than that of ATP have additional stabilizing effects. For example, the hydrol¬  ysis of acyl phosphates (mixed phosphoric-carboxylic anhydrides), such as    is driven by the same competing resonance and differential solvation effects  that influence the hydrolysis of phosphoanhydrides (Fig. 14-6). Apparently,  these effects are more pronounced for acyl phosphates than for phosphoanhy¬  drides, as the rankings in Table 14-3 indicate.    which are below ATP in Table 14-3, have no significantly different resonance  stabilization or charge separation compared to their hydrolysis products. Their   The high phosphoryl group-transfer potentials of phosphoguanidines,  such as phosphocreatine and phosphoarginine, largely result from the com¬  peting resonances in the guanidino group, which are even more pronounced  than they are in the phosphate group of phosphoanhydrides:   Consequently, phosphocreatine can transfer its phosphoryl group to ADP to  form ATP.   Muscle and nerve cells, which have a high ATP turnover, rely on phospho-  guanidines to regenerate ATP rapidly. In vertebrates, phosphocreatine is syn¬  thesized by the reversible phosphorylation of creatine by ATP catalyzed by   Note that this reaction is endergonic under standard conditions; however, the  intracellular concentrations of its reactants and products are such that it operates  close to equilibrium (A G ~ 0). Accordingly, when the cell is in a resting state,  so that [ATP] is relatively high, the reaction proceeds with net synthesis of  phosphocreatine, whereas at times of high metabolic activity, when [ATP] is  low, the equilibrium shifts so as to yield net synthesis of ATP from phospho¬  creatine and ADP. Phosphocreatine thereby acts as an ATP “buffer” in cells that  contain creatine kinase. A resting vertebrate skeletal muscle normally has suffi¬  cient phosphocreatine to supply its free energy needs for several minutes (but  for only a few seconds at maximum exertion). In the muscles of some inver¬  tebrates, such as lobsters, phosphoarginine performs the same function. These  phosphoguanidines are collectively named phosphagens.   Nucleoside Triphosphates Are Freely Interconverted. Many biosynthetic  processes, such as the synthesis of proteins and nucleic acids, require nucleo¬  side triphosphates other than ATP. For example, RNA synthesis requires the  ribonucleotides CTP, GTP, and UTP, along with ATP, and DNA synthesis re¬  quires dCTP, dGTP, dTTP, and dATP (Section 3-1). All these nucleoside  triphosphates (NTPs) are synthesized from ATP and the corresponding nucle¬  oside diphosphate (NDP) in a reaction catalyzed by the nonspecific enzyme  nucleoside diphosphate kinase:   The A G or values for these reactions are nearly 0, as might be expected from the  structural similarities among the NTPs. These reactions are driven by the deple¬  tion of the NTPs through their exergonic utilization in subsequent reactions.   Other kinases reversibly convert nucleoside monophosphates to their  diphosphate forms at the expense of ATP. One of these phosphoryl group-  transfer reactions is catalyzed by adenylate kinase:   Chapter 14 Introduction to Metabolism This enzyme is present in all tissues, where it functions to maintain equilib¬   rium concentrations of the three nucleotides. When AMP accumulates, it is  converted to ADP, which can be used to synthesize ATP through substrate-  level phosphorylation, oxidative phosphorylation, or photophosphorylation.  The reverse reaction helps restore cellular ATP as rapid consumption of ATP  increases the level of ADP.   The X-ray structure of adenylate kinase, determined by Georg Schulz, re¬  veals that, in the reaction catalyzed by the enzyme, two ~ 30-residue domains  of the enzyme close over the substrates (Fig. 14-10), thereby tightly binding  them and preventing water from entering the active site (which would lead to  hydrolysis rather than phosphoryl group transfer). The movement of one of  the domains depends on the presence of four invariant charged residues.  Interactions between those groups and the bound substrates apparently trig¬  ger the rearrangements around the substrate-binding site (Fig. 14-10 b).   Once the adenylate kinase reaction is complete, the tightly bound prod¬  ucts must be rapidly released to maintain the enzyme’s catalytic efficiency. Yet  since the reaction is energetically neutral (the net number of phosphoanhy-  dride bonds is unchanged), another source of free energy is required for rapid  product release. Comparison of the X-ray structures of unliganded adenylate  kinase and adenylate kinase in complex with the bisubstrate model compound  Ap 5 A (AMP and ATP connected by a fifth phosphate) show how the enzyme  avoids the kinetic trap of tight-binding substrates and products: On binding  substrate, a portion of the protein remote from the active site increases its  chain mobility and thereby consumes some of the free energy of substrate  binding. The region “resolidifies” when the binding site is opened and the  products are released. This mechanism is thought to act as an “energetic coun¬  terweight” to help adenylate kinase maintain a high reaction rate.   FIG. 14-10 Conformational changes in E. coli adenylate  kinase on binding substrate, (a) The unliganded enzyme, (b) The  enzyme with the bound bisubstrate analog Ap 5 A. The Ap 5 A is shown  in stick form (C green, N blue, 0 red, and P yellow). The protein’s  cyan and blue domains undergo extensive conformational changes on    ligand binding, whereas the remainder of the protein (magenta), whose  orientation is the same in Parts a and b, largely maintains its  conformation. [Based on X-ray structures by Georg Schulz, Institut  fur Organische Chemie und Biochemie, Freiburg, Germany. PDBids  (a) 4AKE and (b) 1AKE] © See Interactive Exercise 14.   The ubiquity of phosphorylated compounds in metabolism is consistent with  their early evolutionary appearance. Yet phosphate is (and was) scarce in the  abiotic world, which suggests that other kinds of molecules might have served  as energy-rich compounds even before metabolic pathways became specialized  for phosphorylated compounds. One candidate for a primitive “high-energy”  compound is the thioester, which offers as its main recommendation its occur¬  rence in the central metabolic pathways of all known organisms. Notably, the  thioester bond is involved in substrate-level phosphorylation, an ATP-generating  process that is independent of—and presumably arose before—oxidative  phosphorylation.   The thioester bond appears in modern metabolic pathways as a reaction  intermediate (involving a Cys residue in an enzyme active site) and in the form  of acetyl-CoA (Fig. 14-11), the common product of carbohydrate, fatty acid,  and amino acid catabolism. Coenzyme A (CoASH or CoA) consists of a  (3-mercaptoethylamine group bonded through an amide linkage to the vita¬  min pantothenic acid, which, in turn, is attached to a 3'-phosphoadenosine  moiety via a pyrophosphate bridge. The acetyl group of acetyl-CoA is bonded  as a thioester to the sulfhydryl portion of the (3-mercaptoethylamine group.  CoA thereby functions as a carrier of acetyl and other acyl groups (the A of CoA  stands for “acetylation”). Thioesters also take the form of acyl chains bonded  to a phosphopantetheine residue that is linked to a Ser OH group in a pro¬  tein (Section 20-4C) rather than to 3'-phospho-AMP, as in CoA.   FIG. 1 4-1 1 The chemical structure of acetyl-CoA. The thioester bond is drawn with  a ~ to indicate that it is a “high-energy” bond (has a high negative free energy of  hydrolysis). In CoA, the acetyl group is replaced by hydrogen.   • What kinds of molecules do cells use as  energy currency?   • Describe the ways an exergonic process  can drive an endergonic process.   • Why is the activity of inorganic pyrophos¬  phatase metabolically indispensible?   • What are the cellular roles of nucleoside  diphosphate kinase and adenylate kinase?    (1 kj • mol x ) more exergonic than ATP hydrolysis. The hydrolysis of  thioesters is more exergonic than that of ordinary esters because the thioester  is less stabilized by resonance. This destabilization is a result of the large atomic  radius of S, which reduces the electronic overlap between C and S compared  to that between C and O.   The formation of a thioester bond in a metabolic intermediate conserves  a portion of the free energy of oxidation of a metabolic fuel. That free energy  can then be used to drive an exergonic process. In the citric acid cycle, for ex¬  ample, cleavage of a thioester (succinyl-CoA) releases sufficient free energy to  synthesize GTP from GDP and P z (Section 17-3E).    • The electron carriers NAD + and FAD accept electrons from reduced metabolites  and transfer them to other compounds.   • The reduction potential describes the tendency for an oxidized compound to gain  electrons (become reduced); the change in reduction potential for a reaction  describes the tendency for a given oxidized compound to accept electrons from a  given reduced compound.   • Free energy and reduction potential are negatively related: the greater the reduction  potential, the more negative the free energy and the more favorable the reaction.    As metabolic fuels are oxidized to C0 2 , electrons are transferred to molecular  carriers that, in aerobic organisms, ultimately transfer the electrons to molecu¬  lar oxygen. The process of electron transport results in a transmembrane  proton concentration gradient that drives ATP synthesis (oxidative phosphory¬  lation; Section 18-3). Even obligate anaerobes, which do not carry out oxida¬  tive phosphorylation, rely on the oxidation of substrates to drive ATP synthesis.  In fact, oxidation-reduction reactions (also known as redox reactions) supply  living things with most of their free energy. In this section, we examine the ther¬  modynamic basis for the conservation of free energy during substrate oxidation.   Two of the most widely occurring electron carriers are the nucleotide coen¬  zymes nicotinamide adenine dinucleotide (NAD + ) and flavin adenine dinu-  cleotide (FAD). The nicotinamide portion of NAD + (and its phosphorylated  counterpart NADP + ; Fig. 11-4) is the site of reversible reduction, which for¬  mally occurs as the transfer of a hydride ion (H ; a proton with two elec¬  trons) as indicated in Fig. 14-12. The terminal electron acceptor in aerobic  organisms, 0 2 , can accept only unpaired electrons (because each of its two   FIG. 1 4-1 2 Reduction of NAD + to NADH. R represents the ribose-pyrophosphoryl-  adenosine portion of the coenzyme. Only the nicotinamide ring is affected by reduction,  which is formally represented here as occurring by hydride transfer.   available lowest energy molecular orbitals is already occupied by one electron);  that is, electrons must be transferred to 0 2 one at a time. Electrons that are re¬  moved from metabolites as pairs (e.g., with the two-electron reduction of NAD + )  must be transferred to other carriers that can undergo both two-electron and one-  electron redox reactions. FAD (Fig. 14-13) is such a coenzyme.   The conjugated ring system of FAD can accept one or two electrons to  produce the stable radical (semiquinone) FADH* or the fully reduced (hydro-  quinone) FADH 2 (Fig. 14-14). The change in the electronic state of the ring  system on reduction is reflected in a color change from brilliant yellow (in  FAD) to pale yellow (in FADH 2 ). The metabolic functions of NAD + and  FAD demand that they undergo reversible reduction so that they can accept  electrons, pass them on to other electron carriers, and thereby be regenerated  to participate in additional cycles of oxidation and reduction.   Humans cannot synthesize the flavin moiety of FAD but, rather, must ob¬  tain it from their diets, for example, in the form of riboflavin (vitamin B 2 ;  Fig. 14-13). Nevertheless, riboflavin deficiency is quite rare in humans, in part  because of the tight binding of flavin prosthetic groups to their apoenzymes.  The symptoms of riboflavin deficiency, which are associated with general mal¬  nutrition or bizarre diets, include an inflamed tongue, lesions in the corner of  the mouth, and dermatitis.   Oxidation-reduction reactions resemble other types of group-transfer reactions  except that the “groups” transferred are electrons, which are passed from an   electron donor (reductant or reducing agent) to an electron acceptor  (oxidant or oxidizing agent).   Cu + , the reductant, is oxidized to Cu 2+ while Fe 3+ , the oxidant, is reduced  to Fe 2+ .    FIG. 14-14 Reduction of FAD to FADH 2 . R represents the ribitol—pyrophosphoryl—  adenosine portion of the coenzyme. The conjugated ring system of FAD can undergo two  sequential one-electron reductions or a two-electron transfer that bypasses the   half-cell undergoing oxidation (here Cu + —»  Cu 2+ + e“) passes the liberated electrons  through the wire to the half-cell undergoing  reduction (here e~ + Fe 3+ —> Fe 2+ ).  Electroneutrality in the two half-cells is  maintained by the transfer of ions through the  electrolyte-containing salt bridge.   whose sum is the whole reaction above. These particular half-reactions occur  during the oxidation of cytochrome c oxidase in the mitochondrion (Section  18-2F). Note that for electrons to be transferred, both half-reactions must  occur simultaneously. In fact, the electrons are the two half-reactions 5 com¬  mon intermediate.   A half-reaction consists of an electron donor and its conjugate electron ac¬  ceptor; in the oxidative half-reaction shown above, Cu + is the electron donor  and Cu 2+ is its conjugate electron acceptor. Together these constitute a redox  couple or conjugate redox pair analogous to a conjugate acid-base pair (HA  and A ; Section 2-2B). An important difference between redox pairs and  acid-base pairs, however, is that the two half-reactions of a redox reaction , each  consisting of a conjugate redox pair ; can be physically separated to form an elec¬  trochemical cell (Fig. 14-15). In such a device, each half-reaction takes place  in its separate half-cell, and electrons are passed between half-cells as an elec¬  tric current in the wire connecting their two electrodes. A salt bridge is nec¬  essary to complete the electrical circuit by providing a conduit for ions to  migrate and thereby maintain electrical neutrality.   The free energy of an oxidation-reduction reaction is particularly easy to  determine by simply measuring the voltage difference between its two half¬  cells. Consider the general reaction   in which n electrons per mole of reactants are transferred from reductant (B red )  to oxidant (A^). The free energy of this reaction is expressed as   where w is non-pressure—volume work. In this case, w is equivalent to w e \,  the electrical work required to transfer the n moles of electrons through the  electrical potential difference, [where the units of % are volts (V), the  number of joules (J) of work required to transfer 1 coulomb (C) of charge].  This, according to the laws of electrostatics, is    where 9% the faraday, is the electrical charge of 1 mol of electrons (1 =   96,485 C • mol 1 = 96,485 J * V 1 • mol x ), and n is the number of moles  of electrons transferred per mole of reactant converted. Thus, substituting Eq.  14-6 into Eq. 14-5,    Combining Eqs. 14-4 and 14-7, and making the analogous substitution for  A G°, yields the Nernst equation:    which was originally formulated in 1881 by Walther Nernst. Here % is the  reduction potential, the tendency for a substance to undergo reduction (gain  electrons). the electromotive force (emf), can be described as the “elec¬  tron pressure” that the electrochemical cell exerts. The quantity g°, the reduction  potential when all components are in their standard states, is called the stan¬  dard reduction potential. If these standard states refer to biochemical stan¬  dard states (Section 1-3D), then %° is replaced by % ot . Note that a positive  A% in Eq. 14-7 results in a negative AG; in other words, a positive A^ indi¬  cates a spontaneous reaction , one that can do work.   Equation 14-7 shows that the free energy change of a redox reaction can be de¬  termined by directly measuring its change in reduction potential with a voltmeter  (Fig. 14-15). Such measurements make it possible to determine the order of spon¬  taneous electron transfers among a set of electron carriers such as those of the  electron-transport pathway that mediates oxidative phosphorylation in cells.    where, by convention, both half-reactions are written as reductions. These half¬  reactions can be assigned reduction potentials, % A and in accordance with    For the overall redox reaction involving the two half-reactions, the difference  in reduction potential, A%°\ is defined as   Thus, when the reaction proceeds with A as the electron acceptor and B as  the electron donor, A%°' — %° A — %% and A% = % A —   Reduction potentials, like free energies, must be defined with respect to some  arbitrary standard, in this case, the hydrogen half-reaction   in which H + is in equilibrium with H 2 (g) that is in contact with a Pt elec¬  trode. This half-cell is arbitrarily assigned a standard reduction potential %° of  0 V (1 V = 1 J • C _1 ) at pH 0, 25°C, and 1 atm. Under the biochemical   Chapter 14 Introduction to Metabolism convention, where the standard state is pH 7.0, the hydrogen half-reaction has   When A.% is positive, AG is negative (Eq. 14-7), indicating a spontaneous  process. In combining two half-reactions under standard conditions, the di¬  rection of spontaneity therefore involves the reduction of the redox couple  with the more positive standard reduction potential. In other words, the more  positive the standard reduction potential, the higher the affinity of the redox  couples oxidized form for electrons, that is, the greater the tendency for the redox  couples oxidized form to accept electrons and thus become reduced.   Biochemical Half-Reactions Are Physiologically Significant. The biochem¬  ical standard reduction potentials (%°') of some biochemically important half¬  reactions are listed in Table 14-4. The oxidized form of a redox couple with  a large positive standard reduction potential has a high affinity for electrons  and is a strong electron acceptor (oxidizing agent), whereas its conjugate re-  ductant is a weak electron donor (reducing agent). For example, 0 2 is the  strongest oxidizing agent in Table 14-4, whereas H 2 0, which tightly holds its   Cytochrome a 3 (Fe 3+ ) + e \ cytochrome a 3 (Fe 2+ ) 0.385   0 2 + 2 H + + 2 e~ H 2 0 2 0.295   Cytochrome a (Fe 3+ ) + e \ cytochrome a (Fe 2+ ) 0.29   Cytochrome c (Fe 3+ ) + e v cytochrome c (Fe 2+ ) 0.235   Cytochrome c 1 (Fe 3+ ) + e \ cytochrome c 1 (Fe 2+ ) 0.22   Cytochrome b (Fe 3+ ) + e \ cytochrome b (Fe 2+ ) ( mitochondrial) 0.077   Ubiquinone + 2 H + + 2 e v ubiquinol 0.045    Source: Mosdy from Loach, P.A., In Fasman, G.D. (Ed.), Handbook of Biochemistry and Molecular  Biology (3rd ed.), Physical and Chemical Data, Vol. I, pp. 123-130, CRC Press (1976).   NADH + FAD + H + -^NAD + + FADH 2   Next, calculate the electromotive force (A% of ) from the standard reduction poten¬  tials given in Table 14-4, using one of the following methods.   Since FAD (% of = —0.219 V) is the electron acceptor, and NADH fS 0 ' =  — 0.315 V) is the electron donor,   Write the net reaction as a sum of the two relevant half-reactions. For FAD, the  half-reaction is the same as the reductive half-reaction given in Table 14-4, and its  value is —0.219 V. For NADH, which undergoes oxidation rather than reduc¬  tion, the half-reaction is the reverse of the one given in Table 14-4, and its  value is +0.315 V, the reverse of the reduction potential given in the table. The  two half-reactions are added to give the net oxidation-reduction reaction, and the  %°' values are also added:   NADH + FAD + H + —» NADH + + FADH 2 A%°' = +0.096 V   Next, use Eq. 14-7 to calculate A G°' . Because two moles of electrons are trans¬  ferred for every mole of NADH oxidized to NAD + , n = 2.   A G of = - (2) (96,485 J-V _1 -mol -1 ) (0.096 V) = -18.5 kj • mol -1   Since electrons spontaneously flow from low to high reduction potentials,  they are transferred, under standard conditions, from the reduced products in  any half-reaction in Table 14-4 to the oxidized reactants of any half-reaction  above it (see Sample Calculation 14-2). However, such a reaction may not oc¬  cur at a measurable rate in the absence of a suitable enzyme. Note that Fe 3+  ions of the various cytochromes listed in Table 14-4 have significantly differ¬  ent reduction potentials. This indicates that the protein components of redox en¬  zymes play active roles in electron-transfer reactions by modulating the reduction  potentials of their bound redox-active centers.   Electron-transfer reactions are of great biological importance. For exam¬  ple, in the mitochondrial electron-transport chain (Section 18-2), electrons are  passed from NADH along a series of electron acceptors of increasing reduc¬  tion potential (including FAD and others listed in Table 14-4) to 0 2 . ATP is  generated from ADP and P z by coupling its synthesis to this free energy cas¬  cade. NADH thereby functions as an energy-rich electron-transfer coenzyme. In  fact, the oxidation by 0 2 of one NADH to NAD + supplies sufficient free en¬  ergy to generate almost three ATPs. NAD + is an electron acceptor in many  exergonic metabolite oxidations. In serving as the electron donor in ATP syn¬  thesis, it fulfills its cyclic role as a free energy conduit in a manner analogous  to ATP (Fig. 14-9).   • What are the metabolic roles of the coen¬  zymes NAD + and FAD?   • Explain why NADH and FADH 2 are a type  of energy currency in the cell.   • When two half-reactions are combined,  how can you predict which compound will  be oxidized and which will be reduced?    • Metabolic pathways are often studied by tracing metabolites labeled with radionu¬  clides or NMR-active isotopes.   • The steps of a pathway can be identified by examining how metabolic inhibitors  and genetic defects lead to the accumulation of pathway intermediates.   • DNA microarrays and proteomics techniques are used to determine the genetic  expression of metabolic enzymes.    particles are electrons, |3 + particles are positrons,  and 7 rays are photons.   Source: Holden, N.E., in Lide, D.R. (Ed.),  Handbook of Chemistry and Physics (90th ed.),  pp. 11-57 to 266, CRC Press (2009-2010).   1. In terms of the sequence of reactions by which a specific nutrient is  converted to end products, and the energetics of the conversions.   2. In terms of the mechanisms by which each intermediate is converted  to its successor. Such an analysis requires the isolation and characteri¬  zation of the specific enzymes that catalyze each reaction.   3. In terms of the control mechanisms that regulate the flow of metabolites  through the pathway. These mechanisms include the interorgan relation¬  ships that adjust metabolic activity to the needs of the entire organism.   Elucidating a metabolic pathway on all these levels is a complex process, of¬  ten requiring contributions from a variety of disciplines.   The outlines of the major metabolic pathways have been known for  decades, although in many cases, the enzymology behind various steps of the  pathways remains unclear. Likewise, the mechanisms that regulate pathway ac¬  tivity under different physiological conditions are not entirely understood.  These areas are of great interest because of their potential to yield information  that could be useful in improving human health and curing metabolic diseases.  In addition, the unexplored metabolisms of unusual organisms, including re¬  cently discovered “extremophiles,” hold the promise of novel biological mate¬  rials and enzymatic processes that can be exploited for the environmentally  sensitive production of industrial materials, foods, and therapeutic drugs.   Early metabolic studies used whole organisms, often yeast, but also mam¬  mals. For example, Frederick Banting and Charles Best established the role of  the pancreas in diabetes in 1921; they surgically removed that organ from dogs  and observed that the animals then developed the disease (Box 22-2).  Techniques for studying metabolic processes have since become more refined,  progressing from whole-organ preparations and thin tissue slices to cultured  cells and isolated organelles. The most recent approaches include identifying  active genes and cataloguing their protein products.   A metabolic pathway in which one compound is converted to another can be  followed by tracing a specifically labeled metabolite. Franz Knoop formulated  this technique in 1904 to study fatty acid oxidation. He fed dogs fatty acids  chemically labeled with phenyl groups and isolated the phenyl-substituted end  products from the dogs’ urine. From the differences in these products, depend¬  ing on whether the phenyl-substituted starting material contained odd or even  numbers of carbon atoms, Knoop deduced that fatty acids are degraded in  two-carbon units (Section 20-2).   Chemical labeling has the disadvantage that the chemical properties of  labeled metabolites differ from those of normal metabolites. This problem is  largely eliminated by labeling molecules with isotopes. The fate of an isotopi-  cally labeled atom in a metabolite can therefore be elucidated by following its  progress through the metabolic pathway of interest. The advent of isotopic label¬  ing and tracing techniques in the 1940s revolutionized the study of metabo¬  lism. Some of the most common radioactive isotopes (radionuclides) used in  biochemistry are listed in Table 14-5, along with their half-lives and the type    of radioactivity emitted by the spontaneously disintegrating atomic nuclei.  Radioactive compounds can be detected by their ability to expose photo¬  graphic film. Alternatively, p particles and y rays can excite fluorescent com¬  pounds, and the emitted light can be measured.   One of the early advances in metabolic understanding resulting from the  use of isotopic tracers was the demonstration, by David Shemin and David  Rittenberg in 1945, that the nitrogen atoms of heme (Fig. 7-2) are derived  from glycine rather than from ammonia, glutamic acid, proline, or leucine  (Section 21-6A). They showed this by feeding rats the 13 N-labeled nutrients,  isolating the heme in their blood, and analyzing it by mass spectrometry for  15 N content. Only when the rats were fed [ 15 N] glycine did the heme contain  15 N. This technique was also used with the radioactive isotope l4 C to demon¬  strate that all of cholesterols carbon atoms are derived from acetyl-CoA  (Section 20-7A). Radioactive isotopes have become virtually indispensable for  establishing the metabolic origins of complex metabolites.   Another method for tracing the fates of labeled metabolites is nuclear  magnetic resonance (NMR), which detects specific isotopes, including X H, 13 C,  15 N, and 31 P, by their characteristic nuclear spins. Since the NMR spectrum  of a particular nucleus varies with its immediate environment, it is possible to  identify the peaks corresponding to specific atoms even in relatively complex  mixtures. The development of magnets large enough to accommodate animals  and humans, and to localize spectra to specific organs, has made it possible to  study metabolic pathways noninvasively by NMR techniques. For example, 31 P  NMR can be used to study energy metabolism in muscle by monitoring the  levels of phosphorylated compounds such as ATP, ADP, and phosphocreatine.   Iso topically labeling specific atoms of metabolites with 13 C (which is only  1.10% naturally abundant) permits the metabolic progress of the labeled  atoms to be followed by 13 C NMR. Figure 14-16 shows in vivo 13 C NMR  spectra of a rat liver before and after an injection of D-[1- 13 C]glucose. The 13 C  can be seen entering the liver and then being incorporated into glycogen (the  storage form of glucose; Section 16-2).   FIG. 14-16 The conversion of  [1- 13 C]glucose to glycogen as observed by  localized in vivo 13 C NMR. (a) The natural  abundance 13 C NMR spectrum of the liver of a  live rat. Note the resonance corresponding to Cl  of glycogen, (b) The 13 C NMR spectrum of the  liver of the same rat —5 min after it was  intravenously injected with 100 mg of  [1- 13 C]glucose (90% enriched). The resonances  of the Cl atom of both the a and p anomers of  glucose are clearly distinguishable from each  other and from the resonance of the Cl atom of  glycogen, (c) The 13 C NMR spectrum of the  liver of the same rat —30 min after the  [1- 13 C]glucose injection. The Cl resonances of  both the a- and p-glucose anomers are much  reduced while the Cl resonance of glycogen  has increased. [After Reo, N.V., Siegfried, B.A.,  and Acherman, J.J.H., J. Biol. Chem. 259, 13665  (1984).]   Many of the techniques used to elucidate the intermediates and enzymes of  metabolic pathways involve perturbing the system in some way and observing  how this affects the activity of the pathway One way to perturb a pathway is  to add certain substances, called metabolic inhibitors, that block the pathway  at specific points, thereby causing the preceding intermediates to build up. This  approach was used in elucidating the conversion of glucose to ethanol in yeast  by glycolysis (Section 15-2). Similarly, the addition of substances that block  electron transfer at different sites was used to deduce the sequence of electron  carriers in the mitochondrial electron-transport chain (Section 18-2B).   Archibald Garrods realization, in the early 1900s, that human genetic diseases  are the consequence of deficiencies in specific enzymes also contributed to the  elucidation of metabolic pathways. For example, upon the ingestion of either  phenylalanine or tyrosine, individuals with the largely harmless inherited con¬  dition known as alcaptonuria, but not normal subjects, excrete homogentisic  acid in their urine (Box 21-2). This is because the liver of alcaptonurics lacks  an enzyme that catalyzes the breakdown of homogentisic acid (Fig. 14-17).   FIG. 1 4-1 7 Pathway for phenylalanine  degradation. Alcaptonurics lack the enzyme that  breaks down homogentisate; therefore, this  intermediate accumulates and is excreted in the  urine.    Genetic Manipulation Alters Metabolic Processes. Early studies of metab¬  olism led to the astounding discovery that the basic metabolic pathways in most  organisms are essentially identical. This metabolic uniformity has greatly facili¬  tated the study of metabolic reactions. Thus, although a mutation that inac¬  tivates or deletes an enzyme in a pathway of interest may be unknown in  higher organisms, it can be readily generated in a rapidly reproducing microor¬  ganism through the use of mutagens (chemical agents that induce genetic  changes; Section 25-4A), X-rays, or, more recently, through genetic engineer¬  ing techniques (Section 3-5). The desired mutants, which cannot synthesize  the pathways end product, can be identified by their requirement for that  product in their culture medium.   Higher organisms that have been engineered to lack particular genes (i.e.,  gene “knockouts”; Section 3-5D) are useful, particularly in cases in which the  absence of a single gene product results in a metabolic defect but is not lethal.  Genetic engineering techniques have advanced to the point that it is possible  to selectively “knock out” a gene only in a particular tissue. This approach is  necessary in cases in which a gene product is required for development and  therefore cannot be entirely deleted. In the opposite approach, techniques for  constructing transgenic animals make it possible to express genes in tissues in  which they were not originally present.   Metabolism has traditionally been studied by hypothesis-driven research: iso¬  lating individual enzymes and metabolites and assembling them into meta¬  bolic pathways as guided by experimentally testable hypotheses. A new  approach, systems biology, has emerged with the advent of complete genome  sequences, the development of rapid and sensitive techniques for analyzing  large numbers of gene transcripts, proteins, and metabolites all at once, and  the development of new computational and mathematical tools. Systems bi¬  ology is discovery-based: collecting and integrating enormous amounts of data  in searchable databases so that the properties and dynamics of entire biologi¬  cal networks can be analyzed. As a result, our understanding of the path from  genotype to phenotype has expanded. In addition to the central dogma  (Section 3-3B) that a single gene composed of DNA is transcribed to mRNA  which is translated to a single protein that influences metabolism, we are   FIG. 1 4-1 8 The relationship between genotype and phenotype. The path from  genetic information (genotype) to metabolic function (phenotype) has several steps.  Portions of the genome are transcribed to produce the transcriptome, which directs the  synthesis of the proteome, whose various activities are responsible for synthesizing and  degrading the components of the metabolome.    increasingly taking into account the genome, transcriptome, proteome, and  metabolome and their interrelationships (Fig. 14-18). The term bibliome  (Greek: biblion , book) has even been coined to denote the systematic incor¬  poration of preexisting information about reaction mechanisms and metabolic  pathways. Dozens of pathways are catalogued in Internet-accessible databases  that list the structures and names of the intermediates and the enzymes that  catalyze their interconversion, along with links to gene sequences and three-  dimensional protein structures (see Bioinformatics Project 8). Two examples  of such databases are the Kyoto Encyclopedia of Genes and Genomes (KEGG)  Pathway Database: www.genome.jp/kegg/metabolism.html; and BRENDA  (BRaunschweig ENzyme DAtabase): www.brenda-enzymes.org/. In the follow¬  ing paragraphs we discuss some of the emerging technologies used in systems  biology.   Genomics Examines the Entire Complement of an Organism’s DNA  Sequences. The overall metabolic capabilities of an organism are encoded by  its genome (its entire complement of genes). In theory, it should be possible  to reconstruct a cells metabolic activities from its DNA sequences. At pres¬  ent, this can be done only in a general sense. For example, the sequenced  genome of Vibrio cholerae, the bacterium that causes cholera, reveals a large  repertoire of genes encoding transport proteins and enzymes for catabolizing  a wide range of nutrients. This is consistent with the complicated lifestyle of  V cholerae, which can live on its own, in association with zooplankton, or in  the human gastrointestinal tract (where it causes cholera). Of course, a sim¬  ple catalog of an organisms genes does not reveal how the genes function.  Thus, some genes are expressed continuously at high levels, whereas others are  expressed rarely, for example, only when the organism encounters a particular  metabolite.   Creating an accurate picture of gene expression is the goal of transcriptomics,  the study of a cells transcriptome (which, in analogy with the word “genome,”  is the entire collection of RNAs that the cell transcribes). Identifying and  quantifying all the transcripts from a single cell type reveals which genes are  active. Cells transcribe thousands of genes at once so this study requires the  use of new techniques, including DNA microarray technology.   DNA microarrays or DNA chips are made by depositing numerous (up  to several hundred thousand) different DNA segments of known gene    FIG. 14-19 DNA chips, (a) Schematic diagram of an experiment  showing the differences in yeast gene expression in the presence and  absence of glucose: (1) Yeast cells are grown in medium containing  glucose and in glucose-depleted medium. (2) mRNA is isolated from  each population of yeast. (3) Reverse transcriptase copies the mRNA  to cDNA, incorporating a red fluorescent dye into the cDNA from the  cells grown in glucose, and a green fluorescent dye for the cells  harvested after glucose depletion. (4) The cDNAs are mixed. (5) The  labeled cDNAs hybridize with DNA segments immobilized on a gene  chip, and the bound red and green fluorescent cDNAs are detected.    (b) An —6000-gene array DNA chip containing most of the genes  from baker’s yeast, one per spot. The red and green spots,  respectively, reveal those genes that are transcriptionally activated by  the presence or absence of glucose, whereas the yellow spots (red  plus green ) indicate genes whose expression is unaffected by the  level of glucose. [Courtesy of Patrick Brown, Stanford University  School of Medicine.] (c) A DNA microarray assembly. It protects its  enclosed DNA chip and provides a convenient hybridization chamber.  Interrogation requires a specialized fluorescence measurement device.  [Courtesy of Affymetrix, Inc., Santa Clara, California.]    sequences in a precise array on a solid support such as a coated glass surface.  These DNAs are often PCR-amplified cDNA clones derived from mRNAs  (PCR is discussed in Section 3-5C) or their robotically synthesized counter¬  parts. The mRNAs extracted from cells, tissues, or other biological sources  grown under differing conditions are then reverse-transcribed to cDNA,  labeled with a fluorescent dye (a different color for each growth condition),  and allowed to hybridize with the DNAs on the DNA microarray. After the  unhybridized cDNA is washed away, the resulting fluorescence intensity and  color at each site on the DNA microarray indicates how much cDNA (and  therefore how much mRNA) has bound to a particular complementary DNA  sequence for each growth condition. Figure 14-19 shows a DNA chip that  indicates the changes in yeast gene expression when yeast grown on glucose  have depleted their glucose supply.   Differences in the expression of particular genes have been correlated with  many developmental processes or growth patterns. For example, DNA microar¬  rays have been used to profile the patterns of gene expression in tumor cells be¬  cause different types of tumors synthesize different types and amounts of proteins  (Fig. 14-20). This information is useful in choosing how best to treat a cancer.   Proteomics Studies All the Cell’s Proteins. Unfortunately, the correlation  between the amount of a particular mRNA and the amount of its protein  product is imperfect. This is because the various mRNAs and their correspon¬  ding proteins are synthesized and degraded at different rates. Furthermore,  many proteins are posttranslationally modified, sometimes in several different  ways (e.g., by phosphorylation or glycosylation). Consequently, the number of  unique proteins in a cell exceeds the number of unique mRNAs.   A more reliable way than transcriptomics to assess gene expression is to  examine a cells proteome, the complete set of proteins that the cell synthe¬  sizes. This proteomics approach requires that the proteins first be separated,  usually by two-dimensional (2D) gel electrophoresis (a technique that sepa¬  rates proteins by isoelectric point in one direction and by mass in the perpen¬  dicular direction; Section 5-2D). Individual proteins are then identified by  using tandem mass spectrometry to obtain amino acid sequence information  (Section 5-3D) and correlating it with protein sequence databases. Because  many peptides are generated from a single protein, the technique enables the  redundant and unambiguous identification of that protein from the database.  In this way we can catalogue all the proteins that are contained in a cell or  tissue under a given set of conditions.   Can we compare all the proteins synthesized by a cell under two different sets  of conditions as is done for mRNA? The answer is yes, by using different isotopi-  cally labeled reagents that are either contained in the growth medium (e.g., deuter-  ated amino acids) or that are reacted with the cell extract. One technique for  labeling cellular proteins uses isotope-coded affinity tags (ICAT), which are anal¬  ogous to the different fluorescent dyes that label cDNA.   An ICAT contains three functional elements: an iodoacetyl group to re¬  act with cysteine residues, a linker that contains either 8 hydrogen (light) or  8 deuterium (heavy) atoms, and biotin, a coenzyme (Section 16-4A) that is  also used as a biotechnology tool because of its extremely tight binding to  the protein avidin (K = 10 15 M; Fig. 14-21^). Avidin is immobilized on a  chromatographic resin so that the ICAT-labeled peptides can be isolated by  biotin/avidin affinity chromatography (Section 5-2C).   The ICAT procedure is illustrated in Fig. \4-2\b. Two protein mixtures rep¬  resenting two different growth conditions are treated with light (dO) or heavy  (d8) versions of the ICAT reagent. The labeled protein mixtures are then com¬  bined and digested with trypsin to form Cys-containing labeled peptides, which  are then purified by biotin/avidin affinity chromatography. Individual peptides  are separated by liquid chromatography and detected by mass spectrometry  (LC/MS). The ratio of the intensities of the light and heavy peptide signals   FIG. 1 4-20 The relative transcriptional  activities of the genes in hepatocellular  carcinoma (HCC) tumors as determined  using DNA microarrays. The data are  presented in matrix form with each column  representing one of 156 tissue samples [82  HCC tumors (the most common human liver  cancer and among the five leading causes of  cancer deaths in the world) and 74 nontumor  liver tissues] and each row representing one of  3180 genes (those of the —17,400 genes on  the DNA microarray with the greatest variation in  transcriptional activity among the various tissue  samples). The data are arranged so as to group  the genes as well as the tissue samples on the  basis of similarities of their expression patterns.  The color of each cell indicates the expression  level of the corresponding gene in the  corresponding tissue relative to its mean  expression level in all the tissue samples, with  bright red, black, and bright green indicating  expression levels of 4, 1, and 1/4 times that of  the mean for that gene (as indicated on the  scale below). The dendrogram at the top of the  matrix indicates the similarities in expression  patterns among the various tissue samples.  [Courtesy of David Botstein and Patrick Brown,  Stanford University School of Medicine.]   Two cell states:  Reduce and label  cysteine-containing  proteins with  hydrogen (dO) or  deuterium (d8) ICAT  reagent.    Determine amino acid  sequence of each peptide  by MS/MS to identify the  protein of origin.    FIG. 14-21 The isotope-coded affinity tag (ICAT) method for residue. X denotes the position of hydrogen (dO) or deuterium (d8).  quantitative proteome analysis, (a) An example of an ICAT reagent ( b ) The ICAT strategy for differential labeling of proteins expressed  that contains an iodoacetyl reactive group, a linker, and a biotin by cells under two different sets of conditions. See text for details.   • Which of the isotopes listed in Table 14-5  could be used to specifically label a pro¬  tein? A nucleic acid?   • Explain how the buildup of a metabolite  when an enzyme is blocked can shed light  on the steps of a metabolic pathway.   • Describe how information about an organ¬  ism’s genome can be used to assess and  manipulate its metabolic activities.   • What is the difference between hypothesis-  driven and discovery-based research?   • Summarize the relationships between an  organism’s metabolome, proteome, tran-  scriptome, and genome.   • Why might a DNA chip and an ICAT analy¬  sis reveal different information about the  metabolic activity of a particular tissue?    indicates the relative peptide abundance in the two samples. Tandem mass spec¬  trometry (MS/MS) is then used to determine the amino acid sequence of each  peptide and identify the protein from which it is derived by comparing the se¬  quence to a database of all known proteins. This method was used to identify  many of the yeast proteins whose mRNA concentrations increased or decreased  when glucose was depleted from the growth medium (Fig. 14-19^). A hope for  the future is that samples from diseased and normal subjects can be compared  in this manner to find previously undetected disease markers that would allow  early diagnosis of various diseases.   Metabolomics Analyzes All of a Cell’s Metabolites. In order to describe a  cells functional state (its phenotype) we need, in addition to the cells genome,  transcriptome, and proteome, a quantitative description of all of the metabo¬  lites it contains under a given set of conditions, its metabolome. However, a  cell or tissue contains thousands of metabolites with vastly different proper¬  ties, so that identifying and quantifying all these substances is a daunting task,  requiring many different analytical tools. Consequently, this huge undertak¬  ing is often subdivided. For example, lipidomics is the subsection of  metabolomics aimed at characterizing all lipids in a cell under a particular set  of conditions, including how these lipids influence membrane structure, cell  signaling, gene expression, cell-cell interactions, and so on.   • The free energy released from catabolic oxidation reactions is used  to drive endergonic anabolic reactions.   • Nutrition is the intake and utilization of food to supply free en¬  ergy and raw materials.   • Heterotrophic organisms obtain their free energy from compounds  synthesized by chemolithotrophic or photoauto trophic organisms.   • Metabolic pathways are sequences of enzyme-catalyzed reactions  that occur in different cellular locations.   • Near-equilibrium reactions are freely reversible, whereas reactions  that function far from equilibrium serve as regulatory points and  render metabolic pathways irreversible.   • Flux through a metabolic pathway is controlled by regulating the  activities of the enzymes that catalyze its rate-determining steps.   • The free energy of the “high-energy” compound ATP is made  available through cleavage of one or both of its phosphoanhydride  bonds.   • An exergonic reaction such as ATP or PP Z hydrolysis can be cou¬  pled to an endergonic reaction to make it more favorable.   • Substrate-level phosphorylation is the synthesis of ATP from ADP  by phosphoryl group transfer from another compound.    • The common product of carbohydrate, lipid, and protein catabo¬  lism, acetyl-CoA, is a “high-energy” thioester.   • The coenzymes NAD + and FAD are reversibly reduced during the  oxidation of metabolites.   • The Nernst equation relates the electromotive force of a redox re¬  action to the standard reduction potentials and concentrations of  the electron donors and acceptors.   • Electrons flow spontaneously from the reduced member of a re¬  dox couple with the more negative reduction potential to the ox¬  idized member of a redox couple with the more positive reduction  potential.   • Studies of metabolic pathways determine the order of metabolic  transformations, their enzymatic mechanisms, their regulation,  and their relationships to metabolic processes in other tissues.   • Metabolic pathways are studied using isotopic tracers, enzyme in¬  hibitors, natural and engineered mutations, DNA microarrays,  and proteomics techniques.   • Systems biology endeavors to quantitatively describe the properties  and dynamics of biological networks as a whole through the inte¬  gration of genomic, transcriptomic, proteomic, and metabolomic  information.    metabolism 436  catabolism 436  anabolism 436  nutrition 437  autotrophy 437  chemolithotroph 437  photoautotroph 437  heterotroph 437  anaerobic 437  aerobic 437  macronutrient 437  micronutrient 437   phosphorylation 452  oxidative phosphorylation 452  photophosphorylation 452  kinase 452  phosphagen 453  reducing agent 457  oxidizing agent 457  half-reaction 458  redox couple 458  conjugate redox pair 458   systems biology 464  genomics 465  transcrip tomics 465  DNA microarray 465  proteomics 467  metabolomics 468    1. Explain why a heterotrophic organism may require vitamins  whereas an autotroph does not.   2. Methanogens are prokaryotes that produce methane according to  the net equation   4 H 2 + C0 2 ->CH 4 + 2 H 2 0    (a) Classify these two types of bacteria as autotrophic or  heterotrophic.   (b) Explain why the two types of bacteria are often found  associated with each other.   3. A strain of bacteria isolated from an alkaline lake with a high  concentration of arsenic is able to incorporate As into biological  molecules. What class of molecules is most likely to contain As  as part of its structure?   4. Explain why cadmium and mercury are toxic to most organisms.   5. Rank the following compounds in order of increasing oxidation  state.   6. In the partial reactions shown below, is the reactant undergoing  oxidation or reduction?   The standard free energy change for the reaction is —31.5  kj • mol 1 . (a) Calculate the equilibrium constant for this re¬  action at 37 °C. (b) Would you expect this reaction to serve as  a control point for its pathway (the citric acid cycle)?   8 . A certain metabolic reaction takes the form A —> B. Its standard  free energy change is 7.5 kj • mol \ (a) Calculate the equilib¬  rium constant for the reaction at 25°C. (b) Calculate A Cat 37 °C  when the concentration of A is 0.5 mM and the concentration  of B is 0.1 mM. Is the reaction spontaneous under these condi¬  tions? (c) How might the reaction proceed in the cell?   10 . Cells carry out anabolic as well as catabolic pathways, with some  enzymes functioning in both types of pathways, (a) Explain why  these enzymes catalyze near-equilibrium reactions, (b) Explain  why opposing anabolic and catabolic pathways must have differ¬  ent enzymes for at least one of the steps.   11. Assuming 100% efficiency of energy conservation, how many  moles of ATP can be synthesized under standard conditions by  the complete oxidation of 1 mol of glucose?   12. Assuming 100% efficiency of energy conservation, how many  moles of ATP can be synthesized under standard conditions by  the complete oxidation of 1 mol of palmitate?   13. Does the magnitude of the free energy change for ATP hydroly¬  sis increase or decrease as the pH increases from 5 to 6 ?   14. The A G°' for hydrolytically removing a phosphoryl group from    ATP is about twice as large as the A G°' for hydrolytically remov¬  ing a phosphoryl group from AMP (-14 kj • mol x ). Explain the  discrepancy.   has A G°' = +4.6 kj • mol 1 . What is the thermodynamic driv¬  ing force for this reaction?   Malate + NAD + —> oxaloacetate + NADH + H +  has a A G°' value of +29.7 kj 'mol \ (a) Would this reaction  occur spontaneously in a cell? (b) How does the citrate synthase  reaction (described in Problem 7) promote the malate dehydro¬  genase reaction in the cell? What is the overall change in free en¬  ergy for the two reactions?   17. Predict whether creatine kinase will operate in the direction  of ATP synthesis or phosphocreatine synthesis at 25°C when  [ATP] = 4 mM, [ADP] = 0.15 mM, [phosphocreatine] =  2.5 mM, and [creatine] = 1 mM.   18. If intracellular [ATP] = 5 mM, [ADP] = 0.5 mM, and [PJ =  1.0 mM, calculate the concentration of AMP at pH 7 and 25°C  under the condition that the adenylate kinase reaction is at equi¬  librium.   19. List the following substances in order of their increasing oxidiz¬  ing power: (a) acetoacetate, (b) cytochrome b (Fe 3+ ), (c) NAD + ,  (d) SO 4 , and (e) pyruvate.   20. Is the reduced form of cytochrome c more likely to give up its  electron to oxidized cytochrome a or cytochrome b ?   21. In a mixture of NAD + , NADH, ubiquinone, and ubiquinol,  which compound will be oxidized? Which will be reduced?   22 . Aerobic organisms transfer electrons from reduced fuel molecules  to 0 2 , forming H 2 0. Some anaerobic organisms use nitrate  (N0 3 ) as an acceptor for electrons from reduced fuel molecules.  Use the information in Table 14-4 to explain why aerobic organ¬  isms can harvest more free energy from a fuel molecule than can  a nitrate-using anaerobe.   23. Under standard conditions, will the following reaction proceed  spontaneously as written?   Fumarate + NADH + H + v succinate + NAD +   Cyto ^ (Fe 2 + ) + cyto £(Fe 3+ )   cyto a (Fe 3+ ) + cyto b (Fe 2+ )   25. Write a balanced equation for the oxidation of ubiquinol by cy¬  tochrome c. Calculate A G°' and A^°' for the reaction.   26. Under standard conditions, is the oxidation of free FADH 2 by  ubiquinone sufficiently exergonic to drive the synthesis of ATP?   27. A hypothetical three-step metabolic pathway consists of interme¬  diates W, X, Y, and Z and enzymes A, B, and C. Deduce the or¬  der of the enzymatic steps in the pathway from the following  information:   1 . Compound Q, a metabolic inhibitor of enzyme B, causes  Z to build up.   3. An inhibitor of enzyme A causes W, Y, and Z to accumu¬  late.   4. Compound P, a metabolic inhibitor of enzyme C, causes  W and Z to build up.   where A, B, C, and D are the intermediates, and X, Y, and Z are  the enzymes that catalyze the reactions. The physiological free en¬  ergy changes for the reactions are   X —0.2 kj • moP 1  Y —12.3 kj • mol 1  Z — 1.2 kj • mol 1   (a) Which reaction is likely to be a major regulatory point for  the pathway? (b) If your answer in Part a was in fact the case, in  the presence of an inhibitor that blocks the activity of enzyme Z,  would the concentrations of A, B, C, and D increase, decrease,  or not be affected?   29. Would gene chips containing bacterial DNA segments be useful  for monitoring gene expression in a mammalian cell?   30. Why do DNA chips often contain segments derived from cDNA  rather than genomic DNA segments?   1. Metabolic Enzymes. Use the KEGG and Enzyme Structure  databases to obtain information about dihydrofolate reductase.   2. Microarrays. Learn about microarray technology and its use in  studying disease.   5. Two-Dimensional Gel Electrophoresis. Obtain data about  dihydrofolate reductase from the Swiss-2D PAGE resource.   1. The Metabolomics Society. Learn about biomarkers-how they  are identified, followed analytically, and used to assess disease  states.   2. The Human Metabolome Database. See how an individual  metabolite (UDP-Glucose) is followed by NMR and Mass  Spectrometry and follow links to the many pathways that involve  UDP-Glucose.   3. The PubChem Project. Explore one of the newest databases  at NCBI, that contains compound, substance, and bioassay  information about small molecule metabolites.   Focus concept: An enzyme involved in nucleotide synthesis is sub¬  ject to regulation by a variety of combinations of nucleotides.    Access the Human Metabo¬  lome Database (http://www.hmdb.ca/) and search for information  on uric acid. In what metabolic pathway is uric acid an intermedi¬  ate? Does the pathway differ among species? What compounds  are the precursors of uric acid? To which compounds can uric acid  be converted? What is the normal concentration of uric acid in  body fluids? How do diseases affect uric acid levels?    Aebersold, R., Quantitative proteome analysis: Methods and applications,  J. Infect. Dis. 182 (supplement 2), S315-S320 (2003).   Alberty, R.A., Calculating apparent equilibrium constants of enzyme-  catalyzed reactions at pH 7, Biochem. Ed. 28, 12-17 (2000).   Campbell, A.M. and Heyer, L.J., Discovering Genomics, Proteomics and  Bioinformatics (2nd ed.), Pearson Benjamin Cummings, New York  (2007). [An interactive introduction to these subjects.]   Duarte, N.C., Becker, S.A., Jamshidi, N., Thiele, I., Mo, M.L., Vo, T.D.,  Srivas, R., and Palsson, B. 0., Global reconstruction of the human  metabolic network based on genomic and bibliomic data, Proc. Natl.  Acad. Sci, 104, 1777-1782 (2007).   Go, V.L.W., Nguyen, C.T.H., Harris, D.M., and Lee, W.-N.P., Nutrient-  gene interaction: Metabolic genotype-phenotype relationship,  J. Nutr. 135, 2016s-3020s (2005).   Hanson, R.W., The role of ATP in metabolism, Biochem. Ed. 17, 86-92  (1989). [Provides an excellent explanation of why ATP is an energy  transducer rather than an energy store.]   Lassila, J.K., Zalatan, J.G., and Herschlag, D., Biological phosphoryl  transfer reactions: understanding mechanism and catalysis, Annu. Rev.  Biochem. 80, 669-702 (2011).    Schulman, R.G. and Rothman, D.L., 13 C NMR of intermediary metab¬  olism: Implications for systematic physiology, Annu. Rev. Physiol. 63,  15-48 (2001).   Smolin, L.A. and Grosvenor, M.B, Nutrition: Science and Applications ,  Wiley (2008). [A good text for those interested in pursuing nutri¬  tional aspects of metabolism.]   Staughton,, R.B., Applications of DNA microarrays in biology, Annu.  Rev. Biochem. 74, 53-82 (2005).   Valle, D. (Ed.), The Online Metabolic & Molecular Bases of Inherited Dis¬  ease, http://www.ommbid.com/ [Most chapters in this encyclopedic  work include a review of a normal metabolic process that is dis¬  rupted by disease; access to this website requires a subscription  (often available through university and college libraries).]   Xia, Y., Yu, H., Jansen, R., Seringhaus, M., Baxter, S., Greenbaum, D.,  Zhao, H., and Gerstein, M., Analyzing cellular biochemistry in terms  of molecular networks, Annu. Rev. Biochem. 73, 1051-1087 (2004).   A Hexokinase Uses the First ATP  B Phosphoglucose Isomerase Converts Glucose-  6-Phosphate to Fructose-6-Phosphate  C Phosphofructokinase Uses the Second ATP  D Aldolase Converts a 6-Carbon Compound to  Two 3-Carbon Compounds  E Those Phosphate Isomerase Interconverts  Dihydroxyacetone Phosphate and  Glyceraldehyde-3-Phosphate  F Glyceraldehyde-3-Phosphate Dehydrogenase  Forms the First “High-Energy” Intermediate  G Phosphoglycerate Kinase Generates the  First ATP   H Phosphoglycerate Mutase Interconverts  3-Phosphoglycerate and 2-Phosphoglycerate  I Enolase Forms the Second “High-Energy”  Intermediate   A Phosphofructokinase Is the Major Flux-  Controlling Enzyme of Glycolysis in Muscle  B Substrate Cycling Fine-Tunes Flux Control   A Fructose Is Converted to Fructose-6-Phosphate  or Glyceraldehyde-3-Phosphate  B Galactose Is Converted to Glucose-6-Phosphate  C Mannose Is Converted to Fructose-6-Phosphate   A Oxidative Reactions Produce NADPH in Stage 1  B Isomerization and Epimerization of Ribulose-5-  Phosphate Occur in Stage 2  C Stage 3 Involves Carbon-Carbon Bond  Cleavage and Formation  D The Pentose Phosphate Pathway Must Be  Regulated    The protozoan Trypanosoma brucei (purple), which causes trypanosomiasis, or sleeping  sickness, relies almost entirely on glucose catabolism to extract free energy while it  travels through the bloodstream. Although the parasite is a eukaryote, it differs enough  from its host that its glycolytic enzymes offer targets for developing drugs that will not  affect human enzymes. [© Eye of Science/Photo Researchers, Inc.]    Glucose is a major source of metabolic energy in many cells. The fermenta¬  tion (anaerobic breakdown) of glucose to ethanol and C0 2 by yeast has been  exploited for many centuries in baking and brewing. However, scientific in¬  vestigation of the chemistry of this catabolic pathway began only in the mid¬  nineteenth century, with the experiments of Louis Pasteur and others. Nearly  a century would pass before the complete pathway was elucidated. During that  interval, several important features of the pathway came to light:   1. In 1905, Arthur Harden and William Young discovered that phosphate  is required for glucose fermentation.   2. Certain reagents, such as iodoacetic acid and fluoride ion, inhibit the  formation of pathway products, thereby causing pathway intermediates  to accumulate. Different substances caused the buildup of different in¬  termediates and thereby revealed the sequence of molecular intercon¬  versions.   3. Studies of how different organisms break down glucose indicated that,  with few exceptions, all of them do so the same way.   The efforts of many investigators came to fruition in 1940, when the com¬  plete pathway of glucose breakdown was described. This pathway, which is  named glycolysis (Greek: glykus , sweet + lysis , loosening), is alternately known  as the Embden-Meyerhof-Parnas pathway to commemorate the work of  Gustav Embden, Otto Meyerhof, and Jacob Parnas in its elucidation. The dis¬  covery of glycolysis came at a time when other significant inroads were being  made in the area of metabolism (Box 15-1).   Glycolysis, which is among the most completely understood biochemical  pathways, is a sequence of 10 enzymatic reactions in which one molecule of  glucose is converted to two molecules of the three-carbon compound pyruvate  with the concomitant generation of 2 ATP. It plays a key role in energy me¬  tabolism by providing a significant portion of the free energy used by most  organisms and by preparing glucose and other compounds for further oxida-   the great figures in biochemistry—by virtue  of his own contributions and his influence  on younger researchers—is the German  biochemist Otto Warburg. His long career  spanned a period during which studies of  whole organisms and crude extracts gave  way to molecular explanations of biological  structure and function. Like others of his  generation, he earned a doctorate in  chemistry at an early age and went on to obtain a medical degree,  although he spent the remainder of his career in scientific research  rather than in patient care. He became interested primarily in three  subjects related to the chemistry of oxygen and carbon dioxide:  respiration, photosynthesis, and cancer.   One of Warburg’s first accomplishments was to develop a  technique for studying metabolic reactions in thin slices of animal  tissue. This method produced more reliable results than the  alternative practice of chopping or mincing tissues (such  manipulations tend to release lysosomal enzymes that degrade  enzymes and other macromolecules). Warburg was also largely  responsible for refining manometry, the measurement of gas  pressure, as a technique for analyzing the consumption and  production of 0 2 and C0 2 by living tissues.   Warburg received a Nobel prize in 1931 for his discovery of the  catalytic role of iron porphyrins (heme groups) in biological oxidation  (the subject was the reaction carried out by the enzyme complex  now known as cytochrome c oxidase; Section 18-2F). Warburg also    identified nicotinamide as an active part of some enzymes. In 1944,  he was offered a second Nobel prize for his work with enzymes, but  he was unable to accept the award, owing to Hitler’s decree that  Germans could not accept Nobel prizes. In fact, Warburg’s apparent  allegiance to the Nazi regime incensed some of his colleagues in  other countries and may have contributed to their resistance to  some of his more controversial scientific pronouncements. In any  case, Warburg was not known for his warm personality. He was  never a teacher and tended to recruit younger research assistants  who were expected to move on after a few years.   In addition to the techniques he developed, which were widely  adopted, and a number of insights into enzyme action, Warburg  formulated some wide-reaching theories about the growth of cancer  cells. He showed that cancer cells could live and develop even in the  absence of oxygen. Moreover, he came to believe that anaerobiosis  triggered the development of cancer, and he rejected the notion that  viruses could cause cancer, a principle that had already been  demonstrated in animals but not in humans. In the eyes of many,  Warburg was guilty of equating the absence of evidence with the  evidence of absence in the matter of virus-induced human cancer.  Nevertheless, Warburg’s observations of cancer cell metabolism, which  is generally characterized by a high rate of glycolysis, were sound.   Even today, the oddities of tumor metabolism offer opportunities for  chemotherapy. Warburg’s dedication to his research in cancer and  other areas is revealed by the fact that he continued working in his  laboratory until just a few days before his death at age 87.    Warburg, 0., On the origin of cancer cells, Science 123, 309-314 (1956).   tive degradation. Thus, it is fitting that we begin our discussion of specific  metabolic pathways by considering glycolysis. We examine the sequence of  reactions by which glucose is degraded, along with some of the relevant en¬  zyme mechanisms. We will then examine the features that influence glycolytic  flux and the ultimate fate of its products. Finally, we will discuss the catabo¬  lism of other hexoses and the pentose phosphate pathway, an alternative path¬  way for glucose catabolism that functions to provide biosynthetic precursors.    Glycolysis involves the breakdown of glucose to pyruvate while using the free  energy released in the process to synthesize ATP from ADP and P,.   The 10-reaction sequence of glycolysis is divided into two stages: energy invest¬  ment and energy recovery.    Before beginning our detailed discussion of glycolysis, let us first take a mo¬  ment to survey the overall pathway as it fits in with animal metabolism as a  whole. Glucose usually appears in the blood as a result of the breakdown of  polysaccharides (e.g., liver glycogen or dietary starch and glycogen) or from  its synthesis from noncarbohydrate precursors (gluconeogenesis; Section 16-4).  Glucose enters most cells by a specific carrier that transports it from the ex¬  terior of the cell into the cytosol (Section 10-2E). The enzymes of glycolysis  are located in the cytosol, where they are only loosely associated, if at all, with  each other or with other cell structures.    FIG. 15-1 Glycolysis. In its first stage (Reactions 1-5), one  molecule of glucose is converted to two glyceraldehyde-3-phosphate  (GAP) molecules in a series of reactions that consumes 2 ATP. In the  second stage of glycolysis (Reactions 6-10), the two  glyceraldehyde-3-phosphate molecules are converted to two   Without looking at the text, write the overall equation for  glycolysis.   Glycolysis converts glucose to two C 3 units (pyruvate). The free energy released Section 2 The Reactions of Glycolysis   in the process is harvested to synthesize ATP from ADP and P z . Thus, glycolysis  is a pathway of chemically coupled phosphorylation reactions (Section 14-2B).   The 10 reactions of glycolysis are diagrammed in Fig. 15-1. Note that ATP  is used early in the pathway to synthesize phosphorylated compounds  (Reactions 1 and 3) but is later resynthesized twice over (Reactions 7 and 10).   Stage I Energy investment (Reactions 1-5). In this preparatory stage, the  hexose glucose is phosphorylated and cleaved to yield two mol¬  ecules of the triose glyceraldehyde-3-phosphate. This process  consumes 2 ATP.   Stage II Energy recovery (Reactions 6-10). The two molecules of glycer-  aldehyde-3-phosphate are converted to pyruvate, with concomi¬  tant generation of 4 ATP. Glycolysis therefore has a net “profit”  of 2 ATP per glucose: Stage I consumes 2 ATP; Stage II pro¬  duces 4 ATP.    The phosphoryl groups that are initially transferred from ATP to the hexose  do not immediately result in “high-energy” compounds. However, subsequent  enzymatic transformations convert these “low-energy” products to compounds  with high phosphoryl group-transfer potentials, which are capable of phospho-  rylating ADP to form ATP. The overall reaction is   2 pyruvate + 2 NADH + 2 ATP + 2H 2 0 + 4H +   Hence, the NADH formed in the process must be continually reoxidized to  keep the pathway supplied with its primary oxidizing agent, NAD + . In Section  15-3, we examine how organisms do so under aerobic or anaerobic conditions.   • How many ATP are invested and how many  are recovered from each molecule of glu¬  cose that follows the glycolytic pathway?   • Compare the oxidation states of glucose  and pyruvate. Explain why glycolysis  generates NADH.    The 10 steps of glycolysis can be described in terms of their substrates, prod¬  ucts, and enzymatic mechanisms.   Glycolytic enzymes catalyze phosphorylation reactions, isomerizations,  carbon-carbon bond cleavage, and dehydration.   ATP is consumed in Steps 1 and 3 but regenerated in Steps 7 and 10 for a net  yield of 2 ATP per glucose.    In this section, we examine the reactions of glycolysis more closely, describing  the properties of the individual enzymes and their mechanisms. As we study  the individual glycolytic enzymes, we will encounter many of the catalytic  mechanisms described in Section 11-3.    Reaction 1 of glycolysis is the transfer of a phosphoryl group from ATP to  glucose to form glucose-6-phosphate (G6P) in a reaction catalyzed by hex¬  okinase.   The enzyme is represented by a transparent  molecular surface with its backbone shown as a  purple or yellow ribbon, (a) Free hexokinase.  Note its prominent bilobal appearance.   (b) Hexokinase in complex with glucose drawn in  space-filling form with C green and 0 red. In the  enzyme-substrate complex, the two domains  have swung together by a 17° rotation to engulf  the substrate. [Based on an X-ray structure by  Igor Polikarpov, Instituto de Fisica em Sao  Carlos, Brazil. PDBids 11G8 and 3B8A.] O See  Interactive Exercise 15.    A kinase is an enzyme that transfers phosphoryl groups between ATP and  a metabolite (Section 14-2C). The metabolite that serves as the phosphoryl  group acceptor is indicated in the prefix of the kinase name. Hexokinase is a  ubiquitous, relatively nonspecific enzyme that catalyzes the phosphorylation of  hexoses such as D-glucose, D-mannose, and D-fructose. Liver cells also contain  the isozyme glucokinase, which catalyzes the same reaction but which is pri¬  marily involved in maintaining blood glucose levels (Section 22-ID).   The second substrate for hexokinase, as for other kinases, is an Mg 2+ -ATP  complex. In fact, uncomplexed ATP is a potent competitive inhibitor of  hexokinase. Although we do not always explicitly mention the par¬  ticipation of Mg 2 + , it is essential for kinase activity The Mg 2+  shields the negative charges of the ATP s a- and |3- or (3- and y-  phosphate oxygen atoms, making the y-phosphorus atom more  accessible for nucleophilic attack by the C6-OH group of glucose:    Comparison of the X-ray structures of yeast hexokinase and the glucose-  hexokinase complex indicates that glucose induces a large conformational change in  hexokinase (Fig. 15-2). The two lobes that form its active site cleft swing together  by up to 8 A so as to engulf the glucose in a manner that suggests the  closing of jaws. This movement places the ATP close to the — C6H 2 0H  group of glucose and excludes water from the active site (catalysis by prox¬  imity effects; Section 11-3D). If the catalytic and reacting groups were  in the proper position for reaction while the enzyme was in the open  position (Fig. 15-2 d), ATP hydrolysis (i.e., phosphoryl group transfer  to water, which is thermodynamically favored; Fig. 14 -7a) would  almost certainly be the dominant reaction.   Clearly, the substrate-induced conformational change in hex¬  okinase is responsible for the enzymes specificity. In addition,  the active site polarity is reduced by exclusion of water, thereby  facilitating the nucleophilic reaction. Other kinases have the same  deeply clefted structure as hexokinase and undergo conformational  changes on binding their substrates (e.g., protein kinase A, Fig. 13-21;  adenylate kinase, Fig. 14-10).    Reaction 2 of glycolysis is the conversion of G6P to fructose-6-phosphate  (F6P) by phosphoglucose isomerase (PGI).    Since G6P and F6P both exist predominantly in their cyclic forms, the  reaction requires ring opening followed by isomerization and subsequent ring  closure (the interconversions of cyclic and linear forms of hexoses are shown  in Fig. 8-3).   A proposed reaction mechanism for the PGI reaction involves general  acid-base catalysis by the enzyme (Fig. 15-3):   Step 2 An enzymatic acid, probably the 8-amino group of a conserved  Lys residue, catalyzes ring opening.   Step 3 A base, thought to be a His imidazole group, abstracts the acidic  proton from C2 to form a aTenediolate intermediate (the proton  is acidic because it is a to a carbonyl group).   Step 4 The proton is replaced on Cl in an overall proton transfer.  Protons abstracted by bases rapidly exchange with solvent protons.  Nevertheless, Irwin Rose confirmed this step by demonstrating  that [2- 3 H]G6P is occasionally converted to [1- 3 H]F6P by   FIG. 1 5-3 The reaction mechanism of phosphoglucose isomerase. The active site  catalytic residues, BH + and B', are thought to be Lys and His, respectively.    intramolecular proton transfer before the 3 H has had a chance to  exchange with the medium.   Step 5 The ring closes to form the product, which is subsequently released  to yield free enzyme, thereby completing the catalytic cycle.   In Reaction 3 of glycolysis, phosphofructokinase (PFK) phosphorylates F6P  to yield fructose- 1,6-bisphosphate (FBP or F1,6P).    (The product is a ^phosphate rather than a ^phosphate because its two phos¬  phate groups are not attached directly to each other.)   The PFK reaction is similar to the hexokinase reaction. The enzyme cat¬  alyzes the nucleophilic attack by the Cl-OH group of F6P on the electrophilic  y-phosphorus atom of the Mg 2+ —ATP complex.   Phosphofructokinase plays a central role in control of glycolysis because it cat¬  alyzes one of the pathways rate-determining reactions. In many organisms, the  activity of PFK is enhanced allosterically by several substances, including AMP,  and inhibited allosterically by several other substances, including ATP and cit¬  rate. The regulatory properties of PFK are examined in Section 15-4A.   Aldolase catalyzes Reaction 4 of glycolysis, the cleavage of FBP to form the  two trioses glyceraldehyde-3-phosphate (GAP) and dihydroxyacetone phos¬  phate (DHAP):    FIG. 1 5-4 The mechanism of base-catalyzed aldol cleavage. Aldol condensation occurs by the reverse mechanism.    Note that at this point in the pathway, the atom numbering system changes.  Atoms 1, 2, and 3 of glucose become atoms 3, 2, and 1 of DHAP, thus re¬  versing order. Atoms 4, 5, and 6 become atoms 1, 2, and 3 of GAP.   Reaction 4 is an aldol cleavage (retro aldol condensation) whose nonen-  zymatic base-catalyzed mechanism is shown in Fig. 15-4. The enolate inter¬  mediate is stabilized by resonance, as a result of the electron-withdrawing  character of the carbonyl oxygen atom. Note that aldol cleavage between C3  and C4 of FBP requires a carbonyl at C2 and a hydroxyl at C4. Hence, the  “logic” of Reaction 2 in the glycolytic pathway, the isomerization of G6P to  F6P, is clear. Aldol cleavage of G6P would yield products of unequal carbon  chain length, while aldol cleavage of FBP results in two interconvertible C 3 com¬  pounds that can therefore enter a common degradative pathway.   Aldol cleavage is catalyzed by stabilizing its enolate intermediate through  increased electron delocalization. In animals and plants, the reaction occurs as  follows (Fig. 15-5):   Step 2 The FBP carbonyl group reacts with the 8-amino group of the ac¬  tive site Lys to form an iminium cation, that is, a protonated  Schiff base.   Step 3 The C3—C4 bond is cleaved, forming an enamine intermediate  and releasing GAP. The iminium ion is a better electron-withdraw¬  ing group than the oxygen atom of the precursor carbonyl group.  Thus, catalysis occurs because the enamine intermediate (Fig. 15-5,  Step 3) is more stable than the corresponding enolate intermediate  of the base-catalyzed aldol cleavage reaction (Fig. 15-4, Step 2).   Step 4 Protonation and tautomerization of the enamine yields the  iminium cation from the Schiff base.   Step 5 Hydrolysis of the iminium cation releases DHAP and regenerates  the free enzyme.   Only one of the products of the aldol cleavage reaction, GAP, continues along  the glycolytic pathway (Fig. 15-1). However, DHAP and GAP are ketose-  aldose isomers (like F6P and G6P). They are interconverted by an isomeriza¬  tion reaction with an enediol (or enediolate) intermediate. Triose phosphate    FIG. 1 5-5 The enzymatic mechanism of aldolase. See the text for details. O See the Animated Figures.    isomerase (TIM) catalyzes this process in Reaction 5 of glycolysis, the final  reaction of Stage I:    Support for this reaction scheme comes from the use of the transition state  analogs phosphoglycohydroxamate and 2-phosphoglycolate, stable com¬  pounds whose geometry resembles that of the proposed enediol or enediolate  intermediate:    Enzymes catalyze reactions by binding the transition state complex more  tightly than the substrate (Section 11-3E), and, in fact, phosphoglycohydrox¬  amate and 2-phosphoglycolate bind 155- and 100-fold more tightly to TIM  than does either GAP or DHAP.    Glu 165 and His 95 Act as General Acids and Bases. Mechanistic consider¬  ations suggest that the conversion of GAP to the enediol intermediate is catalyzed  by a general base, which abstracts a proton from C2 of GAP, and by a general  acid, which protonates its carbonyl oxygen atom. X-Ray studies reveal that the  Glu 165 side chain of TIM is ideally situated to abstract the C2 proton from  GAP (Fig. 15-6). In fact, the mutagenic replacement of Glu 165 by Asp, which  X-ray studies show withdraws the carboxylate group only ~ 1 A farther away from  the substrate than its position in the wild-type enzyme, reduces TIM s catalytic  activity 1000-fold. X-Ray studies similarly indicate that His 95 is hydrogen  bonded to and hence is properly positioned to protonate GAPs carbonyl oxy¬  gen. The positively charged side chain of Lys 12 is thought to electrostatically  stabilize the negatively charged transition state in the reaction. In  conversion of the enediol intermediate to DHAP, Glu 165 acts a  a general acid to protonate Cl and His 95 acts as a general base  to abstract the proton from the OH group, thereby restoring the  catalytic groups to their initial protonation states.   A Flexible Loop Closes over the Active Site. The comparison  of the X-ray structure of TIM (Fig. 6-30 c) with that of the en-  zyme-2-phosphoglycolate complex reveals that when substrate   FIG. 1 5-6 Ribbon diagram of yeast TIM in complex with its transition  state analog 2-phosphoglycolate. A single subunit of this homodimeric enzyme  is viewed roughly along the axis of its a/|3 barrel. The enzyme’s flexible loop is  cyan, and the side chains of the catalytic Lys, His, and Glu residues are purple,  magenta, and red, respectively. The 2-phosphoglycolate is represented by a  space-filling model colored according to atom type (C green, 0 red, P orange).   [Based on an X-ray structure by Gregory Petsko, Brandeis University. PDBid 2YPI.]    binds to TIM, a conserved 10-residue loop closes over the active site like a  hinged lid, in a movement that involves main chain shifts of >7 A (Fig. 15-6).  A four-residue segment of the loop makes a hydrogen bond with the phos¬  phate group of the substrate. Mutagenic excision of these four residues does  not significantly distort the protein, so substrate binding is not greatly im¬  paired. However, the catalytic power of the mutant enzyme is reduced 10 5 -  fold, and it only weakly binds phosphoglycohydroxamate. Evidently, loop  closure preferentially stabilizes the enzymatic reactions enediol-like transition  state.   Loop closure in the TIM reaction also supplies a striking example of the  so-called stereoelectronic control that enzymes can exert on a reaction. In so¬  lution, the enediol intermediate readily breaks down with the elimination of  the phosphate at C3 to form the toxic compound methylglyoxal (at left). On  the enzymes surface, however, that reaction is prevented because the phos¬  phate group is held by the flexible loop in a position that disfavors phosphate  elimination. In the mutant enzyme lacking the flexible loop, the enediol is  able to escape: ~85% of the enediol intermediate is released into solution  where it rapidly decomposes to methylglyoxal and P z . Thus, the flexible loop  closure assures that substrate is efficiently transformed to product.   FIG. 1 5-7 Schematic diagram of the first  stage of glycolysis. In this series of five  reactions, a hexose is phosphorylated, isomerized,  phosphorylated again, and then cleaved to two  interconvertible triose phosphates. Two ATP are  consumed in the process.    a/p Barrel Enzymes May Have Evolved by Divergent Evolution. TIM was   the first protein found to contain an a/p barrel (also known as a TIM bar¬  rel), a cylinder of eight parallel P strands surrounded by eight parallel a he¬  lices (Fig. 6-30 c). This striking structural motif has since been found in  numerous different proteins, essentially all of which are enzymes (including  the glycolytic enzymes aldolase, enolase, and pyruvate kinase). Intriguingly, the  active sites of nearly all known a/p barrel enzymes are located in the mouth  of the barrel at the end that contains the C-terminal ends of the p strands,  although there is no obvious structural rationale for this. Despite the fact that  few of these proteins exhibit significant sequence similarity, it has been pos¬  tulated that all of them have evolved from a common ancestor (divergent evo¬  lution). However, it has also been argued that the a/p barrel is a particularly  stable arrangement that nature has independently discovered on several occa¬  sions (convergent evolution).   Triose Phosphate Isomerase Is a Catalytically Perfect Enzyme. Jeremy  Knowles demonstrated that TIM has achieved catalytic perfection. By this it  is meant that the rate of the bimolecular reaction between enzyme and sub¬  strate is diffusion controlled, so product formation occurs as rapidly as enzyme  and substrate can collide in solution. Any increase in TIM s catalytic efficiency  therefore would not increase its reaction rate.   GAP and DHAP are interconverted so efficiently that the concentrations  of the two metabolites are maintained at their equilibrium values:  K= [GAP]/[DHAP] = 4.73 X 10“ 2 . At equilibrium, [DHAP] » [GAP],  However, under the steady state conditions in a cell, GAP is consumed in the  succeeding reactions of the glycolytic pathway. As GAP is siphoned off' in this  manner, more DHAP is converted to GAP to maintain the equilibrium ratio. In  effect, DHAP follows GAP into the second stage of glycolysis, so a single path¬  way accounts for the metabolism of both products of the aldolase reaction.   Taking Stock of Glycolysis So Far. At this point in the glycolytic pathway,  one molecule of glucose has been transformed into two molecules of GAP.  This completes the first stage of glycolysis (Fig. 15-7). Note that 2 ATP have  been consumed in generating the phosphorylated intermediates. This energy  investment has not yet paid off, but with a little chemical artistry, the “low-  energy” GAP can be converted to “high-energy” compounds whose free ener¬  gies of hydrolysis can be coupled to ATP synthesis in the second stage of  glycolysis.   Reaction 6 of glycolysis is the oxidation and phosphorylation of GAP by  NAD + and P 7 as catalyzed by glyceraldehyde-3-phosphate dehydrogenase  (GAPDH; Fig. 6-31).    This is the first instance of the chemical artistry alluded to above. In this re¬  action ., aldehyde oxidation ., an exergonic reaction ., drives the synthesis of the “high-  energy” acyl phosphate 1,3-bisphosphogly cerate (1,3-BPG). Recall that acyl  phosphates are compounds with high phosphoryl group-transfer potential  (Section 14-2C).   Several key enzymological experiments have contributed to the elucidation  of the GAPDH reaction mechanism:   1. GAPDH is inactivated by alkylation with stoichiometric amounts of  iodoacetate. The presence of carboxymethylcysteine in the hydrolysate  of the resulting alkylated enzyme (Fig. 15-8^) suggests that GAPDH  has an active site Cys sulfhydryl group.   2. GAPDH quantitatively transfers 3 H from Cl of GAP to NAD +  (Fig. 15-8^), thereby establishing that this reaction occurs via direct  hydride transfer.    FIG. 1 5-8 Reactions that were used to  elucidate the enzymatic mechanism of   GAPDH. (a) The reaction of iodoacetate with  an active site Cys residue, (b) Quantitative  tritium transfer from substrate to NAD + .   (c) The enzyme-catalyzed exchange of 32 P  from phosphate to acetyl phosphate.    3. GAPDH catalyzes exchange of 32 P between P, and the product analog  acetyl phosphate (Fig. 15-8c). Such isotope exchange reactions are in¬  dicative of an acyl—enzyme intermediate; that is, the acetyl group forms  a covalent complex with the enzyme, similar to the acyl—enzyme inter¬  mediate in the serine protease reaction mechanism (Section 11-5C).   David Trentham has proposed a mechanism for GAPDH based on this  information and the results of kinetic studies (Fig. 15-9):   Step 2 The essential sulfhydryl group, acting as a nucleophile, attacks the  aldehyde to form a thiohemiacetal.   Step 3 The thiohemiacetal undergoes oxidation to an acyl thioester by  direct hydride transfer to NAD + . This intermediate, which has  been isolated, has a large free energy of hydrolysis. Thus, the en-  ergy of aldehyde oxidation has not been dissipated but has been con¬  served through the synthesis of the thioester and the reduction of  NAD + to NADH.    FIG. 1 5-9 The enzymatic mechanism of GAPDH. See the text for details. O See the Animated Figures.   Step 5 The thioester intermediate undergoes nucleophilic attack by P z to  form the “high-energy” mixed anhydride 1,3-BPG product (an  acyl phosphate), which then dissociates from the enzyme followed  by replacement of NADH by another molecule of NAD + to re¬  generate the active enzyme.   Reaction 7 of the glycolytic pathway yields ATP together with 3-phospho-  glycerate (3PG) in a reaction catalyzed by phosphoglycerate kinase (PGK).    (Note that this enzyme is called a “kinase” because the reverse reaction is phos-  phoryl group transfer from ATP to 3PG.)   PGK (Fig. 15-10) is conspicuously bilobal in appearance. The  Mg 2+ -ADP-binding site is located on one domain, ~10 A from the 1,3-BPG-  binding site, which is on the other domain. Physical measurements suggest  that, on substrate binding, the two domains of PGK swing together to per¬  mit the substrates to react in a water-free environment, as occurs in hexoki-  nase (Section 15-2A). Indeed, the appearance of PGK is remarkably similar to  that of hexokinase (Fig. 15-2), although the structures of the proteins are oth¬  erwise unrelated.   The GAPDH and PGK Reactions Are Coupled. As described in Section 14-2B,  a slightly unfavorable reaction can be coupled to a highly favorable reaction  so that both reactions proceed in the forward direction. In the case of the sixth   FIG. 15-10 X-Ray structure of yeast  phosphoglycerate kinase (PGK) in complex  with 3PG and ATP. The enzyme is represented  by a transparent molecular surface with its  embedded ribbon diagram colored with the  N-terminal domain yellow and the C-terminal  domain purple. The Mg 2+ -ATP and 3PG are  drawn in space-filling form colored according to  atom type (ATP C green, 3PG C cyan, N blue, 0  red, P orange, and Mg 2+ pale green). Note the  similar overall appearance of PGK and  hexokinase (Fig. 15-2). [Based on an X-ray  structure by Herman Watson, University of  Bristol, U.K. PDBid 3PGK.]   FIG. 1 5-1 1 The active site region of  yeast phosphoglycerate mutase (dephospho  form). The substrate, 3PG, which is drawn in  ball-and-stick form with C green, 0 red, and P  orange, binds to an ionic pocket whose side  chains are drawn in stick form with C green, N  blue, and 0 red. His 8 is phosphorylated in the  active enzyme. [Based on an X-ray  structure by Jennifer Littlechild, University of  Exeter, U.K. PD Bid 1QHFJ   GAP + P, + NAD + + ADP -> 3PG + NADH + ATP   Although the GAPDH reaction is endergonic, the strongly exergonic nature  of the transfer of a phosphoryl group from 1,3-BPG to ADP makes the over¬  all synthesis of NADH and ATP from GAP, P z , NAD + , and ADP favorable.  This production of ATP, which does not involve 0 2 > is an example of substrate-  level phosphorylation. The subsequent oxidation of the NADH produced in this  reaction by 0 2 generates additional ATP by oxidative phosphorylation, as we  will see in Section 18-3.   In Reaction 8 of glycolysis, 3PG is converted to 2-phosphoglycerate (2PG)  by phosphoglycerate mutase (PGM):    A mutase catalyzes the transfer of a functional group from one position to an¬  other on a molecule. This more or less energetically neutral reaction is necessary  preparation for the next reaction in glycolysis, which generates a “high-energy”  phosphoryl compound.   At first sight, the reaction catalyzed by phosphoglycerate appears to be a  simple intramolecular phosphoryl group transfer. This is not the case, however.  The active enzyme has a phosphoryl group at its active site, attached to His 8.    The phosphoryl group is transferred to the substrate to form a bisphospho in¬  termediate. This intermediate then rephosphorylates the enzyme to form the  product and regenerate the active phosphoenzyme. The enzymes X-ray struc¬  ture shows the proximity of His 8 to the substrate (Fig. 15-11).   Step 1 3PG binds to the phosphoenzyme in which His 8 is phosphorylated.   Step 2 The enzymes phosphoryl group is transferred to the substrate, result¬  ing in an intermediate 2,3-bisphosphoglycerate—enzyme complex.   Step 3 The enzyme is rephosphorylated by the substrate’s 3-phospho group.    FIG. 15-12 A proposed reaction mechanism for phosphoglycerate mutase. The active  form of the enzyme contains a phospho-His residue at the active site. See the text for details.    The phosphoryl group of 3PG therefore ends up on C2 of the next 3PG to  undergo reaction.   Occasionally, 2,3-bisphosphoglycerate (2,3-BPG) formed in Step 2 of the  reaction dissociates from the dephosphoenzyme, leaving it in an inactive form.  Trace amounts of 2,3-BPG must therefore always be available to regenerate the  active phosphoenzyme by the reverse reaction. 2,3-BPG also specifically binds  to deoxyhemoglobin, thereby decreasing its oxygen affinity (Section 7-ID).  Consequently, erythrocytes require much more 2,3-BPG (5 mM) than the trace  amounts that are used to prime phosphoglycerate mutase (Box 15-2).   In Reaction 9 of glycolysis, 2PG is dehydrated to phosphoenolpyruvate  (PEP) in a reaction catalyzed by enolase:    Box 15-2 Perspectives Synthesis of 2,3-Bisphosphoglycerate in Erythrocytes and  in Biochemistry Its Effect on the Oxygen Carrying Capacity of the Blood    The specific binding of 2,3-bisphosphoglycerate (2,3-BPG) to  deoxyhemoglobin decreases the oxygen affinity of hemoglobin  (Section 7-ID). Erythrocytes synthesize and degrade 2,3-BPG by  a detour from the glycolytic pathway.    Bisphosphoglycerate mutase catalyzes the transfer of a phos-  phoryl group from Cl to C2 of 1,3-BPG. The resulting 2,3-BPG is  hydrolyzed to 3PG by 2,3-bisphosphoglycerate phosphatase.   The level of available 2,3-BPG regulates hemoglobin’s oxygen  affinity. Consequently, inherited defects of glycolysis in erythro¬  cytes alter the ability of the blood to carry oxygen as is indicated  by the oxygen-saturation curve of its hemoglobin.   For example, in hexokinase-deficient erythrocytes, the concen¬  trations of all the glycolytic intermediates are low (since hexoki-  nase catalyzes the first step of glycolysis), thereby resulting in a  diminished 2,3-BPG concentration and an increased hemoglobin  oxygen affinity (green curve). Conversely, a deficiency in pyruvate   kinase (which catalyzes the final reaction of glycolysis; Fig. 15-1)  decreases hemoglobin’s oxygen affinity (purple curve ) through an  increase in 2,3-BPG concentration resulting from this blockade.  Thus, although erythrocytes, which lack nuclei and other  organelles, have only a minimal metabolism, this metabolism is  physiologically significant.   [Oxygen-saturation curves after Delivoria-Papadopoulos, M., Oski, F.A.,  and Gottlieb, A.J., Science 165, 601 (1969).]    The enzyme forms a complex with a divalent cation such as Mg 2+ before the  substrate binds. Fluoride ion inhibits glycolysis by blocking enolase activity  (F was one of the metabolic inhibitors used in elucidating the glycolytic path¬  way). In the presence of P z , F blocks substrate binding to enolase by form¬  ing a bound complex with Mg 2+ at the enzymes active site. Enolases  substrate, 2PG, therefore builds up, and, through the action of PGM, 3PG  also builds up.   In Reaction 10 of glycolysis, its final reaction, pyruvate kinase (PK) couples  the free energy of PEP cleavage to the synthesis of ATP during the formation  of pyruvate:    FIG. 15-13 The mechanism of the reaction catalyzed by pyruvate kinase. See the text for details.    The PK reaction, which requires both monovalent (K + ) and divalent (Mg 2+ )  cations, occurs as follows (Fig. 15-13):   Step 1 A (Tphosphoryl oxygen of ADP nucleophilically attacks the PEP  phosphorus atom, thereby displacing enolpyruvate and forming  ATP   The PK reaction is highly exergonic, supplying more than enough free energy  to drive ATP synthesis (another example of substrate-level phosphorylation). At  this point, the “logic” of the enolase reaction becomes clear. The standard free  energy of hydrolysis of 2PG is only —16 kj • mol x , which is insufficient to  drive ATP synthesis from ADP (AG°' = 30.5 kj • mol x ). However, the  dehydration of 2PG results in the formation of a “high-energy” compound  capable of such synthesis. The high phosphoryl group-transfer potential of PEP re¬  flects the large release of free energy on converting the product enolpyruvate to its  keto tautomer. Consider the hydrolysis of PEP as a two-step reaction (Fig. 15-14).   value is much more negative than that required  to provide the AG°' for ATP synthesis from ADP  and P/.    FIG. 1 5-1 5 Schematic diagram of the  second stage of glycolysis. In this series of  five reactions, GAP undergoes phosphorylation  and oxidation, followed by molecular rearrange¬  ments so that both phosphoryl groups have  sufficient free energy to be transferred to ADP  to produce ATP Two molecules of GAP are  converted to pyruvate for every molecule of  glucose that enters Stage I of glycolysis.    The tautomerization step supplies considerably more free energy than the phos¬  phoryl group transfer step.   Assessing Stage II of Glycolysis. The energy investment of the first stage  of glycolysis (2 ATP consumed) is doubly repaid in the second stage of gly¬  colysis because two phosphorylated C 3 units are transformed to two pyruvates  with the coupled synthesis of 4 ATP. This process is shown schematically in   2 pyruvate + 2 NADH + 2 ATP + 2 H 2 0 + 4 H +   1. ATP. The initial investment of 2 ATP per glucose in Stage I and the  subsequent generation of 4 ATP by substrate-level phosphorylation  (two for each GAP that proceeds through Stage II) gives a net yield of  2 ATP per glucose. In some tissues and organisms for which glucose  is the primary metabolic fuel, ATP produced by glycolysis satisfies most  of the cells energy needs.   2. NADH. During its catabolism by the glycolytic pathway, glucose is ox¬  idized to the extent that two NAD + are reduced to two NADH. As  described in Section 14-3C, reduced coenzymes such as NADH rep¬  resent a source of free energy than can be recovered by their subse¬  quent oxidation. Under aerobic conditions, electrons pass from reduced  coenzymes through a series of electron carriers to the final oxidizing  agent, 0 2 , in a process known as electron transport (Section 18-2).    The free energy of electron transport drives the synthesis of ATP from  ADP (oxidative phosphorylation; Section 18-3). In aerobic organisms,  this sequence of events also serves to regenerate oxidized NAD + that  can participate in further rounds of catalysis mediated by GAPDH.  Under anaerobic conditions, NADH must be reoxidized by other  means in order to keep the glycolytic pathway supplied with NAD +  (Section 15-3).   3. Pyruvate . The two pyruvate molecules produced through the partial  oxidation of each glucose are still relatively reduced molecules. Under  aerobic conditions, complete oxidation of the pyruvate carbon atoms  to C0 2 is mediated by the citric acid cycle (Chapter 17). The energy  released in that process drives the synthesis of much more ATP than  is generated by the limited oxidation of glucose by the glycolytic path¬  way alone. In anaerobic metabolism, pyruvate is metabolized to a lesser  extent to regenerate NAD + , as we will see in the following section.    • NADH, a substrate for the GAPDH reaction, must be reoxidized for glycolysis to  continue.   • In muscle, pyruvate is reduced to lactate to regenerate NAD + .   • Yeast decarboxylates pyruvate to produce C0 2 and ethanol, in a process that  requires the cofactor TPP.    The three common metabolic fates of pyruvate produced by glycolysis are out¬  lined in Fig. 15-16.   • Write the reactions of glycolysis, showing  the structural formulas of the intermediates  and the names of the enzymes that cat¬  alyze the reactions.   • Summarize the types of catalytic mecha¬  nisms involved. Do any glycolytic enzymes  require cofactors?   • Explain the chemical logic of converting  glucose to fructose before aldolase splits  the sugar in two.   • Why is triose phosphate isomerase con¬  sidered to be catalytically perfect?   • How does phosphorylation differ between  the kinase-catalyzed reactions and the  reaction catalyzed by GAPDH?   • What compounds with high phosphate  group-transfer potential are synthesized  during glycolysis?   • Explain the chemical logic of dehydrating  2-phosphoglycerate before its phosphoryl  group is transferred.   • Explain how chemical coupling of ender-  gonic and exergonic reactions is used to  generate ATP during glycolysis.   • Which products of glycolysis are reduced  molecules that the cell can oxidize to  recover free energy?    1. Under aerobic conditions , the pyruvate is completely oxidized via the cit¬  ric acid cycle to C0 2 and H 2 0.   2. Under anaerobic conditions , pyruvate must be converted to a reduced end  product in order to reoxidize the NADH produced by the GAPDH reac¬  tion. This occurs in two ways:    (a) Under anaerobic conditions in muscle, pyruvate is reduced to  lactate to regenerate NAD + in a process known as homolactic  fermentation (a fermentation is an anaerobic biological reaction  process).   (b) In yeast, pyruvate is decarboxylated to yield C0 2 and acetalde¬  hyde, which is then reduced by NADH to yield NAD + and  ethanol. This process is known as alcoholic fermentation.   Thus, in aerobic glycolysis, NADH acts as a “high-energy” compound, whereas  in anaerobic glycolysis, its free energy of oxidation is dissipated as heat.    FIG. 1 5-1 6 Metabolic fate of pyruvate. Under aerobic  conditions (left), the pyruvate carbons are oxidized to C0 2 by  the citric acid cycle and the electrons are eventually trans¬  ferred to 0 2 to yield H 2 0 in oxidative phosphorylation. Under  anaerobic conditions in muscle, pyruvate is reversibly con¬  verted to lactate (middle), whereas in yeast, it is converted to  C0 2 and ethanol (right).   In muscle, during vigorous activity, when the demand for ATP is high and  oxygen is in short supply, ATP is largely synthesized via anaerobic glycolysis,  which rapidly generates ATP, rather than through the slower process of oxida¬  tive phosphorylation. Under these conditions, lactate dehydrogenase (LDH)  catalyzes the oxidation of NADH by pyruvate to yield NAD + and lactate.    This reaction is often classified as Reaction 11 of glycolysis. The lactate dehy¬  drogenase reaction is freely reversible, so pyruvate and lactate concentrations arc  readily equilibrated.   In the proposed mechanism for pyruvate reduction by LDH, a hydride  ion is stereospecifically transferred from C4 of NADH to C2 of pyruvate with  concomitant transfer of a proton from the imidazolium moiety of His 195:    Both His 195 and Arg 171 interact electrostatically with the substrate to ori¬  ent pyruvate (or lactate, in the reverse reaction) in the enzyme active site.  The overall process of anaerobic glycolysis in muscle can be represented as   Glucose + 2 ADP + 2 P z —» 2 lactate + 2 ATP + 2 H 2 0 + 2 H +   Lactate represents a sort of dead end for anaerobic glucose metabolism. The  lactate can either be exported from the cell or converted back to pyruvate.  Much of the lactate produced in skeletal muscle cells is carried by the blood  to the liver, where it is used to synthesize glucose (Section 22-IF).   Contrary to widely held belief, it is not lactate buildup in the muscle per  se that causes muscle fatigue and soreness but the accumulation of glycolyti-  cally generated acid (muscles can maintain their workload in the presence of  high lactate concentrations if the pH is kept constant).   Under anaerobic conditions in yeast, NAD + for glycolysis is regenerated in a  process that has been valued for thousands of years: the conversion of pyru¬  vate to ethanol and C0 2 . Ethanol is, of course, the active ingredient of wine  and spirits; C0 2 so produced leavens bread.   Yeast (Fig. 15-17) produces ethanol and C0 2 via two consecutive reac¬  tions (Fig. 15-18):    1. The decarboxylation of pyruvate to form acetaldehyde and C0 2 as cat¬  alyzed by pyruvate decarboxylase (an enzyme not present in animals).   2. The reduction of acetaldehyde to ethanol by NADH as catalyzed  by alcohol dehydrogenase (Section 11-1C), thereby regenerating  NAD + for use in the GAPDH reaction of glycolysis.   TPP Is an Essential Cofactor of Pyruvate Decarboxylase. Pyruvate decar¬  boxylase contains the coenzyme thiamine pyrophosphate (TPP; also called  thiamin diphosphate, ThDP):   TPP, which is synthesized from thiamine (vitamin binds tightly but non-  covalently to pyruvate decarboxylase (Fig. 15-19).   The enzyme uses TPP because uncatalyzed decarboxylation of an a-keto  acid such as pyruvate requires the buildup of negative charge on the carbonyl  carbon atom in the transition state, an unstable situation:   FIG. 1 5-1 9 TPP binding to pyruvate  decarboxylase from Saccharomyces uvarum  (brewer’s yeast). The TPP and the side chain  of Glu 51 are shown in stick form with C green,  N blue, 0 red, S yellow, and P orange. The TPP  binds in a cavity situated between the dimer’s  two subunits (cyan and magenta) where it  hydrogen bonds to Glu 51. [Based on an X-ray  structure by William Furey and Martin Sax,  Veterans Administration Medical Center and  University of Pittsburgh. PDBid 1 PYD.] Q See  Interactive Exercise 17.   Chapter 15 Glucose Catabolism This transition state can be stabilized by delocalizing the developing negative   TP P’s catalytically active functional group is the thiazolium ring. The  C2-H atom of this group is relatively acidic because of the adjacent positively  charged quaternary nitrogen atom, which electrostatically stabilizes the carban-  ion formed when the proton dissociates. This dipolar carbanion (or ylid) is  the active form of the coenzyme. Pyruvate decarboxylase operates as follows  (Fig. 15-20):   Step 1 The ylid form of TPP, a nucleophile, attacks the carbonyl carbon  of pyruvate.   Step 2 C0 2 departs, generating a resonance-stabilized carbanion adduct in  which the thiazolium ring of the coenzyme acts as an electron sink.   Step 4 The TPP ylid is eliminated to form acetaldehyde and regenerate  the active enzyme.   This mechanism has been corroborated by the isolation of the hydroxy-  ethylthiamine pyrophosphate intermediate.   T Vitamin Deficiency Causes Beriberi. The ability of TPP s thiazolium  ring to add to carbonyl groups and act as an electron sink makes it the    FIG. 1 5-20 The reaction mechanism of pyruvate decarboxylase. See the text for details.   occur in all organisms, not just yeast. Consequently, thiamine (vitamin Bj), which  is neither synthesized nor stored in significant quantities by the tissues of most  vertebrates, is required in their diets. Thiamine deficiency in humans results in  an ultimately fatal condition known as beriberi (Singhalese for weakness) that  is characterized by neurological disturbances causing pain, paralysis, and atro¬  phy (wasting) of the limbs and/or edema (accumulation of fluid in tissues and  body cavities). Beriberi was particularly prevalent in the late eighteenth and  early nineteenth centuries in the rice-consuming areas of Asia after the intro¬  duction of steam-powered milling machines that polished the rice grains to re¬  move their coarse but thiamine-containing outer layers (the previously used  milling procedures were less efficient and hence left sufficient thiamine on the  grains). Parboiling rice before milling, a process common in India, causes the  rice kernels to absorb nutrients from their outer layers, thereby decreasing the  incidence of beriberi. Once thiamine deficiency was recognized as the cause of  beriberi, enrichment procedures were instituted so that today it has ceased to  be a problem except in areas undergoing famine. However, beriberi occasion¬  ally develops in alcoholics due to their penchant for drinking but not eating.   Reduction of Acetaldehyde and Regeneration of NAD + . Yeast alcohol de¬  hydrogenase (YADH), the enzyme that converts acetaldehyde to ethanol, is a  tetramer, each subunit of which binds one Zn 2+ ion. The Zn 2+ polarizes the  carbonyl group of acetaldehyde to stabilize the developing negative charge in  the transition state of the reaction.    This facilitates the stereospecific transfer of a hydrogen from NADH to ac¬  etaldehyde.   Mammalian liver alcohol dehydrogenase (LADH) metabolizes the alco¬  hols anaerobically produced by the intestinal flora as well as those from exter¬  nal sources (the direction of the alcohol dehydrogenase reaction varies with  the relative concentrations of ethanol and acetaldehyde). Mammalian LADH  is a dimer with significant amino acid sequence similarity to YADH, although  LADH subunits each contain a second Zn 2+ ion that presumably has a struc¬  tural role.   Thermodynamics permits us to dissect the process of fermentation into its  component parts and to account for the free energy changes that occur. This  enables us to calculate the efficiency with which the free energy of glucose  catabolism is used in the synthesis of ATP. For homolactic fermentation,   Each of these processes is coupled to the net formation of 2 ATP, which re¬  quires AG°' = +61 kj • mol -1 of glucose consumed. Dividing AG°' of ATP    Skeletal muscle consists of both slow-twitch (Type I) and fast-  twitch (Type II) fibers. Fast-twitch fibers, so called because they  predominate in muscles capable of short bursts of rapid activity,  are nearly devoid of mitochondria (where oxidative phosphorylation  occurs). Consequently, they must obtain nearly all of their ATP  through anaerobic glycolysis, for  which they have a particularly large  capacity. Muscles designed to con¬  tract slowly and steadily, in contrast,  are enriched in slow-twitch fibers  that are rich in mitochondria and  obtain most of their ATP through  oxidative phosphorylation.   Fast- and slow-twitch fibers  were originally known as white and  red fibers, respectively, because  otherwise pale-colored muscle  tissue, when enriched with mito¬  chondria, takes on the red color  characteristic of their heme-    containing cytochromes. However, fiber color is an imperfect  indictor of muscle physiology.   In a familiar example, the flight muscles of migratory birds such  as ducks and geese, which need a continuous energy supply, are  rich in slow-twitch fibers. Therefore, these birds have dark breast   meat. In contrast, the flight muscles  of less ambitious fliers, such as  chickens and turkeys, which are  used only for short bursts (often to  escape danger), consist mainly of  fast-twitch fibers that form white  meat. In humans, the muscles of  sprinters are relatively rich in fast-  twitch fibers, whereas distance  runners have a greater proportion of  slow-twitch fibers (although their  muscles have the same color).   • Compare homolactic and alcoholic fermen¬  tation in terms of the products and the  cofactors required.   • Compare the ATP yields and rates of ATP  production for anaerobic and aerobic  degradation of glucose.    formation by that of lactate formation indicates that homolactic fermentation  is 31% “efficient”; that is, 31% of the free energy released by the process un¬  der standard biochemical conditions is sequestered in the form of ATP. The  rest is dissipated as heat, thereby making the process irreversible. Likewise, al¬  coholic fermentation is 26% efficient under biochemical standard state con¬  ditions. Under physiological conditions, where the concentrations of reactants and  products differ from those of the standard state, these reactions have thermody¬  namic efficiencies of >50%.   Anaerobic fermentation uses glucose in a profligate manner compared to  oxidative phosphorylation: Fermentation results in the production of 2 ATP  per glucose, whereas oxidative phosphorylation yields up to 32 ATP per glu¬  cose (Section 18-3C). This accounts for Pasteurs observation that yeast con¬  sume far more sugar when growing anaerobically than when growing  aerobically (the Pasteur effect). However, the rate of ATP production by anaer¬  obic glycolysis can be up to 100 times faster than that of oxidative phosphoryla¬  tion. Consequently, when tissues such as muscle are rapidly consuming ATP, they  regenerate it almost entirely by anaerobic glycolysis. (Homolactic fermentation  does not really “waste” glucose since the lactate can be aerobically reconverted  to glucose by the liver; Section 22-IF.) Certain muscles are specialized for the  rapid production of ATP by glycolysis (Box 15-3).    • Enzymes that function with large negative free energy changes are candidates for  flux-control points.   • Phosphofructokinase, the major regulatory point for glycolysis in muscle, is  allosterically inhibited by ATP and activated by AMP and ADR   • Substrate cycling allows the rate of glycolysis to respond rapidly to changing needs.    Under steady state conditions, glycolysis operates continuously in most tissues,  although the glycolytic flux must vary to meet the needs of the organism.   Elucidation of the flux control mechanisms of a given pathway, such as gly¬  colysis, commonly involves three steps:    1. Identification of the rate-determining step(s) of the pathway by meas¬  uring the in vivo AG for each reaction. Enzymes that operate far from  equilibrium are potential control points (Section 14-ID).   2. In vitro identification of allosteric modifiers of the enzymes catalyzing  the rate-determining reactions. The mechanisms by which these com¬  pounds act are determined from their effects on the enzymes’ kinetics.   3. Measurement of the in vivo levels of the proposed regulators under var¬  ious conditions to establish whether the concentration changes are con¬  sistent with the proposed control mechanism.   Let us examine the thermodynamics of glycolysis in muscle tissue with an  eye toward understanding its control mechanisms (keep in mind that differ¬  ent tissues control glycolysis in different ways). Table 15-1 lists the standard  free energy changes (AG°') and the actual physiological free energy change  (AG) associated with each reaction in the pathway. It is important to realize  that the free energy changes associated with the reactions under standard con¬  ditions may differ dramatically from the actual values in vivo.   Only three reactions of glycolysis, those catalyzed by hexokinase, phosphofruc-  tokinase, and pyruvate kinase, function with large negative free energy changes in  heart muscle under physiological conditions (Fig. 15 - 21 ). These nonequilibrium  reactions of glycolysis are candidates for flux-control points. The other glycolytic  reactions function near equilibrium: Their forward and reverse rates are much  faster than the actual flux through the pathway. Consequently, these equilib¬  rium reactions are very sensitive to changes in the concentration of pathway  intermediates and readily accommodate changes in flux generated at the rate¬  determining step(s) of the pathway.   FIG. 1 5-21 Diagram of free energy  changes in glycolysis. This “waterfall” diagram  illustrates the actual free energy changes for the  glycolytic reactions in heart muscle (Table 15-1).  Reactions 1, 3, and 10 are irreversible. The other  reactions operate near equilibrium and can  mediate flux in either direction.    A Phosphofructokinase Is the Major Flux-Controlling Enzyme of  Glycolysis in Muscle   In vitro studies of hexokinase, phosphofructokinase, and pyruvate kinase indi¬  cate that each is controlled by a variety of compounds. Yet when the G6P  source for glycolysis is glycogen, rather than glucose, as is often the case in  skeletal muscle, the hexokinase reaction is not required (Section 16-1). Pyruvate  kinase catalyzes the last reaction of glycolysis and is therefore unlikely to be the  primary point for regulating flux through the entire pathway. Evidently, PFK,  an elaborately regulated enzyme functioning far from equilibrium, is the major  control point for glycolysis in muscle under most conditions.   FIG. 1 5-22 X-Ray structure of PFK from E. coli. Two subunits of the tetrameric  enzyme are shown in ribbon form (the other two subunits, which have been omitted for  clarity, are related to those shown by a twofold rotation about the vertical axis). Each  subunit binds its products, FBP ( near the center of each subunit) and Mg 2+ -ADP ( lower  right and upper left), together with its activator Mg 2+ -ADP ( upper right and lower left, in the  rear), all in space-filling form with atoms colored according to type (ADP C green, FBP C  cyan, N blue, 0 red, P orange, and Mg purple). Note the close proximity of the product  ADP’s |3-phosphate group to the phosphoryl group at FBP’s 1-position, the group that  PFK transfers from ATP to F6P. [Based on an X-ray structure by Philip Evans,  Cambridge University. PDBid 1 PFK.] O See Kinemage Exercise 13-1.    PFK (Fig. 15-22) is a tetrameric enzyme with two conformational  states, R and T, that are in equilibrium. ATP is both a substrate and  an allosteric inhibitor of phosphofructokinase. Other compounds, in¬  cluding ADP, AMP, and fructose-2,6-bisphosphate (F2,6P), reverse  the inhibitory effects of ATP and are therefore activators of PFK. Each  PFK subunit has two binding sites for ATP: a substrate site and an  inhibitor site. The substrate site binds ATP equally well in either con¬  formation, but the inhibitor site binds ATP almost exclusively in the T  state. The other substrate of PFK, F6P, preferentially binds to the R state.  Consequently, at high concentrations, ATP acts as an allosteric inhibitor of  PFK by binding to the T state, thereby shifting the T R equilibrium in  favor of the T state and thus decreasing PFKs affinity for F6P (this is similar  to the action of 2,3-BPG in decreasing the affinity of hemoglobin for 0 2 ;  Section 7-ID).   In graphical terms, high concentrations of ATP shift the curve of PFK  activity versus [F6P] to the right and make it even more sigmoidal (cooper¬  ative) (Fig. 15-23). For example, when [F6P] = 0.5 mM (the dashed line in  Fig. 15-23), the enzyme is nearly maximally active, but in the presence of  1 mM ATP, the activity drops to 15% of its original level, a nearly sevenfold  decrease. An activator such as AMP or ADP counters the effect of ATP by  binding to R-state PFK, thereby shifting the T R equilibrium toward the  R state. (Actually, the most potent allosteric effector of PFK is F2,6P, which  we discuss in Section 16-4C.)   The X-ray structures of PFK from several organisms have been determined in  both the R and the T states by Philip Evans. The R state of PFK is stabilized  by the binding of its substrate F6P. In the R state of Bacillus stearothermophilus  PFK, the side chain of Arg 162 forms an ion pair with the phosphoryl group  of an F6P bound in the active site of another subunit (Fig. 15-24). However,  Arg 162 is located at the end of a helical turn that unwinds on transition to  the T state. The positively charged side chain of Arg 162 thereby swings away    FIG. 1 5-23 PFK activity versus F6P  concentration. The various conditions are as  follows: purple, no inhibitors or activators; green,   1 mM ATP; and red, 1 mM ATP + 0.1 mM AMP.  [After data from Mansour, T.E. and Ahlfors, C.E.,   FIG. 15-24 Allosteric  changes in PFK from Bacillus  stearothermophilus. Segments  of the T state (blue) are superimposed  on segments of the R state (pink) that  undergo a large conformational rearrangement  on the T^R allosteric transition (indicated by the arrows).   Residues of the R-state structure are marked by a prime. Note  that in the R state, Arg 162' forms an attractive ionic interaction with  F6P', whereas in the T state, F6P' is repelled by Glu 161. Also shown are  bound ligands: the nonphysiological inhibitor 2-phosphoglycolate (PGC; a PEP  analog) for the T state, and the cooperative substrate F6P and the activator ADP  for the R state. [After Schirmer, T. and Evans, P.R., Nature 343, 142 (1990). PDBids  4PFK and 6PFK.] Q See Kinemage Exercise 13-2.    and is replaced by the negatively charged side chain of Glu 161. As a conse¬  quence, the doubly negative phosphoryl group of F6P has a greatly diminished  affinity for the T-state enzyme. The unwinding of this helical turn, which is  obligatory for the R —> T transition, is prevented by the binding of the acti¬  vator ADP to its effector site on the enzyme. Presumably, ATP can bind to this  site only when the helical turn is in its unwound conformation (the T state).   AMP Overcomes the ATP Inhibition of PFK. Direct allosteric regulation of  PFK by ATP may at first appear to be the means by which glycolytic flux is  controlled. After all, when [ATP] is high as a result of low metabolic demand,  PFK is inhibited and flux through glycolysis is low; conversely when [ATP] is  low, flux through the pathway is high and ATP is synthesized to replenish the  pool. Consideration of the physiological variation in ATP concentration, how¬  ever, indicates that the situation must be more complex. The metabolic flux  through glycolysis may vary by 100-fold or more, depending on the metabolic  demand for ATP. However, measurements of [ATP] in vivo at various levels of  metabolic activity indicate that [ATP] varies <10% between rest and vigorous  exertion. Yet there is no known allosteric mechanism that can account for a  100-fold change in flux of a nonequilibrium reaction with only a 10% change  in effector concentration. Thus, some other mechanism(s) must be responsi¬  ble for controlling glycolytic flux.   The inhibition of PFK by ATP is relieved by AMP as well as ADP. This  results from AMPs preferential binding to the R state of PFK. If a PFK so¬  lution containing 1 mM ATP and 0.5 mM F6P is brought to 0.1 mM in  AMP, the activity of PFK rises from 15 to 50% of its maximal activity, a three¬  fold increase (Fig. 15-23).   The [ATP] decreases by only 10% in going from a resting state to one of  vigorous activity because it is buffered by the action of two enzymes: creatine    kinase and adenylate kinase (Section 14-2C). Adenylate kinase catalyzes the  reaction   which rapidly equilibrates the ADP resulting from ATP hydrolysis in muscle  contraction with ATP and AMP   In muscle, [ATP] is —50 times greater than [AMP] and ~10 times greater  than [ADP]. Consequently, a change in [ATP] from, for example , 1 to 0.9 mM ’  a 10% decrease , can result in a 100% increase in [ADP] (from 0.1 to 0.2 mM)  as a result of the adenylate kinase reaction , and a > 400% increase in [AMP]  (from 0.02 to ~0.1 mM). Therefore, a metabolic signal consisting of a decrease  in [ATP] too small to relieve PFK inhibition is amplified significantly by the  adenylate kinase reaction, which increases [AMP] by an amount that produces  a much larger increase in PFK activity   Even a finely tuned allosteric mechanism like that of PFK can account for  only a fraction of the 100-fold alterations in glycolytic flux. Additional con¬  trol may be achieved by substrate cycling. Recall from Section 14-ID that only  a near-equilibrium reaction can undergo large changes in flux because, in a  near-equilibrium reaction, zy — v r ~ 0 (where zy and v r are the forward and  reverse reaction rates) and hence a small change in zy will result in a large  fractional change in zy — v r . However, this is not the case for the PFK reac¬  tion because, for such nonequilibrium reactions, v r is negligible.   Nevertheless, such equilibrium-like conditions may be imposed on a nonequi¬  librium reaction if a second enzyme (or series of enzymes) catalyzes the regenera¬  tion of its substrate from its product in a thermodynamically favorable manner.  This can be diagrammed as    Since two different enzymes catalyze the forward (f) and reverse (r) reactions,  zy and v r may be independently varied and v r is no longer negligible com¬  pared to zy. Note that the forward process (e.g., formation of FBP from F6P)  and the reverse process (e.g., breakdown of FBP to F6P) must be carried out  by different enzymes since the laws of thermodynamics would otherwise be  violated (i.e., for a single reaction, the forward and reverse reactions cannot  simultaneously be favorable).   is highly exergonic (AG = — 25.9 kj • mol 1 ). Consequently, the back reac¬  tion has a negligible rate compared to the forward reaction. Fructose-1,6-  bisphosphatase (FBPase), however, which is present in many mammalian  tissues (and which is an essential enzyme in gluconeogenesis; Section 16-4B),  catalyzes the exergonic hydrolysis of FBP (AG = —8.6 kj • mol 1 ).   Note that the combined reactions catalyzed by PFK and FBPase result in net  ATP hydrolysis:    FIG. 1 5-25 Substrate cycling in the  regulation of PFK. (a) In resting muscle, both  enzymes in the F6P/FBP substrate cycle are  active, and glycolytic flux is low. (b) In active  muscle, PFK activity increases while FBPase  activity decreases. This dramatically increases the  flux through PFK and therefore results in high  glycolytic flux.    Such a set of opposing reactions (Section 14-IE) is known as a substrate  cycle because it cycles a substrate to an intermediate and back again. When  this set of reactions was discovered, it was referred to as a futile cycle since  its net result seemed to be the useless consumption of ATP.   Eric Newsholme has proposed that substrate cycles are not at all “futile” but,  rather, have a regulatory function. The combined effects of allosteric effectors on  the opposing reactions of a substrate cycle can produce a much greater fractional ef¬  fect on pathway flux (vp~ v r ) than is possible through allosteric regulation of a sin¬  gle enzyme. For example, the allosteric effector F2,6P activates the PFK reaction  while inhibiting the FBPase reaction (this regulatory mechanism is important  for balancing glycolysis and gluconeogenesis in liver cells; Section 16-4C).   Substrate cycling does not increase the maximum flux through a path¬  way. On the contrary, it functions to decrease the minimum flux. In a sense,  the substrate is put into a “holding pattern.” In the PFK/FBPase example  (Fig. 15-25), the cycling of substrate appears to be the energetic “price” that  a muscle must pay to be able to change rapidly from a resting state (where  Vf— v r is small), in which substrate cycling is maximal, to one of sustained  high activity (where iy — v r is large). The rate of substrate cycling itself may  be under hormonal or neuronal control so as to increase the sensitivity of the  metabolic system under conditions when high activity (fight or flight) is  anticipated.   Substrate cycling and other mechanisms that control PFK activity in vivo  are part of larger systems that regulate all the cell’s metabolic activities. At one  time, it was believed that because PFK is the controlling enzyme of glycoly¬  sis, increasing its level of expression via genetic engineering would increase flux  through glycolysis. However, this is not the case, because the activity of PFK,  whatever its concentration, is ultimately controlled by factors that reflect the  cells demand for the products supplied by glycolysis and all other metabolic  pathways.    T Substrate Cycling Is Related to Thermogenesis and Obesity. Many an¬  imals, including adult humans, are thought to generate much of their body  heat, particularly when it is cold, through substrate cycling in muscle and liver,  a process known as nonshivering thermogenesis (the muscle contractions of  shivering or any other movement also produce heat). Substrate cycling is stim¬  ulated by thyroid hormones (which stimulate metabolism in most tissues) as  is indicated, for example, by the observation that rats lacking a functional thy¬  roid gland do not survive at 5°C. Chronically obese individuals tend to have  lower than normal metabolic rates, which is probably due, in part, to a re¬  duced rate of nonshivering thermogenesis. Such individuals therefore tend to  be cold sensitive. Indeed, whereas normal individuals increase their rate of thy¬  roid hormone activation on exposure to cold, genetically obese animals and  obese humans fail to do so.   • Why is ATP alone not an effective  allosteric regulator of enzyme activity?   • What is the metabolic advantage of a  substrate cycle? What is its cost?    The commonly available hexoses fructose, galactose, and mannose are converted  to glycolytic intermediates for further metabolism.    Together with glucose, the hexoses fructose, galactose, and mannose are promi¬  nent metabolic fuels. After digestion, these monosaccharides enter the blood¬  stream, which carries them to various tissues. Fructose, galactose, and mannose  are converted to glycolytic intermediates that are then metabolized by the gly¬  colytic pathway (Fig. 15-26).    Fructose is a major fuel source in diets that contain large amounts of fruit or  sucrose (a disaccharide of fructose and glucose; Section 8-2A). There are two  pathways for the metabolism of fructose; one occurs in muscle and the other  occurs in liver. This dichotomy results from the different enzymes present in  these tissues.   Fructose metabolism in muscle differs little from that of glucose.  Hexokinase (Section 15-2A), which converts glucose to G6P, also phosphory-  lates fructose, yielding F6P (Fig. 15-27, left). The entry of fructose into  glycolysis therefore involves only one reaction step.   Liver contains a hexokinase homolog known as glucokinase, which phos-  phorylates only glucose (Section 22-ID). Fructose metabolism in liver must  therefore differ from that in muscle. In fact, liver converts fructose to glycolytic  intermediates through a pathway that involves seven enzymes (Fig. 15-27,  right):   1. Fructokinase catalyzes the phosphorylation of fructose by ATP at Cl  to form fructose-1-phosphate. Neither hexokinase nor PFK can phos-  phorylate fructose-1-phosphate at C6 to form the glycolytic interme¬  diate FBP.   2. Aldolase (Section 15-2D) has several isozymic forms. Muscle contains  Type A aldolase, which is specific for FBP. Liver, however, contains  Type B aldolase, for which fructose-1-phosphate is also a substrate  (Type B aldolase is sometimes called fructose-1-phosphate aldolase).  In liver, fructose-1-phosphate therefore undergoes an aldol cleavage:   3. Direct phosphorylation of glyceraldehyde by ATP through the action  of glyceraldehyde kinase forms the glycolytic intermediate GAP.   4. Alternatively, glyceraldehyde is converted to the glycolytic intermedi¬  ate DHAP, beginning with its NADH-dependent reduction to glycerol  as catalyzed by alcohol dehydrogenase.   6. DHAP is produced by NAD + -dependent oxidation catalyzed by glyc¬  erol phosphate dehydrogenase.   7. The DHAP is then converted to GAP by triose phosphate isomerase.    FIG. 1 5-26 Entry of other hexoses into glycolysis. Fructose (in muscle) and  mannose are converted to F6P; liver fructose is converted to GAP; and galactose is  converted to G6P.    FIG. 1 5-27 The metabolism of fructose. In muscle (left), the  conversion of fructose to the glycolytic intermediate F6P involves only  one enzyme, hexokinase. In liver (right), seven enzymes participate in  the conversion of fructose to glycolytic intermediates.    Write an equation for the conversion of fructose to GAP in  the liver.    The two pathways leading from glyceraldehyde to GAP have the same net  cost: Both consume ATP, and although NADH is oxidized in Reaction 4, it  is reduced again in Reaction 6. The longer pathway, however, produces glycerol-  3-phosphate, which (along with DHAP) can become the glycerol backbone of  glycerophospholipids and triacylglycerols (Section 20-6A).   T Is Excess Fructose Harmful? The consumption of fructose in the United  States has increased at least 10-fold in the last quarter century, in large part  due to the use of high-fructose corn syrup as a sweetener in soft drinks and  other foods. Fructose has a sweeter taste than sucrose (Box 8-2) and is inex¬  pensive to produce. One possible hazard of excessive fructose intake is that  fructose catabolism in liver bypasses the PFK-catalyzed step of glycolysis and  thereby avoids a major metabolic control point. This could potentially disrupt    fuel metabolism so that glycolytic flux is directed toward lipid synthesis in the  absence of a need for ATP production. This hypothesis suggests a link between  the increase in both fructose consumption and the recently increasing inci¬  dence of obesity in the United States.   At the opposite extreme are individuals with fructose intolerance, which  results from a deficiency in Type B aldolase. In the absence of the aldolase,  fructose-1-phosphate may accumulate enough to deplete the livers store of P z .  Under these conditions, [ATP] drops, which causes liver damage. In addition,  the increased [fructose-1-phosphate] inhibits both glycogen phosphorylase  (an essential enzyme in the breakdown of glycogen to glucose; Section 16-1 A)  and fructose-1,6-bisphosphatase (an essential enzyme in gluconeogenesis;  Section 16-4B), thereby causing severe hypoglycemia (low levels of blood glu¬  cose), which can reach life-threatening proportions. However, fructose intol¬  erance is self-limiting: Individuals with the condition rapidly develop a strong  distaste for anything sweet.   Galactose is obtained from the hydrolysis of lactose (a disaccharide of galac¬  tose and glucose; Section 8-2A) in dairy products. Galactose and glucose are  epimers that differ only in their configuration at C4.   Although hexokinase phosphorylates glucose, fructose, and mannose, it does  not recognize galactose. An epimerization reaction must therefore occur be¬  fore galactose enters glycolysis. This reaction takes place after the conversion  of galactose to its uridine diphosphate derivative (the role of UDP-sugars  and other nucleotidyl—sugars is discussed in more detail in Section 16-5). The  entire pathway converting galactose to a glycolytic intermediate requires four  reactions (Fig. 15-28):   1. Galactose is phosphorylated at Cl by ATP in a reaction catalyzed by   2. Galactose-1-phosphate uridylyl transferase transfers the uridylyl  group of UDP-glucose to galactose-1-phosphate to yield glucose-1-  phosphate (G1P) and UDP-galactose by the reversible cleavage of  UDP-glucose’s pyrophosphoryl bond.   3. UDP-galactose-4-epimerase converts UDP-galactose back to UDP-  glucose. This enzyme has an associated NAD + , which suggests that the  reaction involves the sequential oxidation and reduction of the hexose  C4 atom:   4. G1P is converted to the glycolytic intermediate G6P by the action of  phosphoglucomutase (Section 16-1C).   FIG. 1 5-28 The metabolism of galactose. Four enzymes participate in the conversion of galactose to the glycolytic intermediate G6P.   Galactosemia is a genetic disease characterized by the inability to convert  galactose to glucose. Its symptoms include failure to thrive, mental retarda¬  tion, and, in some instances, death from liver damage. Most cases of galac¬  tosemia involve a deficiency in the enzyme catalyzing Reaction 2 of the  interconversion, galactose-1-phosphate uridylyl transferase. Formation of  UDP—galactose from galactose-1-phosphate is thus prevented, leading to a  buildup of toxic metabolic by-products. For example, the increased galactose  concentration in the blood results in a higher galactose concentration in the  lens of the eye, where the sugar is reduced to galactitol.    The presence of this sugar alcohol in the lens eventually causes cataract for¬  mation (clouding of the lens).   Galactosemia is treated by a galactose-free diet. Except for the mental re¬  tardation, this reverses all symptoms of the disease. The galactosyl units that   FIG. 1 5-29 The metabolism of mannose. Two enzymes are required to convert mannose to the glycolytic intermediate F6P.   • Describe how fructose, galactose, and  mannose enter the glycolytic pathway.    are essential for the synthesis of glycoproteins (Section 8-3C) and glycolipids  (Section 9-ID) can be synthesized from glucose by a reversal of the epimerase  reaction. These syntheses therefore do not require dietary galactose.   Mannose, a product of digestion of polysaccharides and glycoproteins, is the  C2 epimer of glucose.   Mannose enters the glycolytic pathway after its conversion to F6P via a two-  reaction pathway (Fig. 15-29):   2. Phosphomannose isomerase then converts this aldose to the glycolytic  intermediate F6P in a reaction whose mechanism resembles that of  phosphoglucose isomerase (Section 15-2B).    • The pentose phosphate pathway consists of three stages, in which NADPH is  produced, pentoses undergo isomerization, and glycolytic intermediates are  recovered.   • The pathway provides NADPH for reductive biosynthesis and ribose-5-phosphate  for nucleotide biosynthesis in the quantities that the cell requires.    ATP is the cells “energy currency”; its exergonic cleavage is coupled to many  otherwise endergonic cell functions. Cells also have a second currency, reducing  power. Many endergonic reactions, notably the reductive biosynthesis of fatty  acids (Section 20-4) and cholesterol (Section 20-7A), require NADPH in ad¬  dition to ATP. Despite their close chemical resemblance, NADPH and NADH  are not metabolically interchangeable. Whereas NADH uses the free energy of  metabolite oxidation to synthesize ATP (oxidative phosphorylation), NADPH  uses the free energy of metabolite oxidation for reductive biosynthesis. This  differentiation is possible because the dehydrogenases involved in oxidative and  reductive metabolism are highly specific for their respective coenzymes.  Indeed, cells normally maintain their [NAD + ]/[NADH] ratio near 1000,  which favors metabolite oxidation, while keeping their [NADP + ]/[NADPH]  ratio near 0.01, which favors reductive biosynthesis.   NADPH is generated by the oxidation of glucose-6-phosph ate via an alter - Section 6 The Pentose Phosphate Pathway   native pathway to glycolysis, the pentose phosphate pathway (also called the hex-  ose monophosphate shunt; Fig. 15-30). Tissues most heavily involved in  lipid biosynthesis (liver, mammary gland, adipose tissue, and adrenal cortex)  are rich in pentose phosphate pathway enzymes. Indeed, some 30% of the  glucose oxidation in liver occurs via the pentose phosphate pathway rather  than glycolysis.   FIG. 1 5-30 The pentose phosphate pathway. The number of  lines in an arrow represents the number of molecules reacting in one  turn of the pathway so as to convert 3 G6P to 3 C0 2 , 2 F6P, and 1  GAR For the sake of clarity, sugars from Reaction 3 onward are  shown in their linear forms. The carbon skeleton of R5P and the  atoms derived from it are drawn in red, and those from Xu5P are    drawn in green. The C 2 units transferred by transketolase are shaded  in green, and the C 3 units transferred by transaldolase are shaded in  blue. Double-headed arrows indicate reversible reactions.   3 G6P + 6 NADP + + 3 H 2 0   6 NADPH + 6 H + + 3 C0 2 + 2 F6P + GAP   Stage 1 Oxidative reactions (Fig. 15-30, Reactions 1—3), which yield  NADPH and ribulose-5-phosphate (Ru5P):   3 G6P + 6 NADP + + 3 H 2 0 ->   6 NADPH + 6 H + + 3 C0 2 + 3 Ru5P   4 and 5), which transform Ru5P either to ribose-5-phosphate  (R5P) or to xylulose-5-phosphate (Xu5P):   15-30, Reactions 6—8) that convert two molecules of Xu5P and  one molecule of R5P to two molecules of F6P and one molecule  of GAP   The reactions of Stages 2 and 3 are freely reversible, so the products of the  pathway vary with the needs of the cell (see below). In this section, we dis¬  cuss the three stages of the pentose phosphate pathway and how the pathway  is controlled.   G6P is considered the starting point of the pentose phosphate pathway. This  metabolite may arise through the action of hexokinase on glucose (Reaction  1 of glycolysis; Section 15-2A) or from glycogen breakdown (which produces  G6P directly; Section 16-1). Only the first three reactions of the pentose phos¬  phate pathway are involved in NADPH production (Fig. 15-30):   1. Glucose-6-phosphate dehydrogenase (G6PD) catalyzes net transfer  of a hydride ion to NADP + from Cl of G6P to form 6-phospho-  glucono-8-lactone:    G6P, a cyclic hemiacetal with Cl in the aldehyde oxidation state, is  thereby oxidized to a cyclic ester (lactone). The enzyme is specific for  NADP + and is strongly inhibited by NADPH.   2. 6-Phosphogluconolactonase increases the rate of hydrolysis of 6-  phosphoglucono-8-lactone to 6-phosphogluconate (the nonenzymatic  reaction occurs at a significant rate).   3. 6-Phosphogluconate dehydrogenase catalyzes the oxidative decar¬  boxylation of 6-phosphogluconate, a (3-hydroxy acid, to Ru5P and   FIG. 15-31 The 6-phosphogluconate dehydrogenase reaction. Oxidation of the  OH group forms an easily decarboxylated p-keto acid (although the proposed intermediate  has not been isolated).    C0 2 (Fig. 15-31). This reaction is thought to proceed via the forma¬  tion of a (3-keto acid intermediate. The keto group presumably facili¬  tates decarboxylation by acting as an electron sink.   Formation of Ru5P completes the oxidative portion of the pentose phos¬  phate pathway. It generates two molecules of NADPH for each molecule of G6P  that enters the pathway.   Ru5P is converted to R5P by ribulose-5-phosphate isomerase (Fig. 15-30,  Reaction 4) or to Xu5P by ribulose-5-phosphate epimerase (Fig. 15-30,  Reaction 5). These isomerization and epimerization reactions, like the reaction  catalyzed by triose phosphate isomerase (Section 15-2E), are thought to occur  via enediolate intermediates.   The relative amounts of R5P and Xu5P produced from Ru5P depend on the  needs of the cell. For example, R5P is an essential precursor in the biosynthe¬  sis of nucleotides (Chapter 23). Accordingly, R5P production is relatively high  (in fact, the entire pentose phosphate pathway activity may be elevated) in  rapidly dividing cells, in which the rate of DNA synthesis is increased. If the  pathway is being used solely for NADPH production, Xu5P and R5P are pro¬  duced in a 2:1 ratio for conversion to glycolytic intermediates in the third  stage of the pentose phosphate pathway as is discussed below.   How is a five-carbon sugar transformed to a six-carbon sugar such as F6P?  The rearrangements of carbon atoms in the third stage of the pentose phos¬  phate pathway are easier to follow by considering the stoichiometry of the  pathway. Every three G6P molecules that enter the pathway yield three Ru5P  molecules in Stage 1. These three pentoses are then converted to one R5P and  two Xu5P (Fig. 15-30, Reactions 4 and 5). The conversion of these three C 5  sugars to two sugars and one C 3 sugar involves a remarkable “juggling act”  catalyzed by two enzymes, transaldolase and transketolase. These enzymes  have mechanisms that involve the generation of stabilized carbanions and their  addition to the electrophilic centers of aldehydes.   Transketolase Catalyzes the Transfer of C 2 Units. Transketolase, which has  a thiamine pyrophosphate cofactor (TPP; Section 15-3B), catalyzes the   Chapter 15 Glucose Catabolism transfer of a C 2 unit from Xu5P to R5P, yielding GAP and sedoheptulose-   7-phosphate (S7P; Fig. 15-30, Reaction 6). The reaction intermediate is a co¬  valent adduct between Xu5P and TPP (Fig. 15 -32). The X-ray structure of  the dimeric enzyme shows that the TPP binds in a deep cleft between the sub¬  units so that residues from both subunits participate in its binding, just as in  pyruvate decarboxylase (another TPP-requiring enzyme; Fig. 15-19). In fact,  the structures are so similar that the enzymes likely diverged from a common  ancestor.   Transaldolase Catalyzes the Transfer of C 3 Units. Transaldolase catalyzes  the transfer of a C 3 unit from S7P to GAP yielding erythrose-4-phosphate  (E4P) and F6P (Fig. 15-30, Reaction 7). The reaction occurs by aldol cleav¬  age (Section 15-2D), which begins with the formation of a Schiff base    C2-C3 bond cleavage  yields GAP and a  resonance-stabilized (2  carbanion of the C 2  unit.    FIG. 1 5-32 Mechanism of transketolase. Transketolase formed on cleavage of the C2—C3 bond of Xu5P. The reaction   (represented by E) uses the coenzyme TPP to stabilize the carbanion occurs in 4 steps as shown.   between an 8-amino group of an essential Fys residue and the carbonyl group    A Second Transketolase Reaction Yields Glyceraldehyde-3-Phosphate  and a Second Fructose-6-Phosphate Molecule. In a second transketolase  reaction, a C 2 unit is transferred from a second molecule of Xu5P to E4P to  form GAP and another molecule of F6P (Fig. 15-30, Reaction 8). The third  stage of the pentose phosphate pathway thus transforms two molecules of  Xu5P and one of R5P to two molecules of F6P and one molecule of GAP To  summarize, a series of carbon-carbon bond formations and cleavages convert  three C 5 sugars to two sugars and one C 3 sugar. Here, the number to the  left of each reaction is keyed to the corresponding reaction in Fig. 15-30.    FIG. 1 5-33 Mechanism of transaldolase. Transaldolase  contains an essential Lys residue that facilitates an aldol cleavage  reaction as follows: (1) The £-amino group of Lys forms a Schiff base  with the carbonyl group of S7P; (2) a Schiff base-stabilized C3  carbanion is formed in an aldol cleavage reaction between C3 and C4    that eliminates E4P; (3) the enzyme-bound resonance-stabilized  carbanion adds to the carbonyl C atom of GAP, forming F6P linked to  the enzyme via a Schiff base; (4) the Schiff base hydrolyzes,  regenerating active enzyme and releasing F6P.    NADPH is required for several reductive processes in addition to  biosynthesis. For example, erythrocytes require a plentiful supply  of reduced glutathione (GSH), a Cys-containing tripeptide (Sec¬  tion 4-3B).   A major function of GSH in the erythrocyte is to reductively elimi¬  nate H 2 0 2 and organic hydroperoxides, which are reactive oxygen  metabolites that can irreversibly damage hemoglobin and cleave  the C—C bonds in the phospholipid tails of cell membranes. The  unchecked buildup of peroxides results in premature cell lysis.  Peroxides are eliminated by reaction with glutathione, catalyzed by  glutathione peroxidase:   GSSG represents oxidized glutathione (two GSH molecules linked  through a disulfide bond between their sulfhydryl groups).    Reduced GSH is subsequently regenerated by the reduction of  GSSG by NADPH as catalyzed by glutathione reductase:   GSSG + NADPH + H + -- 2 GSH + NADP +   A steady supply of NADPH is therefore vital for erythrocyte  integrity.   The erythrocytes in individuals who are deficient in glucose-  6-phosphate dehydrogenase (G6PD) are particularly sensitive to  oxidative damage, although clinical symptoms may be absent.  This enzyme deficiency, which is common in African, Asian, and  Mediterranean populations, came to light through investigations  of the hemolytic anemia that is induced in these individuals  when they ingest drugs such as the antimalarial compound  primaquine   or eat fava beans (broad beans, Vicia faba), a staple Middle  Eastern vegetable. Primaquine stimulates peroxide formation, thereby  increasing the demand for NADPH to a level that the mutant cells  cannot meet. Certain toxic glycosides present in small amounts in fava  beans have the same effect, producing a condition known as favism.   • Compare the transketolase and transal¬  dolase reactions in terms of substrates,  products, mechanism, and cofactor re¬  quirement.   • How does flux through the pentose phos¬  phate pathway change in response to the  need for NADPH or ribose-5-phosphate?   The principal products of the pentose phosphate pathway are R5P and  NADPH. The transaldolase and transketolase reactions convert excess R5P to  glycolytic intermediates when the metabolic need for NADPH exceeds that of  R5P in nucleotide biosynthesis. The resulting GAP and F6P can be consumed  through glycolysis and oxidative phosphorylation or recycled by gluconeogen-  esis (Section 16-4) to form G6P.   When the need for R5P outstrips the need for NADPH, F6P and GAP  can be diverted from the glycolytic pathway for use in the synthesis of R5P  by reversal of the transaldolase and transketolase reactions. The relationship  between glycolysis and the pentose phosphate pathway is diagrammed in   Flux through the pentose phosphate pathway and thus the rate of NADPH  production is controlled by the rate of the glucose-6-phosphate dehydrogenase reac¬  tion (Fig. 15-30, Reaction 1). The activity of the enzyme, which catalyzes the  pathways first committed step (AG — —17.6 kj • mol 1 in liver), is regu¬  lated by the NADP + concentration (i.e., regulation by substrate availability).  When the cell consumes NADPH, the NADP + concentration rises, increas¬  ing the rate of the G6PD reaction and thereby stimulating NADPH regener¬  ation. In some tissues, the amount of enzyme synthesized also appears to be  under hormonal control. A deficiency in G6PD is the most common clini¬  cally significant enzyme defect of the pentose phosphate pathway (Box 15-4).    The major reason for low enzymatic activity in affected cells ap¬  pears to be an accelerated rate of breakdown of the mutant enzyme.  This explains why patients with relatively mild forms of G6PD defi¬  ciency react to primaquine with hemolytic anemia but recover within  a week despite continued primaquine treatment. Mature erythrocytes  lack a nucleus and protein synthesizing machinery and therefore  cannot synthesize new enzyme molecules to replace degraded ones  (they likewise cannot synthesize new membrane components, which  is why they are so sensitive to membrane damage in the first place).  The initial primaquine treatments result in the lysis of old red blood  cells whose defective G6PD has been largely degraded. Lysis prod¬  ucts stimulate the release of young cells that contain more enzyme  and are therefore better able to cope with primaquine stress.   It is estimated that over 400 million people are deficient in  G6PD, which makes this condition the most common human  enzyme deficiency. Indeed, —400 G6PD variants have been re¬  ported and at least 140 of them have been characterized at the  molecular level. G6PD is active in a dimer-tetramer equilibrium.  Many of the mutation sites in individuals with the most severe  G6PD deficiency are at the dimer interface, shifting the equilib¬  rium toward the inactive and unstable monomer.   The high prevalence of defective G6PD in malarial areas of the  world suggests that such mutations confer resistance to the  malarial parasite, Plasmodium falciparum. Indeed, erythrocytes with  G6PD deficiency appear to be less suitable hosts for plasmodia  than normal cells. Thus, like the sickle-cell trait (Section 7-1 E), a  defective G6PD confers a selective advantage on individuals living  where malaria is endemic.   The G6PD deficiency primarily affects erythrocytes, in which the  lack of a nucleus prevents replacement of the unstable mutant en¬    zyme. However, the importance of NADPH in cells other than erythro¬  cytes has been demonstrated through the development of mice in  which the G6PD gene has been knocked out. All the cells in these  animals are extremely sensitive to oxidative stress, even though they  contain other mechanisms for eliminating reactive oxygen species.   Photo of red blood cells containing intracellular Plasmodium falciparum  (the malarial parasite), here seen as yellow bodies (named schizonts),  which generate the small red cells (called merozoites). [© Dr. Gopal  Murti/Photo Researchers, Inc.]    FIG. 1 5-34 Relationship between  glycolysis and the pentose phosphate  pathway. The pentose phosphate pathway,  which begins with G6P produced in Step 1 of  glycolysis, generates NADPH for use in  reductive reactions and R5P for nucleotide  synthesis. Excess R5P is converted to glycolytic  intermediates by a sequence of reactions that  can operate in reverse to generate additional  R5P, if needed.   • Glycolysis is a sequence of 10 enzyme-catalyzed reactions by which  one molecule of glucose is converted to two molecules of pyru¬  vate, with the net production of 2 ATP and the reduction of 2  NAD + to 2 NADH.   • In the first stage of glycolysis, glucose is phosphorylated by hexokin-  ase, isomerized by phosphoglucose isomerase (PGI), phosphory¬  lated by phosphofructokinase (PFK), and cleaved by aldolase to yield  the trioses glyceraldehyde-3-phosphate (GAP) and dihydroxyacetone  phosphate (DHAP), which are interconverted by triose phosphate  isomerase (TIM). These reactions consume 2 ATP per glucose.   • In the second stage of glycolysis, GAP is oxidatively phosphory¬  lated by glyceraldehyde-3-phosphate dehydrogenase (GAPDH),  dephosphorylated by phosphoglycerate kinase (PGK) to produce  ATP, isomerized by phosphoglycerate mutase (PGM), dehydrated  by enolase, and dephosphorylated by pyruvate kinase to produce  a second ATP and pyruvate. This stage produces 4 ATP per glu¬  cose for a net yield of 2 ATP per glucose.   • Under anaerobic conditions, pyruvate is reduced to regenerate  NAD + for glycolysis. In homolactic fermentation, pyruvate is re¬  versibly reduced to lactate.   • In alcoholic fermentation, pyruvate is decarboxylated by a thi¬  amine pyrophosphate (TPP)-dependent mechanism, and the re¬  sulting acetaldehyde is reduced to ethanol.   • The glycolytic reactions catalyzed by hexokinase, phosphofructo¬  kinase, and pyruvate kinase are metabolically irreversible.   • Phosphofructokinase is the primary flux control point for glycol¬  ysis. ATP inhibition of this allosteric enzyme is relieved by AMP  and ADP, whose concentrations change more dramatically than  those of ATP.   • The opposing reactions of the fructose-6-phosphate (F6P)/  fructose-1,6-bisphosphate (FBP) substrate cycle allow large  changes in glycolytic flux.   • Fructose, galactose, and mannose are enzymatically converted to  glycolytic intermediates for catabolism.   • In the pentose phosphate pathway, glucose-6-phosphate (G6P) is  oxidized and decarboxylated to produce two NADPH, C0 2 , and  ribulose-5-phosphate (Ru5P).   • Depending on the cell’s needs, ribulose-5-phosphate may be iso¬  merized to ribose-3-phosphate (R5P) for nucleotide synthesis or  converted, via ribose-5-phosphate and xylulose-5-phosphate  (Xu5P), to fructose-6-phosphate and glyceraldehyde-3-phosphate,  which can re-enter the glycolytic pathway.    1. Which of the 10 reactions of glycolysis are (a) phosphorylations,  (b) isomerizations, (c) oxidation-reductions, (d) dehydrations,  and (e) carbon-carbon bond cleavages?   2. When the pathogen Salmonella typhimurium infects mammalian  cells, the host cell protease caspase-1 is enabled to cleave and  thereby activate signaling proteins that instigate the immune re¬  sponse. The targets of caspase-1 also include aldolase and eno¬  lase. What effect would this have on the infected cell and why  might this be advantageous to the organism?   3. The aldolase reaction can proceed in reverse as an enzymatic al¬  dol condensation. If the enzyme were not stereospecific, how  many different products would be obtained?   4. Bacterial aldolase does not form a Schiff base with the substrate.  Instead, it has a divalent Zn 2+ ion in the active site. How does  the ion facilitate the aldolase reaction?    5. Arsenate (AsO| ), a structural analog of phosphate, can act as  a substrate for any reaction in which phosphate is a substrate.  Arsenate esters, unlike phosphate esters, are kinetically as well as  thermodynamically unstable and hydrolyze almost instanta¬  neously. Write a balanced overall equation for the conversion  of glucose to pyruvate in the presence of ATP, ADP, NAD + ,  and either (a) phosphate or (b) arsenate, (c) Why is arsenate a  poison?   6. The enzyme phosphoglucomutase interconverts glucose-1-phos¬  phate and glucose-6-phosphate. Why is this enzyme likely to in¬  clude a side chain such as Ser in its active site?   8. The compound you identified in Problem 7 is a metabolic reg¬  ulator. Predict its effect on (a) hexokinase and (b) PFK.   9. The pyruvate —> lactate reaction in animals is reversible, but the  pyruvate —> ethanol fermentation in yeast is irreversible. Explain.   10. The half-reactions involved in the lactate dehydrogenase (LDH)  reaction and their standard reduction potentials are   Calculate AC at pH 7.0 for the LDH-catalyzed reduction of  pyruvate under the following conditions:   (d) Discuss the effect of the concentration ratios in parts a-c on  the direction of the reaction.   11. Why is it possible for the AC values in Table 15-1 to differ from  the A G of values?   12. If a reaction has a AC°' value of at least —30.5 kj • mol 1 , suf¬  ficient to drive the synthesis of ATP (A G°’ = 30.5 kj • mol ! ),  can it still drive the synthesis of ATP in vivo when its AC is only  —10 kj • mol x ? Explain.   13. AC°' for the aldolase reaction is 22.8 kj • mol 1 . In the cell at  37°C, [DHAP]/[GAP] = 5.5. Calculate the equilibrium ratio of  [FBP]/[GAP] when [GAP] = 10“ 4 M.   14. The catalytic behavior of liver and brain phosphofructokinase-1  (PFK-1) was observed in the presence of AMP, P z , and fructose-  2,6-bisphosphate. The following table lists the concentrations of  each effector required to achieve 50% of the maximal velocity.  Compare the response of the two isozymes to the three effec¬  tors and discuss the possible implications of their different  responses.    15. Although it is not the primary flux-control point for glycolysis,  pyruvate kinase is subject to allosteric regulation. What is the  metabolic importance of regulating flux through the pyruvate  kinase reaction?   16. What is the advantage of activating pyruvate kinase with fruc¬  tose-1,6-bisphosphate?   17. Consider the pathway for catabolizing galactose. What are the  potential control points for this pathway?   18. Yeast take up and metabolize galactose, using the pathway out¬  lined in Fig. 15-28. Galactose-1-phosphate, an intermediate of  this pathway, inhibits phosphoglucomutase. Can this explain why  galactose catabolism is much slower than glucose catabolism in  yeast?   19. (a) Describe how glycerol enters the glycolytic pathway, (b) What  is the ATP yield for the conversion of glycerol to pyruvate?   20. Some organisms can anaerobically convert glycerol to pyruvate.  Could homolactic or alcoholic fermentation regenerate sufficient  NAD + to support this pathway?   21. Compare the ATP yield of three glucose molecules that enter gly¬  colysis and are converted to pyruvate with that of three glucose  molecules that proceed through the pentose phosphate pathway   such that their carbon skeletons (as two F6P and one GAP) reenter  glycolysis and are metabolized to pyruvate.   22. If G6P is labeled at its C2 position, where will the label appear  in the products of the pentose phosphate pathway?   23. Draw the enediolate intermediate of the ribulose-5-phosphate  isomerase reaction (Ru5P—»R5P).   24. Draw the enediolate intermediate of the ribulose-5-phosphate  epimerase reaction (Ru5P —>Xu5P).   25. Describe the lengths of the products of the transketolase reaction  when the two substrates are both five-carbon sugars.   26. Describe the products of the transketolase reaction when the sub¬  strates are a five-carbon aldose and a six-carbon ketose. Does it  matter which of the substrates binds to the enzyme first?   27. Explain why some tissues continue to produce C0 2 in the pres¬  ence of high concentrations of fluoride ion, which inhibits gly¬  colysis.   28. Some bacteria catabolize glucose by the Entner-Doudoroff  pathway, a variant of glycolysis in which glucose-6-phosphate is  converted to 6-phosphogluconate (as in the pentose phosphate  pathway) and then to 2-keto-3-deoxy-6-phosphogluconate  (KDPG).   Next, an aldolase acts on KDPG. (a) Draw the structures of the  products of the KDPG aldolase reaction, (b) Describe how these  reaction products are further metabolized by glycolytic enzymes,  (c) What is the ATP yield when glucose is metabolized to pyru¬  vate by the Entner-Doudoroff pathway? How does this compare  to the ATP yield of glycolysis?   29. A proposed pathway for ascorbic acid (vitamin C) biosynthesis in  plants takes the form   Match each of the following enzymes to one of the pathway  steps.   Case 18 Purification of Phosphofructokinase 1-C  Focus concept: The purification of the C isozyme of PFK-1 is  presented and the kinetic properties of the purified enzyme are  examined.   Case 20 NAD + -Dependent Glyceraldehyde-3-Phos-  phate Dehydrogenase from Therrmoproteus tenax  Focus concept: Glycolytic enzymes from T. tenax are regulated in  an unusual manner.    Plant cell walls, which contain  significant amounts of cellulose, are a source of glucose that can  be converted to ethanol by microbial fermentation. Outline the  metabolic pathways involved in the conversion of cellulose to  ethanol. In order for the production of cellulosic biofuels to become  economically viable, what steps of the process must be optimized?  Why are genetically engineered organisms part of this effort?    Bernstein, B.E., Michels, P.A.M., and Hoi, W.G.J., Synergistic effects of  substrate-induced conformational changes in phosphoglycerate acti¬  vation, Nature 385, 275-278 (1997).   Cappellini, M.D. and Fiorelli, G., Glucose-6-phosphate dehydrogenase  deficiency, Lancet 375, 64-74 (2008).   Dalby, A., Dauter, Z., and Littlechild, J.A., Crystal structure of human  muscle aldolase complexed with fructose 1,6-bisphosphate: Mecha¬  nistic implications, Protein Sci. 8, 291-297 (1999).   Depre, C., Rider, M.H., and Hue, L., Mechanisms of control of heart  glycolysis, Eur. J. Biochem. 258, 277-290 (1998). [Discusses how  the control of glycolysis in heart muscle is distributed among sev¬  eral enzymes, transporters, and other pathways.]   Frey, P.A., The Leloir pathway: a mechanistic imperative for three en¬  zymes to change the stereochemical configuration of a single carbon  in galactose, FASEB J. 10, 461-470 (1996).   Gefflaut, T., Blonski, C., Perie, J., and Wilson, M., Class I aldolases: sub¬  strate specificity, mechanism, inhibitors and structural aspects, Prog.  Biophys. Molec. Biol. 63, 301-340 (1995).    Hofmeyr, J.-H.S. and Cornish-Bowden, A., Regulating the cellular econ¬  omy of supply and demand, FEBS Lett. 476, 47-51 (2000).  Lindqvist, Y. and Schneider, G., Thiamin diphosphate dependent en¬  zymes: transketolase, pyruvate oxidase and pyruvate decarboxylase,  Curr. Opin. Struct. Biol. 3, 896-901 (1993).   Muirhead, H. and Watson, H., Glycolytic enzymes; from hexose to pyru¬  vate, Curr. Opin. Struct. Biol. 2, 870-876 (1992). [A brief summary  of the structures of glycolytic enzymes.]   Schirmer, T. and Evans, P.R., Structural basis of the allosteric behaviour  of phosphofructokinase, Nature 343, 140-145 (1990).   Valle, D. (Ed.), The Online Metabolic & Molecular Bases of Lnherited Dis¬  ease, http://www.ommbid.com/ [Chapters 70 and 72 discuss fruc¬  tose and galactose metabolism and their genetic disorders. Chapter  179 discusses glucose-6-phosphate dehydrogenase deficiency]    The muscles of animals contain glycogen, a storage form of the metabolic fuel  glucose. In living animals, the balance between glycogen synthesis and utilization is  carefully regulated. Measurement of glycogen levels in lobster has been used to  determine environmental conditions, since animals that have been undernourished or  stressed have depleted glycogen stores. [Andrew J. Martinez/Photo Researchers.]   B Glycogen Debranching Enzyme Acts as a  GI ucosy Itransferase  C Phosphoglucomutase Interconverts  Glucose-1-Phosphate and Glucose-6-Phosphate   B Glycogen Synthase Extends Glycogen Chains  C Glycogen Branching Enzyme Transfers  Seven-Residue Glycogen Segments    Glycogen (in animals, fungi, and bacteria) and starch (in plants) function to  stockpile glucose for later metabolic use. In animals, a constant supply of glu¬  cose is essential for tissues such as the brain and red blood cells, which de¬  pend almost entirely on glucose as an energy source (other tissues can also  oxidize fatty acids or amino acids for energy; Sections 20-2 and 21-4). The  mobilization of glucose from glycogen stores, primarily in the liver, provides  a constant supply of glucose (~5 mM in blood) to all tissues. When glucose  is plentiful, such as immediately after a meal, glycogen synthesis accelerates.  Yet the livers capacity to store glycogen is sufficient to supply the brain with  glucose for only about half a day. Under fasting conditions, most of the body’s  glucose needs are met by gluconeogenesis (literally, new glucose synthesis)  from noncarbohydrate precursors such as amino acids. Not surprisingly, the reg¬  ulation of glucose synthesis, storage, mobilization, and catabolism by glycoly¬  sis (Section 15-2) or the pentose phosphate pathway (Section 15-6) is elaborate  and is sensitive to the immediate and long-term energy needs of the organism.   The importance of glycogen for glucose storage is plainly illustrated by  the effects of deficiencies of the enzymes that release stored glucose. McArdle’s  disease, for example, is an inherited condition whose major symptom is  painful muscle cramps on exertion. The muscles in afflicted individuals lack  the enzyme required for glycogen breakdown to yield glucose. Although glyco¬  gen is synthesized normally, it cannot supply fuel for glycolysis to keep up  with the demand for ATP.   Figure 16-1 summarizes the metabolic uses of glucose. Glucose-6-  phosphate (G6P), a key branch point, is derived from free glucose through  the action of hexokinase (Section 15-2A) or is the product of glycogen break¬  down or gluconeogenesis. G6P has several possible fates: It can be used to syn¬  thesize glycogen; it can be catabolized via glycolysis to yield ATP and carbon  atoms (as acetyl-CoA) that are further oxidized by the citric acid cycle; and it   B Glycogen Phosphorylase and Glycogen  Synthase Undergo Control by Covalent  Modification    FIG. 1 6-1 Overview of glucose  metabolism. Glucose-6-phosphate (G6P) is  produced by the phosphorylation of free  glucose, by glycogen degradation, and by  gluconeogenesis. It is also a precursor for  glycogen synthesis and the pentose phosphate  pathway. The liver can hydrolyze G6P to glucose.  Glucose is metabolized by glycolysis to pyruvate,  which can be further broken down to acetyl-CoA  for oxidation by the citric acid cycle. Lactate and  amino acids, which are reversibly converted to  pyruvate, are precursors for gluconeogenesis.    can be shunted through the pentose phosphate pathway to generate NAD PH  and/or ribose-5-phosphate. In the liver and kidney, G6P can be converted to  glucose for export to other tissues via the bloodstream.   The opposing processes of glycogen synthesis and degradation, and of gly¬  colysis and gluconeogenesis, are reciprocally regulated; that is, one is largely  turned on while the other is largely turned off. In this chapter, we examine  the enzymatic steps of glycogen metabolism and gluconeogenesis, paying par¬  ticular attention to the regulatory mechanisms that ensure efficient operation  of opposing metabolic pathways.   Glucose mobilization in the liver involves a series of conversions from glycogen to  glucose-1-phosphate to glucose-6-phosphate and finally to glucose.    Glycogen is a polymer of a(l—>4)-linked D-glucose with a(l—>6)-linked  branches every 8-14 residues (Fig. \G-2a y b and Section 8-2C). Glycogen  occurs as intracellular granules of 100- to 400-A-diameter spheroidal mole¬  cules that each contain up to 120,000 glucose units (Fig. l6-2c). The gran¬  ules are especially prominent in the cells that make the greatest use of glycogen:  muscle (up to 1-2% glycogen by weight) and liver cells (up to 10% glycogen  by weight; Fig. 8-11). Glycogen granules also contain the enzymes that cat¬  alyze glycogen synthesis and degradation as well as many of the proteins that  regulate these processes.   Glucose units are mobilized by their sequential removal from the nonre¬  ducing ends of glycogen (the ends lacking a Cl-OH group). Whereas glyco¬  gen has only one reducing end, there is a nonreducing end on every branch.  Glycogens highly branched structure therefore permits rapid glucose mobiliza¬  tion through the simultaneous release of the glucose units at the end of every  branch.    FIG. 1 6-2 The structure of  glycogen, (a) Molecular formula. In the  actual molecule, there are — 12 residues  per chain, (b) Schematic diagram of  glycogen’s branched structure. Note that  the molecule has many nonreducing ends  but only one reducing end. In vivo , the  reducing end is covalently linked to the  protein glycogenin. (c) Electron micrograph  of a glycogen granule from rat skeletal  muscle. Each granule (labeled a) consists  of several spherical glycogen molecules  (p) and associated proteins. [From Calder,  P.C., Int. J. Biochem. 23, 1339 (1991).  Copyright Elsevier Science. Used with  permission.]   1. Glycogen phosphorylase (or simply phosphorylase) catalyzes glyco¬  gen phosphorolysis (bond cleavage by the substitution of a phosphate  group) to yield glucose-1-phosphate (G1P):   The enzyme releases a glucose unit only if it is at least five units away  from a branch point.   2. Glycogen debranching enzyme removes glycogens branches, thereby  making additional glucose residues accessible to glycogen phosphorylase.   3. Phosphoglucomutase converts G1P to G6P, which has several meta¬  bolic fates (Fig. 16-1).   Several key features of glycogen metabolism were discovered by the team of  Carl and Gerty Cori (Box 16-1).   Glycogen phosphorylase is a dimer of identical 842-residue (97-kD) subunits  that catalyzes the rate-controlling step in glycogen breakdown. It is regulated  both by allosteric interactions and by covalent modification (phosphorylation   A lifelong collaboration commenced with  the marriage of Carl and Gerty Cori in  1920. Although the Coris began their  professional work in Austria, they fled the  economic and social hardships of Europe  in 1922 and moved to Buffalo, New York.  They later made their way to Washington  University School of Medicine in St. Louis,  where Carl served as chair of the  Pharmacology Department and, later, chair  of the Biochemistry Department. Despite  her role as an equal partner in their  research work, Gerty officially remained a  research associate.   The Coris’ research focused primarily  on the metabolism of glucose. One of  their first discoveries was the connection  between glucose metabolism in muscle  and glycogen metabolism in the liver. The “Cori cycle” (Section  22-1 F) describes how lactate produced by glycolysis in active  muscle is transported to the liver, where it is used to synthesize  glucose that is stored as glycogen until needed.   After describing the interorgan movement of glucose in intact  animals, the Coris turned their attention to the metabolic fate of  glucose, specifically, the intermediates and enzymes of glucose  metabolism. In 1936, using a preparation of minced frog muscle,  the Coris found glucose in the form of a phosphate ester (called  the Cori ester, now known as glucose-1-phosphate). They traced  the presence of the Cori ester to the activity of a phosphorylase  (glycogen phosphorylase). This was a notable discovery, because    the enzyme used phosphate, rather than water, to split glucose  residues from the ends of glycogen chains. Even more remarkably,  the enzyme could be made to work in reverse to elongate a  glycogen polymer by adding glucose residues (from glucose-1-  phosphate). For the first time, a large biological molecule could be  synthesized in vitro.   During the 1940s, the Coris unraveled many of the secrets of  glycogen phosphorylase. For example, they found that the enzyme  exists in two forms, one that requires the activator AMP and one  that is active in the absence of an allosteric activator. Although it  was not immediately appreciated that the differences between the  two forms resulted from the presence of covalently bound  phosphate, this work laid the foundation for subsequent research on  enzyme regulation through phosphorylation and dephosphorylation.   Carl and Gerty Cori also described phosphoglucomutase, the  enzyme that converts glucose-1-phosphate to glucose-e-  phosphate so that it can participate in other pathways of glucose  metabolism. Over time, the Cori lab became a magnet for  scientists interested in purifying and characterizing other enzymes  of glucose metabolism.   Perhaps because of their experience with discrimination and—  especially for Gerty-the lack of equal opportunity, the Cori lab  welcomed a more diverse group of scientists than was typical of labs  of that era. The Coris received the 1947 Nobel Prize in Physiology  or Medicine. Several of their junior colleagues, Arthur Kornberg (see  Box 25-1), Severo Ochoa, Luis Leloir, Earl Sutherland, Christian de  Duve, and Edwin G. Krebs, later earned Nobel prizes of their own,  quite possibly reflecting the work ethic, broad view of science and  medicine, and meticulous work habits instilled by Carl and Gerty Cori.   Cori, G.T., Colowick, S.P., and Cori, C.F., The activity of the phosphorylating enzyme in  muscle extracts, J. Biol. Chem. 127 , 771-782 (1939).    and dephosphorylation). Phosphorylases allosteric inhibitors (ATP, G6P, and  glucose) and its allosteric activator (AMP) interact differently with the phospho-  and dephosphoenzymes, resulting in an extremely sensitive regulation process.  The structure of glycogen phosphorylase as well as its regulation by phospho¬  rylation are discussed in Section 12-3B.   An ~30-A-long crevice on the surface of the phosphorylase monomer  connects the glycogen storage site to the active site. Since this crevice can ac¬  commodate four or five sugar residues in a chain but is too narrow to admit  branched oligosaccharides , it provides a clear physical rationale for the inability of  phosphorylase to cleave glycosyl residues closer than five units from a branch point.  Presumably, the glycogen storage site increases the catalytic efficiency of phos¬  phorylase by permitting it to phosphorylyze many glucose residues on the same  glycogen particle without having to dissociate and reassociate completely be¬  tween catalytic cycles.   Phosphorylase binds the cofactor pyridoxal-5'-phosphate (PLP; at left),  which it requires for activity. This prosthetic group, a vitamin B 6 derivative,  is covalently linked to the enzyme via a Schiff base (imine) formed between  its aldehyde group and the 8-amino group of Lys 680. PLP also occurs in a  variety of enzymes involved in amino acid metabolism, where PLP’s conju¬  gated ring system functions catalytically to delocalize electrons (Sections 21-  2A and 21-4A). In phosphorylase, however, only the phosphate group  participates in catalysis, where it acts as a general acid-base catalyst.    Phosphorolysis of glycogen proceeds by a Random mechanism (Secdon 12-ID)  involving an enzyme • P z • glycogen ternary complex. An oxonium ion in¬  termediate forms during Cl—Ol bond cleavage, similar to the transition state  that forms in the reaction catalyzed by lysozyme (Section 11-4B). The phos-  phorylase reaction mechanism is diagrammed in Fig. 16-3, which shows how  PLP s phosphate group functions as a general acid-base catalyst.    Shielded oxonium ion intermediate  2 ) formation from the a-linked terminal  glucosyl residue involving acid  catalysis by P, as facilitated by proton  transfer from PLP The oxonium ion  half-chair conformation.    Reaction of P, with the oxonium ion with  overall retention of configuration about  3 Cl to form a-D-glucose-l-phosphate. The  glycogen, which has one less residue  than before, cycles back to step 1.    FIG. 1 6-3 The reaction mechanism of glycogen  phosphorylase. PL is an enzyme-bound pyridoxal group; BH + is a  positively charged amino acid side chain, probably that of Lys 568,    necessary for maintaining PLP electrical neutrality. The reaction occurs  via a shielded oxonium ion intermediate and involves acid catalysis by  P/ as facilitated by proton transfer from PLP.   structural differences between the active (R) and inactive (T) conformations  of phosphorylase (Fig. 12-16) are fairly well understood in terms of the sym¬  metry model of allosterism (Section 7-ID). The T-state enzyme has a buried  active site and hence a low affinity for its substrates, whereas the R-state  enzyme has an accessible catalytic site and a high-affinity phosphate-binding  site.   AMP promotes phosphorylase s T ( inactive) —> R ( active ) conformational  shift by binding to the R state of the enzyme at its allosteric effector site. This  conformational change results in increased access of the substrate to the active  site by disordering a loop of residues (282-286) that otherwise block the  active site. The conformational change also causes the Arg 569 side chain,  which is located in the active site near the PLP and the P z -binding site, to  rotate in a way that increases the enzymes binding affinity for its anionic P z  substrate (Fig. 12-16).   ATP also binds to the allosteric effector site, but in the T state, so that it  inhibits rather than promotes the T —» R conformational shift. This is because  the (3- and y-phosphate groups of ATP prevent the alignment of its ribose  and a-phosphate groups that are required for the conformational changes  elicited by AMP.   Phosphorylation and dephosphorylation alter enzymatic activity in a man¬  ner reminiscent of allosteric regulation. The phosphate group has a double  negative charge (a property not shared by naturally occurring amino acid  residues) and its covalent attachment to Ser 14 causes dramatic tertiary and  quaternary changes as the N-terminal segment moves to allow the phospho-  Ser to ion pair with two cationic Arg residues. The presence of the Ser 14—  phosphoryl group causes conformational changes similar to those triggered by AMP  binding, thereby shifting the enzyme's T R equilibrium in favor of the R state.  This accounts for the observation that phosphorylase b (the unphosphorylated  enzyme) requires AMP for activity and that the a form (the phosphorylated  enzyme) is active without AMP. We return to the regulation of phosphorylase  activity when we discuss the mechanisms that balance glycogen synthesis  against glycogen degradation (Section 16-3).   Phosphorolysis proceeds along a glycogen branch until it approaches to  within four or five residues of an a(1^6) branch point, leaving a “limit  branch.” Glycogen debranching enzyme acts as an Gt(l— >4) transglycosy-  lase (glycosyltransferase) by transferring an a(l—»4)-linked trisaccharide unit  from a limit branch of glycogen to the nonreducing end of another branch  (Fig. 16-4). This reaction forms a new a(l—>4) linkage with three more  units available for phosphorylase-catalyzed phosphorolysis. The a(l—>6)  bond linking the remaining glycosyl residue in the branch to the main chain  is hydrolyzed (not phosphorolyzed) by the same debranching enzyme to yield  glucose and debranched glycogen. About 10% of the residues in glycogen  (those at the branch points) are therefore converted to glucose rather than  G1P. Debranching enzyme has separate active sites for the transferase and the  a(1^6)-glucosidase reactions. The presence of two independent catalytic ac¬  tivities on the same enzyme no doubt improves the efficiency of the de¬  branching process.   The maximal rate of the glycogen phosphorylase reaction is much greater  than that of the glycogen debranching reaction. Consequently, the outermost  branches of glycogen, which constitute nearly half of its residues, are degraded  in muscle in a few seconds under conditions of high metabolic demand.  Glycogen degradation beyond this point requires debranching and hence  occurs more slowly. This, in part, accounts for the fact that a muscle can sus¬  tain its maximum exertion for only a few seconds.    FIG. 1 6-4 The reactions catalyzed by  debranching enzyme. The enzyme transfers the  terminal three a(1—»4)-linked glucose residues  from a “limit branch” of glycogen to the  nonreducing end of another branch. The a(1—>6)  bond of the residue remaining at the branch  point is hydrolyzed by further action of  debranching enzyme to yield free glucose. The  newly elongated branch is subject to degradation  by glycogen phosphorylase.   Phosphorylase converts the glucosyl units of glycogen to G1P, which, in turn,  is converted by phosphoglucomutase to G6P. The phosphoglucomutase reac¬  tion is similar to that catalyzed by phosphoglycerate mutase (Section 15-2H).  A phosphoryl group is transferred from the active phosphoenzyme to G1P,  forming glucose- 1,6-bisphosphate (G1,6P), which then rephosphorylates the  enzyme to yield G6P (Fig. 16-5; this near-equilibrium reaction also functions  in reverse). An important difference between this enzyme and phosphoglycerate    Glycogen storage diseases are inherited disorders that affect  glycogen metabolism, producing glycogen that is abnormal in  either quantity or quality. Studies of the genetic defects that  underlie these diseases have helped elucidate the complexities of  glycogen metabolism (e.g., McArdle’s disease). Conversely, the  biochemical characterization of the pathways affected by a genetic  disease often leads to useful strategies for its treatment. The  table on p. 525 lists the enzyme deficiencies associated with each  type of glycogen storage disease.   Glycogen storage diseases that mainly affect the liver generally  produce hepatomegaly (enlarged liver) and hypoglycemia (low  blood sugar), whereas glycogen storage diseases that affect the  muscles cause muscle cramps and weakness. Both types of  disease may also cause cardiovascular and renal disturbances.   Type I: Glucose-6-Phosphatase Deficiency (von Gierke’s  Disease). Glucose-6-phosphatase catalyzes the final step lead¬  ing to the release of glucose into the bloodstream by the liver.  Deficiency of the enzyme results in an increase of intracellular  [G6P], which leads to a large accumulation of glycogen in the liver  and kidney (because G6P activates glycogen synthase) and to an  inability to increase blood glucose concentration in response to  the hormones glucagon or epinephrine. The symptoms of Type I  glycogen storage disease include severe hepatomegaly and hypo¬  glycemia and a general failure to thrive. Treatment of the disease  has included drug-induced inhibition of glucose uptake by the liver  (to increase blood [glucose]), continuous intragastric feeding  overnight (again to increase blood [glucose]), surgical transposition  of the portal vein, which ordinarily feeds the liver directly from the  intestines (to allow this glucose-rich blood to reach peripheral  tissues before it reaches the liver), and liver transplantation.   a-1,4-Glucosidase deficiency is the most devastating of the glyco¬  gen storage diseases. It results in a large accumulation of glyco¬  gen of normal structure in the lysosomes of all cells and causes  death by cardiorespiratory failure, usually before the age of 1 year.  a-1,4-Glucosidase is not involved in the main pathways of glyco¬  gen metabolism. It occurs in lysosomes, where it hydrolyzes malt¬  ose (a glucose disaccharide) and other linear oligosaccharides, as  well as the outer branches of glycogen, thereby yielding free  glucose. Normally, this alternative pathway of glycogen metabolism   Type III: Amylo-1,6-Glucosidase (Debranching Enzyme) Defi¬  ciency (Cori’s Disease). In Cori’s disease, glycogen of abnormal  structure containing very short outer chains accumulates in both  liver and muscle since, in the absence of debranching enzyme, the  glycogen cannot be further degraded. The resulting hypoglycemia is  not as severe as in von Gierke’s disease (Type I) and can be  treated with frequent feedings and a high-protein diet (to offset the  loss of amino acids used for gluconeogenesis). For unknown rea¬  sons, the symptoms of Cori’s disease often disappear at puberty.   Type IV: Amylo-(1,4—>1,6)-Transglycosylase (Branching  Enzyme) Deficiency (Andersen’s Disease). Andersen’s disease  is one of the most severe glycogen storage diseases; victims  rarely survive past the age of 4 years because of liver dysfunction.  Liver glycogen is present in normal concentrations, but it contains  long unbranched chains that greatly reduce its solubility. The liver  dysfunction may be caused by a “foreign body” immune reaction  to the abnormal glycogen.   Type V: Muscle Phosphorylase Deficiency (McArdle’s  Disease). The symptoms of McArdle’s disease, painful muscle  cramps on exertion, typically do not appear until early adulthood  and can be prevented by avoiding strenuous exercise. This condi¬  tion affects glycogen metabolism in muscle but not in liver, which  contains normal amounts of a different phosphorylase isozyme.   Patients with a deficiency of liver glycogen phosphorylase have  symptoms similar to those with mild forms of Type I glycogen  storage disease. The hypoglycemia in this case results from the  inability of liver glycogen phosphorylase to respond to the need  for circulating glucose.   Type VII: Muscle Phosphofructokinase Deficiency (Tarui’s  Disease). The result of a deficiency of the glycolytic enzyme  PFK in muscle is an abnormal buildup of the glycolytic metabo¬  lites G6P and F6P. High concentrations of G6P increase the  activities of glycogen synthase and UDP-glucose pyrophosphory-  lase (G6P is in equilibrium with G1P, which is a substrate for    mutase is that the phosphoryl group in phosphoglucomutase is covalently  bound to a Ser hydroxyl group rather than to a His imidazole nitrogen.   Glucose-6-Phosphatase Generates Glucose in the Liver. The G6P pro¬  duced by glycogen breakdown can continue along the glycolytic pathway or  the pentose phosphate pathway (note that the glucose is already phosphory-  lated, so that the ATP-consuming hexokinase-catalyzed phosphorylation of  glucose is bypassed). In the liver, G6P is also made available for use by other  tissues. Because G6P cannot pass through the cell membrane, it is first hy¬  drolyzed by glucose-6-phosphatase (G6Pase):    UDP-glucose pyrophosphorylase, an enzyme required for glyco¬  gen synthesis) so that glycogen accumulates in muscle. Other  symptoms are similar to those of muscle phosphorylase deficiency,  since PFK deficiency prevents glycolysis from keeping up with the  ATP demand in contracting muscle.   Some individuals with symptoms of Type VI glycogen storage  disease have normal phosphorylase enzymes but a defective phos¬  phorylase kinase, which results in their inability to convert phos¬  phorylase b to phosphorylase a. The a subunit of phosphorylase  kinase is encoded by a gene on the X chromosome, so Type VIII  disease is X-linked rather than autosomal recessive, as are the  other glycogen storage diseases.   Type IX: Phosphorylase Kinase Deficiency. Phosphorylase  kinase deficiency, an autosomal recessive disease, results from a    mutation in one of the genes that encode the p, 7 , and 8 sub¬  units of phosphorylase kinase. Because different tissues contain  different phosphorylase kinase isozymes, the symptoms and  severity of the disease vary according to the affected organs.  Techniques for identifying genetic lesions are therefore more  reliable than clinical symptoms for diagnosing a particular  glycogen storage disease.   Type 0: Liver Glycogen Synthase Deficiency. Liver glycogen  synthase deficiency is the only disease of glycogen metabolism  in which there is a deficiency rather than an overabundance  of glycogen. The activity of liver glycogen synthase is  extremely low in individuals with Type 0 disease, who exhibit  hyperglycemia after meals and hypoglycemia at other times.  Some individuals, however, are asymptomatic, which suggests  that there may be multiple forms of this autosomal recessive  disorder.    Although G6P is produced in the cytosol, G6Pase resides in the endoplasmic  reticulum (ER) membrane. Consequently G6P must be imported into the ER  by a G6P translocase before it can be hydrolyzed. The resulting glucose and  P z are then returned to the cytosol via specific transport proteins. A defect in  any of the components of this G6P hydrolysis system results in type I glyco¬  gen storage disease (Box 16-2). Glucose leaves the liver cell via a specific glu¬  cose transporter named GLUT2 and is carried by the blood to other tissues.  Muscle and other tissues lack G6Pase and therefore retain their G6P.   • Describe the structure of a glycogen  molecule. What is the advantage of its  branched structure?   • List the three enzymes involved in glyco¬  gen degradation and describe the type of  reactions they catalyze.   • List the activators and inhibitors of glyco¬  gen phosphorylase. How does phosphory¬  lation affect its activity?    • Liver glycogen synthesis involves a series of conversions from glucose to  glucose-6-phosphate, to UDP-glucose, and finally to glycogen.   • Glycogen is extended from a primer built on and by the protein glycogenin.    The A G°' for the glycogen phosphorylase reaction is +3.1 kj • mol \ but  under physiological conditions, glycogen breakdown is exergonic (AG°' = — 5  to — 8 kj • mol x ). The synthesis of glycogen from G1P under physiological  conditions is therefore thermodynamically unfavorable without free energy in¬  put. Consequently, glycogen synthesis and breakdown must occur by separate path¬  ways. This recurrent metabolic strategy—that biosynthetic and degradative  pathways of metabolism are different—is particularly important when both  pathways must operate under similar physiological conditions. This situation  is thermodynamically impossible if one pathway is just the reverse of the other.   It was not thermodynamics, however, that led to recognition of the sepa¬  ration of synthetic and degradative pathways for glycogen, but McArdle’s dis¬  ease. Individuals with the disease lack muscle glycogen phosphorylase activity  and therefore cannot break down glycogen. Yet their muscles contain moder¬  ately high quantities of normal glycogen. Clearly, glycogen synthesis does not  require glycogen phosphorylase. In this section, we describe the three enzymes  that participate in glycogen synthesis: UDP-glucose pyrophosphorylase,  glycogen synthase, and glycogen branching enzyme. The opposing reactions  of glycogen synthesis and degradation are diagrammed in Fig. 16-6.   Since the direct conversion of G1P to glycogen and P z is thermodynamically  unfavorable (positive A G) under physiological conditions, glycogen biosynthe¬  sis requires an exergonic step. This is accomplished, as Luis Leloir discovered  in 1957, by combining G1P with uridine triphosphate (UTP) in a reaction  catalyzed by UDP-glucose pyrophosphorylase (Fig. 16-7). The product of this  reaction, uridine diphosphate glucose (UDP-glucose or UDPG) , is an  “activated” compound that can donate a glucosyl unit to the growing glycogen  chain. The formation of UDPG itself has A G or ~ 0 (it is a phosphoanhy-  dride exchange reaction), but the subsequent exergonic hydrolysis of PP Z by   FIG. 1 6-6 Opposing pathways of glycogen synthesis and degradation. The exer¬  gonic process of glycogen breakdown is reversed by a process that uses UTP to generate  a UDP-glucose intermediate.   FIG. 16-7 The reaction catalyzed by UDP-glucose  pyrophosphorylase. This is a phosphoanhydride exchange reaction.  The reaction is driven to completion by PP, hydrolysis.   This is an example of the common biosynthetic strategy of cleaving a nucle¬  oside triphosphate to form PP,. The free energy of PP, hydrolysis can then be  used to drive an otherwise unfavorable reaction to completion (Section 14-2B);  the near total elimination of the PP, by the highly exergonic (irreversible)  pyrophosphatase reaction prevents the reverse of the PP,-producing reaction  from occurring.   In the next step of glycogen synthesis, the glycogen synthase reaction, the glu-  cosyl unit of UDPG is transferred to the C4-OH group on one of glycogens   is —13.4 kj • mol \ making the overall reaction spontaneous under the same  conditions that glycogen breakdown by glycogen phosphorylase is also spon¬  taneous. However, glycogen synthesis does have an energetic price. Combining  the first two reactions of glycogen synthesis gives   Thus, one molecule of UTP is cleaved to UDP for each glucose residue incorpo¬  rated into glycogen. The UTP is replenished through a phosphoryl-transfer re¬  action mediated by nucleoside diphosphate kinase (Section 14-2C):   The transfer of a glucosyl unit from UDPG to a growing glycogen chain in¬  volves the formation of a glycosyl oxonium ion by the elimination of UDP, a  good leaving group (Fig. 16-8). The enzyme is inhibited by 1,5-ghiconolactone  (at left), an analog that mimics the oxonium ions half-chair geometry. The  same analog inhibits both glycogen phosphorylase (Section 16-1 A) and  lysozyme (Section 11-4), which have similar mechanisms.   Human muscle glycogen synthase is a homotetramer of 737-residue sub¬  units (the liver isozyme has 703-residue subunits). Like glycogen phosphory¬  lase, it has two enzymatically interconvertible forms; in this case, however, the  phosphorylated b form is less active, and the original (dephosphorylated) a  form is more active. (Note: For enzymes subject to covalent modification, “a*  refers to the more active form and refers to the less active form.)   Glycogen synthase is under allosteric control; it is strongly inhibited by  physiological concentrations of ATP, ADP, and P z . In fact, the phosphorylated   enzyme is almost totally inactive in vivo. The dephosphorylated enzyme, how- Section 2 Glycogen Synthesis   ever, can be activated by G6P, so the cells glycogen synthase activity varies  with [G6P] and the fraction of the enzyme in its dephosphorylated form. The  mechanistic details of the interconversion of phosphorylated and dephospho¬  rylated forms of glycogen synthase are complex and are not as well understood  as those of glycogen phosphorylase (for one thing, glycogen synthase has mul¬  tiple phosphorylation sites). We discuss the regulation of glycogen synthase  further in Section 16-3B.   Glycogenin Primes Glycogen Synthesis. Glycogen synthase cannot simply  link together two glucose residues; it can only extend an already existing  a(l—>4)-linked glucan chain. How, then, is glycogen synthesis initiated? In  the first step of this process, a 349-residue protein named glycogenin, acting  as a glycosyltransferase, attaches a glucose residue donated by UDPG to the  OH group of its Tyr 194. Glycogenin then extends the glucose chain by up  to seven additional UDPG-donated glucose residues to form a glycogen  “primer.” Only at this point does glycogen synthase commence glycogen  synthesis by extending the primer. Analysis of glycogen granules suggests that  each glycogen molecule is associated with only one molecule each of glycogenin  and glycogen synthase.   Branching to form glycogen is accomplished by a separate enzyme, amylo-  (1,4—»l,6)-transglycosylase (branching enzyme), which is distinct from  glycogen debranching enzyme (Section 16-IB). A branch is created by trans¬  ferring a 7-residue segment from the end of a chain to the C6-OH group of  a glucose residue on the same or another glycogen chain (Fig. 16-9). Each    • Why must opposing biosynthetic and  degradative pathways differ in at least one  enzyme?   • List the three enzymes involved in glyco¬  gen synthesis and describe the types of  reactions they catalyze.    transferred segment must come from a chain of at least 11 residues, and the  new branch point must be at least 4 residues away from other branch points.  The branching pattern of glycogen has been optimized by evolution for the  efficient storage and mobilization of glucose (Box 16-3).   • The opposing processes of glycogen breakdown and synthesis are reciprocally  regulated by allosteric interactions and the covalent modification of key enzymes.   • Glycogen metabolism is ultimately under the control of hormones such as insulin,  glucagon, and epinephrine.    If glycogen synthesis and breakdown proceed simultaneously, all that is accom¬  plished is the wasteful hydrolysis of UTP. Glycogen metabolism must there¬  fore be controlled according to cellular needs. The regulation of glycogen  metabolism involves allosteric control as well as hormonal control by covalent mod¬  ification of the pathways regulatory enzymes.   As we saw in Sections 14-IE and 15-4B, the net flux, J, of reactants through  a step in a metabolic pathway is the difference between the forward and re¬  verse reaction velocities, zy and v r . However, the flux varies dramatically with  substrate concentration as the reaction approaches equilibrium (zy ~ zy). The  flux through a near-equilibrium reaction is therefore all but uncontrollable.  Precise flux control of a pathway is possible when an enzyme functioning far from  equilibrium is opposed by a separately controlled enzyme. Then , z yand v r vary in¬  dependently and v r can be larger or smaller than zy, allowing control of both rate  and direction. Exactly this situation occurs in glycogen metabolism through  the opposition of the glycogen phosphorylase and glycogen synthase reactions.   Both glycogen phosphorylase and glycogen synthase are under allosteric  control by effectors that include ATP, G6P, and AMP. Muscle glycogen phos¬  phorylase is activated by AMP and inhibited by ATP and G6P. Glycogen syn¬  thase, on the other hand, is activated by G6P. This suggests that when there  is high demand for ATP (low [ATP], low [G6P], and high [AMP]), glycogen  phosphorylase is stimulated and glycogen synthase is inhibited, which favors  glycogen breakdown. Conversely, when [ATP] and [G6P] are high, glycogen  synthesis is favored.   In vivo , this allosteric scheme is superimposed on an additional control  system based on covalent modification. For example, phosphorylase a is active  even without AMP stimulation (Section 16-1A), and glycogen synthase is  essentially inactive (Section 16-2B) unless it is dephosphorylated and G6P is  present. Thus, covalent modification (phosphorylation and dephosphorylation) of  glycogen phosphorylase and glycogen synthase provides a more sophisticated control  system that modulates the responsiveness of the enzymes to their allosteric effectors.    B Glycogen Phosphorylase and Glycogen Synthase Undergo Control  by Covalent Modification   The interconversion of the a and b forms of glycogen synthase and glycogen  phosphorylase is accomplished through enzyme-catalyzed phosphorylation and  dephosphorylation (Section 12-3B), a process that is under hormonal control  (Section 13-2). Enzymatically interconvertible enzyme systems can there¬  fore respond to a greater number of effectors than simple allosteric systems.    The function of glycogen in animal cells is to store the metabolic  fuel glucose and to release it rapidly when needed. Glucose must  be stored as a polymer, because glucose itself could not be  stored without a drastic increase in intracellular osmotic pressure  (Section 2-1 D). It has been estimated that the total concentration  of glucose residues stored as glycogen in a liver cell is —0.4 M,  whereas the concentration of glycogen is only —10 nM. This huge  difference mitigates osmotic stress.   To fulfill its biological function, the glycogen polymer must store  the largest amount of glucose in the smallest possible volume  while maximizing both the amount of glucose available for release  by glycogen phosphorylase and the number of nonreducing ends  (to maximize the rate at which glucose residues can be mobilized).  These criteria must be met by optimizing just two variables: the  degree of branching and chain length.   the glycogen chains, beginning with the innermost chain attached  to glycogenin (G), each have two branches (the outermost chains  are unbranched). The entire molecule is roughly spherical and is  organized in tiers. There are an estimated 12 tiers in mature  glycogen (only 4 are shown above).   With two branches per chain, the number of chains in a given  tier is twice the number of the preceding tier, and the outermost  tier contains about half of the total glucose residues (regardless  of the number of tiers). When the degree of branching  increases, for example, to three branches per chain, the propor¬  tion of residues in the outermost tier increases, but so does the  density of glucose residues. This severely limits the maximum  size of the glycogen particle and the number of glucose  residues it can accommodate. Thus, glycogen has around two  branches per chain.   Mathematical analysis of the other variable, chain length, yields  an optimal value of 13, which is in good agreement with the actual  length of glycogen chains in cells (8-14 residues). Consider the  two simplified glycogen molecules shown below, which contain the  same number of glucose residues (the same total length of line  segments) and the same branching pattern:    The molecule with the shorter chains packs more glucose in a  given volume and has more points for phosphorylase attack, but  only about half the amount of glucose can be released before  debranching must occur (debranching is much slower than phos-  phorolysis). In the less dense molecule, the longer chains increase  the number of residues that can be continuously phosphorylyzed;  however, there are fewer points of attack. Thirteen residues is  apparently a compromise for mobilizing the largest amount of  glucose in the shortest time.   Amylopectin (Section 8-2C), which is chemically similar to  glycogen, is a much larger molecule and has longer chains.  Amylose lacks branches altogether. Evidently, starch, unlike glyco¬  gen, is not designed for rapid mobilization of metabolic fuel.   [Figures adapted from Melendez-Hevia, E., Waddell, T.G., and Shelton,  E.D., Biochem. J. 295, 477-483 (1993).]   Furthermore, a set of kinases and phosphatases linked in cascade fashion has  enormous potential for signal amplification and flexibility in response to dif¬  ferent metabolic signals. Note that the correlation between phosphorylation  and enzyme activity varies with the enzyme. For example, glycogen phospho¬  rylase is activated by phosphorylation (b —> a), whereas glycogen synthase is  inactivated by phosphorylation (a —> b). Conversely, dephosphorylation inac¬  tivates glycogen phosphorylase and activates glycogen synthase.    Glycogen Phosphorylase Is Activated by Phosphorylation. The cascade  that governs the enzymatic interconversion of glycogen phosphorylase involves  three enzymes (Fig. 16-10):   1 . Phosphorylase kinase , which specifically phosphorylates Ser 14 of  glycogen phosphorylase b.   2. Protein kinase A ( PKA ; Section 13-3C), which phosphorylates and  thereby activates phosphorylase kinase.   3. Phosphoprotein phosphatase-1 ( PP1 ; Section 13-2D), which dephos-  phorylates and thereby deactivates both glycogen phosphorylase a and  phosphorylase kinase.   Phosphorylase b is sensitive to allosteric effectors, but phosphorylase a is  much less so, as discussed in Section 12-3B (Fig. 12-17). In the resting cell,  the concentrations of ATP and G6P are high enough to inhibit phosphory¬  lase b. The level of phosphorylase activity is therefore largely determined by  the fraction of enzyme present as phosphorylase a. The steady state fraction  of phosphorylated enzyme depends on the relative activities of phosphorylase  kinase, PKA, and PP1. Recall that PKA is activated by cAMP, a second mes¬  senger that is made by adenylate cyclase on hormone-stimulated activation  of a heterotrimeric G protein (Section 13-3C). Let us examine the factors that  regulate the activities of phosphorylase kinase and PP1 before we return to the  regulation of glycogen synthase activity.   Phosphorylase Kinase Is Activated by Phosphorylation and by Ca 2+ .   Phosphorylase kinase is a 1300-kD protein with four nonidentical subunits,  known as a, (3, y, and 8. The y subunit contains the catalytic site, and the  other three subunits have regulatory functions. Phosphorylase kinase is maxi¬  mally activated by Ca 2+ and by the phosphorylation of its a and (3 subunits by  PKA.    FIG. 16-10 The glycogen phosphorylase  interconvertible enzyme system. Conversion  of phosphorylase b (the less active form) to  phosphorylase a (the more active form) is  accomplished through phosphorylation catalyzed  by phosphorylase kinase, which is itself subject  to activation through phosphorylation by protein  kinase A (PKA). Both glycogen phosphorylase a  and phosphorylase kinase are dephosphorylated  by phosphoprotein phosphatase-1.    FIG. 1 6-1 1 X-Ray structure of the y subunit of rabbit muscle phosphorylase  kinase in complex with ATP and a substrate analog. The N-terminal domain is  pink, the C-terminal domain is cyan, the activation loop is light blue, and the  heptapeptide substrate analog is orange, with its residue to be phosphorylated (Ser)  white. The ATP is shown in space-filling form and the side chains of the catalytically  essential Arg 148, Asp 149, and Glu 182 are shown in stick form, all colored according  to atom type (C green, N blue, 0 red, and P yellow). Note the structural similarities and  differences between this protein and other protein kinases, for example, the C subunit of  protein kinase A (Fig. 13-21) and the tyrosine kinase domain of the insulin receptor (Fig.  13-5). [After an X-ray structure by Louise Johnson, Oxford University, U.K. PDBid 2PHK.]    The 386-residue y subunit of phosphorylase kinase is 36% identical  in sequence to the PKA C subunit (Fig. 13-21) and has a similar struc¬  ture (Fig. 16-11). The 7 subunit is not subject to phosphorylation, as are  many other protein kinases, because the Ser, Thr, or Tyr residue that is  phosphorylated to activate those other kinases is replaced by a Glu residue  in the 7 subunit. The negative charge of the Glu is thought to mimic the  presence of a phosphate group and interact with a conserved Arg residue near  the active site. However, full catalytic activity of the 7 subunit is prevented  by an autoinhibitory C-terminal segment, which binds to and blocks the  kinases active site, much like the R subunit blocks the activity of the  C subunit of protein kinase A. An inhibitory segment in the (3 subunit  may also block the activity of the 7 subunit. f   Autoinhibition of phosphorylase kinase is relieved by PKA-catalyzed  phosphorylation of both the a and (3 subunits. This presumably causes the (3  inhibitor segment to move aside (the way in which phosphorylation of the a  subunit modulates the enzymes behavior is not understood). However, full ac¬  tivity of the 7 subunit also requires Ca 2+ binding to the 8 subunit, which is  calmodulin (CaM; Section 13-4B; CaM functions both as a free-floating pro¬  tein and as a subunit of other proteins). Ca 2+ concentrations as low as 10 7  M activate phosphorylase kinase by inducing a conformational change in CaM  that causes it to bind to and extract the 7 subunits autoinhibitor segment  from its catalytic site (Fig. 13-29).   The conversion of glycogen phosphorylase b to glycogen phosphorylase a  through the action of phosphorylase kinase increases the rate of glycogen  breakdown. The physiological significance of the Ca 2+ trigger for this activa¬  tion is that muscle contraction is also triggered by a transient increase in the  level of cytosolic Ca 2+ (Section 7 - 2 B). The rate of glycogen breakdown is  thereby linked to the rate of muscle contraction. This is critical because glyco¬  gen breakdown provides fuel for glycolysis to generate the ATP required for  muscle contraction. Since Ca 2+ release occurs in response to nerve impulses,  whereas the phosphorylation of phosphorylase kinase ultimately occurs in re¬  sponse to the presence of certain hormones, these two signals act synergisti-  cally in muscle cells to stimulate glycogenolysis.   A steady state for many phosphorylated enzymes is maintained by a balance  between phosphorylation, as catalyzed by a corresponding kinase, and hy¬  drolytic dephosphorylation, as catalyzed by a phosphatase. Phosphoprotein  phosphatase -1 (PP 1 ) removes the phosphoryl groups from glycogen phos¬  phorylase a and the a and (3 subunits of phosphorylase kinase (Fig. 16-10), as  well as those of other proteins involved in glycogen metabolism (see below).   PP 1 is controlled differently in muscle and in liver. In muscle, the cat¬  alytic subunit of PP1 (called PPlc) is active only when it is bound to glyco¬  gen through its glycogen-binding G M subunit. The activity of PPlc and its  affinity for the G M subunit are regulated by phosphorylation of the G M subunit   FIG. 1 6-1 2 Regulation of phosphoprotein phosphatase-1 in  muscle. The antagonistic effects of insulin and epinephrine on  glycogen metabolism in muscle occur through their effects on the    phosphoprotein phosphatase-1 catalytic subunit, PPIc, via its  glycogen-bound G M subunit. Green circles and dashed arrows indicate  activation.    at two separate sites (Fig. 16-12). Phosphorylation of site 1 by an insulin-  stimulated protein kinase (a homolog of PKA and the y subunit of phos-  phorylase kinase) activates PPIc, whereas phosphorylation of site 2 by PKA  (which can also phosphorylate site 1) causes PPIc to be released into the cy¬  toplasm, where it cannot dephosphorylate the glycogen-bound enzymes of  glycogen metabolism.   In the cytosol, phosphoprotein phosphatase-1 is also inhibited by its bind¬  ing to the protein phosphoprotein phosphatase inhibitor 1 . The latter  protein provides yet another example of control by covalent modification: It  too is activated by PKA and deactivated by phosphoprotein phosphatase-1  (Fig. 16-13, lower left). The concentration ofcAMP therefore controls the fraction  of an enzyme in its phosphorylated form, not only by increasing the rate at which  it is phosphorylated, but also by decreasing the rate at which it is dephosphorylated.  In the case of glycogen phosphorylase, an increase in [cAMP] not only increases  the enzymes rate of activation, but also decreases its rate of deactivation.   In liver, PP1 is also bound to glycogen, but through the intermediacy of  a glycogen-binding subunit named G L . In contrast to G M , G L is not subject  to control via phosphorylation. The activity of the PP1 • G L complex is  controlled by its binding to phosphorylase a. Both the R and T forms of  phosphorylase a strongly bind PP1, but only in the T state is the Ser 14 phos-  phoryl group accessible for hydrolysis (in the R state, the Ser 14 phosphoryl  group is buried at the dimer interface; Fig. 12-16). Consequently, when phos¬  phorylase a is in its active R form, it effectively sequesters PP1. However,  under conditions where phosphorylase a shifts to the T state (see below), PP1  hydrolyzes the now exposed Ser 14 phosphoryl group, thereby converting  phosphorylase a to phosphorylase b, which has only a low affinity for the  PP1 • G l complex. One effect of phosphorylase a dephosphorylation, there¬  fore, is to relieve the inhibition of PP1. Since liver cells contain 10 times more  glycogen phosphorylase than PP1, the phosphatase is not released until more  than ~90% of the glycogen phosphorylase is in the b form. Only then can  PP1 dephosphorylate its other target proteins, including glycogen synthase.    Glucose is an allosteric inhibitor of phosphorylase a (Fig. 12-17).  Consequently, when the concentration of glucose is high, phosphorylase a con¬  verts to its T form, thereby leading to its dephosphorylation and the dephos¬  phorylation of glycogen synthase. Glucose is therefore thought to be important  in the control of glycogen metabolism in the liver.   Glycogen Synthase Is Elaborately Regulated. Phosphorylase kinase, which  activates glycogen phosphorylase, also phosphorylates and thereby inactivates  glycogen synthase. Eight other protein kinases, including PKA and phos¬  phorylase kinase, are known to at least partially deactivate human muscle glyco¬  gen synthase by phosphorylating one or more of the nine Ser residues on each  of its subunits (Fig. 16-13). The reason for this elaborate regulation of glyco¬  gen synthase is unclear.   The balance between net synthesis and degradation of glycogen as well as  the rates of these processes depend on the relative activities of glycogen syn¬  thase and glycogen phosphorylase. To a large extent, the rates of the phos¬  phorylation and dephosphorylation of these enzymes control glycogen synthesis    and breakdown. The two processes are linked by PKA and phosphorylase ki¬  nase, which, through phosphorylation, activate glycogen phosphorylase as they  inactivate glycogen synthase (Fig. 16-13). They are also linked by PP1, which  in liver is inhibited by phosphorylase a and therefore unable to activate (de-  phosphorylate) glycogen synthase unless it first inactivates (also by dephos¬  phorylation) phosphorylase a. Of course, control by allosteric effectors is  superimposed on control by covalent modification so that, for example, the  availability of the substrate G6P (which activates glycogen synthase) also in¬  fluences the rate at which glucose residues are incorporated into glycogen.  Inherited deficiencies of enzymes can disrupt the fine control of glycogen me¬  tabolism, leading to various diseases (Box 16-2).   Glycogen metabolism in the liver is largely controlled by the polypeptide hor¬  mones insulin (Fig. 5-1) and glucagon acting in opposition. Glucagon,   like insulin, is synthesized by the pancreas in response to the concentration of  glucose in the blood. In muscles and various tissues, control is exerted by in¬  sulin and by the adrenal hormones epinephrine and norepinephrine (Section  13-IB). These hormones affect metabolism in their target tissues by ultimately  stimulating covalent modification (phosphorylation) of regulatory enzymes.  They do so by binding to transmembrane receptors on the surface of cells.  Different cell types have different complements of receptors and therefore re¬  spond to different sets of hormones. The responses involve the release inside the  cell of molecules collectively known as second messengers, that is, intracellular  mediators of the externally received hormonal message. Different receptors cause  the release of different second messengers. cAMP, identified by Earl Sutherland  in the 1950s, was the first second messenger discovered. Ca 2+ , as released from  intracellular reservoirs into the cytosol, is also a common second messenger.  Receptors and second messengers are discussed in greater depth in Chapter 13.   When hormonal stimulation increases the intracellular cAMP concentra¬  tion, PKA activity increases, increasing the rates of phosphorylation of many  proteins and decreasing their dephosphorylation rates as well. Because of the  cascade nature of the regulatory system diagrammed in Fig. 16-13, a small  change in [cAMP] results in a large change in the fraction of phosphorylated en¬  zymes. When a large fraction of the glycogen metabolism enzymes are phos¬  phorylated, the metabolic flux is in the direction of glycogen breakdown, since  glycogen phosphorylase is active and glycogen synthase is inactive. When  [cAMP] decreases, phosphorylation rates decrease, dephosphorylation rates in¬  crease, and the fraction of enzymes in their dephospho forms increases. The  resulting activation of glycogen synthase and inhibition of glycogen phos¬  phorylase cause the flux to shift to net glycogen synthesis.   Glucagon binding to its receptor on liver cells, which generates intracel¬  lular cAMP, results in glucose mobilization from stored glycogen (Fig. 16-14).  Glucagon is released from the pancreas when the concentration of circulating  glucose decreases to less than ~5 mM, such as during exercise or several hours  after a meal has been digested. Glucagon is therefore critical for the livers  function in supplying glucose to tissues that depend primarily on glycolysis  for their energy needs. Muscle cells do not respond to glucagon because they  lack the appropriate receptor.   flight” hormones, are released into the bloodstream by the adrenal glands in  response to stress. There are two types of receptors for these hormones: the  P-adrenergic receptor, which is linked to the adenylate cyclase system, and  the ot-adrenergic receptor, whose second messenger causes intracellular  [Ca 2+ ] to increase (Section 13-4A). Muscle cells, which have the (3-adrenergic  receptor (Fig. 16-14), respond to epinephrine by breaking down glycogen for  glycolysis, thereby generating ATP and helping the muscles cope with the stress  that triggered the epinephrine release.   Liver cells respond to epinephrine directly and indirectly. Epinephrine pro¬  motes the release of glucagon from the pancreas, and glucagon binding to its  receptor on liver cells stimulates glycogen breakdown as described above.   Epinephrine also binds directly to both a- and (3-adrenergic receptors on the  surfaces of liver cells (Fig. 16-14). Binding to the |3-adrenergic receptor results  in increased intracellular cAMP, which leads to glycogen breakdown.   Epinephrine binding to the a-adrenergic receptor stimulates an increase in in¬  tracellular [Ca 2+ ], which reinforces the cells’ response to cAMP (recall that  phosphorylase kinase, which activates glycogen phosphorylase and inactivates  glycogen synthase, is fully active only when phosphorylated and in the  presence of increased [Ca 2 ]). In addition, glycogen synthase is inactivated  through phosphorylation catalyzed by several Ca 2+ -dependent protein kinases.   Insulin and Epinephrine Are Antagonists. Insulin is released from the pancreas  in response to high levels of circulating glucose (e.g., immediately after a meal).   Hormonal stimulation by insulin increases the rate of glucose transport into  the many types of cells that have both insulin receptors and insulin-sensitive   Epinephrine binding to p-adrenergic receptors on liver and muscle  cells increases intracellular [cAMP], which promotes glycogen  degradation to G6P for glycolysis (in muscle) or to glucose for export  (in liver). The liver responds similarly to glucagon. Epinephrine binding  to a-adrenergic receptors on liver cells leads to increased cytosolic  [Ca 2+ ], which also promotes glycogen degradation. When circulating    glucose is plentiful, insulin stimulates glucose uptake and glycogen  synthesis in muscle cells. The liver responds both to insulin and  directly to increased glucose by increasing glycogen synthesis.    Explain why a given hormone elicits different responses in  different tissues.   • Summarize the effects of AMP and G6P  on glycogen phosphorylase and glycogen  synthase.   • Summarize the effects of phosphorylation  and dephosphorylation on glycogen phos¬  phorylase and glycogen synthase.   • Why does a phosphorylation/dephosphory¬  lation system allow more sensitive regula¬  tion of a metabolic process than a simple  allosteric system?   • Draw a simple diagram, similar to Fig.  16-13, showing how a kinase and a phos¬  phatase can regulate the activities of two  enzymes that catalyze opposing processes.   • How does regulation of glycogen metabo¬  lism differ between liver and muscle?   • Summarize the effects of insulin,  glucagon, and epinephrine on glycogen  metabolism.   • What are the intracellular effects of cAMP  and Ca 2+ ?    glucose transporters called GLUT4 on their surfaces (e.g., muscle and fat cells,  but not liver and brain cells). In addition, [cAMP] decreases, causing glycogen  metabolism to shift from glycogen breakdown to glycogen synthesis (Fig. 16-14).  The mechanism of insulin action is very complex (Sections 13-4D and 22-2),  but one of its target enzymes appears to be PP1. As outlined in Fig. 16-12,  insulin activates insulin-stimulated protein kinase in muscle to phosphorylate  site 1 on the glycogen-binding G M subunit of PP1 so as to activate this  protein and thus dephosphorylate the enzymes of glycogen metabolism. The  storage of glucose as glycogen is thereby promoted through the inhibition of  glycogen breakdown and the stimulation of glycogen synthesis.   In liver, insulin stimulates glycogen synthesis as a result of the inhibition  of glycogen synthase kinase 3P (GSK3[5; Fig. 13-31). This action decreases  the phosphorylation of glycogen synthase, thus increasing its activity. In addi¬  tion, it is thought that glucose itself may be a messenger to which the glyco¬  gen metabolism system responds. Glucose inhibits phosphorylase a by binding to  the enzyme's inactive T state and thereby shifting the T ^ R equilibrium toward  the T state (Fig. 12-17). This conformational shift exposes the Ser 14 phos-  phoryl group to dephosphorylation. An increase in glucose concentration  therefore promotes inactivation of glycogen phosphorylase a through its  conversion to phosphorylase b. The subsequent release of phosphoprotein  phosphatase-1 activates glycogen synthase. Thus when glucose is plentiful, the  liver stores the excess as glycogen.    • The liver and kidney can synthesize glucose from lactate, pyruvate, and amino  acids.   • Gluconeogenesis is mostly the reverse of glycolysis with the pyruvate kinase  reaction bypassed by the pyruvate carboxylase and phosphoenolpyruvate  carboxykinase reactions, and the phosphofructokinase and hexokinase reactions  bypassed by phosphatase reactions.   • Glycolysis and gluconeogenesis are reciprocally regulated by allosteric effects,  phosphorylation, and changes in enzyme synthesis rates.    When dietary sources of glucose are not available and when the liver has ex¬  hausted its supply of glycogen, glucose is synthesized from noncarbohydrate  precursors by gluconeogenesis. In fact, gluconeogenesis provides a substantial  fraction of the glucose produced in fasting humans, even within a few hours  of eating. Gluconeogenesis occurs in liver and, to a lesser extent, in kidney.   The noncarbohydrate precursors that can be converted to glucose include the  glycolysis products lactate and pyruvate, citric acid cycle intermediates, and the car¬  bon skeletons of most amino acids. First, however, all these substances must be  converted to the four-carbon compound oxaloacetate (at left), which itself is  a citric acid cycle intermediate (Section 17-1). The only amino acids that can¬  not be converted to oxaloacetate in animals are leucine and lysine because their  breakdown yields only acetyl-CoA (Section 21-4E) and because there is no  pathway in animals for the net conversion of acetyl-CoA to oxaloacetate. Likewise,  fatty acids cannot serve as glucose precursors in animals because most fatty  acids are degraded completely to acetyl-CoA (Section 20-2). However, fatty  acid breakdown generates much of the ATP that powers gluconeogenesis.   For convenience, we consider gluconeogenesis to be the pathway by which  pyruvate is converted to glucose. Most of the reactions of gluconeogenesis are  glycolytic reactions that proceed in reverse (Fig. 16-15). However, the gly¬  colytic enzymes hexokinase, phosphofructokinase, and pyruvate kinase catalyze  reactions with large negative free energy changes. These reactions must therefore   FIG. 1 6-1 5 Comparison of the pathways of gluconeogenesis and glycolysis. The   red arrows represent the steps that are catalyzed by different enzymes in gluconeogenesis.  The other seven reaction steps of gluconeogenesis are catalyzed by glycolytic enzymes that  function near equilibrium. O See the Animated Figures.    be replaced in gluconeogenesis by reactions that make glucose synthesis  thermodynamically favorable.   We begin our examination of the reactions unique to gluconeogenesis with  the conversion of pyruvate to phosphoenolpyruvate (PEP). Because this step  is the reverse of the highly exergonic reaction catalyzed by pyruvate kinase  (Section 15-2J), it requires free energy input. This is accomplished by  first converting the pyruvate to oxaloacetate. Oxaloacetate is a “high-energy”    FIG. 16-16 The conversion of pyruvate to carboxylase to convert pyruvate to oxaloacetate and (2) PEP   phosphoenolpyruvate (PEP). This process requires (1) pyruvate carboxykinase (PEPCK) to convert oxaloacetate to PEP    intermediate because its exergonic decarboxylation provides the free energy  necessary for PEP synthesis. The process requires two enzymes (Fig. 16-16):   1. Pyruvate carboxylase catalyzes the ATP-driven formation of oxalo¬  acetate from pyruvate and HC0 3 .   2. PEP carboxykinase (PEPCK) converts oxaloacetate to PEP in a reac¬  tion that uses GTP as a phosphoryl-group donor.   Pyruvate Carboxylase Has a Biotin Prosthetic Group. Pyruvate carboxylase is a  tetrameric protein of identical 160-residue subunits, each of which has a bi¬  otin prosthetic group. Biotin (Fig. 16-17a) functions as a C0 2 carrier by form¬  ing a carboxyl substituent at its ureido group (Fig. 16-17 b). Biotin is covalently  bound to an enzyme Lys residue to form a biocytin (alternatively, biotinyllysine)  residue (Fig. 16-17^). The biotin ring system is therefore at the end of a 14-A-  long flexible arm. Biotin, which was first identified in 1935 as a growth factor in  yeast, is an essential human nutrient. Its nutritional deficiency is rare, however, be¬  cause it occurs in many foods and is synthesized by intestinal bacteria.   Phase I The cleavage of ATP to ADP acts to dehydrate bicarbonate via  the formation of a “high-energy” carboxyphosphate intermedi¬  ate. The reaction of the resulting C0 2 with biotin is exergonic.  The biotin-bound carboxyl group is therefore “activated” rela¬  tive to bicarbonate and can be transferred to another molecule  without further free energy input.   Phase II The activated carboxyl group is transferred from carboxybiotin  to pyruvate in a three-step reaction to form oxaloacetate.   These two reaction phases occur on different subsites of the same enzyme; the  14-A arm of biocytin transfers the biotin ring between the two sites.   Oxaloacetate is both a precursor for gluconeogenesis and an intermediate of  the citric acid cycle (Section 17-3). When the citric acid cycle substrate acetyl-CoA  accumulates, it allosterically activates pyruvate carboxylase, thereby increasing the    FIG. 16-17 Biotin and carboxybiotinyl-enzyme. (a) Biotin  consists of an imidazoline ring that is cis-fused to a  tetrahydrothiophene ring bearing a valerate side chain. Positions 1, 2,  and 3 constitute a ureido group, (b) Biotin is covalently attached to   carboxylases by an amide linkage between its valeryl carboxyl group  and an e-amino group of an enzyme Lys side chain. The  carboxybiotinyl-enzyme forms when N1 of the biotin ureido group is  carboxylated.   C0 2 is produced at  the active site via  the elimination of  the biotinyl-enzyme.   Pyruvate donates a  proton to the biotinyl  group to generate  pyruvate enolate.    FIG. 16-18 The two-phase reaction mechanism of pyruvate carboxylase. [After Knowles, J.R., Annu. Rev. Biochem. 58, 217 (1989).]    amount of oxaloacetate that can participate in the citric acid cycle. When citric  acid cycle activity is low, oxaloacetate instead enters the gluconeogenic pathway.   PEP Carboxykinase Catalyzes the Formation of PEP. PEPCK, a monomeric  ~610-residue enzyme, catalyzes the GTP-requiring decarboxylation/  phosphorylation of oxaloacetate to form PEP and GDP (Fig. 16-19). Note that  the C0 2 that carboxylates pyruvate to yield oxaloacetate is eliminated in the    FIG. 16-19 The PEPCK mechanism. Decarboxylation of  oxaloacetate (a p-keto acid) forms a resonance-stabilized enolate    anion whose oxygen atom attacks the 7 -phosphoryl group of GTP,  forming PEP and GDP   Chapter 16 Glycogen Metabolism and formation of PEE The favorable decarboxylation reaction drives the formation of   Gluconeogenesis Requires Metabolite Transport between Mitochondria  and Cytosol. The generation of oxaloacetate from pyruvate or citric acid cy¬  cle intermediates occurs only in the mitochondrion, whereas the enzymes that  convert PEP to glucose are cytosolic. The cellular location of PEPCK varies: In  some species, it is mitochondrial; in some, it is cytosolic; and in some (includ¬  ing humans) it is equally distributed between the two compartments. In order  for gluconeogenesis to occur, either oxaloacetate must leave the mitochondrion  for conversion to PEP or the PEP formed there must enter the cytosol.   PEP is transported across the mitochondrial membrane by specific mem¬  brane transport proteins. There is, however, no such transport system for ox¬  aloacetate. In species with cytosolic PEPCK, oxaloacetate must first be converted  either to aspartate (Fig. 16-20, Route 1) or to malate (Fig. 16-20, Route 2),  for which mitochondrial transport systems exist. The difference between the  two routes involves the transport of NADH reducing equivalents (in the trans¬  port of reducing equivalents, the electrons—but not the electron carrier—cross  the membrane). The malate dehydrogenase route (Route 2) results in the  transport of reducing equivalents from the mitochondrion to the cytosol, since  it uses mitochondrial NADH and produces cytosolic NADH. The aspartate  aminotransferase route (Route 1) does not involve NADH. Cytosolic NADH  is required for gluconeogenesis, so, under most conditions, the route through  malate is a necessity. However, when the gluconeogenic precursor is lactate, its    FIG. 1 6-20 The transport of PEP and  oxaloacetate from the mitochondrion to the  cytosol. PEP is directly transported between the  compartments. Oxaloacetate, however, must first  be converted either to aspartate through the  action of aspartate aminotransferase (Route 1)  or to malate by malate dehydrogenase (Route 2).  Route 2 involves the mitochondrial oxidation of  NADH followed by the cytosolic reduction of  NAD + and therefore also transfers NADH  reducing equivalents from the mitochondrion to  the cytosol. Q See the Animated Figures.   can then be used. All the reactions shown in Fig. 16-20 are freely reversible, so  that under appropriate conditions, the malate-aspartate shuttle system also op¬  erates to transport NADH reducing equivalents into the mitochondrion for ox¬  idative phosphorylation (Section 18-IB). Liver has a variation of Route 1 in  which aspartate entering the cytosol is deaminated via the urea cycle before un¬  dergoing a series of reactions that yield oxaloacetate (Section 21-3A).    The route from PEP to fructose-1,6-bisphosphate (FBP) is catalyzed by the  enzymes of glycolysis operating in reverse. However, the glycolytic reactions cat¬  alyzed by phosphofructokinase (PFK) and hexokinase are endergonic in the gluco¬  neogenesis direction and hence must be bypassed by different gluconeogenic enzymes.  FBP is hydrolyzed by fructose-1,6-bisphosphatase (FBPase). The resulting  fructose-6-phosphate (F6P) is isomerized to G6P, which is then hydrolyzed by  glucose-6-phosphatase, the same enzyme that converts glycogen-derived G6P  to glucose (Section 16-1C) and which is present only in liver and kidney.  Note that these two hydrolytic reactions release P z rather than reversing the  ATP —» ADP reactions that occur at this point in the glycolytic pathway.   The net energetic cost of converting two pyruvate molecules to one glucose  molecule by gluconeogenesis is six ATP equivalents: two each at the steps cat¬  alyzed by pyruvate carboxylase, PEPCK, and phosphoglycerate kinase. Since the  energetic profit of converting one glucose molecule to two pyruvate molecules  via glycolysis is two ATP (Section 15-1), the energetic cost of the futile cy¬  cle in which glucose is converted to pyruvate and then resynthesized is four ATP  equivalents. Such free energy losses are the thermodynamic price that must be  paid to maintain the independent regulation of two opposing pathways.   Although glucose is considered the endpoint of the gluconeogenic path¬  way, it is possible for pathway intermediates to be directed elsewhere, for ex¬  ample, through the transketolase and transaldolase reactions of the pentose  phosphate pathway (Section 15-6C) to produce ribose-5-phosphate. The G6P  produced by gluconeogenesis may not be hydrolyzed to glucose but may in¬  stead be converted to G1P for incorporation into glycogen.   The opposing pathways of gluconeogenesis and glycolysis, like glycogen syn¬  thesis and degradation, do not proceed simultaneously in vivo. Instead, the  pathways are reciprocally regulated to meet the needs of the organism. There  are three substrate cycles and therefore three potential points for regulating  glycolytic versus gluconeogenic flux (Fig. 16-21).    Fructose-2,6-Bisphosphate Activates Phosphofructokinase and Inhibits  Fructose-1,6-Bisphosphatase. The net flux through the substrate cycle created  by the opposing actions of PFK and FBPase (described in Section  15-4B) is determined by the concentration of fructose-2,6-bisphosphate (F2,6P).   F2,6P, which is not a glycolytic intermediate, is an extremely potent allosteric  activator of PFK and an inhibitor of FBPase.   The concentration of F2,6P in the cell depends on the balance between  its rates of synthesis and degradation by phosphofructokinase-2 (PFK-2) and    FIG. 16-21 Substrate cycles in glucose  metabolism. The interconversions of glucose  and G6P, F6P and FBP, and PEP and pyruvate  are catalyzed by different enzymes in the forward  and reverse directions so that all reactions are  exergonic (the AG values for the reactions in  liver are given in kJ • mol -1 ). [AG’s obtained  from Newsholme, E.A. and Leech, A.R.,  Biochemistry for the Medical Sciences, p. 448,  Wiley (1983).]    FIG. 1 6-22 The formation and degradation of (FBPase-2) occur on different domains of the same protein molecule.   p-D-fructose-2,6-bisphosphate (F2,6P). The enzymatic activities of The phosphorylation of the liver enzyme inactivates PFK-2 while  phosphofructokinase-2 (PFK-2) and fructose bisphosphatase-2 activating FBPase-2.    fructose bisphosphatase-2 (FBPase-2), respectively (Fig. 16-22). These en¬  zyme activities are located on different domains of the same ~100-kD ho-  modimeric protein. The bifunctional enzyme is regulated by a variety of  allosteric effectors and by phosphorylation and dephosphorylation as catalyzed  by PKA and a phosphoprotein phosphatase. Thus, the balance between glu-  coneogenesis and glycolysis is under hormonal control.   For example, when [glucose] is low, glucagon stimulates the production of  cAMP in liver cells. This activates PKA to phosphorylate the bifunctional en¬  zyme at a specific Ser residue, which inactivates the enzymes PFK-2 activity and  activates its FBPase-2 activity. The net result is a decrease in [F2,6P], which shifts  the balance between the PFK and FBPase reactions in favor of FBP hydrolysis  and hence increases gluconeogenic flux (Fig. 16-23). The concurrent increases  in gluconeogenesis and glycogen breakdown allow the liver to release glucose  into the circulation. Conversely, when the blood [glucose] is high, cAMP levels  decrease, and the resulting increase in [F2,6P] promotes glycolysis.   In muscle, which is not a gluconeogenic tissue, the F2,6P control system  functions quite differently from that in liver due to the presence of different  PFK-2/FBPase-2 isozymes. For example, hormones that stimulate glycogen  breakdown in heart muscle lead to phosphorylation of a site on the bifunc¬  tional enzyme that activates rather than inhibits PFK-2. The resulting increase  in F2,6P stimulates glycolysis so that glycogen breakdown and glycolysis are  coordinated. The skeletal muscle isozyme lacks a phosphorylation site alto¬  gether and is therefore not subject to cAMP-dependent control.   Other Allosteric Effectors Influence Gluconeogenic Flux. Acetyl-CoA acti¬  vates pyruvate carboxylase (Section 16-4A), but there are no known allosteric  effectors of PEPCK, which together with pyruvate carboxylase reverses the  pyruvate kinase reaction. Pyruvate kinase, however, is allosterically inhibited  in the liver by alanine, a major gluconeogenic precursor. Alanine is converted  to pyruvate by the transfer of its amino group to an a-keto acid to yield a  new amino acid and the a-keto acid pyruvate,    a process termed transamination (which is discussed in Section 21-2A). Liver  pyruvate kinase is also inactivated by phosphorylation, further increasing glu¬  coneogenic flux. Since phosphorylation also activates glycogen phosphorylase,  the pathways of gluconeogenesis and glycogen breakdown both flow toward  G6P, which is converted to glucose for export from the liver.   FIG. 1 6-23 Sequence of metabolic  events linking low blood [glucose] to  gluconeogenesis in liver.    The activity of hexokinase (or glucokinase, the liver isozyme) is also con¬  trolled, as we will see in Section 22-ID. The activity of glucose-6-phosphatase  is controlled as well but the process is complex and poorly understood.   The regulation of glucose metabolism occurs not only through allosteric effectors,  but also through long-term changes in the amounts of enzymes synthesized. Pancreatic  and adrenal hormones influence the rates of transcription and the stabilities of  the mRNAs encoding many of the regulatory proteins of glucose metabolism.  For example, insulin inhibits transcription of the gene for PEPCK, whereas high  concentrations of intracellular cAMP promote the transcription of the genes for  PEPCK, FBPase, and glucose-6-phosphatase, and repress transcription of the  genes for glucokinase, PFK, and the PFK-2/FBPase-2 bifunctional enzyme.    • The formation of glycosidic bonds in carbohydrates requires the energy of acti¬  vated nucleotide sugars.   • O-Linked oligosaccharides are synthesized by the sequential addition of sugars to  a protein.    The liver, by virtue of its mass and its metabolic machinery, is primarily respon¬  sible for maintaining a constant level of glucose in the circulation. Glucose pro¬  duced by gluconeogenesis or from glycogen breakdown is released from the liver  for use by other tissues as an energy source. Of course, glucose has other uses  in the liver and elsewhere, for example, in the synthesis of lactose (Box 16-4).   • What are the substrates for gluconeogen¬  esis? What role do fatty acids play in  gluconeogenesis?   • Describe the reactions of gluconeogene¬  sis. Which reactions are not shared with  glycolysis?   • What is the net energy cost for synthesiz¬  ing a molecule of glucose from two mole¬  cules of pyruvate?   • Describe the role of fructose-2,6-bisphos-  phate in regulating gluconeogenesis and  glycolysis.    Like sucrose in plants, lactose is a disaccharide that is synthe¬  sized for later use as a metabolic fuel, in this case, after digestion  by very young mammals. Lactose, or milk sugar, is produced in the  mammary gland by lactose synthase. In this reaction, the donor  sugar is UDP-galactose, which is formed by the epimerization of  UDP-glucose (Fig. 15-28). The acceptor sugar is glucose:    Thus, both saccharide units of lactose are ultimately derived from  glucose.   1. Galactosyltransferase, the catalytic subunit, occurs in many  tissues, where it catalyzes the reaction of UDP-galactose and  A/-acetylglucosamine to yield JV-acetyllactosamine, a  constituent of many complex oligosaccharides.   2. a-Lactalbumin, a mammary gland protein with no catalytic  activity, alters the specificity of galactosyltransferase so that it  uses glucose as an acceptor, rather than /V-acetylglucosamine,  to form lactose instead of A/-acetyllactosamine.   Synthesis of a-lactalbumin, whose sequence is —37% identical to  that of lysozyme (which also participates in reactions involving  sugars), is triggered by hormonal changes at parturition (birth),  thereby promoting lactose synthesis for milk production.   FIG. 1 6-24 Role of nucleotide sugars. These compounds are the glycosyl donors in  Nucleotide sugar oligosaccharide biosynthetic reactions as catalyzed by glycosyltransferases.    Nucleotide Sugars Power the Formation of Glycosidic Bonds. Glucose and  other monosaccharides (principally mannose, TV-acetylglucosamine, fucose, galac¬  tose, TV-acetylneuraminic acid, and ^acetylgalactosamine) occur in glycoproteins  and glycolipids. Formation of the glycosidic bonds that link sugars to each other  and to other molecules requires free energy input under physiological conditions  (A G ot = 16 kj • mol x ). This free energy, as we have seen in glycogen synthe¬  sis (Section 16-2A), is acquired through the synthesis of a nucleotide sugar from  a nucleoside triphosphate and a monosaccharide, thereby releasing PP Z , whose ex-  ergonic hydrolysis drives the reaction. The nucleoside diphosphate at the sugars  anomeric carbon atom is a good leaving group and thereby facilitates formation  of a glycosidic bond to a second sugar in a reaction catalyzed by a glycosyltrans-  ferase (Fig. 16-24). In mammals, most glycosyl groups are donated by UDP-sug-  ars, but fucose and mannose are carried by GDP, and sialic acid by CMP. In  plants, starch is built from glucose units donated by ADP-glucose, and cellulose  synthesis relies on ADP-glucose or CDP-glucose.    O-Linked Oligosaccharides Are Posttranslationally Formed. Nucleotide sug¬  ars are the donors in the synthesis of O-linked oligosaccharides and in the pro¬  cessing of the TV-linked oligosaccharides of glycoproteins (Section 8-3C).  O-Linked oligosaccharides are synthesized in the Golgi apparatus by the serial  addition of monosaccharide units to a completed polypeptide chain (Fig. 16-25).  Synthesis begins with the transfer, as catalyzed by GalNAc transferase, of  TV-acetylgalactosamine (GalNAc) from UDP-GalNAc to a Ser or Thr residue  on the polypeptide. The location of the glycosylation site is thought to be spec¬  ified only by the secondary or tertiary structure of the polypeptide. Glycosylation  continues with the stepwise addition of sugars such as galactose, sialic acid, TV-  acetylglucosamine, and fucose. In each case, the sugar residue is transferred from  its nucleotide sugar derivative by a corresponding glycosyltransferase.   FIG. 1 6-25 Synthesis of an O-linked  oligosaccharide chain. This pathway shows  the proposed steps in the assembly of a  carbohydrate moiety in canine submaxillary  mucin. SA is sialic acid.   synthesis of TV-linked oligosaccharides is more complicated than that of  O-linked oligosaccharides. In the early stages of TVdinked oligosaccharide syn¬  thesis, sugar residues are sequentially added to a lipid carrier, dolichol pyrophos¬  phate (Fig. 16-26). Dolichol is a long-chain polyisoprenol containing 17 to 21  isoprene units in animals and 14 to 24 units in fungi and plants. It anchors  the growing oligosaccharide to the endoplasmic reticulum membrane, where  the initial glycosylation reactions take place.   Although nucleotide sugars are the most common monosaccharide donors  in glycosyltransferase reactions, several mannosyl and glucosyl residues are  transferred to growing dolichol-PP-oligosaccharides from their corresponding  dolichol-P derivatives. Dolichol phosphate “activates” a sugar residue for sub¬  sequent transfer, as does a nucleoside diphosphate.   The construction of an TV-linked oligosaccharide begins, as is described in  Section 8-3C, by the synthesis of an oligosaccharide with the composition  (TV^acetylglucosamine) 2 (mannose) 9 (glucose) 3 . This occurs on a dolichol carrier in  a 12-step process catalyzed by a series of specific glycosyltransferases (Fig. 16-27).    FIG. 1 6-26 Dolichol pyrophosphate glycoside. The carbohydrate precursors of  /V-linked glycosides are synthesized as oligosaccharides attached to dolichol, a long-chain  polyisoprenol (n = 14-24) in which the a-isoprene unit is saturated.    FIG. 1 6-27 The pathway of dolichol-PP-oligosaccharide  synthesis. (1) Addition of A/-acetylglucosamine-1-P and a second  A/-acetylglucosamine to dolichol-P. (2) Addition of five mannosyl residues  from GDP-mannose in reactions catalyzed by five different  mannosyltransferases. (3) Membrane translocation of  dolichol-PP-(/V-acetylglucosamine) 2 (mannose) 5 to the lumen of the  endoplasmic reticulum (ER). (4) Cytosolic synthesis of  dolichol-P-mannose from GDP-mannose and dolichol-P.   (5) Membrane translocation of dolichol-P-mannose to the lumen of  the ER. ( 6 ) Addition of four mannosyl residues from  dolichol-P-mannose in reactions catalyzed by four different    mannosyltransferases. (7) Cytosolic synthesis of dolichol-P-glucose  from UDPG and dolichol-P. (8) Membrane translocation of  dolichol-P-glucose to the lumen of the ER. (9) Addition of three  glucosyl residues from dolichol-P-glucose. (10) Transfer of the  oligosaccharide from dolichol-PP to the polypeptide chain at an Asn  residue in the sequence Asn-X-Ser/Thr, releasing dolichol-PP.   (11) Translocation of dolichol-PP to the cytoplasmic surface of the ER  membrane. (12) Hydrolysis of dolichol-PP to dolichol-P.   (13) Dolichol-P can also be formed by phosphorylation of dolichol by  CTP. [Modified from Abeijon, C. and Hirschberg, C.B., Trends Biochem.  Sci. 17, 34 (1992).] Q See the Animated Figures.   FIG. 1 6-28 The chemical structure of bacitracin. Note that this dodecapeptide has  four D-amino acid residues and two unusual intrachain linkages. “Orn” represents the  nonstandard amino acid ornithine (Fig. 21-9).   • List some of the synthetic pathways that  use nucleotide sugars.   • Explain the group transfer reaction of  glycosidic bond formation by describing  the attacking nucleophile, the atom at¬  tacked, and the leaving group.   • Explain why a lipid, dolichol, is involved in  glycoprotein synthesis.   • Describe the types of enzymatic reactions  that occur during the synthesis of A/-linked  oligosaccharides.    Note that some of these reactions take place on the lumenal surface of the en¬  doplasmic reticulum, whereas others occur on its cytoplasmic surface. Hence, on  four occasions (Reactions 3, 5, 8, and 11 in Fig. 16-27), dolichol and its attached  hydrophilic group are translocated, via unknown mechanisms, across the  endoplasmic reticulum membrane. In the final steps of the process, the oligosac¬  charide is transferred to the Asn residue in a segment of sequence Asn-X-Ser/Thr  (where X is any residue except Pro and only rarely Asp, Glu, Leu, or Trp) on a  growing polypeptide chain. The resulting dolichol pyrophosphate is hydrolyzed  to dolichol phosphate and P z , a process similar to the pyrophosphatase cleavage  of PP Z to 2 P z . Further processing of the oligosaccharide takes place, as described  in Section 8-3C, first in the endoplasmic reticulum and then in the Golgi appa¬  ratus (Fig. 8-19), where certain monosaccharide residues are trimmed away by  specific glycosylases and others are added by specific nucleotide sugar—requiring  glycosyl transferases.   Pyrophosphate. A number of compounds block the actions of specific gly-  cosylation enzymes, including bacitracin (Fig. 16-28), a cyclic polypeptide  that is a widely used antibiotic. Bacitracin forms a complex with dolichol py¬  rophosphate that inhibits its dephosphorylation (Fig. 16-27, Reaction 12),  thereby preventing the synthesis of glycoproteins from dolichol-linked  oligosaccharide precursors. Bacitracin is clinically useful because it inhibits bac¬  terial cell wall synthesis (which also involves dolichol-linked oligosaccharides)  but does not affect animal cells since it cannot cross cell membranes (bacter¬  ial cell wall synthesis is an extracellular process).   • Glycogen breakdown requires three enzymes. Glycogen phos-  phorylase converts the glucosyl units at the nonreducing ends of  glycogen to glucose-1-phosphate (G1P).   • Debranching enzyme transfers an a( 1^-4)-linked trisaccharide to  a nonreducing end and hydrolyzes the a(l—>6) linkage.   • Phosphoglucomutase converts G1P to glucose-6-phosphate  (G6P). In liver, G6P is hydrolyzed by glucose-6-phosphatase to  glucose for export to the tissues.   • Glycogen synthesis requires a different pathway in which G1P is  activated by reaction with UTP to form UDP-glucose.    • Glycogen synthase adds glucosyl units to the nonreducing ends of  a growing glycogen molecule that has been primed by glycogenin.   • Branching enzyme removes an a( 1^4)-linked 7-residue segment  and reattaches it through an a(l—>6) linkage to form a branched  chain.   • Glycogen metabolism is controlled in part by allosteric effectors  such as AMP, ATP, and G6P. Covalent modification of glycogen  phosphorylase and glycogen synthase shifts their T R equilib¬  ria and therefore alters their sensitivity to allosteric effectors.   • The ratio of phosphorylase a (more active) to phosphorylase b (less  active) depends on the activity of phosphorylase kinase, which is   regulated by the activity of protein kinase A (PKA), a cAMP-  dependent enzyme, and on the activity of phosphoprotein phos¬  phatase-1 (PP1). Glycogen phosphorylase is activated by  phosphorylation, whereas glycogen synthase is activated by  dephosphorylation.   • Hormones such as glucagon, epinephrine, and insulin control  glycogen metabolism. Hormone signals that generate cAMP as a  second messenger or that elevate intracellular Ca 2+ , which binds  to the calmodulin subunit of phosphorylase kinase, promote  glycogen breakdown. Insulin stimulates glycogen synthesis in part  by activating phosphoprotein phosphatase-1.   • Compounds that can be converted to oxaloacetate can subsequently  be converted to glucose. The conversion of pyruvate to glucose by  gluconeogenesis requires enzymes that bypass the three exergonic  steps of glycolysis: Pyruvate carboxylase and PEP carboxykinase   (PEPCK) bypass pyruvate kinase, fructose-1,6-bisphosphatase  (FBPase) bypasses phosphofructokinase, and glucose-6-phosphatase  bypasses hexokinase.   • Gluconeogenesis is regulated by changes in enzyme synthesis and  by allosteric effectors, including fructose-2,6-bisphosphate  (F2,6P), which inhibits FBPase and activates phosphofructokinase  (PFK) and whose synthesis depends on the phosphorylation state  of the bifunctional enzyme phosphofructokinase-2/fructose bis-  phosphatase-2 (PFK-2/FBPase-2).   • O-Finked oligosaccharides are synthesized by adding monosaccha¬  rides to a protein.   • 77-Finked oligosaccharides are assembled on a dolichol carrier and  then transferred to a protein.    1. Explain why the glycogen phosphorylase reaction (AG 0 ' =  3.1 kj • moF 1 2 3 4 * 6 7 8 ) is exergonic in the cell.   2. Glucose binds to glycogen phosphorylase and competitively in¬  hibits the enzyme. What is the physiological advantage of this?   3. Phosphoglucokinase catalyzes the phosphorylation of the C6-OH  group of G1P. Why is this enzyme important for the normal  function of phosphoglucomutase?   4. Glucose-6-phosphatase is located inside the endoplasmic reticu¬  lum. Describe the probable symptoms of a defect in G6P trans¬  port across the endoplasmic reticulum membrane.   3. Individuals with McArdle’s disease often experience a “second  wind” resulting from cardiovascular adjustments that allow glu¬  cose mobilized from liver glycogen to fuel muscle contraction.  Explain why the amount of ATP derived in the muscle from cir¬  culating glucose is less than the amount of ATP that would be  obtained by mobilizing the same amount of glucose from mus¬  cle glycogen.   6. A sample of glycogen from a patient with liver disease is incu¬  bated with P z , normal glycogen phosphorylase, and normal de¬  branching enzyme. The ratio of G1P to glucose formed in the  reaction mixture is 100. What is the patient’s most probable en¬  zymatic deficiency?   7. One molecule of dietary glucose can be oxidized through glycol¬  ysis and the citric acid cycle to generate a maximum of 32 mol¬  ecules of ATP. Calculate the fraction of this energy that is lost  when the glucose is stored as glycogen before it is catabolized.   8. The free energy of hydrolysis of an a(l—>4) glycosidic bond is  — 13.3 kj • mol \ whereas that of an a(l—>6) glycosidic bond  is —7.1 kj • mol 1 . Use these data to explain why glycogen de¬  branching includes three reactions [breaking and re-forming    a(l—>4) bonds and hydrolyzing a(l—>6) bonds], whereas glyco¬  gen branching requires only two reactions [breaking a(l—>4)  bonds and forming a(l—>6) bonds].   9. Calculations based on the volume of a glucose residue and the  branching pattern of cellular glycogen indicate that a glycogen  molecule could have up to 28 branching tiers before becoming  impossibly dense. What are the advantages of such a molecule  and why is it not found in vivo ?   10. Subjects who consumed a meal containing a high amylose:amy-  lopectin ratio showed a lower increase in blood glucose levels  than subjects who consumed a meal with a low amylose:amy-  lopectin ratio. Explain.   11. Many diabetics do not respond to insulin because of a deficiency  of insulin receptors on their cells. How does this affect (a) the  levels of circulating glucose immediately after a meal and (b) the  rate of glycogen synthesis in muscle?   12. Why does it make metabolic sense for the same hormone signal  to stimulate glycogenolysis and inhibit glycolysis in the liver while  stimulating both glycogenolysis and glycolysis in muscles?   13. Write the balanced equation for the sequential conversion of glu¬  cose to pyruvate and of pyruvate to glucose.   14. Write the balanced equation for the catabolism of six molecules  of G6P by the pentose phosphate pathway followed by conver¬  sion of ribulose-5-phosphate back to G6P by gluconeogenesis.   16. In the Cori cycle, the lactate product of glycolysis in muscle is  transformed back into glucose by the liver, (a) List the enzymes  involved in the lactate —> glucose pathway, (b) What is the net  gain/loss of ATP for one round of the Cori cycle (glucose —» lac¬  tate —> glucose)?   17. (a) What amino acid, other than alanine, can be converted by  transamination to a gluconeogenic precursor? (b) Calculate the  cost, in ATP equivalents, of converting two molecules of this  amino acid to one molecule of glucose.   18. The pathways of carbohydrate metabolism described in Chapters  15 and 16 include two reactions involving the transfer of one-  carbon groups, (a) Name the enzymes and the pathways where  these reactions occur, (b) What cofactors are required for the  transfer reactions?   19. Predict the effect of a fructose-1,6-bisphosphatase deficiency on  blood glucose levels (a) before and (b) after a 24-hour fast.   20. Explain why an individual with a fructose-1,6-bisphosphatase de¬  ficiency would have elevated levels of pyruvate in the blood.   21. Would you expect the allosteric regulator AMP to increase or de¬  crease fructose-1,6-bisphosphatase activity?   22. Explain why it makes metabolic sense for acetyl-CoA, which is not  a substrate for gluconeogenesis, to activate pyruvate carboxylase.   23. Fill in the blanks in the following diagram representing sucrose  synthesis in plants.    24. Starch synthase in plants contains a single active site that accom¬  modates ADP-glucose. However, cellulose synthase may contain  two UDP-glucose-binding active sites. Given the structural dif¬  ferences between starch and cellulose, explain why two substrate¬  binding sites would promote efficient cellulose synthesis. [Hint:  See Fig. 8-9.]   Focus concept: The structural characteristics of the lactate trans¬  port protein in hepatocytes are determined.   Case 26 The Role of Specific Amino Acids in the  Peptide Hormone Glucagon in Receptor Binding  and Signal Transduction   Focus concept: Amino acid side chains important in glucagon bind¬  ing and signal transduction are identified.    Use the Online Mendelian  Inheritance in Man (OMIM) database (http://www.ncbi.nlm.nih.  gov/omim) to explore one of the glycogen storage diseases de¬  scribed in Box 16-2. At what age is the disease typically noticed?  What are the clinical symptoms? How is a diagnosis made? How  is the disease treated? Are there variant forms of the disease, and  what is the genetic basis for the variation?    Bollen, M., Keppens, S., and Stalmans, W., Specific features of glycogen  metabolism in the liver, Biochem. J. 336, 19-31 (1998). [Describes  the activities of the enzymes involved in glycogen synthesis and  degradation and discusses the mechanisms for regulating the  processes.]   Brosnan, J.T., Comments on metabolic needs for glucose and the role of  gluconeogenesis, Eur. J. Clin. Nutr. 53, S107—S 111 (1999). [A read¬  able review that discusses possible reasons why carbohydrates are  used universally as metabolic fuels and why glucose is stored as  glycogen.]   Browner, M.E and Fletterick, R.J., Phosphorylase: a biological transducer,  Trends Biochem. Sci. 17, 66-71 (1992).   Burda, P. and Aebi, M., The dolichol pathway of N-linked glycosylation,  Biochim. Biophys. Acta 1426, 239-257 (1999).   Croniger, C.M., Olswang, Y., Reshef, L., Kalhan, S.C., Tilghman, S.M.,  and Hanson, R.W., Phosphoenolpyruvate carboxykinase revisited.  Insights into its metabolic role, Biochem. Mol. Biol. Educ. 30, 14-20  (2002); and Croniger, C.M., Chakravarty, K., Olswang, Y., Cassuto,  H., Reshef, L., and Hanson, R.W., Phosphoenolpyruvate carboxy¬  kinase revisited. II. Control of PEPCK-C gene expression, Biochem.  Mol. Biol. Educ. 30, 353-362 (2002).    Kishnani, P.S., Koeberi, D., and Chen, Y.T., Glycogen storage diseases,  in Valle, D. (Ed.), The Online Metabolic & Molecular Bases of Inher¬  ited Disease, Chapter 71. http://www.ommbid.com/. [Begins with a  review of glycogen metabolism.]   Melendez-Hevia, E., Waddell, T.G., and Shelton, E.D., Optimization of  molecular design in the evolution of metabolism: the glycogen mol¬  ecule, Biochem. J. 295, 477-483 (1993).   Nordlie, R.C., Foster, J.D., and Lange, A.J., Regulation of glucose pro¬  duction by the liver, Annu. Rev. Nutr. 19, 379-406 (1999).   Okar, D.A., Manzano, A., Navarro-Sabate, A., Riera, L., Bartrons, R.,  and Lange, A.J., PFK-2/FBPase-2: Maker and breaker of the essen¬  tial biofactor fructose-2,6-bisphosphate, Trends Biochem. Sci. 26,  30-35 (2001).   Roach, P.J. and Skurat, A.V., Self-glucosylating initiator proteins and their  role in glycogen biosynthesis, Prog. Nucleic Acid Res. Mol. Biol. 57,  289-316 (1997). [Discusses glycogenin.]   Whelan, W.J., Why the linkage of glycogen to glycogenin was so hard to  determine. Biochem. Mol. Biol. Educ. 35, 313-315 (2007).   B Aconitase Interconverts Citrate and Isocitrate  C NAD + -Dependent Isocitrate Dehydrogenase  Releases C0 2   D a-Ketoglutarate Dehydrogenase Resembles  Pyruvate Dehydrogenase  E Succinyl-CoA Synthetase Produces GTP  F Succinate Dehydrogenase Generates FADH 2  G Fumarase Produces Malate  H Malate Dehydrogenase Regenerates  Oxaloacetate    The metabolic activity of this germinating seed is geared to generate the energy and  carry out the biosynthetic reactions necessary for growth. Among the most active  pathways in this and most organisms is the citric acid cycle, a series of reactions that  participate in both catabolic and anabolic processes. [© Michael P. Gadomski/Photo  Researchers, Inc.]    In the preceding two chapters, we examined the catabolism of glucose and its  biosynthesis, storage, and mobilization. Although glucose is a source of energy  for nearly all cells, it is not the only metabolic fuel, nor is glycolysis the only  energy-yielding catabolic pathway. Cells that rely exclusively on glycolysis to  meet their energy requirements actually waste most of the chemical potential  energy of carbohydrates. When glucose is converted to lactate or ethanol, a  relatively reduced product leaves the cell. If the end product of glycolysis is  instead further oxidized, the cell can recover considerably more energy.   The oxidation of an organic compound requires an electron acceptor, such  as N0 3 , SO4 , Fe 3+ , or 0 2 , all of which are exploited as oxidants in dif¬  ferent organisms. In aerobic organisms, the electrons produced by oxidative  metabolism are ultimately transferred to 0 2 . Oxidation of metabolic fuels is  carried out by the citric acid cycle, a sequence of reactions that arose some¬  time after levels of atmospheric oxygen became significant, about 3 billion  years ago. As the reduced carbon atoms of metabolic fuels are oxidized to C0 2 ,  electrons are transferred to electron carriers that are subsequently reoxidized  by 0 2 . In this chapter, we examine the oxidation reactions of the citric acid  cycle itself. In the following chapter, we examine the fate of the electrons and  see how their energy is used to drive the synthesis of ATP.   A Pyruvate Dehydrogenase Is Regulated by  Product Inhibition and Covalent Modification   B Three Enzymes Control the Rate of the Citric  Acid Cycle   C The Glyoxylate Cycle Shares Some Steps with  the Citric Acid Cycle   FIG. 1 7-1 Overview of oxidative fuel metabolism. Acetyl groups derived from  carbohydrates, amino acids, and fatty acids enter the citric acid cycle, where they are  oxidized to C0 2 . © See the Animated Figures.    It is sometimes convenient to think of the citric acid cycle as an ad¬  dendum to glycolysis. Pyruvate derived from glucose can be split into C0 2  and a two-carbon fragment that enters the cycle for oxidation as acetyl-CoA  (Fig. 17-1). However, it is really misleading to think of the citric acid  cycle as merely a continuation of carbohydrate catabolism. The citric acid  cycle is a central pathway for recovering energy from several metabolic fuels,  including carbohydrates, fatty acids, and amino acids, that are broken down to  acetyl-CoA for oxidation. In fact, under some conditions, the principal func¬  tion of the citric acid cycle is to recover energy from fatty acids. We will  also see that the citric acid cycle supplies the reactants for a variety of  biosynthetic pathways.   We begin this chapter with an overview of the citric acid cycle. Next, we  explore how acetyl-CoA, its starting compound, is formed from pyruvate.  After discussing the reactions catalyzed by each of the enzymes of the cycle,  we consider the regulation of these enzymes. Finally, we examine the links be¬  tween citric acid cycle intermediates and other metabolic processes.   • The citric acid cycle is a multistep catalytic process that converts acetyl groups  derived from carbohydrates, fatty acids, and amino acids to C0 2 , and produces  NADH, FADH 2 , and GTP.    The citric acid cycle (Fig. 17-2) is an ingenious series of eight reactions that  oxidizes the acetyl group of acetyl-CoA to two molecules of C0 2 in a man¬  ner that conserves the liberated free energy in the reduced compounds NADH  and FADH 2 . The cycle is named after the product of its first reaction, citrate.  One complete round of the cycle yields two molecules of C0 2 , three NADH,  one FADH 2 , and one “high-energy” compound (GTP or ATP).   The citric acid cycle first came to light in the 1930s, when Hans Krebs,  building on the work of others, proposed a circular reaction scheme for the  interconversion of certain compounds containing two or three carboxylic acid  groups (that is, di- and tricarboxylates). At the time, many of the citric acid  cycle intermediates were already well known as plant products: citrate from  citrus fruit, aconitate from monkshood (. Aconitum ), succinate from amber  (succinum), fumarate from the herb Fumaria, and malate from apple (. Malus ).  Two other intermediates, oi-ketoglutarate and oxaloacetate, are known by  their chemical names because they were synthesized before they were identi¬  fied in living organisms. Krebs was the first to show how the metabolism of  these compounds was linked to the oxidation of metabolic fuels. His discov¬  ery of the citric acid cycle, in 1937, ranks as one of the most important  achievements of metabolic chemistry (Box 17-1). Although the enzymes and  intermediates of the citric acid cycle are now well established, many investi¬  gators continue to explore the molecular mechanisms of the enzymes and how  the enzymes are regulated for optimal performance under varying metabolic  conditions in different organisms.   Before we examine each of the reactions in detail, we should emphasize  some general features of the citric acid cycle:   1 . The circular pathway, which is also called the Krebs cycle or the  tricarboxylic acid (TCA) cycle, oxidizes acetyl groups from many    FIG. 1 7-2 The reactions of the citric acid cycle. The reactants  and products of this catalytic cycle are boxed. The pyruvate —> acetyl-  CoA reaction (top) supplies the cycle’s substrate via carbohydrate  metabolism but is not considered to be part of the cycle. An isotopic  label at C4 of oxaloacetate  becomes Cl of a-ketoglutarate and is  released as C0 2 in Reaction 4. An isotopic label at Cl of acetyl-CoA     becomes C5 of a-ketoglutarate and is scrambled in Reaction 5  between Cl and C4 of succinate (V 2 +). O See the Animated  Figures.    Without looking at the text, write an equation for the citric  acid cycle.    sources, not just pyruvate. Because it accounts for the major portion  of carbohydrate, fatty acid, and amino acid oxidation, the citric acid  cycle is often considered the “hub” of cellular metabolism.   2 . The net reaction of the citric acid cycle is   3 NADH + FADH 2 + GTP + CoA + 2 C0 2   The oxaloacetate that is consumed in the first step of the citric acid  cycle is regenerated in the last step of the cycle. Thus, the citric acid  cycle acts as a multistep catalyst that can oxidize an unlimited number of  acetyl groups.   3. In eukaryotes, all the enzymes of the citric acid cycle are located in the  mitochondria, so all substrates, including NAD + and GDP, must be   Hans Krebs worked in Otto Warburg’s  laboratory from 1926 until 1930 and later  declared that he had learned more from  Warburg (see Box 15-1) than from any other  teacher. Krebs applied Warburg’s tissue-slice  technique to the study of biosynthetic  reactions (Warburg himself was interested  primarily in oxidative and degradative  reactions). Over a period of years, Krebs investigated synthetic  pathways for urea, uric acid, and purines as well as oxidative  pathways. He was forced to leave his native Germany for England  in 1933, but unlike many other German emigrant scientists, he  was able to bring much of his laboratory equipment with him. In  England, Krebs continued to work on a series of metabolic  reactions that he named the citric acid cycle.   The cycle was discovered not through sudden inspiration but  through a series of careful experiments spanning the years 1932  to 1937. Krebs became interested in the “combustion” phase of  fuel use, namely, what occurs after the fermentation of glucose to  lactate. Until the 1930s, the mechanism of glucose oxidation and  its relationship to cellular respiration (oxygen uptake) was a  mystery. Krebs understood that the stoichiometry of the overall  process (glucose + 6 0 2 —» 6 C0 2 + 6 H 2 0) required a  multistep pathway. He was also aware that other researchers had  examined the ability of muscle tissue to rapidly oxidize various  dicarboxylates (a-ketoglutarate, succinate, and malate) and a  tricarboxylate (citrate), but none of these substances had a clear  relationship to any foodstuffs.   In 1935, Albert Szent-Gyorgyi found that cellular respiration  was dramatically accelerated by small amounts of succinate,  fumarate, malate, or oxaloacetate. In fact, the addition of any of  these compounds stimulated 0 2 uptake and C0 2 production far in  excess of what would be expected for their oxidation. In other  words, the compounds acted catalytically to boost the combustion  of other compounds in the cell. At about the same time, Carl  Martius and Franz Knoop showed that citrate could be converted  to a-ketoglutarate. Soon, Krebs had an entire sequence of  reactions for converting citrate to oxaloacetate: citrate —>  aconitate —» isocitrate —» a-ketoglutarate —» succinate —»  fumarate —» malate —> oxaloacetate. However, in order for this    sequence of reactions to work catalytically, it must repeatedly  return to its starting point; that is, the first compound must be  regenerated. And there was still no obvious link to glucose  metabolism!   Krebs believed that citrate and the other intermediates were  involved in glucose combustion because they appeared to burn at  the same rate as foodstuffs and were the only substances that  did so. In addition, earlier work had shown that the three-carbon  compound malonate not only blocks the conversion of succinate  to fumarate, it blocks all combustion by living cells. In 1937,   Martius and Knoop provided Krebs with a key piece of  information: oxaloacetate and pyruvate could be converted to  citrate in the presence of hydrogen peroxide.   Krebs now had the missing link: Pyruvate is a product of  glucose metabolism, and its reaction with oxaloacetate to form  citrate closed off the linear series of reactions to form a cycle.   The idea of a circular pathway was not new to Krebs. He and  Kurt Henseleit had elucidated the four-step urea cycle in 1932  (Section 21-3). Krebs quickly showed that the reaction of pyruvate  with oxaloacetate to form citrate took place in living tissue and  that the rates of citrate synthesis and breakdown were high  enough to account for the observed fuel combustion in a variety  of tissue types.   Remarkably, Krebs’ initial report on the citric acid cycle was  rejected by Nature, a leading journal, before being accepted for  publication in the less prestigious Enzymologia. In his report, Krebs  established the major outlines of the pathway, although some  details were later revised. For example, the mechanism of citrate  formation (which involves acetyl-CoA rather than pyruvate) and the  participation of succinyl-CoA in the cycle were not immediately  appreciated. Coenzyme A was not discovered until 1945, and only  in 1951 was acetyl-CoA shown to be the intermediate that  condenses with oxaloacetate to form citrate. Work by Krebs and  others established that the citric acid cycle plays a major role in  the oxidation of amino acids and fatty acids. In fact, the pathway  accounts for approximately two-thirds of the energy derived from  metabolic fuels. Krebs also recognized the role of the citric acid  cycle in supplying precursors for synthetic reactions.   [Krebs, H.A. and Johnson, W.A., The role of citric acid in intermediate metabolism in  animal tissues, Enzymologia 4, 148-156 (1937).]    generated in the mitochondria or be transported into mitochondria  from the cytosol. Similarly all the products of the citric acid cycle must  be consumed in the mitochondria or transported into the cytosol.   4. The carbon atoms of the two molecules of C0 2 produced in one round  of the cycle are not the two carbons of the acetyl group that began the  round (Fig. 17-2). These acetyl carbon atoms are lost in subsequent  rounds of the cycle. However, the net effect of each round of the cy¬  cle is the oxidation of one acetyl group to 2 C0 2 .   5. Citric acid cycle intermediates are precursors for the biosynthesis of other  compounds (e.g., oxaloacetate for gluconeogenesis; Section 16-4).   6 . The oxidation of an acetyl group to 2 C0 2 requires the transfer of  four pairs of electrons. The reduction of 3 NAD + to 3 NADH  accounts for three pairs of electrons; the reduction of FAD to FADH 2  accounts for the fourth pair. Much of the free energy of oxidation of  the acetyl group is conserved in these reduced coenzymes. Energy is  also recovered as GTP (or ATP). In Section 18-3C, we will see that  approximately 10 ATP are formed when the four pairs of electrons are  eventually transferred to 0 2 .   • Explain why the citric acid cycle is consid¬  ered to be the hub of cellular metabolism.   • What are the substrates and products of  the net reaction corresponding to one turn  of the citric acid cycle?    • Pyruvate dehydrogenase is a multienzyme complex that catalyzes a five-part reac¬  tion in which pyruvate releases C0 2 and the remaining acetyl group becomes  linked to coenzyme A.   • The reaction sequence requires the cofactors TPP, lipoamide, coenzyme A, FAD,  and NAD + .    Acetyl groups enter the citric acid cycle as part of the “high-energy” com¬  pound acetyl-CoA (recall that thioesters have high free energies of hydroly¬  sis; Section 14-2D). Although acetyl-CoA can also be derived from fatty acids  (Section 20 - 2 ) and some amino acids (Section 21-4), we focus here on the  production of acetyl-CoA from pyruvate derived from carbohydrates.   As we saw in Section 15-3, the end product of glycolysis under anaerobic  conditions is lactate or ethanol. However, under aerobic conditions, when the  NADH generated by glycolysis is reoxidized in the mitochondria, the final  product is pyruvate. A transport protein imports pyruvate along with H + (i.e.,  a pyruvate—H + symport) into the mitochondrion for further oxidation.   Multienzyme complexes are groups of noncovalently associated enzymes that  catalyze two or more sequential steps in a metabolic pathway. Virtually all or¬  ganisms contain multienzyme complexes, which represent a step forward in the  evolution of catalytic efficiency because they offer the following advantages:   1 . Enzymatic reaction rates are limited by the frequency with which en¬  zymes collide with their substrates (Section 11-3D). When a series of  reactions occurs within a multienzyme complex, the distance that sub¬  strates must diffuse between active sites is minimized, thereby enhanc¬  ing the reaction rate.   2 . The channeling (passing) of metabolic intermediates between successive  enzymes in a metabolic pathway reduces the opportunity for these inter¬  mediates to react with other molecules, thereby minimizing side reactions.   3. The reactions catalyzed by a multienzyme complex can be coordinately  controlled.    FIG. 1 7-3 Electron micrographs of the  E. coli pyruvate dehydrogenase multienzyme  complex, (a) The intact complex, (b) The  dihydrolipoyl transacetylase (E2) core complex.  [Courtesy of Lester Reed, University of Texas at  Austin.]    Acetyl-CoA is formed from pyruvate through oxidative decarboxylation by  a multienzyme complex named pyruvate dehydrogenase. This complex con¬  tains multiple copies of three enzymes: pyruvate dehydrogenase (El), dihy¬  drolipoyl transacetylase (E2), and dihydrolipoyl dehydrogenase (E3).   The E. coli pyruvate dehydrogenase complex is an ~4600-kD particle  with a diameter of about 300 A (Fig. 17-3 d). The core of the particle is made  of 24 E2 proteins arranged in a cube (Figs. 17-3 ^ and 17-4#), which is sur¬  rounded by 24 El proteins and 12 E3 proteins (Fig. 17-4/,c). In mammals,  yeast, and some bacteria, the pyruvate dehydrogenase complex is even larger  and more complicated, although it catalyzes the same reactions using homolo¬  gous enzymes and similar mechanisms. In these ~10,000-kD complexes, the  largest known multienzyme complexes, the E2 core consists of 60 subunits  arranged with dodecahedral symmetry [Fig. 17-5; a dodecahedron is a regular  polyhedron with / symmetry (Fig. 6-34 c) that has 20 vertices and 12 pentago¬  nal faces] surrounded by a shell consisting of ~45 El a 2 (3 2 heterotetramers and  ~9 E3 homodimers. El and E3 competitively bind to mutually exclusive sites  on E2 in a random distribution. Mammalian complexes, in addition, contain  ~12 copies of E3 binding protein, which facilitates the binding of E3 to the  E2 core, and several copies of a kinase and a phosphatase that function to reg¬  ulate the activity of the complex (Section 17-4A).   FIG. 1 7-4 Structural organization of the E. coli pyruvate dehydrogenase  multienzyme complex, (a) The dihydrolipoyl transacetylase (E2) core. The 24 E2 proteins  (green spheres ) associate as trimers at the corners of a cube, (b) The 24 pyruvate  dehydrogenase (El) proteins (orange spheres) form dimers that associate with the E2 core  (.shaded cube ) along its 12 edges. The 12 dihydrolipoyl dehydrogenase (E3) proteins  (purple spheres ) form dimers that attach to the six faces of the E2 cube, (c) Parts a and b  combined form the entire 60-subunit complex.    FIG. 1 7-5 Model of the Bacillus stearothermophilus pyruvate dehydrogenase  complex. This ~500-A-diameter cutaway surface diagram is based on cryoelectron  microscopic images of a complex consisting of a core of 60 E2 subunits ( gray-green )  surrounded by an outer shell of 60 El subunits ( purple ) separated by an ~90-A-wide  annular gap. Three full-length E2 subunits are schematically represented in red, green, and  yellow to indicate their catalytic domains (embedded in the E2 core), their peripheral  subunit-binding domains (embedded in the E3 shell) that bind the El (and E3) subunits,  and their lipoyl domains, all connected by flexible polypeptide linkers (Section 17-2B). Here  the red lipoyl domain is shown visiting an El active site (white dot), whereas the green and  yellow lipoyl domains occupy intermediate positions in the annular region between the E2  core and El shell. In a normal pyruvate dehydrogenase complex, E3 subunits would take  the place of some of the El subunits. Indeed, a complex of 60 E3 subunits and 60 E2  subunits closely resembles the E1-E2 complex. [Courtesy of Jacqueline Milne, National  Institutes of Health, Bethesda, MD.]   The pyruvate dehydrogenase complex catalyzes five sequential reactions with  the overall stoichiometry    Pyruvate + CoA + NAD + —> acetyl-CoA + C0 2 + NADH   Five different coenzymes are required: thiamine pyrophosphate (TPP;  Section 15-3B), lipoamide, coenzyme A (Fig. 14-11), FAD (Fig. 14-13),  and NAD + (Fig. 11-4). The coenzymes and their mechanistic functions are  listed in Table 17-1. The sequence of reactions catalyzed by the pyruvate  dehydrogenase complex is as follows (Fig. 17-6):   1. Pyruvate dehydrogenase (El), a TPP-requiring enzyme, decarboxylates  pyruvate with the formation of a hydroxyethyl-TPP intermediate:   This reaction is identical to that catalyzed by yeast pyruvate decarboxylase  (Fig. 15-20). Recall (Section 15-3B) that the ability of TPP s thiazolium    FIG. 1 7-6 The five reactions of the pyruvate dehydrogenase  multienzyme complex. El (pyruvate dehydrogenase) contains TPP  and catalyzes Reactions 1 and 2. E2 (dihydrolipoyl transacetylase)  contains lipoamide and catalyzes Reaction 3. E3 (dihydrolipoyl    dehydrogenase) contains FAD and a redox-active disulfide and  catalyzes Reactions 4 and 5.    What type of reaction occurs in each of the five steps?    ring to add to carbonyl groups and act as an electron sink makes it the  coenzyme most utilized in a-keto acid decarboxylation reactions.   2 . Unlike pyruvate decarboxylase, however, pyruvate dehydrogenase does  not convert the hydroxyethyl-TPP intermediate into acetaldehyde and  TPP. Instead, the hydroxyethyl group is transferred to the next en¬  zyme, dihydrolipoyl transacetylase (E2), which contains a lipoamide  group. Lipoamide consists of lipoic acid linked via an amide bond to  the 8-amino group of a Lys residue (Fig. 17-7). The reactive center  of lipoamide is a cyclic disulfide that can be reversibly reduced to yield  dihydrolipoamide. The hydroxyethyl group derived from pyruvate at¬  tacks the lipoamide disulfide, and TPP is eliminated, thus regenerating    FIG. 1 7-7 Interconversion of lipoamide  and dihydrolipoamide. Lipoamide consists of  lipoic acid covalently joined to the e-amino group  of a Lys residue via an amide bond.   active El. The hydroxyethyl carbanion is oxidized to an acetyl group  as the lipoamide disulfide is reduced:    3. E2 then catalyzes a transesterification reaction in which the acetyl group  is transferred to Co A, yielding acetyl-CoA and dihydrolipoamide-E2:   4. Acetyl-CoA has now been formed, but the lipoamide group of E2 must  be regenerated. Dihydrolipoyl dehydrogenase (E3) reoxidizes dihy-  drolipoamide to complete the catalytic cycle of E2. Oxidized E3 con¬  tains a reactive Cys—Cys disulfide group and a tightly bound FAD. The  oxidation of dihydrolipoamide is a disulfide interchange reaction:    is reoxidized. The sulfhydryl groups are reoxidized  which FAD funnels electrons to NAD + , yielding    FIG. 1 7-8 Active site of dihydrolipoamide  dehydrogenase (E3). In this X-ray structure of  the enzyme from Pseudomonas putida, the  redox-active portions of the bound NAD + and  FAD cofactors, the side chains of Cys 43 and  Cys 48 forming the redox-active disulfide bond,  and the side chain of Tyr 181 are shown in  ball-and-stick form with FAD C green, NAD + C  cyan, protein side chain C magenta, N blue, 0  red, P orange, and S yellow. Note that the side  chain of Tyr 181 is interposed between the flavin  and the nicotinamide rings. Portions of the  homodimeric protein’s two subunits are drawn in  ribbon form in purple and light orange. [Based  on an X-ray structure by Wim Hoi, University of  Washington. PDBid 1 LVL]   • Describe the five reactions of the pyruvate  dehydrogenase multienzyme complex.   • What cofactors are required? Which of  these are prosthetic groups?    The X-ray structure of dihydrolipoyl dehydrogenase together with mech¬  anistic information indicate that the reaction catalyzed by dihydrolipoamide  dehydrogenase (E3) is more complex than Reactions 4 and 5 in Fig. 17-6 sug¬  gest. The enzymes redox-active disulfide bond occurs between Cys 43 and Cys  48, which reside on a highly conserved segment of the enzymes polypeptide  chain. The disulfide bond links successive turns in a distorted segment of an  a helix (in an undistorted helix, the C a atoms of Cys 43 and Cys 48 would  be too far apart to permit the disulfide bond to form). The enzymes flavin  group is almost completely buried in the protein, which prevents the surround¬  ing solution from interfering with the electron-transfer reaction catalyzed by  the enzyme. The nicotinamide ring of NAD + binds on the side of the flavin  opposite the disulfide. In the absence of NAD + , the phenol side chain of Tyr  181 covers the nicotinamide-binding pocket so as to shield the flavin from  contact with the solution (Fig. 17-8). The Tyr side chain apparently moves  aside to allow the nicotinamide ring to bind near the flavin ring.   FAD prosthetic groups in proteins have standard reduction potentials of  around 0 V (Table 14-4), which makes FADH 2 unsuitable for donating elec¬  trons to NAD + ( < g°' = —0.315 V). Evidence suggests that the FAD group in  dihydrolipoamide dehydrogenase never becomes fully reduced as FADH 2 . Due  to the precise positioning of the flavin and nicotinamide ring, electrons are  rapidly transferred from the enzyme disulfide through FAD to NAD + , so a re¬  duced flavin anion (FADH ) has but a transient existence. Thus, FAD appears  to function more as an electron conduit than as a source or sink of electrons.   A Swinging Arm Transfers Intermediates. How are reaction intermediates chan¬  neled between E2 (the core of the pyruvate dehydrogenase complex) and the El  and E3 proteins on the outside? The key is the lipoamide group of E2. The lipoic  acid residue and the side chain of the Fys residue to which it is attached have a  combined length of about 14 A. This lipoyllysyl arm ( at left) is linked to the  rest of the E2 protein by a highly flexible and >l40-A-long, Pro- and Ala-rich  segment, which together function as a long tether that swings the disulfide group  from El (where it acquires a hydroxyethyl group), to the E2 active site (where  the hydroxyethyl group is transferred to form acetyl-CoA), and from there to E3  (where the reduced disulfide is reoxidized). Because of the flexibility and reach of  this assembly, one El protein can acetylate numerous E2 proteins, and one E3  protein can reoxidize several dihydrolipoamide groups. The flexible linkers pro¬  trude into the space between the E2 core and the E1-E3 outer shell (Fig. 17-5)  and swing around in order to “visit” the active sites of El, E2, and E3. The en¬  tire pyruvate dehydrogenase complex can be inactivated by the reaction of the  lipoamide group with certain arsenic-containing compounds (see Box 17-2).    The toxicity of arsenic has been known since ancient times. As(lll)  compounds such as arsenite (AsOl - ) and organic arsenicals   are toxic because they bind to sulfhydryl compounds (including  lipoamide) that can form bidentate adducts:   The inactivation of lipoamide-containing enzymes by arsenite,  especially the pyruvate dehydrogenase and a-ketoglutarate  dehydrogenase complexes, brings respiration to a halt. However,  organic arsenicals are more toxic to microorganisms than they are  to humans, apparently because of differences in the sensitivities  of their various enzymes to the compounds. This differential  toxicity is the basis for the early twentieth century use of organic  arsenicals in the treatment of syphilis (a bacterial disease) and  trypanosomiasis (a parasitic disease). These compounds were  actually the first antibiotics, although, not surprisingly, they  produced severe side effects.   Arsenic is often suspected as a poison in untimely deaths. It  was long thought that Napoleon Bonaparte died from arsenic    poisoning while in exile on the  island of St. Helena, a suspicion  that is strongly supported by the  recent finding that a lock  of his hair contains high levels  of arsenic. But was it murder  or environmental pollution? Ar¬  senic-containing dyes were used  in wallpaper at the time, and it  was eventually determined that  in damp weather, fungi convert  the arsenic to a volatile com¬  pound. Surviving samples of the  wallpaper from Napoleon’s room  in fact contain arsenic.  Napoleon’s arsenic poisoning  may therefore have been unin¬  tentional.   Charles Darwin may also  have been an unwitting victim of  chronic arsenic poisoning. In the  years following his epic voyage  on the Beagle, Darwin was  plagued by eczema, vertigo,  headaches, gout, and nausea-  all symptoms of arsenic poison¬  ing. Fowler’s solution, a widely  used nineteenth century “tonic,”  contained 10 mg of arsenite per  mL. Many individuals, quite  possibly Darwin himself, took  this “medication” for years.    • The eight enzymes of the citric acid cycle catalyze condensation, isomerization,  oxidation-reduction, phosphorylation, and hydration reactions.   • Two reactions produce C0 2 , one reaction produces GTP, and four reactions  generate the reduced coenzymes NADH or FADH 2 .    In this section, we discuss the eight enzymes of the citric acid cycle. The elu¬  cidation of the mechanisms for each of these enzymes is the result of an enor¬  mous amount of experimental work. Even so, there remain questions about  the mechanistic details of the enzymes and their regulatory properties.   Citrate synthase catalyzes the condensation of acetyl-CoA and oxaloacetate.  This initial reaction of the citric acid cycle is the point at which carbon atoms  (from carbohydrates, fatty acids, and amino acids) are “fed into the furnace”  as acetyl-CoA. The citrate synthase reaction proceeds with an Ordered  Sequential kinetic mechanism in which oxaloacetate binds before acetyl-CoA.   (a) The open conformation, (b) The closed, substrate-binding  conformation. In both forms, the homodimeric protein is viewed along  its twofold axis and is represented by its transparent molecular  surface with its polypeptide chains drawn in worm form colored in  rainbow order from their N-termini (blue) to their C-termini (red). The  reaction product citrate, which is bound to both enzymatic forms, and  coenzyme A, which is also bound to the closed form, are shown in   space-filling form with citrate C cyan, CoA C green, N blue, 0 red,  and P orange. The large conformational shift between the open and  closed forms entails 18° rotations of the small domains (upper left and  lower right) relative to the large domains resulting in relative  interatomic movements of up to 15 A. [Based on X-ray structures by  James Remington and Robert Huber, Max-Planck-lnstitut fur  Biochemie, Martinsried, Germany. PDBids 1CTS and 2CTS.] ©See  Interactive Exercise 18.    X-Ray studies show that the free enzyme (a homodimer) is in an “open”  form, with two domains that form a cleft containing the substrate-binding site  (Fig. 17-9 d). When substrate binds, the smaller domain undergoes a remark¬  able 18° rotation, which closes the cleft (Fig. 17-9 b). The existence of the  “open” and “closed” forms explains the enzymes Ordered Sequential kinetic  behavior. The conformational change generates the acetyl-CoA binding site and  seals the oxaloacetate binding site so that solvent cannot reach the bound substrate.  We have seen similar conformational changes in adenylate kinase (Fig. 14-10)  and hexokinase (Fig. 15-2), which prevent ATP hydrolysis.   In the reaction mechanism proposed by James Remington, three ionizable  side chains of citrate synthase participate in catalysis (Fig. 17-10):   1 . The enol of acetyl-CoA is generated in the rate-limiting step of the  reaction when Asp 375 (a base) removes a proton from the methyl  group. His 274 forms a hydrogen bond with the enolate oxygen.   2. Citryl-CoA is formed in a concerted acid—base catalyzed step, in which  the acetyl-CoA enolate (a nucleophile) attacks oxaloacetate. His 320  (an acid) donates a proton to oxaloacetate’s carbonyl group. The  citryl-CoA intermediate remains bound to the enzyme. Citrate synthase  is one of the few enzymes that directly forms a carbon-carbon bond  without the assistance of a metal ion cofactor.    Nucleophilic  attack of  acetyl-CoA  enolate on  oxaloacetate’s  carbonyl  carbon to  produce  citryl-CoA.    FIG. 1 7-10 The mechanism of the citrate synthase reaction. implicated as general acid-base catalysts. [Mostly after Remington,   His 274 and His 320 in their neutral forms and Asp 375 have been J.S., Curr. Opin. Struct. Biol. 2, 732 (1992).]    3. Citryl-CoA is hydrolyzed to citrate and CoA. This hydrolysis provides  the reactions thermodynamic driving force (AG°' = — 31.5 kj • mol l ).  We will see later why this reaction requires such a large, seemingly waste¬  ful, expenditure of free energy.   Aconitase catalyzes the reversible isomerization of citrate and isocitrate, with  czV-aconitate as an intermediate:    The reaction begins with a dehydration step in which a proton and an OH  group are removed. Since citrate has two carboxymethyl groups substituent to  its central C atom, it is prochiral rather than chiral. Thus, although water  might conceivably be eliminated from either of the two carboxymethyl arms,  aconitase removes water only from citrates lower ( pro-R ) arm (i.e., such that  the product molecule has the R configuration; Box 4-2).   Aconitase contains a [4Fe-4S] iron-sulfur cluster (an arrangement of four  iron atoms and four sulfur atoms, Section 18-2C) that presumably coordinates    the OH group of citrate to facilitate its elimination. Iron-sulfur clusters nor¬  mally participate in redox processes; aconitase is an intriguing exception.   The second stage of the aconitase reaction is rehydration of the double  bond of aTaconitate to form isocitrate. Although addition of water across the  double bond of aTaconitate could potentially yield four stereoisomers, aconi¬  tase catalyzes the stereospecific addition of OH and H + to produce only one  isocitrate stereoisomer. The ability of an enzyme to differentiate its substrates  pro-R and pro-S groups was not appreciated until 1948, when Alexander Ogston  pointed out that aconitase can distinguish between the two —CH 2 COO  groups of citrate when it is bound to the enzyme (Section 11-IB).   Isocitrate dehydrogenase catalyzes the oxidative decarboxylation of isocitrate to  a-ketoglutarate (also known as 2-oxoglutarate). This reaction produces the first  C0 2 and NADH of the citric acid cycle. Note that this C0 2 began the citric  acid cycle as a component of oxaloacetate, not of acetyl-CoA (Fig. 17-2).  (Mammalian tissues also contain an isocitrate dehydrogenase isozyme that uses  NADP + as a cofactor.)   NAD + -dependent isocitrate dehydrogenase, which also requires a Mn 2+  or Mg 2+ cofactor, catalyzes the oxidation of a secondary alcohol (isocitrate)  to a ketone (oxalosuccinate) followed by the decarboxylation of the carboxyl  group (3 to the ketone (Fig. 17-11). Mn 2+ helps polarize the newly formed  carbonyl group. The isocitrate dehydrogenase reaction mechanism is similar to  that of phosphogluconate dehydrogenase in the pentose phosphate pathway  (Section 15-6A).   The oxalosuccinate intermediate of the isocitrate dehydrogenase reaction  exists only transiently, and its existence was therefore difficult to confirm.  However, an enzymatic reaction can be slowed by mutating catalytically im¬  portant residues—in this case, Tyr 160 and Lys 230—to create kinetic “bot¬  tlenecks” so that reaction intermediates accumulate. Accordingly, crystals of  the mutant isocitrate dehydrogenase were exposed to the substrate isocitrate  and immediately visualized via X-ray crystallography using rapid measurement  techniques that require the highly intense X-rays generated by a synchrotron.  These studies revealed the oxalosuccinate intermediate in the active site of the  enzyme.   a-Ketoglutarate dehydrogenase catalyzes the oxidative decarboxylation of an  a-keto acid (a-ketoglutarate). This reaction produces the second C0 2 and  NADH of the citric acid cycle:   Again, this C0 2 entered the citric acid cycle as a component of oxaloacetate  rather than of acetyl-CoA (Fig. 17-2). Thus, although each round of the cit¬  ric acid cycle oxidizes two C atoms to C0 2 , the C atoms of the entering acetyl  groups are not oxidized to C0 2 until subsequent rounds of the cycle.   FIG. 1 7-1 1 The reaction mechanism of isocitrate dehydrogenase. Oxalosuccinate  is shown in brackets because it does not dissociate from the enzyme.    The a-ketoglutarate dehydrogenase reaction chemically resembles the re¬  action catalyzed by the pyruvate dehydrogenase multienzyme complex.  a-Ketoglutarate dehydrogenase is a multienzyme complex containing  a-ketoglutarate dehydrogenase (El), dihydrolipoyl transsuccinylase (E2),  and dihydrolipoyl dehydrogenase (E3). Indeed, this E3 is identical to the  E3 of the pyruvate dehydrogenase complex (a third member of the 2-keto  acid dehydrogenase family of multienzyme complexes is branched-chain  a-keto acid dehydrogenase, which participates in the degradation of  isoleucine, leucine, and valine; Section 21-4). The reactions catalyzed by the  a-ketoglutarate dehydrogenase complex occur by mechanisms identical to  those of the pyruvate dehydrogenase complex. Again, the product is a “high-  energy” thioester, in this case, succinyl-CoA.   Succinyl-CoA synthetase (also called succinate thiokinase) couples the  cleavage of the “high-energy” succinyl-CoA to the synthesis of a “high-energy”  nucleoside triphosphate (both names for the enzyme reflect the reverse re¬  action). Mammalian enzymes usually synthesize GTP from GDP + P z ,  whereas plant and bacterial enzymes usually synthesize ATP from ADP + P z .  These reactions are nevertheless energetically equivalent since ATP and GTP  are rapidly interconverted through the action of nucleoside diphosphate  kinase (Section 14-2C):   How does succinyl-CoA synthetase couple the exergonic cleavage of  succinyl-CoA (A G of = — 32.6 kj • mol X ) to the endergonic formation of  a nucleoside triphosphate (AG°' = 30.5 kj • mol x ) from the corresponding  nucleoside diphosphate and P z ? This question was answered by an experi¬  ment with isotopically labeled ADP. In the absence of succinyl-CoA, the  spinach enzyme catalyzes the transfer of the y-phosphoryl group from ATP  to [ 14 C]ADP, producing [ 14 C]ATP. Such an isotope-exchange reaction sug¬  gests the participation of a phosphoryl-enzyme intermediate that mediates  the reaction sequence    This information led to the isolation of a kinetically active phosphoryl-enzyme  in which the phosphoryl group is covalently linked to the N3 position of a  His residue. A three-step mechanism for succinyl-CoA synthetase is shown in   1. Succinyl-CoA reacts with P z to form succinyl-phosphate and CoA.   2 . The phosphoryl group is then transferred from succinyl-phosphate to  a His residue on the enzyme, releasing succinate.   3. The phosphoryl group on the enzyme is transferred to GDP, forming  GTP.   Note that in each of these steps, the energy of succinyl-CoA is conserved  through the formation of “high-energy” compounds: first, succinyl-phosphate,  then a 3-phospho-His residue, and finally GTP. The process is reminiscent of  passing a hot potato. The reaction catalyzed by succinyl-CoA synthetase is  another example of substrate-level phosphorylation (ATP synthesis that does  not directly depend on the presence of oxygen).   By this point in the citric acid cycle, one acetyl equivalent has been com¬  pletely oxidized to two C0 2 . Two NADH and one GTP (equivalent to one  ATP) have also been generated. In order to complete the cycle, succinate must  be converted back to oxaloacetate. This is accomplished by the cycle’s remain¬  ing three reactions.   This enzyme is strongly inhibited by malonate (at right ), a structural analog  of succinate and a classic example of a competitive inhibitor. When Krebs was  formulating his theory of the citric acid cycle, the inhibition of cellular respi¬  ration by malonate provided one of the clues that succinate plays a catalytic  role in oxidizing substrates and is not just another substrate.   Succinate dehydrogenase contains an FAD prosthetic group that is co¬  valently linked to the enzyme via a His residue (Fig. 17-13; in most other  FAD-containing enzymes, the FAD is held tightly but noncovalently). In  general, FAD functions biochemically to oxidize alkanes (such as succinate)  to alkenes (such as fumarate), whereas NAD + participates in the more ex-  ergonic oxidation of alcohols to aldehydes or ketones (e.g., in the reaction  catalyzed by isocitrate dehydrogenase). The dehydrogenation of succinate  produces FADH 2 , which must be reoxidized before succinate dehydrogenase  can undertake another catalytic cycle. The reoxidation of FADH 2 occurs  when its electrons are passed to the mitochondrial electron transport chain,  which we will examine in Section 18-2. Succinate dehydrogenase is the only  membrane-bound enzyme of the citric acid cycle (the others are components  of the mitochondrial matrix), so it is positioned to funnel electrons directly  into the electron transport machinery of the mitochondrial membrane.   of fumarate to form malate. The hydration reaction proceeds via a carbanion  transition state. OH addition occurs before H + addition:   Malate dehydrogenase catalyzes the final reaction of the citric acid cycle, the  regeneration of oxaloacetate. The hydroxyl group of malate is oxidized in an  NAD + -dependent reaction:   • Draw the structures of the eight intermedi¬  ates of the citric acid cycle and name the  enzymes that catalyze their interconversions.   • Which steps of the citric acid cycle re¬  lease C0 2 as a product? Which steps  produce NADH or FADH 2 ? Which step  produces GTP?   • Write the net equations for oxidation of  pyruvate, the acetyl group of acetyl-CoA,  and glucose to C0 2 and H 2 0.    Transfer of the hydride ion to NAD + occurs by the same mechanism used for  hydride ion transfer in lactate dehydrogenase and alcohol dehydrogenase (Section  15-3). X-Ray crystallographic comparisons of the NAD + -binding domains of  these three enzymes indicate that they are remarkably similar, consistent with the  proposal that all NAD + -binding domains evolved from a common ancestor.   The A G°' value for the malate dehydrogenase reaction is +29.7 kj • mol 1 ;  therefore, the concentration of oxaloacetate at equilibrium (and under cellu¬  lar conditions) is very low relative to malate. Recall, however, that the reac¬  tion catalyzed by citrate synthase, the first reaction of the citric acid cycle, is  highly exergonic (AG°' = — 31.5 kj • mol x ) because of the cleavage of the  thioester bond of citryl-CoA. We can now understand the necessity for such  a seemingly wasteful process. It allows citrate formation to be exergonic even  at the low oxaloacetate concentrations present in cells and thus helps keep the  citric acid cycle rolling.   FIG. 1 7-1 4 Products of the citric acid  cycle. For every two carbons that are oxidized to  C0 2 , electrons are recovered in the form of  three NADH and one FADH 2 . One GTP (or ATP)  is also produced.    • The need for energy regulates the citric acid cycle capacity at the pyruvate dehy¬  drogenase step and at the three rate-controlling steps of the cycle.   • Regulatory mechanisms depend on substrate availability, product inhibition, cova¬  lent modification, and allosteric effects.   The capacity of the citric acid cycle to generate energy for cellular needs is closely  regulated. The availability of substrates, the need for citric acid cycle intermedi¬  ates as biosynthetic precursors, and the demand for ATP all influence the oper¬  ation of the cycle. There is some evidence that the enzymes of the citric acid  cycle are physically associated, which might contribute to their coordinated  regulation. Before we examine the various mechanisms for regulating the  citric acid cycle, let us briefly consider the energy-generating capacity of  the cycle.   The oxidation of one acetyl group to two molecules of  C0 2 is a four-electron pair process (but keep in mind that it  is not the carbon atoms of the incoming acetyl group that  are oxidized). For every acetyl-CoA that enters the cycle,  three molecules of NAD + are reduced to NADH, which  accounts for three of the electron pairs, and one mole¬  cule of FAD is reduced to FADH 2 , which accounts for  the fourth electron pair. In addition, one GTP (or ATP)  is produced (Fig. 17-14).   The electrons carried by NADH and FADH 2 are fun-  neled into the electron-transport chain ,, which culminates  with the reduction of 0 2 to H 2 0. The energy of electron  transport is conserved in the synthesis of ATP by oxidative  phosphorylation (Section 18-3). For every NADH that passes    its electrons on, approximately 2.5 ATP are produced from  ADP + P z . For every FADH 2 , approximately 1.5 ATP are pro¬  duced. Thus, one turn of the citric acid cycle ultimately generates ap¬  proximately 10 ATP. We will see in Section 18-3C why these values are only  approximations.   When glucose is converted to two molecules of pyruvate by glycol¬  ysis, two molecules of ATP are generated and two molecules of NAD + are re¬  duced (Section 15-1). The NADH molecules yield approximately 5 molecules  of ATP on passing their electrons to the electron-transport chain. When the  two pyruvate molecules are converted to two acetyl-CoA by the pyruvate de¬  hydrogenase complex, the two molecules of NADH produced in that process   also eventually give rise to ~5 ATP. Two turns of the citric acid cycle (one for Section 4 Regulation of the Citric Acid Cycle   each acetyl group) generate ~20 ATP. Thus, one molecule of glucose can po¬  tentially yield ~32 molecules of ATP under aerobic conditions, when the cit¬  ric acid cycle is operating. In contrast, only 2 molecules of ATP are produced  per glucose molecule under anaerobic conditions.   A Pyruvate Dehydrogenase Is Regulated by Product Inhibition and  Covalent Modification   Given the large amount of ATP that can potentially be generated from car¬  bohydrate catabolism via the citric acid cycle, it is not surprising that the en¬  try of acetyl units derived from carbohydrate sources is regulated. The  decarboxylation of pyruvate by the pyruvate dehydrogenase complex is irre¬  versible, and since there are no other pathways in mammals for the synthesis  of acetyl-CoA from pyruvate, it is crucial that the reaction be precisely con¬  trolled. Two regulatory systems are used:    1. Product inhibition by NADH and acetyl-CoA . These compounds  compete with NAD + and CoA for binding sites on their respective en¬  zymes. They also drive the reversible transacetylase (E2) and dihy-  drolipoyl dehydrogenase (E3) reactions backward (Fig. 17-6). High  [NADH]/[NAD + ] and [acetyl-CoA]/[CoA] ratios therefore maintain  E2 in the acetylated form, incapable of accepting the hydroxy-   ethyl group from the TPP on El. This, in turn, ties up the  TPP on the El subunit in its hydroxyethyl form, decreasing  the rate of pyruvate decarboxylation.   2. Covalent modification by phosphorylation!dephosphoryla¬  tion of El. In eukaryotes, the products of the pyruvate de¬  hydrogenase reaction, NADH and acetyl-CoA, also activate  the pyruvate dehydrogenase kinase associated with the en¬  zyme complex. The resulting phosphorylation of a specific dehydroge¬  nase Ser residue inactivates the pyruvate dehydrogenase complex (Fig.  17-15). Insulin, the hormone that signals fuel abundance, reverses the  inactivation by activating pyruvate dehydrogenase phosphatase, which  removes the phosphate groups from pyruvate dehydrogenase. Recall  that insulin also activates glycogen synthesis by activating phosphopro-  tein phosphatase (Section 16-3B). Thus, in response to increases in   inactivated by the specific phosphorylation of  one of its Ser residues in a reaction catalyzed by  pyruvate dehydrogenase kinase. This phosphoryl  group is hydrolyzed through the action of  pyruvate dehydrogenase phosphatase, thereby  reactivating El.    blood [glucose], insulin promotes the synthesis of acetyl-CoA as well  as glycogen.   Other regulators of the pyruvate dehydrogenase system include pyruvate and  ADP, which inhibit pyruvate dehydrogenase kinase, and Ca 2+ , which inhibits  pyruvate dehydrogenase kinase and activates pyruvate dehydrogenase phosphatase.  In contrast to the glycogen metabolism control system (Section 16-3B), pyruvate  dehydrogenase activity is unaffected by cAMP.   B Three Enzymes Control the Rate of the Citric Acid Cycle   To understand how a metabolic pathway is controlled, we must identify the  enzymes that catalyze its rate-determining steps, the in vitro effectors of the  enzymes, and the in vivo concentrations of these substances. A proposed mech¬  anism of flux control must operate within the physiological concentration ranges  of the effectors.   Identifying the rate-determining steps of the citric acid cycle is more diffi¬  cult than it is for glycolysis because most of the cycles metabolites are present in  both mitochondria and cytosol and we do not know their distribution between  these two compartments (recall that identifying a pathways rate-determining steps  requires determining the A G of each of its reactions from the concentrations of  its substrates and products). However, we will assume that the compartments are  in equilibrium and use the total cell concentrations of these substances to esti¬  mate their mitochondrial concentrations. Table 17-2 gives the standard free en¬  ergy changes for the eight citric acid cycle enzymes and estimates of the  physiological free energy changes for the reactions in heart muscle or liver tissue.  We can see that three of the enzymes are likely to function far from equilibrium  under physiological conditions (negative A G): citrate synthase, NAD + -dependent iso¬  citrate dehydrogenase, and a-ketoglutarate dehydrogenase. These are therefore the  rate-determining enzymes of the cycle.   In heart muscle, where the citric acid cycle is active, the flux of metabolites  through the citric acid cycle is proportional to the rate of cellular oxygen con¬  sumption. Because oxygen consumption, NADH reoxidation, and ATP production  are tightly coupled (Section 18-3), the citric acid cycle must be regulated by feed¬  back mechanisms that coordinate NADH production with energy expenditure.  Unlike the rate-limiting enzymes of glycolysis and glycogen metabolism, which  regulate flux by elaborate systems of allosteric control, substrate cycles, and co¬  valent modification, the regulatory enzymes of the citric acid cycle seem to con¬  trol flux primarily by three simple mechanisms: (1) substrate availability, (2)  product inhibition, and (3) competitive feedback inhibition by intermediates    TABLE 1 7-2 Standard Free Energy Changes (AG°') and Physiological  Free Energy Changes (AG) of Citric Acid Cycle Reactions   FIG. 1 7-1 6 Regulation of the citric acid cycle. This diagram of the citric acid cycle,  which includes the pyruvate dehydrogenase reaction, indicates points of inhibition (red  octagons ) and the pathway intermediates that function as inhibitors (dashed red arrows).  ADP and Ca 2+ (green dots ) are activators. Q See the Animated Figures.    further along the cycle. Some of the major regulatory mechanisms are diagramed  in Fig. 17-16. There is no single flux-control point in the citric acid cycle; rather,  flux control is distributed among several enzymes.   Perhaps the most crucial regulators of the citric acid cycle are its substrates ,  acetyl-CoA and oxaloacetate , and its product , NADH. Both acetyl-CoA and  oxaloacetate are present in mitochondria at concentrations that do not saturate  citrate synthase. The metabolic flux through the enzyme therefore varies with  substrate concentration and is controlled by substrate availability. We have  already seen that the production of acetyl-CoA from pyruvate is regulated by  the activity of pyruvate dehydrogenase. The concentration of oxaloacetate,  which is in equilibrium with malate, fluctuates with the [NADH]/[NAD + ]  ratio according to the equilibrium expression   If, for example, the muscle workload and respiration rate increase, mitochon¬  drial [NADH] decreases. The consequent increase in [oxaloacetate] stimulates  the citrate synthase reaction, which controls the rate of citrate formation.   Aconitase functions close to equilibrium, so the rate of citrate consump¬  tion depends on the activity of NAD + -dependent isocitrate dehydrogenase,  which is strongly inhibited in vitro by its product NADH. Citrate synthase is  also inhibited by NADH but is less sensitive than isocitrate dehydrogenase to  changes in [NADH].   Other instances of product inhibition in the citric acid cycle are the inhi¬  bition of citrate synthase by citrate (citrate competes with oxaloacetate) and the   Chapter 17 Citric Acid Cycle inhibition of a-ketoglutarate dehydrogenase by NADH and succinyl-CoA.   Succinyl-CoA also competes with acetyl-CoA in the citrate synthase reaction  (competitive feedback inhibition). This interlocking system helps keep the cit¬  ric acid cycle coordinately regulated and the concentrations of its intermediates  within reasonable bounds.   • How much ATP can be generated from  glucose when the citric acid cycle is  operating?   • Which steps of the citric acid cycle  regulate flux through the cycle?   • Describe the role of ADP, Ca 2+ , acetyl-  CoA, and NADH in regulating pyruvate  dehydrogenase and the citric acid cycle.    Additional Regulatory Mechanisms. In vitro studies of citric acid cycle en¬  zymes have identified a few allosteric activators and inhibitors. ADP is an  allosteric activator of isocitrate dehydrogenase, whereas ATP inhibits the en¬  zyme. Ca 2+ , in addition to its many other cellular functions, regulates the  citric acid cycle at several points. It activates pyruvate dehydrogenase phos¬  phatase (Fig. 17-15), which in turn activates the pyruvate dehydrogenase  complex to produce acetyl-CoA. Ca 2+ also activates both isocitrate dehy¬  drogenase and a-ketoglutarate dehydrogenase (Fig. 17-16). Thus Ca 2+ , the  signal that stimulates muscle contraction, also stimulates the production of  the ATP to fuel it.    The citric acid cycle provides metabolites for gluconeogenesis, fatty acid synthe¬  sis, and amino acid synthesis.   Some organisms use the glyoxylate cycle, a variant of the citric acid cycle, for the  net conversion of acetyl-CoA to oxaloacetate.    At first glance, a metabolic pathway appears to be either catabolic, with the  release and conservation of free energy, or anabolic, with a requirement for  free energy. The citric acid cycle is catabolic, of course, because it involves  degradation and is a major free-energy conservation system in most organisms.  Cycle intermediates are required in only catalytic amounts to maintain the  degradative function of the cycle. However, several biosynthetic pathways  use citric acid cycle intermediates as starting materials for anabolic reactions.  The citric acid cycle is therefore amphibolic (both anabolic and catabolic). In  this section, we examine some of the reactions that feed intermediates into the  citric acid cycle or draw them off; we also examine the glyoxylate cycle, a  variation of the citric acid cycle that occurs only in plants and converts acetyl-  CoA to oxaloacetate. Some of the reactions that use and replenish citric acid  cycle intermediates are summarized in Fig. 17-17.   Reactions that utilize and therefore drain citric acid cycle intermediates are called  cataplerotic reactions (emptying; Greek: cata, down + plerotikos , to fill). These  reactions serve not only to synthesize important products but also to avoid the  inappropriate buildup of citric acid cycle intermediates in the mitochondrion,  for example, when there is a high rate of breakdown of amino acids to citric  acid cycle intermediates. Cataplerotic reactions occur in the following pathways:   1. Glucose biosynthesis (gluconeogenesis) utilizes oxaloacetate  (Section 16-4). Because gluconeogenesis takes place in the cytosol,  oxaloacetate must be converted to malate or aspartate for transport out  of the mitochondrion (Fig. 16-20). Since the citric acid cycle is a cycli¬  cal pathway, any of its intermediates can be converted to oxaloacetate  and used for gluconeogenesis.   2. Fatty acid biosynthesis is a cytosolic process that requires acetyl-CoA.  Acetyl-CoA is generated in the mitochondrion and is not transported    FIG. 1 7-1 7 Amphibolic functions of the  citric acid cycle. The diagram indicates the  positions at which intermediates are drawn off  by cataplerotic reactions for use in anabolic  pathways (red arrows ) and the points where  anaplerotic reactions replenish cycle  intermediates (green arrows). Reactions involving  amino acid transamination and deamination are  reversible, so their direction varies with metabolic  demand. O See the Animated Figures.    across the mitochondrial membrane. Cytosolic acetyl-CoA is therefore  generated by the breakdown of citrate, which can cross the membrane, in  a reaction catalyzed by ATP-citrate lyase (Section 20-4A). This reaction  uses the free energy of ATP to “undo” the citrate synthase reaction:   3. Amino acid biosynthesis uses a-ketoglutarate and oxaloacetate as  starting materials. For example, a-ketoglutarate is converted to gluta¬  mate by reductive amination catalyzed by a glutamate dehydrogenase  that utilizes either NADH or NADPH:   In aerobic organisms, the citric acid cycle is the major source of free energy,  and hence the catabolic function of the citric acid cycle cannot be interrupted:  Cycle intermediates that have been siphoned off must be replenished. The re¬  plenishing reactions are called anaplerotic reactions (filling up; Greek: ana,  up + plerotikos, to fill). The most important of these reactions is catalyzed by  pyruvate carboxylase, which produces oxaloacetate from pyruvate:   Pyruvate + C0 2 + ATP + H 2 0 —> oxaloacetate + ADP + P z   (This is also one of the first steps of gluconeogenesis; Section 16-4A). Pyruvate  carboxylase “senses” the need for more citric acid cycle intermediates through its  activator, acetyl-CoA. Any decrease in the rate of the cycle caused by insufficient ox¬  aloacetate or other intermediates allows the concentration of acetyl-CoA to rise. This  activates pyruvate carboxylase, which replenishes oxaloacetate. The reactions of  the citric acid cycle convert the oxaloacetate to citrate, a-ketoglutarate, succinyl-  CoA, and so on, until all the intermediates are restored to appropriate levels.   An increase in the concentrations of citric acid cycle intermediates sup¬  ports increased flux of acetyl groups through the cycle. For example, flux  through the citric acid cycle may increase as much as 60- to 100-fold in mus¬  cle cells during intense exercise. Not all of this increase is due to elevated  concentrations of cycle intermediates (which only increase about fourfold),  because other regulatory mechanisms (as described in Section 17-4B) also  promote flux through the rate-controlling steps of the cycle.   During exercise, some of the pyruvate generated by increased  glycolytic flux is directed toward oxaloacetate synthesis as catalyzed  by pyruvate carboxylase. Pyruvate can also accept an amino group  from glutamate (a transamination reaction) to generate alanine (the  amino acid counterpart of pyruvate) and the citric acid cycle inter¬  mediate a-ketoglutarate (the ketone counterpart of glutamate). Both  of these mechanisms help the citric acid cycle efficiently catabolize  the acetyl groups derived—also from pyruvate—by the reactions of the  pyruvate dehydrogenase complex. These reactions are sum¬  marized at left. The end result is increased production of ATP  to power muscle contraction.   Other metabolites that feed into the citric acid cycle are  succinyl-CoA, a product of the degradation of odd-chain fatty acids (Section  20-2E) and certain amino acids (Section 21-4), and a-ketoglutarate and ox¬  aloacetate, which are formed by the reversible transamination of certain amino  acids, as indicated above. The links between the citric acid cycle and other  metabolic pathways offer some clues to its evolution (Box 17-3).    C The Glyoxylate Cycle Shares Some Steps with the  Citric Acid Cycle   Plants , bacteria, and fungi, but not animals, possess enzymes that mediate the net  conversion of acetyl-CoA to oxaloacetate, which can be used for gluconeogenesis. In  plants, these enzymes constitute the glyoxylate cycle (Fig. 17-18), which oper¬  ates in two cellular compartments: the mitochondrion and the glyoxysome, a  membrane-bounded plant organelle that is a specialized peroxisome. Most of the  enzymes of the glyoxylate cycle are the same as those of the citric acid cycle.   Reactions 1 and 2. Glyoxysomal oxaloacetate is condensed with acetyl-  CoA to form citrate, which is isomerized to isocitrate  as in the citric acid cycle. Since the glyoxysome con¬  tains no aconitase, Reaction 2 presumably takes place  in the cytosol.   Glyoxysomal isocitrate lyase cleaves the isocitrate to Section 5 Reactions Related to the Citric Acid Cycle  succinate and glyoxylate (hence the cycles name).   Malate synthase, a glyoxysomal enzyme, condenses  glyoxylate with a second molecule of acetyl-CoA to  form malate.   Glyoxysomal malate dehydrogenase catalyzes the ox¬  idation of malate to oxaloacetate by NAD + .   FIG. 17-18 The glyoxylate cycle. The cycle results in the net   conversion of two acetyl-CoA to succinate in the glyoxysome, which  can be converted to malate in the mitochondrion for use in  gluconeogenesis. Isocitrate lyase and malate synthase, enzymes unique  to glyoxysomes (which occur only in plants), are indicated in blue.   (1) Glyoxysomal citrate synthase catalyzes the condensation of  oxaloacetate with acetyl-CoA to form citrate. (2) Cytosolic aconitase  catalyzes the conversion of citrate to isocitrate. (3) Isocitrate lyase  catalyzes the cleavage of isocitrate to succinate and glyoxylate.    (4) Malate synthase catalyzes the condensation of glyoxylate with  acetyl-CoA to form malate. (5) Glyoxysomal malate dehydrogenase  catalyzes the oxidation of malate to oxaloacetate, completing the cycle.  Succinate is transported to the mitochondrion, where it is converted  to malate via the citric acid cycle. This malate is transported to the  cytosol, where malate dehydrogenase catalyzes its oxidation to  oxaloacetate, which can then be used in gluconeogenesis.   Alternatively, malate can continue in the citric acid cycle, making the  glyoxylate cycle anaplerotic.    The citric acid cycle is ubiquitous in aerobic organisms and plays  a central role in energy metabolism in these cells. However, an  eight-step catalytic cycle such as the citric acid cycle is unlikely to  have arisen all at once and must have evolved from a simpler set  of enzyme-catalyzed reactions. Clues to its origins can be found  by examining the metabolism of cells that resemble early life-  forms. Such organisms emerged before significant quantities of  atmospheric oxygen became available some 3 billion years ago.  These cells may have used sulfur as their terminal oxidizing agent,  reducing it to H 2 S. Their modern-day counterparts are anaerobic  autotrophs that harvest free energy by pathways that are  independent of the pathways that oxidize carbon-containing   Methanococcus jannaschii, an organism without a citric acid cycle.  [Courtesy of B. Boonyaratanakornkit, D.S. Clark, and G. Vrdoljak,  University of California at Berkeley.]    compounds. These organisms therefore do not use the citric acid  cycle to generate reduced cofactors that are subsequently oxi¬  dized by molecular oxygen. However, all organisms must synthe¬  size the small molecules from which they can build proteins,  nucleic acids, carbohydrates, and lipids.   The task of divining an organism’s metabolic capabilities  has been facilitated through bioinformatics. By comparing the  sequences of prokaryotic genomes and assigning functions to  various homologous genes, it is possible to reconstruct the  central metabolic pathways for the organisms. This approach  has been fruitful because many “housekeeping” genes, which  encode enzymes that make free energy and molecular building  blocks available to the cell, are highly conserved among  different species and hence are relatively easy to recognize.   Genomic analysis reveals that many prokaryotes lack the citric  acid cycle. However, these organisms do contain genes for some  citric acid cycle enzymes. The last four reactions of the cycle,  leading from succinate to oxaloacetate, appear to be the most  highly conserved. This pathway fragment constitutes a mechanism  for accepting electrons that are released during sugar fermentation.  For example, the reverse of this pathway could regenerate NAD +  from the NADH produced by the glyceraldehyde-3-phosphate  dehydrogenase step of glycolysis.    The glyoxylate cycle therefore results in the net conversion of two acetyl-CoA to  succinate instead of to four molecules of CO 2 as would occur in the citric acid  cycle. The succinate produced in Reaction 3 is transported to the mitochon¬  drion where it enters the citric acid cycle and is converted to malate, which  has two alternative fates: (1) It can be converted to oxaloacetate in the mi¬  tochondrion, continuing the citric acid cycle and thereby making the gly¬  oxylate cycle an anaplerotic process (Section 17-5B); or (2) it can be  transported to the cytosol, where it is converted to oxaloacetate for entry  into gluconeogenesis.   The overall reaction of the glyoxylate cycle can be considered to be the  formation of oxaloacetate from two molecules of acetyl-CoA:   oxaloacetate + 2 CoA + 2 NADH + FADH 2 + 2 H +    The resulting succinate could then be used as a starting material  for the biosynthesis of other compounds.   that converts pyruvate to acetyl-CoA (but without producing  NADH). In a primitive cell, the resulting acetyl groups could  have condensed with oxaloacetate (by the action of a citrate  synthase), eventually giving rise to an oxidative sequence of  reactions resembling the first few steps of the modern citric  acid cycle.   The a-ketoglutarate produced in this way can be converted to  glutamate and other amino acids.   The reductive and oxidative branches of the citric acid cycle  outlined so far function in modern bacterial cells such as E. coli  cells when they are growing anaerobically, suggesting that similar  pathways could have filled the metabolic needs of early cells. The  evolution of a complete citric acid cycle in which the two branches  are linked and both proceed in an oxidative (clockwise) direction  would have required an enzyme such as a-ketoglutarate:ferredoxin  reductase (a homolog of pyruvate:ferredoxin oxidoreductase) to  link a-ketoglutarate and succinate.    Interestingly, a primitive citric acid cycle that operated in the  reverse (counterclockwise) direction could have provided a  route for fixing C0 2 (that is, incorporating C0 2 into biological  molecules).    The genes encoding enzymes that catalyze the steps of such a  pathway have been identified in several modern autotrophic  bacteria. This reductive pathway, which occurs in some deeply  rooted archaeal species, possibly predates the C0 2 -fixing pathway  used in some photosynthetic bacteria and in the chloroplasts of  green plants (Section 19-3A).    Isocitrate lyase and malate synthase do not occur in animals. In plants, these  enzymes enable germinating seeds to convert their stored triacylglycerols,  through acetyl-CoA, to glucose. It had long been assumed that this was a re¬  quirement of germination. However, a mutant of Arabidopsis thaliana (an  oilseed plant) lacking isocitrate lyase, and hence unable to convert lipids to  carbohydrate, nevertheless germinated. This process was only inhibited when  the mutant plants were subjected to low light conditions. Therefore, it now  appears that the glyoxylate cycles importance in seedling growth is its  anaplerotic function in providing four-carbon units to the citric acid cycle,  which can then oxidize the triacylglycerol-derived acetyl-CoA.   Organisms that lack the glyoxylate pathway cannot undertake the net syn¬  thesis of glucose from acetyl-CoA. This is the reason humans cannot convert  fats (that is, fatty acids, which are catabolized to acetyl-CoA) to carbohydrates  (that is, glucose).    • Explain how a catalytic cycle can supply  precursors for other metabolic pathways  without depleting its own intermediates.   • Which citric acid cycle intermediates can  be directly used in gluconeogenesis?  Which can be used for fatty acid synthe¬  sis? Which can be directly converted to  amino acids?   • Describe the reactions of the glyoxylate  cycle. Which two enzymes are unique to  this pathway? What does the pathway  accomplish?    T Some human pathogens use the glyoxylate cycle, sometimes to great  advantage. For example, Mycobacterium tuberculosis, which causes tubercu¬  losis, can persist for years in the lung without being attacked by the immune  system. During this period, the bacterium subsists largely on lipids, using  the citric acid cycle to produce precursors for amino acid synthesis and us¬  ing the glyoxylate cycle to produce carbohydrate precursors. Drugs that are  designed to inhibit the bacterial isocitrate lyase can therefore potentially  limit the pathogens survival. The virulence of the yeast Candida albicans ,  which often infects immunosuppressed individuals, may also depend on ac¬  tivation of the glyoxylate cycle when these yeast cells take up residence in¬  side macrophages.   • The eight enzymes of the citric acid cycle function in a multistep  catalytic cycle to oxidize an acetyl group to two C0 2 molecules  with the concomitant generation of three NADH, one FADH 2 ,  and one GTP.   • The free energy released when the reduced coenzymes ultimately  reduce 0 2 is used to generate ATP.   • Acetyl groups enter the citric acid cycle as acetyl-CoA. The pyru¬  vate dehydrogenase multienzyme complex, which contains three  types of enzymes and five types of coenzymes, generates acetyl-  CoA from the glycolytic product pyruvate.   • The lipoyllysyl arm of E2 acts as a tether that swings reactive  groups between enzymes in the complex.   • Citrate synthase catalyzes the condensation of acetyl-CoA and  oxaloacetate in a highly exergonic reaction.   • Aconitase catalyzes the isomerization of citrate to isocitrate, and  isocitrate dehydrogenase catalyzes the oxidative decarboxylation of  isocitrate to a-ketoglutarate to produce the citric acid cycle’s first  C0 2 and NADH.   • a-Ketoglutarate dehydrogenase catalyzes the oxidative decarboxy¬  lation of a-ketoglutarate to produce succinyl-CoA and the citric  acid cycle’s second C0 2 and NADH.   • Succinyl-CoA synthetase couples the cleavage of succinyl-CoA to  the synthesis of GTP (or in some organisms, ATP) via a phos-  phoryl-enzyme intermediate.    • The citric acid cycle’s remaining three reactions, catalyzed by suc¬  cinate dehydrogenase, fumarase, and malate dehydrogenase, regen¬  erate oxaloacetate to continue the citric acid cycle.   • Neither of the C0 2 molecules released in a given turn of the cit¬  ric acid cycle are derived from the acetyl group that entered the  same turn of the cycle. Instead, they are derived from the oxalo¬  acetate that was synthesized from the acetyl groups that entered  previous turns of the cycle.   • Entry of glucose-derived acetyl-CoA into the citric acid cycle is  regulated at the pyruvate dehydrogenase step by product inhibi¬  tion (by NADH and acetyl-CoA) and by covalent modification.   • The citric acid cycle itself is regulated at the steps catalyzed by cit¬  rate synthase, NAD + -dependent isocitrate dehydrogenase, and a-  ketoglutarate dehydrogenase.   • Regulation is accomplished mainly by substrate availability, prod¬  uct inhibition, and feedback inhibition.   • Cataplerotic reactions deplete citric acid cycle intermediates. Some  citric acid cycle intermediates are substrates for gluconeogenesis,  fatty acid biosynthesis, and amino acid biosynthesis.   • Anaplerotic reactions such as the pyruvate carboxylase reaction re¬  plenish citric acid cycle intermediates.   • The glyoxylate cycle, which operates only in plants, bacteria, and  fungi, requires the glyoxysomal enzymes isocitrate lyase and malate  synthase. This variation of the citric acid cycle permits net syn¬  thesis of glucose from acetyl-CoA.    1. The C0 2 produced in one round of the citric acid cycle does not  originate in the acetyl carbons that entered that round. If acetyl-  CoA is labeled with l4 C at its carbonyl carbon, how many rounds  of the cycle are required before l4 C0 2 is released?   2. If acetyl-CoA is labeled with l4 C at its methyl group, how many  rounds of the cycle are required before l4 C0 2 is released?   3. List four possible metabolic fates for pyruvate in a mammalian  cell. Indicate the type of reaction that occurs. What additional  reaction occurs in yeast?   4. Explain why metabolic acidosis (Box 2-2) may result from the  accumulation of some citric acid cycle intermediates.   3. Which one of the five steps of the pyruvate dehydrogenase com¬  plex reaction is most likely to be metabolically irreversible?  Explain.   6. Explain why an individual with a deficiency of pyruvate dehy¬  drogenase phosphatase (PDP) is unable to tolerate exercise.   7. Some human brain cancer cells contain a mutated form of iso¬  citrate dehydrogenase that has lost its normal activity and instead  catalyzes the NADP + -dependent reduction of the carbonyl group  of a-ketoglutarate. Draw the structure of the reaction product.   8. The compound you identified in Problem 7 resembles glutamate  and might therefore competitively inhibit glutamate transamina¬  tion. How would this affect citric acid cycle activity?   9. The branched-chain a-keto acid dehydrogenase complex, which  participates in amino acid catabolism, contains the same three  types of enzymes as are in the pyruvate dehydrogenase and the  a-ketoglutarate dehydrogenase complexes. Draw the reaction  product when valine is deaminated as in the glutamate  a-ketoglutarate reaction (Section 17-3A) and then is acted on by  the branched-chain a-keto acid dehydrogenase.   10. Which compounds would accumulate in an individual with  beriberi (caused by thiamine deficiency)?   11. Refer to Table 14-4 to explain why FAD rather than NAD + is  used in the succinate dehydrogenase reaction.   12. Certain microorganisms with an incomplete citric acid cycle decar-  boxylate a-ketoglutarate to produce succinate semialdehyde. A  dehydrogenase then converts succinate semialdehyde to succinate.   These reactions can be combined with other standard citric acid  cycle reactions to create a pathway from citrate to oxaloacetate.  Compare the ATP and reduced cofactor yield of the standard and  alternate pathways.   13. Malonate is a competitive inhibitor of succinate in the succinate  dehydrogenase reaction. Explain why increasing the oxaloacetate  concentration can overcome malonate inhibition.    14. What citric acid cycle intermediates would accumulate in the  presence of malonate?   15. What is the A G°' value for the portion of the citric acid cycle  that converts malate and acetyl-CoA to citrate?   16. Given the following information, calculate the physiological A G  of the isocitrate dehydrogenase reaction at 25°C and pH 7.0:  [NAD + ]/[NADH] = 8, [a-ketoglutarate] =0.1 mM, and  [isocitrate] = 0.02 mM. Assume standard conditions for C0 2  (A G°' is given in Table 17-2). Is this reaction a likely site for  metabolic control?   17. Calculate the ratio of [fumarate] to [succinate] under cellular  conditions at 37°C.   18. What is the ratio of [isocitrate] to [citrate] under cellular condi¬  tions at 37°C?   19. Why is it advantageous for citrate, the product of Reaction 1 of  the citric acid cycle, to inhibit phosphofructokinase, which  catalyzes the third reaction of glycolysis?   20. When blood [glucose] is high and a cell’s energy needs are met,  insulin stimulates glycogen synthesis. Is it counterproductive for  insulin to also promote the conversion of pyruvate to acetyl-CoA?  Explain.   21. (a) Explain why obligate anaerobes contain some citric acid cy¬  cle enzymes, (b) Why don’t these organisms have a complete cit¬  ric acid cycle?   22. The first organisms on earth may have been chemoautotrophs in  which the citric acid cycle operated in reverse to “fix” atmospheric  C0 2 in organic compounds. Complete a catalytic cycle that be¬  gins with the hypothetical overall reaction succinate + 2 C0 2 —>  citrate.   23. The catabolism of several amino acids generates succinyl-CoA.  Describe the series of reactions that are required to convert mi¬  tochondrial succinyl-CoA to cytosolic oxaloacetate that can be  used for gluconeogenesis.   24. Many amino acids are broken down to intermediates of the cit¬  ric acid cycle, (a) Why can’t these amino acid “remnants” be di¬  rectly oxidized to C0 2 by the citric acid cycle? (b) Explain why  amino acids that are broken down to pyruvate can be completely  oxidized by the citric acid cycle.   25. Anaplerotic reactions permit the citric acid cycle to supply inter¬  mediates to biosynthetic pathways while maintaining the proper  levels of cycle intermediates. Write the equation for the net  synthesis of citrate from pyruvate.   26. Although animals cannot synthesize glucose from acetyl-CoA, if  a rat is fed l4 C-labeled acetate, some of the label appears in glyco¬  gen extracted from its muscles. Explain.   27. Helicobacter pylori , which causes gastric ulcers, does not operate  a citric acid cycle but contains many of the citric acid cycle en¬  zymes. H. pylori can convert oxaloacetate to succinate for biosyn¬  thetic processes. Write an equation for this conversion.   28. The malaria parasite Plasmodium falciparum does not carry out  oxidative phosphorylation and therefore does not use the citric  acid cycle to generate reduced cofactors. Instead, the parasite con¬  verts amino acid-derived a-ketoglutarate to succinate. Write an  equation for the a-ketoglutarate —> succinate conversion that fol¬  lows (a) the oxidative (clockwise) path of the citric acid cycle or  (b) the reductive (counterclockwise) path of the cycle.   Focus concept: Pyruvate carboxylase is discovered in a bacterium  that was previously thought not to contain the enzyme.    Plasmodium falciparum, the  parasite that causes malaria, spends part of its life cycle inside  erythrocytes, an environment that is rich in nutrients and oxygen.  However, the parasite consumes very little oxygen. How does it  generate most of its ATP? The parasite contains a pyruvate dehy¬  drogenase, but this enzyme does not generate acetyl-CoA for the  citric acid cycle. Where is the dehydrogenase located, and what is  the resulting acetyl-CoA used for?    Eastmond, P.J. and Graham, I.A., Re-examining the role of the glyoxy¬  late cycle in oilseeds, Trends Plant Sci. 6, 72-77 (2001).   Huynen M.A., Dandekar, T., and Bork, P., Variation and evolution of the  citric-acid cycle: a genomic perspective, Trends Microbiol. 7,  281-291 (1999). [Discusses how genome studies can allow recon¬  struction of metabolic pathways, even when some enzymes appear  to be missing.]   Lengyel, J.S., Stott, K.M., Wu, X., Brooks, B.R., Balbo, A., Schuck, P.,  Perham, R.N., Subramaniam, S., and Milne, J.L.S., Extended  polypeptide linkers establish the spatial architecture of a pyruvate de¬  hydrogenase multienzyme complex, Structure 16, 93-103 (2008).   Milne, J.L.S., Wu, X., Borgnia, M.J., Lengyel, J.S., Brooks, B.R., Shi, D.,  Perham, R.N., and Subramaniam, S., Molecular structure of a    9-MDa icosahedral pyruvate dehydrogenase subcomplex containing  the E 2 and E 3 enzymes using cryoelectron microscopy, J. Biol. Chem.  281, 4364-4370 (2006).   Owen, O.E., Kalhan, S.C., and Hanson, R.W., The key role of anaplero-  sis and cataplerosis for citric acid cycle function, J. Biol. Chem. 277,  30409-30412 (2002). [Describes the influx (anaplerosis) and efflux  (cataplerosis) of citric acid cycle intermediates in different organ sys¬  tems.]   Perham, R.N., Swinging arms and swinging domains in multifunctional  enzymes: catalytic machines for multistep reactions, Annu. Rev.  Biochem. 69, 961-1004 (2000). [An authoritative review on mul¬  tienzyme complexes.]    Energy can take many forms, including light and sound. The mitochondria in eukaryotic  cells facilitate energy transformations involving chemical energy, mechanical energy,  and the energy of a proton gradient during the process of ATP synthesis. [Jeremiah  Barber/iStockphoto.]    Aerobic organisms consume oxygen and generate carbon dioxide in the process  of oxidizing metabolic fuels. The complete oxidation of glucose (C^H^O^),  for example, by molecular oxygen   C 6 H 12 O 6 + 6 O 2 —> 6 CO 2 + 6 H 2 0   can be broken down into two half-reactions that the metabolic machinery  carries out. In the first, glucose carbon atoms are oxidized:   C 6 H 12 0 6 + 6 H 2 0 -> 6 C0 2 + 24 H + + 24 e~   A Electron Transport Is an Exergonic Process  B Electron Carriers Operate in Sequence  C Complex I Accepts Electrons from NADH  D Complex II Contributes Electrons to  Coenzyme Q   A The Chemiosmotic Theory Links Electron  Transport to ATP Synthesis  B ATP Synthase Is Driven by the Flow of Protons  C The P/O Ratio Relates the Amount of ATP  Synthesized to the Amount of Oxygen Reduced  D Oxidative Phosphorylation Can Be Uncoupled  from Electron Transport   A The Rate of Oxidative Phosphorylation Depends  on the ATP and NADH Concentrations  B Aerobic Metabolism Has Some Disadvantages    We have already seen that the first half-reaction is mediated by the enzymatic  reactions of glycolysis and the citric acid cycle (the breakdown of fatty acids—  the other major type of metabolic fuel—also requires the citric acid cycle). In  this chapter, we describe the pathway by which the electrons from reduced fuel  molecules are transferred to molecular oxygen in eukaryotes. We also examine  how the energy of fuel oxidation is conserved and used to synthesize ATP.   As we have seen, the 12 electron pairs released during glucose oxidation are  not transferred directly to 0 2 . Rather, they are transferred to the coenzymes    FIG. 18-1 The sites of electron transfer  that form NADH and FADH 2 in glycolysis and  the citric acid cycle.    Point out the electron carriers that  account for all 24 of the electrons  derived from the oxidation of glucose   NAD + and FAD to form 10 NADH and 2 FADH 2 (Fig. 18-1) in the reactions  catalyzed by the glycolytic enzyme glyceraldehyde-3-phosphate dehydrogenase  (Section 15-2F), pyruvate dehydrogenase (Section 17-2B), and the citric acid cy¬  cle enzymes isocitrate dehydrogenase, a-ketoglutarate dehydrogenase, succinate  dehydrogenase, and malate dehydrogenase (Section 17-3). The electrons then pass  into the mitochondrial electron-transport chain , a system of linked electron car¬  riers. The following events occur during the electron-transport process:   1. By transferring their electrons to other substances, the NADH and  FADH 2 are reoxidized to NAD + and FAD so that they can partici¬  pate in additional substrate oxidation reactions.   2. The transferred electrons participate in the sequential oxidation-  reduction of multiple redox centers (groups that undergo oxidation-  reduction reactions) in four enzyme complexes before reducing 0 2 to H 2 0.   3. During electron transfer, protons are expelled from the mitochondrion,  producing a proton gradient across the mitochondrial membrane. The  free energy stored in this electrochemical gradient drives the synthesis of  ATP from ADP and P t through oxidative phosphorylation .   • A highly folded, protein-rich inner membrane separates the mitochondrial matrix  from the outer membrane.   • Transport proteins are required to import reducing equivalents, ADP, and P, into  the mitochondria.   The mitochondrion (Greek: mitos, thread + chondros , granule) is the site of  eukaryotic oxidative metabolism. Mitochondria contain pyruvate dehydroge¬  nase, the citric acid cycle enzymes, the enzymes catalyzing fatty acid oxidation  (Section 20-2), and the enzymes and redox proteins involved in electron trans¬  port and oxidative phosphorylation. It is therefore with good reason that the  mitochondrion is often described as the cells “power plant.”   Mitochondria vary in size and shape, depending on their source and meta¬  bolic state, but they are often ellipsoidal with dimensions of around 0.5 X  1.0 [Jim, about the size of a bacterium. A eukaryotic cell typically contains  800 to 2000 mitochondria, which occupy roughly one-fifth of its total cell  volume. A mitochondrion is bounded by a smooth outer membrane and con¬  tains an extensively invaginated inner membrane (Fig. 18-2). The number of  invaginations, called cristae (Latin: crests), reflects the respiratory activity of  the cell. The proteins mediating electron transport and oxidative phosphory¬  lation are bound in the inner mitochondrial membrane, so the respiration rate  varies with membrane surface area.   The inner membrane divides the mitochondrion into two compartments, the  intermembrane space and the internal matrix. The matrix is a gel-like solution  that contains extremely high concentrations of the soluble enzymes of oxidative  metabolism as well as substrates, nucleotide cofactors, and inorganic ions. The  matrix also contains the mitochondrial genetic machinery—DNA, RNA, and ri¬  bosomes—that generates several (but by no means all) mitochondrial proteins.   Two-dimensional electron micrographs of mitochondria such as Fig. 18 - 2/2  suggest that mitochondria are discrete kidney-shaped organelles. In fact, some  mitochondria adopt a tubular shape that extends throughout the cytosol.  Furthermore, mitochondria are highly variable structures. For example, the  cristae may not resemble baffles and the intercristal spaces may not commu¬  nicate freely with the mitochondrion’s intermembrane space. Electron   FIG. 1 8-2 The mitochondrion, (a) An electron micrograph of an animal  mitochondrion. [© K.R. Porter/Photo Researchers.] (b) Cutaway diagram of a mitochondrion.   FIG. 1 8-3 Electron microscopy-based  three-dimensional image reconstruction of a  rat liver mitochondrion. The outer membrane  (OM) is red, the inner membrane (IM) is yellow,  and the cristae (C) are green. The arrowheads  point to tubular regions of the cristae that  connect them to the inner membrane and to  each other. [Courtesy of Carmen Mannella,  Wadsworth Center, Albany, New York.]    microscopy-based three-dimensional image reconstruction methods have  revealed that cristae can range in shape from simple tubular entities to more  complicated lamellar assemblies that merge with the inner membrane via nar¬  row tubular structures (Fig. 18-3). Evidently cristae form microcompartments  that restrict the diffusion of substrates and ions between the intercristal and  intermembrane spaces. This has important functional implications because it  would result in a locally greater pH gradient across cristal membranes than across  inner membranes that are not part of cristae, thereby significantly influencing  the rate of oxidative phosphorylation (Section 18-3).   Like bacterial outer membranes, the outer mitochondrial membrane contains  porins, proteins that permit the free diffusion of molecules of up to 10 kD  (Section 10-2B). The intermembrane space is therefore equivalent to the cytosol in  its concentrations of metabolites and ions. The inner membrane, which is ~75%  protein by mass, is considerably richer in proteins than is the outer membrane  (Fig. 18-4). It is freely permeable only to 0 2 , C0 2 , and H 2 0 and contains, in  addition to respiratory chain proteins, numerous transport proteins that control  the passage of metabolites such as ATP, ADP, pyruvate, Ca 2+ , and phosphate.  The controlled impermeability of the inner mitochondrial membrane to most ions and  metabolites permits the generation of ion gradients across this barrier and results in  the compartmentalization of metabolic functions between cytosol and mitochondria.   The NADH produced in the cytosol by glycolysis must gain access to the mi¬  tochondrial electron-transport chain for aerobic oxidation. However, the inner  mitochondrial membrane lacks an NADH transport protein. Only the electrons  from cytosolic NADH are transported into the mitochondrion by one of several in¬  genious “shuttle” systems. We have already discussed the malate—aspartate shuttle  (Fig. 16-20), in which, when run in reverse, cytosolic oxaloacetate is reduced  to malate for transport into the mitochondrion. When malate is reoxidized in  the matrix, it gives up the reducing equivalents that originated in the cytosol.   In the glycerophosphate shuttle (Fig. 18-5) of insect flight muscle (the tis¬  sue with the largest known sustained power output—about the same power-to-  weight ratio as a small automobile engine), 3-phosphoglycerol dehydrogenase  catalyzes the oxidation of cytosolic NADH by dihydroxyacetone phosphate to  yield NAD + , which re-enters glycolysis. The electrons of the resulting 3-phos¬  phoglycerol are transferred to flavoprotein dehydrogenase to form  FADH 2 . This enzyme, which is situated on the inner mitochondrial  membranes outer surface, supplies electrons  transport chain (Section 18-2D).   A Translocator Exchanges ADP and ATP. Most of the ATP gener¬  ated in the mitochondrial matrix through oxidative phosphorylation  is used in the cytosol. The inner mitochondrial membrane contains  an ADP-ATP translocator (also called the adenine nucleotide  translocase) that transports ATP out of the matrix in exchange for  ADP produced in the cytosol by ATP-consuming reactions.   Several natural products inhibit the ATP-ADP translocator.  Atractyloside (a poison produced by the Mediterranean thistle   FIG. 1 8-4 Electron micrographs of the inner and outer mitochondrial  membranes that have been split to expose the inner surfaces of their  bilayer leaflets. Note that the inner membrane contains about twice the density  of embedded particles as does the outer membrane. The particles are the  portions of integral membrane proteins that were exposed when the bilayers  were split. [Courtesy of Lester Packer, University of California at Berkeley.]   The electrons of cytosolic NADH are transported  to the mitochondrial electron-transport chain in  three steps (shown in red as hydride transfers):  (1) Cytosolic oxidation of NADH by  dihydroxyacetone phosphate catalyzed by  3-phosphoglycerol dehydrogenase. (2) Oxidation  of 3-phosphoglycerol by flavoprotein  dehydrogenase with reduction of FAD to FADH 2 .  (3) Reoxidation of FADH 2 with passage of  electrons into the electron-transport chain.    Atractylis gummifera that was known to the ancient Egyptians) and its derivative  carboxyatractyloside (CATR) inhibit the process only from the external surface  of the inner mitochondrial membrane; bongkrekic acid (a product of the bac¬  terium Pseudomonas cocovenenans) exerts its effects only on the internal surface.    These differentially acting inhibitors have been valuable tools in the isolation of  the ATP-ADP translocator and in the elucidation of its mechanism of action.  For example, the translocator has been purified by affinity chromatography  (Section 5-2C) using atractyloside derivatives as affinity ligands. Atractyloside  binding is also a convenient means of identifying the translocator.   The ATP-ADP translocator, a dimer of identical ~300-residue subunits,  has characteristics similar to those of other transport proteins. Each subunit  has one binding site for which ADP and ATP compete. It has two major con¬  formations, one with its ATP-ADP binding site facing the matrix, and the    FIG. 1 8-6 X-Ray structure of bovine heart ATP-ADP  translocator in complex with carboxyatractyloside. (a) A subunit  of the homodimeric protein, viewed from the plane of the inner  membrane with the intermembrane space above, is drawn in ribbon  form colored in rainbow order from its N-terminus (blue) to its  C-terminus (red) and embedded in its semitransparent surface  diagram. The carboxyatractyloside is drawn in space-filling form with C  cyan, 0 red, and S yellow. Three cardiolipin molecules (Table 9-2) that   cocrystallized with the protein are drawn in stick form with C green, 0  red, and P orange. ( b) The molecular surface as viewed from the  intermembrane space and colored according its surface charge with  blue positive, white neutral, and red negative. Note the deep, positively  charged cavity in which the anionic ATP binds. [Based on an X-ray  structure by Eva Pebay-Peyroula, Universite Joseph Fourier, Grenoble,  France. PDBid 2C3E.]   other with this site facing the intermembrane space. In its X-ray structure in  complex with CATR (Fig. 18-6), determined by Eva Pebay-Peyroula, each  subunit’s six transmembrane helices surround a deep, cone-shaped cavity open  to the matrix that is occupied by the CATR.   The translocator must bind ligand to change from one conformation to  the other at a physiologically reasonable rate. Thus it functions as an exchanger  by importing one ADP for every ATP that is exported. In this respect, it dif¬  fers from the glucose transporter (Fig. 10-13), which can change its conforma¬  tion in the absence of ligand. Note that the export of ATP (net charge ~4)  and the import of ADP (net charge —3) results in the export of one negative  charge per transport cycle. This electrogenic antiport is driven by the mem¬  brane potential difference, A'T', across the inner mitochondrial membrane (pos¬  itive outside), which is a consequence of the transmembrane proton gradient.   • Draw a simple diagram of a mitochondrion  and identify its structural features.   • Describe how shuttle systems transport  reducing equivalents into the mitochondria.   • Explain how the free energy of the proton  gradient drives the transport of ATP, ADP,  and P/.    Phosphate Must Be Imported into the Mitochondrion. ATP is synthesized  from ADP + P z in the mitochondrion but is utilized in the cytosol. The P z  is returned to the mitochondrion by the phosphate carrier, an electroneutral  P z —H + symport that is driven by ApH. The transmembrane proton gradient  generated by the electron-transport machinery of the inner mitochondrial  membrane thus not only provides the thermodynamic driving force for ATP  synthesis (Section 18-3), it also motivates the transport of the raw materials—  ADP and P z —required for the process.   • The free energy of electron transport from NADH to 0 2 can drive the synthesis of  approximately 2.5 ATP.   • Electron carriers are arranged in the mitochondrial membrane so that electrons  travel from Complexes I and II via coenzyme Q to Complex III, and from there via  cytochrome c to Complex IV.   • The L-shaped Complex I transfers electrons from NADH to CoQ via a series of  iron-sulfur clusters and translocates four protons to the intermembrane space.   • Complex II transfers electrons from succinate to the CoQ pool but does not  contribute to the transmembrane proton gradient.   • Electrons from Complex III are transferred to cytochrome c and two protons are  translocated during the operation of the Q cycle in Complex III.   • Complex IV accepts electrons from cytochrome c to reduce 0 2 to H 2 0 and  translocates four protons for every two electrons transferred.   The electron carriers that ferry electrons from NADH and FADH 2 to 0 2 are as¬  sociated with the inner mitochondrial membrane. Some of these redox centers are  highly mobile, and others are less mobile components of integral membrane pro¬  tein complexes. The sequence of electron carriers roughly reflects their relative  reduction potentials, so that the overall process of electron transport is exergonic.  We begin this section by examining the thermodynamics of electron transport. We  then consider the molecular characteristics of the various electron carriers.   We can estimate the thermodynamic efficiency of electron transport by in¬  specting the standard reduction potentials of the redox centers. As we saw  in our thermodynamic considerations of oxidation—reduction reactions  (Section 14-3), an oxidized substrates affinity for electrons increases with  its standard reduction potential, %°' (Table 14-4 lists the standard reduc¬  tion potentials of some biologically important half-reactions). The standard  reduction potential difference, A%°\ for a redox reaction involving any two  half-reactions is expressed   For the reaction that occurs in mitochondria, that is, the oxidation of  NADH by 0 2 , the relevant half-reactions are   Since the 0 2 /H 2 0 half-reaction has the greater standard reduction potential  (0 2 has a higher affinity for electrons than NAD + ), we write the NADH half¬  reaction in reverse so that NADH is the electron donor in the couple and 0 2  the electron acceptor. The overall reaction is   \ 0 2 + NADH + H + H 2 0 + NAD +   A% of = 0.815 V - (-0.315 V) = 1.130 V   The standard free energy change for the reaction can then be calculated from   For NADH oxidation, A G or — —218 kj • mol 1 . In other words, the oxida¬  tion of 1 mol of NADH by 0 2 (the transfer of 2 mol e ) under standard bio¬  chemical conditions is associated with the release of 218 kj of free energy.    Because the standard free energy required to synthesize 1 mol of ATP from  ADP + P z is 30.5 kj • mol \ the oxidation of NADH by 0 2 is theoreti¬  cally able to drive the formation of several moles of ATP In mitochondria,  the coupling of NADH oxidation to ATP synthesis is achieved by an elec¬  tron-transport chain in which electrons pass through three protein com¬  plexes. This allows the overall free energy change to be broken into three smaller  parcels, each of which contributes to ATP synthesis by oxidative phosphoryla¬  tion. Oxidation of one NADH results in the synthesis of approximately 2.5  ATP (we will see later why the relationship is not strictly stoichiometric).  The thermodynamic efficiency of oxidative phosphorylation is therefore  2.5 X 30.5 kj • mol 1 X 100/218 kj • mol 1 = 35% under standard  biochemical conditions. However, under physiological conditions in active  mitochondria (where the reactant and product concentrations as well as  the pH deviate from standard conditions), the thermodynamic efficiency  is thought to be ~70%. In comparison, the energy efficiency of a typical  automobile engine is <30%.   Oxidation of NADH and FADH 2 is carried out by the electron-transport chain, a  set of protein complexes containing redox centers with progressively greater affinities  for electrons (increasing standard reduction potentials). Electrons travel through the  chain from lower to higher standard reduction potentials (Fig. 18-7). Electrons  are carried from Complexes I and II to Complex III by coenzyme Q (CoQ  or ubiquinone; so named because of its ubiquity in respiring organisms), and  from Complex III to Complex IV by the peripheral membrane protein  cytochrome c .    FIG. 1 8-7 Overview of electron transport  in the mitochondrion. The standard reduction  potentials of its most mobile components (green)  are indicated, as are the points where sufficient  free energy is released to synthesize ATP (blue)  and the sites of action of several respiratory  inhibitors (red). Complexes I, III, and IV do not  directly synthesize ATP but sequester the free  energy necessary to do so by pumping protons  outside the mitochondrion to form a proton   for the passage of electrons through  Complexes I, III, and IV?    Complex I  AT' = 0.360 V  (AG°' = -69.5 kJ • mol" 1 )    Complex IV  A%°' = 0.580 V  (A G°' = -112 kJ • mol" 1 )   Complex III catalyzes oxidation of Co Q (reduced) by cytochrome c:   Complex IV catalyzes oxidation of reduced cytochrome c by 0 2 , the terminal elec¬  tron acceptor of the electron-transport process:   2 Cytochrome c [reduced) + \ 0 2 —>2 cytochrome c [oxidized) + H 2 0   As an electron pair successively traverses Complexes I, III, and IV, sufficient  free energy is released at each step to power ATP synthesis.   Complex II catalyzes the oxidation of FAD H 2 by CoQ:   This redox reaction does not release sufficient free energy to synthesize ATP; it  functions only to inject the electrons from FADH 2 into the electron-transport chain.    Inhibitors Reveal the Workings of the Electron-Transport Chain. The se¬  quence of events in electron transport was elucidated largely through the use  of specific inhibitors and later corroborated by measurements of the standard  reduction potentials of the redox components. The rate at which 0 2 is con¬  sumed by a suspension of mitochondria is a sensitive measure of the activity  of the electron-transport chain. Compounds that inhibit electron transport, as  judged by their effect on 0 2 consumption, include rotenone (a plant toxin  used by Amazonian Indians to poison fish and which is also used as an insec¬  ticide), amytal (a barbiturate), antimycin A (an antibiotic), and cyanide.   Adding rotenone or amytal to a suspension of mitochondria blocks electron  transport in Complex I; antimycin A blocks Complex III, and CN blocks elec¬  tron transport in Complex IV (Fig. 18-7). Each of these inhibitors also halts 0 2  consumption. Oxygen consumption resumes following addition of a substance  whose electrons enter the electron-transport chain “downstream” of the block. For  example, the addition of succinate to rotenone-blocked mitochondria restores  electron transport and 0 2 consumption. Experiments with inhibitors of electron  transport thus reveal the points of entry of electrons from various substrates.   Each of the four respiratory complexes of the electron-transport chain con¬  sists of several protein components that are associated with a variety of    TABLE 18-1 Reduction Potentials of Electron-Transport Chain  Components in Resting Mitochondria   Complex IV (cytochrome c oxidase; —410 kD dimer, 8-13 unique subunits):    Source: Mainly Wilson, D.F., Erecinska, M., and Dutton, P.L., Annu. Rev. Biophys. Bioeng. 3, 205  and 208 (1974); and Wilson, D.E, in Bittar, E.E. (Ed.), Membrane Structure and Function, Vol. 1,  p. 160, Wiley (1980).    redox-active prosthetic groups with successively increasing reduction potentials  (Table 18-1). The complexes are all laterally mobile within the inner mito¬  chondrial membrane and may associate to form “supercomplexes” with variable  composition, such as III 2 IV 2 and I 1 III 2 IV 1 . In the following sections, we ex¬  amine the structures of Complexes I through IV and the molecules that trans¬  fer electrons between them. Their relationships are summarized in Fig. 18-8.   Complex I (NADH-coenzyme Q oxidoreductase), which passes electrons  from NADH to CoQ, may be the largest protein complex in the inner mito¬  chondrial membrane, containing, in mammals, 45 subunits with a total mass  of ~900 kD. In both prokaryotes and eukaryotes, the complex is L-shaped,    FIG. 1 8-8 The mitochondrial electron-transport chain. This  diagram indicates the pathways of electron transfer (blue) and proton  translocation (red). Electrons are transferred between Complexes I  and III by the membrane-soluble coenzyme Q (Q) and between    Complexes III and IV by the peripheral membrane protein cytochrome  c. Complex II transfers electrons from succinate to coenzyme Q.    with one arm embedded in the inner mitochondrial membrane (or plasma  membrane in prokaryotes) and the other extending into the matrix (or extra¬  cellular space in prokaryotes; Fig. 18-9). Proton translocation occurs in the  membrane-embedded arm of the complex, whereas redox reactions take place  in the peripheral arm. In mammals, 7 of its 45 subunits, its most hydropho¬  bic subunits, which form the core of its transmembrane region, are encoded  by mitochondrial genes. The remainder are encoded by nuclear genes and  hence must be imported into the mitochondrion.   Complex I Contains Multiple Coenzymes. Complex I contains one molecule  of flavin mononucleotide (FMN, a redox-active prosthetic group that differs  from FAD only by the absence of the AMP group) and eight or nine iron—sulfur  clusters. Iron-sulfur clusters occur as the prosthetic groups of iron-sulfur  proteins (also called nonheme iron proteins). The two most common types,  designated [2Fe-2S] and [4Fe-4S] clusters (at right, ), consist of equal numbers  of iron and sulfide ions and are both coordinated to four protein Cys sulfhydryl  groups. Note that the Fe atoms in both types of clusters are each coordinated  by four S atoms, which are more or less tetrahedrally disposed around the Fe.   Iron-sulfur clusters can undergo one-electron oxidation and reduction.  The oxidized and reduced states of all iron-sulfur clusters differ by one formal    FIG. 1 8-9 Complex I from the bacterium Thermus  thermophilus. The protein is viewed parallel to the plane of the  plasma membrane with the cytoplasm (equivalent to the  mitochondrial matrix) above and the periplasm (outside) below. The  15 subunits are drawn in ribbon form in different colors with the  110-A-long horizontal helix of subunit Nqo12 purple. The loops  connecting the transmembrane helices are not visible in this  low-resolution (4.5 A) X-ray structure. The nine Fe-S clusters in  the peripheral (cytoplasmic) arm are drawn in space-filling form  with S yellow and Fe red-brown. [Courtesy of Leonid Sazanov,  Medical Research Council, Cambridge, U.K. PDBid 3M9S.]    FIG. 18-10 The oxidation states of FMN  and coenzyme Q. Both (a) FMN and    charge regardless of their number of Fe atoms. This is because the Fe atoms in  each cluster form a conjugated system and thus can have oxidation states be¬  tween the normal +2 and +3 values for individual Fe ions.   FMN and CoQ can each adopt three oxidation states (Fig. 18-10). They  are capable of accepting and donating either one or two electrons because their  semiquinone forms are stable (these semiquinones are stable free radicals, mol¬  ecules with an unpaired electron). FMN is tightly bound to proteins; however,  CoQ has a hydrophobic tail that makes it soluble in the inner mitochondrial  membranes lipid bilayer. In mammals, this tail consists of 10 C 5 isoprenoid  units (Section 9-IF) and hence the coenzyme is designated Qio* I n other or¬  ganisms, CoQ may have only 6 (Q^) or 8 (Q 8 ) isoprenoid units.   Electrons Follow a Multistep Path through Complex I. The X-ray structure  of the peripheral arm of the Thermus thermophilus Complex I, determined by  Leonid Sazanov, reveals a Y-shaped assembly that is 140 A high (Fig. 18-11 a). This  subcomplex contains all of the enzymes redox centers: an FMN, seven [4Fe—4S]  clusters, and two [2Fe-2S] clusters. The FMN is located at the end of a sol-  vent-exposed cavity that presumably forms the NADH-binding site. The CoQ-  binding site is located near the end of a chain of [4Fe-4S] clusters. X-Ray  analysis of a yeast mitochondrial Complex I suggests that the CoQ must diffuse  out of the lipid bilayer and move some 25 A through a hydrophobic cavity in  the protein in order for its quinone head to be positioned to accept electrons.   FIG. 1 8-1 1 X-Ray structure of the peripheral arm of  Complex I from Thermus thermophilus. (a) The protein is viewed  parallel to the plane of the membrane but turned —180° about the  vertical axis relative to the structure shown in Fig. 18-9. Its eight  subunits are drawn in semitransparent ribbon form in different colors  (which differ from those in Fig. 18-9). The FMN (upper left) and the  nine iron-sulfur clusters are shown in space-filling form with C green,  N blue, 0 red, P orange, S yellow, and Fe red-brown. The likely  CoQ-binding site (Q) is indicated by a gray arrow, (b) The   arrangement of redox groups viewed similarly to Part a. The FMN  (stick model with C yellow) and the two [Fe-S] and seven [4Fe-4S]  clusters (space-filling models) are shown together with their  center-to-center distances in angstroms (shortest edge-to-edge  distances are indicated in parentheses). Blue arrows represent the  ~94-A-long main path of electrons after their transfer from NADH to  FMN. [Part a based on an X-ray structure by and Part b courtesy of  Leonid Sazanov, Medical Research Council, Cambridge, U.K. PDBid  2FUG.]    The transit of electrons from NADH to CoQ presumably occurs by a step¬  wise mechanism, according to the reduction potential of the various redox cen¬  ters in Complex I (Table 18-1). This process involves the transient reduction of  each group as it binds electrons and its reoxidation when it passes the electrons  to the next group. The spatial arrangement of the groups indicates the likely  path of the electrons (Fig. 18-11 b). Note that redox centers do not need to come  into contact in order to transfer an electron. An electrons quantum mechani¬  cal properties enable it to quickly “tunnel” (jump) between protein-embed¬  ded redox groups that are separated by less than ~ 14 A. Because electron transfer  rates decrease exponentially with the distance between redox centers (they ex¬  hibit an ~ 10-fold decrease for each 1.7 A increase in distance), electron trans¬  fers over distances longer than ~14 A always involve chains of redox centers.   NADH can participate in only a two-electron transfer reaction. In contrast,  the cytochromes of Complex III (see below), to which reduced CoQ passes its  electrons, are capable of only one-electron reactions. FMN and CoQ, which can  transfer one or two electrons at a time, therefore provide an electron conduit between  the two-electron donor NADH and the one-electron acceptors, the cytochromes.   Complex I Translocates Four Protons. As electrons are transferred between the  redox centers of Complex I, four protons are translocated from the matrix to the  intermembrane space. The fate of protons donated by NADH is uncertain:  They may be among those pumped across the membrane or they may be used  in the reduction of CoQ to its hydroquinone form, CoQH 2 . Proton pump¬  ing in Complex I is apparently driven by conformational changes induced by  changes in the redox state of the protein. These conformational changes alter  the p K values of ionizable side chains so that protons are taken up or released  as electrons are transferred. Coupling between electron transport in the  peripheral arm and proton pumping in the transmembrane arm requires   FIG. 1 8-1 2 Proton translocation in  bacteriorhodopsin. The retinal prosthetic group  in Schiff base linkage to Lys 216 of the  seven-transmembrane helix protein is shown in  purple. The side chains of amino acids that  participate in light-driven proton translocation are  shown in stick form with C gray, N blue, and 0  red. The arrows with their associated numbers  indicate the order of proton-transfer steps during  the photochemical cycle: (1) deprotonation of the  Schiff base and protonation of Asp 85;   (3) reprotonation of the Schiff base and  deprotonation of Asp 96; (4) reprotonation of  Asp 96 from the cytoplasmic surface; and   (5) deprotonation of Asp 85 and reprotonation of  the proton release site. [Courtesy of Janos Lanyi,  University of California at Irvine. PDBid 1C3W.]    communication over a distance of ~ 100 A. An "^llO-A-long a helix embed¬  ded in the membrane near the matrix-facing surface (purple in Fig. 18-9) is  postulated to function as a sort of piston to transmit redox-sensitive confor¬  mational changes in the peripheral arm to the proton-transport site.   Because a proton is simply an atomic nucleus, it cannot be transported  across a membrane in the same way as ions such as Na + and K + . However,  a proton can be translocated by “hopping” along a chain of hydrogen-bonded  groups in a transmembrane channel, just as it “jumps” between hydrogen-  bonded water molecules in solution (Fig. 2-15). Such an arrangement of  hydrogen-bonded groups in the protein has been described as a proton wire  and may include water molecules. Apparently, Complex I contains several  proton wires that collectively pump four protons across the membrane for  every pair of electrons that passes from NADH to CoQ.    Bacteriorhodopsin Is a Model Proton Pump. A useful model for proton-  translocating complexes is bacteriorhodopsin, an integral membrane protein  from Halobacterium salinarium that contains seven transmembrane helical seg¬  ments surrounding a central polar channel (Fig. 9-22). Bacteriorhodopsin is a  light-driven proton pump: It obtains the free energy required for pumping pro¬  tons through the absorbance of light by its retinal prosthetic group. The retinal  is linked to the protein via a protonated Schiff base to the side chain of Lys 216.   On absorbing light, the all-tom-retinal isomerizes to its 13 -cis configuration:    This structural change initiates a sequence of protein conformational adjustments  that restore the system to its ground state over a period of ~ 10 ms. These con¬  formational changes alter the p K’s of several amino acid side chains (Fig. 18-12).  Specifically, the p K of Asp 85 increases so that it can receive a proton from the  Schiff base. Asp 85 then transfers the proton to the extracellular medium via a  hydrogen-bonded network that includes Arg 82, Glu 194, Glu 204, and several  water molecules. Water molecules also move into position to form a hydrogen-  bonded network that reprotonates the Schiff base with an intracellular proton via  Asp 96, whose piLdecreases. The net result is that a proton appears to move from  the cytosol to the cell exterior (the proton that leaves the cytosol is not the same  proton that enters the extracellular space).   The various amino acid side chains involved in proton transport in bac¬  teriorhodopsin move by ~1 A or less, but this is enough to alter their piLval-  ues and to sequentially make and break hydrogen bonds so as to allow a proton  to pass along the proton wire. The vectorial (one-way) nature of the process  arises from the unidirectional series of conformational changes made by the  photoexcited retinal as it relaxes to its ground state. Electron transfers between    the various redox cofactors of Complex I likely motivate a similar sequence of  conformational and p K changes.   Complex II (succinate-coenzyme Q oxidoreductase), which contains the cit¬  ric acid cycle enzyme succinate dehydrogenase (Section 17-3F), passes electrons  from succinate to CoQ. Its redox groups include succinate dehydrogenase’s cova¬  lently bound FAD (Fig. 17-13) to which electrons are initially passed, one  [4Fe-4S] cluster, a [3Fe-4S] cluster (essentially a [4Fe-4S] complex that lacks  one Fe atom), one [2Fe—2S] cluster, and one cytochrome b (cytochromes are  discussed in Box 18-1). All of its four subunits are encoded by nuclear genes.   The free energy for electron transfer from succinate to CoQ (Fig. 18-7)  is insufficient to drive ATP synthesis. The complex is nevertheless important  because it allows relatively high-potential electrons to enter the electron-  transport chain by bypassing Complex I.   Note that Complexes I and II, despite their names, do not operate in se¬  ries. But both accomplish the same result: the transfer of electrons to CoQ  from reduced substrates (NADH or succinate). CoQ, which diffuses in the lipid  bilayer among the respiratory complexes, therefore serves as a sort of collection point  for electrons. As we will see in Section 20-2C, the first step in fatty acid oxi¬  dation also generates electrons that enter the electron-transport chain at the  level of CoQ. CoQ also collects electrons from the FADH 2 produced by the  glycerophosphate shuttle (Fig. 18-5).   Complex II Contains a Linear Chain of Redox Cofactors. The X-ray structure  of chicken mitochondrial Complex II has been determined (Fig. 18-13).  Complex II is a mushroom-shaped homotrimer whose protomers each consist of  two hydrophilic subunits, a flavoprotein (Fp) and an iron-sulfur subunit (Ip), that  project into the mitochondrial matrix, and two hydrophobic membrane-anchor  subunits, CybL and CybS, which each have three transmembrane helices and   semitransparent ribbon diagram viewed  parallel to the inner mitochondrial  membrane with the matrix above. The  enzyme’s four subunits are drawn in  different colors. The inhibitor  oxaloacetate (OAA, which marks the  succinate-binding site), FAD, three Fe-S  clusters, CoQ, and heme b are shown  in space-filling form with oxaloacetate  C magenta, FAD and heme b C green,  CoQ C cyan, N blue, 0 red, P orange,   S yellow, and Fe red-brown. The  inferred position of the inner  mitochondrial membrane is indicated.   (b) The ligands and redox cofactors  viewed and colored as in Part a but  drawn in stick form. Their closest  edge-to-edge distances are indicated.  [Based on an X-ray structure by  Edward Berry, Lawrence Berkeley  National Laboratory, Berkeley, California.  PDBid 1YQ3J    Cytochromes, whose function was elucidated in 1925 by David  Keilin, are redox-active proteins that occur in all organisms except  a few types of obligate anaerobes. These proteins contain heme  groups that alternate between their Fe(ll) and Fe(lll) oxidation  states during electron transport.   The heme groups of the reduced Fe(ll) cytochromes have  prominent visible absorption spectra consisting of three peaks: the  a , (3, and 7 (Soret) bands. The spectrum for cytochrome c is  shown in Fig. a. The wavelength of the a peak, which varies  characteristically with the reduced cytochrome species (it is  absent in oxidized cytochromes), is used to differentiate the vari¬   ous cytochromes in mitochondrial membranes (top right of Fig. a  and Fig. b).   Each group of cytochromes contains a differently substituted  heme group coordinated with the redox-active iron atom. The b-type  cytochromes contain protoporphyrin IX, which also occurs in myo¬  globin and hemoglobin (Section 7-1 A). The heme group of c-type  cytochromes differs from protoporphyrin IX in that its vinyl groups  have added Cys sulfhydryls across their double bonds to form  thioether linkages to the protein. Heme a contains a long hydropho¬  bic tail of isoprene units attached to the porphyrin, as well as a  formyl group in place of a methyl substituent in hemes b and c. The   which collectively bind one 6-type heme and one ubiquinone. Fp binds both the  substrate (whose binding site in the X-ray structure is occupied by the inhibitor ox-  aloacetate) and the FAD prosthetic group, whereas Ip binds the complex’s three  iron—sulfur clusters. The substrate- and ubiquinone-binding sites are connected  by a >40-A-long chain of redox centers with the sequence substrate—FAD—  [2Fe-2S]—[4Fe-4S]—[3Fe-4S]—CoQ (top to bottom in Fig. 18-13). The  heme b , which is not located in this direct electron-transfer pathway, apparently  fine-tunes the systems electronic properties so as to suppress side reactions that  form damaging reactive oxygen species such as H 2 0 2 (Section 18-4B). The  [2Fe-2S] cluster labeled Nla in Complex I (Fig. 18-11 b) may have a similar role.   Complex III (also known as coenzyme Q-cytochrome c oxidoreductase or  cytochrome be x ) passes electrons from reduced CoQ to cytochrome c. It con¬  tains two 6-type cytochromes, one cytochrome c ly and one [2Fe-2S] clus¬  ter in which one of the Fe atoms is coordinated by two His residues rather  than two Cys residues (and which is known as a Rieske center after its dis¬  coverer, John Rieske). Complex III from yeast mitochondria is a 419-kD ho¬  modimer whose protomers each consist of 9 subunits (11 in the 485-kD  bovine heart mitochondrial Complex III). Its X-ray structure (Fig. 18-14)    axial ligands of the heme iron also vary with the cytochrome type. In  cytochromes a and b, both ligands are His residues, whereas in  cytochromes c, one is His and the other is the S atom of Met.   Within each group of cytochromes, different heme group envi¬  ronments may be characterized by slightly different a peak wave¬  lengths. For this reason, it is convenient to identify cytochromes by  the wavelength (in nanometers) at which its a band absorbance is  maximal (e.g., cytochrome b 560 in Complex II). Cytochromes are  also identified nondescriptively with either numbers or letters.   Reduced heme groups are highly reactive entities; they can  transfer electrons over distances of 10 to 20 A at physiologically    significant rates. Hence cytochromes, in a sense, have the  opposite function of enzymes: Instead of persuading unreactive  substrates to react, they must prevent their hemes from trans¬  ferring electrons nonspecifically to other cellular components.  This, no doubt, is why the hemes are almost entirely enveloped  by protein. However, cytochromes must also provide a path for  electron transfer to an appropriate partner. Since electron  transfer occurs far more efficiently through bonds than through  space, protein structure appears to be an important determinant  of the rate of electron transfer between proteins.    FIG. 18-14 X-Ray structure of yeast Complex III in  complex with cytochrome c and the inhibitor stigmatellin.   The homodimeric complex is viewed from within the plane of the  membrane with the intermembrane space above. The nine different  subunits in each protomer, which collectively have 12  transmembrane helices, are differently colored with cytochrome b  pale yellow, cytochrome light purple, the ISP tan, and  cytochrome c pink. The portions of the protein that extend into the  matrix are colored cyan and orange. The four different heme  groups, the [2Fe-2S] cluster, and stigmatellin are drawn in  space-filling form with heme C green, stigmatellin C magenta,   N blue, 0 red, S yellow, and Fe red-brown. The horizontal lines  delineate the inferred position of the membrane. Note that only  one cytochrome c is bound to the homodimeric Complex III.   [Based on an X-ray structure by Carola Hunte, Max Planck  Institute for Biophysics, Frankfurt am Main, Germany. PDBid  1 KYO.] Q See Interactive Exercise 19.   reveals a pear-shaped homodimer whose widest part extends ~75 A into the  mitochondrial matrix. The ~40-A-thick transmembrane portion consists of  12 transmembrane helices per protomer (14 in bovine heart Complex III),  most of which are tilted with respect to the plane of the membrane. Eight of  the helices belong to the cytochrome b subunit, which binds both b-type cy¬  tochrome hemes, b 5 ^2 ( or for high potential, which lies near the matrix)   and b 5 66 (or b L , for low potential, which lies near the intermembrane space),  and is the only subunit encoded by a mitochondrial gene. The cytochrome c x  subunit is anchored by a single transmembrane helix, with its globular head,  which contains a c-type heme, extending into the intermembrane space. The  iron-sulfur protein (ISP), which contains the Rieske center, is similarly an¬  chored by a single transmembrane helix and extends into the intermembrane  space. The two ISPs of the dimeric complex are intertwined so that the  [2Fe-2S] cluster in the ISP of one protomer interacts with the cytochrome b   Electrons from Coenzyme Q Follow Two Paths. Complex III functions to  permit one molecule of CoQH 2 , a two-electron carrier, to reduce two mole¬  cules of cytochrome c, a one-electron carrier. This occurs by a surprising  bifurcation of the flow of electrons from CoQH 2 to cytochrome c x and to  cytochrome b (in which the flow is cyclic). It is this so-called Q cycle that  permits Complex III to pump protons from the matrix to the intermembrane  space.   The essence of the Q cycle is that CoQH 2 undergoes a two-cycle reoxidation  in which the semiquinone, CoQ*, is a stable intermediate. This involves two in¬  dependent binding sites for coenzyme Q: Q 0 , which binds CoQH 2 and is  located between the Rieske [2Fe-2S] center and heme in proximity to the   intermembrane space; and Qj, which binds both CoQ* and CoQ and is lo¬  cated near heme b H in proximity to the matrix. In the first cycle (Fig. 18-15,  top), CoQH 2 from Complex I (1 and 2) binds to the Q 0 site, where it trans¬  fers one of its electrons to the ISP (3), releasing its two protons into the inter¬  membrane space and yielding CoQ* The ISP goes on to reduce cytochrome c x ,  whereas the CoQ* transfers its remaining electron to cytochrome (4), yield¬  ing fully oxidized CoQ. Cytochrome then reduces cytochrome b H (6). The   CoQ from Step 4 is released from the Q 0 site and rebinds to the Qi site (5),  where it picks up the electron from cytochrome b H (7), reverting to the semi¬  quinone form, CoQ* . Thus, the reaction for the first cycle is   CoQ* + cytochrome c x (Fe 2+ ) + 2 H + ( intermembrane )   In the second cycle (Fig. 18-15, bottom ), another CoQH 2 from Complex I  repeats Steps 1 through 6: One electron reduces the ISP and then cytochrome c x ,  and the other electron sequentially reduces cytochrome b L and then cytochrome  b H . This second electron then reduces the CoQ* at the Qi site produced in the  first cycle (8), yielding CoQH 2 . The protons consumed in this last step originate  in the mitochondrial matrix. The reaction for the second cycle is therefore   CoQH 2 + CoQ 7 + cytochrome c x (Fe 3+ ) + 2 H + ( matrix ) —>   CoQ + CoQH 2 + cytochrome c x (Fe 2+ ) + 2 H + ( intermembrane )   For every two CoQH 2 that enter the Q cycle, one CoQH 2 is regenerated.  The combination of both cycles, in which two electrons are transferred from  CoQH 2 to cytochrome c x , results in the overall reaction   CoQH 2 + 2 cytochrome c x (Fe 3+ ) + 2 H + ( matrix ) —>   CoQ + 2 cytochrome c x (Fe 2+ ) + 4 H + ( intermembrane )    FIG. 1 8-1 5 The Q cycle. The Q cycle,  which is mediated by Complex III, results in the  translocation of H + from the matrix to the  intermembrane space as driven by the transport  of electrons from cytochrome b to cytochrome c.  The overall cycle is actually two cycles, the first  requiring Reactions 1 through 7 and the second  requiring Reactions 1 through 6 and 8.   (1) Coenzyme QH 2 is supplied by Complex I on  the matrix side of the membrane. (2) QH 2  diffuses to the cytosolic side of the membrane,  where it binds in the Q 0 site on the cytochrome  b subunit of Complex III. (3) QH 2 reduces the  Rieske iron-sulfur protein (ISP), forming Q 7  semiquinone and releasing 2 H + . The ISP goes  on to reduce heme c-|. (4) Q 7 reduces heme b L  to form coenzyme Q. (5) Q diffuses to the matrix  side, where, in Cycle 1 only, it binds in the Q,  site on cytochrome b. ( 6 ) Heme b L reduces  heme b H . (7, Cycle 1 only) Q is reduced to Q 7  by heme b H . ( 8 , Cycle 2 only) Q 7 bound in the  Q, site is reduced to QH 2 by heme d H . The net  reaction is the transfer of two electrons from  QH 2 to cytochrome c-| and the translocation of  four protons from the matrix to the  intermembrane space. [After Trumpower, B.L,    How does the structure of Complex III support the operation of the Q  cycle? First, X-ray studies provide direct evidence for the independent exis¬  tence of the Q 0 and Qj sites. The antifungal agent stigmatellin,   which is known to inhibit electron flow from CoQH 2 to the ISP and to heme  (Steps 3 and 4 of both cycles), binds in a pocket within cytochrome b mid¬  way between the iron positions of the Rieske [2Fe-2S] center and heme b L .  Thus, this binding pocket is likely to overlap the Q 0 site. Similarly, antimycin   FIG. 18-16 Ribbon diagram of  cytochrome c showing the Lys residues  involved in intermolecular complex formation.   Dark and light blue balls, respectively, mark the  position of Lys residues whose £-amino groups  are strongly and less strongly protected by  cytochrome or cytochrome c oxidase against  acetylation. Note that the Lys residues form a  ring around the heme (solid bar) on one face of  the protein. [After Mathews, F.S., Prog. Biophys.  Mol. Biol. 45, 45 (1986).] O See Interactive  Exercise 20 and Kinemage Exercise 5.    A (Section 18-2B), which has been shown to block electron flow from heme  b H to CoQ or CoQ* (Step 7 of Cycle 1 and Step 8 of Cycle 2), binds in a  pocket near heme b H , thereby identifying this pocket as site Q>   The circuitous route of electron transfer in Complex III is tied to the abil¬  ity of coenzyme Q to diffuse within the hydrophobic core of the membrane  in order to bind to both the Q 0 and Qj sites. In fact, the mitochondrial mem¬  brane likely contains a pool of CoQ, CoQ*, and CoQH 2 , so that the  ubiquinone molecule released from the Q 0 site may not be the same one that  rebinds to the Qj site in Cycle 1 (Fig. 18-15).   X-Ray structures of cytochrome bc x also explain why Q 0 -bound CoQ* ex¬  clusively reduces heme rather than the Rieske [2Fe-2S] cluster of the ISP,  despite the greater reduction potential difference (A'S) favoring the latter reac¬  tion (Table 18-1). The globular domain of the ISP can swing via an ~20-A  hinge motion between the Q 0 site and cytochrome c 1 . Consequently, the ISP ac¬  quires an electron from CoQH 2 in the (f site and mechanically delivers it to the  heme group. The CoQ* product cannot reduce the ISP (after it has reduced  cytochrome c x ) because the ISP has moved too far away for this to occur.   The net reaction for the Q cycle indicates that when CoQH 2 is oxidized,  two reduced cytochrome c molecules and four protons appear on the outer side of  the membrane. Proton transport by the Q cycle thus differs from the proton¬  pumping mechanism of Complexes I and IV (see below): In the Q cycle, a  redox center itself (CoQ) is the proton carrier.   Cytochrome c Is a Soluble Electron Carrier. The electrons that flow to cy¬  tochrome Ci are transferred to cytochrome c, which, unlike the other cy¬  tochromes of the respiratory electron-transport chain, is a peripheral  membrane protein. It shuttles electrons between Complexes III and IV on the  outer surface of the inner mitochondrial membrane. The evolution and struc¬  ture of cytochrome c are discussed in Sections 5-4A and 6-2D. Several highly  conserved Lys residues in cytochrome c lie in a ring around the exposed edge  of its otherwise buried heme group (Fig. 18-16). These positively charged  residues constitute binding sites for complementary negatively charged groups  on cytochrome c 1 and cytochrome c oxidase. Such interactions presumably  serve to align redox groups for optimal electron transfer.   The X-ray structure shown in Fig. 18-14 reveals that the association be¬  tween cytochrome bc x and cytochrome c is particularly tenuous, because its  interfacial area (880 A 2 ) is significantly less than that exhibited by protein-  protein complexes known to have low stability (typically < 1600 A 2 ). Such a  small interface is well suited for fast binding and release. This interface  involves only two cytochrome c Lys residues, Lys 86 and Lys 79 (vertebrate  numbering; Table 5-6), which respectively contact Glu 235 and Ala 164 of  cytochrome c 1 . Other pairs of charged and often conserved residues surround  the contact site but they are not close enough for direct polar interactions.  Perhaps these interactions are mediated by water molecules that are not seen  in the X-ray structure. The closest approach between the heme groups of the  contacting proteins is 4.5 A between atoms of their respective thioether-  bonded substituents, which accounts for the rapid rate of electron transfer be¬  tween the two redox centers.   Cytochrome c oxidase (Complex IV) catalyzes the one-electron oxidations of  four consecutive reduced cytochrome c molecules and the concomitant four-  electron reduction of one 0 2 molecule:   4 Cytochrome c (Fe 2+ ) + 4 H + +0 2 —>4 cytochrome c (Fe 3+ ) + 2 H 2 0   Mammalian Complex IV is an ~4lO-kD homodimer whose component  protomers are each composed of 13 subunits. The X-ray structure of Complex  IV from bovine heart mitochondria, determined by Shinya Yoshikawa, reveals   FIG. 1 8-1 7 X-Ray structure of the bovine  heart cytochrome c oxidase homodimer. The   homodimeric complex is viewed from within the  plane of the membrane with the intermembrane  space at the top. The 13 different subunits in  each protomer, which collectively have 28 trans¬  membrane helices, are differently colored. The  protein’s bound heme groups and Cu ions are  drawn in space-filling form with C green, N blue,  0 red, Fe red-brown, and Cu cyan. The horizontal  black lines delineate the inferred position of the  inner mitochondrial membrane. [Based on an  X-ray structure by Shinya Yoshikawa, Himeji  Institute of Technology, Hyogo, Japan. PDBid  1V54J © See Interactive Exercise 21.    that ten of its subunits are transmembrane proteins that contain a total of 28  membrane-spanning a helices (Fig. 18-17). The core of Complex IV consists  of its three largest and most hydrophobic subunits, I, II, and III (pale green,  pink, and pale yellow in Fig. 18-17), which are the only subunits encoded by  mitochondrial genes. A concave area on the surface of the protein that faces  the intermembrane space contains numerous acidic amino acids that can po¬  tentially interact with the ring of Lys residues on cytochrome c, the electron  donor for Complex IV.   Complex IV contains four redox centers: cytochrome (V> cytochrome a 3 ,  a copper atom known as Cu B > and a pair of copper atoms known as the Cua  center (Fig. 18-18). The Cua center, which is bound to Subunit II, lies 8 A  above the membrane surface. Its two copper ions are bridged by the sulfur  atoms of two Cys residues, giving it a geometry similar to that of a [ 2 Fe- 2 S]  cluster. The other redox groups—Cu B and cytochromes a and a 3 —all bind to  Subunit I and lie ~13 A below the membrane surface.   Spectroscopic studies have shown that electron transfer in Complex IV is  linear, proceeding from cytochrome c to the Cua center, then to heme a, and  finally to heme and Cu B . The Fe of heme a 3 lies only 4.9 A from Cu B ;  these redox groups really form a single binuclear complex. Electrons appear to  travel between the redox centers of Complex IV via a hydrogen-bonded net¬  work, involving amino acid side chains, the polypeptide backbone, and the  propionate side chains of the heme groups.    Cytochrome c Oxidase Catalyzes a Four-Electron Redox Reaction. The re¬  duction of 0 2 to 2 H 2 0 by cytochrome c oxidase takes place at the cy¬  tochrome a 3 -C u B binuclear complex and requires the nearly simultaneous  input of four electrons. However, the fully reduced Fe(II)-Cu(I) binuclear  complex can readily contribute only three electrons to its bound 0 2 in reach¬  ing its fully oxidized Fe(IV)-Cu(II) state [cytochrome a 3 assumes its Fe(IV)  or ferryl oxidation state during the reduction of 0 2 ; see below]. What is the  source of the fourth electron?   X-Ray structures of cytochrome c oxidase clearly indicate that the His 240  ligand of Cu B is covalently bonded to the side chain of a conserved Tyr residue    FIG. 18-18 The redox centers of bovine heart cytochrome c oxidase.   The view is similar to that of the left protomer in Fig. 18-17. The Fe and Cu  ions are represented by orange and cyan spheres. Their liganding heme and  protein groups are drawn in stick form colored according to atom type  (heme C magenta, protein C green, N blue, 0 red, and S yellow). The peroxy  group that bridges the Cu B and heme a 3 Fe ions is shown in ball-and-stick  form in red. Coordination bonds are drawn as gray lines. Note that the side  chains of His 240 and Tyr 244 are joined by a covalent bond (lower right).  [Based on an X-ray structure by Shinya Yoshikawa, Himeji Institute of  Technology, Hyogo, Japan. PDBid 20CC.]    FIG. 18-19 Proposed reaction sequence  for cytochrome c oxidase. A total of four  electrons ultimately donated by four cytochrome  c molecules, together with four protons, are  required to reduce 0 2 to H 2 0 at the cytochrome  a 3 -Cu B binuclear complex. The numbered steps  are discussed in the text The entire reaction is  extremely fast; it goes to completion in — 1 ms  at room temperature. [Modified from Babcock,  G.T., Proc. Natl. Acad Sci. 96, 12971 (1999).]    How does Tyr 244 function in the reduc¬  tion of 0 2 ?    (Tyr 244; Fig. 18-18, lower right). This places the Tyr phenolic —OH group  close to the heme a^— ligated 0 2 such that Tyr 244 can supply the fourth elec¬  tron by transiently forming a tyrosyl radical (TyrO*)- Tyrosyl radicals have been  implicated in several other enzyme-mediated redox processes, including the  generation of 0 2 from H 2 0 in photosynthesis (Section 19-2C) and in the  ribonucleotide reductase reaction (which converts NDP to dNDP; Section  23-3A). In cytochrome c oxidase, the Tyr phenolic —OH group is within  hydrogen-bonding distance of the enzyme-bound 0 2 and hence is a likely H +  donor during O—O bond cleavage. The formation of the covalent cross-link  is expected to lower both the reduction potential and the p K of Tyr 244,  thereby facilitating both radical formation and proton donation.   A proposed reaction sequence for cytochrome c oxidase, which was elucidated  through the use of a variety of spectroscopic techniques, is shown in Fig. 18-19:   1 and 2. The O (for oxidized) state binuclear complex [Fe(III )^ 3 —OH  H 2 0—Cu(II) B ], in which Tyr 244 is in its phenolate state  (Y—O ), is reduced by two consecutive one-electron transfers  from cytochrome c via Cu A and cytochrome a, each accompanied  by the acquisition of a proton from the matrix and the release of  an H 2 0. This yields the R (for reduced) complex [Fe(II) Cu(I)]  in which Tyr 244 has assumed its phenolic state (Y—OH).   3. 0 2 binds to the R state binuclear complex so as to ligand its  Fe(II )^ 3 atom. It binds to the heme with much the same con¬  figuration it has in oxymyoglobin (Fig. 7-3). Tyr 244 remains  in its phenolic state.   4. Internal electron redistribution rapidly yields the oxyferryl com¬  plex [Fe(IV) = 0 2 HO —Cu(II)] in which Tyr 244 has do¬  nated an electron and a proton to the complex and thereby  assumed its neutral radical state (Y—O*)- This is known as the  P state because it was once thought to be a y?eroxy compound.   5. A third one-electron transfer from cytochrome c, together with  the acquisition of a proton from the matrix, converts Tyr 244 to  its phenolate state (Y—O ), yielding the F (for /erryl) state in  which an H 2 0 remains liganded to Cu(II) B .    Cytochrome c Oxidase Has Two Proton-Translocating Channels. The reac¬  tion catalyzed by cytochrome c oxidase contributes to the transmembrane pro¬  ton gradient in two ways. First, four so-called chemical or scalar protons are  taken up from the matrix during the reduction of 0 2 by cytochrome c oxidase  to yield 2 H 2 0, thereby depleting the matrix [H + ]. Second, the four-electron  reduction reaction is coupled to the translocation of four so-called pumped or  vectorial protons from the matrix to the intermembrane space, one each in steps  3, 4, 5, and 6 of Fig. 18-19. Note that for each turnover of the enzyme,   a total of eight positive charges are lost from the matrix, thus contributing to  the membrane potential difference that drives ATP synthesis (Section 18-3).   X-Ray structures of cytochrome c oxidase reveal the presence of two chan¬  nels that lead from the matrix to the vicinity of the 0 2 -reducing center, which  could potentially transport protons via a proton wire mechanism. The  K-channel (so named because it contains an essential Lys residue) leads from  the matrix side of the protein to Tyr 244, the residue that forms a free radical  as described above. Since this channel does not appear to be connected to the  intermembrane space, it is thought to supply the first two chemical protons  for 0 2 reduction (steps 1 and 2 of Fig. 18-19). The D-channel (named for a  key Asp residue) extends from the matrix to the vicinity of the heme a 3 -C u B  center, where it connects to the so-called exit channel, which communicates  with the intermembrane space. Apparently, the D-channel, in series with the  exit channel, functions to pump vectorial protons from the matrix to the in¬  termembrane space. Moreover, the D-channel also functions as a conduit for  the other two chemical protons that are required for the reaction cycle (steps  5 and 6 of Fig. 18-19). Despite the foregoing, the mechanism that couples  0 2 reduction to proton pumping in Complex IV remains largely an enigma.   • Describe the route followed by electrons  from glucose to 0 2 .   • Assuming 100% efficiency, calculate the  maximum amount of ATP that could be  synthesized as a result.   • For each of the electron-transport com¬  plexes, write the relevant redox half¬  reactions.   • Position the four electron-transport com¬  plexes on a graph showing their relative  reduction potentials, and indicate the path  of electron flow.   • List the types of prosthetic groups in  Complexes I, II, III, and IV and state  whether they are one- or two-electron  carriers.   • What does the Q cycle accomplish in  each of its two rounds?   • Describe the different mechanisms for  translocating protons during electron  transport.   • What is the significance of the Tyr radical  in cytochrome c oxidase?    • The chemiosmotic theory explains how a proton gradient links electron transport  to ATP synthesis.   • ATP synthase consists of an F-, component that catalyzes ATP synthesis by a  binding change mechanism.   • The F 0 component of ATP synthase includes a c-ring whose rotation is driven by  the dissipation of the proton gradient and drives conformational changes in the F-,  component.   • For every two electrons that enter the electron-transport chain as NADH and  reduce one oxygen atom, approximately 2.5 ATP molecules are produced, giving a  P/O ratio of 2.5.   • Agents that dissipate the proton gradient can uncouple electron transport and  ATP synthesis.    The endergonic synthesis of ATP from ADP and P z in mitochondria is cat¬  alyzed by an ATP synthase (also known as Complex V) that is driven by the  electron-transport process. The free energy released by electron transport through  Complexes I-IV must be conserved in a form that the ATP synthase can use. Such  energy conservation is referred to as energy coupling.   The physical characterization of energy coupling proved to be surprisingly  elusive; many sensible and often ingenious ideas failed to withstand the test  of experimental scrutiny. For example, one theory—now abandoned—was that  electron transport yields a “high-energy” intermediate, such as phospho-  enolpyruvate (PEP) in glycolysis (Section 15-2J), whose subsequent breakdown  drives ATP synthesis. No such intermediate has ever been identified. In fact,   Chapter 18 Electron Transport and Oxidative ATP synthesis is coupled to electron transport through the formation of a   Phosphorylation transmembrane proton gradient during electron transport by Complexes I, III,   and IV. In this section, we explore this coupling mechanism and the opera¬  tion of ATP synthase.   The chemiosmotic theory, proposed in 1961 by Peter Mitchell, spurred con¬  siderable controversy before becoming widely accepted (Box 18-2). Mitchells  theory states that the free energy of electron transport is conserved by pumping  H + from the mitochondrial matrix to the intermembrane space to create an elec¬  trochemical H + gradient across the inner mitochondrial membrane. The electro¬  chemical potential of this gradient is harnessed to synthesize ATP (Fig. 18-20).  Several key observations are explained by the chemiosmotic theory:   2 . The inner mitochondrial membrane is impermeable to ions such as  H + , OH , K + , and Cl , whose free diffusion would discharge an elec¬  trochemical gradient.   3. Electron transport results in the transport of H + out of intact mito¬  chondria (the intermembrane space is equivalent to the cytosol),  thereby creating a measurable electrochemical gradient across the inner  mitochondrial membrane.   4. Compounds that increase the permeability of the inner mitochon¬  drial membrane to protons, and thereby dissipate the electrochemical  gradient, allow electron transport (from NADH and succinate oxida¬  tion) to continue but inhibit ATP synthesis; that is, they “uncouple”  electron transport from oxidative phosphorylation. Conversely, in¬  creasing the acidity outside the inner mitochondrial membrane stim¬  ulates ATP synthesis.    FIG. 1 8-20 The coupling of electron transport and ATP  synthesis. Electron transport (blue arrow) generates a proton  electrochemical gradient across the inner mitochondrial membrane.  H + is pumped out of the mitochondrion during electron transport    (red arrows) and its exergonic return powers the synthesis of ATP  (green arrows). Note that the intermembrane space is topologically  equivalent to the cytosol because the outer mitochondrial membrane is  permeable to H + . Q See the Animated Figures.   the most dramatic paradigm shifts in  biochemistry came about through the work  of Peter Mitchell, whose chemiosmotic  hypothesis linked biological electron  transport to ATP synthesis. Mitchell was  primarily a theoretical biochemist, although  he also generated experimental data to  support his hypothesis. He once compared  the human mind to a garden planted with  facts and ideas that are constantly being rearranged. However, he  promoted his own ideas, which were highly controversial, with great  tenacity and very little flexibility.   Mitchell graduated from the University of Cambridge in 1942  and conducted research there, laying the groundwork for the  chemiosmotic theory. Mitchell continued his work after 1955 at  the University of Edinburgh. He was captivated by the idea of  compartmentation in living cells and by the vectorial, or one-way,  aspect of metabolic processes. Initially, he chose to study  phosphate transport in bacteria because this process was linked  both to metabolism and to transmembrane transport. He realized  that the vectorial nature of membrane transport must be due to  the presence of membrane-associated systems that were driven  by chemical forces.   Mitchell also became interested in the respiratory chain, an  idea formulated by David Keilin at Cambridge, because this too  was a clearly vectorial biological phenomenon. Mitchell reasoned  that there must be enzymes that, like transporters, convert a  substrate on one side of a membrane to a product on the other  side. Other researchers had already discovered that the activity of  the respiratory chain generated a pH gradient. Mitchell’s genius  was to explain how the pH gradient could drive ATP synthesis. In  his seminal paper of 1961, Mitchell proposed that the respiratory  chain, associated with the cristae in the mitochondrion, generates  a protonmotive force due to electrical and pH differences across  the membrane. This force drives an ATPase, working in reverse, to  catalyze the condensation of phosphate with ADP to produce ATP.   Mitchell’s hypothesis, elegant as it was, met strong resistance  from other biochemists for several reasons. First, chemiosmosis  was a theoretical notion without direct experimental evidence.  Second, the study of oxidative phosphorylation was dominated by  a few powerful laboratories that were not inclined to welcome  new theories. In particular, metabolic studies since the mid-1940s  had been focused on the activities of soluble enzymes. Mitchell’s  theory was not a product of this classic biochemical approach but  instead came through an understanding of membrane physiology.  Furthermore, the prevailing theories about the connection between  electron transport and ATP synthesis centered on a phosphorylated    compound as a high-energy intermediate. Fritz Lipmann (Box 14-2)  had proposed that a high-energy phosphate group might become  attached to some component of the respiratory chain. This  hypothesis was later amended to invoke a soluble phosphorylated  intermediate. The search for the elusive compound lasted some  20 years, and the investment of time and money may have made  some researchers hesitant to abandon the theory in favor of  Mitchell’s seemingly outrageous proposal. In the meantime, a third  theory was proposed, in which electron transport was coupled to  ATP synthesis through protein conformational changes, with the  observed pH gradient presumed to be simply a by-product of the  process.   Mitchell’s chemiosmotic hypothesis did not garner significant  support until about 10 years after its publication. During this  period of sometimes acrimonious debate, Mitchell became ill,  moved to Cornwall, and renovated a manor house, part of which  became a private laboratory, known as Glynn Research, that was  funded by Mitchell’s family fortune. Here, he and his lifelong  collaborator, Jennifer Moyle, produced experimental evidence to  support the chemiosmotic theory. Ultimately, Mitchell was proven  correct by other researchers who demonstrated the proton¬  pumping activity of purified mitochondrial components  reconstituted in liposomes. Mitchell was awarded a Nobel Prize in  Chemistry in 1978.   Mitchell succeeded in changing the prevailing views of a  central feature of aerobic metabolism, although it was a long  battle. He later expressed sadness that his work was taken for  granted, as if it had been “self-evident from the beginning.” Oddly,  Mitchell stubbornly resisted altering any of his own ideas. For  example, he never wavered in his belief that protons participate  directly in ADP phosphorylation in the active site of ATP synthase.  And for many years, Mitchell refused to acknowledge proton  pumping (as we now know occurs in Complexes I and IV). Instead,  he insisted that the source of the proton gradient was a “redox  loop” in which two electrons are transferred from the positive to  the negative side of the membrane and combine with two protons  to reduce a quinone to a quinol. The quinol then diffuses back  across the bilayer to be reoxidized at the positive side, where the  protons are released. The redox loop mechanism, which requires  two “active sites,” does occur during the Q cycle in mitochondrial  Complex III and in certain bacterial systems, but it cannot account  for the entire protonmotive force that is the heart of the  chemiosmotic mechanism.    Mitchell, P., Coupling of phosphorylation to electron and hydrogen transfer by a  chemiosmotic type of mechanism, Nature 191 , 144-148 (1961).   Prebble, J., Peter Mitchell and the ox phos wars, Trends Biochem. Sci. 27 , 209-212  ( 2002 ).    An entirely analogous process occurs in bacteria, whose electron-transporting  machinery is located in their plasma membranes (Box 18-3).   Electron Transport Generates a Proton Gradient. Electron transport , as we have  seen ., causes Complexes I, III\ and IV to transport protons across the inner mitochon¬  drial membrane from the matrix , a region of low /7 / + ], to the intermembrane space  (which is in contact with the cytosol), a region of high [H + ] (Fig. 18-8). The free    It comes as no surprise that aerobic bacteria (such as those  pictured below), whose ancestors gave rise to mitochondria,  use similar machinery to oxidize reduced coenzymes and con¬  serve their energy in ATP synthesis. In bacteria, the compo¬  nents of the respiratory electron-transport chain are located in  the plasma membrane, and protons are pumped from the  cytosol to the outside of the plasma membrane. Protons flow  back into the cell via an ATP synthase, whose catalytic compo¬  nent is oriented toward the cytosol. This is exactly the arrange¬  ment expected if bacteria and mitochondria are evolutionarily  related.   The oxidation of CoQH 2 is universal in aerobic organisms. In  mitochondria, CoQ collects electrons donated by NADH (via  Complex I), succinate (via Complex II), and fatty acids. In aerobic  bacteria, CoQ is the collection point for electrons extracted by  dehydrogenases specific for a wide variety of substrates. In  bacteria, as in mitochondria, electrons flow from CoQ through  cytochrome-based oxidoreductases before reaching 0 2 (below,  right). In some species, two protein complexes (analogous to mito¬  chondrial Complexes III and IV) carry out this process. In other  species, including E. coli, a single type of enzyme, quinol  oxidase, uses the electrons donated by CoQ to reduce 0 2 .    The advantage of a multicomplex electron-transport pathway is  that it affords more opportunities for proton translocation across  the bacterial membrane, so the ATP yield per electron is greater.  However, the shorter electron-transport pathways may confer a  selective advantage in the presence of toxins that inactivate the  bacterial counterpart of mitochondrial Complex III. Multiple routes  for electron transport probably also allow bacteria to adjust oxida¬  tive phosphorylation to the availability of different energy sources  and to balance ATP synthesis against the regeneration of various  reduced coenzymes. For example, in facultative anaerobic bacteria  (which can grow in either the absence or presence of 0 2 ), when  energy needs are met through anaerobic fermentation, electron  transport can be adjusted to regenerate NAD + without synthesiz¬  ing ATP by oxidative phosphorylation.   A variety of cytochrome-containing protein complexes occur in  bacterial plasma membranes. Some of these proteins represent  more streamlined versions of the mitochondrial complexes since  they lack the additional subunits encoded by the nuclear genome  of eukaryotes. However, this is not a universal feature of respira¬  tory complexes, and many bacterial proteins (e.g., cytochrome d)  have no counterparts encoded by either the mitochondrial or  nuclear genomes.    energy sequestered by the resulting electrochemical gradient (also called the  protonmotive force; pmf) powers ATP synthesis.   The free energy change of transporting a proton from one side of the  membrane to the other has a chemical as well as an electrical component, since  H + is an ion (Section 10-1). AG is therefore expressed by Eq. 10-3, which in  terms of pH is   where Z is the charge on the proton (including sign), is the Faraday con¬  stant, and A' V E is the membrane potential. The sign convention for A'T' is that  when a proton is transported from a negative region to a positive region, A'T'  is positive. Since the pH outside the mitochondrion {side 2) is less than the  pH of the matrix {side 1), the export of protons from the mitochondrial matrix  (against the proton gradient) is an endergonic process.    Calculate the free energy change for transporting a proton out of the mitochon¬  drial matrix when the matrix pH is 8 and the cytosolic pH is 7. Assume that  A^P = 0.168 V (inside negative) and T= 37°C.   A G = 2.3 RT [pH {side 1) - pH {side 2 )] + Z^AT  A G= 2.3(8.3143 J-K _1 -mor 1 )(310K)(8-7) +  (1)(96,485 J • V -1 • m O r 1 )(0.168 V)   = 5900 J-mol -1 + 16,200 J- mol -1  = 22,100 J-mol -1 = 22.1 kJ-moU 1    The measured membrane potential across the inner membrane of a liver  mitochondrion, for example, is 0.168 V (inside negative). The pH of its ma¬  trix is 0.75 units higher than that of its intermembrane space. AG for proton  transport out of this mitochondrial matrix is therefore 21.5 kj • mol 1 (see  Sample Calculation 18-1). Because formation of the proton gradient is an en-  dergonic process, discharge of the gradient is exergonic. This free energy is  harnessed by ATP synthase to drive the phosphorylation of ADP.   An ATP molecule’s estimated physiological free energy of synthesis, around  + 40 to +50 kj • mol \ is too large for ATP synthesis to be driven by the  passage of a single proton back into the mitochondrial matrix; at least two  protons are required. In fact, most experimental measurements (which are dif¬  ficult to precisely quantitate) indicate that around three protons are required  per ATP synthesized.    ATP synthase, also known as proton-pumping ATP synthase and F 1 F 0 -  ATPase, is a multisubunit transmembrane protein with a total molecular mass  of 450 kD. Efraim Racker discovered that mitochondrial ATP synthase is com¬  posed of two functional units, F 0 and F x . F 0 is a water-insoluble transmembrane  protein containing as many as eight different types of subunits. F x is a water-  soluble peripheral membrane protein, composed of five types of subunits, that  is easily and reversibly dissociated from F 0 by treatment with urea. Solubilized  F x hydrolyzes ATP but cannot synthesize it (hence the name ATPase).   Electron micrographs of the inner mitochondrial membrane reveal that its  matrix surface is studded with molecules of ATP synthase whose F x compo¬  nent is connected to the membrane-embedded F 0 component by a protein  stalk, thereby giving F x a lollipop-like appearance (Fig. 18-21). Similar enti¬  ties have been observed lining the inner surface of the bacterial plasma mem¬  brane and in chloroplasts (Section 19-2D).   The F a Component Has Pseudo-Threefold Symmetry. The F x component  of mitochondrial ATP synthase has the subunit composition a 3 (3 3 y88. The  X-ray structure of the 3440-residue (371-kD) F x subunit from bovine heart  mitochondria, determined by John Walker and Andrew Leslie, reveals that it  consists of a 100-A-high and 100-A-wide spheroid that is mounted on a    FIG. 18-21 Electron micrograph of mitochondrial cristae. The “lollipops” projecting  into the matrix are the F^ components of ATP synthase. [From Parsons, D.F., Science 140,  985 (1963). Copyright © 1963 American Association for the Advancement of Science.  Used by permission.]   FIG. 1 8-22 X-Ray structure of Ft-ATP synthase from bovine  heart mitochondria, (a) The protein is drawn as a semitransparent  ribbon viewed parallel to the membrane with the matrix above. The a,  p, 7 , 8 , and 8 subunits are pink, yellow, blue, orange, and green,  respectively. Bound nucleotides are shown in space-filling form with  ATP C green, ADP C cyan, N blue, 0 red, and P orange, (b) The    structure in Part a rotated 90° about the horizontal axis such that the  protein is viewed along its pseudo-three-fold axis from the matrix. The  8 and 8 subunits have been deleted for clarity. [Based on an X-ray  structure by Andrew Leslie and John Walker, MRC Laboratory of  Molecular Biology, Cambridge, U.K. PDBid 1 E79J Q See Interactive  Exercise 22.    50-A-long stem (Fig. 18-22^). The a and P subunits, which are 20% identi¬  cal in sequence and have nearly identical folds, are arranged alternately, like the  segments of an orange, around the upper portion of a 114-A-long a helix  formed by the y subunit (Fig. 18-22 A). The lower portion of the helix forms  an antiparallel coiled coil with the N-terminal segment of the y subunit. This  coiled coil, along with the 8 and 8 subunits that are wrapped around it, links  F x to F 0 .   The cyclic arrangement and structural similarities of F x s a and p sub¬  units give it both pseudo-threefold and pseudo-sixfold rotational symmetry  (Fig. 18-22 A- Nevertheless, the protein is asymmetric due to the presence of  the y subunit but, more importantly, because each pair of a and P subunits  adopts a different conformation, each with a different substrate affinity.  Although the a subunits can bind ADP or ATP, only the P subunits catalyze  ATP synthesis. Thus one p subunit (designated Pdp) binds a molecule of the  substrate ADP, the second (Ptp) binds the product ATP, and the third (P E )  has an empty and distorted binding site. These conformational states are la¬  beled in Figure 18-22 A although p TP contains a bound ADP rather than ATP  in this model.   The F 0 Component Includes a Transmembrane Ring. The F 0 component  of bacterial and mitochondrial F^-ATPases consists of multiple subunits. In  E. colU three transmembrane subunits— a, A and c —form an cL x b 2 c X2 com¬  plex. Mitochondrial F 0 contains additional subunits whose functions are un¬  clear. The c subunits, each of which contains two a helices, associate to form   E. coli F-i F 0 -ATPase. The model is based on the  X-ray structure of the E. coli Ft subunit (PDBid  1JNV), which resembles that of bovine Ft (Fig.  18-22), and X-ray and NMR structures of 8 and  the transmembrane segment of the b protein  from E. coli (PDBids 2A7U and 1 B9U). The  structures of a and the so-called hinge region of  b are unknown. [Courtesy of Peter Dimroth, ETH,  Zurich, Switzerland.]    a ring that is embedded in the membrane. Studies of F 0 from different species  indicate that the number of c subunits varies from 10 to 15. The sequence of  the mitochondrial a subunit suggests that this highly hydrophobic 271 -residue  protein forms five transmembrane helices.   In E. coli , the two b subunits plus the 8 protein (which, due to confusing  nomenclature, is not a counterpart of the eukaryotic 8 protein) form a periph¬  eral stalk that links the a subunit of F 0 to the top of F x . In vertebrates, this stalk  consists of at least four proteins. Figure 18-23 shows a composite model of the  E. coli ATP synthase, based on X-ray and NMR structures of its various compo¬  nents, which of course resemble their mitochondrial counterparts. The c ring is  largely embedded in the membrane, with the 8 subunit (the counterpart of the  mitochondrial 8 subunit) forming extensive contacts with the top of the c ring  and the y subunit of F x . The y subunit passes through the center of the a 3 (3 3  assembly.   ATP Is Synthesized by the Binding Change Mechanism. The mechanism  of ATP synthesis by proton-translocating ATP synthase can be conceptually  broken down into three phases:   2 . Catalysis of formation of the phosphoanhydride bond of ATP carried  out by F x .   3. Coupling of the dissipation of the proton gradient with ATP synthe¬  sis, which requires interaction of F x and F 0 .   Considerable evidence supports a mechanism for ATP formation proposed  by Paul Boyer. According to this binding change mechanism, F x has three  interacting catalytic protomers (a|3 units), each in a different conformational  state: one that binds substrates and products loosely (L state), one that binds  them tightly (T state), and one that does not bind them at all (open or O  state). The free energy released on proton translocation is harnessed to intercon¬  vert these three states. The phosphoanhydride bond of ATP is synthesized only   FIG. 1 8-24 The binding change mechanism for ATP  synthase. F ^ has three chemically identical but conformationally  distinct interacting a|3 protomers: 0, the open conformation, has very  low affinity for ligands and is catalytically inactive; L binds ligands  loosely and is catalytically inactive; T binds ligands tightly and is  catalytically active. ATP synthesis occurs in three steps: (1) ADP and  P/ bind to site L ( 2 ) An energy-dependent conformational change  converts binding site L to T, T to 0, and 0 to L (3) ATP is  synthesized at site T and ATP is released from site 0. The enzyme    returns to its initial state after two more passes of the reaction  sequence. The free energy that drives the conformational change is  transmitted to the catalytic a 3 p 3 assembly via the rotation of the 78  assembly (78 in E. coli), here represented by the centrally located  asymmetric object (green). [After Cross, R.L., Annu. Rev. Biochem. 50,  687 (1980).] Q See the Animated Figures.    How far does the y subunit rotate during the reaction  shown here?   FIG. 1 8-25 The mitochondrial t* 3 p 3  assembly viewed from the matrix. The protein  surface is colored according to its electrical  potential, with positive potentials blue, negative  potentials red, and neutral potentials white. Note  the absence of charge on the inner surface of  the sleeve. The portion of the y subunit’s  C-terminal helix that contacts the sleeve is  similarly devoid of charge. [From Abrahams, J.P.,  Leslie, A.G.W., Lutter, R., and Walker, J.E., Nature  370, 621 (1994). PDBid 1BMFJ    in the T state, and ATP is released only in the O state. The reaction involves  three steps (Fig. 18-24):   1. ADP and P z bind to the loose (L) binding site ((3 DP in Fig. 18-22^).   2 . A free energy-driven conformational change converts the L site to a  tight (T) binding site (pxp) that catalyzes the formation of ATP This  step also involves conformational changes of the other two protomers  that convert the ATP-containing T site to an open (O) site ((3 E ) and  convert the O site to an L site.   3. ATP is synthesized at the T site on one subunit while ATP dissociates  from the O site on another subunit. The reaction forming ATP is es¬  sentially at equilibrium under the conditions at the enzymes active site.  The free energy supplied by the proton flow primarily facilitates the  release of the newly synthesized ATP from the enzyme; that is, it drives  the T —» O transition, thereby disrupting the enzyme-ATP interac¬  tions that had previously promoted the spontaneous formation of ATP  from ADP + P z in the T site.   How is the free energy of proton transfer coupled to the synthesis of ATP?  The cyclic nature of the binding change mechanism led Boyer to propose that  the binding changes are driven by the rotation of the catalytic assembly ; ct 3 f3 3 ,  with respect to other portions of the FjF 0 -ATPase. This hypothesis is supported  by the X-ray structure of F x . Thus, the closely fitting nearly circular arrange¬  ment of the a and (3 subunits’ inner surface about the y subunits helical  C-terminus is reminiscent of a cylindrical bearing rotating in a sleeve. Indeed,  the contacting hydrophobic surfaces in this assembly are devoid of the hydrogen¬  bonding and ionic interactions that would interfere with their free rotation  (Fig. 18-25); that is, the bearing and sleeve appear to be “lubricated.”  Moreover, the central cavity in the a 3 (3 3 assembly (Figs. 18-22^ and 18-23)  would permit the passage of the y subunits N-terminal helix within the core  of the particle during rotation. Finally, the conformational differences between  F x s three catalytic sites appear to be correlated with the position of the y sub¬  unit. Apparently the y subunit, which rotates within the fixed oi 3 f> 3 assembly, acts  as a molecular camshaft in linking the proton gradient—driven F 0 engine to the  conformational changes in the catalytic sites of Fj.    The F-|Fo-ATPase Is a Rotary Engine. The proposed rotation of the a 3 (3 3 as¬  sembly with respect to the y subunit engendered by the binding change mech¬  anism has led to the model of the F^o-ATPase diagrammed in Fig. 18-26.  A rotational engine must have both a rotor (which rotates) and a stator (which  is stationary). In the F^o-ATPase, the rotor is proposed to be an assembly of   FIG. 1 8-26 Model of the E. coli  F-|F 0 -ATPase. The 78 -c 12 ring complex is the  rotor and the ab 2 -a 3 p 3 8 complex is the stator.  Rotational motion is imparted to the rotor by the  passage of protons from the outside (periplasmic  space, bottom ; equivalent to the mitochondrial  intermembrane space) to the inside (cytoplasm,  top ; equivalent to the mitochondrial matrix).  Protons entering from the outside bind to a c  subunit where it interacts with the a subunit, and  exit to the inside after the c-ring has made a  nearly full rotation as indicated (black arrows ), so  that the c subunit again contacts the a subunit.  The b 2 8 complex presumably functions to  prevent the a 3 p 3 assembly from rotating with  the 7 subunit. [After a drawing by Richard Cross,  State University of New York, Syracuse, New York.]    the c-ring with the y and (E. coli ) £ subunits, whereas the ab 2 unit and the  (E. coli ) 8 subunit together with the a 3 (3 3 spheroid form the stator. The ro¬  tation of the c-ring in the membrane relative to the stationary a subunit is  driven by the migration of protons from the outside to the inside as we dis¬  cuss below (here “outside” refers to the mitochondrial intermembrane space  or the bacterial exterior, whereas “inside” refers to the mitochondrial matrix  or the bacterial cytoplasm). The b 2 h assembly presumably functions to hold  the a 3 (S 3 spheroid in position while the 7 subunit rotates inside it.   In the model for proton-driven rotation of the F 0 subunit that is dia¬  grammed in Fig. 18-26, protons from the outside enter a hydrophilic chan¬  nel between the a subunit and the c-ring, where they bind to a c subunit. The  c-ring then rotates nearly a full turn (while protons bind to successive c sub¬  units as they pass this input channel) until the subunit reaches a second hy¬  drophilic channel between the a subunit and the c-ring that opens into the  inside, where the proton is released (in an alternative model, the protons are  released through putative channels between the C-terminal helices of adjacent  c subunits, channels that are occluded when the a and c subunits are in  contact). Thus, the F^Q-ATPase, which generates 3 ATP per turn and (at least  in E. coli ) has 12 c subunits in its F 0 assembly, ideally forms 3/12 = 0.25  ATP for every proton it passes from outside to inside.   How does the passage of protons through this system induce the rotation  of the c-ring and hence the synthesis of ATP? Each c subunit consists of two  a helices of different lengths that are connected by a four-residue polar loop  and arranged in an antiparallel coiled coil. Protons most likely bind to Asp 61  of each c subunit, an invariant residue whose protonation and deprotonation  alter the subunits conformation (Fig. 18-27). Evidently, when a c subunit  binds a proton as it passes the input channel, its conformation changes, which  causes it to mechanically push against the a subunit so as to induce the c-ring  to rotate in the direction indicated in Fig. 18-26. This process is augmented   FIG. 1 8-27 NMR structure of the c  subunit of E. coli F^o-MPase. (a) At pH 8 ,   Asp 61 (D61) is deprotonated. (b) At pH 5, D61  is protonated. Selected side chains are shown to  aid in the comparison of the two structures.   Note that the C-terminal helix in the pH 8  structure is rotated 140° clockwise, as viewed  from the top of the drawing, relative to that in  the pH 5 structure. [Courtesy of Mark Girvin,  Albert Einstein College of Medicine. PDBid 1COV.]    FIG. 1 8-28 The rotation of the c-ring in  E. coli F-|F 0 -ATPase. (a) The experimental  system used to observe the rotation. See the  text for details. The blue arrow indicates the  observed direction of rotation of the fluorescently  labeled actin filament that was linked to the  c-ring. (b) The rotation of a 3.6-fxm-long actin  filament in the presence of 5 mM MgATP as  seen in successive video images taken through a  fluorescence microscope. [Courtesy of Masamitsu  Futai, Osaka University, Osaka, Japan.]    by the interaction between Asp 61 on the c subunit and the in¬  variant Arg 210 on the a subunit. These two residues have been  shown to become juxtaposed at some point during the c-ring s  rotation cycle. It has therefore been proposed that the electro¬  static attraction between the cationic Arg 210 and the anionic  Asp 61 helps rotate the c-ring so as to bring the two residues  into opposition but, as this occurs, Asp 61 becomes protonated,  thereby forcing the c-ring to continue its rotation.   The rotation of the y8-c-ring rotor with respect to the  stator has been ingeniously demonstrated by  Masamitsu Futai (Fig. 18-2 8 a). The a 3 (3 3 spheroid of E. coli  FiFo-ATPase was fixed, head down, to a glass surface as follows.  Six consecutive His residues (a so-called His tag) were muta-  genically appended to the N-terminus of the a subunit, which  is located at the top of the a 3 (3 3 spheroid as it is drawn in Fig.  18-22 a. The His-tagged assembly was applied to a glass surface  coated with horseradish peroxidase (which, like most proteins,  sticks to glass) conjugated with Ni 2+ -nitriloacetic acid [Ni 2+ -  N(CH 2 COOH) 3 , which tightly binds His tags], thereby bind¬  ing the F^Q-ATPase to the surface with its F 0 side facing away  from the surface. The Glu 2 residues of this assembly’s c sub¬  units, which are located on the side of the c-ring facing away  from F 1? had been mutagenically replaced by Cys residues,  which were then covalently linked to biotin (Section 16-4A).  A fluorescently labeled and biotinylated (at one end) filament  of the muscle protein actin (Section 7-2C) was then attached  to the c subunit through the addition of a bridging molecule of streptavidin,  a bacterial protein that avidly binds biotin to each of four binding sites.   The E. coli F^Q-ATPase can work in reverse, that is, it can pump pro¬  tons from the inside to the outside at the expense of ATP hydrolysis (this en¬  ables the bacterium to maintain its proton gradient under anaerobic  conditions, which it uses to drive various processes). Thus, the foregoing  preparation was observed under a fluorescence microscope as a 5 mM MgATP  solution was infused over it. Many of the actin filaments were seen to rotate  (Fig. 18-28b), and always in a counterclockwise direction when viewed looking  down on the glass surface (from the outside). This would permit the y subunit  to sequentially interact with the (3 subunits in the direction    (3 E (O state) —» (3 DP (L state) —» (3 TP (T state)   (Figs. 18-22 a and 18-24), the direction expected for ATP hydrolysis. Similar  experiments revealed that the y subunit rotates mostly in increments of 120°.  Presumably, electrostatic interactions between the y and (3 subunits of F x act  as a catch that temporarily holds the y subunit in place. As the c-ring rotates,  strain builds up and causes the y subunit to snap to the next (3 subunit. In  addition, when a magnetic bead was instead attached to the y subunit of im¬  mobilized Fx-ATPase and an external magnetic field was rotated in the clock¬  wise direction so as to force the y subunit to follow, ATP was synthesized from  ADP + P z ,    C The P/0 Ratio Relates the Amount of ATP Synthesized to the  Amount of Oxygen Reduced   ATP synthesis is tightly coupled to the proton gradient; that is, ATP synthesis re¬  quires the discharge of the proton gradient, and the proton gradient cannot  be discharged without the synthesis of ATP. The proton gradient is established  through the activity of the electron-transporting complexes of the inner mito¬  chondrial membrane. Therefore, it is possible to express the amount of ATP    synthesized in terms of substrate molecules oxidized. Experiments with iso¬  lated mitochondria show that the oxidation of NADH is associated with the  synthesis of approximately 3 ATP, and the oxidation of FADH 2 with approx¬  imately 2 ATP. Oxidation of the nonphysiological compound tetramethyl-/>-  phenylenediamine,   which donates an electron pair directly to Complex IV, yields approximately  1 ATP. These stoichiometric relationships are called P/O ratios because they  relate the amount of ATP synthesized (P) to the amount of oxygen reduced   Experimentally determined P/O ratios are compatible with the chemios-  motic theory and the known structure of ATP synthase. The flow of two elec¬  trons through Complexes I, III, and IV results in the translocation of  10 protons into the intermembrane space (Fig. 18-8). Influx of these 10 protons  through the FiFo-ATPase, which contains 10 c subunits in eukaryotes, results  in one complete rotation of the c-ring—y subunit rotor relative to the a 3 |33  spheroid, enough to drive the synthesis of ~3 ATP. Electrons that enter the  electron-transport chain as FADH 2 at Complex II bypass Complex I and  therefore lead to the transmembrane movement of only 6 protons, enough to  synthesize ~2 ATP (corresponding to approximately two-thirds of a full rota¬  tion of the ATP synthase rotary engine). The transit of two electrons through  Complex IV alone contributes 2 protons to the gradient, enough for ~ 1 ATP  (around one-third of a rotation).   In actively respiring mitochondria, P/O ratios are almost certainly not in¬  tegral numbers. This is also consistent with the chemiosmotic theory. Oxidation  of a physiological substrate contributes to the transmembrane proton gradient  at several points, but the gradient is tapped at only a single point, the F^q-  ATPase. Therefore, the number of protons translocated out of the mitochon¬  drion by any component of the electron-transport chain need not be an  integral multiple of the number of protons required to synthesize ATP from  ADP + P z . Moreover, the proton gradient is dissipated to some extent by the  nonspecific leakage of protons back into the matrix and by the consumption  of protons for other purposes, such as the transport of P z into the matrix  (Section 18-IB). Taking P z transport into account gives a stoichiometry of four  protons consumed per ATP synthesized from ADP + P z . Thus, as Peter Hinkle  has experimentally demonstrated, the P/O ratios above are actually closer to  2.5, 1.5, and 1. Consequently, the number of ATPs that are synthesized per  molecule of glucose oxidized is 2.5 ATP/NADH X 10 NADH/glucose +  1.5 ATP/FADH 2 X 2 FADH 2 /glucose + 2 ATP/glucose from the citric acid  cycle + 2 ATP/glucose from glycolysis = 32 ATP/glucose (see page 569).   Electron transport (the oxidation of NADH and FADH 2 by 0 2 ) and oxida¬  tive phosphorylation (the proton gradient-driven synthesis of ATP) are nor¬  mally tightly coupled. This coupling depends on the impermeability of the inner  mitochondrial membrane, which allows an electrochemical gradient to be estab¬  lished across the membrane by H + translocation during electron transport. Virtually  the only way for H + to re-enter the matrix is through the F 0 portion of ATP  synthase. In the resting state, when oxidative phosphorylation is minimal, the  electrochemical gradient across the inner mitochondrial membrane builds up   FIG. 1 8-29 Action of 2,4-dinitrophenol. A proton-transporting ionophore such as  DNP uncouples oxidative phosphorylation from electron transport by discharging the  electrochemical proton gradient generated by electron transport.   • Explain why an intact, impermeable mito¬  chondrial membrane is essential for ATP  synthesis.   • Describe the overall structure of the F-,  and F 0 components of ATP synthase.  Which parts move? Which are stationary?  Which are mostly stationary but undergo  conformational changes?   • Describe how protons move from the  intermembrane space into the matrix. How  is proton translocation linked to ATP syn¬  thesis?   • Explain why the P/O ratio for a given  substrate is not necessarily an integer.   • Explain how oxidative phosphorylation is  linked to electron transport and how the  two processes can be uncoupled.    to the extent that it prevents further proton pumping and therefore inhibits  electron transport. When ATP synthesis increases, the electrochemical gradient  dissipates, allowing electron transport to resume.   Over the years, compounds such as 2,4-dinitrophenol (DNP) have been  found to “uncouple” electron transport and ATP synthesis. DNP is a lipophilic  weak acid that readily passes through membranes in its neutral, protonated  state. In a pH gradient, it binds protons on the acidic side of the membrane,  diffuses through the membrane, and releases the protons on the membranes  alkaline side, thereby acting as a proton-transporting ionophore (Section  10-2A) and dissipating the gradient (Fig. 18-29). The chemiosmotic theory pro¬  vides a rationale for understanding the action of such uncouplers. The pres¬  ence in the inner mitochondrial membrane of an agent that increases its  permeability to Pf uncouples oxidative phosphorylation from electron transport  by providing a route for the dissipation of the proton electrochemical gradient that  does not require ATP synthesis. Uncoupling therefore allows electron transport  to proceed unchecked even when ATP synthesis is inhibited. Consequently, in  the 1920s, DNP was used as a “diet pill,” a practice that was effective in in¬  ducing weight loss but often caused fatal side effects. Under physiological con¬  ditions, the dissipation of an electrochemical H + gradient , which is generated by  electron transport and uncoupled from ATP synthesis, produces heat (see Box 18-4).    The rate of oxidative phosphorylation is coordinated with the cell’s other oxidative  pathways.   Although aerobic metabolism is efficient, it leads to the production of reactive  oxygen species.    An adult woman requires some 1500 to 1800 kcal (6300-7500 kj) of meta¬  bolic energy per day. This corresponds to the free energy of hydrolysis of over  200 mol of ATP to ADP and P z . Yet the total amount of ATP present in the  body at any one time is <0.1 mol; obviously, this sparse supply of ATP must  be continually recycled. As we have seen, when carbohydrates serve as the en¬  ergy supply and aerobic conditions prevail, this recycling involves glycogenol-  ysis, glycolysis, the citric acid cycle, and oxidative phosphorylation.   Of course, the need for ATP is not constant. There is a 100-fold change  in the rate of ATP consumption between sleep and vigorous activity. The   Heat generation is the physiological function of brown adipose  tissue (brown fat). This tissue is unlike typical (white) adipose  tissue in that it contains numerous mitochondria whose  cytochromes cause its brown color. Newborn mammals that lack  fur, such as humans, as well as hibernating mammals, all contain  brown fat in their neck and upper back that generates heat by  nonshivering thermogenesis. Other sources of heat are the ATP  hydrolysis that occurs during muscle contraction (in shivering or  any other movement) and the operation of ATP-hydrolyzing  substrate cycles (see Section 15-4B).   The mechanism of heat generation in brown fat involves the  regulated uncoupling of oxidative phosphorylation. Brown fat mito¬  chondria contain a proton channel known as uncoupling protein  (UCP1, also called thermogenin; see below). In cold-adapted  animals, UCP1 constitutes up to 15% of the protein in the inner  mitochondrial membranes of brown fat. The flow of protons  through UCP1 is inhibited by physiological concentrations of  purine nucleotides (ADP, ATP, GDP, GTP), but this inhibition can  be overcome by free fatty acids.    Thermogenesis in brown fat mitochondria is under hormonal  control (see diagram). Norepinephrine (1; noradrenaline) induces  the production of the second messenger cAMP ( 2 ) and thereby  activates protein kinase A (3; Section 13-3C). The kinase then  activates hormone-sensitive triacylglycerol lipase (4) by  phosphorylating it. The activated lipase hydrolyzes triacylglycerols  (5) to yield free fatty acids that counteract the inhibitory effect of  the purine nucleotides on UCP1 ( 6 ). The resulting flow of protons  through UCP1 dissipates the proton gradient across the inner  mitochondrial membrane. This allows substrate oxidation to  proceed (and generate heat) without the synthesis of ATP.   Adult humans have relatively little brown fat, but the mitochon¬  dria of ordinary adipose tissue and muscle appear to contain  uncoupling proteins known as UCP2 and UCP3. These proteins  may help regulate metabolic rates, and variations in UCP levels or  activity might explain why some people seem to have a “fast” or  “slow” metabolism. UCPs are being studied as targets for treating  obesity, since increasing the activity of UCPs could uncouple  respiration from ATP synthesis, thus permitting stored metabolic  fuels (especially fat) to be metabolized.    activities of the pathways that produce ATP are under strict coordinated control  so that ATP is never produced more rapidly than necessary. We have already dis¬  cussed the control mechanisms of glycolysis, glycogenolysis, and the citric acid  cycle (Sections 15-4, 16-3, and 17-4). In this section, we discuss the mecha¬  nisms that control the rate of oxidative phosphorylation.    A The Rate of Oxidative Phosphorylation Depends on the ATP  and NADH Concentrations   In our discussions of metabolic pathways, we have seen that most of their re¬  actions function close to equilibrium. The few irreversible reactions constitute  potential control points of the pathways and usually are catalyzed by regula¬  tory enzymes that are under allosteric control. In the case of oxidative phos¬  phorylation, the pathway from NADH to cytochrome c functions near  equilibrium:   \ NADH + cytochrome c(Fe 3+ ) + ADP + P z v   NAD + + cytochrome c(Fe 2+ ) + ATP AG' ~ 0   This pathway is therefore readily reversed by the addition of its product, ATP.  However, the cytochrome c oxidase reaction (the terminal step of the electron-  transport chain) is irreversible and is therefore a potential control site. Cytochrome  c oxidase, in contrast to most regulatory enzyme systems, appears to be con¬  trolled primarily by the availability of one of its substrates, reduced cytochrome  c (c 2+ ). Since c 2+ is in equilibrium with the rest of the coupled oxidative phos¬  phorylation system, the concentration of c 2+ ultimately depends on the in-  tramitochondrial ratios of [NADH]/[NAD + ] and [ATP]/[ADP] [PJ (the latter  quantity is known as the ATP mass action ratio). We can see, by rearrang¬  ing the foregoing equilibrium expression,   [c 3 + ] \ [NAD + ] / [ATP] eq   that the higher the [NAD H]/[NAD + ] ratio and the lower the ATP mass ac¬  tion ratio, the higher the concentration of reduced cytochrome c and thus the  higher the cytochrome c oxidase activity.   How is this system affected by changes in physical activity? In an individ¬  ual at rest, the rate of ATP hydrolysis to ADP and P z is minimal and the ATP  mass action ratio is high; the concentration of reduced cytochrome c is there¬  fore low and the rate of oxidative phosphorylation is minimal. Increased ac¬  tivity results in hydrolysis of ATP to ADP and P z , thereby decreasing the ATP  mass action ratio and increasing the concentration of reduced cytochrome c.  This results in an increase in the rate of electron transport and ADP phos¬  phorylation.   The concentrations of ATP, ADP, and P z in the mitochondrial matrix de¬  pend on the activities of the transport proteins that import these substances from  the cytosol. Thus, the ADP-ATP translocator and the P z transporter may play  a part in regulating oxidative phosphorylation. There is also some evidence that  Ca 2+ stimulates the electron-transport complexes and possibly ATP synthase  itself. This is consistent with the many other instances in which Ca 2+ directly  stimulates oxidative metabolic processes.   Mitochondria contain an 84-residue protein, called IF X that functions to  regulate ATP synthase. In actively respiring mitochondria, in which the ma¬  trix pH is relatively high, IF X exists as an inactive tetramer. However, below   pH 6.5, the protein dissociates into dimers and in this form inhibits the  ATPase activity of the F x component by binding to the interface between its  a DP and (3 DP subunits so as to trap the ATP bound to the (3 DP subunit. This  appears to be a mechanism to prevent ATP hydrolysis when respiratory ac¬  tivity (and therefore the proton gradient) is temporarily interrupted by  lack of 0 2 . Otherwise, the FiFo-ATPase would reverse its direction of  rotation as driven by the hydrolysis of ATP (then generated by glycol¬  ysis), thus depriving the cell of its remaining energy resources.   The primary sources of the electrons that enter the mitochondrial  electron-transport chain are glycolysis, fatty acid degradation, and  the citric acid cycle. For example, 10 molecules of NAD + are con¬  verted to NADH per molecule of glucose oxidized (Fig. 18-1). Not  surprisingly, the control of glycolysis and the citric acid cycle is co¬  ordinated with the demand for oxidative phosphorylation. An ade¬  quate supply of electrons to feed the electron-transport chain is  provided by regulation of the control points of glycolysis and the  citric acid cycle (phosphofructokinase, pyruvate dehydrogenase, cit¬  rate synthase, isocitrate dehydrogenase, and a-ketoglutarate dehy¬  drogenase) by adenine nucleotides or NADH or both, as well as by  certain metabolites (Fig. 18-30).   One particularly interesting regulatory effect is the inhibition  of phosphofructokinase (PFK) by citrate. When demand for ATP  decreases, [ATP] increases and [ADP] decreases. Because isocitrate  dehydrogenase is activated by ADP and a-ketoglutarate dehydro¬  genase is inhibited by ATP, the citric acid cycle slows down. This  causes the citrate concentration to build up. Citrate leaves the mi¬  tochondrion via a specific transport system and, once in the cytosol  acts to restrain further carbohydrate breakdown by inhibiting PFK.  The citrate concentration also builds up when the acetyl-CoA con¬  centration increases, which occurs, as we will see in Chapter 20,  during the oxidation of fatty acids. The inhibition of glycolysis by  fatty acid oxidation is called the glucose-fatty acid cycle or  Randle cycle (after its discoverer, Phillip Randle) although it is  not, in fact, a cycle. The Randle cycle allows fatty acids to be uti¬  lized as the major fuel for oxidative metabolism in heart muscle,  while conserving glucose for organs such as the brain, which  require it.   Not all organisms carry out oxidative phosphorylation. However, those  that do are able to extract considerably more energy from a given  amount of a metabolic fuel. This principle is illustrated by the Pasteur  effect (Section 15-3C): When anaerobically growing yeast are exposed  to oxygen, their glucose consumption drops precipitously. An analo¬  gous effect is observed in mammalian muscle; the concentration of  lactic acid (the anaerobic product of muscle glycolysis; Section 15-3A)  drops dramatically when cells switch to aerobic metabolism. These  effects are easily understood by examining the stoichiometries of    FIG. 1 8-30 The coordinated control of glycolysis and the citric acid  cycle. The diagram shows the effects of ATP, ADP, AMP, P,-, Ca 2+ , and the  [NADH]/[NAD + ] ratio (the vertical arrows indicate increases in this ratio).  Here a green dot signifies activation and a red octagon represents inhibition.  [After Newsholme, E. A. and Leech, A. R., Biochemistry for the Medical  Sciences, pp. 316, 320, Wiley (1983).] Q See the Animated Figures.   Q 5 H 12 O <5 + 2 ADP + 2 P z — >2 lactate + 2 H + +2 H 2 0 + 2 ATP   Thus, aerobic metabolism is up to 16 times more efficient than anaerobic glycol¬  ysis in producing ATP.   Aerobic metabolism has its drawbacks, however. Many organisms and tis¬  sues depend exclusively on aerobic metabolism and suffer irreversible damage  during oxygen deprivation (Box 18-5). Oxidative metabolism is also accom¬  panied by the production of low levels of reactive oxygen metabolites that,  over time, may damage cellular components. Evidently, the organisms that  have existed during the last 3 billion years (the period in which the earths at¬  mosphere has contained significant amounts of 0 2 ) exhibit physiological and  biochemical adaptations that permit them to take advantage of the oxidizing  power of 0 2 while minimizing the potential dangers of oxygen itself.   Partial Oxygen Reduction Produces Reactive Oxygen Species. Although  the four-electron reduction of 0 2 by cytochrome c oxidase is nearly always  orchestrated with great rapidity and precision, 0 2 is occasionally only partially  reduced, yielding oxygen species that readily react with a variety of cellular  components. Complexes I, II, and III also occasionally generate these reactive  oxygen species (ROS), presumably by leaking electrons. The best known ROS  is the superoxide radical:   Superoxide radical is a precursor of other reactive species. Protonation of 0 2 *  yields H0 2 # , a much stronger oxidant than 0 2 \ The most potent oxygen  species in biological systems is probably the hydroxyl radical, which forms  from the relatively harmless hydrogen peroxide (H 2 0 2 ):   H 2 0 2 + Fe 2+ -> -OH + OH - + Fe 3 +   The hydroxyl radical also forms through the reaction of superoxide with H 2 0 2 :   Although most free radicals are extremely short-lived (the half-life of 0 2 # is  1 X 10 6 s, and that of *OH is 1 X 10 9 s), they readily extract electrons  from other molecules, converting them to free radicals and thereby initiating  a chain reaction.   The random nature of free-radical attacks makes it difficult to characterize  their reaction products, but all classes of biological molecules are susceptible to  oxidative damage caused by free radicals. The oxidation of polyunsaturated lipids  in cells may disrupt the structures of biological membranes, and oxidative dam¬  age to DNA may result in point mutations. Enzyme function may also be com¬  promised through radical reactions with amino acid side chains. Because the  mitochondrion is the site of the bulk of the cells oxidative metabolism, its lipids,  DNA, and proteins probably bear the brunt of free radical—related damage.   r Several degenerative diseases, including Parkinson’s, Alzheimer’s, and  Huntington’s diseases, are associated with oxidative damage to mitochon¬  dria. Such observations have led to the free-radical theory of aging, which holds  that free-radical reactions arising during the course of normal oxidative metabolism  are at least partially responsible for the aging process. In fact, individuals with con¬  genital defects in their mitochondrial DNA suffer from a variety of symptoms  typical of old age, including neuromotor difficulties, deafness, and dementia.  Their genetic defects may make their mitochondria all the more susceptible to  the reactive oxygen species generated by the electron-transport machinery.    As outlined in Box 7-1, organisms larger than 1 mm thick require  circulatory systems to deliver nutrients to cells and dispose of  cellular wastes. In addition, the circulatory fluid in most larger  organisms contains proteins specialized for oxygen transport (e.g.,  hemoglobin). The sophistication of oxygen-delivery systems and  their elaborate regulation are consistent with their essential nature  and their long period of evolution.   What happens during oxygen deprivation? Consider two  common causes of human death, myocardial infarction (heart  attack) and stroke, which result from interruption of the blood  (0 2 ) supply to a portion of the heart or the brain, respectively. In  the absence of 0 2 , a cell, which must then rely only on glycolysis  for ATP production, rapidly depletes its stores of phosphocreatine  (a source of rapid ATP production; Section 14-2C) and glycogen.  As the rate of ATP production falls below the level required by  membrane ion pumps for maintaining proper intracellular ion  concentrations, osmotic balance is disrupted so that the cell and  its membrane-enveloped organelles begin to swell. The resulting  overstretched membranes become permeable, thereby leaking  their enclosed contents. For this reason, a useful diagnostic  criterion for myocardial infarction is the presence in the blood of  heart-specific enzymes, such as the H-type isozyme of lactate  dehydrogenase (Section 14-1C) and creatine kinase, which leak  out of necrotic (dead) heart tissue. Moreover, the decreased  intracellular pH that accompanies anaerobic glycolysis (because of    lactic acid production) permits the released lysosomal enzymes  (which are active only at acidic pH) to degrade the cell contents.  Thus, 0 2 deprivation leads not only to cessation of cellular activity  but to irreversible cell damage and cell death. Rapidly respiring  tissues, such as the heart and brain, are particularly susceptible to  damage by oxygen deprivation.   Cells Are Equipped with Antioxidant Mechanisms. Antioxidants destroy  oxidative free radicals such as 0 2 * and *OH. In 1969, Irwin Fridovich dis¬  covered that the enzyme superoxide dismutase (SOD), which is present in  nearly all cells, catalyzes the conversion of 0 2 * to H 2 0 2 .    Mitochondrial and bacterial SOD are both Mn-containing tetramers; eukary¬  otic cytosolic SOD is a dimer containing copper and zinc ions. The rate of  nonenzymatic superoxide breakdown is ~2 X 10 5 M 1 • s \ whereas the  rate of the Cu,Zn-SOD-catalyzed reaction is ~2 X 10 9 M 1 • s 1 .   This rate enhancement, which is close to the diffusion-controlled limit  (Section 12-IB), is apparently accomplished by electrostatic guidance of  the negatively charged superoxide substrate into the enzyme’s active site  (Fig. 18 -31). The active-site Cu ion lies at the bottom of a deep pocket  in each enzyme subunit. A hydrogen-bonded network of Glu 123, Glu  133, Lys 136, and Thr 137 at the entrance to the pocket facilitates the  diffusion of 0 2 * to a site between the Cu ion and Arg 143.   SOD is considered a first-line defense against reactive oxygen  species. The H 2 0 2 produced in the reaction, which can potentially   this cross section of the active site channel of Cu,Zn-SOD, the molecular surface is  represented by a dot surface that is colored according to charge: red, most negative;  yellow, negative; green, neutral; light blue, positive; dark blue, most positive. The  electrostatic field vectors are represented by similarly colored arrows. The Cu and Zn  ions at the active site are represented by orange and silver dotted spheres. The  Oy* -binding site is located between the Cu ion and the side chain of Arg 143.  [Courtesy of Elizabeth Getzoff, The Scripps Research Institute, La Jolla, California.]   • How do the ATP mass action ratio and  the IF-, protein regulate ATP synthesis?   • What control mechanisms link glycolysis,  the citric acid cycle, and oxidative phos¬  phorylation?    react to yield other reactive oxygen species, is degraded to water and oxygen  by enzymes such as catalase, which catalyzes the reaction   2 H 2 0 2 —> 2 H 2 0 + 0 2   and glutathione peroxidase (Box 15-4), which uses glutathione (GSH) as the  reducing agent:   2 GSH + H 2 0 2 -> GSSG + 2 H 2 0   The latter enzyme also catalyzes the breakdown of organic hydroperoxides.  Some types of glutathione peroxidase require Se for activity; this is one rea¬  son why Se appears to have antioxidant activity.   Other potential antioxidants are plant-derived compounds such as ascor¬  bate (vitamin C; Section 6-1C) and a-tocopherol (vitamin E; Section 9-IF).  These compounds may help protect plants from oxidative damage during pho¬  tosynthesis, a process in which H 2 0 is oxidized to 0 2 . Their efficacy as an¬  tioxidants in humans, however, is unproven.   • Electrons from the reduced coenzymes NADH and FADH 2 pass  through a series of redox centers in the electron-transport chain  before reducing 0 2 .   • During electron transfer, protons are translocated out of the mi¬  tochondrion to form an electrochemical gradient whose free en¬  ergy drives ATP synthesis.   • The mitochondrion contains soluble and membrane-bound en¬  zymes for oxidative metabolism.   • Reducing equivalents are imported from the cytosol via a shuttle  system. Specific transporters mediate the transmembrane move¬  ments of ADP, ATP, and P z .   • Electrons flow from redox centers with more negative reduction po¬  tentials to those with more positive reduction potentials. Inhibitors  have been used to reveal the sequence of electron carriers and the  points of entry of electrons into the electron-transport chain.   • Electron transport is mediated by one-electron carriers (Fe-S  clusters, cytochromes, and Cu ions) and two-electron carriers  (CoQ, FMN, FAD).   • Complex I transfers two electrons from NADFI to CoQ while  translocating four protons to the intermembrane space.   • Complex II transfers electrons from succinate through FAD to CoQ.   • Complex III transfers two electrons from CoQH 2 to two mole¬  cules of cytochrome c. The concomitant operation of the Q cycle  translocates four protons to the intermembrane space.    • Complex IV reduces 0 2 to 2 H 2 0 using four electrons donated  by four cytochrome c and four protons from the matrix. Two pro¬  tons are translocated to the intermembrane space for every two  electrons that reduce oxygen.   • As explained by the chemiosmotic theory, protons translocated  into the intermembrane space during electron transport through  Complexes I, III, and IV establish an electrochemical gradient  across the inner mitochondrial membrane.   • The influx of protons through the F 0 component of ATP synthase  (FjFo'ATPase) drives its F x component to synthesize ATP from  ADP + P z via the binding change mechanism, a process that is  mechanically driven by the F 0 -mediated rotation of Ffs y subunit  with respect to its catalytic a 3 (3 3 assembly.   • The P/O ratio, the number of ATPs synthesized per oxygen re¬  duced, need not be an integral number.   • Agents that discharge the proton gradient can uncouple oxidative  phosphorylation from electron transport.   • Oxidative phosphorylation is controlled by the ratio [NADH]/  [NAD + ] and by the ATP mass action ratio. Glycolysis and the  citric acid cycle are coordinately regulated according to the need  for oxidative phosphorylation.   • Aerobic metabolism is more efficient than anaerobic metabolism.  However, aerobic organisms must guard against damage caused by  reactive oxygen species.    energy coupling 603  chemiosmotic theory 604  protonmotive force (pmf) 606  FTo-ATPase 607  binding change    uncoupler 614  ATP mass action ratio 616  glucose-fatty acid cycle 617  reactive oxygen species  (ROS) 618  superoxide radical 618  antioxidant 619    1. Explain why a liver cell mitochondrion contains fewer cristae  than a mitochondrion from a heart muscle cell.   2. Certain genetic diseases caused by defective mitochondrial pro¬  teins are inherited only maternally, in contrast to most genetic  diseases, in which either parent can pass on the defect. Explain.   3. How many ATPs are synthesized for every cytoplasmic NADH that  participates in the glycerophosphate shuttle in insect flight muscle?   4. How many ATPs are synthesized for every cytoplasmic NADH  reducing equivalent that is transferred into the matrix via the  malate-aspartate shuttle?   3. Calculate AC 0 ' for the oxidation of free FADH 2 by 0 2 . What is  the maximum number of ATPs that can be synthesized, assum¬  ing standard conditions and 100% conservation of energy?   6. Show that the free energy change for the succinate dehydroge¬  nase reaction catalyzed by Complex II is insufficient to drive ATP  synthesis under standard conditions.   7. Some anaerobic prokaryotes use nitrate (N0 3 ) as the terminal  electron acceptor for energy metabolism. Assuming 100% effi¬  ciency, how much ATP could be synthesized by the oxidation of  NADH by nitrate?   8. Some anaerobic prokaryotes reduce elemental sulfur to H 2 S.  Assuming 100% efficiency, how much ATP could be synthesized  by the oxidation of acetate by S under standard conditions?   9. The 0 2 -consumption curve of a dilute, well-buffered suspension  of mitochondria containing an excess of ADP and P z takes the form   Sketch the curves obtained when (a) amytal is added at time t = 1  and (b) amytal is added at t = 1 and succinate is added at t = 2.   10. Sketch the 0 2 -consumption curves obtained for the mitochon¬  dria in Problem 9 when (a) CN is added at t = 1 and succi¬  nate is added at t = 2 and (b) oligomycin (which binds to F 0  and prevents ATP synthesis) is added at t = 1 and DNP is added  at t = 2.   11. Why is it possible for electrons to flow from a redox center with  a more positive %°' to one with a more negative within an  electron-transfer complex?   12. A family of proteins known as cupredoxins contain a single  redox-active Cu ion coordinated by a Cys, a Met, and two His  residues. The reduction potentials of cupredoxins range from  about 0.13 V to 0.68 V. What does this information reveal about  the role of the protein component of the cupredoxins?   13. Bombarding a suspension of mitochondria with high-frequency  sound waves (sonication) produces submitochondrial particles  derived from the inner mitochondrial membrane. These mem¬  branous vesicles seal inside out, so that the intermembrane space  of the mitochondrion becomes the lumen of the submitochon¬  drial particle. Diagram the process of electron transfer and oxida¬  tive phosphorylation in these particles.    14. For the experimental system described in Problem 13, and as¬  suming all the substrates for oxidative phosphorylation are pres¬  ent in excess, does ATP synthesis increase or decrease with an  increase in the pH of the fluid in which the submitochondrial  particles are suspended?   15. The difference in pH between the internal and external surfaces  of the inner mitochondrial membrane is 1.4 pH units (external  side acidic). If the membrane potential is 0.06 V (inside nega¬  tive), what is the free energy change on transporting 1 mol of  protons across the membrane from outside to inside at 25°C?   16. For the mitochondrial membrane described in Problem 15, how  many protons must be transported to provide enough free energy  for the synthesis of 1 mol of ATP (assuming standard biochem¬  ical conditions)?   17. Consider the mitochondrial ADP-ATP translocator and the  P z —H + symport protein. How do the activities of the two trans¬  porters affect the electrochemical gradient across the mitochon¬  drial membrane?   18. What thermodynamic force drives the transport of ADP and P z  into the mitochondrial matrix for ATP synthesis?   19. How do the P/O ratios for NADH differ in ATP synthases that  contain 10 and 15c subunits?   20. How many protons are required to synthesize one ATP by F^q-  ATPase containing (a) 10 or (b) 15c subunits?   21. Dicyclohexylcarbodiimide (DCCD) is a reagent that reacts with  Asp or Glu residues.    Explain why the reaction of DCCD with the c subunits of F^q-  ATPase blocks its ATP-synthesizing activity.   22. The antibiotic oligomycin B blocks proton transport through F 0 .  Explain why lactate concentrations build up in rats who have  been treated with oligomycin B.   24. What is the advantage of hormones activating a lipase to stimu¬  late nonshivering thermogenesis in brown fat rather than activat¬  ing UCP1 directly (see Box 18-4)?   25. Describe the changes in [NADH]/[NAD + ] and [ATP]/[ADP]  that occur during the switch from anaerobic to aerobic metabo¬  lism. How do these ratios influence the activity of glycolysis and  the citric acid cycle?   26. During cell signaling events that increase cytosolic calcium con¬  centrations, Ca 2+ enters the mitochondria via Ca 2+ channels in  the inner membrane, (a) Explain why it is necessary for these  channels to be highly specific for Ca 2+ . (b) How would increased  matrix [Ca 2+ ] affect oxidative phosphorylation?   27. In coastal marine environments, high concentrations of nutrients  from terrestrial runoff often lead to algal blooms. When the nu¬  trients are depleted, the algae die and sink and are degraded by  other microorganisms. The algal die-off may be followed by a  sharp drop in oxygen in the depths, which can kill fish and bot¬  tom-dwelling invertebrates. How do these “dead zones” form?   28. Chromium is most toxic and highly soluble in its oxidized Cr(VI)  state but is less toxic and less soluble in its more reduced Cr(III)  state. Efforts to detoxify Cr-contaminated groundwater have in¬  volved injecting chemical reducing agents underground. Another  approach is bioremediation, which involves injecting molasses or  cooking oil into the contaminated groundwater. Explain how these  substances would promote the reduction of Cr(VI) to Cr(III).   29. Activated neutrophils and macrophages (types of white blood  cells) fight invading bacteria by releasing superoxide. These cells  contain an NADPH oxidase that catalyzes the reaction   20 2 + NADPH-> 20 2* + NADP + + H +   Explain why flux through the glucose-6-phosphate dehydrogenase  reaction increases in these cells.   30. Mutations in SOD are associated with the neurodegenerative dis¬  ease amyotrophic lateral sclerosis (ALS). Explain why researchers  initially believed that loss of SOD activity would damage  neurons.    Case 27 Regulation of Sugar and Alcohol  Metabolism in Ssiccha.romyces cerevisiae  Focus concept: The regulation of carbohydrate metabolic pathways  in yeast serves as a good model for regulation of the same path¬  ways in multicellular organisms.   • The major pathways associated with carbohydrate metabolism,  including glycolysis, the citric acid cycle, oxidative phosphory¬  lation, pentose phosphate pathway, and gluconeogenesis   • The various fates of pyruvate via alcoholic fermentation and  aerobic respiration   Focus concept: Modification of Lys 43 in the mitochondrial ATPase,  once thought to be the structural basis of Batten disease, has been  found in bovine mitochondria, indicating that this modification is  completely normal.    Anaerobic organisms produce  ATP by pathways that do not require molecular oxygen. However,  all organisms carry out oxidation-reduction reactions in the normal  course of metabolism. What compounds other than 0 2 can serve  as electron acceptors in anaerobes? What is the fate of the result¬  ing reduced compounds?    Boyer, P.D., Catalytic site forms and controls in ATP synthase catalysis,  Biochim. Biophys. Acta 1458, 252-262 (2000). [A description of the  steps of ATP synthesis and hydrolysis, along with experimental evi¬  dence and alternative explanations, by the author of the binding  change mechanism.]   Brzezinski, P. and Johansson, A.-L., Variable proton-pumping stoichi¬  ometry in structural variants of cytochrome c oxidase, Biochim.  Biophys. Acta 1797, 710-723 (2010).   Crofts, A.R., The cytochrome hc x complex: Function in the context of  structure, Anna. Rev. Physiol. 66, 689-733 (2004).   Efremov, R.G., Baradaran, R., and Sazanov, L.A., The architecture of res¬  piratory complex I, Nature 465, 441-445 (2010).   Frey, T.G. and Mannella, C.A., The internal structure of mitochondria,  Trends Biochem. Sci. 23, 319-324 (2000).   Goodsell, D.S., Mitochondrion, Biochem. Mol. Biol. Educ. 38, 134-140  (2010). [An illustrated guide to the mitochondrion.]   Hinkle, P.C., P/O ratios of mitochondrial oxidative phosphorylation,  Biochim. Biophys. Acta 1706, 1-11 (2005).   Hosier, J.P., Ferguson-Miller, S., and Mills, D.A., Energy transduction:  Proton transfer through the respiratory complexes, Annu. Rev.  Biochem. 75, 165-187 (2006). [A review that focuses on cytochrome  c oxidase.]   Johnson, D.C., Dean, D.R., Smith, A.D., and Johnson, M.K., Structure,  function, and formation of biological iron-sulfur clusters, Annu. Rev.  Biochem. 74, 247-281 (2005)   Nicholls, D.G. and Ferguson, S.J., Bioenergetics 3, Academic Press (2002).  [An authoritative monograph devoted almost entirely to the mechanism  of oxidative phosphorylation and the techniques used to elucidate it.]    Noji, H. and Yoshida, M., The rotary machine in the cell, ATP synthase,  ]. Biol. Chem. 276, 1665-1668 (2001).   Pebay-Peyroula, E., Dahout-Gonzalez, C., Kahn, R., Trezeguet, V.,  Lauquin, G.J.-M., and Brandolin, G., Structure of mitochondrial  ADP/ATP carrier in complex with carboxyatractyloside, Nature 426,  39-44 (2003).   Schultz, B.E. and Chan, S.I., Structures and proton-pumping strategies  of mitochondrial respiratory enzymes, Annu. Rev. Biophys. Biomol.  Struct. 30, 23—65 (2001).   Solmaz, S.R.N. and Hunte, C., Structure of Complex III with bound cy¬  tochrome c in reduced state and definition of a minimal core inter¬  face for electron transfer, J. Biol. Chem. 283, 17542-17549 (2008)   Stock, D., Gibbons, C., Arechaga, I., Leslie, A.G.W., and Walker, J.E.,  The rotary mechanism of ATP synthase, Curr. Opin. Struct. Biol. 10 ,  692-679 (2000).   von Ballamoos, C., Wiedenmann, A., and Dimroth, P., Essentials for  ATP synthesis by FTq ATP synthases, Annu. Rev. Biochem. 78,  649-672 (2009)   Vonck, J., and Schafer, E., Supramolecular organization of protein com¬  plexes in the mitochondrial inner membrane, Biochim. Biophys. Acta  1793, 117-124 (2009).   Walker, J.E. (Ed.), The Mechanism of F 1 F 0 -ATPase, Biochim. Biophys.  Acta 1458, 221-514 (2002). [A series of authoritative reviews.]   Yoshikawa, S., Muramoto, K., and Shinzawa-Itoh, K., Proton-pumping  mechanism of cylochrome c oxidase, Annu. Rev. Biophys. 40,  205-223 (2011).   Yankovskaya, V., Horsefield, R., Tornroth, S., Luna-Chavez, C.,  Miyoshi, H., Leger, C., Byrne, B., Cecchini, G., and Iwata, S.,  Architecture of succinate dehydrogenase and reactive oxygen  species generation, Science 299, 700-704 (2003). [X-Ray struc¬  ture of E. coli Complex II.]    Although terrestrial plants are the most familiar examples of photosynthetic organisms,  marine cyanobacteria carry out just as much photosynthesis and generate, as a by¬  product, as much oxygen. In fact, the appearance of atmospheric oxygen that made  possible the evolution of large organisms such as animals is attributed to the  photosynthetic activity of cyanobacteria. [Michael Abbey/Photo Researchers, Inc.]   A The Light Reactions Take Place in the  Thylakoid Membrane  B Pigment Molecules Absorb Light   B Electron Transport in Photosynthetic Bacteria  Follows a Circular Path  C Two-Center Electron Transport Is a Linear  Pathway That Produces 0 2 and NADPH  D The Proton Gradient Drives ATP Synthesis  by Photophosphorylation   A The Calvin Cycle Fixes C0 2  B Calvin Cycle Products Are Converted to Starch,  Sucrose, and Cellulose  C The Calvin Cycle Is Controlled Indirectly  by Light   light and air was not validated until the eighteenth century. Evidence that  plants produce a vital substance—0 2 —was not obtained until 1771 when  Joseph Priestly noted that the air in a jar in which a candle had burnt out  could be “restored” by introducing a small plant into the jar. In the presence  of sunlight, plants and cyanobacteria consume C0 2 and H 2 0 and produce  0 2 and “fixed” carbon in the form of carbohydrate:    Photosynthesis, in which light energy drives the reduction of carbon, is  essentially the reverse of oxidative carbohydrate metabolism. Photosynthetically  produced carbohydrates therefore serve as an energy source for the organism  that produces them as well as for nonphotosynthetic organisms that directly or  indirectly consume photosynthetic organisms. It is estimated that photosynthe¬  sis annually fixes ~10 n tons of carbon, which represents the storage of over  10 18 kj of energy. The process by which light energy is converted to chemical  energy has its roots early in evolution, and its complexity is consistent with its  long history. Our discussion focuses first on purple photosynthetic bacteria, be¬  cause of the relative simplicity of their photosynthetic machinery, and then on  plants, whose chloroplasts are their site of photosynthesis.   Early in the twentieth century, it was mistakenly thought that light ab¬  sorbed by photosynthetic pigments directly reduced C0 2 , which then  combined with water to form carbohydrate. In fact, photosynthesis in plants  is a two-stage process in which light energy is harnessed to oxidize H 2 0:   Chapter 19 Photosynthesis The two stages of photosynthesis are traditionally referred to as the light   1. In the light reactions, specialized pigment molecules capture light en¬  ergy and are thereby oxidized. A series of electron-transfer reactions,  which culminate with the reduction of NADP + to NADPH, generate  a transmembrane proton gradient whose energy is tapped to synthesize  ATP from ADP + P z . The oxidized pigment molecules are reduced by  H 2 0, thereby generating 0 2 .   2. The dark reactions use NADPH and ATP to reduce C0 2 and incor¬  porate it into the three-carbon precursors of carbohydrates.   As we will see, both processes occur in the light and are therefore better described  as light-dependent and light-independent reactions. After describing the chloro-  plast and its contents, we consider the light reactions and dark reactions in turn.    • The chloroplast thylakoid membrane is the site of light absorption.   • Pigment molecules, some arranged in light-harvesting complexes, absorb visible light.    The site of photosynthesis in eukaryotes (algae and higher plants) is the chloro¬  plast. Cells contain 1 to 1000 chloroplasts, which vary considerably in size  and shape but are typically ~5-fJim-long ellipsoids. These organelles presum¬  ably evolved from photosynthetic bacteria.   Like mitochondria, which they resemble in many ways, chloroplasts have a  highly permeable outer membrane and a nearly impermeable inner membrane  separated by a narrow intermembrane space (Fig. 19-1). The inner membrane  encloses the stroma, a concentrated solution of enzymes, including those re¬  quired for carbohydrate synthesis. The stroma also contains the DNA, RNA,  and ribosomes involved in the synthesis of several chloroplast proteins.   The stroma, in turn, surrounds a third membranous compartment, the  thylakoid (Greek: thylakos, a sac or pouch). The thylakoid is probably a   FIG. 1 9-1 Chloroplast from corn, (a) An electron micrograph.  [Dr. Jeremy Burgess/Photo Researchers, Inc.] (b) Schematic diagram.   of stacks of disklike sacs named grana (singular, granum), which are intercon¬  nected by unstacked stromal lamellae. A chloroplast usually contains 10 to  100 grana. Thylakoid membranes arise from invaginations in the inner mem¬  brane of developing chloroplasts and therefore resemble mitochondrial cristae.   The thylakoid membrane contains protein complexes involved in harvesting  light energy, transporting electrons, and synthesizing ATP. In photosynthetic  bacteria, the machinery for the light reactions is located in the plasma  membrane, which often forms invaginations or multilamellar structures that  resemble grana.   The principal photoreceptor in photosynthesis is chlorophyll. This cyclic  tetrapyrrole, like the heme group of globins and cytochromes (Section 7-1A  and Box 18-1), is derived biosynthetically from protoporphyrin IX.   Chlorophyll molecules, however, differ from heme in several respects (Fig. 19-2).   In chlorophyll, the central metal ion is Mg 2+ rather than Fe(II) or Fe(III),  and a cyclopentanone ring, Ring V, is fused to pyrrole Ring III. The major   molecular formulas of chlorophylls a and b and  bacteriochlorophylls a and b are compared to  that of iron-protoporphyrin IX (heme). The  isoprenoid phytyl and geranylgeranyl tails  presumably increase the chlorophylls’ solubility in  nonpolar media.    FIG. 1 9-3 Absorption spectra of various  photosynthetic pigments. The chlorophylls  have two absorption bands, one in the red  (long wavelength) and one in the blue (short  wavelength). Phycoerythrin absorbs blue and  green light, whereas phycocyanin absorbs yellow  light. Together, these pigments absorb most of  the visible light in the solar spectrum. [After a  drawing by Govindjee, University of Illinois.]    chlorophyll forms in plants and cyanobacteria, chlorophyll a (Chi a) and  chlorophyll b (Chi &), and the major forms in photosynthetic bacteria, bac-  teriochlorophyll a (BChl a) and bacteriochlorophyll b (BChl b ), also dif¬  fer from heme and from each other in the degree of saturation of Rings II  and IV and in the substituents of Rings I, II, and IV.   The highly conjugated chlorophyll molecules, along with other photosyn¬  thetic pigments, strongly absorb visible light (the most intense form of the solar  radiation reaching the earths surface; Fig. 19-3). The relatively small chemical  differences among the various chlorophylls greatly affect their absorption spectra.   Light-Harvesting Complexes Contain Multiple Pigments. The primary reac¬  tions of photosynthesis, as is explained in Section 19-2B, take place at  photosynthetic reaction centers. Yet photosynthetic assemblies contain far more  chlorophyll molecules than are contained in reaction centers. This is because most  chlorophyll molecules do not participate directly in photochemical reactions but  function to gather light; that is, they act as light-harvesting antennas. These antenna  chlorophylls pass the energy of absorbed photons (units of light) from molecule  to molecule until it reaches a photosynthetic reaction center (Fig. 19-4).   Transfer of energy from the antenna system to a reaction center (RC) oc¬  curs in <10 10 s with an efficiency of >90%. This high efficiency depends  on the chlorophyll molecules having appropriate spacings and relative orien-    FIG. 1 9-4 Flow of energy through a  photosynthetic antenna complex. The energy  of an absorbed photon randomly migrates among  the molecules of the antenna complex (light  green circles ) until it reaches a reaction center  chlorophyll (dark green circles ) or, less frequently,  is reemitted (fluorescence).   FIG. 1 9-5 X-Ray structure of the light-harvesting complex  LH-2 from Rs. molischianum. (a) View perpendicular to the  photosynthetic membrane showing that the a subunits (blue; 56  residues) and the |3 subunits (magenta; 45 residues), as represented  by their C a backbones, are arranged in two concentric eightfold  symmetric rings. Twenty-four bacteriochlorophyll a (BChl a; green ) and  eight lycopene (a carotenoid; yellow) molecules are sandwiched  between the protein rings, (b) View from the plane of the membrane,  using the same colors as in Part a, in which the a-helical portions of  the proteins are represented by cylinders and the Mg 2+ ions are    represented by white spheres. Note that eight of the BChl a  molecules are bound near the top of the complex with their ring  systems nearly parallel to the plane of the membrane, whereas the  remaining sixteen BChl a molecules are bound near the bottom of the  complex with their ring systems approximately perpendicular to the  plane of the membrane. This arrangement, together with that of the  lycopene molecules, presumably optimizes the light-absorbing and  excitation-transmitting capability of the antenna system. [Courtesy of  Juergen Koepke and Hartmut Michel, Max-Planck Institut fur  Biochemie, Germany. PDBid 1LGH.] © See Interactive Exercise 23.    tations. Even in bright sunlight, an RC directly intercepts only ~1 photon  per second, a metabolically insignificant rate. Hence, a complex of antenna  pigments, or light-harvesting complex (LHC), is essential.   LHCs consist of arrays of membrane-bound hydrophobic proteins that  each contain numerous, often symmetrically arranged pigment molecules. For  example, LH-2 from the purple photosynthetic bacterium Rho do spirillum  molischianum is an integral membrane protein that consists of eight a sub¬  units and eight (3 subunits arranged in two eightfold symmetric concentric  rings between which are sandwiched 32 pigment molecules (Fig. 19-5). Other  LHCs vary widely in their structure and complement of light-harvesting pig¬  ments. The number and arrangement of pigment molecules in each LHC have  presumably been optimized for efficient energy transfer throughout the LHC.  Indeed, the ring of 16 BChl a molecules in LH-2 (lower part of Fig. 19-5^)  are so strongly coupled that they absorb radiation almost as a single unit.   Most LHCs contain other light-absorbing substances besides chlorophyll. These  accessory pigments “fill in” the absorption spectra of the antenna complexes,  covering the spectral regions where chlorophylls do not absorb strongly  (Fig. 19-3). For example, carotenoids, which are linear polyenes such as  (5-carotene,   Chapter 19 Photosynthesis are components of all green plants and many photosynthetic bacteria and are   therefore the most common accessory pigments. They are largely responsible  for the brilliant fall colors of deciduous trees as well as for the orange color  of carrots (after which they are named).   Water-dwelling photosynthetic organisms, which carry out at least half of  the photosynthesis on earth, additionally contain other types of accessory pig¬  ments. This is because light outside the wavelengths 450 to 550 nm (blue and  green light) is absorbed almost completely by passage through more than 10 m  of water. In red algae and cyanobacteria, Chi a therefore is replaced as an  antenna pigment by a set of linear tetrapyrroles, notably the red phyco-  erythrobilin and the blue phycocyanobilin (their spectra when linked to pro¬  tein are shown in Fig. 19-3).   • Summarize the events of the light reac¬  tions and light-independent reactions.   • Why do photosynthetic organisms contain  several types of pigment molecules?    • Absorbed light energy can be dissipated by internal conversion, fluorescence,  exciton transfer, or photooxidation.   • The special pair of the purple bacterial photosynthetic reaction center undergoes  photooxidation, and an electron-transport chain returns an electron to the special  pair.   • In plants and in cyanobacteria, two photosystems, cytochrome b 6 /, and mobile  electron carriers form an electron-transport chain described by the Z-scheme.   • Photosystem II reduces its photooxidized special pair with electrons derived from  water.   • Electrons traveling from photosystem II through the cytochrome b 6 f complex un¬  dergo a Q cycle that generates a transmembrane proton gradient.   • Electrons liberated by photooxidation of photosystem I reduce NADP + or return  to the cytochrome b 6 f complex, whose activity contributes to the proton gradient.    Photosynthesis is a process in which electrons from excited chlorophyll molecules are  passed through a series of acceptors that convert electronic energy to chemical en¬  ergy. We can thus ask two questions: (1) What is the mechanism of energy  transduction; and (2) How do photooxidized chlorophyll molecules regain  their lost electrons?   Electromagnetic radiation is propagated as discrete quanta (photons) whose  energy E is given by Planck’s law:   (2.998 X 10 8 m • s 1 in vacuum), v is the frequency of the radiation, and X  is its wavelength.   When a molecule absorbs a photon, one of its electrons is promoted from  its ground (lowest energy) state molecular orbital to one of higher energy.  However, a given molecule can absorb photons of only certain wavelengths be¬  cause , as is required by the law of conservation of energy, the energy difference be¬  tween the two states must exactly match the energy of the absorbed photon.   An electronically excited molecule can dissipate its excitation energy in  several ways (Fig. 19-6):   1. Internal conversion, a common mode of decay in which electronic  energy is converted to the kinetic energy of molecular motion, that is,  to heat. Many molecules relax in this manner to their ground states.  Chlorophyll molecules, however, usually relax only to their lowest ex¬  cited states. Consequently, the photosynthetically applicable excitation  energy of a chlorophyll molecule that has absorbed a photon in its  short-wavelength band, which corresponds to its second excited state,  is no different than if it had absorbed a photon in its less energetic  long-wavelength band.   2. Fluorescence, in which an electronically excited molecule decays to its  ground state by emitting a photon. A fluorescently emitted photon  generally has a longer wavelength (lower energy) than that initially ab¬  sorbed. Fluorescence accounts for the dissipation of only 3 to 6% of  the light energy absorbed by living plants.   3. Exciton transfer (also known as resonance energy transfer), in which  an excited molecule directly transfers its excitation energy to nearby  unexcited molecules with similar electronic properties. This process oc¬  curs through interactions between the molecular orbitals of the partic¬  ipating molecules. Light energy is funneled to RCs through exciton  transfer among antenna pigments. The energy (excitation) is trapped at  the RC chlorophylls because they have slightly lower excited state en¬  ergies than the antenna chlorophylls (Fig. 19-7). This energy differ¬  ence is lost as heat.   4. Photooxidation, in which a light-excited donor molecule is oxidized  by transferring an electron to an acceptor molecule, which is thereby  reduced. This process occurs because the transferred electron is less  tightly bound to the donor in its excited state than it is in the ground  state. In photosynthesis, excited chlorophyll (Chi*) is such a donor.  The energy of the absorbed photon is thereby chemically transferred to the  photosynthetic reaction system. Photooxidized chlorophyll, Chl + , a  cationic free radical, eventually returns to its reduced state by oxidiz¬  ing some other molecule.   FIG. 1 9-6 An energy diagram indicating  the electronic states of chlorophyll and their  most important modes of interconversion.   The wiggly arrows represent the absorption of  photons or their fluorescent emission. Excitation  energy may also be dissipated in radiationless  processes such as internal conversion (heat  production) and chemical reactions. Q See the  Animated Figures.    FIG. 1 9-7 Excitation energy trapping by  the photosynthetic reaction center. Light  energy that has been passed among pigment  molecules by exciton transfer is trapped by the  reaction center chlorophyll because its lowest  excited state has a lower energy than those of  the antenna pigment molecules.   FIG. 19-8 X-Ray structure of the  photosynthetic reaction center from Rb.  sphaeroides. (a) The H, M, and L subunits of  the protein, as viewed from within the plane of  the plasma membrane (cytoplasm below) are  magenta, cyan, and orange, respectively. The  prosthetic groups are drawn in stick form with C  green, N blue, and 0 red. The Fe(ll) atom is  represented by a yellow sphere. The 11 largely  vertical helices that form the central portion of  the protein constitute its transmembrane region.  (b) View from the extracellular side of the  membrane. Note how the transmembrane  portions of the M and L subunits are related by  a pseudotwofold axis passing through the Fe(ll)  ion and that the prosthetic groups are  sandwiched between the two subunits. [Based  on an X-ray structure by Marianne Schiffer,  Argonne National Laboratory. PDBid 2RCR.]    In purple photosynthetic bacteria, a membrane-bound bacteriochlorophyll  complex undergoes photooxidation when illuminated with red light. The ex¬  cited electron is transferred along a series of carriers until it returns to the orig¬  inal bacteriochlorophyll complex. During the electron-transfer process,  cytoplasmic protons are translocated across the plasma membrane. Dissipation  of the resulting proton gradient drives ATP synthesis. The relatively simple  RC of purple photosynthetic bacteria (PbRC) illustrates some general princi¬  ples of the photochemical events that occur in the more complicated photo¬  synthetic apparatus of plants and cyanobacteria (Section 19-2C).   The PbRC Is a Transmembrane Protein. The RCs from several species of  purple photosynthetic bacteria each contain three hydrophobic subunits  known as H, L, and M. The L and M subunits collectively bind four mol¬  ecules of bacteriochlorophyll, two molecules of bacteriopheophytin  (BPheo; bacteriochlorophyll in which the Mg 2+ ion is replaced by two pro¬  tons), one Fe(II) ion, and two molecules of the redox coenzyme ubiquinone  (Fig. 18-10^) or one molecule of ubiquinone and one of the related  menaquinone:   The PbRC from Rhodobacter (Rb.) sphaeroides is a transmembrane protein  with 11 membrane-spanning helices (Fig. 19-8). The disposition of prosthetic  groups in the homologous protein from Rhodopseudomonas (Rps.) viridis is  shown in Fig. 19-9. The most striking aspect of the PbRC is that the groups are  arranged with nearly perfect twofold symmetry. Two of the BChl molecules, the  so-called special pair, are closely associated; they are nearly parallel and have  an Mg—Mg distance of ~7 A. The special pair is named for the wavelength  (in nanometers) at which its absorbance maximally decreases on photooxida-    FIGURE 19-9 Disposition of prosthetic groups in the photosynthetic reaction  center of Rps. viridis. Note that their rings, but not their aliphatic side chains, are  arranged with close to twofold symmetry. The prosthetic groups bound by the L subunit are  on the right and those bound by the M subunit are on the left. Photons are absorbed by  the special pair of BChl b molecules (red). © See Kinemage Exercise 8-2.   tion [P870 or P960 depending on whether it consists of Bchl a or BChl b; Section 2 The Light Reactions   photosynthetic bacteria tend to inhabit murky stagnant ponds where visible  light (400-800 nm) does not penetrate; they require a near infrared-absorbing  species of chlorophyll]. Each member of the special pair, here P960, contacts  another BChl molecule that, in turn, associates with a BPheo molecule. The  menaquinone is close to the L subunit BPheo (Fig. 19-9, right ), whereas the  ubiquinone associates with the M subunit BPheo b (Fig. 19-9, left). The Fe(II)  is positioned between the menaquinone and the ubiquinone rings. Curiously,  the two symmetry-related sets of prosthetic groups are not functionally equiv¬  alent; electrons are almost exclusively transferred through the F subunit (the  right sides of Figs. 19-8 and 19-9). This effect is generally attributed to sub¬  tle structural and electronic differences between the F and M subunits.   1. The primary photochemical event of bacterial photosynthesis is the ab¬  sorption of a photon by the special pair (e.g., P960). The excited elec¬  tron is delocalized over both its BChl molecules.   2. P960*, the excited state of P960, has but a fleeting existence. Within  ~3 picoseconds (ps; 10 12 s), P960* transfers an electron to the BPheo  on the right in Fig. 19-9 to yield P960 + BPheo b (the intervening  BChl group probably plays a role in conveying electrons, although it  is not itself reduced; it is therefore known as the accessory BChl).   3. During the next 200 ps, the electron migrates to the menaquinone (or,  in many species, the second ubiquinone), designated Qa, to form the  anionic semiquinone radical Qa* . All these electron transfers, as dia¬  grammed in Fig. 19-10, are to progressively lower energy states, which  makes the process all but irreversible.   Rapid removal of the excited electron from the vicinity of P960 + is an essen¬  tial feature of the PbRC; this prevents return of the electron to P960 + , which    FIG. 19-10 The photosynthetic electron-transport system of purple photosyn¬  thetic bacteria. Electrons liberated by the absorption of photons by P960 pass through  BPheo b and Q A before reaching Q B , which exchanges with a pool of free ubiquinone.  Electrons from QH 2 pass through cytochrome be^ to cytochrome c 2 , which then reduces  P960 + . Note that two photons are required for the two-electron reduction of Q to QH 2  and that cytochrome c 2 carries one electron at a time back to the PbRC. The overall  process is essentially irreversible because electrons are transferred to progressively lower  energy states (more positive standard reduction potentials).    Estimate the change in reduction potential of P960 that occurs when it absorbs  light.   Chapter 19 Photosynthesis would lead to the wasteful internal conversion of its excitation energy to heat.   In fact, electron transfer in the PbRC is so efficient that its overall quantum yield  (ratio of molecules reacted to photons absorbed) is virtually 100%. No man-made  device has yet approached this level of efficiency.   Electrons Are Returned to the Photooxidized Special Pair via an Electron-  Transport Chain. Qf* , which occupies a hydrophobic pocket in the PbRC,  transfers its excited electron to the more solvent-exposed ubiquinone, Qb, to  form Q b * (the Fe ion positioned between Q A and Qb does not directly par¬  ticipate in these redox reactions). Q A never becomes fully reduced; it shuttles  between its oxidized and semiquinone forms.   When the PbRC is again excited, it transfers a second electron to Q B * to  form the fully reduced Qb . This anionic quinol takes up two protons from  the cytoplasmic side of the plasma membrane to form QbH 2 . Thus, is a  molecular transducer that converts two light-driven one-electron excitations to a  two-electron chemical reduction.   The electrons taken up by are eventually returned to P960 + via an   electron-transport chain (Fig. 19-10). The details of this process are species  dependent. The available redox carriers include a membrane-bound pool of  ubiquinone molecules, a cytochrome be j complex, and cytochrome c 2  (whose structure is drawn in Fig. 6-32 b). The electron-transport pathway  leads from Q B H 2 through the ubiquinone pool, with which Q B H 2 ex¬  changes, to cytochrome bc\, and then to cytochrome c 2 . The reduced cy¬  tochrome c 2 , which closely resembles mitochondrial cytochrome c, carries an  electron back to P960 + . The PbRC is thereby reduced and prepared to ab¬  sorb another photon.   Since electron transport in purple photosynthetic bacteria is a cyclic  process (Fig. 19-10), it results in no net oxidation—reduction. However, when  QH 2 transfers its electrons to cytochrome bc\ , its protons are translocated  across the plasma membrane. Cytochrome bc\ is a transmembrane protein  complex containing a [2Fe—2S] iron—sulfur protein, cytochrome c\, and a  cytochrome b that contains two hemes, b\\ and b L (H and L for /ugh and  /ow potential). Note that cytochrome bc\ is strikingly similar to the pro¬  ton-translocating Complex III of mitochondria (which is also called cy¬  tochrome bc\\ Section 18-2E). In fact, electron transfer from QH 2 (a  two-electron carrier) to the one-electron acceptor cytochrome c 2 occurs in a  two-stage Q cycle, exactly as occurs in mitochondrial electron transport  (Fig. 18-15). The net result is that for every two electrons transferred from  QH 2 to cytochrome c 2 , four protons enter the periplasmic space. Thus, pho¬  ton absorption by the PbRC generates a transmembrane H + gradient. Light-  dependent synthesis of ATP is driven by the dissipation of this gradient  (Section 19-2D).   C Two-Center Electron Transport Is a Linear Pathway That Produces  0 2 and NADPH   In plants and cyanobacteria, photosynthesis is a noncyclic process that uses the re¬  ducing power generated by the light-driven oxidation ofH 2 0 to produce NADPH.  This multistep process involves two photosynthetic reaction centers (RCs) that  each bear considerable resemblance to PbRCs. These RCs are photosystem II  (PSII), which oxidizes H 2 0, and photosystem I (PSI), which reduces  NADP + . Each photosystem is independently activated by light, with electrons  flowing from PSII to PSI. PSII and PSI therefore operate in electrical series to  couple H 2 0 oxidation with NADP + reduction.   Evidence for the existence of two photosystems came from observations  that in the presence of both red light (which activates only PSI) and yellow-  green light (which also activates PSII), plants produce 0 2 (i.e., oxidize H 2 0)    FIG. 1 9-1 1 A model of the thylakoid membrane. The   electron-transport system consists of three protein complexes: PSII,  the cytochrome b 6 f complex, and PSI, which are electrically  “connected” by the diffusion of the electron carriers plastoquinone (Q)  and plastocyanin (PC). Light-driven transport of electrons (blue arrows )  from H 2 0 to NADP + motivates the transport of protons (red arrows)  into the thylakoid lumen. Additional protons are split off from water  by the oxygen-evolving center (OEC), yielding 0 2 . The resulting    proton gradient powers the synthesis of ATP by the CFtCFq  proton-translocating ATP synthase. The membrane also contains  light-harvesting complexes (not shown) whose component pigments  transfer their excitations to PSII and PSI. Fd represents ferredoxin.  [After Ort, D.R. and Good, N.E., Trends Biochem. Sci. 13, 469 (1988).]   How many photon-absorption events are required to reduce  NADP + to NADPH? How many protons are translocated  across the thylakoid membrane during the process?    at a greater rate than the sum of the rates for each light acting alone. The her¬  bicide 3-(3,4-dichlorophenyl)-l,l-dimethylurea (DCMU)   blocks electron flow from PSII to PSI so that even with adequate illumina¬  tion (i.e., activation of both PSI and PSII), PSI is not supplied with electrons,  PSII cannot be reoxidized, and photosynthetic oxygen production ceases.   The pathway of electron transport in the chloroplast is more elaborate  than in photosynthetic bacteria. The components involved in electron transport  from H 2 0 to NADPH are largely organized into three thylakoid membrane-  bound particles (Fig. 19-11): PSII, a cytochrome b 6 f complex , and PSI  Electrons are transferred between the complexes via mobile electron carriers,  much as occurs in the respiratory electron-transport chain. The ubiquinone  analog plastoquinone (Q), via its reduction to plastoquinol (QH 2 ; at righi ),  links PSII to the cytochrome ^/complex, which, in turn, interacts with PSI  through the mobile peripheral membrane protein plastocyanin (PC).   FIG. 1 9-1 2 The Z-scheme of photosynthesis. Electrons  ejected from P680 in PSII by the absorption of photons are  replaced with electrons abstracted from H 2 0 by an Mn-containing  complex (the oxygen-evolving center; OEC), thereby forming 0 2 and  4 H + . Each ejected electron passes through a chain of electron  carriers to a pool of plastoquinone molecules (Q p00 i)- The resulting  plastoquinol, in turn, reduces the cytochrome b Q f complex (green box)  with the concomitant translocation of protons into the thylakoid lumen.  Cytochrome b 6 f then transfers the electrons to plastocyanin (PC). The  plastocyanin regenerates photooxidized P700 in PSI. The electron    ejected from P700, through the intermediacy of a chain of electron  carriers, reduces NADP + to NADPH in noncyclic electron transport.  Alternatively, the electron may return to the cytochrome b 6 f complex in  a cyclic process that translocates additional protons into the thylakoid  lumen. The reduction potentials increase downward so that electrons  flow spontaneously in this direction.    Electrons eventually reach ferredoxin-NADP + reductase (FNR), where they  are used to reduce NADP + . The oxidation of water and the passage of elec¬  trons through a Q cycle generate a transmembrane proton gradient, with the  greater [H + ] in the thylakoid lumen. The free energy of the proton gradient  is tapped by chloroplast ATP synthase.   The various prosthetic groups of the photosynthetic apparatus of plants  can be arranged in a diagram known as the Z-scheme (Fig. 19-12). As in  other electron-transport systems, electrons flow from low to high reduction  potentials. The zigzag nature of the Z-scheme reflects the two loci for photo¬  chemical events (one at PSII, one at PSI) that are required to drive electrons  from H 2 0 to NADP + .   PSII Resembles the PbRC. PSII from the thermophilic cyanobacterium  Thermosynechococcus elongatus consists of 19 subunits, 14 of which occupy the  photosynthetic membrane. These transmembrane subunits include the reac¬  tion center proteins D1 (PsbA) and D2 (PsbD), the chlorophyll-containing  inner-antenna subunits CP43 (PsbC) and CP47 (PsbB), and cytochrome  £ 559 . The X-ray structure of this PSII (Fig. 19-13), independently determined by  James Barber and So Iwata and by Wolfram Saenger, reveals that the ~340-kD   The PSII dimer is viewed from within the plane of the membrane. Its  transmembrane subunits include D1 (yellow), D2 (orange), CP47 (red),  CP43 (green), and cytochrome b 55 g (magenta). Other transmembrane  subunits are colored light blue and blue-gray. Its extrinsic proteins are  PsbO (dark blue), PsbU (purple), and PsbV (light green). The various  cofactors are drawn in stick form with the chlorophylls of the D1/D2  reaction center light green, those of the antenna complexes dark  green, pheophytins dark blue, hemes red, p-carotenes orange, Q A and  Q b purple, and the nonheme Fe represented by a red sphere. The  inferred position of the membrane is indicated by the light blue band.  (b) View of a PSII protomer perpendicular to the membrane from the  thylakoid lumen showing only the transmembrane portions of the  complex and colored as in Part a. A portion of the other protomer in  the PSII dimer is shown in muted colors with the dashed line  indicating the region of monomer-monomer interactions. The  pseudotwofold axis, which is perpendicular to the membrane and  passes through the nonheme Fe, relates the transmembrane helices  of the D1/D2 heterodimer, CP43 and CP47, and Psbl and PsbX as  emphasized by the black lines encircling the subunits. [Courtesy of  James Barber and So Iwata, Imperial Colleqe London, U.K. PDBid  1S5L]   protein is a symmetric dimer, whose protomeric units each contain 35 trans¬  membrane helices. Each protomer, which has pseudotwofold symmetry, binds  36 Chi as, 2 pheophytin a’s (Pheo dfs; Chi a with its Mg 2+ replaced by two  protons), 1 heme b, 1 heme c, 2 plastoquinones, 1 nonheme Fe, 7 all-trans  carotenoids presumed to be (3-carotene, 2 HC0 3 ions, and 1 M^CaC^ com¬  plex known as the oxygen-evolving center (OEC). In higher plants, the PSII  protomer contains ~25 subunits and forms an ~1000-kD transmembrane su¬  percomplex with several antenna proteins. The arrangement of the 5 trans¬  membrane helices in both D1 and D2 resembles that in the L and M subunits  of the PbRC (Fig. 19-8). Indeed, the two sets of subunits have similar  sequences, thereby indicating that they arose from a common ancestor.   The complex is viewed along the membrane  plane with the thylakoid lumen below. The  cofactors are colored as in Fig. 19-13 but with  Mg 2+ yellow, N blue, and 0 red. The phytyl tails  of the chlorophylls and pheophytins have been  removed for clarity. The side chains of Tyr z (D1  Tyr 161) and D1 His 190 are yellow, and those  of Tyr D (D2 Tyr 160) and D2 His 189 are  orange. The OEC is drawn in space-filling form  with Mn purple, Ca 2+ cyan, and 0 red. The  numbers indicate the center-to-center distances,  in angstroms, between the cofactors spanned by  the accompanying thin black lines. Compare this  figure to Fig. 19-9 (which is drawn upside down  relative to this figure). [Courtesy of James  Barber and So Iwata, Imperial College London,  U.K. PDBid 1S5L]    The cofactors of PSIIs RC (Fig. 19-14) are organized similarly to those of  the bacterial system (Fig. 19-9): They have essentially the same components  (with Chi a, Pheo a, and plastoquinone replacing BChl b, BPheo b, and  menaquinone, respectively) and are symmetrically organized along the complex’s  pseudo two fold axis. The two Chi a rings labeled P D1 and P D2 in Fig. 19-14  are positioned analogously to the BChl tfs of P960 s “special pair” and are there¬  fore presumed to form PSIIs primary electron donor, P680 (named after the  wavelength at which its absorbance maximally decreases on photooxidation).  The electron ejected from P680 follows a similar asymmetric course as that in  the PbRC even though the two systems operate over different ranges of reduc¬  tion potential (compare Figs. 19-10 and 19-12). As indicated in the central part  of Fig. 19-12, the electron is transferred to a molecule of Pheo a (Pheo D1 in  Fig. 19-14), probably via a Chi a molecule (Chl D1 ), and then to a bound plas¬  toquinone (Qa)- The electron is subsequently transferred to a second plasto¬  quinone molecule, Q B , which after it receives a second electron in a like manner  takes up two protons at the stromal (cytoplasmic in cyanobacteria) surface of  the thylakoid membrane. The resulting plastoquinol, Q B H 2 , then exchanges  with a membrane-bound pool of plastoquinone molecules. DCMU as well as  many other commonly used herbicides compete with plastoquinone for the Q B -  binding site on PSII, which explains how they inhibit photosynthesis.   0 2 Is Generated by a Five-Stage Water-Splitting Reaction. The electron  ejected by photooxidation of P680 is replaced by an electron derived from H 2 0  via the OEC. The OEC of PSII is also known as the water-splitting enzyme  because it breaks down two water molecules to 0 2 , four protons, and four elec¬  trons. Insight into this process was garnered by Pierre Joliet and Bessel Kok, who  analyzed the production of 0 2 by dark-adapted chloroplasts that were exposed  to a series of short flashes of light. 0 2 was evolved with a peculiar oscillatory  pattern (Fig. 19-15). There is virtually no 0 2 evolved by the first two flashes.  The third flash results in the maximum 0 2 yield. Thereafter, the amount of 0 2  produced peaks with every fourth flash until the oscillations damp out to a    FIG. 1 9-1 5 The 0 2 yield per flash in dark-adapted spinach chloroplasts. Note  that the yield peaks on the third flash and then on every fourth flash thereafter until the  curve eventually damps out to its average value. [After Forbush, B., Kok, B., and McGloin,  M.P., Photochem. Photobiol. 14, 309 (1971).]    FIG. 19-16 Schematic mechanism of 0 2 generation in chloroplasts. Four  electrons are stripped, one at a time, in light-driven reactions (S 0 —> S 4 ), from two bound  H 2 0 molecules. In the recovery step (S 4 —>• S 0 ), which is light independent, 0 2 is  released and two more H 2 0 molecules are bound. Three of the five steps release  protons into the thylakoid lumen.   The OEC is thought to cycle through five different states, 5 0 through S 4  (Fig. 19-16). 0 2 is released in the transition between S 4 and 5 0 . The obser¬  vation that 0 2 evolution peaks at the third rather than the fourth flash indi¬  cates that the OECs resting state is predominantly Si rather than S 0 . The  oscillations gradually damp out because a small fraction of the RCs fail to be  excited or become doubly excited by a given flash of light, so that the RCs  eventually lose synchrony. The complete reaction sequence releases a total of  four water-derived protons into the inner thylakoid space in a stepwise man¬  ner. These protons contribute to the transmembrane proton gradient.   Since the OEC abstracts electrons from H 2 0, its five states must have ex¬  traordinarily high reduction potentials (recall from Table 14-4 that the  0 2 /H 2 0 half-reaction has a standard reduction potential of 0.815 V). PSII  must also stabilize the highly reactive intermediates for extended periods (as  much as minutes) in close proximity to water.   The OEC, which is located at the lumenal surface of the D1 subunit  (Fig. 19-14), is a Mn 4 Ca 04 or Mn4Ca0 5 complex in which the O atoms  bridge neighboring Mn atoms. The structure of the OEC remains elusive due  to PSIEs relatively poorly resolved X-ray structures and the observation that  the OEC decomposes when illuminated with X-rays at the intensities used  in X-ray structure determinations. However, the use of X-ray spectroscopy  techniques of lower intensity that are sensitive to bond lengths have led to the  formulation of several related models for the OEC that are compatible with  the X-ray structure of PSII. One of these models is shown in Fig. 19-17.   The water-splitting reaction is driven by the excitation of the PSII RC. A  variety of evidence indicates that the Mn ions in the OECs various S states  (Fig. 19-16) cycle through specific combinations of Mn(II), Mn(III), Mn(IV),  and Mn(V) while abstracting protons and electrons from two H 2 0 molecules  to yield 0 2 , which is released into the thylakoid lumen. However, the mech¬  anism whereby this occurs, that is, the nature of the five S states, remains un¬  known due to the lack of structural information concerning the states.   The electrons abstracted from water by the OEC are relayed, one at a  time, to P680 + by an entity originally named Z (Fig. 19-12). Spectroscopic  measurements have identified Z as a transient neutral tyrosyl radical (TyrO*)  located on D1 Tyr 161 (known as Tyr z ), which is situated between the OEC  and P680 (Fig. 19-14). Recall that a tyrosyl radical has also been implicated  in the reduction of 0 2 to 2 H 2 0 by cytochrome c oxidase (Complex IV) in  the respiratory electron-transport chain (Section 18-2F).   Electron Transport through the Cytochrome b 6 f Complex Generates a  Proton Gradient. From the plastoquinone pool, electrons pass through the cy¬  tochrome ^/complex. This integral membrane assembly resembles cytochrome  bci, its purple bacterial counterpart (Section 19-2B), as well as Complex III of  the mitochondrial electron-transport chain (Section 18-2E). Electron flow  through the cytochrome ^/complex occurs through a Q cycle (Fig. 18-15).  Accordingly, two protons are translocated across the thylakoid membrane for  every electron transported. The four electrons abstracted from 2 H 2 0 by the  OEC therefore lead to the translocation of eight H + from the stroma to the  thylakoid lumen. Electron transport via the cytochrome b^f complex generates much  of the electrochemical proton gradient that drives the synthesis of ATP in chloroplasts.   Mn 4 Ca0 5 complex is shown in ball-and-stick  form with Mn ions magenta, the Ca 2+ ion cyan,  and 0 red. The bonds between the Ca and Mn  ions are drawn in gray to indicate that the  position of the Ca ion is relatively poorly defined.  Presumably, numerous protein side chains and  water molecules ligand the Ca and Mn ions.  Several related models are also compatible with  the structural data. [Based on a model by Vittal  Yachandra, Lawrence Berkeley National  Laboratory, Berkeley, CA.]    FIG. 19-18 X-Ray structure of the  cytochrome b 6 f complex from the thermophilic  cyanobacterium Mastigocladus laminosus. A   ribbon diagram of the dimeric complex is drawn  on the left with cytochrome b 6 blue, subunit IV  purple, cytochrome f red, the iron-sulfur protein  (ISP) yellow, and the other subunits green. The  inferred position of the lipid bilayer is indicated  by a yellow band. The paths of electron and  proton transfer through the complex and the  distances, in angstroms, between redox centers  are shown on the right. [Modified from a drawing  by William A. Cramer and Janet Smith, Purdue  University. PDBid 1UM3J    Compare this figure to Fig. 18-14 (which  is upside down relative to this figure)  and summarize the similarities and  differences.    The X-ray structure of cytochrome b 6 f{ Fig. 19-18) was independently de¬  termined by Janet Smith and William Cramer and by Jean-Luc Polpot and  Daniel Picot. Cytochrome b G f is a dimer of ~ 109-kD protomers, each con¬  taining four large subunits (18-32 kD) that have counterparts in cytochrome  bq: cytochrome b& a homolog of the N-terminal half of cytochrome b; sub¬  unit IV, a homolog of the C-terminal half of cytochrome b; cytochrome f(f  for feuille, French for leaf), a c-type cytochrome that is a functional analog of  cytochrome q, although the two are unrelated in structure or sequence; and a  Rieske iron-sulfur protein (ISP), which is also present in cytochrome bq. In ad¬  dition, cytochrome b 6 f has four small hydrophobic subunits that have no equiv¬  alents in cytochrome bq. Each protomer contains 13 transmembrane helices,  four in cytochrome b 6 , three in subunit IV, and one each in the remaining sub¬  units. Cytochrome b 6 f binds cofactors that are the equivalents of all of those  in cytochrome bq : heme f y a r-type heme bound by cytochrome f; a [2Fe-2S]  cluster bound by the ISP; hemes and b L ; a plastoquinone molecule that oc¬  cupies either the Qj site (the quinone-binding site at which fully reduced  quinone is regenerated during the Q cycle; Section 18-2E) or the Q 0 site. In  addition, cytochrome b 6 f binds several cofactors that have no counterparts in  cytochrome bq: a Chi a, a (3-carotene, and, unexpectedly, a novel heme named  heme x (alternatively, heme q), which is covalently linked to the protein via a  single thioether bond to Cys 35 of cytochrome b 6 , and whose only axial ligand  is a water molecule (compare with hemes a, b, and c in Box 18-1).    Plastocyanin Transports Electrons from Cytochrome b 6 f to PSI. Electron  transfer between cytochrome f, the terminal electron carrier of the cytochrome  ^/complex, and PSI is mediated by plastocyanin (PC), a peripheral membrane  protein located on the thylakoid lumenal surface (Fig. 19-11). The Cu-contain-  ing redox center of this mobile monomer cycles between its Cu(I) and Cu(II)  oxidation states. The X-ray structure of PC from poplar leaves shows that the  Cu atom is coordinated with distorted tetrahedral geometry by a Cys, a Met,  and two His residues (Fig. 19-19). Cu(II) complexes with four ligands normally  adopt a square planar coordination geometry, whereas those of Cu(I) are usu¬  ally tetrahedral. Evidently, the strain of Cu(II) s protein-imposed tetrahedral co¬  ordination in PC promotes its reduction to Cu(I). This hypothesis accounts for  PCs high standard reduction potential (0.370 V) compared to that of the normal    FIG. 19-19 X-Ray structure of plastocyanin (PC) from poplar leaves. This  99-residue monomeric protein, a member of the family of blue copper proteins, folds  into a |3 sandwich. Its Cu atom (orange sphere), which alternates between its Cu(l) and  Cu(ll) oxidation states, is tetrahedrally coordinated by the side chains of His 37, Cys 84, His  87, and Met 92, which are shown in stick form with their C, N, and S atoms green, blue,  and yellow. Seven conserved Asp and Glu residues that form a negatively charged patch on  the protein’s surface are red. [Based on an X-ray structure by Mitchell Guss and Hans  Freeman, University of Sydney, Australia. PDBid 1 PLCJ    Cu(II)/Cu(I) half-reaction (0.158 V) and illustrates how proteins can modulate  the reduction potentials of their redox centers. In the case of plastocyanin, this  facilitates electron transfer from the cytochrome ^/complex to PSI.   The structures of cytochrome f and PC suggest how the proteins associ¬  ate. Cytochrome fs Lys 187, a member of a conserved group of five positively  charged residues on the proteins surface, can be cross-linked to Asp 44 on  PC, which occupies a conserved negatively charged surface patch. Quite pos¬  sibly the two proteins associate through electrostatic interactions, much like  PCs functional analog cytochrome c interacts with its redox partners in the  mitochondrial electron-transport chain (Section 18-2E).   The PSI RC Resembles Both the PSII RC and the PbRC. Cyanobacterial PSIs  are trimers of protomers that each consist of at least 11 different protein subunits  coordinating >100 cofactors. The X-ray structure of PSI from T. elongatus (Fig.  19-20), determined by Saenger, reveals that each of its 356-kD protomers con¬  tains nine transmembrane subunits (PsaA, PsbB, PsaF, Psal-M, and PsaX) and  three stromal (cytoplasmic in cyanobacteria) subunits (PsaC-E), which collectively   FIG. 1 9-20 X-Ray structure of PSI from T. elongatus. (a) View  of the trimeric complex perpendicular to the membrane from its stromal  side. The stromal subunits have been removed for clarity. PSI’s threefold  axis of symmetry is represented by the small black triangle. Different  structural elements are shown for each of the three protomers (I, II, and  III). I shows the arrangement of transmembrane helices ( cylinders ),  which are differently colored for each subunit. The transmembrane  helices of both PsaA (blue) and PsaB (red) are named a through k  from their N- to C-termini. The six helices in extramembranous loop  regions are drawn as spirals. II shows the transmembrane helices as  cylinders with the stromal and lumenal loop regions drawn in ribbon   form. Ill shows the transmembrane helices as cylinders together with all  cofactors. The RC Chi a’s and quinones, drawn in stick form, are blue,  the Fe and S atoms of the [4Fe-4S] clusters are drawn as orange and  yellow spheres, the antenna system Chi a’s (whose side chains have  been removed for clarity) are yellow, the carotenoids are black, and the  bound lipids are light green, (b) One protomer as viewed parallel to the  membrane along the arrow in Part a with the stroma above. The  transmembrane subunits are colored as in Part a with the stromal  subunits PsaC, PsaD, and PsaE pink, cyan, and light green. The vertical  line and triangle mark the trimer’s threefold axis of symmetry. [Courtesy  of Wolfram Saenger, Freie Universitat Berlin, Germany. PDBid 1JB0.]    FIG. 1 9-21 The cofactors of the PSI RC and PsaC. The view is  parallel to the membrane plane with the stroma above. The Chi a and  phylloquinone molecules are arranged in two pseudosymmetric branches.   The Chi a’s are labeled A or B to indicate that their Mg 2+ ions are liganded  by the side chains of PsaA or PsaB, respectively. The phylloquinones are  named Q K -A and Q K -B. PsaC is shown in ribbon form with those portions  resembling segments in bacterial 2[4Fe-4S] ferredoxins pink and with  insertions and extensions green. The three [4Fe-4S] clusters are shown in  ball-and-stick form and are labeled according to their spectroscopic  identities F x , F A , and F B . The center-to-center distances between cofactors  (vertical black lines) are given in angstroms. [Courtesy of Wolfram Saenger,  Freie Universitat Berlin, Germany. PDBid 1JB0.]    bind 127 cofactors that comprise 30% of PSIs mass. The cofactors forming the  PSI RC are all bound by the homologous subunits PsaA (755 residues) and PsaB  (740 residues), whose 11 transmembrane helices each are arranged in a manner  resembling those in the L and M subunits of the PbRC (Fig. 19-8) and the D1  and D2 subunits of PSII (Fig. 19-13), thus supporting the notion that all RCs  arose from a common ancestor. PsaA and PsaB, together with other transmem¬  brane subunits, also bind the cofactors of the core antenna system (see below).   Figure 19-21 indicates that PSIs RC consists of 6 Chi as and two mole¬  cules of phylloquinone (at left), which has the same phytyl side chain as do  chlorophylls (Fig. 19-2), all arranged in two pseudosymmetrically related  branches, followed by three [4Fe—4S] clusters. The primary electron donor of  this system, P700, consists of a pair of parallel Chi as, A1 and Bl, whose  Mg 2+ ions are separated by 6.3 A, and thus resembles the special pair in the  PbRC. A1 is followed in the left branch of Fig. 19-21 by two more Chi a rings,  B2 and A3, and Bl is followed by A2 and B3 in the right branch. One or both  of the third pair of Chi a molecules, A3 and B3, probably form the spectro¬  scopically identified primary electron acceptor Aq (right side of Fig. 19-12).  The Mg 2+ ions of A3 and B3 are each axially liganded by the S atoms of a  Met residue rather than by His side chains (thereby forming the only known  biological examples of Mg 2+ —S coordination). Electrons are passed from A3  and B3 to the phylloquinones, Q K -A and Q K -B, which almost certainly corre¬  spond to the spectroscopically identified electron acceptor A l . Spectroscopic in¬  vestigations indicate that, in contrast to the case for the PbRC, electrons pass  through both branches of the PSI RC, although at different rates. Indeed, the  PSI RC is most closely related to the RC of green sulfur bacteria (a second  class of photosynthetic bacteria), which is a true homodimer.   Up until this point, PSIs RC resembles those of PSII and purple photosyn¬  thetic bacteria. However, rather than the reduced forms of either Q K -A or Q K -  B dissociating from PSI, both of the quinones directly pass their photoexcited  electron to a chain of three spectroscopically identified [4Fe-4S] clusters desig¬  nated F x , F a , and F B (right side of Fig. 19-12). F x , which lies on the pseudo¬  twofold axis relating PsaA and PsaB, is coordinated by two Cys residues from each  of the subunits. F A and F B are bound to the stromal subunit PsaC, which struc¬  turally resembles bacterial ferredoxins that contain two [4Fe—4S] clusters (Fig.  19-22). The observation that both branches of PSIs electron-transfer pathways  are active, in contrast to only one active branch in PSII and the PbRC, is ration¬  alized by the observation that the two quinones at the ends of each branch are  functionally equivalent in PSI but functionally different in PSII and the PbRC.    in the core antenna subunits CP43 and CP47 of PSII (Fig. 19-13). Indeed, the  N-terminal domains of PsaA and PsaB are similar in sequence to those of CP43  and CP47 and fold into similar structures containing six transmembrane helices  each. The carotenoids, which are mostly (3-carotenes, are deeply buried in the  membrane, where they are in van der Waals contact with Chi a rings. This  permits efficient energy transfer from photoexcited carotenoids to Chi a.   PSIs from higher plants are monomers rather than trimers as are cyanobac-  terial PSIs. Nevertheless, the X-ray structure of PSI from peas reveals that the  positions and orientations of the chlorophylls in both species of PSIs are nearly  identical, a remarkable finding considering the > 1 billion years since chloro-  plasts diverged from their cyanobacterial ancestors. However, pea PSI has  four antenna proteins not present in cyanobacterial PSI that are arranged in  a crescent-shaped transmembrane belt around one side of its RC and which  collectively bind 56 chlorophyll molecules.   PSI-Activated Electrons May Reduce NADP + or Motivate Proton Gradient  Formation. Electrons ejected from F B in PSI may follow either of two alterna¬  tive pathways (Fig. 19-12):   1. Most electrons follow a noncyclic pathway by reducing an ~ 100-  residue, [2Fe-2S]-containing, soluble protein called ferredoxin (Fd;  Fig. 19-22) that is located in the stroma. Reduced Fd, in turn, reduces  NADP + in a reaction mediated by the ~310-residue, monomeric,  FAD-containing ferredoxin-NADP + reductase (FNR, Fig. 19-23) to  yield the final product of the chloroplast light reactions, NADPH. Two  reduced Fd molecules successively deliver one electron each to the FAD  of FNR, which thereby sequentially assumes the neutral semiquinone  and fully reduced states before transferring the two electrons and a pro¬  ton to the NADP + via what is formally a hydride ion transfer.    FIG. 1 9-22 X-Ray structure of ferredoxin  from Peptococcus aerogenes. This monomeric  54-residue protein contains two [4Fe-4S]  clusters. The C p atoms of the four Cys residues  liganding each cluster are green, the Fe atoms  are orange, and the S atoms are yellow. [Based  on an X-ray structure by Elinor Adman, Larry  Sieker, and Lyle Jensen, University of  Washington. PDBid 1 FDX.] © See Interactive  Exercise 25.   FIG. 1 9-23 X-Ray structure of pea ferredoxin-NADP + reductase  (FNR) in complex with FAD and NADP + . This 308-residue protein has  two domains: The N-terminal domain (gold), which forms the FAD-binding site,  folds into an antiparallel |3 barrel, whereas the C-terminal domain (magenta), which  provides the NADP + -binding site, forms a dinucleotide-binding fold (Section 6-2C).   The FAD and NADP + are shown in stick form with NADP + C green, FAD C cyan,   N blue, 0 red, and P yellow. The flavin and nicotinamide rings are in opposition with  C4 of the nicotinamide ring and C5 of the flavin ring 3.0 A apart, an arrangement that  is consistent with direct hydride transfer as also occurs in dihydrolipoyl dehydrogenase  (Fig. 17-8). [Based on an X-ray structure by Andrew Karplus, Cornell University. PDBid  1QFYJ © See Interactive Exercise 26.    2. Some electrons are returned from PSI, via cytochrome h 6 , to the plas-  toquinone pool, thereby traversing a cyclic pathway that translocates  protons across the thylakoid membrane. A mechanism that has been  proposed for this process is that Fd transfers an electron to heme x of  cytochrome b 6 (Fig. 19-18) rather than to FNR. Since heme x con¬  tacts heme A L at the periphery of cytochrome h 6 f s Qj site, an electron  injected into heme x would be expected to reduce plastoquinone via a  Q cycle-like mechanism (Fig. 18-15). Note that the cyclic pathway is  independent of the action of PSII and hence does not result in the  evolution of 0 2 . This accounts for the observation that chloroplasts  absorb more than eight photons per 0 2 molecule evolved.    The cyclic electron flow presumably functions to increase the amount of  ATP produced relative to that of NADPH and thus permits the cell to adjust  the relative amounts of the two substances produced according to its  needs. However, the mechanism that apportions electrons between  the cyclic and noncyclic pathways is unknown. Fine-tuning of the  light reactions also depends on the segregation of PSI and PSII  in distinct portions of the thylakoid membrane (Box 19-1).    Electron microscopy has revealed that the protein complexes of the  thylakoid membrane have characteristic distributions (see figure).   1. PSI occurs mainly in the unstacked stromal lamellae, in contact  with the stroma, where it has access to NADP + .   2. PSII is located almost exclusively between the closely stacked  grana, out of direct contact with the stroma.   3. Cytochrome b 6 f is uniformly distributed throughout the  membrane.   The high mobilities of plastoquinone and plastocyanin, the electron  carriers that shuttle electrons between these particles, permit  photosynthesis to proceed at a reasonable rate.   What function is served by the segregation of PSI and PSII? If  the two photosystems were in close proximity, the higher excitation  energy of PSII (P680 versus P700) would cause it to pass a large  fraction of its absorbed photons to PSI via exciton transfer; that is,  PSII would act as a light-harvesting antenna for PSI. The separation  of the particles by around 100 A eliminates this difficulty.   The physical separation of PSI and PSII also permits the  chloroplast to respond to changes in illumination. The relative    amounts of light absorbed by the two photosystems vary with how  the light-harvesting complexes are distributed between the  stacked and unstacked portions of the thylakoid membrane. Under  high illumination (normally direct sunlight, which contains a high  proportion of short-wavelength blue light), PSII absorbs more light  than PSI. PSI is then unable to take up electrons as fast as PSII  can supply them, so the plastoquinone is predominantly in its  reduced state. The reduced plastoquinone activates a protein  kinase to phosphorylate specific Thr residues of the LHCs, which,  in response, migrate to the unstacked regions of the thylakoid  membrane, where they associate with PSI. A greater fraction of  the incident light is thereby tunneled to PSI.   Under low illumination (normally shady light, which contains a  high proportion of long-wavelength red light), PSI takes up  electrons faster than PSII can provide them so that plastoquinone  predominantly assumes its oxidized form. The LHCs are  consequently dephosphorylated and migrate to the stacked  portions of the thylakoid membrane, where they associate with  PSII. The chloroplast therefore maintains the balance between its  two photosystems by a light-activated feedback mechanism.   Chloroplasts generate ATP in much the same way as mitochondria ,, that is , by cou¬  pling the dissipation of a proton gradient to the enzymatic synthesis of ATP  (Section 18-3). This light-dependent process is known as photophosphoryla-  tion. Like oxidative phosphorylation, it requires an intact thylakoid membrane  and can be uncoupled from light-driven electron transport by compounds such  as 2,4-dinitrophenol (Fig. 18-29).   Electron micrographs of thylakoid membrane stromal surfaces and bacterial  plasma membrane inner surfaces reveal lollipop-shaped structures (Fig. 19-24).  These closely resemble the F x units of the proton-translocating ATP synthase  in mitochondria (Fig. 18-21). In fact, the chloroplast ATP synthase, which is   FIG. 1 9-24 Electron micrograph of thylakoids. The CF^ “lollipops” of their ATP  synthases project from their stromal surfaces. [Courtesy of Peter Hinkle, Cornell University.]    also known as the CFjCFq complex (C for chloroplast), is remarkably similar  to the mitochondrial F 1 F 0 complex. For example,   1. Both the F 0 and the CF 0 units are hydrophobic transmembrane pro¬  teins that contain a proton-translocating channel.   2. Both the F x and the CF X units are hydrophilic peripheral membrane  proteins of subunit composition a 3 (3 3 y8£, of which (3 is a reversible  ATPase.   3. Both ATP synthases are inhibited by oligomycin and by dicyclohexyl-  carbodiimide (DCCD).   Clearly, proton-translocating ATP synthase must have evolved very early in the  history of cellular life. Note, however, that whereas chloroplast ATP synthase  translocates protons out of the thylakoid space into the stroma (Fig. 19-11),  mitochondrial ATP synthase conducts them from the intermembrane space  into the matrix space (Fig. 18-20). This is because the stroma is topologically  analogous to the mitochondrial matrix.   Photosynthesis with Noncyclic Electron Transport Produces Around One  ATP per Absorbed Photon. At saturating light intensities, chloroplasts gen¬  erate proton gradients of ~3.5 pH units across their thylakoid membranes as  a result of two processes:   1. The evolution of a molecule of 0 2 from two H 2 0 molecules releases  four protons into the thylakoid lumen.   2. The transport of the liberated four electrons through the cytochrome  ^/complex occurs with the translocation of eight protons from the  stroma to the thylakoid lumen.   Altogether, ~ 12 protons enter the lumen per molecule of 0 2 produced by non¬  cyclic electron transport.   The thylakoid membrane, in contrast to the inner mitochondrial membrane,  is permeable to ions such as Mg 2+ and Cl . Translocation of protons and elec¬  trons across the thylakoid membrane is consequently accompanied by the pas¬  sage of these ions so as to maintain electrical neutrality (Mg 2+ out and Cl in).  This all but eliminates the membrane potential. The electrochemical gradient in  chloroplasts is therefore almost entirely a result of the pH (concentration) gradient.   Chloroplast ATP synthase, according to most estimates, produces one ATP  for every three protons it transports from the thylakoid lumen to the stroma.  Noncyclic electron transport in chloroplasts therefore results in the produc¬  tion of ~ 12/3 — 4 molecules of ATP per molecule of 0 2 evolved (cyclic elec¬  tron transport generates more ATP because more protons are translocated to  the thylakoid lumen via the Q cycle mediated by cytochrome b&f).   Noncyclic electron transport, of course, also yields NADPH (2 NADPH  for every 4 electrons liberated from 2 H 2 0 by the OEC). Each NADPH has  the free energy to produce 2.5 ATP (Section 18-3C; although NADPH is not  used to drive ATP synthesis), for a total of 5 more ATP equivalents per 0 2  produced. Consequently, a total of 9 ATP equivalents are generated per 0 2  produced. A minimum of two photons is required for each electron travers¬  ing the system from H 2 0 to NADPH, that is, eight photons per 0 2 pro¬  duced. This is confirmed by experimental measurements which indicate that  plants and algae require 8 to 10 photons of visible light to produce one mol¬  ecule of 0 2 . Thus, the overall efficiency of the light reactions is 9 ATP/8-10  photons, or approximately one ATP per absorbed photon.   • How do molecules dissipate absorbed  light energy? Which mechanism is most  important for photosynthesis?   • Describe the events that occur after the  special pair of PbRC absorbs a photon.   • Explain why electron transport in purple  photosynthetic bacteria follows a circular  path.   • Describe the effect of photooxidation on  the redox reactions summarized in the  Z-scheme.   • What is the importance of the water¬  splitting reaction for photosynthesis?   • What does the Q cycle accomplish in  bacterial and plant photosynthetic electron  transport?   • What chloroplast protein is the functional  counterpart of mitochondrial cytochrome c?   • What are the implications of cyclic and  noncyclic electron transfer in PSI?   • Explain how the energy of one photon is  transformed to the energy of one ATP.   • What is the relationship between the  number of photons absorbed and the  amount of 0 2 produced?    • The Calvin cycle carboxylates a pentose, converts the products to glyceraldehyde-  3-phosphate, and regenerates the pentose, using the ATP and NADPH produced  by the light reactions.   • The products of the Calvin cycle are converted to carbohydrates (glucose  polymers).   • The Calvin cycle enzymes are more reactive in the light.   • Plants undergo photorespiration, which consumes 0 2 and generates C0 2 .    In the previous section we saw how plants harness light energy to generate  ATP and NADPH. In this section we discuss how these products are used to  synthesize carbohydrates and other substances from C0 2 .   FIG. 1 9-25 Overview of the dark  reactions. The products of the light reactions,  ATP and NADPH, are consumed in converting  C0 2 into carbohydrates in a process called the  Calvin cycle.    The metabolic pathway by which plants incorporate C0 2 into carbohy¬  drates was elucidated between 1946 and 1953 by Melvin Calvin, James  Bassham, and Andrew Benson. They did so by tracing the metabolic fate  of the radioactive label from l4 C0 2 in cultures of algal cells. Some of  Calvins earliest experiments indicated that algae exposed to l4 C0 2 for a  minute or more synthesize a complex mixture of labeled metabolites, in¬  cluding sugars and amino acids. Analysis of the algae within 5 s of their  exposure to l4 C0 2 , however, showed that the first stable radioactive com¬  pound formed is 3-phosphoglycerate (3PG) y which is initially labeled only  in its carboxyl group. This result immediately suggested that the 3PG was  formed by the carboxylation of a C 2 compound. Yet no such precursor was  found. The actual carboxylation reaction involves a pentose derived from  ribulose-5-phosphate (Ru5P; at left). The resulting product splits into two  C 3 compounds, both of which turned out to be 3PG. ATP and NADPH, the  products of the light reactions, are required to convert 3PG to glyceraldehyde-  3-phosphate (GAP), which is used to synthesize carbohydrates as well as  re-form Ru5P (Fig* 19-25). The entire pathway, which involves the carboxy¬  lation of a pentose, the formation of carbohydrate products, and the regen¬  eration of the pentose, is known as the Calvin cycle or the reductive pentose  phosphate cycle.   During the search for the carboxylation substrate, several other photosyn¬  thetic intermediates had been identified and their labeling patterns elucidated.   For example, the hexose fructose-1,6-bisphosphate (FBP) is ini¬  tially labeled only at its C3 and C4 positions but later becomes  labeled to a lesser degree at its other atoms. A consideration of  the flow of labeled carbon through the various tetrose, pentose,  hexose, and heptose phosphates led, in what is a milestone of metabolic bio¬  chemistry, to the deduction of the Calvin cycle as is diagrammed in Fig. 19-26.  The existence of many of its postulated reactions was eventually confirmed by in  vitro studies using purified enzymes.    FIG. 1 9-26 The Calvin cycle. ( Opposite) The number of lines in an arrow indicates  the number of molecules reacting in that step for a single turn of the cycle that converts  three C0 2 molecules to one GAP molecule. For the sake of clarity, the sugars are all  shown in their linear forms, although the hexoses and heptoses predominantly exist in their  cyclic forms. The 14 C-labeling patterns generated in one turn of the cycle through the use  of 14 C0 2 are indicated in red. Note that two of the product Ru5Ps are labeled only at C3,  whereas the third Ru5P is equally labeled at Cl, C2, and C3. Q See the Animated  Figures.    The Calvin Cycle Generates GAP from C0 2 via a Two-Stage Process. The   Stage 1 . The production phase (top line of Fig. 19-26), in which three  molecules of Ru5P react with three molecules of C0 2 to yield  six molecules of glyceraldehyde-3-phosphate (GAP) at the ex¬  pense of nine ATP and six NADPH molecules. The cyclic na¬  ture of the entire pathway makes this process equivalent to the  synthesis of one GAP from three C0 2 molecules. At this point, GAP  can be bled off from the cycle for use in biosynthesis.   Stage 2. The recovery phase (bottom lines of Fig. 19-26), in which the  carbon atoms of the remaining five GAPs are shuffled in a re¬  markable series of reactions, similar to those of the pentose phos¬  phate pathway (Section 15-6), to re-form the three Ru5Ps with  which the cycle began. This stage can be conceptually decom¬  posed into four sets of reactions (with the numbers keyed to the  corresponding reactions in Fig. 19-26):   8 . C 3 + C (5 —> C 5 + C 4   11 . C 3 + C 7 —> C 3 + C 3   Note that this stage of the Calvin cycle occurs without further  input of free energy (ATP) or reducing equivalents (NADPH).   The first reaction of the Calvin cycle is the phosphorylation of Ru5P by  phosphoribulokinase to form ribulose-l,5-bisphosphate (RuBP). Following  the carboxylation of RuBP (Reaction 2 ; discussed below), the resulting 3PG  is converted first to 1 , 3 -bisphosphoglycerate (BPG) and then to GAP. The lat¬  ter sequence is the reverse of two consecutive glycolytic reactions (Section 15-  2G and 15-2F) except that the Calvin cycle reaction uses NADPH rather than  NADH.   The second stage of the Calvin cycle begins with the reverse of a famil¬  iar glycolytic reaction, the isomerization of GAP to dihydroxyacetone phos¬  phate (DHAP) by triose phosphate isomerase (Section 15-2E). Following  this, DHAP is directed along two analogous paths: Reactions 6 to 8 or  Reactions 9 to 11. Reactions 6 and 9 are aldolase-catalyzed aldol condensa¬  tions in which DHAP is linked to an aldehyde. Reaction 6 is also the re¬  verse of a glycolytic reaction (Section 15-2D). Reactions 7 and 10 are  phosphate hydrolysis reactions that are catalyzed, respectively, by fructose  bisphosphatase (FBPase; Section 15-4B) and sedoheptulose bisphosphatase  (SBPase). The remaining Calvin cycle reactions are catalyzed by enzymes  that also participate in the pentose phosphate pathway. In Reactions 8 and  11 , both catalyzed by transketolase, a C 2 keto unit (shaded green in Fig. 19-26)  is transferred from a ketose to GAP to form xylulose- 5 -phosphate (Xu5P),  leaving the aldoses erythrose-4-phosphate (E4P) in Reaction 8 and ribose-  5-phosphate (R5P) in Reaction 11 . The E4P produced by Reaction 8 feeds  into Reaction 9. The Xu5Ps produced by Reactions 8 and 11 are converted  to Ru5P by phosphopentose epimerase in Reaction 12 . The R5P from  Reaction 11 is also converted to Ru5P by ribose phosphate isomerase in  Reaction 13, thereby completing a turn of the Calvin cycle. Only 3 of the  11 Calvin cycle enzymes—phosphoribulokinase, the carboxylation enzyme  ribulose bisphosphate carboxylase, and SBPase—have no equivalents in  animal tissues.   FIG. 1 9-27 X-Ray structure of tobacco RuBP carboxylase in complex with the  transition state inhibitor 2-carboxyarabinitol-1,5-bisphosphate. The L 8 S 8 protein has  D 4 symmetry (the symmetry of a square prism; Fig. 6-346). (a) Surface diagram viewed  along the protein’s fourfold axis and (b) along its twofold axis. Parts a and b are related by  a 90° rotation about the horizontal axis. The elongated L subunits can be considered to  associate as two interdigitated tetramers, with that extending from the top in Part b cyan  and that extending from the bottom green. The S subunits, which form square tetramers  that cap the top and bottom of the complex, are alternately yellow and orange. The  2-carboxyarabinitol-1,5-bisphosphate is drawn in stick form with C green, 0 red, and P  orange, (c) Ribbon diagram of an L subunit oriented as is the central green subunit in Part  b and colored in rainbow order from its N-terminus (blue) to its C-terminus (red). The  2-carboxyarabinitol-1,5-bisphosphate is bound in the substrate-binding site at the mouth of the  enzyme’s a/|3 barrel. [Based on an X-ray structure by David Eisenberg, UCLA. PDBid 1RLC.]    RuBP Carboxylase Catalyzes C0 2 Fixation. The enzyme that catalyzes  CO 2 fixation, ribulose bisphosphate carboxylase (RuBP carboxylase), is ar¬  guably the world s most important enzyme since nearly all life on earth ul¬  timately depends on its action. This protein, presumably as a consequence  of its low catalytic efficiency (£ cat ~ 3 s l ), accounts for up to 50% of leaf  proteins and is therefore the most abundant protein in the biosphere. RuBP  carboxylase from higher plants and most photosynthetic microorganisms  consists of eight large (L) subunits (477 residues in tobacco leaves) encoded  by chloroplast DNA, and eight small (S) subunits (123 residues) specified  by a nuclear gene (the RuBP carboxylase from certain photosynthetic bac¬  teria is an L 2 dimer whose L subunit has 28% sequence identity with and  is structurally similar to that of the L 8 S 8 enzyme). X-Ray studies by Carl-  Ivar Branden and by David Eisenberg demonstrated that the L 8 S 8 enzyme  has the symmetry of a square prism (Fig. 19-27 a,b). The L subunit is made  of a (3 sheet domain and an a/(3 barrel domain that contains the enzymes  catalytic site (Fig. 19-27c). The function of the S subunit is unknown; at¬  tempts to show that is has a regulatory role, in analogy with other enzymes,  have been unsuccessful.   The accepted mechanism of RuBP carboxylase, which was largely formu¬  lated by Calvin, is indicated in Fig. 19-28. Abstraction of the C3 proton of  RuBP, the reactions rate-determining step, generates an enediolate that nucle-  ophilically attacks C0 2 . The resulting (3-keto acid is rapidly attacked at its C3  position by H 2 0 to yield an adduct that splits, by a reaction similar to aldol  cleavage, to yield the two product 3PG molecules. The driving force for the  overall reaction ., which is highly exergonic (AG of = ~35.1 kj • mol 2 ), is pro¬  vided by the cleavage of the (3-keto acid intermediate to yield an additional reso¬  nance-stabilized carboxylate group.    FIG. 19-28 Mechanism of the RuBP  carboxylase reaction. The reaction proceeds via  an enediolate intermediate that nucleophilically  attacks C0 2 to form a p-keto acid. The  intermediate reacts with water to yield two  molecules of 3PG. © See the Animated  Figures.    RuBP carboxylase activity requires Mg 2+ , which probably stabilizes devel¬  oping negative charges during catalysis. The Mg 2+ is, in part, bound to the  enzyme by a catalytically important carbamate group (—NH—COO ) that  is generated by the reaction of a nonsubstrate C0 2 with the £-amino group  of Lys 201. This essential reaction is catalyzed in vivo by the enzyme RuBP  carboxylase activase in an ATP-driven process.   B Calvin Cycle Products Are Converted to Starch, Sucrose,  and Cellulose   GAP, the primary product of photosynthesis, is used in a variety of biosyn¬  thetic pathways, both inside and outside the chloroplast. For example, it can  be converted to fructose-6-phosphate by the further action of Calvin cycle en¬  zymes and then to glucose-1-phosphate (G1P) by phosphoglucose isomerase  and phosphoglucomutase (Section 16-1C). G1P is the precursor of the higher  order carbohydrates characteristic of plants.   The polysaccharide a-amylose, a major component of starch (Section 8-2C),  is synthesized in the chloroplast stroma as a temporary storage depot for glucose  units. It is also synthesized as a long-term storage molecule elsewhere in the  plant, including leaves, seeds, and roots. G1P is first activated by its reaction  with ATP to form ADP-glucose as catalyzed by ADP-glucose pyrophosphor-  ylase. Starch synthase then transfers the glucose residue to the nonreducing end  of an a-amylose molecule, forming a new glycosidic linkage (Fig. 19-29). The  overall reaction is driven by the exergonic hydrolysis of the PP Z released in the  formation of ADP-glucose. A similar reaction sequence occurs in glycogen  synthesis, which uses UDP-glucose (Section 16-2).   FIG. 19-29 Starch synthesis. ADP-glucose is formed from G1P and ATP in a  phosphoanhydride exchange reaction (1). The PP, product is rapidly hydrolyzed.   ADP-glucose is the substrate for starch synthase (2), which adds the glucose residue  to an existing polysaccharide, releasing ADP.    Sucrose, a disaccharide of glucose and fructose (Section 8-2A), is the ma¬  jor transport sugar for delivering carbohydrates to nonphotosynthesizing cells  and hence is the major photosynthetic product of green leaves. Since sucrose  is synthesized in the cytosol, either glyceraldehyde-3-phosphate or dihydroxy-  acetone phosphate is transported out of the chloroplast by an antiporter that  exchanges phosphate for a triose phosphate. Two trioses combine to form fruc-  tose-6-phosphate (F6P) and subsequently glucose-1-phosphate (G1P), which  is then activated by UTP to form UDP—glucose. Next, sucrose-6-phosphate is  produced in a reaction catalyzed by sucrose-phosphate synthase. Finally, su-  crose-6-phosphate is hydrolyzed by sucrose-phosphate phosphatase to yield  sucrose (at right ), which is then exported to other plant tissues.   Cellulose, which consists of long chains of (3(1 —>4)-linked glucose units  and is the major polysaccharide of plants, is also synthesized from  UDP—glucose. Plant cell walls consist of nearly crystalline cables containing  ~36 cellulose chains, all embedded in an amorphous matrix of other polysac¬  charides and lignin (Section 8-2B). Unlike starch in plants or glycogen in  mammals, cellulose is synthesized by multisubunit enzyme complexes in the  plant plasma membrane and extruded into the extracellular space.   During the day, plants satisfy their energy needs via the light and dark reac¬  tions of photosynthesis. At night, however, like other organisms, they must  use their nutritional reserves to generate ATP and NADPH through glycoly¬  sis, oxidative phosphorylation, and the pentose phosphate pathway. Since the  stroma contains the enzymes of glycolysis and the pentose phosphate pathway   TABLE 19-1 Standard and Physiological Free Energy Changes for the Reactions of the Calvin Cycle   Source: Bassham, J.A. and Buchanan, B.B., in Govindjee (Ed.), Photosynthesis, Vol. II, p. 155, Academic Press (1982).    as well as those of the Calvin cycle, plants must have a light-sensitive control  mechanism to prevent the Calvin cycle from consuming this catabolically produced  ATP and NAD PH in a wasteful futile cycle.   As we have seen, the control of flux in a metabolic pathway occurs at en¬  zymatic steps that are far from equilibrium (large negative value of AG).  Inspection of Table 19-1 indicates that the three best candidates for flux con¬  trol in the Calvin cycle are the reactions catalyzed by RuBP carboxylase, FBPase,  and SBPase (Reactions 2 , 7, and 10 of Fig. 19-26). In fact, the catalytic effi¬  ciencies of the three enzymes all vary in vivo with the level of illumination.   1. pH. On illumination, the pH of the stroma increases from ~7.0 to  ~ 8.0 as protons are pumped from the stroma into the thylakoid lu¬  men. RuBP carboxylase has a sharp pH optimum near pH 8.0.   2 . [Mg 2+ ]. Recall that the light-induced influx of protons to the thy¬  lakoid lumen is accompanied by the efflux of Mg 2+ to the stroma  (Section 19-2D). This Mg 2+ stimulates RuBP carboxylase.   Many plants synthesize this compound, which inhibits RuBP carboxy¬  lase, only in the dark. RuBP carboxylase activase facilitates the release  of the tight-binding CA1P from RuBP carboxylase as well as catalyzing  its carbamoylation (Section 19-3A).    FIG. 1 9-30 The light-activation mechanism of FBPase and SBPase. Photoactivated  PSI reduces soluble ferredoxin (Fd), which reduces ferredoxin-thioredoxin reductase, which, in  turn, reduces the disulfide linkage of thioredoxin. Reduced thioredoxin reacts with the inactive  bisphosphatases by disulfide interchange, thereby activating the flux-controlling Calvin cycle  enzymes.    FBPase and SBPase are also activated by increased pH and [Mg 2+ ], and  by NADPH as well. The effect of these factors is complemented by a second  regulatory system that responds to the redox potential of the stroma.  Thioredoxin, an ~ 105-residue protein that occurs in many types of cells, con¬  tains a reversibly reducible disulfide group. Reduced thioredoxin activates both  FBPase and SBPase by a disulfide interchange reaction (Fig. 19-30). The re¬  dox level of thioredoxin is maintained by a second disulfide-containing enzyme,  ferredoxin-thioredoxin reductase, which directly responds to the redox state  of the soluble ferredoxin in the stroma. This in turn varies with the illumina¬  tion level. The thioredoxin system also deactivates phosphofructokinase (PFK),  the main flux-generating enzyme of glycolysis (Section 15-4A). Thus, in plants,  light stimulates the Calvin cycle while deactivating glycolysis, whereas darkness has  the opposite effect (that is, the so-called dark reactions do not occur in the dark).   It has been known since the 1960s that illuminated plants consume 0 2 and  evolve CO 2 in a pathway distinct from oxidative phosphorylation. In fact, at low  CO 2 and high 0 2 levels, this photorespiration process can outstrip photosynthetic  CO 2 fixation. The basis of photorespiration was unexpected: 0 2 competes with  CO 2 as a substrate for RuBP carboxylase (RuBP carboxylase is therefore also  called RuBP carboxylase-oxygenase or RuBisCO). In the oxygenase reac¬  tion, 0 2 reacts with the enzymes other substrate, RuBP, to form 3PG and  2-phosphoglycolate (Fig. 19-31). The 2-phosphoglycolate is hydrolyzed to  glycolate by phosphoglycolate phosphatase and, as described below, is par¬  tially oxidized to yield C0 2 by a series of enzymatic reactions that occur in  the peroxisome and the mitochondrion. Thus, photorespiration is a seemingly  wasteful process that undoes some of the work of photosynthesis. In this sec¬  tion we discuss the biochemical basis of photorespiration and how certain  plants manage to evade its deleterious effects.    FIG. 19-31 Probable mechanism of the  oxygenase reaction catalyzed by RuBP  carboxylase-oxygenase. Note the similarity  of this mechanism to that of the carboxylase  reaction catalyzed by the same enzyme  (Fig. 19-28).    FIG. 1 9-32 Photorespiration. This  pathway metabolizes the phosphoglycolate  produced by the RuBP carboxylase-catalyzed  oxidation of RuBP. The reactions occur, as  indicated, in the chloroplast, the peroxisome, the  mitochondrion, and the cytosol. Note that two  glycines are required to form serine + C0 2 .    Photorespiration Dissipates ATP and NADPH. The photorespiration path¬  way is outlined in Fig. 19-32. Glycolate is exported from the chloroplast to  the peroxisome (also called the glyoxysome; Section 17-5C), where it is oxi¬  dized by glycolate oxidase to glyoxylate and H 2 0 2 . The H 2 0 2 , a potentially  harmful oxidizing agent, is converted to H 2 0 and 0 2 by the heme-containing  enzyme catalase (Section 18-4B). The glyoxylate can be converted to glycine  in a transamination reaction, as is discussed in Section 21-2A, and exported  to the mitochondrion. There, two molecules of glycine are converted to one  molecule of serine and one of C0 2 . This is the origin of the C0 2 generated by   transamination reaction converts it to hydroxypyruvate. This substance is re¬  duced to glycerate and phosphorylated in the cytosol to 3PG, which  reenters the chloroplast and is reconverted to RuBP in the Calvin cycle. The  net result of this complex photorespiration cycle is that some of the ATP and  NADPH generated by the light reactions is uselessly dissipated.   Although photorespiration has no known metabolic function, the RuBP  carboxylases from the great variety of photosynthetic organisms so far tested all  exhibit oxygenase activity. Yet, over the eons, the forces of evolution must have  optimized the function of this important enzyme. Photorespiration may con¬  fer a selective advantage by protecting the photosynthetic apparatus from pho-  tooxidative damage when insufficient C0 2 is available to otherwise dissipate its  absorbed light energy. This hypothesis is supported by the observation that  when chloroplasts or leaf cells are brightly illuminated in the absence of both  C0 2 and 0 2 , their photosynthetic capacity is rapidly and irreversibly lost.   C 4 Plants Concentrate C0 2 . On a hot, bright day, when photosynthesis has  depleted the level of C0 2 at the chloroplast and raised that of 0 2 , the rate of  photorespiration approaches the rate of photosynthesis. This phenomenon is  a major limitation on the growth of many plants (and is therefore an impor¬  tant agricultural problem that is being attacked through genetic engineering  studies—none of which has yet been successful). However, certain species of  plants , such as sugarcane , corn, and most important weeds , have a metabolic cy¬  cle that concentrates C0 2 in their photosynthetic cells , thereby almost totally pre¬  venting photorespiration. The leaves of plants that have this so-called C 4 cycle  have a characteristic anatomy. Their fine veins are concentrically surrounded  by a single layer of so-called bundle-sheath cells, which in turn are sur¬  rounded by a layer of mesophyll cells.   The C 4 cycle (Fig. 19-33) was elucidated in the 1960s by Marshall Hatch  and Rodger Slack. It begins when mesophyll cells, which lack RuBP carboxy¬  lase, take up atmospheric C0 2 by condensing it as HCO3 with phospho-  enolpyruvate (PEP) to yield oxaloacetate. The oxaloacetate is reduced by   FIG. 1 9-33 The C 4 pathway. C0 2 is concentrated in the mesophyll cells and  transported to the bundle-sheath cells for entry into the Calvin cycle.   • How many different types of reactions  (e.g., phosphorylation, isomerization) occur  in the Calvin cycle?   • Which Calvin cycle steps are similar to  those of the pentose phosphate pathway?   • Describe how GAP produced by the  Calvin cycle is converted to sucrose,  starch, and cellulose.   • How do pH, Mg 2+ , and ferredoxin link the  light reactions with control of the Calvin  cycle?    NADPH to malate, which is exported to the bundle-sheath cells (the name C 4  refers to these four-carbon acids). There, the malate is oxidatively decarboxy-  lated by NADP to form C0 2 , pyruvate, and NADPH. The C0 2 , which has  been concentrated by this process, enters the Calvin cycle. The pyruvate is re¬  turned to the mesophyll cells, where it is phosphorylated to regenerate PEP.  The enzyme that mediates this reaction, pyruvate-phosphate dikinase, has the  unusual action of simultaneously phosphorylating pyruvate and P z with ATP s  (3 and 7 phosphoryl groups, respectively. The C 4 pathway thereby concentrates  CO 2 in the bundle-sheath cells at the expense of 2 ATP equivalents. Consequently\  photosynthesis in C 4 plants consumes a total of 5 ATP per C0 2 fixed versus the 3  ATP required by the Calvin cycle alone.   C4 plants occur largely in tropical regions because they grow faster under  hot and sunny conditions than other, so-called C 3 plants (so named because  they initially fix C0 2 in the form of three-carbon acids). In cooler climates,  where photorespiration is less of a burden, C 3 plants have the advantage be¬  cause they require less energy to fix C0 2 .   CAM Plants Store C0 2 through a Variant of the C 4 Cycle. A variant of the  C 4 cycle that separates C0 2 acquisition and the Calvin cycle in time rather  than in space occurs in many desert-dwelling succulent plants. If these plants  opened their stomata (pores in the leaves) by day to acquire C0 2 , as most  plants do, they would lose unacceptable amounts of water by evaporation. To  minimize this loss, these succulents absorb C0 2 only at night and use the re¬  actions of the C 4 pathway (Fig. 19-33) to store it as malate. This process is  known as crassulacean acid metabolism (CAM), so named because it was  first discovered in plants of the family Crassulaceae. The large amount of PEP  necessary to store a days supply of C0 2 is obtained by the breakdown of starch  via glycolysis. During the course of the day, the malate is broken down to  C0 2 , which enters the Calvin cycle, and pyruvate, which is used to resynthe¬  size starch. CAM plants are thus able to carry out photosynthesis with mini¬  mal water loss.   • Photosynthesis is the process whereby light energy drives the re¬  duction of C0 2 to yield carbohydrates. In plants and cyanobac¬  teria, photosynthesis oxidizes water to 0 2 .   • In plants, the photosynthetic machinery consists of protein com¬  plexes embedded in the thylakoid membrane and enzymes dis¬  solved in the stroma of chloroplasts.   • Chlorophyll and other light-absorbing pigments are organized in  light-harvesting complexes that funnel light energy to photosyn¬  thetic reaction centers (RCs).   • The purple bacterial photosynthetic reaction center (PbRC) un¬  dergoes photooxidation when it absorbs a photon. The excited  electron passes through a series of electron carriers before reduc¬  ing ubiquinone. The reduced ubiquinone is reoxidized by cy¬  tochrome bci, which in the process translocates four protons from  the cytosol to the periplasmic space via a Q cycle. The electron is  then returned to the PbRC via an electron-transport chain result¬  ing in no net oxidation-reduction.    • In plants and cyanobacteria, photosystems I and II (PSI and PSII)  operate in electrical series in an arrangement known as the Z-  scheme. The oxidation of water by the Mn-containing oxygen-  evolving center (OEC) is driven by the photooxidation of PSII.   • The electrons released by the photooxidation of PSII are trans¬  ferred, via plastoquinone, to the cytochrome ^/complex, which  mediates a proton-translocating Q cycle while passing the elec¬  trons to plastocyanin.   • The photooxidation of PSI drives the electrons obtained from  plastocyanin to ferredoxin and then to NADP + to produce  NADPH. In cyclic electron flow, however, electrons return to cy¬  tochrome b^f thereby bypassing the need for PSII photooxidation.   • The reaction centers of the PbRC, PSII, and PSI have similar  structures and mechanisms and therefore appear to have arisen  from a common ancestor.   • In photophosphorylation, the protons released by the oxidation of  H 2 0 and proton translocation into the thylakoid lumen generate  a transmembrane proton gradient that is tapped by chloroplast  ATP synthase to drive the phosphorylation of ADP. A similar  process occurs in purple photosynthetic bacteria.   • The dark reactions use the ATP and NADPH produced in the  light reactions to power the synthesis of carbohydrates from C0 2 .  In the first phase of the Calvin cycle, C0 2 reacts with ribulose-  1,5-bisphosphate (RuBP) to ultimately yield glyceraldehyde-3-  phosphate (GAP). The remaining reactions of the cycle regenerate  the RuBP acceptor of C0 2 .   • RuBP carboxylase, the key enzyme of the dark reactions, is regu¬  lated by pH, [Mg 2+ ], and the inhibitory compound 2-carboxyara-    binitol- 1 -phosphate (CA 1 P). The two bisphosphatases of the  Calvin cycle are controlled by the redox state of the chloroplast  via disulfide interchange reactions mediated in part by thiore-  doxin.   • Photorespiration, in which plants consume 0 2 and evolve C0 2 ,  uses the ATP and NADPH produced by the light reactions. C 4  plants minimize the oxygenase activity of RuBP carboxylase  (RuBisCO) by concentrating C0 2 in their photosynthetic cells.  CAM plants use a related mechanism to conserve water.   1 . The net equation for oxidative phosphorylation can be written  as   2 NADH + 2 H + + 0 2 ^2H 2 0 + 2 NAD +   Write an analogous equation for the light reactions of photosyn¬  thesis.   2 . Green sulfur bacteria use H 2 S as an electron donor for photo¬  synthesis. Write an equation that describes photosynthesis in  these organisms.   3. The “red tide” is a massive proliferation of certain algal species  that causes seawater to become visibly red. Describe the spectral  characteristics of the dominant photosynthetic pigments in the  algae.   4. Explain why the chlorophylls in light-harvesting complexes must  absorb light of shorter wavelength than the light that would di¬  rectly excite the special pair of PSI or PSII,   3. Calculate the energy of one mole of photons of red light  (X = 700 nm).   6 . How many moles of ATP could theoretically be synthesized un¬  der standard conditions using the energy of the photons in  Problem 3?   7. The three electron-transporting complexes of the thylakoid  membrane can be called plastocyanin-ferredoxin oxidoreduc-  tase, plastoquinone-plastocyanin oxidoreductase, and water-  plastoquinone oxidoreductase. What are the common names of  these enzymes and in what order do they act?   8 . H 2 1s O is added to a suspension of chloroplasts capable of pho¬  tosynthesis. Where does the label appear when the suspension is  exposed to sunlight?   9. Describe the functional similarities between the purple bacterial  photosynthetic reaction center and PSI.   10 . Calculate the change in free energy for the transit of two electrons  from the quinone pool to cytochrome c 2 in bacterial photosynthetic    electron transport. Assume that the reduction potential of cy¬  tochrome c 2 is similar to those of other c-type cytochromes.   11 . Estimate the change in free energy when P960 undergoes  photooxidation.   12 . Calculate \%°' and AC 0 ' for the light reactions in plants, that  is, the four-electron oxidation of H 2 0 by NADP + .   13. Use the solutions of Problems 19-5 and 19-12 to calculate how  many moles of photons of red light (X = 700 nm) are theoret¬  ically required to drive the four-electron oxidation of H 2 0 by  NADP + under standard conditions to produce one mole of 0 2 .   14. Use the solution of Problem 19-12 to calculate how many moles  of photons of UV light (X = 220 nm) would be required to drive  the four-electron oxidation of H 2 0 by NADP + under standard  conditions to produce one mole of 0 2 .   15. Under conditions of very high light intensity, excess absorbed so¬  lar energy is dissipated by the action of photoprotective proteins  in the thylakoid membrane. Explain why it is advantageous for  these proteins to be activated by buildup of the proton gradient  across the membrane.   16. Which of the mechanisms for dissipating light energy shown in  Fig. 19-6 would best protect the photosystems from excess light  energy?   17. Although the net equation for photosynthesis indicates that the  process can be measured in terms of either 0 2 produced or C0 2  fixed, in practice, these measurements are not necessarily equiv¬  alent. Explain.   18. Why is it possible for chloroplasts to absorb much more than  8-10 photons per 0 2 molecule evolved?   19. Predict the effect of adding myxothiazol, an inhibitor of electron  transport in mitochondrial Complex III, to a suspension of  chloroplasts exposed to light. Would either ATP or NADPH pro¬  duction be affected?   20. Why would “knocking out” a gene for a chloroplast fatty acid  desaturase (an enzyme involved in synthesizing fatty acids con¬  taining three double bonds) increase the rate of photosynthe¬  sis at 40°C, a temperature at which photosynthesis is normally  impaired?   21. Calculate the free energy change for moving a proton from the  thylakoid lumen to the stroma when ApH = 3.4, A^P = 0 , and  T = 23°C.   22 . Chloroplast ATP synthase contains 14 c subunits. How many  protons must be translocated to the thylakoid lumen to support  the synthesis of one ATP?   23. Predict the effect of an uncoupler such as dinitrophenol (Fig. 18-29)  on production of ATP in a chloroplast.   24. Predict the effect of an uncoupler such as dinitrophenol (Fig. 18-29)  on production of NADPH in a chloroplast.   23. Chloroplasts are illuminated until the levels of the Calvin cycle  intermediates reach a steady state. The light is then turned off.  How does the level of RuBP vary after this point?   26. For the chloroplasts described in Problem 25, how does the level  of 3PG vary after the light is turned off?   27. Calculate the energy cost of the Calvin cycle combined with  glycolysis and oxidative phosphorylation, that is, the ratio of  the energy spent synthesizing starch from C0 2 and photosyn-  thetically produced NADPH and ATP to the energy generated  by the complete oxidation of starch. Assume that each NADPH  is energetically equivalent to 2.5 ATP and that starch biosyn¬  thesis and breakdown are mechanistically identical to glycogen  synthesis and breakdown.   28. Oil is a major energy-storage molecule in most seeds. During seed  development in some plants, oil (triacylglycerols) is synthesized  using acetyl-CoA derived from sucrose. The developing seed also    contains RuBP carboxylase, although the Calvin cycle is not  active, (a) How does the carboxylase maximize the efficiency  of converting starch into oil? (b) The seeds that contain RuBP  carboxylase are green, suggesting that they harvest some light  energy. Explain why some energy collection would be useful in  the developing seed.   29. Cyanobacteria contain carboxysomes, which consist of a protein  shell that encloses RuBP carboxylase and is permeable to small  anions such as HC0 3 . Explain why carbonic anhydrase is also  a component of the carboxysome.   30. The shell of the cyanobacterial carboxysome, which contains  RuBP carboxylase, is relatively impermeable to 0 2 . Why would  this be an advantage for the bacteria?   31. Describe the effects of an increase in oxygen pressure on the dark  reactions of photosynthesis.   32. The leaves of some species of desert plants taste sour in the early  morning, but, as the day wears on, they become tasteless and  then bitter. Explain.   33. Plants must obtain C0 2 but avoid the loss of H 2 0 by evapora¬  tion. What would be the effect of increased atmospheric [C0 2 ]  on photosynthesis? Would the effect be the same for C 3 and C 4  plants?    Five distinct pathways for fixing  C0 2 , that is, incorporating it into biological molecules, have been de¬  scribed. In what types of organisms do the different pathways oc¬  cur? What is the free energy source for carbon fixation in each case?  Which of the processes can be considered to be photosynthesis?    Amunts, A., Drory, O., and Nelson, N., The structure of plant photo¬  system I at 3.4 A resolution, Nature 447, 58-63 (2007).   Blankenship, R.E., Molecular Mechanisms of Photosynthesis, Blackwell  Science ( 2002 ).   Chitnis, P.R., Photosystem I: Function and physiology, Annu. Rev. Plant  Physiol. Plant Biol. 52, 593-626 (2001).   Ferreira, K.N., Iverson, T.M., Maghlaoui, K., Barber, J., and Iwata, S.,  Architecture of the photosynthetic oxygen-evolving center, Science  303, 1831-1838 (2004)   Fromme, P. (Ed.), Photosynthetic Protein Complexes. A Structural Approach,  Wiley-Blackwell (2008).   Guskov, A., Kern, J., Gabdulkhakov, A., Broser, M., Zouni, A., and  Saenger, W., Cyanobacterial photosystem II at 2 . 9 -A resolution and  the roles of quinones, lipids, channels, and chloride, Nature Struct.  Mol. Biol. 16, 334-342 (2009). [The X-ray structure of PSII.]   Heathcote, P., Fyfe, P.K., and Jones, M.R., Reaction centres: the struc¬  ture and evolution of biological solar power, Trends Biochem. Sci. 27,  79-87 ( 2002 ).    Jordan, P., Fromme, P., Witt, H.T., Klukas, O., Saenger, W., and Krauss,  N., Three-dimensional structure of cyanobacterial photosystem I at  2.5 A resolution, Nature AW, 909-917 ( 2001 ).   Koepke, J., Hu, X., Muenke, C., Schulen, K., and Michel, H., The crys¬  tal structure of the light-harvesting complex II (B800-850) from  Rho do spirillum molischianum, Structure A, 581-597 (1996).   Kurisu, G., Zhang, H., Smith, J.L., and Cramer, W.A., Structure of the  cytochrome £ 6 /complex of oxygenic photosynthesis: Tuning the cav¬  ity, Science 302, 1009-1014 (2003); and Stroebel, D., Choquet, Y.,  Popot, J.-L., and Picot, D., An atypical haem in the cytochrome h^f  complex, Nature 426, 413-418 (2003).   Spreitzer, R.J. and Salvucci, M.E., Rubisco: structure, regulatory interac¬  tions, and possibilities for a better enzyme, Annu. Rev. Plant  Biochem. 53, 449-475 ( 2002 ).   Yano, J., et al., Where water is oxidized to dioxygen: Structure of the pho¬  tosynthetic Mn 4 Ca cluster, Science 314, 821-825 (2006).   Just before pupation, the larvae of some insects synthesize large quantities of fatty  acids in order to stockpile them as triacylglycerols to fuel the structural changes that  occur during metamorphosis. [iStockphoto.]    Most cells contain a wide variety of lipids, but many of these structurally dis¬  tinct molecules are functionally similar. For example, most cells can tolerate  variations in the lipid composition of their membranes, provided that mem¬  brane fluidity, which is largely a property of their component fatty acid chains,  is maintained (Section 9-2B).   Even greater variation is exhibited in the cellular content of lipids stored  as energy reserves. Triacylglycerol stores are built up and gradually depleted in  response to changing physiological demands. The camels hump is a well-  known example of a fat depot that supplies energy (the catabolism of the fat  also generates water). Other organisms that undergo dramatic changes in body  fat content are hibernating mammals and birds that migrate long distances  without refueling. Lipid metabolism in those cases is notable for the sheer  amount of material that flows through the few relatively simple biosynthetic  and degradative pathways.   In this chapter, we acknowledge the central function of lipids in energy  metabolism by first examining the absorption and transport of their compo¬  nent fatty acids and the oxidation of fatty acids to produce energy. The sec¬  ond part of this chapter examines the synthesis of fatty acids and other lipids,  including glycerophospholipids, sphingolipids, and cholesterol.   A Fatty Acids Are Activated by Their Attachment  to Coenzyme A   C C 2 o Patty Acids Are the Precursors of  Prostaglandins    • Triacylglycerols are broken down by lipases, and the products are absorbed by  the intestine.   • Lipoproteins transport lipids between the intestines, liver, and other tissues.   A Cholesterol Is Synthesized from Acetyl-CoA  B HMG-CoA Reductase Controls the Rate of  Cholesterol Synthesis    Triacylglycerols (also called fats or triglycerides) constitute ~90% of dietary  lipids and are the major form of metabolic energy storage in humans.    FIG. 20-1 Structures of the major bile  acids and their glycine and taurine  conjugates.    Triacylglycerols consist of glycerol triesters of fatty acids such as palmitic and  oleic acids   (the names and structural formulas of some biologically common fatty acids  are listed in Table 9-1). The mechanisms for digesting, absorbing, and trans¬  porting triacylglycerols from the intestine to the tissues must accommodate  their inherent hydrophobicity.   Since triacylglycerols are water insoluble , whereas digestive enzymes are water sol¬  uble , triacylglycerol digestion takes place at lipid—water interfaces. The rate of tri-  acylglycerol digestion therefore depends on the surface area of the interface,  which is greatly increased by the churning peristaltic movements of the intes¬  tine combined with the emulsifying action of bile acids. The bile acids (also  called bile salts) are amphipathic detergent-like molecules that act to solubi¬  lize fat globules. Bile acids are cholesterol derivatives that are synthesized by  the liver and secreted as glycine or taurine conjugates (Fig. 20-1) into the  gallbladder for storage. From there, they are secreted into the small intestine,  where lipid digestion and absorption mainly take place.   Lipases Act at the Lipid-Water Interface. Pancreatic lipase (triacylglycerol li¬  pase) catalyzes the hydrolysis of triacylglycerols at their 1 and 3 positions to form  sequentially 1,2-diacylglycerols and 2-acylglycerols, together with the Na +  and K + salts of fatty acids (soaps). The enzymatic activity of pancreatic lipase  greatly increases when it contacts the lipid-water interface, a phenomenon known  as interfacial activation. Binding to the lipid-water interface requires mixed  micelles of phosphatidylcholine and bile acids, as well as pancreatic colipase, a  90-residue protein that forms a 1:1 complex with lipase. The X-ray structures,  determined by Christian Cambillau, of pancreatic lipase-colipase complexes  reveal the structural basis of the interfacial activation of lipase as well as how col¬  ipase and micelles help lipase bind to the lipid-water interface (Fig. 20-2).   The active site of the 449-residue pancreatic lipase, which is contained in  the enzymes N-terminal domain (residues 1-336), contains a catalytic triad that   FIG. 20-2 X-Ray structures of pancreatic lipase in complex  with colipase, (a) In aqueous solution, and (b) cocrystallized with  mixed micelles of phosphatidylcholine and bile salts. The lipase is  drawn in ribbon form with its N-terminal domain (residues 1-336)  cyan, its C-terminal domain (residues 337-449) green, the lid  (residues 237-262) magenta, and the p5 loop (residues 76-85)    orange. The colipase is yellow. A phosphatidylcholine molecule that is  bound in the lipase active site in Part b is shown in stick form with  C green, 0 red, and P orange. The micelles, which have irregular  structures, are not visible. [Based on X-ray structures by Christian  Cambillau, LCCMB-CNRS, Marseille, France. PDBids 1 N8S and  1 LPBJ    closely resembles that in serine proteases (Section 11-5B; recall that ester hydrol¬  ysis is mechanistically similar to peptide hydrolysis). In the absence of lipid mi¬  celles, lipases active site is covered by a 26-residue helical “lid.” However, in the  presence of the micelles, the lid undergoes a complex structural reorganization  that exposes the active site. Simultaneously, a 10-residue loop, called the (35 loop,  changes conformation in a way that forms the active enzymes oxyanion hole and  generates a hydrophobic surface near the entrance to the active site.   Colipase binds to the C-terminal domain of lipase (residues 337-449)  such that the hydrophobic tips of its three loops extend from the complex.  This creates a continuous hydrophobic plateau, extending >50 A past the  lipase active site, that presumably helps bind the complex to the lipid surface.  Colipase also forms three hydrogen bonds to the opened lid, thereby stabiliz¬  ing it in that conformation.   Other lipases, such as phospholipase A 2 (Fig. 9-6), also preferentially cat¬  alyze reactions at interfaces. Instead of changing its conformation, however,  phospholipase A 2 contains a hydrophobic channel that provides the substrate  with direct access from the phospholipid aggregate (micelle or membrane)  surface to the bound enzymes active site (Fig. 20-3). Hence, on leaving its  micelle to bind to the enzyme, the substrate need not become solvated and  then desolvated. In contrast, soluble and dispersed phospholipids must first  surmount those significant kinetic barriers in order to bind to the enzyme.   Bile Acids and Fatty Acid-Binding Protein Facilitate the Intestinal  Absorption of Lipids. The mixture of fatty acids and mono- and diacylglyc-  erols produced by lipid digestion is absorbed by the cells lining the small in¬  testine (the intestinal mucosa). Bile acids not only aid lipid digestion; they are  essential for the absorption of the digestion products. The micelles formed by the  bile acids take up the nonpolar lipid degradation products so as to permit their  transport across the unstirred aqueous boundary layer at the intestinal wall.  The importance of this process is demonstrated in individuals with obstructed  bile ducts: They absorb little of their ingested lipids but, rather, eliminate them  in hydrolyzed form in their feces. Bile acids are likewise required for the effi¬  cient intestinal absorption of the lipid-soluble vitamins A, D, E, and K.   Inside the intestinal cells, fatty acids form complexes with intestinal fatty  acid-binding protein (I-FABP), a cytoplasmic protein that increases the ef¬  fective solubility of the water-insoluble substances and also protects the cell  from their detergent-like effects. The X-ray structure of rat I-FABP, determined  by James Sacchettini, shows that this monomeric, 131-residue protein consists   FIG. 20-3 Substrate binding to  phospholipase A 2 . (a) Hypothetical model of  phospholipase A 2 in complex with a micelle of  lysophosphatidylethanolamine as shown in cross  section. The protein is drawn in cyan, the  phospholipid head groups are yellow, and their  hydrocarbon tails are blue. The calculated atomic  motions of the assembly are indicated through a  series of superimposed images taken at 5-ps  intervals. [Courtesy of Raymond Salemme, E. I.  du Pont de Nemours & Company.] (b) Schematic  diagram of a phospholipid contained in a micelle  entering the hydrophobic phospholipid channel in  phospholipase A 2 (red arrow).   FIG. 20-4 X-Ray structure of rat intestinal  fatty acid-binding protein in complex with  palmitate. The protein is colored in rainbow  order from its N-terminus (blue) to its  C-terminus (red). The palmitate is drawn in  space-filling form with C green and 0 red.   [Based on an X-ray structure by James  Sacchettini, Texas A&M University. PDBid 21FB.]    largely of 10 antiparallel (3 strands stacked in two approximately orthogonal  (3 sheets (Fig. 20 -4). A fatty acid molecule occupies a gap between two of the  (3 strands, lying between the (3 sheets so that it is more or less parallel to the  gapped (3 strands (a structure that has been dubbed a “(3-clam”). The fatty  acids carboxyl group interacts with Arg 106, Gin 115, and two bound water  molecules, whereas its tail is encased by the side chains of several hydropho¬  bic, mostly aromatic residues.   The fatty acid products of lipid digestion that are absorbed by the intestinal  mucosa make their way to other tissues for catabolism or storage. But because  they are only sparingly soluble in aqueous solution, lipids are transported by  the circulation in complex with proteins.   Lipoproteins Are Complexes of Lipid and Protein. Lipoproteins are globu¬  lar micelle-like particles that consist of a nonpolar core of triacylglycerols and cho-  lesteryl esters surrounded by an amphiphilic coating of protein, phospholipid, and  cholesterol. There are five classes of lipoproteins (Table 20-1), which vary in  composition and physiological function.   Intestinal mucosal cells convert dietary fatty acids to triacylglycerols and pack¬  age them, along with dietary cholesterol, into lipoproteins called chylomicrons.  These particles are released into the intestinal lymph and are transported through  the lymphatic vessels before draining into the large veins. (The lymphatic circu¬  lation serves to collect extracellular fluid that leaks out of blood vessels under high  pressure.) The bloodstream then delivers chylomicrons throughout the body.  Other lipoproteins known as very low density lipoproteins (VLDL), interme¬  diate density lipoproteins (IDL), and low density lipoproteins (LDL) are syn¬  thesized by the liver to transport endogenous (internally produced) triacylglycerols  and cholesterol from the liver to the tissues. High density lipoproteins (HDL)  transport cholesterol and other lipids from the tissues back to the liver.   Each lipoprotein contains just enough protein, phospholipid, and choles¬  terol to form an ~20-A-thick monolayer of these substances on the particle  surface (Fig. 20-5). Lipoprotein densities increase with decreasing particle  diameter because the density of their outer coating is greater than that of their  inner core. Thus, the HDL, which are the most dense of the lipoproteins, are  also the smallest.    TABLE 20-1 Characteristics of the Major Classes of Lipoproteins in Human Plasma   Apolipoproteins Coat Lipoprotein Surfaces. The protein components of  lipoproteins are known as apolipoproteins or just apoproteins. At least nine  apolipoproteins are distributed in different amounts in the human lipoproteins  (Table 20-1). For example, LDL contain apolipoprotein B-100 (apoB-100).  This protein, a 4536-residue monomer (and thus one of the largest monomeric  proteins known), has a hydrophobicity approaching that of integral membrane  proteins. Each LDL particle contains only one molecule of apoB-100, which  appears to cover at least half of the particle surface (Fig. 20-5).   Unlike apoB-100, the other apolipoproteins are water soluble and associ¬  ate rather weakly with lipoproteins. These apolipoproteins also have a high  helix content, which increases when they are incorporated into lipoproteins.  Contact with a hydrophobic surface apparently favors formation of helices,  which satisfy the hydrogen bonding potential of the proteins polar backbone  groups. In addition, the helices in apolipoproteins have hydrophilic and hydropho¬  bic side chains on opposite sides of the helical cylinder ; suggesting that lipoprotein  a helices are amphipathic and float on phospholipid surfaces , much like logs on  water. The charged head groups of the lipids presumably bind to oppositely  charged residues on the helix while the first few methylene groups of their  fatty acyl chains associate with the nonpolar face of the helix.   Apolipoprotein A-I (apoA-I), which occurs in chylomicrons and HDL,  is a 243-residue, 29-kD polypeptide. It consists largely of tandem 22-residue  segments of similar sequence. The X-ray structure of a truncated apolipopro¬  tein A-I (lacking residues 1-43) reveals that the polypeptide chain forms a  pseudocontinuous a helix that is punctuated by kinks at Pro residues spaced  about every 22 residues. Four monomers associate to form the structure shown  in Fig. 20 -Ga. The size and twisted elliptical shape of the complex seem ideal  for wrapping around an HDL particle. Figure 20-6 b contains a helical wheel  representation of a portion of apoA-I, illustrating the amphipathic nature of  the helix.    FIG. 20-5 Diagram of LDL, the major  cholesterol carrier of the bloodstream. This  spheroidal particle consists of some 1500  cholesteryl ester molecules surrounded by an  amphiphilic coat of —800 phospholipid  molecules, —500 cholesterol molecules, and a  single 4536-residue molecule of apolipoprotein  B-100.   Chylomicrons adhere to binding sites on the inner surface (endothe¬  lium) of the capillaries in skeletal muscle and adipose tissue. The  chylomicrons component triacylglycerols are hydrolyzed through  the action of the extracellular enzyme lipoprotein lipase. The tis¬  sues then take up the liberated monoacylglycerol and fatty acids.   The chylomicrons shrink as their triacylglycerols are progressively  hydrolyzed until they are reduced to cholesterol-enriched  chylomicron remnants. The remnants dissociate from the capillary  endothelium and re-enter the circulation to be taken up by the ^    FIG. 20-6 Structure of human apolipoprotein A-I. (a) The X-ray  structure of the D 2 -symmetric homotetramer with its four subunits, which  lack their N-terminal 43 residues, drawn in different colors. The complex,  which has a twisted ellipsoidal shape, is viewed along one of its twofold  axes. [Based on an X-ray structure by David Borhani, Southern Research  Institute, Birmingham, Alabama, and Christie Brouillette, University of  Alabama Medical Center, Birmingham, Alabama. PDBid 1AV1.] (b) A helical  wheel projection of the amphipathic a helix constituting residues 148 to  164 of apolipoprotein A-I (in a helical wheel representation, the side chain  positions are projected down the helix axis onto a plane). Note the  segregation of nonpolar and polar residues to different sides of the helix as  well as the segregation of the basic residues to the outer edges of the polar  surface, where they can interact with the anionic head groups of membrane  lipids. Other apolipoprotein helices have similar polarity distributions. [After  Kaiser, E.T., in Oxender, D.L and Fox, C.F. (Eds.), Protein Engineering, p. 194,  Liss (1987).]    Trace the path of dietary lipids from the stomach to other tissues. What functions does the liver perform?    liver. Chylomicrons therefore deliver dietary triacylglycerols to muscle and adipose  tissue\ and dietary cholesterol to the liver (Fig. 20-7, blue arrows ).   VLDL Are Gradually Degraded. Very low density lipoproteins (VLDL), which  transport endogenous triacylglycerols and cholesterol, are also degraded by  lipoprotein lipase in the capillaries of adipose tissue and muscle (Fig. 20-7,  red arrows ). The released fatty acids are taken up by the cells and oxidized for  energy or used to resynthesize triacylglycerols. The glycerol backbone of tri¬  acylglycerols is transported to the liver or kidneys and converted to the glycolytic  intermediate dihydroxyacetone phosphate. Oxidation of this three-carbon unit,  however, yields only a small fraction of the energy available from oxidizing the  three fatty acyl chains of a triacylglycerol.   After giving up their triacylglycerols, the VLDL remnants, which have also  lost some of their apolipoproteins, appear in the circulation first as IDL and  then as LDL. About half of the VLDL, after degradation to IDL and LDL,  are taken up by the liver.   Cells Take Up LDL by Receptor-Mediated Endocytosis. Animal cells acquire  cholesterol, an essential component of cell membranes (Section 9-2B), either    by synthesizing it or by taking up LDL, which are rich in cholesterol and cho-  lesteryl esters. The latter process, as Michael Brown and Joseph Goldstein have  demonstrated, occurs by receptor-mediated endocytosis (engulfment) of the  LDL (Fig. 20-8). The LDL particles are sequestered by LDL receptors, cell-  surface transmembrane glycoproteins that specifically bind apoB-100. LDL  receptors cluster into clathrin-coated pits, which gather the cell-surface recep¬  tors that are destined for endocytosis while excluding other cell-surface proteins.  The coated pits invaginate from the plasma membrane to form clathrin-  coated vesicles (Section 9-4E). Then, after divesting themselves of their  clathrin coats, the vesicles fuse with vesicles known as endosomes, whose   receptor is synthesized on the endoplasmic reticulum (a), processed in  the Golgi apparatus (b), and inserted into the plasma membrane (c),  where it becomes a component of clathrin-coated pits (d). The  apolipoprotein B-100 (apoB-100) component of LDL specifically binds  to LDL receptors on clathrin-coated pits (1). These bud into the cell  ( 2 ) to form clathrin-coated vesicles, whose clathrin coats depolymerize  to form triskelions (3), which are recycled to the cell surface. The  uncoated vesicles fuse with vesicles called endosomes (4), which have  an internal pH of —5.0. The acidity induces the LDL particle to disso¬  ciate from its receptor. The LDL accumulates in the vesicular portion  of the endosome, whereas the LDL receptors concentrate in the  membrane of an attached tubular structure, which then separates from  the endosome (5) and subsequently recycles the LDL receptors to the    plasma membrane ( 6 ). The vesicular portion of the endosome fuses  with a lysosome (7), yielding a secondary lysosome in which the  apoB-100 component of the LDL and the cholesteryl esters are  hydrolyzed. This releases cholesterol ( 8 ), which either is converted to  cholesteryl esters through the action of acyl-CoA:cholesterol acyl-  transferase (ACAT) and sequestered in a droplet (9) or proceeds to  the endoplasmic reticulum (10). Increased cholesterol concentration in  the endoplasmic reticulum decreases the rate of synthesis of HMG-  CoA reductase (the enzyme that catalyzes the rate-limiting step of  cholesterol biosynthesis; Section 20-7B) and LDL receptors (down  arrows ), while increasing that of ACAT (up arrow). [After Brown, M.S.  and Goldstein, J.L, Curr. Topics Cell. Reg. 26, 7 (1985).] O See the  Animated Figures.   Chapter 20 Lipid Metabolism internal pH is ~5.0. Under these conditions, the LDL particle dissociates from   its receptor. The receptors are recycled back to the cell surface, while the en-  dosome with its enclosed LDL fuses with a lysosome. In the lysosome, LDLs  apoB-100 is rapidly degraded to its component amino acids, and the choles-  teryl esters are hydrolyzed to yield cholesterol and fatty acids.   Receptor-mediated endocytosis is a general mechanism whereby cells take up large  molecules, each through a corresponding specific receptor. Many cell-surface receptors  recycle between the plasma membrane and the endosomal compartment as does  the LDL receptor, even in the absence of ligand. The LDL receptor cycles in and  out of the cell membrane about every 10 minutes. New receptors are synthesized  on the endoplasmic reticulum and travel via vesicles through the Golgi complex  to the cell surface (Fig. 20-8; Section 9-4E). The intracellular concentration of free  cholesterol controls the rate of LDL receptor synthesis (Section 20-7). However,  defects in the LDL receptor system lead to abnormally high levels of circulating  cholesterol with the attendant increased risk of heart disease (Section 20-7C).   • Explain the purpose of bile acids and fatty  acid-binding protein in the digestion and  absorption of lipids.   • Describe the overall structure of lipopro¬  teins. What distinguishes the different  types?   • Summarize the roles of the various  lipoproteins in transporting triacylglycerols  and cholesterol.    HDL Transport Cholesterol from the Tissues to the Liver. HDL have  essentially the opposite function of LDL: They remove cholesterol from the tissues.  HDL are assembled in the plasma from components largely obtained through  the degradation of other lipoproteins. A circulating HDL particle acquires its  cholesterol by extracting it from cell-surface membranes. The cholesterol is then  converted to cholesteryl esters by the HDL-associated enzyme lecithin-  cholesterol acyltransferase (LCAT), which is activated by apoA-I. HDL  therefore function as cholesterol scavengers.   The liver is the only organ capable of disposing of significant quantities of choles¬  terol (by its conversion to bile acids; Fig. 20-1). About half of the VLDL, after  their degradation to IDL and LDL, are taken up by the liver via LDL receptor-  mediated endocytosis (Fig. 20-7). However, liver cells take up HDL by an entirely  different mechanism: Rather than being engulfed and degraded, an HDL particle  binds to a cell-surface receptor named SR-BI (for scavenger receptor class B type  I) and selectively transfers its component lipids to the cell. The lipid-depleted  HDL particle then dissociates from the cell and re-enters the circulation.    • Fatty acids to be degraded are linked to CoA in an ATP-dependent reaction.   • Fatty acyl groups are transported into the mitochondrion via a carnitine shuttle for  oxidation.   • Each round of mitochondrial p oxidation produces FADH 2 , NADH, and acetyl-CoA.   • The propionyl-CoA produced by the oxidation of odd-chain fatty acids is con¬  verted to succinyl-CoA.   • Peroxisomes oxidize long-chain fatty acids, producing H 2 0 2 .    The triacylglycerols stored in adipocytes are mobilized in times of metabolic need  by the action of hormone-sensitive lipase (Section 20-5). The free fatty acids  are released into the bloodstream, where they bind to albumin, a soluble 66-kD  monomeric protein. In the absence of albumin, the maximum solubility of fatty  acids is ~ 10 6 M; the effective solubility of fatty acids in complex with albu¬  min is as high as 2 mM. Nevertheless, those rare individuals with analbumine-  mia (severely depressed levels of albumin) suffer no apparent adverse symptoms;  evidently, their fatty acids are transported in complex with other serum proteins.   Fatty acids are catabolized by an oxidative process that releases free energy.  The biochemical strategy of fatty acid oxidation was understood long before  the oxidative enzymes were purified. In 1904, Franz Knoop, in the first use   w-Phenyl-labeled fatty acids containing an odd number of carbon  atoms are oxidized to the phenyl-labeled product, benzoic acid,  whereas those with an even number of carbon atoms are oxidized   to the phenyl-labeled C 2 product, phenylacetic acid. These products  are excreted as their respective glycine amides, hippuric and  phenylaceturic acids. The vertical arrows indicate the deduced sites of  carbon oxidation. The intermediate C 2 products are oxidized to C0 2  and H 2 0 and were therefore not isolated.    of chemical labels to trace metabolic pathways, fed dogs fatty acids labeled at  their 0 ) (last) carbon atom by a benzene ring and isolated the phenyl-containing  metabolic products from their urine. Dogs fed labeled odd-chain fatty acids  excreted hippuric acid, the glycine amide of benzoic acid, whereas those fed  labeled even-chain fatty acids excreted phenylaceturic acid, the glycine amide  of phenylacetic acid (Fig. 20-9). Knoop therefore deduced that fatty acids are  progressively degraded by two-carbon units and that the process involves the  oxidation of the carbon atom (3 to the carboxyl group. Otherwise, the phenyl¬  acetic acid would be further oxidized to benzoic acid. Knoop’s |3 oxidation hy¬  pothesis was only confirmed in the 1950s. The (3-oxidation pathway is a series  of enzyme-catalyzed reactions that operates in a repetitive fashion to  progressively degrade fatty acids by removing two-carbon units.   A Fatty Acids Are Activated by Their Attachment to Coenzyme A   Before fatty acids can be oxidized, they must be “primed” for reaction in an  ATP-dependent acylation reaction to form fatty acyl-CoA. The activation  process is catalyzed by a family of at least three acyl-CoA synthetases (also  called thiokinases) that differ in their chain-length specificities. These en¬  zymes, which are associated with either the endoplasmic reticulum or the outer  mitochondrial membrane, all catalyze the reaction   Fatty acid + CoA + ATP v acyl-CoA + AMP + PP Z   This reaction proceeds via an acyladenylate mixed anhydride intermediate  that is attacked by the sulfhydryl group of CoA to form the thioester product  (Fig. 20-10), thereby preserving the free energy of ATP hydrolysis in the  “high-energy” thioester bond (Section 14-2D). The overall reaction is driven  to completion by the exergonic hydrolysis of pyrophosphate catalyzed by in¬  organic pyrophosphatase.   Although fatty acids are activated for oxidation in the cytosol, they are oxi¬  dized in the mitochondrion, as Eugene Kennedy and Albert Lehninger estab¬  lished in 1950. We must therefore consider how fatty acyl-CoA is transported  across the inner mitochondrial membrane. A long-chain fatty acyl-CoA    FIG. 20-10 The mechanism of fatty acid activation catalyzed by acyl-CoA  synthetase. Formation of acyl-CoA involves an intermediate acyladenylate mixed anhydride.    cannot directly cross the inner mitochondrial membrane. Instead, its acyl por¬  tion is first transferred to carnitine, a compound that occurs in both plant  and animal tissues.   This transesterification reaction has an equilibrium constant close to 1, which  indicates that the O-acyl bond of acyl-carnitine has a free energy of hydrol¬  ysis similar to that of acyl-CoA’s thioester bond. Carnitine palmitoyl trans¬  ferases I and II, which can transfer a variety of acyl groups (not just palmitoyl  groups), are located, respectively, on the external and internal surfaces of the  inner mitochondrial membrane. The translocation process itself is mediated  by a specific carrier protein that transports acyl-carnitine into the mitochon¬  drion while transporting free carnitine in the opposite direction. The acyl-CoA  transport system is diagrammed in Fig. 20-11.    FIG. 20-1 1 The transport of fatty acids  into the mitochondrion. (1) The acyl group of  a cytosolic acyl-CoA is transferred to carnitine,  thereby releasing the CoA to its cytosolic pool.   ( 2 ) The resulting acyl-carnitine is transported into  the mitochondrial matrix by the carrier protein.   ( 3 ) The acyl group is transferred to a CoA  molecule from the mitochondrial pool. ( 4 ) The  product carnitine is returned to the cytosol.   The degradation of fatty acyl-CoA via (3 oxidation occurs in four reactions   1. Formation of a trans-a,(3 double bond through dehydrogenation by  the flavoenzyme acyl-CoA dehydrogenase (AD).   2. Hydration of the double bond by enoyl-CoA hydratase (EH) to form  a 3-L-hydroxyacyl-CoA.   3. NAD + -dependent dehydrogenation of the (3-hydroxyacyl-CoA by  3-L-hydroxyacyl-CoA dehydrogenase (HAD) to form the correspon¬  ding (3-ketoacyl-CoA.   4. C a —Cp cleavage in a thiolysis reaction with CoA as catalyzed by  (3-ketoacyl-CoA thiolase (KT; also called just thiolase) to form acetyl-  CoA and a new acyl-CoA containing two fewer C atoms than the  original one.    ^ CH 3 - (CH 2 ) n -Cp-C a -C-SCoA    FIG. 20-12 The p-oxidation pathway of fatty acyl-CoA. Reactions 1-4 degrade fatty acyl-CoA to acetyl-CoA. Reactions 5-8 are  electron transfers. Q See the Animated Figures.    FIG. 20-13 Ribbon diagram of the active site region of  medium-chain acyl-CoA dehydrogenase. The enzyme, from pig liver  mitochondria, is a tetramer of identical 385-residue subunits, each of  which binds an FAD prosthetic group (green) and an octanoyl-CoA  substrate (whose octanoyl and CoA moieties are blue and white) in  largely extended conformations. The octanoyl-CoA binds such that its  C a —C p bond is sandwiched between the carboxylate group of Glu 376  (red) and the flavin ring (green), consistent with the proposal that Glu  376 is the general base that abstracts the a proton in the a,|3  dehydrogenation reaction catalyzed by the enzyme. [Based on an X-ray  structure by Jung-Ja Kim, Medical College of Wisconsin. PDBid 3MDE.]  ©• See Interactive Exercise 27.   Mitochondria contain four acyl-CoA dehydrogenases, with specificities for  short (C 4 to Q), medium (C 6 to C 10 ), long (between medium and very long),  and very long chain (C 12 to C 18 ) fatty acyl-CoAs. The reaction catalyzed by  these enzymes is thought to involve removal of a proton at C a and transfer  of a hydride ion equivalent from Cp to FAD (Fig. 20-12, Reaction 1). The  X-ray structure of medium-chain acyl-CoA dehydrogenase (MCAD) in  complex with octanoyl-CoA, determined by Jung-Ja Kim, clearly shows how  the enzyme orients a basic group (Glu 376), the substrate C a —Cp bond, and  the FAD prosthetic group for reaction (Fig. 20-13).   r A deficiency of MCAD has been identified in —10% of cases of sudden  infant death syndrome (SIDS). Glucose is the principal metabolic fuel  just after eating, but when the glucose level later decreases, the rate of fatty  acid oxidation must correspondingly increase. The sudden death of infants  lacking MCAD may be caused by the imbalance between glucose and fatty  acid oxidation.   The FADH 2 resulting from the oxidation of the fatty acyl-CoA substrate  is reoxidized by the mitochondrial electron-transport chain through a series of  electron-transfer reactions. Electron-transfer flavoprotein (ETF) transfers an  electron pair from FADH 2 to the flavo-iron-sulfur protein ETF:ubiquinone  oxidoreductase, which in turn transfers an electron pair to the mitochondrial  electron-transport chain by reducing coenzyme Q (CoQ; Fig. 20-12,  Reactions 5-8). Reduction of 0 2 to H 2 0 by the electron-transport chain  beginning at the CoQ stage results in the synthesis of approximately 1.5 ATP  per electron pair transferred (Section 18-3C).   Long-Chain Enoyl-CoAs Are Converted to Acetyl-CoA and a Shorter Acyl-  CoA by Mitochondrial Trifunctional Protein. The product of acyl-CoA dehy¬  drogenases are 2-enoyl-CoAs. Depending on their chain lengths, their  processing is continued by one of three systems (Fig. 20-12): the short chain,  medium chain, or long chain 2-enoyl-CoA hydratases (EHs), hydroxyacyl-CoA  dehydrogenases (HADs), and 3-ketoacyl-CoA thiolases (KTs). The long chain  (LC) versions of these enzymes are contained on one a 4 p 4 heterooctameric,  trifunctional protein, located in the inner mitochondrial membrane. LCEH  and LCHAD are contained on the a subunits while LCKT is located on the  (3 subunits. The protein is therefore both a multifunctional protein (more than  one enzyme activity on a single polypeptide chain) and a multienzyme  complex (a complex of polypeptides catalyzing more than one reaction). The  advantage of such a trifunctional enzyme is the ability to channel the inter-    mediates toward the final product. Indeed, no long-chain hydroxyacyl-CoA  or ketoacyl-CoA intermediates are released into solution by this system.   The Thiolase Reaction Occurs via Claisen Ester Cleavage. The fourth  step of (3 oxidation is the thiolase reaction (Fig. 20-12, Reaction 4), which  yields acetyl-Co A and a new acyl-CoA that is two carbon atoms shorter than  the one that began the cycle (Fig. 20-14):   1. An active site thiol group adds to the (3-keto group of the sub¬  strate acyl-Co A.   2. Carbon—carbon bond cleavage yields a thioester between the  acyl-CoA substrate and the active site thiol group, together with  an acetyl-CoA carbanion intermediate that is stabilized by elec¬  tron withdrawal into the thioester s carbonyl group.   This type of reaction is known as a Claisen ester  cleavage (the reverse of a Claisen condensation).   The citric acid cycle enzyme citrate synthase also  catalyzes a reaction that involves a stabilized acetyl-  CoA carbanion intermediate (Section 17-3A).   3. An enzyme acidic group protonates the acetyl-CoA  carbanion, yielding acetyl-CoA.   enzyme—thioester intermediate, yielding an acyl-CoA that is  shortened by two C atoms.    Fatty Acid Oxidation Is Highly Exergonic. The function of fatty acid oxida¬  tion is, of course, to generate metabolic energy. Each round of (3 oxidation pro¬  duces one NADH, one FADH 2 , and one acetyl-CoA. Oxidation of acetyl-CoA  via the citric acid cycle generates an additional FADH 2 and 3 NADH, which  are reoxidized through oxidative phosphorylation to form ATP. Complete oxi¬  dation of a fatty acid molecule is therefore a highly exergonic process that yields  numerous ATPs. For example, oxidation of palmitoyl-CoA (which has a C 1( 5  fatty acyl group) involves seven rounds of (3 oxidation, yielding 7 FADH 2 , 7  NADH, and 8 acetyl-CoA. Oxidation of the 8 acetyl-CoA, in turn, yields 8  GTP, 24 NADH, and 8 FADH 2 . Since oxidative phosphorylation of the 31  NADH molecules yields 77.5 ATP and that of the 15 FADH 2 yields 22.5 ATP,  subtracting the 2 ATP equivalents required for fatty acyl-CoA formation (Section  20-2A), the oxidation of one palmitate molecule has a net yield of 106 ATP.   Almost all unsaturated fatty acids of biological origin contain only cis double  bonds, which most often begin between C9 and CIO (referred to as a A 9 or  9-double bond; Table 9-1). Additional double bonds, if any, occur at three-  carbon intervals and are therefore never conjugated. Two examples of unsatu¬  rated fatty acids are oleic acid and linoleic acid.    FIG. 20-14 Mechanism of action of  p-ketoacyl-CoA thiolase. An active site Cys  residue participates in the formation of an  enzyme-thioester intermediate.    FIG. 20-1 5 The oxidation of unsaturated fatty acids. ( Opposite ) p oxidation of an  unsaturated fatty acid such as linoleic acid presents three problems. The first problem, the  presence of a p/y double bond, seen in the left-hand pathway, is solved by the bond’s  conversion to a trans-a,p double bond. The second problem in the left-hand pathway, that  a 2,4-dienoyl-CoA is a poor substrate for enoyl-CoA hydratase, is eliminated by the  NADPH-dependent reduction of the A 4 bond by 2,4-dienoyl-CoA reductase to yield the  p-oxidation substrate trans- 2-enoyl-CoA. This step requires one enzyme in E. coli and two  in mammals. The third problem, the isomerization of 2,5-dienoyl-CoA (originating from the  oxidation of unsaturated fatty acids with double bonds at odd-numbered C atoms) to  3,5-dienoyl-CoA, is solved by converting the 3,5-dienoyl-CoA to 2,4-dienoyl-CoA, a  substrate for 2,4-dienoyl-CoA reductase.    Note that one of the double bonds in linoleic acid is at an odd-numbered car¬  bon atom and the other is at an even-numbered carbon atom. The double bonds  in fatty acids such as linoleic acid pose three problems for the (3-oxidation path¬  way that are solved through the actions of four additional enzymes (Fig. 20-15).   Problem Is A (3,y Double Bond. The first enzymatic difficulty occurs  after the third round of (3 oxidation: The resulting cis-(3,y  double bond-containing enoyl-CoA is not a substrate for  enoyl-CoA hydratase. Enoyl-CoA isomerase, however, con¬  verts the cis-A 3 double bond to the trans-A 2 form. The A 2  compound is the normal substrate of enoyl-CoA hydratase,  so (3 oxidation can continue.   Problem 2: A A 4 Double Bond Inhibits Enoyl-CoA Hydratase. The   next difficulty arises in the fifth round of (3 oxidation: The  presence of a double bond at an even-numbered carbon atom  results in the formation of 2,4-dienoyl-CoA, which is a poor  substrate for enoyl-CoA hydratase. However, NADPH-  dependent 2,4-dienoyl-CoA reductase reduces the A 4 double  bond. The E. coli reductase produces trans- 2-enoyl-CoA, a  normal substrate of (3 oxidation. The mammalian reductase,  however, yields trans-3-c noyl-CoA, which, to proceed along  the (3-oxidation pathway, must first be isomerized to trans- 2-  enoyl-CoA by 3,2-enoyl-CoA isomerase.   Problem 3: The Unanticipated Isomerization of 2,5-Enoyl-CoA by  3,2-Enoyl-CoA Isomerase. Mammalian 3,2-enoyl-CoA isom¬  erase catalyzes a reversible reaction that interconverts A 2  and A 3 double bonds. A carbonyl group is stabilized by be¬  ing conjugated to a A 2 double bond. However, the presence  of a A 5 double bond (originating from an unsaturated fatty  acid with a double bond at an odd-numbered C atom such  as the A 9 double bond of linoleic acid) is likewise stabilized  by being conjugated with a A 3 double bond. If a 2,5-enoyl-  CoA is converted by 3,2-enoyl-CoA isomerase to 3,5-enoyl  CoA, which occurs up to 20% of the time, another enzyme  is necessary to continue the oxidation: 3,5-2,4-Dienoyl-  CoA isomerase isomerizes the 3,5 diene to a 2,4 diene,  which is then reduced by 2,4-dienoyl-CoA reductase and  isomerized by 3,2-enoyl-CoA isomerase as in Problem 2 above.  After two more rounds of (3 oxidation, the cis- A 4 double  bond originating from the cis-A 12 double bond of linoleic  acid is also dealt with as in Problem 2.   Most fatty acids, for reasons explained in Section 20-4, have even numbers of  carbon atoms and are therefore completely converted to acetyl-CoA. Some    plants and marine organisms, however, synthesize fatty acids with an odd num¬  ber of carbon atoms. The final round of (3 oxidation of these fatty acids yields  propionyl-CoA , which is converted to succinyl-CoA for entry into the citric acid  cycle. Propionate and propionyl-CoA are also produced by the oxidation of the  amino acids isoleucine, valine, and methionine (Section 21-4D).    a biotin prosthetic group and is driven by the hydrolysis of ATP to  ADP + P r The reaction resembles that of pyruvate carboxylase  (Fig. 16-18).   The (S)-methylmalonyl-CoA product of the carboxylase reaction is  converted to the R form by methylmalonyl-CoA racemase.   (W)-Methylmalonyl-CoA is a substrate for methylmalonyl-CoA  mutase, which catalyzes an unusual carbon skeleton rearrangement.    Methylmalonyl-CoA mutase uses a 5'-deoxyadenosylcobalamin prosthetic  group (AdoCbl; also called coenzyme B 12 , a derivative of cobalamin, or  vitamin B 12 ; Box 20-1). Dorothy Hodgkin determined the structure of this  complex molecule (Fig. 20-17) in 1956 through X-ray crystallographic analy¬  sis combined with chemical degradation studies, a landmark achievement  (Box 20-2).   5 r -Deoxyadenosylcobalamin contains a hemelike corrin ring whose four  pyrrole N atoms each ligand a six-coordinate Co ion. The fifth Co ligand is  an N atom of a 5,6-dimethylbenzimidazole (DMB) nucleotide that is cova¬  lently linked to the corrin D ring. The sixth ligand is a 5 r -deoxyadenosyl group    The existence of vitamin B 12 came to light in 1926 when George  Minot and William Murphy discovered that pernicious anemia,  an often fatal disease of the elderly characterized by decreased  numbers of red blood cells, low hemoglobin levels, and progressive  neurological deterioration, can be treated by the daily consumption  of large amounts of raw liver. Nevertheless, the antipernicious  anemia factor—vitamin B 12 —was not isolated until 1948.   Vitamin B 12 is synthesized by neither plants nor animals but  only by a few species of bacteria. Herbivores obtain their vitamin  B 12 from the bacteria that inhabit their gut (in fact, some animals,  such as rabbits, must periodically eat some of their feces to  obtain sufficient amounts of this essential substance). Humans,  however, obtain almost all their vitamin B 12 directly from their diet,  particularly from meat. In the intestine, the glycoprotein intrinsic    factor, which is secreted by the stomach, specifically binds  vitamin B 12 , and the protein-vitamin complex is absorbed via a  receptor in the intestinal mucosa. The complex dissociates and  the liberated vitamin B 12 is transported to the bloodstream. At  least three different plasma proteins, called transcobalamins,  bind the vitamin and facilitate its uptake by the tissues.   Pernicious anemia is not usually a dietary deficiency disease  but, rather, results from insufficient secretion of intrinsic factor,  often due to an autoimmune attack against the cells, that produce  it. The normal human requirement for cobalamin is very small,   ~3 |xg • day -1 , and the liver stores a 3- to 5-year supply of the  vitamin. This accounts for the insidious onset of pernicious anemia  and the fact that true dietary deficiency of vitamin B 12 , even  among strict vegetarians, is extremely rare.    in which the deoxyribose C5' atom forms a covalent C—Co bond, one of only  two carbon—metal bonds known in biology (the other is a C—Ni bond in the  bacterial enzyme carbon monoxide dehydrogenase). In some cobalamin-  dependent enzymes, the sixth ligand instead is a CH 3 group that likewise forms  a C—Co bond. There are only about a dozen known cobalamin-dependent  enzymes, which catalyze molecular rearrangements or methyl-group transfer  reactions.   Methylmalonyl-CoA Mutase Has a Unique a/p Barrel. The X-ray structure  of methylmalonyl-CoA mutase from Propionibacterium shermanii, an ap het¬  erodimer, in complex with the substrate analog 2-carboxypropyl-CoA (which  lacks methylmalonyl-CoAs thioester oxygen atom) was determined by Philip  Evans. Its AdoCbl cofactor is sandwiched between the catalytically active  a subunits two domains: a 559-residue N-terminal a/p barrel (TIM barrel,    Dorothy Crowfoot Hodgkin (1910—  1994) The third woman to receive a  Nobel Prize in Chemistry, after Marie  Curie in 1911 and her daughter Irene  Joliot-Curie in 1935, was Dorothy  Crowfoot Hodgkin, who received the  award in 1964 for determining the  structures of biological molecules by  X-ray crystallography. Hodgkin’s career  as a scientist is notable not only for her  accomplishments-elucidating the  structures of sterols, penicillin, vitamin  B 12 , and insulin—but also for her contributions to the methodology  of X-ray crystallography.   Hodgkin’s fascination with crystals reportedly began at age 10  with a simple experiment involving alum. Her scientific tendencies  were encouraged by her parents, particularly her mother, who was  an accomplished amateur botanist. Hodgkin attributed her  independent spirit to the fact that her parents were absent for  long periods, as her father worked as an archaeologist in Egypt  and the Middle East.   Hodgkin’s formal training in crystallography took place at  Oxford during her undergraduate years and continued as she  earned a doctorate under the guidance of J.D. Bernal at  Cambridge. Unlike many of his colleagues at the time, Bernal was  generous in giving credit to his junior associates. Consequently,  Hodgkin received recognition along with Bernal in 1934 when  they reported the first X-ray diffraction pattern of a protein,  pepsin. Bernal and Hodgkin had thereby shown that proteins are  not amorphous colloids but have discrete structures. They also  advanced the field of macromolecular crystallography by noting  the necessity of examining crystals surrounded by their mother  liquor (the solution from which the molecules crystallize) rather  than air-dried, as was standard procedure at the time.   After returning to Oxford, Hodgkin established her own  laboratory, a challenging task for someone striking out in a new  field, and even more difficult for a woman trying to balance work  with marriage and motherhood in the late 1930s. At Oxford,  Hodgkin undertook studies of crystallized cholesterol and other  steroids, showing that calculations based on X-ray diffraction in  three rather than two dimensions could disclose considerable  information about molecular structure, including the stereochemistry  of each carbon atom. Hodgkin also began studying crystals of  insulin, although an atomic model of the 777-atom protein took  34 years to complete (the sequence of insulin, determined by  Frederick Sanger, was not known until 1955; Box 5-1). During  World War II, Hodgkin turned her attention to the structure of  penicillin and used X-ray crystallography to elucidate its  unexpected ring structure (three carbons and a nitrogen;   Box 8-3). This discovery helped pave the way for the synthesis of    penicillin and its derivatives. Hodgkin’s work also marked the  beginning of an electronic age in biochemical research, as she  used one of the first analog computers from IBM to help with the  required calculations.   In 1955, a visitor brought Hodgkin crystals of cyanocobalamin,  whose deep-red color may have enticed Hodgkin to immediately  examine their X-ray diffraction. The cobalamin structure, four times  larger than that of penicillin, had not yet yielded its structural  secrets to conventional chemical approaches. Hodgkin’s success  in determining the structure of cobalamin was the result of  experience as well as intuition. She was reportedly able to discern  features of molecular structure simply by examining the diffraction  pattern. However, she also took advantage of powerful computers  available to her collaborator Kenneth Trueblood at the University  of California at Los Angeles, who, due to the great distance  between them, communicated with her through letters and  telegrams. Hodgkin bravely decided to use the cobalt atom  naturally present in cobalamin to solve the “phase problem” that  prevented the straightforward determination of X-ray structures  (Box 7-2). Although others believed that the cobalt atom would  not scatter X-rays strongly enough relative to the lighter atoms in  the structure, Hodgkin was able to derive the necessary phase  information and thus was spared having to produce cobalamin  crystals containing a heavier atom.   Hodgkin and her collaborators identified the unusual porphyrin  structure of cyanocobalamin, in which rings A and D are directly  linked and ring A is fully saturated (Fig. 20-17). They  subsequently identified cobalamin, with its Co—C bond, as the  first known biological organometallic compound. Hodgkin later  determined the structure of the physiological form of vitamin B 12 ,  adenosylcobalamin, in 1961. Now that the structure of the vitamin  was known, it could be synthesized and used to treat pernicious  anemia (Box 20-1). In addition to its obvious implications for  human health, Hodgkin’s work with cobalamin encouraged other  researchers to pursue the structures of large compounds  previously thought to be too complicated for crystallographic  analysis. Hodgkin herself finally completed a three-dimensional  model of insulin in 1969.   After receiving the Nobel prize in 1964, along with many other  honors, Hodgkin became increasingly involved in international  organizations, including the International Union of Crystallography  and other groups dedicated to promoting scientific cooperation as  a means for decreasing international tensions during the Cold  War. Her own laboratory over the years hosted scientists from  around the globe. By the end of her life, her efforts to foster  international goodwill were no less notable than her  accomplishments as a crystallographer.   Ferry, G., Dorothy Hodgkin: A Life, Cold Spring Harbor Laboratory Press (2000).  Hodgkin, D.C., Kamper, J., Mackay, M., Pickworth, J., Trueblood, K.N. and White, J.G.,  Structure of vitamin B-12, Nature 178 , 64 (1956).   the most common enzymatic motif; Sections 6-2C and 15-2) and a 169-residue  C-terminal a/(3 domain that resembles a Rossmann fold (Section 6-2C). The  structure of the a/(3 barrel contains several surprising features (Fig. 20-18):   1. The active sites of nearly all a/(3 barrel enzymes are located at the  C-terminal ends of the barrels (3 strands. In methylmalonyl-CoA mutase,   FIG. 20-1 8 X-Ray structure of P. shermanii methylmalonyl-CoA mutase in  complex with 2-carboxypropyl-CoA and AdoCbl. (a) The catalytically active a subunit in  which the N-terminal domain is cyan, the |3 strands of its a/|3 barrel are orange, and the  C-terminal domain is pink. The substrate analog 2-carboxypropyl-CoA ( magenta ) and  AdoCbl (green) are drawn in space-filling form. The 2-carboxypropyl-CoA passes through  the center of the a/|3 barrel and is oriented such that the methylmalonyl group of  methylmalonyl-CoA would contact the corrin ring of the AdoCbl, which is sandwiched  between the enzyme’s N- and C-terminal domains, (b) The arrangement of the AdoCbl and  2-carboxypropyl-CoA molecules which, together with the side chain of His 610, are  represented in stick form colored according to atom type (2-carboxypropyl-CoA and His C  green, AdoCbl C cyan, N blue, 0 red, P magenta, and S yellow). The corrin ring’s Co atom  is represented by a lavender sphere and the a/p barrel’s p strands are represented by  orange ribbons. The view is similar to that in Part a. Note that the DMB group (bottom) has  swung away from the corrin ring (seen edgewise) to be replaced by the side chain of His  610 from the C-terminal domain and that the 5'-deoxyadenosyl group is unseen (due to  disorder). [Based on an X-ray structure by Philip Evans, MRC Laboratory of Molecular  Biology, Cambridge, U.K. PDBid 7REQ.] Q See Interactive Exercise 28.    however, the AdoCbl is packed against the N-terminal ends of the bar-  rels P strands.   2. In free AdoCbl, the Co atom is axially liganded by an N atom of its  DMB group and by the adenosyl residues 5'-CH 2 group (Fig. 20-17).  In the enzyme, however, the DMB has swung aside to bind in a sep¬  arate pocket and has been replaced by the side chain of His 610 from  the C-terminal domain. The adenosyl group is not visible in the struc¬  ture due to disorder and hence has probably also swung aside.   3. In nearly all other a/p barrel—containing enzymes, the center of the  barrel is occluded by large, often branched, hydrophobic side chains.  In methylmalonyl-CoA mutase, however, the 2-carboxypropyl-CoAs  pantetheine group binds in a narrow tunnel along the center of the  a/|3 barrel so as to put the methylmalonyl group of an intact substrate  in close proximity to the unliganded face of the cobalamin ring. The  tunnel provides the only direct access to the active site cavity, thereby  protecting the reactive free radical intermediates that are produced in  the catalytic reaction from side reactions (see below). The tunnel is  lined by small hydrophilic residues (Ser and Thr).    Methylmalonyl-CoA mutase’s substrate-binding mode resembles that of sev¬  eral other AdoCbl-containing enzymes of known structure, which are collec¬  tively unique among a /(3 barrel-containing enzymes.   Methylmalonyl-CoA Mutase Stabilizes and Protects Free Radical  Intermediates. The proposed methylmalonyl-CoA mutase reaction mechanism  (Fig. 20-19) begins with the homolytic cleavage of the cobalamin C—Co  bond; in other words, the C and Co atoms each acquire one of the electrons  that formed the cleaved electron pair bond. (Note that a homolytic cleavage    FIG. 20-19 Proposed mechanism of methylmalonyl-CoA  mutase. (1) The homolytic cleavage of the C—Co(lll) bond yields a  5'-deoxyadenosyl (Ado) radical and cobalamin in its Co(ll) oxidation  state. (2) The 5'-deoxyadenosyl radical abstracts a hydrogen atom  from methylmalonyl-CoA, thereby generating a methylmalonyl-CoA    radical. (3) Carbon skeleton rearrangement yields a succinyl-CoA  radical via a proposed cyclopropyloxy radical intermediate.   (4) The succinyl-CoA radical abstracts a hydrogen atom from  5'-deoxyadenosine to regenerate the 5'-deoxyadenosyl radical.   reaction is unusual in biology; most other biological bond-cleavage reactions  occur via heterolytic cleavage in which the electron pair forming the cleaved  bond is fully acquired by one of the separating atoms.) The Co ion therefore  alternates between its Co (III) and Co (II) oxidation states and hence functions  as a reversible free radical generator. The C—Co (III) bond is well suited to this  function because it is inherently weak (dissociation energy 109 kj • mol X )  and is further weakened through steric interactions with the enzyme.   Spectroscopic measurements indicate that the Co atom in methylmalonyl-  CoA mutase is in the Co(II) state, thereby confirming that it has no sixth lig¬  and (as occurs during its catalytic cycle; Fig. 20-19). Protein-induced strain  makes the His N—Co bond extremely long (2.5 A versus 1 . 9 — 2.0 A in vari¬  ous other B 12 -containing structures), stabilizing the Co(II) species relative to  Co (III) and thereby favoring the formation of the adenosyl radical. The rad¬  ical abstracts a hydrogen atom from the methylmalonyl-CoA substrate, thereby  facilitating its rearrangement to succinyl-CoA through the intermediate for¬  mation of a cyclopropyloxy radical.   Methylmalonyl-CoA mutase catalyzes the conversion of a metabolite to a cit¬  ric acid cycle intermediate other than acetyl-Co A. However, such C 4 interme¬  diates are actually catalysts, not substrates, of the citric acid cycle. In order for  succinyl-CoA to undergo net oxidation by the citric acid cycle, it must first be con¬  verted to pyruvate and then to acetyl-CoA. This is accomplished by converting  succinyl-CoA to malate (Reactions 5-7 of the citric acid cycle; Fig. 17-2) fol¬  lowed by its transport to the cytosol and oxidative decarboxylation to pyru¬  vate and C0 2 by malic enzyme    (this enzyme also functions in the C 4 cycle of photosynthesis; Fig. 19-33).  Pyruvate is then completely oxidized via pyruvate dehydrogenase and the cit¬  ric acid cycle.    In mammalian cells, the bulk of (3 oxidation occurs in the mitochondria, but  peroxisomes (Fig. 20-20) also oxidize fatty acids, particularly those with very  long chains or branched chains. In plants, fatty acid oxidation occurs exclu¬  sively in the peroxisomes and glyoxysomes (which are specialized peroxisomes).  In addition to lipid catabolism, mammalian peroxisomes participate in the syn¬  thesis of certain lipids, including bile acids. A variety of human diseases result  from defects in peroxisomal enzymes or proteins that transport intermediates  across the peroxisomal membrane.   Peroxisomal (3 oxidation , which differs only slightly from mitochondrial  (3 oxidation ., shortens very long chain fatty acids (>22 carbon atoms), which are  then fully degraded by the mitochondrial pathway Very long chain fatty acids  are transported into the peroxisomes by a mechanism that does not require   FIG. 20-20 Electron micrograph of a peroxisome from rat liver surrounded by  the rough endoplasmic reticulum. Peroxisomes are membrane-bounded organelles that  perform a variety of metabolic functions, including the oxidation of very long chain fatty acids.  [Donald W. Fawcett/Photo Researchers Inc.]   • Describe the activation of fatty acids.  What is the energy cost for the process?   • How do cytosolic acyl groups enter the  mitochondrion for degradation?   • Summarize the chemical reactions that  occur in each round of p oxidation.   • How is ATP recovered from the products  of p oxidation?   • What additional steps are required to  oxidized unsaturated and odd-chain fatty  acids?   • What is unusual about the methylmalonyl-  CoA mutase structure and the reaction it  catalyzes?   • Describe the differences between peroxi¬  somal and mitochondrial p oxidation.    carnitine, and are activated by a long chain acyl-CoA synthetase. Peroxisomal  (3 oxidation results in the same chemical changes to fatty acids as in the  mitochondrial pathway but requires only three enzymes:   The enzyme uses an FAD cofactor, but the abstracted electrons are  transferred directly to 0 2 rather than passing through the electron-  transport chain with its concomitant oxidative phosphorylation  (Fig. 20-12, Reactions 5-8). Peroxisomal fatty acid oxidation therefore  generates 1.5 fewer ATP per C 2 cycle than mitochondrial fatty acid  oxidation. Catalase converts the H 2 0 2 produced in the oxidase reaction  to H 2 0 + 0 2 .   2. Peroxisomal enoyl-CoA hydratase and 3-L-hydroxyacyl-CoA dehydro¬  genase activities occur on a single polypeptide. The reactions catalyzed  are identical to those of the mitochondrial system (Fig. 20-12,  Reactions 2 and 3).   3. Peroxisomal thiolase catalyzes the final step of oxidation. This enzyme  is almost inactive with acyl-CoAs of length C 8 or less, so peroxisomes  incompletely oxidize fatty acids.   The peroxisome contains both a carnitine acetyltransferase and a transferase  specific for longer chain acyl groups. Acyl-CoAs that have been chain-shortened  by peroxisomal (3 oxidation are thereby converted to their carnitine esters.  These substances, for the most part, passively diffuse out of the peroxisome  to the mitochondrion, where they are oxidized further.    Acetyl-CoA may be reversibly converted to ketone bodies in the liver to be used  as fuel by other tissues.    The acetyl-CoA produced by oxidation of fatty acids can be further oxidized  via the citric acid cycle. In liver mitochondria, however, a significant fraction  of this acetyl-CoA has another fate. By a process known as ketogenesis, acetyl-  CoA is converted to acetoacetate or D-(3-hydroxybutyrate. These compounds  together with acetone are somewhat inaccurately referred to as ketone bodies:    Ketone bodies are important metabolic fuels for many peripheral tissues, par¬  ticularly heart and skeletal muscle. The brain, under normal circumstances,  uses only glucose as its energy source (fatty acids are unable to pass through  the blood-brain barrier), but during starvation, the small, water-soluble ke¬  tone bodies become the brains major fuel source (Section 22-4A).   1 . Two molecules of acetyl-CoA are condensed to acetoacetyl-CoA by  thiolase (also called acetyl-CoA acetyltransferase) working in the re¬  verse direction from the way it does in the final step of (3 oxidation  (Fig. 20-12, Reaction 4).    FIG. 20-21 Ketogenesis. Acetoacetate is  formed from acetyl-CoA in three steps. (1) Two  molecules of acetyl-CoA condense to form  acetoacetyl-CoA. (2) A Claisen ester  condensation of the acetoacetyl-CoA with a third  acetyl-CoA forms p-hydroxy-p-methylglutaryl-CoA  (HMG-CoA). (3) HMG-CoA is degraded to  acetoacetate and acetyl-CoA in a mixed  aldol-Claisen ester cleavage.    2 . Condensation of the acetoacetyl-CoA with a third acetyl-CoA by  HMG-CoA synthase forms P-hydroxy-P-methylglutaryl-CoA  (HMG-CoA). The mechanism of this reaction resembles the reverse of  the thiolase reaction (Fig. 20-14) in that an active site thiol group  forms an acyl-thioester intermediate.   3. HMG-CoA is degraded to acetoacetate and acetyl-CoA in a mixed  aldol-Claisen ester cleavage by HMG-CoA lyase. The mechanism of  this reaction is analogous to the reverse of the citrate synthase reaction  (Fig. 17-10). HMG-CoA is also a precursor in cholesterol biosynthesis  (Section 20 - 7 A). HMG-CoA lyase is present only in liver mitochon¬  dria and therefore does not interfere with cholesterol synthesis in the  cytoplasm.    Acetoacetate, a P-keto acid, also undergoes relatively facile nonenzymatic de¬  carboxylation to acetone and C0 2 . Indeed, in individuals with ketosis, a  pathological condition in which acetoacetate is produced faster than it is me¬  tabolized (a symptom of diabetes; Section 22-4B), the breath has the charac¬  teristic sweet smell of acetone.   The liver releases acetoacetate and P-hydroxybutyrate, which are carried  by the bloodstream to the peripheral tissues for use as alternative fuels. There,    Compare this pathway to the ketogenesis  pathway. Which steps are similar?    these products are converted to two acetyl-Co A as is diagrammed in Fig. 20-22.  Succinyl-CoA, which acts as the CoA donor in this process, can also be con¬  verted to succinate with the coupled synthesis of GTP in the succinyl-CoA  synthetase reaction of the citric acid cycle (Section 17-3E). The “activation”  of acetoacetate bypasses this step and therefore “costs” the free energy of GTP  hydrolysis.    • The tricarboxylate transport system transfers acetyl-CoA into the cytosol for fatty  acid synthesis.   • Fatty acid synthesis begins with the carboxylation of acetyl-CoA to generate malonyl-  CoA.   • Fatty acid synthase carries out seven reactions and lengthens a fatty acid two  carbons at a time.    Fatty acid biosynthesis occurs through condensation of C 2 units, the reverse of the  (3 -oxidation process. Through isotopic labeling techniques, David Rittenberg  and Konrad Bloch demonstrated, in 1945, that these condensation units are  derived from acetic acid. Subsequent research showed that both acetyl-CoA    FIG. 20-23 Comparison of fatty acid p oxidation and fatty hydration/dehydration reaction, and (5) the form in which C 2 units are    and bicarbonate are required, and that a C 3 unit, malonyl-CoA, is an inter¬  mediate of fatty acid biosynthesis.   The pathway of fatty acid synthesis differs from that of fatty acid oxida¬  tion. This situation, as we saw for glycogen metabolism in Section 16-3, is  typical of opposing biosynthetic and degradative pathways because it permits  them both to be thermodynamically favorable and independently regulated  under similar physiological conditions. Figure 20-23 outlines fatty acid oxi¬  dation and synthesis with emphasis on the differences between the pathways,  including the cellular locations of the pathways, the redox coenzymes, and the  manner in which C 2 units are removed or added to the fatty acyl chain.   Acetyl-CoA, the starting material for fatty acid synthesis, is generated in the  mitochondrion by the oxidative decarboxylation of pyruvate as catalyzed by  pyruvate dehydrogenase (Section 17-2B) as well as by the oxidation of fatty  acids. When the demand for ATP is low, so that the oxidation of acetyl-CoA  via the citric acid cycle and oxidative phosphorylation is minimal, this mito¬  chondrial acetyl-CoA may be stored for future use as fat. Fatty acid biosyn¬  thesis occurs in the cytosol, however, and the mitochondrial membrane is  essentially impermeable to acetyl-CoA. Acetyl-CoA enters the cytosol in the form  of citrate via the tricarboxylate transport system (Fig. 20-24). ATP-citrate  lyase then catalyzes the reaction   Citrate + CoA + ATP —> acetyl-CoA + oxaloacetate + ADP + P z   which resembles the reverse of the citrate synthase reaction (Fig. 17-10) ex¬  cept that ATP hydrolysis is required to drive the synthesis of the thioester  bond. Oxaloacetate is then reduced to malate by malate dehydrogenase. Malate  is oxidatively decarboxylated to pyruvate by malic enzyme and returned in this  form to the mitochondrion. This reaction involves the reoxidation of malate    FIG. 20-24 The tricarboxylate transport  system. The overall effect of this sequence of  reactions is the transfer of acetyl groups from  the mitochondrion to the cytosol.    to oxaloacetate, a (3-keto acid, which is then decarboxylated, a reaction remi¬  niscent of the isocitrate dehydrogenase reaction in the citric acid cycle (Section  17-3C). The NADPH produced is used in the reductive reactions of fatty acid  biosynthesis.   Acetyl-CoA carboxylase (ACC) catalyzes the first committed step of fatty acid  biosynthesis and one of its rate-controlling steps. The mechanism of this  biotin-dependent enzyme is similar to those of propionyl-CoA carboxylase  (Section 20-2E) and pyruvate carboxylase (Fig. 16-18). The reaction occurs in  two steps, a C0 2 activation and a carboxylation:    The result is a three-carbon (malonyl) group linked as a thioester to CoA.    Mammalian acetyl-CoA carboxylase, a 230-kD polypeptide, is subject to  allosteric and hormonal control. For example, citrate stimulates acetyl-CoA car¬  boxylase and long chain fatty acyl-CoAs are feedback inhibitors of the enzyme.  Fine-tuning of enzyme activity is accomplished through covalent modification.  Acetyl-CoA carboxylase is a substrate for several kinases. It has six phosphory¬  lation sites, but phosphorylation of only one (Ser 79) is clearly associated with  enzyme inactivation. Ser 79 is phosphorylated by AMP-dependent protein ki¬  nase (AMPK) in a cAMP-independent pathway. However, glucagon as well  as epinephrine, which act through protein kinase A (PKA; Section 16-3B), pro¬  mote the phosphorylation of Ser 79, possibly by inhibiting its dephosphoryla¬  tion (recall that this occurs in glycogen metabolism when the PKA-mediated  phosphorylation of phosphoprotein phosphatase inhibitor-1 inhibits dephos¬  phorylation; Fig. 16-13). Insulin, on the other hand, stimulates dephosphory¬  lation of acetyl-CoA carboxylase and thereby activates the enzyme.   Mammalian Acetyl-CoA Carboxylase Has Two Major Isoforms. There are  two major isoforms of ACC. ACC1 occurs in adipose tissue and ACC2 occurs  in tissues that oxidize but do not synthesize fatty acids, such as heart muscle.  Tissues that both synthesize and oxidize fatty acids, such as liver, contain both  isoforms, which are homologous although the genes encoding them are located  on different chromosomes. What is the function of ACC2? The product of the  ACC-catalyzed reaction, malonyl-CoA, strongly inhibits the mitochondrial im¬  port of fatty acyl-CoA for fatty acid oxidation, the major control point for this  process. Thus it appears that ACC2 has a regulatory function (Section 20-5).   The E. coli acetyl-CoA carboxylase, which is a multisubunit protein, is reg¬  ulated by guanine nucleotides so that fatty acid synthesis is coordinated with  cell growth. In prokaryotes, fatty acids serve primarily as phospholipid precur¬  sors, since these organisms do not synthesize triacylglycerols for energy storage.   The synthesis of fatty acids, mainly palmitic acid, from acetyl-CoA and  malonyl-CoA involves seven enzymatic reactions. These reactions were first  studied in cell-free extracts of E. coli, in which they are catalyzed by independ¬  ent enzymes. Individual enzymes with these activities also occur in chloroplasts  (plant fatty acid synthesis does not occur in the cytosol). In yeast, fatty acid  synthase is a cytosolic, 2500-kD multifunctional enzyme with the composi¬  tion a^(3(5, whereas in animals it is a 534-kD multifunctional enzyme consist¬  ing of two identical polypeptide chains. Presumably, such proteins evolved by  the joining of previously independent genes for the enzymes.   Although fatty acid synthesis begins with the synthesis of a CoA ester,  malonyl-CoA, the growing fatty acid is anchored to acyl-carrier protein  (ACP; Fig. 20-25) . ACP, like CoA, contains a phosphopantetheine group that   FIG. 20-25 The phosphopantetheine group in acyl-carrier protein (ACP) and in CoA.    forms a thioester with an acyl group. The phosphopantetheine phosphoryl  group is esterified with a Ser OH group of ACP, whereas in CoA it is linked  to AMP. In E. coli, ACP is a 10-kD polypeptide, whereas in animals it is part  of the multifunctional fatty acid synthase.   The reactions catalyzed by mammalian fatty acid synthase are diagrammed  in Fig. 20-26:   1 . These are priming reactions in which the synthase is “loaded” with the  precursors for the condensation reaction. In mammals, malonyl/  acetyl-CoA-ACP transacylase (MAT) catalyzes two similar reactions  at a single active site: An acetyl group originally linked as a thioester  in acetyl-CoA is transferred to ACP (la), and a malonyl group is trans¬  ferred from malonyl-CoA to ACP (lb).   2. The (3-ketoacyl-ACP synthase (KS; also known as condensing  enzyme) first transfers the acetyl group from ACP to an enzyme Cys  residue (2a). In the condensation reaction (2b), the malonyl-ACP  is decarboxylated, and the resulting carbanion attacks the acetyl-  thioester to form a four-carbon acetoacetyl-ACP. The decarboxyla¬  tion reaction drives the condensation reaction:    3—5. Two reductions and a dehydration convert the (3-keto group to an  alkyl group. The coenzyme in both reductive steps is NADPH. In (3  oxidation, the analogs of Reactions 3 and 5, respectively, use NAD +  and FAD (Fig. 20-12, Reactions 3 and 1). Moreover, Reaction 4 re¬  quires a D-(3-hydroxyacyl substrate, whereas the analogous reaction in  (3 oxidation forms the corresponding L isomer.   At this point, the acyl group, originally an acetyl group, has been elon¬  gated by a C 2 unit. This butyryl group is then transferred from ACP to the  Cys—SH of the enzyme (a repeat of Reaction 2a) so that it can be extended  by additional rounds of the fatty acid synthase reaction sequence. Note that  the malonyl-CoA synthesized by the acetyl-CoA carboxylase reaction is decar¬  boxylated in the condensation reaction. The formation of a C—C bond is an  endergonic process requiring an activated precursor. Malonyl-ACP is a (3-keto  ester whose exergonic decarboxylation yields the acetyl-ACP carbanion    FIG. 20-26 Reaction sequence for the biosynthesis of fatty acids. In forming palmitate, the pathway is  repeated for seven cycles of C 2 elongation followed by a final hydrolysis step. Q See the Animated Figures.   Chapter 20 Lipid Metabolism required for C — C bond formation. The free energy required for the overall   reaction is supplied by the ATP hydrolysis in the acetyl-CoA carboxylase re¬  action, which generates the malonyl-CoA precursor of malonyl-ACP. This car-  boxylation—decarboxylation sequence is similar to the activation of pyruvate to  oxaloacetate for conversion to phosphoenolpyruvate in gluconeogenesis  (Section 16-4A).   In each reaction cycle, the ACP is “reloaded” with a malonyl group, and  the acyl chain grows by two carbon atoms. Note that each new acetyl unit do¬  nated by malonyl-CoA adds to the growing acyl chain at its point of attach¬  ment to the enzyme. Thus, the fatty acid grows from its thioester end, not  from its methyl end (likewise, fatty acids are catabolized from their thioester  ends).   Seven cycles of elongation are required to form palmitoyl-ACP. The  thioester bond is then hydrolyzed by palmitoyl thioesterase (TE; Fig. 20-26,  Reaction 6), yielding palmitate, the normal product of the fatty acid synthase  pathway, and regenerating the enzyme for a new round of synthesis.   Acetyl-CoA + 7 malonyl-CoA +14 NADPH + 7 H + —>   palmitate + 7 C0 2 + 14 NADP + + 8 CoA + 6 H 2 0   7Acetyl-CoA + 7 C0 2 + 7 ATP -> 7 malonyl-CoA + 7 ADP + 7 P z + 7 H +   palmitate +14 NADP + + 8 CoA + 6 H 2 0 + 7 ADP + 7 P z   The Two Halves of Fatty Acid Synthase Operate in Concert. In animals, the  seven reactions of fatty acid synthesis are localized to six discrete active sites  (MAT carries out two reactions, la and lb). The order of the domains along  the animal fatty acid synthase polypeptide chain is indicated in Fig. 20-27/z.  Three of the domains besides ACP lack enzymatic activity: a linker domain  (LD) that bridges the KS and MAT domains; a pseudo-methyltransferase  OPME) domain, so named because it is homologous to the methyltransferase  family; and a pseudo-ketoreductase (M/XR) domain, so named because it is a  truncated form of the KR domain.   The two subunits of fatty acid synthase form an asymmetric X with two  reaction chambers (Fig. 20-27 B). The subunits associate via an extensive in¬  terface involving the KS, ER, and DH domains. However the upper and lower  portions of the X are only loosely connected. The lower portion of the X con¬  tains the enzymes two condensing activities, MAT and KS (which catalyze re¬  actions 1 and 2 in Fig. 20-26), and the upper portion of the X contains the  enzymes (3-carbon modifying activities, DH, KR, and ER (which catalyze re¬  actions 3-5 in Fig. 20-26). The C-terminal ACP and TE domains are flexi¬  bly tethered to the enzyme and are therefore unobserved.   Each of the two reaction chambers of the enzyme is lined by the full set  of catalytic domains from one subunit, all of which must be visited by the  long, flexible phosphopantetheine arm of an ACP. [The flexible lipoyllysyl  arms of the pyruvate dehydrogenase multienzyme complex (Section 17-2B)  perform a similar function in that enzyme.] The catalytic sites of the fatty acid  synthase domains occur on both the front and back faces of the protein, and  their distribution has no discernable relationship to the order of the reactions  in the catalytic cycle. Mutational studies indicate that a single ACP domain  can service active sites on both subunits. Apparently, the upper portion of the  X can rotate 180° relative to the lower portion, a degree of flexibility that is   FIG. 20-27 Structure of mammalian fatty acid synthase, (a) Domain organization  of porcine fatty acid synthase. The abbreviations for each enzyme activity are given in Fig.  20-26. (b) X-Ray structure of porcine fatty acid synthase. The ~190-A-wide homodimer is  viewed perpendicular to its pseudotwofold axis (vertical black arrow), with its various do¬  mains colored as in Part a. Bound NADP + cofactors are drawn in space-filling form in blue.  The attachment sites of the disordered ACP/TE domains are indicated by black dots. The  domain names for the second subunit include primes, (c) A schematic diagram of the  protein, indicating how the various domains are linked. [Courtesy of Timm Maier and Nenad  Ban, ETH Zurich, Switzerland. PDBid 2VZ9.]    consistent with the tenuous structure that joins the two halves of the enzyme.  Because fatty acid synthase is a dimer, two fatty acids can be synthesized si¬  multaneously. In some organisms, multifunctional enzymes that resemble fatty  acid synthase produce complex molecules (Box 20-3).   r ln well-nourished individuals, fatty acid synthesis proceeds at a low rate.   However, certain tissues, particularly malignancies, express high levels of  fatty acid synthase and produce fatty acids at a high rate. Consequently, in¬  hibitors of fatty acid synthase may act as anticancer agents.    Polyketides are a family of >10,000 diverse and structurally complex  natural products, many of which have antimicrobial, antitumor, and  immunosuppressive properties. They are synthesized in bacteria, fungi,  plants, and certain marine animals by the stepwise condensation of  acetyl-CoA or propionyl-CoA with malonyl-CoA or methylmalonyl-CoA  extender units. Each condensation reaction is driven by the decar¬  boxylation of the extender unit and yields a p-keto functional group  (e.g., an acetoacetyl group; this is why the ultimate product is called a  polyketide). Palmitate is an example of a polyketide since it is formed  by the condensation of one acetyl-CoA primer and seven malonyl-  CoA extender units. Following each condensation reaction, the new p-  keto group may be reduced, dehydrated, and reduced again as with  fatty acids, or may undergo only partial modification.   Polyketides are synthesized by multienzyme polypeptides that  resemble eukaryotic fatty acid synthase but are often much larger.  They include acyl-carrier domains plus an assortment of enzymatic  units that can function in an iterative fashion, repeating the same  elongation/modification steps (as in fatty acid synthesis), or in a  sequential fashion so as to generate more heterogeneous types  of polyketides.   For example, the soil bacterium Saccharopolyspora erythraea  produces the polyketide 6-deoxyerythronolide B (6dEB), which is  the precursor of the antibiotic erythromycin A. Synthesis requires a  propionyl-CoA primer and six (S)-methylmalonyl-CoA extenders. The  enzyme deoxyerythronolide B synthase (DEBS),  a 2000-kD a 2 p 272 complex, contains 28  different active sites that function like  an assembly line. Its three units  (DEBS1, DEBS2, and DEBS3)  each catalyze two elongation Loading,  and modification steps  (see the figure).    Module 4 resembles fatty acid synthase (it contains KS, AT,  ACP, KR, DH, and ER). However, it does not contain TE because  the elongation process is not complete after this phase. Module 3  contains only ACP, KS, and AT and passes its p-ketone conden¬  sation product to module 4 without further modification. Modules  1, 2, 5, and 6 contain only ACP, AT, KS, and KR, the sites neces¬  sary for the condensation and ketone reduction steps, thereby  generating hydroxy products. The overall organization of the mod¬  ules therefore creates a polyhydroxy chain with one keto group.  The chain cyclizes through reaction of the terminal hydroxyl group  with the thioester that anchored the growing chain to the syn¬  thase. As is the case for many polyketides, additional covalent  modifications are required to generate a product with full biologi¬  cal activity. The modular nature of polyketide synthases and the  variety of their reaction products make these enzymes appealing  to genetic engineers trying to design enzymes that can synthesize  novel compounds or more effective drugs.   An example of polyketide biosynthesis: the synthesis of erythromycin  A. [After Pfeifer, B.A., Admiraal, S.J., Gramajo, H., Cane, D.E., and  Khosla, C., Science 291, 1790 (2001).]    DEBS: Deoxyerythronolide B synthase  KS: Ketosynthase  AT: Acyltransferase  ACP: Acyl carrier protein  KR: Keto reductase   Palmitate, a saturated C 1( - fatty acid, is converted to longer chain saturated and  unsaturated fatty acids through the actions of elongases and desaturases.   Elongases are present in both the mitochondria and the endoplasmic reticulum,  but the mechanisms of elongation at the two sites differ. Mitochondrial elonga¬  tion (a process independent of the fatty acid synthase pathway in the cytosol)  occurs by the successive addition and reduction of acetyl units in a reversal of  fatty acid oxidation; the only chemical difference between the two pathways oc¬  curs in the final reduction step in which NAD PH takes the place of FADH 2 as  the terminal redox coenzyme (Fig. 20-28). Elongation in the endoplasmic retic¬  ulum involves the successive condensations of malonyl-CoA with acyl-CoA.   These reactions are each followed by NADPH-dependent reductions similar to  those catalyzed by fatty acid synthase, the only difference being that the fatty  acid is elongated as its CoA derivative rather than as its ACP derivative.   Mammalian systems contain four terminal desaturases of broad chain-length  specificities designated A 9 -, A 6 -, A 5 -, and A 4 -fatty acyl-CoA desaturases.   where x is at least five and where (CH 2 ) X can contain one or more double  bonds. The (CH 2 ) y portion of the substrate is always saturated. Double  bonds are inserted between existing double bonds in the (CH 2 ) X portion  of the substrate and the CoA group such that the new double bond is three  carbon atoms closer to the CoA group than the next double bond (not con¬  jugated to an existing double bond) and, in animals, never at positions be¬  yond C9.   T Certain Polyunsaturated Fatty Acids Must Be Obtained in the Diet.   A variety of unsaturated fatty acids can be synthesized by combinations  of elongation and desaturation reactions. However, since palmitic acid is  the shortest available fatty acid in animals, the above rules preclude the for¬  mation of the A 12 double bond of linoleic acid (A 9,12 -octadecadienoic acid;  a polyunsaturated fatty acid), a required precursor of prostaglandins and  other eicosanoids (Section 9-IF). Linoleic acid must consequently be ob¬  tained in the diet (ultimately from plants that have A 12 - and A 15 -desaturases)  and is therefore an essential fatty acid. Indeed, animals maintained on a  fat-free diet develop an ultimately fatal condition that is initially character¬  ized by poor growth, poor wound healing, and dermatitis. Linoleic acid is  also an important constituent of epidermal sphingolipids that function as  the skins water permeability barrier: Animals deprived of linoleic acid must  drink far more water than those with an adequate diet.    FIG. 20-28 Mitochondrial fatty acid elongation. This process is the reverse of  fatty acid oxidation (Fig. 20-12) except that the final reaction employs NADPH rather  than FADH 2 as its redox coenzyme.   Triacylglycerols are synthesized from fatty acyl-CoA esters and glycerol-3-  phosphate or dihydroxyacetone phosphate (Fig. 20-29) . The initial step in this  process is catalyzed either by glycerol-3-phosphate acyltransferase in mito¬  chondria and the endoplasmic reticulum, or by dihydroxyacetone phosphate  acyltransferase in the endoplasmic reticulum or peroxisomes. In the latter  case, the product acyl-dihydroxyacetone phosphate is reduced to the corre¬  sponding lysophosphatidic acid by an NADPH-dependent reductase. The  lysophosphatidic acid is converted to a triacylglycerol by the successive actions  of l-acylglycerol-3-phosphate acyltransferase, phosphatidic acid phos¬  phatase, and diacylglycerol acyltransferase. The intermediate phosphatidic  acid and 1 , 2 -diacylglycerol can also be converted to phospholipids by the path¬  ways described in Section 20 - 6 A. The acyltransferases are not completely spe¬  cific for particular fatty acyl-CoAs, either in chain length or degree of    unsaturation, but in human adipose tissue triacylglycerols, palmitate tends to  be concentrated at position 1 and oleate at position 2.   Glyceroneogenesis Is Important for Triacylglycerol Biosynthesis. The dihy-  droxyacetone phosphate used to make glycerol-3-phosphate for triacylglycerol syn¬  thesis comes either from glucose via the glycolytic pathway (Fig. 15-1) or from  oxaloacetate via an abbreviated version of gluconeogenesis (Fig. 16-15) termed  glyceroneogenesis. Glyceroneogenesis is necessary in times of starvation, since ap¬  proximately 30% of the fatty acids that enter the liver during a fast are re-esteri-  fied to triacylglycerol and exported in VLDL (Section 20-1). Adipocytes also carry  out glyceroneogenesis in times of starvation. They do not carry out gluconeogen¬  esis but contain the gluconeogenic enzyme phosphoenolpyruvate carboxykinase  (PEPCK), which is upregulated when the glucose concentration is low and par¬  ticipates in the glyceroneogenesis required for triacylglycerol biosynthesis.   At this point, we can appreciate how triacylglycerols synthesized from fatty  acids built from two-carbon acetyl units can be broken back down into acetyl  units. In the liver, the resulting acetyl-CoA may be shunted to the formation  of ketone bodies and later converted back to acetyl-CoA by another tissue.  The acetyl-Co A can then either be used to build fatty acids that are stored as  triacylglycerols or be oxidized by the citric acid cycle to generate considerable  ATP by oxidative phosphorylation. As we will see, the flux of material in the  direction of triacylglycerol synthesis or triacylglycerol degradation depends on  the metabolic energy needs of the organism and the need for synthesis of other  compounds, such as membrane lipids (Section 20-6) and cholesterol (Section  20-7A). These key features of lipid metabolism are summarized in Fig. 20-30.   • Compare the shuttle systems for trans¬  porting fatty acids into the mitochondria  and acetyl-CoA into the cytosol.   • What is the role of ATP-citrate lyase in  fatty acid synthesis?   • Describe the reactions that produce and  consume malonyl-CoA during fatty acid  synthesis.   • Why does it make metabolic sense for the  cell to regulate the activity of acetyl-CoA  carboxylase?   • Summarize the reactions catalyzed by fatty  acid synthase and compare them to the  reactions of fatty acid oxidation.   • Describe the role of multifunctional en¬  zymes in both fatty acid synthesis and  oxidation.   • What are the purposes of elongases and  desaturases? Where are the enzymes  located?   • Describe how newly synthesized fatty  acids are incorporated into triacylglycerols.    Fatty acid oxidation and synthesis are regulated by hormones and cellular factors.    Discussions of metabolic control are usually concerned with the regulation of  metabolite flow through a pathway in response to the differing energy needs  and dietary states of an organism. In mammals, glycogen and triacylglycerols   Chapter 20 Lipid Metabolism serve as primary fuels for energy-requiring processes and are synthesized in   times of plenty for future use. Synthesis and breakdown of glycogen and triacyl-  glycerols are processes that concern the whole organism ., with its organs and tissues  forming an interdependent network connected by the bloodstream. The blood car¬  ries the metabolites responsible for energy production: triacylglycerols in the  form of chylomicrons and VLDL, fatty acids as their albumin complexes, ke¬  tone bodies, amino acids, lactate, and glucose. As in glycogen metabolism  (Section 16-3), hormones such as insulin and glucagon regulate the rates of the  opposing pathways of lipid metabolism and thereby control whether fatty acids will  be oxidized or synthesized. The major control mechanisms are summarized in  Fig. 20-31.   Fatty acid oxidation is regulated largely by the concentration of fatty acids  in the blood, which is, in turn, controlled by the hydrolysis rate of triacylglyc¬  erols in adipose tissue by hormone-sensitive triacylglycerol lipase. This en¬  zyme is so named because it is susceptible to regulation by phosphorylation  and dephosphorylation in response to hormonally controlled cAMP levels.  Glucagon, epinephrine, and norepinephrine, which are released in times of  metabolic need, increase adipose tissue cAMP concentrations. cAMP alloster-  ically activates PKA (Section 13-3C), which in turn phosphorylates certain en¬  zymes. Phosphorylation activates hormone-sensitive lipase, thereby stimulating  lipolysis in adipose tissue, raising blood fatty acid levels, and ultimately acti¬  vating the (3-oxidation pathway in other tissues such as liver and muscle. In  liver, this process leads to the production of ketone bodies that are secreted  into the bloodstream for use as an alternative fuel to glucose by peripheral tis¬  sues. PKA also inactivates acetyl-CoA carboxylase (Section 20-4B), so cAMP-  dependent phosphorylation simultaneously stimulates fatty acid oxidation and  inhibits fatty acid synthesis.   Insulin, a pancreatic hormone released in response to high blood glucose  concentrations (the fed state), has the opposite effect of glucagon and epineph¬  rine: It stimulates the formation of glycogen and triacylglycerols. Insulin de¬  creases cAMP levels, leading to the dephosphorylation and thus the  inactivation of hormone-sensitive lipase. This reduces the amounts of fatty acids  available for oxidation. Insulin also activates acetyl-CoA carboxylase (Section  20-4B). The glucagon:insulin ratio therefore determines the rate and direction of  fatty acid metabolism.   Another mechanism that inhibits fatty acid oxidation when fatty acid  synthesis is stimulated is the inhibition of carnitine palmitoyl transferase I  by malonyl-CoA. This inhibition keeps the newly synthesized fatty acids out  of the mitochondria (Section 20-2B) and thus away from the (3-oxidation  system. In fact, heart muscle, an oxidative tissue that does not carry out fatty  acid biosynthesis, contains an isoform of acetyl-CoA carboxylase, ACC2  (Section 20-4B), whose sole function appears to be the synthesis of malonyl-  CoA to regulate fatty acid oxidation.   AMP-dependent protein kinase (AMPK), which phosphorylates (inacti¬  vates) ACC, may itself be an important regulator of fatty acid metabolism.  The enzyme is activated by AMP and inhibited by ATP and thus has been  proposed to serve as a fuel gauge for the cell. When ATP levels are high, sig¬  naling the fed and rested state, this kinase is inhibited, allowing ACC to be¬  come dephosphorylated (activated) so as to stimulate malonyl-CoA production  for fatty acid synthesis in adipose tissue and for inhibition of fatty acid oxi¬  dation in muscle cells. When activity levels increase, causing ATP levels to de¬  crease with a concomitant increase in AMP levels, AMPK is activated to  phosphorylate (inactivate) ACC. The resulting decrease in malonyl-CoA lev¬  els causes fatty acid biosynthesis to decrease in adipose tissue while fatty acid  oxidation increases in muscle to provide the ATP for continued activity.   Factors such as substrate availability, allosteric interactions, and covalent  modification (phosphorylation) control enzyme activity with response times of  minutes or less. Such short-term regulation is complemented by long-term   enzyme by altering the amount of enzyme present. This is accomplished  through changes in the rates of protein synthesis and/or breakdown.   The long-term regulation of lipid metabolism includes stimulation by in¬  sulin and inhibition by starvation of the synthesis of acetyl-CoA carboxylase   • How do hormones link fuel availability to  fatty acid synthesis and degradation in  humans?    and fatty acid synthase. The amount of adipose tissue lipoprotein lipase, the  enzyme that initiates the entry of lipoprotein-packaged fatty acids into adi¬  pose tissue for storage (Section 20-IB), is also increased by insulin and de¬  creased by starvation. Thus, an abundance of glucose, reflected in the level of  insulin, promotes fatty acid synthesis and the storage of fatty acids by  adipocytes, whereas starvation, when glucose is unavailable, decreases fatty acid  synthesis and the uptake of fatty acids by adipocytes.    The triacylglycerol precursors diacylglycerol and phosphatidic acid are used to  synthesize glycerophospholipids.    Fatty acids are the precursors not just of triacylglycerols but of a variety of  other compounds, including membrane lipids and certain signaling molecules.  Most membrane lipids are dual-tailed amphipathic molecules composed of either  1,2-diacylglycerol or TV-acylsphingosine (ceramide) linked to a polar head  group that is either a carbohydrate or a phosphate ester (Fig. 20-32). In this  section, we describe the biosynthesis of these complex lipids from their sim¬  pler components. These lipids are synthesized in membranes, mostly on the  cytosolic face of the endoplasmic reticulum, and from there are transported in  vesicles to their final cellular destinations (Section 9-4E). Arachidonate groups  released from membrane lipids give rise to prostaglandins of various types.   Glycerophospholipids have significant asymmetry in their Cl- and C2-linked  fatty acyl groups: Cl substituents are mostly saturated fatty acids, whereas  those at C2 are by and large unsaturated fatty acids.   Activated Head Groups or Activated Lipids Are Used to Synthesize  Diacylglycerophospholipids. The triacylglycerol precursors 1,2-diacylglycerol  and phosphatidic acid are also the precursors of glycerophospholipids (Fig. 20-29).  The polar head groups of glycerophospholipids are linked to C3 of the   0 ch- C = C - (CH 2 ) 12 - CH 3    FIG. 20-32 Glycerolipids and sphingolipids. The structures of the common  head groups, X, are presented in Table 9-2. Plant membranes are particularly rich in  glyceroglycolipids.    glycerol via a phosphodiester bond. In mammals, the head groups ethanolamine  and choline are activated before being attached to the lipid (Fig. 20-33):   2. The phosphoryl group of the resulting phosphoethanolamine or  phosphocholine then attacks CTP, displacing PP Z , to form the corre¬  sponding CDP derivatives, which are activated phosphate esters of the  polar head group.   3. The C3-OH group of 1,2-diacylglycerol attacks the phosphoryl group  of the activated CDP-ethanolamine or CDP-choline, displacing CMP  to yield the corresponding glycerophospholipid.   In liver, phosphatidylethanolamine can also be converted to phosphatidyl¬  choline by the addition of methyl groups.   Phosphatidylserine is synthesized from phosphatidylethanolamine by a  head group exchange reaction catalyzed by phosphatidylethanolamine serine   In mammals, CDP-ethanolamine and CDP-choline are the precursors of the head groups.    transferase in which serines OH group attacks the donors phosphoryl group.  The original head group is then eliminated, forming phosphatidylserine:   hydrophobic tail is activated rather than the polar head group. Phosphatidic  acid, the precursor of 1,2-diacylglycerol (Fig. 20-29), attacks the a-phosphoryl  group of CTP to form the activated CDP-diacylglycerol and PP Z (Fig. 20-34).  Phosphatidylinositol results from the attack of inositol on CDP-diacylglycerol.  Phosphatidylglycerol is formed in two reactions: (1) attack of the Cl-OH group  of glycerol-3-phosphate on CDP, yielding phosphatidylglycerol phosphate;  and (2) hydrolysis of the phosphoryl group to form phosphatidylglycerol.   Cardiolipin forms by the condensation of two molecules of phosphatidyl¬  glycerol with the elimination of one molecule of glycerol:    Enzymes that synthesize phosphatidic acid have a general preference for  saturated fatty acids at Cl and for unsaturated fatty acids at C2. Yet this gen¬  eral preference cannot account, for example, for the observations that — 80%  of brain phosphatidylinositol has a stearoyl group (18:0) at Cl and an arachi-  donoyl group (20:4) at C2, and that —40% of lung phosphatidylcholine has  palmitoyl groups (16:0) at both positions (the latter substance is the major  component of the surfactant that prevents the lung from collapsing when air    is expelled; its deficiency in premature infants is responsible for respiratory dis¬  tress syndrome; see Box 9-1). William Lands showed that such side chain speci¬  ficity results from “remodeling” reactions in which specific acyl groups of individual  glycerophospholipids are exchanged by specific phospholipases and acyltransferases.   Glycerophospholipids Include the Plasmalogens and Alkylacyl-  glycerophospholipids. Eukaryotic membranes contain significant amounts of  two other types of glycerophospholipids: plasmalogens, which contain a hy¬  drocarbon chain linked to glycerol Cl via a vinyl ether linkage, and alkyl-  acylglycerophospholipids, in which the alkyl substituent at glycerol Cl is  attached via an ether linkage.    About 20% of mammalian glycerophospholipids are plasmalogens, but the ex¬  act percentage varies both among species and among tissues within a given or¬  ganism. For example, plasmalogens account for only 0.8% of the  phospholipids in human liver but 23% of those in human nervous tissue. The  alkylacylglycerophospholipids are much less abundant than the plasmalogens.  The biosynthetic pathway for these lipids yields the alkylacylglycerophospho-  lipid, and this ether is then oxidized to a vinyl ether by the action of a desat-  urase to yield the plasmalogen.   Most sphingolipids are sphingoglycolipids; that is, their polar head groups con¬  sist of carbohydrate units. Cerebrosides are ceramide monosaccharides, whereas  gangliosides are sialic acid-containing ceramide oligosaccharides (Section 9-ID).    These lipids are synthesized by attaching carbohydrate units to the  Cl-OH group of ceramide (7V-acylsphingosine).   7V-Acylsphingosine is synthesized in four reactions from the  precursors palmitoyl-CoA and serine (Fig. 20-35):   1. 3-Ketosphinganine synthase catalyzes condensation of  palmitoyl-CoA with serine, yielding 3-ketosphinganine.   2. 3-Ketosphinganine reductase catalyzes the NADPH-  dependent reduction of 3-ketosphinganine’s keto group to  form sphinganine (dihydrosphingosine).   3. Dihydroceramide is formed by transfer of an acyl group  from an acyl-CoA to sphinganine s 2-amino group, form¬  ing an amide bond.   4. Dihydroceramide dehydrogenase converts dihydro¬  ceramide to ceramide by an FAD-dependent oxidation  reaction.   The nonglycosylated lipid sphingomyelin, an important struc¬  tural lipid of nerve cell membranes,   is the product of a reaction in which phosphatidylcholine do¬  nates its phosphocholine group to the Cl-OH group of 7V-acyl-  sphingosine.   Cerebrosides, which are most commonly l-(3-galacto-  ceramide or l-p-glucoceramide, are synthesized from ceramide by  the addition of the glycosyl unit from the corresponding  UDP-hexose to ceramide s Cl-OH group. The more elaborate  oligosaccharide head groups of gangliosides (Fig. 9-9) are con¬  structed through the action of a series of glycosyltransferases.  Defects in the pathways for degrading these complex lipids are re¬  sponsible for certain lipid storage diseases (Box 20-4).   Prostaglandins and related compounds (Fig. 9-12) are derivatives  of C 2 o fatty acids such as arachidonate, which is released from  membrane phospholipids in response to hormones and other sig¬  nals. The functions of prostaglandins vary in a tissue-specific  manner, but several of them trigger pain, fever, or inflammation.  The production of prostaglandins begins with the formation of  a cyclopentane ring in the linear fatty acid, as catalyzed by  prostaglandin H 2 synthase (Fig. 20-36). This heme-contain¬  ing enzyme contains two catalytic activities: a cyclooxygenase  that adds two molecules of 0 2 to arachidonate, and a peroxi¬  dase that converts the resulting hydroperoxy group to an OH  group. The enzyme is commonly called COX, after its cyclooxy¬  genase activity (not to be confused with cytochrome c oxidase,  which is also called COX).   The use of aspirin as an analgesic (pain-relieving), antipyretic  (fever-reducing), and anti-inflammatory agent has been wide¬  spread since the nineteenth century. Yet it was not until 1971    FIG. 20-36 The prostaglandin H 2 synthase reaction. A cyclooxygenase activity  (1) catalyzes the additions and rearrangements that generate the cyclopentane ring. The  enzyme’s peroxidase activity ( 2 ) converts the peroxide intermediate to prostaglandin H 2  (PGH 2 ), which is the precursor of other prostaglandins.   that John Vane discovered its mechanism of action: Aspirin inhibits the syn¬  thesis of prostaglandins by acetylating a specific Ser residue of prostaglandin  H 2 synthase, which prevents arachidonate from reaching the cyclooxygenase  active site. Other nonsteroidal anti-inflammatory drugs (NSAIDs) such as  ibuprofen and acetaminophen noncovalently bind to the enzyme so as to  similarly block its active site.   T Low doses of aspirin, ~80 mg (baby aspirin) every day, significantly reduce  the long-term incidence of heart attacks and strokes. Such low doses selec¬  tively inhibit platelet aggregation and thus blood clot formation (Box 11-4) be¬  cause these enucleated cells, which have a lifetime in the circulation of ~10  days, cannot resynthesize their inactivated enzymes. Vascular endothelial cells  are not so drastically affected since, for the most part, they are far from the  site where aspirin is absorbed, are exposed to lesser concentrations of aspirin  and, in any case, can synthesize additional PGH 2 synthase.   COX-2 Inhibitors Lack the Side Effects of Other NSAIDs. Prostaglandin H 2  synthase has two isoforms, COX-1 and COX-2, that share a high degree  (60%) of sequence identity and structural homology. COX-1 is constitutively  (without regulation) expressed in most, if not all, mammalian tissues, thereby  supporting levels of prostaglandin synthesis necessary to maintain organ and  tissue homeostasis. In contrast, COX-2 is only expressed in certain tissues in  response to inflammatory stimuli and hence is responsible for the elevated  prostaglandin levels that cause inflammation.   T Aspirin and ibuprofen are relatively nonspecific and therefore can have ad¬  verse side effects (e.g., gastrointestinal ulceration) when used to treat in¬  flammation or fever. A structure-based drug design program (Section 12-4)  was therefore instituted to create inhibitors that would target COX-2 but not  COX-1. The three-dimensional structures of COX-1 and COX-2 are almost  identical. However, their amino acid differences make COX-2 s active site  channel ~20% larger in volume than that of COX-1. Chemists therefore syn¬  thesized inhibitors, collectively known as coxibs, that could enter the COX-  2 channel but are excluded from that of COX-1. Two of these inhibitors,  rofecoxib (Vioxx) and celecoxib (Celebrex) (at right), became important  drugs for the treatment of inflammatory diseases such as arthritis because they  lack the major side effects of the nonspecific NSAIDs. However, in 2004, Vioxx  was withdrawn from the market because of unanticipated cardiac side effects.   Interestingly, acetaminophen, the most commonly used analgesic/  antipyretic (Section 12-4D), binds poorly to both COX-1 and COX-2 and is not  (despite its inclusion in this category) an anti-inflammatory agent. Its mechanism  of action remained a mystery until Daniel Simmons recent discovery of a third  COX isozyme, COX-3, which is expressed at high levels in the central nervous  system and is apparently the target of drugs that decrease pain and fever.   • Explain how glycerophospholipids are  synthesized by activating either the head  group or the lipid tail.   • Explain the action of drugs such as  aspirin, NSAIDs, and coxibs.    Sphingoglycolipids are lysosomally degraded by a  series of enzymatically mediated hydrolytic reactions.  Below is shown the pathway for the degradation of  ganglioside G M1 and a related globoside and sulfatide.  The abbreviations for the monosaccharide residues are  Gal, galactose; GalNAc, A/-acetylgalactosamine; Glc,  glucose; NANA, A/-acetylneuraminic acid (sialic acid).  Cer represents ceramide.   A hereditary defect in one of the enzymes  (indicated by a red bar) results in a sphingolipid  storage disease. The substrate of the missing  enzyme therefore accumulates, often with disastrous  consequences. In many cases, affected individuals  suffer from mental retardation and die in infancy or  early childhood. One of the most common lipid  storage diseases is Tay-Sachs disease, an autosomal  recessive deficiency in hexosaminidase A, which  hydrolyzes A/-acetylgalactosamine from ganglioside    • The liver converts acetyl-CoA to cholesterol by a multistep pathway.   • Cholesterol synthesis is regulated by the activity and amount of HMG-CoA reduc¬  tase.    Cholesterol is a vital constituent of cell membranes and the precursor of  steroid hormones and bile acids. It is clearly essential to life, yet its deposition  in arteries is associated with cardiovascular disease and stroke, two leading    G M2 . The absence of hexosaminidase A activity results in the  accumulation of G M2 as shell-like inclusions in neuronal cells as  shown below:    Although infants born with Tay-Sachs disease at first appear  normal, by ~ 1 year of age, when sufficient G M2 has accumulated  to interfere with neuronal function, they become progressively  weaker, retarded, and blinded until they die, usually by the age of  3 years. It is possible, however, to screen potential carriers of the  disease by a simple serum assay for hexosaminidase A.   Experiments using a mouse model of Tay-Sachs disease sug¬  gest that G M2 accumulation can be reduced by inhibiting its syn¬  thesis. This is accomplished in mice by administering the imino  sugar JV-butyldeoxynojirimycin,   which inhibits the glycosyltransferase in the first step of gluco-  ceramide synthesis. Similar “substrate deprivation” approaches  may be effective in treating other lipid storage diseases, partic¬  ularly when the defective but essential enzyme has some  residual activity.   causes of death in humans. In a healthy organism, an intricate balance is main¬  tained between the biosynthesis, utilization, and transport of cholesterol, keeping  its harmful deposition to a minimum. In this section, we study the pathways of  cholesterol biosynthesis and transport and how they are controlled.   Cholesterol biosynthesis follows a lengthy pathway, first outlined by Konrad  Bloch, in which acetate (from acetyl-CoA) is converted to isoprene units that  have the carbon skeleton of isoprene:    The isoprene units then condense to form a linear molecule with 30 carbons  that cyclizes to form the four-ring structure of cholesterol.    HMG-CoA Is a Key Cholesterol Precursor. Acetyl-CoA is converted to iso¬  prene units by a series of reactions that begins with formation of hydroxy-  methylglutaryl-CoA (HMG-CoA; this compound is also an intermediate in  ketone body synthesis; Fig. 20-21). HMG-CoA synthesis requires thiolase and  HMG-CoA synthase. In liver mitochondria, these two enzymes form HMG-  CoA for ketone body synthesis. Cytosolic isozymes of the two proteins gen¬  erate the HMG-CoA that is used in cholesterol biosynthesis. Four additional   Chapter 20 Lipid Metabolism reactions convert HMG-CoA to the isoprenoid intermediate isopentenyl    1. The CoA thioester group of HMG-CoA is reduced to an alcohol in  an NADPH-dependent four-electron reduction catalyzed by HMG-  CoA reductase, yielding mevalonate, a compound. This is the rate¬  determining step of cholesterol biosynthesis.   3. The phosphate group is converted to a pyrophosphate by phospho-  mevalonate kinase.    4. The molecule undergoes an ATP-dependent decarboxylation reaction  catalyzed by pyrophosphomevalonate decarboxylase to yield isopen¬  tenyl pyrophosphate:    Squalene Is Formed by the Condensation of Six Isoprene Units. Isopentenyl   pyrophosphate is converted to dimethylallyl pyrophosphate by isopentenyl  pyrophosphate isomerase (at left). Four isopentenyl pyrophosphates and two  dimethylallyl pyrophosphates condense to form the C 30 cholesterol precursor  squalene in three reactions catalyzed by two enzymes (Fig. 20-38):   1. Prenyltransferase catalyzes the head-to-tail condensation of dimethyl¬  allyl pyrophosphate and isopentenyl pyrophosphate to yield the C 10  compound geranyl pyrophosphate.   2. Prenyltransferase catalyzes a second head-to-tail condensation of  geranyl pyrophosphate and isopentenyl pyrophosphate to yield the C 15  compound farnesyl pyrophosphate. The prenyltransferase catalyzes an  S N 1 reaction to form a carbocation intermediate with an  ionization—condensation—elimination mechanism:    3. Squalene synthase then catalyzes the head-to-head condensation of  two farnesyl pyrophosphate molecules to form squalene.   Farnesyl pyrophosphate is also the precursor of other isoprenoid compounds  in mammals, including ubiquinone (Section 9-IF) and the isoprenoid tails of  some lipid-linked membrane proteins (Section 9-3B).    Squalene Cyclization Eventually Yields Cholesterol. Squalene, a linear hy¬  drocarbon, cyclizes to form the tetracyclic steroid skeleton in two steps.    FIG. 20-38 Formation of squalene from isopentenyl pyrophosphate and  dimethylallyl pyrophosphate. The pathway involves two head-to-tail condensations  catalyzed by prenyltransferase and a head-to-head condensation catalyzed by squalene  synthase.    Squalene epoxidase catalyzes oxidation of squalene to form 2,3-oxidosqua-  lene (Fig. 20-39). Oxidosqualene cyclase converts this epoxide to the steroid  lanosterol. The reaction is a chemically complex process involving cyclization  of 2,3-oxidosqualene to a protosterol cation and rearrangement of the cation  to lanosterol by a series of 1,2 hydride and methyl shifts (Fig. 20-40).   Conversion of lanosterol to cholesterol (at left) is a 19-step process that  involves an oxidation and the loss of three methyl groups. The enzymes re¬  quired for the process are embedded in the endoplasmic reticulum membrane.   Cholesterol Has Many Uses. In addition to its function as a membrane com¬  ponent, cholesterol is the precursor of steroid hormones such as cortisol, an¬  drogens, and estrogens (Section 9-IE). The liver converts cholesterol to bile  acids (Fig. 20-1), which act as emulsifying agents in the digestion and absorp¬  tion of fats (Section 20-1 A). An efficient recycling system allows the bile acids  to re-enter the bloodstream and return to the liver for reuse several times each  day. The bile acids that escape this recycling are further metabolized by intes¬  tinal microorganisms and excreted. This is the only route for cholesterol excretion.   Cholesterol synthesized by the liver may be esterified by acyl-CoA:cho-  lesterol acyltransferase (ACAT) to form cholesteryl esters.   These highly hydrophobic compounds are transported throughout the body  in lipoprotein complexes (Section 20-IB).   HMG-CoA reductase, which catalyzes the rate-limiting step of cholesterol  biosynthesis (Fig. 20-37), is the pathways main regulatory site. The enzyme  is subject to short-term control by competitive inhibition, allosteric effects,  and covalent modification involving reversible phosphorylation, hike glycogen  phosphorylase, glycogen synthase, and other enzymes, HMG-CoA reductase  exists in interconvertible more active and less active forms. When phosphory-  lated at Ser 871, the enzyme is less active. Phosphorylation is carried out by  AMP-dependent protein kinase (AMPK), the same enzyme that inactivates  acetyl-CoA carboxylase (Section 20-4B). It appears that this control mecha¬  nism conserves energy when ATP levels fall and AMP levels rise, by generally  inhibiting biosynthetic pathways.    FIG. 20-40 The oxidosqualene cyclase reaction. (1) 2,3-Oxidosqualene is cyclized to the protosterol  cation in a process that is initiated by the enzyme-mediated protonation of the squalene epoxide oxygen.   The opening of the epoxide leaves an electron-deficient center whose migration drives the series of  cyclizations that form the protosterol cation. (2) A series of methyl and hydride migrations followed by  the elimination of a proton from C9 of the sterol to form a double bond ultimately yields neutral lanosterol.    The primary regulatory mechanism for HMG-CoA reductase activity is long¬  term feedback control of the amount of enzyme present in the cell. The amount  of enzyme can rise as much as 200-fold, due to an increase in enzyme syn¬  thesis combined with a decrease in its degradation. In fact, cholesterol itself  regulates the expression of the HMG-CoA reductase gene along with more  than 20 other genes involved in its biosynthesis and uptake, including the gene  encoding the LDL receptor. These genes all contain a specific recognition se¬  quence called a sterol regulatory element (SRE; control of eukaryotic gene  expression is discussed in detail in Chapter 28). The transcription of these  genes, as Brown and Goldstein elucidated, requires the binding to their SRE  of a portion of sterol regulatory element binding protein (SREBP).  However, when cholesterol levels are sufficiently high, SREBP resides in the  endoplasmic reticulum (ER) membrane as an inactive 1160-residue precursor H —O 1  that binds to SREBP cleavage-activating protein (SCAP). SCAP is an  ~ 1276-residue intracellular cholesterol sensor that contains two domains: an  N-terminal transmembrane domain called the sterol-sensing domain that  interacts with sterols, and a C-terminal domain containing five copies of a  protein-protein interaction motif known as a WD repeat that interacts with  the C-terminal so-called regulatory domain of SREBP (Fig. 20-41). When cho¬  lesterol in the ER membrane is depleted, SCAP changes conformation and es¬  corts its bound SREPB to the Golgi apparatus via COPII-coated membranous  vesicles (Section 9-4E). SREBP is then sequentially cleaved by two Golgi pro¬  teases: site-1 protease (SIP), a serine protease that cleaves SREBP only when  it is associated with SCAP, and site-2 protease (S2P), a zinc metalloprotease    FIG. 20-41 The cholesterol-mediated  proteolytic activation of SREBP. When  cholesterol levels in the cell are high, the  SREBP-SCAP complex resides in the ER. When  cholesterol levels are low, SCAP escorts SREBP  via COPII-coated membranous vesicles to the  Golgi apparatus, where SREBP undergoes  sequential proteolytic cleavage by the  membrane-bound proteases SIP and S2P. This  releases SREBP’s N-terminal domain, which  enters the nucleus where it binds to the SREs  of its target genes, thereby inducing their  transcription. [After Goldstein, J., Rawson, R.B.,  and Brown, M., Arch. Biochem. Biophys. 397,   sults in the release of the active fragment, a soluble 480-residue protein that  travels to the nucleus, where it activates the transcription of genes containing  an SRE by binding to the SRE via a DNA-binding motif known as a basic  helix-loop-helix (bHLH; Section 24-4C; Fig. 20-41). The cholesterol level in  the cell thereby rises until SCAP no longer induces the translocation of SREBP  to the Golgi apparatus, a classic case of feedback inhibition.   Statins Inhibit HMG-CoA Reductase. High levels of circulating cholesterol,  a condition known as hypercholesterolemia, can be treated with drugs called  statins that inhibit HMG-CoA reductase (Fig. 20-42). These compounds all  contain an HMG-like group that acts as a competitive inhibitor of HMG-  CoA binding to the enzyme. The statins bind extremely tightly, with K\ val¬  ues in the nanomolar range, whereas the substrate, HMG-CoA, has a K M of  ~4 mM.   The X-ray structures of HMG-CoA reductase in its complexes with six  different statins reveal that the bulky hydrophobic groups of the inhibitors  play a major role in interfering with enzyme activity. The HMG-CoA reduc¬  tase active site normally accommodates NADPH as well as the pantothenate  portion of CoA. Statin binding does not interfere with NADPH binding.  However, the enzyme alters its conformation in order to accommodate the  large hydrophobic groups of the statins. It appears that the inhibitors exploit  the inherent flexibility of the enzyme. Furthermore, despite the structural vari¬  ation among the different statins, they all experience extensive van der Waals  contacts with the enzyme. This high degree of complementarity between the  statins and the active site , a result of the enzymes flexibility\ accounts for the ex¬  tremely low inhibition constants.   The initial decreased cellular cholesterol supply caused by the presence of  statins is met by the induction of the LDL receptor and HMG-CoA reduc¬  tase (Sections 20-IB and 20-7A; Fig. 20-8) so that at the new steady state,  the HMG-CoA level is almost that of the predrug state. However, the in¬  creased number of LDL receptors causes increased removal from the blood of  both LDL and IDL (the apoB-100 containing precursor to LDL), decreasing  serum LDL-cholesterol levels appreciably.    FIG. 20-42 Competitive inhibitors of HMG-CoA reductase  used for the treatment of hypercholesterolemia. The molecular  formulas of lovastatin (Mevacor), pravastatin (Pravachol), simvastatin  (Zocor), and atorvastatin (Lipitor), which are known as statins, are  given. The structures of HMG-CoA and the HMG-CoA reductase    product mevalonate are shown for comparison. Note that lovastatin,  pravastatin, and simvastatin are lactones whereas atorvastatin and  mevalonate are hydroxy acids. The lactones are hydrolyzed  enzymatically in vivo to the active hydroxy-acid forms.   As mentioned above, cellular cholesterol concentrations depend not only on  the rate of cholesterol synthesis but also on the ability of cells to absorb cho¬  lesterol from circulating lipoproteins (Section 20-IB). The role of lipoproteins  in cholesterol metabolism has been studied extensively because an elevated  cholesterol level in the blood, primarily in the form of LDL, is a strong risk  factor for cardiovascular disease.   * Approximately half of all deaths in the United States are linked to the vas¬  cular disease atherosclerosis (Greek: athera, mush + sclerosis, hardness).  Atherosclerosis is a slow progressive disease that begins with the deposition of  lipids in the walls of large blood vessels, particularly the coronary arteries. The  initial event appears to be the association of lipoproteins with vessel wall pro¬  teoglycans (Section 8-3A). The trapped lipids trigger inflammation by inducing  the endothelial cells lining the vessels to express adhesion molecules specific for  monocytes, a type of white blood cell. Once these cells burrow into the vessel  wall, they differentiate into macrophages that take up the accumulated lipids,  becoming so engorged that they are known as “foam cells.” Macrophages do not  normally take up lipids but do so when the lipids are oxidized, as apparently  occurs when LDL particles are trapped for an extended period in blood vessel  walls. Factors released by the foam cells, and possibly the oxidized lipids them¬  selves, recruit more white blood cells, perpetuating a state of inflammation.   The damaged vessel wall forms a plaque with a core of cholesterol, cho-  lesteryl esters, and remnants of dead macrophages, surrounded by proliferat¬  ing smooth muscle cells that may undergo calcification, as occurs in bone  formation (hence the “hardening” of the arteries). Although a very large plaque  can occlude the lumen of the artery (Fig. 20-43), blood flow is usually not  completely blocked unless the plaque ruptures. This triggers formation of a  blood clot that can prevent circulation to the heart, causing myocardial  infarction (heart attack). Stoppage of blood flow to the brain causes stroke.   The development of atherosclerosis is strongly correlated with the concen¬  tration of circulating LDL (often referred to as “bad cholesterol”). Some high-  fat diets may contribute to atherosclerosis by boosting LDL levels, but genetic  factors and infection also increase the risk of atherosclerosis. Smoking con¬  tributes to the disease because cigarette smoke oxidizes LDL, which promotes  their uptake by macrophages. Atherosclerosis is less likely to occur in individ¬  uals who maintain low total cholesterol levels in their blood and who have  high levels of HDL (or “good cholesterol”). These lipoproteins transport ex¬  cess cholesterol to the liver for disposal as bile acids. Women have more HDL  than men and a lower risk of heart disease. Statins also reduce the risk of car¬  diovascular disease by decreasing blood cholesterol levels (Section 20-7B).   T LDL Receptor Deficiency Causes Hypercholesterolemia. LDL receptors  clearly play an important role in the maintenance of plasma LDL levels.  Circulating LDL and IDL (which both contain apolipoproteins that specifi¬  cally bind to the LDL receptor) re-enter the liver through receptor-  mediated endocytosis (Fig. 20-8). Individuals with the inherited disease familial  hypercholesterolemia (FH) are deficient in functional LDL receptors. FH ho¬  mozygotes, who lack the receptors entirely, have such high levels of cholesterol-  rich LDL in their plasma that their cholesterol levels are three to five times  greater than the average level of —200 mg/dL. This situation results in the  deposition of cholesterol in their skin and tendons as yellow nodules known  as xanthomas. However, far greater damage is caused by the development of  atherosclerosis, which causes death from heart attacks as early as age 5. FH  heterozygotes (about 1 person in 500) have about half the normal number of  functional LDL receptors and exhibit plasma LDL levels of about twice the  average. They typically develop symptoms of cardiovascular disease after age 30.   FIG. 20-43 An atherosclerotic plaque  in a coronary artery. The vessel wall is  dramatically thickened as a result of lipid  accumulation and activation of inflammatory  processes. [© Eye of Science/Photo  Researchers.]    FIG. 20-44 The role of LDL and HDL in cholesterol  metabolism, (a) Cells acquire cholesterol and cholesteryl esters via  endocytosis of LDL as mediated by the LDL receptor (purple).  Cholesterol efflux to form HDL is assisted by the ABCA1 transporter  (orange), (b) In familial hypercholesterolemia, a lack of functional LDL    receptors results in high levels of circulating LDL (c) In Tangier  disease, cells become laden with cholesterol and cholesteryl esters  and few HDL are formed because efflux is prevented due to lack of  functional ABCA1.    The long-term ingestion of a high-fat/high-cholesterol diet has an effect simi¬  lar to although not as extreme as FH. Cholesterol regulates the synthesis of the  LDL receptor by the same mechanism that regulates HMG-CoA reductase  synthesis (Section 20-7B). Consequently high intracellular concentrations of  cholesterol suppress LDL receptor synthesis so that more LDL particles remain  in the circulation. Excessive dietary cholesterol, delivered to the tissues via chy¬  lomicrons, therefore contributes to high plasma LDL levels.   • Describe the types of reactions required  to produce a molecule of cholesterol.   • What do familial hypercholesterolemia and  Tangier disease reveal about the develop¬  ment of atherosclerosis?    Cholesterol Exits Cells via a Transport Protein. Most cells do not consume  cholesterol by converting it to steroid hormones or bile acids, for example, but  all cells require cholesterol to maintain membrane fluidity. Cholesterol in ex¬  cess of the requirement can be esterified by the action of ACAT and stored as  cholesteryl esters in intracellular deposits. Cholesterol can also be eliminated  from cells by a mechanism illuminated through studies of individuals with  Tangier disease. In this recessive inherited disorder, almost no HDL are pro¬  duced, because cells have a defective transport protein, known as ATP-cassette  binding protein A1 (ABCA1).   In normal individuals, ABCA1 apparently acts as a flippase (Section 9-4C)  to transfer cholesterol, cholesteryl esters, and other lipids from the inner to  the outer leaflet of the plasma membrane, from which they can be captured  by apolipoprotein A-I to form HDL. Cells lacking ABCA1 are unable to offload  cholesterol and accumulate cholesteryl esters in the cytoplasm. Macrophages  thus engorged with lipids contribute to the development of atherosclerosis, so  that individuals with Tangier disease exhibit symptoms similar to those with  FH. The functions of the LDL receptor and ABCA1 in cholesterol homeosta¬  sis are presented schematically in Fig. 20-44.   • Triacylglycerol digestion depends on the emulsifying activity of  bile acids and the activation of lipases at the lipid-water interface.   • Lipoproteins, complexes of nonpolar lipids surrounded by a coat  of amphipathic lipids and apolipoproteins, transport lipids in the  bloodstream. Cells take up cholesterol and other lipids by the  receptor-mediated endocytosis of LDL.   • Fatty acid oxidation begins with the activation of the acyl group  by formation of a thioester with CoA. The acyl group is trans¬  ferred to carnitine for transport into the mitochondria, where it  is re-esterified to CoA.   • |3 oxidation occurs in four reactions: (1) formation of an a,p dou¬  ble bond, (2) hydration of the double bond, (3) dehydrogenation  to form a P-ketoacyl-CoA, and (4) thiolysis by CoA to produce  acetyl-CoA and an acyl-CoA shortened by two carbons. This  process is repeated until fatty acids with even numbers of carbon  atoms are converted to acetyl-CoA and the fatty acids with odd  numbers of carbon atoms are converted to acetyl-CoA and one  molecule of propionyl-CoA. The acetyl-CoA is oxidized by the cit¬  ric acid cycle and oxidative phosphorylation to generate ATP.  Propionyl-CoA is converted to the citric acid cycle intermediate  succinyl-CoA, in part, by the coenzyme B 12 -containing enzyme  methylmalonyl-CoA mutase.   • The oxidation of unsaturated fatty acids requires an isomerase to  convert A 3 double bonds to A 2 double bonds and a reductase to  remove A 4 double bonds. The oxidation of odd-chain fatty acids  yields propionyl-CoA, which is converted to succinyl-CoA through  a cobalamin (B 12 )-dependent pathway. Very long chain fatty acids  are partially oxidized by a three-enzyme system in peroxisomes.   • The liver uses acetyl-CoA to synthesize the ketone bodies acetoac-  etate and P-hydroxybutyrate, which are released into the blood¬  stream. Tissues that use the ketone bodies for fuel convert them  back to acetyl-CoA.   • In fatty acid synthesis, mitochondrial acetyl-CoA is shuttled to the  cytosol via the tricarboxylate transport system and activated to  malonyl-CoA by the action of acetyl-CoA carboxylase.   • A series of seven enzymatic activities, which in mammals are con¬  tained in a multifunctional homodimeric enzyme, extend acyl-  ACP chains by two carbons at a time. An enzyme-bound acyl  group and malonyl-ACP condense to form a P-ketoacyl interme¬    diate and C0 2 . Two reductions and a dehydration yield an acyl-  ACP in a series of reactions that resemble the reverse of P oxida¬  tion but are catalyzed by separate enzymes in the cytosol.  Palmitate (C^), the normal product of fatty acid biosynthesis, is  synthesized in seven such reaction cycles and is then cleaved from  ACP by a thioesterase.   • Other fatty acids are synthesized from palmitate through the ac¬  tion of elongases and desaturases. Human triacylglycerols synthe¬  sized from fatty acyl-CoA and glycerol-3-phosphate or  dihydroxyacetone phosphate tend to contain saturated fatty acids  at Cl and unsaturated fatty acids at C2.   • The opposing pathways of fatty acid degradation and synthesis are  hormonally regulated. Glucagon and epinephrine activate hormone-  sensitive lipase in adipose tissue, thereby increasing the supply of  fatty acids for oxidation in other tissues, and inactivate acetyl-CoA  carboxylase. Insulin has the opposite effect. Insulin also regulates  the levels of acetyl-CoA carboxylase and fatty acid synthase by  controlling their rates of synthesis.   • Mammalian phosphatidylethanolamine and phosphatidylcholine  are synthesized from 1,2-diacylglycerol and CDP derivatives of the  head groups. Phosphatidylinositol, phosphatidylglycerol, and car-  diolipin syntheses begin with CDP-diacylglycerol.   • Sphingoglycolipids are synthesized from ceramide (A^-acylsphingo-  sine, a derivative of palmitate and serine) by the addition of gly-  cosyl units donated by nucleotide sugars.   • Arachidonate is the precursor of prostaglandins. Certain drugs, in¬  cluding aspirin, block prostaglandin synthesis by inhibiting cy¬  clooxygenase.   • Cholesterol is synthesized from acetyl units that pass through  HMG-CoA and mevalonate intermediates on the way to being  converted to a C 5 isoprene unit. Six isoprene units condense to  form the C 30 compound squalene, which cyclizes to yield lanos-  terol, the steroid precursor of cholesterol.   • A cholesterol sensing system in the endoplasmic reticulum regu¬  lates the synthesis of HMG-CoA reductase and LDL receptor in  the cell. HMG-CoA reductase is competitively inhibited by  statin drugs.   • Hypercholesterolemia, associated with certain genetic defects or a  high-cholesterol diet, contributes to the development of athero¬  sclerosis.   endocytosis 663  |3 oxidation 665  homolytic cleavage 676  heterolytic cleavage 677    1. The removal of fatty acids from triacylglycerols leaves glycerol.  Show how the actions of glycerol kinase and glycerol-3-phosphate  dehydrogenase on glycerol produce an intermediate of glycolysis.   2. Identify the products generated by the action of pancreatic lipase  on the lipid shown on page 638.   4. Adipocytes store fat in phospholipid-coated droplets in the cy¬  tosol. The protein perilipin is also associated with the surface of  the lipid droplet, (a) Describe the likely structure of the perilipin  protein, (b) Explain how phosphorylation of perilipin could help  expose the lipids in the droplet to digestion by lipases.   3. One strategy for converting biomass to conventional hydrocar¬  bon fuels involves the use of naturally occurring bacterial en¬  zymes. In certain cyanobacteria, a fatty acyl-ACP undergoes  conversion to a fatty aldehyde. An aldehyde decarbonylase acts on  the aldehyde, producing CO and an alkane. Identify the alkanes  generated from stearoyl-ACP and palmitoyl-ACP.   6. Biodiesel, a fuel typically derived from plant oils, can be manu¬  factured by treating the oil with a methanol/KOH mixture to  produce fatty acid methyl esters. Indicate the structures of the  products generated by treating the triacylglycerol on page 658  with methanol/KOH.   7. Explain why individuals with a hereditary deficiency of carnitine  palmitoyl transferase II have muscle weakness.   8. Why are the symptoms of carnitine palmitoyl transferase II de¬  ficiency more severe during fasting?   9. The first three steps of (3 oxidation (Fig. 20-12) chemically re¬  semble three successive steps of the citric acid cycle. Which steps  are these?   10. The complete combustion of palmitate and glucose yields 9781  kJ*mol 1 and 2850 kJ*mol 1 of free energy, respectively.  Compare these values to the free energy (as ATP) obtained  though catabolism of palmitate and glucose under standard con¬  ditions. Which process is more efficient?   11. Why are unsaturated fats preferable to saturated fats for an indi¬  vidual whose caloric intake must be limited?   12. Calculate the ATP yield for the complete oxidation of oleate.   13. Explain why the degradation of odd-chain fatty acids can boost  the activity of the citric acid cycle.   14. How many ATP can be produced by the complete oxidation of  each propionyl-CoA product of fatty acid oxidation? Compare  this to the ATP yield for each acetyl-CoA.   15. Digestion of plant materials generates phytanate (derived from  chlorophyll molecules; Fig. 19-2).   (b) Instead, phytanate is linked to CoA, then a peroxisomal  dioxygenase converts phytanoyl-CoA to 2-hydroxyphytanoyl-  CoA. Draw the structure of this intermediate.   (c) 2-Hydroxyphytanoyl-CoA lyase generates an aldehyde (pris-  tanal) and formyl-CoA. Draw the structure of pristanal.   16. Pristanal (see Problem 20-15) can be oxidized to pristanate. Can  this compound be further degraded by (3 oxidation and, if so,  what are the reaction products?   17. Why is it important that liver cells lack 3-ketoacyl-CoA trans¬  ferase (Fig. 20-22)?   18. An animal is fed palmitate with a l4 C-labeled carboxyl group.  Under ketogenic conditions, where would the label appear in  acetoacetate?   19. The tricarboxylate transport system supplies cytosolic acetyl-CoA  for palmitate synthesis. What percentage of the NADPH required  for palmitate synthesis is thereby provided?   20. Is the fatty acid shown below likely to be synthesized in animals?  Explain.   21. On what carbon atoms does the l4 C0 2 used to synthesize malonyl-  CoA from acetyl-CoA appear in palmitate?   inhibits bacterial enoyl-ACP reductase. Explain why triclosan  is useful as a microbicide.   23. Compare the energy cost, in ATP equivalents, of synthesizing  stearate from mitochondrial acetyl-CoA to the energy recovered  by degrading stearate to acetyl-CoA.   24. Compare the energy cost, in ATP equivalents, of synthesizing  stearate from mitochondrial acetyl-CoA to the energy recovered  by degrading stearate to C0 2 .   25. Calculate the cost, in ATP equivalents, of converting 12 acetyl-  CoA to lignocerate (a C 2 4 fatty acid; Table 9-1) when elongation  occurs in the mitochondrion.   26. Calculate the cost, in ATP equivalents, of converting 12 acetyl-  CoA to lignocerate when elongation occurs in the endoplasmic  reticulum.   27. Explain why adipocytes need glucose as well as fatty acids in or¬  der to synthesize triacylglycerols.   28. The antidiabetes drugs known as thiazolidinediones induce the  production of glycerol kinase in adipocytes. Explain why this    would decrease the concentration of serum fatty acids, which are  often elevated in diabetics.   29. Pharmacological inhibition of acetyl-CoA carboxylase has been  proposed as a treatment for obesity. Explain why this enzyme  would appear to be a good anti-obesity drug target.   30. Explain why inhibiting the activity of acetyl-CoA carboxylase  might not affect a person’s body mass.   31. An animal is fed palmitate with a l4 C-labeled carboxyl group.  Under conditions of membrane lipid synthesis, where would the  label appear in sphinganine?   32. Hypercholesterolemic individuals taking statins are sometimes  advised to take supplements of coenzyme Q. Explain.   Case 23 The Role of Uncoupling Proteins in Obesity  Focus concept: The properties of adipose tissue factors that un¬  couple oxidative phosphorylation are discussed, and possible links  between uncoupling proteins and obesity are examined.    Project 10 Drug Design and Cholesterol Medications  1. Bile Acid Sequestrants. The relationship between cholesterol  and bile acids was exploited in the use of a common laboratory  material as a medication.    2. Statins. Follow the history of the development of HMG-CoA re¬  ductase inhibitors and explore related structures on the Protein  Data Bank web site.    Clinical assays are used to  measure the blood levels of triacylglycerols and cholesterol. Why is  it necessary to break the total serum cholesterol level into an LDL  and an HDL component? What levels of LDL cholesterol and HDL  cholesterol are considered normal? What levels are associated with  an increased risk of cardiovascular disease? In addition to the  statins, what types of drugs affect serum cholesterol levels? How  do they work and why are they often prescribed in combination?   Valle, D. (Ed.), The Online Metabolic & Molecular Bases of Inherited Dis-  ease, http://www.ommbid.com/. [Includes numerous chapters on de¬  fects in lipid metabolism.]   Vance, D.E. and Vance J.E. (Eds.), Biochemistry of Lipids, Lipoproteins,  and Membranes (3th ed.), Elsevier (2008).   Ajees, A.A., Anantharamaiah, G.M., Mishra, V.K., Hussain, M.M., and  Murthy, H.M.K., Crystal structure of human apolipoprotein A-I: In¬  sights into its protective effect against cardiovascular diseases, Proc.  Natl. Acad. Sci. 103, 2126-2131 (2006).   Bartlett, K. and Eaton, S., Mitochondrial P-oxidation, Eur. J. Biochem.  271, 462-469 (2004). [Discusses the reactions and enzymes of mi¬  tochondrial P oxidation as well as their regulation.]   Kent, C., Eukaryotic phospholipid synthesis, Anna. Rev. Biochem. 64,  313-342 (1995).   Kim, J.-J.P. and Battaile, K.P., Burning fat: the structural basis of fatty  acid p-oxidation, Curr. Opin. Struct. Biol. 12, 721-728 (2002).   Maier, T., Leibundgut, M., and Ban, N., The crystal structure of a mam¬  malian fatty acid synthase, Science 321, 1315-1322 (2008); and    Leibundgut, M., Maier, T., Jenni, S., and Ban, N., The multienzyme  architecture of eukaryotic fatty acid synthases, Curr. Opin. Struct.  Biol. 18, 714-725 (2008).   Marsh, E.N.G. and Drennan, C.L., Adenosylcobalamin-dependent  isomerases: new insights into structure and mechanism, Curr. Opin.  Chem. Biol. 5, 499-505 (2001).   Kurumbail, R.G., Kiefer, J.R., and Marnett, L.J., Cyclooxygenase en¬  zymes: catalysis and inhibition, Curr. Opin. Struct. Biol. 11,  752-760 (2001).   Ikonen, E., Cellular cholesterol trafficking and compartmentalization,  Nature Rev. Mol. Cell Biol. 9, 125-138 (2008).   Istvan, E.S. and Deisenhofer, J., Structural mechanism for statin inhibi¬  tion of HMG-CoA reductase, Science 292, 1160-1164 (2001).   Steinberg, D., Atherogenesis in perspective: hypercholesterolemia and in¬  flammation as partners in crime, Nature Medicine 8, 1211-1216  (2002). [Summarizes current hypotheses linking cholesterol levels to  the development of atherosclerosis.]   A Lysosomes Degrade Many Proteins  B Ubiquitin Marks Proteins for Degradation  C The Proteasome Unfolds and Hydrolyzes  Ubiquitinated Polypeptides   A Five Enzymes Carry Out the Urea Cycle  B The Urea Cycle Is Regulated by Substrate  Availability   A Alanine, Cysteine, Glycine, Serine, and  Threonine Are Degraded to Pyruvate  B Asparagine and Aspartate Are Degraded  to Oxaloacetate   C Arginine, Glutamate, Glutamine, Histidine, and  Proline Are Degraded to a-Ketoglutarate  D Methionine, Threonine, Isoleucine, and Valine  Are Degraded to Succinyl-CoA  E Leucine and Lysine Are Degraded Only  to Acetyl-CoA and/or Acetoacetate  F Tryptophan Is Degraded to Alanine and  Acetoacetate   A Nonessential Amino Acids Are Synthesized  from Common Metabolites  B Plants and Microorganisms Synthesize  the Essential Amino Acids   A Nitrogenase Reduces N 2 to NH 3  B Fixed Nitrogen Is Assimilated into Biological  Molecules   Many of the neurotransmitters that relay signals between nerve cells are amino  acids or derivatives of amino acids. These small molecules are easily synthesized by  the cells, stockpiled until their release, then rapidly degraded or taken up again.  [Thomas Deerinck, NCMIR/Photo Researchers, Inc.]    The metabolism of amino acids comprises a wide array of synthetic and  degradative reactions by which amino acids are assembled as precursors of  polypeptides or other compounds and broken down to recover metabolic  energy. The chemical transformations of amino acids are distinct from those  of carbohydrates or lipids in that they involve the element nitrogen. We must  therefore examine the origin of nitrogen in biological systems and its disposal.   The bulk of the cells amino acids are incorporated into proteins, which  are continuously being synthesized and degraded. Aside from this dynamic  pool of polymerized amino acids, there is no true storage form of amino acids  analogous to glycogen or triacylglycerols. Mammals synthesize certain amino  acids and obtain the rest from their diets. Excess dietary amino acids are not  simply excreted but are converted to common metabolites that are precursors of  glucose , fatty acids, and ketone bodies and are therefore metabolic fuels.   In this chapter, we consider the pathways of amino acid metabolism, be¬  ginning with the degradation of proteins and the deamination (amino group  removal) of their component amino acids. We then examine the incorpora¬  tion of nitrogen into urea for excretion. Next, we examine the pathways by  which the carbon skeletons of individual amino acids are broken down and  synthesized. We conclude with a brief examination of some other biosynthetic  pathways involving amino acids and nitrogen fixation, a process that converts  atmospheric N 2 to a biologically useful form.   • Extracellular and intracellular proteins may be digested by lysosomal proteases.   • Other proteins to be degraded are first conjugated to the protein ubiquitin.   • The proteasome, a barrel-shaped complex, unfolds ubiquitinated proteins in an  ATP-dependent process and proteolytically degrades them.   The components of living cells are constantly turning over. Proteins have life¬  times that range from as short as a few minutes to weeks or more. In any  case, cells continuously synthesize proteins from and degrade them to amino acids.  This seemingly wasteful process has three functions: (1) to store nutrients in  the form of proteins and to break them down in times of metabolic need,  processes that are most significant in muscle tissue; (2) to eliminate abnor¬  mal proteins whose accumulation would be harmful to the cell; and (3) to  permit the regulation of cellular metabolism by eliminating superfluous en¬  zymes and regulatory proteins. Controlling a proteins rate of degradation is  therefore as important to the cellular and organismal economy as is controlling  its rate of synthesis.   The half-lives of different enzymes in a given tissue vary substantially, as  is indicated for rat liver in Table 21-1. Remarkably, the most rapidly degraded  enzymes all occupy important metabolic control points , whereas the relatively sta¬  ble enzymes have nearly constant catalytic activities under all physiological condi¬  tions. The susceptibilities of enzymes to degradation have evidently evolved  along with their catalytic and allosteric properties so that cells can efficiently  respond to environmental changes and metabolic requirements. The rate of  protein degradation in a cell also varies with its nutritional and hormonal state.  For example, under conditions of nutritional deprivation, cells increase their  rate of protein degradation so as to provide the necessary nutrients for indis¬  pensable metabolic processes.   Lysosomes contain ~50 hydrolytic enzymes, including a variety of proteases  known as cathepsins. The lysosome maintains an internal pH of ~5, and its  enzymes have acidic pH optima. This situation presumably protects the cell  against accidental lysosomal leakage since lysosomal enzymes are largely inac¬  tive at cytosolic pHs.   Lysosomes degrade substances that the cell takes up via endocytosis  (Section 20-IB). They also recycle intracellular constituents that are enclosed  within vesicles that fuse with lysosomes, a process called autophagy (Greek:  autos y self + phagein, to eat). In well-nourished cells , lysosomal protein degrada¬  tion is nonselective. In starving cells, however, this degradation would deplete  essential enzymes and regulatory proteins. Lysosomes therefore also have a se¬  lective pathway, which is activated only after a prolonged fast, that imports  and degrades cytosolic proteins containing the pentapeptide Lys-Phe-Glu-Arg-  Gln (KFERQ) or a closely related sequence. Such KFERQ proteins are selec¬  tively lost from tissues that atrophy in response to fasting (e.g., liver and  kidney) but not from tissues that do not do so (e.g., brain and testes). Many  normal and pathological processes are associated with increased lysosomal ac¬  tivity, for example, the muscle wastage caused by disuse, denervation, or trau¬  matic injury. The regression of the uterus after childbirth, when the muscular  organ reduces its mass from 2 kg to 50 g in nine days, is a striking example  of this process. Many chronic inflammatory diseases such as rheumatoid  arthritis involve the extracellular release of lysosomal enzymes, which break  down surrounding tissues.    FIG. 21-1 X-Ray structure of human ubiquitin. The polypeptide is colored in rainbow  order from its N-terminus (blue) to its C-terminus (red) with the side chain of Lys 48 drawn  in stick form with C green and N blue. [Based on an X-ray structure by Charles Bugg,  University of Alabama at Birmingham. PDBid 1UBQJ © See Interactive Exercise 29.   Protein breakdown in eukaryotic cells also occurs in an ATP-requiring process  that is independent of lysosomes. This process involves ubiquitin (Fig. 21-1),  a 76-residue monomeric protein named for its ubiquity and abundance. It is  one of the most highly conserved eukaryotic proteins known (it is identical in  such diverse organisms as humans, trout, and Drosophila :), suggesting that it  is uniquely suited for some essential cellular function(s).   Proteins are marked for degradation by covalently linking them to ubiquitin.  This process occurs in three steps, elucidated notably by Avram Hershko,  Aaron Ciechanover, and Irwin Rose (Fig. 21-2):   1. In an ATP-requiring reaction, ubiquitin s terminal carboxyl group is  conjugated, via a thioester bond, to ubiquitin-activating enzyme  (El). Most organisms have only one type of El.   2. The ubiquitin is then transferred to a specific Cys sulfhydryl group on  one of numerous homologous proteins named ubiquitin-conjugating  enzymes (E2s; 11 in yeast and >20 in mammals). The various E2s  are characterized by an ~ 150-residue catalytic core containing the ac¬  tive site Cys.   3. Ubiquitin-protein ligase (E3) transfers the activated ubiquitin from  E2 to a Lys 8-amino group of a previously bound protein, thereby  forming an isopeptide bond. Cells contain many species of E3s, each  of which mediates the ubiquitination (alternatively, ubiquitylation) of  a specific set of proteins and thereby marks them for degradation. Each  E3 is served by one or a few specific E2s. The known E3s are mem¬  bers of two unrelated families, those containing a HECT domain  (HECT for homologous to A6AP Oerminus) and those containing a  so-called RING finger (RING for really interesting new gene), al¬  though some E2s react well with members of both families. The hu¬  man genome contains 28 HECT genes and 616 RING genes, more  than its number of protein kinase genes (518), which is indicative of  the E3s’ specialized and varied functions (see below).   In order for a protein to be efficiently degraded, it must be linked to a chain of  at least four tandemly linked ubiquitin molecules in which Lys 48 of each ubiq¬  uitin forms an isopeptide bond with the C-terminal carboxyl group of the fol¬  lowing ubiquitin. These polyubiquitin chains may contain 50 or more ubiquitin  units. Ubiquitinated proteins are dynamic entities, with ubiquitin molecules be¬  ing rapidly attached and removed (the latter by ubiquitin isopeptidases).   Until the mid-1990s, it appeared that the ubiquitin system functioned mainly  in a “housekeeping” capacity to maintain the proper balance among metabolic  proteins and to eliminate damaged proteins. Indeed, as Alexander Varshavsky    FIG. 21-2 Reactions involved in protein ubiquitination. Ubiquitin’s terminal carboxyl  group is first joined, via a thioester linkage, to El in a reaction driven by ATP hydrolysis.  The activated ubiquitin is subsequently transferred to a sulfhydryl group of an E2 and then,  in a reaction catalyzed by an E3, to a Lys e-amino group on a condemned protein, thereby  marking the protein for proteolytic degradation.   discovered, the half-lives of many cytoplasmic proteins vary with the identi- Section 1 Protein Degradation   ties of their N-terminal residues via the so-called N-end rule: Proteins with  the “destabilizing” N-terminal residues Asp, Arg, Leu, Lys, and Phe have half-  lives of only 2 to 3 minutes, whereas those with the “stabilizing” N-terminal  residues Ala, Gly, Met, Ser, Thr, and Val have half-lives of >10 hours in  prokaryotes and >20 hours in eukaryotes. The N-end rule applies in both eu¬  karyotes and prokaryotes, which suggests the system that selects proteins for  degradation is conserved in eukaryotes and prokaryotes, even though prokary¬  otes lack ubiquitin.   In eukaryotes, the N-end rule results from the actions of a RING finger  E3 named E3a whose ubiquitination signals are the destabilizing N-terminal  residues. However, it is now clear that the ubiquitin system is far more so¬  phisticated than a simple garbage disposal system. Thus, the many E3s have  a variety of ubiquitination signals that often occur on a quite limited range of  target proteins, many of which have regulatory functions. For example, it has  long been known that proteins with segments rich in Pro (P), Glu (E), Ser  (S), and Thr (T), the so-called PEST proteins, are rapidly degraded. It is now  realized that this is because these PEST elements often contain phosphoryla¬  tion sites that target their proteins for ubiquitination. The resulting destruc¬  tion of these regulatory proteins, of course, has important regulatory  consequences. Likewise, proteins known as cyclins that control the progres¬  sion of the cell cycle (the general sequence of events that occur over the life¬  times of eukaryotic cells; Section 28-4A) are selectively degraded through their  ubiquitination at specific stages of the cell cycle. Interestingly, reversible  monoubiquitination controls the activities of certain proteins rather than  their degradation, in much the same way that phosphorylation and dephos¬  phorylation alter protein activity (Section 12-3B).   Ubiquitinated proteins are proteolytically degraded in an ATP-dependent  process mediated by a large (—2100 kD, 26S) multiprotein complex named  the 26S proteasome (Fig. 21-3). The 26S proteasome consists of a hollow  cylindrical core, known as the 20S proteasome, which is covered at each end   FIG. 21-3 Electron microscopy-based image of the Drosophila melanogaster  26S proteasome. The complex is around 450 X 190 A. The central portion of the  twofold symmetric multiprotein complex (yellow), the 20S proteasome, consists of four  stacked seven-membered rings of subunits that form a hollow barrel in which the proteolysis  of ubiquitin-linked proteins occurs. The 19S caps (blue), which may attach to one or both  ends of the 20S proteasome, control the access of condemned proteins to the 20S  proteasome (see text). [Courtesy of Wolfgang Baumeister, Max-Planck-lnstitut fur  Biochemie, Martinsried, Germany.]   responding particles 5 sedimentation coefficients in units of Svedbergs (S);  Section 5-2E.]   The yeast 20S proteasome, which is closely similar to other eukaryotic 20S  proteasomes, is composed of seven different types of a-like subunits and seven  different types of (3-like subunits. The X-ray structure of this enormous (6182-  residue, ~670-kD) protein complex, determined by Robert Huber, reveals that  it consists of four stacked rings of subunits with its outer and inner rings, re¬  spectively, consisting of seven different a-type subunits and seven different  (3-type subunits (Fig. 21-4#). The various a-type subunits have folds that are  similar to one another and to the various (3-type subunits. Consequently, the  28-subunit complex has exact twofold rotational symmetry relating the two  pairs of rings, but only pseudosevenfold rotational symmetry relating the sub¬  units within each ring. The 20S proteasomes hollow core consists of three  large chambers (Fig. 21-4£): Two are located at the interfaces between adjoin¬  ing rings of a and (3 subunits, with the third, larger chamber centrally located  between the two rings of (3 subunits.   Although the a-type subunits and the (3-type subunits are structurally sim¬  ilar, only three of the (3-type subunits have proteolytic activity. The X-ray struc¬  ture together with enzymological studies reveal that the three active sites  catalyze peptide bond hydrolysis via a novel mechanism in which the (3 sub¬  units 5 N-terminal Thr residues function as catalytic nucleophiles. The active  sites are located inside the central chamber of the 20S proteasome, thereby pre¬  venting this omnivorous protein-dismantling machine from indiscriminately  hydrolyzing the proteins in its vicinity. It appears that polypeptide substrates  must enter the central chamber of the barrel through the narrow axially located  apertures in the a rings that are lined with hydrophobic residues so that only  unfolded proteins can enter the central chamber. Nevertheless, in the X-ray   (a) The arrangement of the 28 subunits represented as spheres. Four  rings of seven subunits each are stacked to form a barrel (the ends  of the barrel are at the right and left) with the a-type and (3-type  subunits forming the outer and inner rings, respectively. The complex’s  twofold (C 2 ) axis of symmetry is represented by the vertical red line.   (b) Surface view of the proteasome core cut along its cylindrical axis.  Three bound protease inhibitor molecules marking the three active p    subunits are shown in red as space-filling models. The entry channels  at each end of the cylinder are not visible in this model. [Courtesy of  Robert Huber, Max-Planck-lnstitut fur Biochemie, Martinsried, Germany.  PDBid 1RYPJ   Compare this structure to that of the GroEL/ES chaperonin  (Fig 6-44).    structure of the yeast 20S proteasome (Fig. 21-4£), these apertures are blocked  by a plug formed by the interdigitation of the a subunits N-terminal tails.   The three active (3-type subunits have different substrate specificities,  cleaving after acidic residues (the (31 subunit), basic residues (the (32 subunit),  and hydrophobic residues (the (35 subunit). As a result, the 20S proteasome  cleaves its polypeptide substrates into ~ 8-residue fragments, which then dif¬  fuse out of the proteasome. Cytosolic peptidases degrade the peptides to their  component amino acids. The ubiquitin molecules attached to the target  protein are not degraded, however, but are returned to the cell and reused.   r ln addition to facilitating normal protein turnover, proteasomes play a role  in the immune system. Certain antigen-presenting cells display peptide  antigens on their surface in order to initiate an immune response. Receptors  on T lymphocytes (Section 7-3) recognize the peptides, when they are com-  plexed with cell-surface proteins that are members of the major histocompat¬  ibility complex (MHC). During a bacterial or viral infection, signals from  damaged cells or other immune-system cells stimulate the antigen-presenting  cells to increase their synthesis of alternate proteasomal (3 subunits that have  altered substrate specificity. The resulting peptide fragments, which represent  normal cellular proteins as well as pathogen-derived proteins, bind optimally  to the MHC proteins and hence can better alert T lymphocytes to the pres¬  ence of the pathogen.   19S Caps Control the Access of Ubiquitinated Proteins to the 20S  Proteasome. The 20S proteasome probably does not exist alone in vivo; it  is most often in complex with two 19S caps that recognize ubiquitinated pro¬  teins, unfold them, and feed them into the 20S proteasome in an ATP-  dependent manner. The 19S cap, which consists of ~18 different subunits,  is poorly characterized due in large part to its low intrinsic stability. Its so-called  base complex consists of 9 different subunits, 6 of which are ATPases that  form a ring that abuts the a ring of the 20S proteasome (Fig. 21-3). Cecile  Pickart demonstrated, via cross-linking experiments, that one of these  ATPases, named S6', contacts the polyubiquitin signal that targets a con¬  demned protein to the 26S proteasome. This suggests that the recognition of  the polyubiquitin chain as well as substrate protein unfolding are ATP-driven  processes. Moreover, the ring of ATPases must open the otherwise closed  axial aperture of the 20S proteasome so as to permit the entry of the unfolded  substrate protein.   Eight additional subunits form the so-called lid complex, the portion of  the 19S cap that is more distant from the 20S proteasome. The functions of  the lid subunits are largely unknown, although a truncated 26S proteasome  that lacks the lid subunits is unable to degrade polyubiquitinated substrates.  Several other subunits may be transiently associated with the 19S cap and/or  with the 20S proteasome.   Eubacteria Also Contain Self-Compartmentalized Proteases. Although  20S proteasomes occur in all eukaryotes and archaebacteria yet examined, they  are absent in nearly all eubacteria (which provides further evidence that eu¬  karyotes arose from archaea; Fig. 1-9). Nevertheless, eubacteria have ATP-  dependent proteolytic assemblies that share a barrel-shaped architecture with  proteasomes and carry out similar functions. For example, in E. coli , two pro¬  teins known as Lon and Clp carry out up to 80% of the bacterium’s protein  degradation. Thus, all cells appear to contain proteases whose active sites are only  available from the inner cavity of a hollow particle to which access is controlled.  These so-called self-compartmentalized proteases appear to have arisen early  in the history of cellular life, before the advent of eukaryotic membrane-bound  organelles such as the lysosome, which similarly carry out degradative processes  in a way that protects the cell contents from indiscriminant destruction.   FIG. 21 -5 X-Ray structure of E. coli CIpP. This complex of 14  identical subunits has D 7 symmetry (the symmetry of a heptagonal  prism; Section 6-3). (a) View along the protein’s sevenfold axis, drawn  in ribbon form in which the lower ring is pale cyan and the upper ring  is pink with one subunit colored in rainbow order from its N-terminus   (blue) to its C-terminus (red), (b) View along the protein’s twofold axis  (rotated 90° about a horizontal axis with respect to Part a). [Based on  an X-ray structure by John Flannagan, Brookhaven National  Laboratory, Upton, New York. PDBid 1TYF.]    What is the role of the lysosome in  degrading extracellular and intracellular  proteins?    • Describe the steps of protein ubiquitina-  tion What is the difference between  mono- and polyubiquitination?   • Describe the pathway for proteasome-  mediated protein degradation, including  the roles of ubiquitin and ATP.    • What is the advantage of the proteasomal  active sites having different substrate  specificities?   Clp protease consists of two components, the proteolytically active ClpP  and one of several ATPases, which in E. coli are ClpA and ClpX. The X-ray  structure of ClpP reveals that it oligomerizes to form a hollow barrel ~90 A long  and wide that consists of two back-to-back sevenfold symmetric rings of  193-residue subunits and thereby has the same rotational symmetry as does  the 20S proteasome (Fig. 21-5). Nevertheless, the ClpP subunit has a novel  fold that is entirely different from that of the 20S proteasome’s homologous  a and (3 subunits. The ClpP active site, which is only exposed on the inside  of the barrel, contains a catalytic triad composed of its Ser 97, His 122, and  Asp 171, and hence is a serine protease (Fig. 11-26).    Free amino acids originate from the degradation of cellular proteins and from  the digestion of dietary proteins. The gastric protease pepsin, the pancreatic  enzymes trypsin, chymotrypsin, and elastase (discussed in Sections 5-3B and  11-5), and a host of other endo- and exopeptidases degrade polypeptides to  oligopeptides and amino acids. These substances are absorbed by the intestinal  mucosa and transported via the bloodstream to be absorbed by other tissues.   The further degradation of amino acids takes place intracellularly and in¬  cludes a step in which the a-amino group is removed. In many cases, the  amino group is converted to ammonia, which is then incorporated into urea  for excretion (Section 21-3). The remaining carbon skeleton (a-keto acid) of  the amino acid can be broken down to other compounds (Section 21-4). This  metabolic theme is outlined in Fig. 21-6.   FIG. 21-6 Overview of amino acid catabolism. The amino group is removed and  incorporated into urea for disposal. The remaining carbon skeleton (a-keto acid) can be  broken down to C0 2 and H 2 0 or converted to glucose, acetyl-CoA, or ketone bodies.   Most amino acids are deaminated by transamination, the transfer of their  amino group to an a-keto acid to yield the a-keto acid of the original amino  acid and a new amino acid. The predominant amino group acceptor is  a-ketoglutarate, producing glutamate and the new a-keto acid:    Glutamates amino group, in turn, can be transferred to oxaloacetate in a sec¬  ond transamination reaction, yielding aspartate and re-forming a-ketoglutarate:    The enzymes that catalyze transamination, called aminotransferases or  transaminases, require the coenzyme pyridoxal-5'-phosphate (PLP;  Fig. 21-7 a). PLP is a derivative of pyridoxine (vitamin B 6 ; Fig. 21-7^). The  coenzyme is covalently attached to the enzyme via a Schiff base (imine) link¬  age formed by the condensation of its aldehyde group with the 8-amino group  of an enzyme Lys residue (Fig. 21-7c). The Schiff base, which is conjugated to  the pyridinium ring, is the center of the coenzymes activity. When PLP   (b) Pyridoxine (vitamin B 6 ). (c) The Schiff base  that forms between PLP and an enzyme e-amino  group, (d) Pyridoxamine-5'-phosphate (PMP).   Chapter 21 Amino Acid Metabolism accepts the amino group from an amino acid as described below, it becomes   Esmond Snell, Alexander Braunstein, and David Metzler demonstrated  that the aminotransferase reaction occurs via a Ping Pong mechanism (Section  12-ID) whose two stages consist of three steps each (Fig. 21-8):   1. The amino acids nucleophilic amino group attacks the enzyme—PLP  Schiff base carbon atom in a transimination reaction to form an  amino acid-PLP Schiff base (an aldimine), with concomitant release  of the enzymes Lys amino group. This Lys is then free to act as a gen¬  eral base at the active site.   2. The amino acid-PLP Schiff base tautomerizes to an a-keto acid-PMP  Schiff base (a ketimine) by the active site Lys-catalyzed removal of the  amino acid a-hydrogen and protonation of PLP atom C4' via a  resonance-stabilized carbanion intermediate. This resonance stabiliza¬  tion facilitates the cleavage of the C a —H bond.   3. The a-keto acid-PMP Schiff base is hydrolyzed to PMP and an  a-keto acid.   Stage II Converts an a-Keto Acid to an Amino Acid. To complete the  aminotransferases catalytic cycle, the coenzyme must be converted from PMP  back to the enzyme—PLP Schiff base. This involves the same three steps as  above, but in reverse order:   3'. PMP reacts with an a-keto acid to form a Schiff base.   2'. The a-keto acid-PMP Schiff base tautomerizes to form an amino  acid-PLP Schiff base.   1'. The 8-amino group of the active site Lys residue attacks the amino  acid-PLP Schiff base in a transimination reaction to regenerate the ac¬  tive enzyme-PLP Schiff base and release the newly formed amino acid.   Note that removal of the substrate amino acids amino group produces a  resonance-stabilized C a carbanion whose electrons are delocalized all the way  to the coenzymes protonated pyridinium nitrogen atom; that is, PLP func¬  tions as an electron sink. For transamination reactions, this electron-withdraw¬  ing capacity facilitates removal of the a proton (a bond cleavage, top right of  Fig. 21-8) during tautomerization. PLP functions similarly in enzymatic reac¬  tions involving b and c bond cleavage.   Aminotransferases differ in their specificity for amino acid substrates in  the first stage of the transamination reaction, thereby producing the corre¬  spondingly different a-keto acid products. Most aminotransferases, however,  accept only a-ketoglutarate or (to a lesser extent) oxaloacetate as the a-keto  acid substrate in the second stage of the reaction, thereby yielding glutamate  or aspartate as their only amino acid product. The amino groups from most  amino acids are consequently funneled into the formation of glutamate and aspar¬  tate. The transaminase reaction is freely reversible, so transaminases participate  in pathways for amino acid synthesis as well as degradation. Lysine is the only  amino acid that is not transaminated.   T The presence of transaminases in muscle and liver cells makes them useful  markers of tissue damage. Assays of the enzymes’ activities in the blood are  the basis of the commonly used clinical measurements known as SGOT (serum  glutamate-oxaloacetate transaminase, also known as aspartate transaminase,  AST) and SGPT (serum glutamate-pyruvate transaminase, or alanine  transaminase, ALT). The concentrations of these enzymes in the blood increase  after a heart attack, when damaged heart muscle leaks its intracellular contents.  Liver damage is also monitored by SGOT and SGPT levels.    FIG. 21-8 The mechanism of PLP-dependent enzyme-  catalyzed transamination. The first stage of the reaction, in which  the a-amino group of an amino acid is transferred to PLP yielding an  a-keto acid and PMP, consists of three steps: (l) transimination,   ( 2 ) tautomerization, in which the Lys released during the transimination  reaction acts as a general acid-base catalyst, and (3) hydrolysis. The  second stage of the reaction, in which the amino group of PMP is    transferred to a different a-keto acid to yield a new a-amino acid and  PLP, is essentially the reverse of the first stage: Steps 3', 2', and 1'  are, respectively, the reverse of Steps 3, 2, and 1. © See the    Describe the products that would result from cleavage of  the amino acid’s b and c bonds indicated at the upper right.   Transamination, of course, does not result in any net deamination. Glutamate,  however, can be oxidatively deaminated by glutamate dehydrogenase  (GDH), yielding ammonia and regenerating a-ketoglutarate for use in addi¬  tional transamination reactions.   Glutamate dehydrogenase, a mitochondrial enzyme, is the only known  enzyme that can accept either NAD + or NADP + as its redox coenzyme.  Oxidation is thought to occur with transfer of a hydride ion from glutamates  C a to NAD(P) + , thereby forming a-iminoglutarate, which is hydrolyzed to  a-ketoglutarate and ammonium ion:   • Describe how a-ketoglutarate and  oxaloacetate participate in amino acid  catabolism.   • Summarize the reactions that release an  amino acid’s amino group as ammonia.    The enzyme is aliosterically inhibited by GTP and NADH (signaling  abundant metabolic energy) and activated by ADP and NAD + (signaling the  need to generate ATP). Because the product of the reaction, a-ketoglutarate,  is an intermediate of the citric acid cycle, activation of glutamate dehydroge¬  nase can stimulate flux through the citric acid cycle, leading to increased ATP  production by oxidative phosphorylation.   The equilibrium position of the glutamate dehydrogenase reaction  (A G of ~ 30 kj • mol ) favors glutamate synthesis, the reverse of the reaction  written above. At one time, it was believed that this reaction represented a  route for the body to remove free ammonia, which is toxic at high concentra¬  tions. Under physiological conditions, the enzyme was thought to function  close to equilibrium so changes in ammonia concentration could, in princi¬  ple, cause a shift in equilibrium toward glutamate synthesis to remove the ex¬  cess ammonia. However, a form of hyperinsulinism that is characterized by  hypoglycemia and hyperammonemia (HI/HA; hyperammonemia is elevated  levels of ammonia in the blood) is caused by mutations in GDH resulting in  decreased sensitivity to GTP inhibition and therefore increased GDH activity.  Since HI/HA patients have increased GDH activity but higher levels of NH 3  than normal, this accepted role of GDH functioning close to equilibrium and  preventing ammonia toxicity cannot be correct. Indeed, if GDH functioned  close to equilibrium, changes in its activity resulting from allosteric interac¬  tions would not result in significant flux changes (recall that enzymes control¬  ling flux must function far from equilibrium).   The ammonia liberated in the GDH reaction as written above is eventu¬  ally excreted in the form of urea. Thus, the glutamate dehydrogenase reaction  functions to eliminate amino groups from amino acids that undergo transamina¬  tion reactions with a- ketoglutarate.   The rate of the urea cycle changes with the rate of amino acid breakdown.    Living organisms excrete the excess nitrogen arising from the metabolic break¬  down of amino acids in one of three ways. Many aquatic animals simply ex¬  crete ammonia. Where water is less plentiful, however, processes have evolved    that convert ammonia to less toxic waste products that require less water for  excretion. One such product is urea, which is produced by most terrestrial  vertebrates; another is uric acid, which is excreted by birds and terrestrial  reptiles.   In this section, we focus our attention on urea formation. Uric acid biosyn¬  thesis is discussed in Section 23-4.   Urea is synthesized in the liver by the enzymes of the urea cycle. It is  then secreted into the bloodstream and sequestered by the kidneys for excre¬  tion in the urine. The urea cycle was outlined in 1932 by Hans Krebs and  Kurt Henseleit (the first known metabolic cycle; Krebs did not elucidate the  citric acid cycle until 1937; Box 17-1). Its individual reactions were later de¬  scribed in detail by Sarah Ratner and Philip Cohen. The overall urea cycle re¬  action is    Thus, ureas two nitrogen atoms are contributed by ammonia and aspartate,  whereas its carbon atom comes from HC0 3 .   Five enzymatic reactions are involved in the urea cycle, two of which are mi¬  tochondrial and three cytosolic (Fig. 21-9).   1. Carbamoyl Phosphate Synthetase Acquires the First Urea Nitrogen  Atom. Carbamoyl phosphate synthetase (CPS) is technically not a member  of the urea cycle. It catalyzes the condensation and activation of NH 3 and  HC0 3 to form carbamoyl phosphate, the first of the cycles two nitrogen-  containing substrates, with the concomitant cleavage of 2 ATP. Eukaryotes  have two forms of CPS: Mitochondrial CPS I uses ammonia as its nitrogen  donor and participates in urea biosynthesis, whereas cytosolic CPS II uses glu¬  tamine as its nitrogen donor and is involved in pyrimidine biosynthesis    FIG. 21-9 The urea cycle. Five enzymes participate in the  urea cycle: (l) carbamoyl phosphate synthetase, (2) ornithine  transcarbamoylase, (3) argininosuccinate synthetase, (4) argininosuccinase,  and (5) arginase. Enzymes 1 and 2 are mitochondrial and enzymes  3-5 are cytosolic. Ornithine and citrulline must therefore be  transported across the mitochondrial membrane by specific transport  systems (yellow circles ).The urea amino groups arise from the  deamination of amino acids. One amino group (green) originates as  ammonia that is generated by the glutamate dehydrogenase reaction    (top). The other amino group (red) is obtained from aspartate (right),  which is formed via the transamination of oxaloacetate by an amino  acid. The fumarate product of Reaction 4 is converted to the  gluconeogenic precursor oxaloacetate by the action of cytosolic  fumarase and malate dehydrogenase (bottom). Q See the Animated  Figures.    What factors make the reactions of the cycle irreversible  under physiological conditions?    FIG. 21-10 The mechanism of action of CPS I. (l) Phosphorylation activates HC0 3  to form the intermediate carboxyphosphate. ( 2 ) NH 3 attacks carboxyphosphate to form  carbamate. (3) ATP phosphorylates carbamate, yielding carbamoyl phosphate.    (Section 23-2A). CPS I catalyzes an essentially irreversible reaction that is the  rate-limiting step of the urea cycle (Fig. 21-10):   3. A second ATP phosphorylates carbamate to form carbamoyl phosphate  and ADP    In E. coli , a single CPS with the subunit structure (a( 3)4 generates car¬  bamoyl phosphate, using glutamine as its nitrogen donor. The enzymes small  subunit is a glutaminase that hydrolyzes glutamine, and its large subunit cat¬  alyzes Reactions 1, 2, and 3 of Fig. 21-10. The X-ray structure of E. coli CPS,  determined by Hazel Holden and Ivan Rayment, reveals the astonishing fact  that although the three active sites are widely separated in space, they are con¬  nected by a narrow 96-A-long tunnel that runs nearly the entire length of the  protein molecule (Fig. 21-11). The NH 3 produced by the glutaminase active  site travels —45 A in order to react with carboxyphosphate. The resulting car¬  bamate then travels —35 A to reach the carbamoyl phosphate synthesis site.  This phenomenon, in which the intermediate of two reactions is directly trans¬  ferred from one enzyme active site to another, is called channeling.   Channeling increases the rate of a metabolic pathway by preventing the  loss of its intermediate products as well as protecting the intermediates from  degradation. Channeling is critical for CPS because the intermediates car¬  boxyphosphate and carbamate are extremely reactive, having half-lives of 28  and 70 ms, respectively, at neutral pH. Also, channeling allows the local con¬  centration of NH 3 to reach a higher value than is present in the cellular  medium. We will encounter several other examples of channeling in our stud¬  ies of metabolic enzymes, but the CPS tunnel is the longest known.   2. Carbamoylation of Ornithine Produces Citrulline. Ornithine transcar-  bamoylase transfers the carbamoyl group of carbamoyl phosphate to  ornithine, yielding citrulline (Fig. 21-9, Reaction 2). Note that both of the    FIG. 21-11 X-Ray structure of E. coli carbamoyl phosphate synthetase. The   protein is represented by its C a backbone. The small subunit ( magenta ) contains the  glutamine-binding site where NH 3 is produced. The large subunit consists of several  domains (green, yellow, blue, and orange ) and contains the other two active sites. The  96-A-long tunnel connecting the three active sites is outlined in red. [Courtesy of Hazel  Holden and Ivan Rayment, University of Wisconsin. PDBid 1JDB.]    FIG. 21-12 The mechanism of action of argininosuccinate synthetase, (l) The    latter compounds are “nonstandard” a-amino acids that do not occur in pro¬  teins. The transcarbamoylase reaction occurs in the mitochondrion, so  ornithine, which is produced in the cytosol, must enter the mitochondrion via  a specific transport system. Likewise, since the remaining urea cycle reactions  occur in the cytosol, citrulline must be exported from the mitochondrion.   3. Argininosuccinate Synthetase Acquires the Second Urea Nitrogen  Atom. Ureas second nitrogen atom is introduced by the condensation of cit-  rulline’s ureido group with an aspartate amino group by argininosuccinate  synthetase (Fig. 21-12). ATP activates the ureido oxygen atom as a leaving  group through formation of a citrullyl-AMP intermediate, and AMP is sub¬  sequently displaced by the aspartate amino group. The PP Z formed in the re¬  action is hydrolyzed to 2 P z , so the reaction consumes two ATP equivalents.   4. Argininosuccinase Produces Fumarate and Arginine. With the formation  of argininosuccinate, all of the urea molecule components have been assembled.  However, the amino group donated by aspartate is still attached to the aspartate  carbon skeleton. This situation is remedied by the argininosuccinase-catalyzed  elimination of fumarate, leaving arginine (Fig. 21-9, Reaction 4). Arginine is  ureas immediate precursor. The fumarate produced in the argininosuccinase re¬  action is converted to oxaloacetate by the action of fumarase and malate dehy¬  drogenase. These two reactions are the same as those that occur in the citric acid  cycle, although they take place in the cytosol rather than in the mitochondrion.  The oxaloacetate is then used for gluconeogenesis (Section 16-4).   5. Arginase Releases Urea. The urea cycles final reaction is the arginase-  catalyzed hydrolysis of arginine to yield urea and regenerate ornithine  (Fig. 21-9, Reaction 5). Ornithine is then returned to the mitochondrion for  another round of the cycle.   The urea cycle thus converts two amino groups, one from ammonia and  one from aspartate, and a carbon atom from HCO 3 to the relatively nontoxic  product, urea, at the cost of four “high-energy” phosphate bonds. The energy  spent is more than recovered, however, by the oxidation of the carbon skele¬  tons of the amino acids that have donated their amino groups, via transami¬  nation, to glutamate and aspartate. Indeed, half the oxygen that the liver  COO - consumes is used to provide this energy.   l 11 B The Urea Cycle Is Regulated by Substrate Availability   Carbamoyl phosphate synthetase I, which catalyzes the first committed step of  the urea cycle, is allosterically activated by A^acetylglutamate (at left). This  metabolite is synthesized from glutamate and acetyl-CoA by A^acetylglutamate    synthase. When amino acid breakdown rates increase, the concentration of glu¬  tamate increases as a result of transamination. The increased glutamate stimu¬  lates TV-acetylglutamate synthesis. The resulting activation of carbamoyl  phosphate synthetase increases the rate of urea production. Thus, the excess ni¬  trogen produced by amino acid breakdown is efficiently excreted. Note that the  urea cycle, like gluconeogenesis (Section 16-4) and ketogenesis (Section 20-3),  is a pathway that occurs in the liver but serves the needs of the body as a whole.   The remaining enzymes of the urea cycle are controlled by the concentra¬  tions of their substrates. In individuals with inherited deficiencies in urea cy¬  cle enzymes other than arginase, the corresponding substrate builds up,  increasing the rate of the deficient reaction so that the rate of urea produc¬  tion is normal (the total lack of a urea cycle enzyme, however, is lethal). The  anomalous substrate buildup is not without cost, however. The substrate con¬  centrations become elevated all the way back up the cycle to ammonia, result¬  ing in hyperammonemia. Although the root cause of ammonia toxicity is not  completely understood, it is clear that the brain is particularly sensitive to high  ammonia concentrations (symptoms of urea cycle enzyme deficiencies include  mental retardation and lethargy).   • Summarize the steps of the urea cycle.  How do the amino groups of amino acids  enter the cycle?   • How is the rate of amino acid deamination  linked to the rate of the urea cycle?    Alanine, cysteine, glycine, serine, and threonine are broken down to pyruvate.   • a-Ketoglutarate is produced by the degradation of arginine, glutamate, glutamine,  histidine, and proline.   Amino acids are degraded to compounds that can be metabolized to C0 2 and  H 2 0 or used in gluconeogenesis. Indeed, oxidative breakdown of amino acids  typically accounts for 10 to 15% of the metabolic energy generated by ani¬  mals. In this section we consider how the carbon skeletons of the 20 “stan¬  dard” amino acids are catabolized. We do not describe in detail all of the many  reactions involved. Rather, we consider how the pathways are or¬  ganized and focus on a few reactions of chemical and/or medical  interest.   “Standard” amino acids are degraded to one of seven meta¬  bolic intermediates: pyruvate, a-ketoglutarate, succinyl-CoA, fu¬  marate, oxaloacetate, acetyl-CoA, or acetoacetate (Fig. 21-13).   The amino acids can therefore be divided into two groups based  on their catabolic pathways:    1. Glucogenic amino acids, which are degraded to pyru¬  vate, a-ketoglutarate, succinyl-CoA, fumarate, or oxaloac¬  etate and are therefore glucose precursors (Section 16-4).   2. Ketogenic amino acids, which are broken down to  acetyl-Co A or acetoacetate and can thus be converted to  fatty acids or ketone bodies (Section 20-3).    FIG. 21-13 Degradation of amino acids to one of seven common  metabolic intermediates. Glucogenic and ketogenic degradations are  indicated in green and red, respectively.   Chapter 21 Amino Acid Metabolism Some amino acids are precursors of both carbohydrates and ketone bodies.   Since animals lack any metabolic pathways for the net conversion of acetyl-  CoA or acetoacetate to gluconeogenic precursors, no net synthesis of carbohy¬  drates is possible from the purely ketogenic amino acids Lys and Leu.   In studying the specific pathways of amino acid breakdown, we will or¬  ganize the amino acids into groups that are degraded to each of the seven  metabolites mentioned above.   A Alanine, Cysteine, Glycine, Serine, and Threonine Are  Degraded to Pyruvate   Five amino acids—alanine, cysteine, glycine, serine, and threonine—are bro¬  ken down to yield pyruvate (Fig. 21-14). Alanine is straightforwardly transam-  inated to pyruvate. Serine is converted to pyruvate through dehydration by  serine—threonine dehydratase. This PLP-dependent enzyme, like the amino¬  transferases (Section 21-2A), forms a PLP-amino acid Schiff base, which  facilitates the removal of the amino acids a-hydrogen atom. In the  serine-threonine dehydratase reaction, however, the C a carbanion breaks down  with the elimination of the amino acids Cp OH, rather than with tautomer-  ization (Fig. 21-8, Step 2), so that the substrate undergoes a,|3 elimination of  H 2 0 rather than deamination (Fig. 21-15). The product of the dehydration,    FIG. 21-14 The pathways converting  alanine, cysteine, glycine, serine, and  threonine to pyruvate. The enzymes involved are  (l) alanine aminotransferase, (2) serine-threonine  dehydratase, (3) glycine cleavage system, (4 and  5) serine hydroxymethyltransferase, (6) threonine  dehydrogenase, and (7) a-amino-p-ketobutyrate  lyase.   FIG. 21-15 The serine-threonine dehydratase reaction. This  PLP-dependent enzyme catalyzes the elimination of water from serine  in six steps: (l) formation of a serine-PLP Schiff base, (2) removal of  the a-H atom of serine to form a resonance-stabilized carbanion,   (3) p elimination of OH - , (4) hydrolysis of the Schiff base to yield the   PLP-enzyme and aminoacrylate, (5) nonenzymatic tautomerization to  the imine, and ( 6 ) nonenzymatic hydrolysis to form pyruvate and  ammonia. Threonine undergoes an analogous series of reactions to  yield a-ketobutyrate.    the enamine aminoacrylate, tautomerizes nonenzymatically to the corre¬  sponding imine, which spontaneously hydrolyzes to pyruvate and ammonia.   Cysteine can be converted to pyruvate via several routes in which the  sulfhydryl group is released as H 2 S, SO 3 , or SCN .   Glycine is converted to pyruvate by first being converted to serine by the  enzyme serine hydroxymethyltransferase, another PLP-containing enzyme  (Fig. 21-14, Reaction 4). This enzyme uses Af 5 ,Af 10 -methylenetetrahydro-  folate (A^,7V 10 -methylene-THF) as a one-carbon donor (the structure and  chemistry of THF cofactors are described in Section 21-4D). The methylene  group of the THF cofactor is obtained from a second glycine in Reaction 3  of Fig. 21-14, which is catalyzed by the glycine cleavage system (called the  glycine decarboxylase multienzyme system in plants). This enzyme is a mul¬  tiprotein complex that resembles pyruvate dehydrogenase (Section 17-2). The  glycine cleavage system mediates the major route of glycine degradation in  mammalian tissues. An inherited deficiency of the glycine cleavage system  causes the disease nonketotic hyperglycinemia, which is characterized by men¬  tal retardation and accumulation of large amounts of glycine in body fluids.   In mammals other than humans, threonines major route of breakdown is  through threonine dehydrogenase (Fig. 21-14, Reaction 6 ), producing  a-amino-(5-ketobutyrate, which is converted to acetyl-CoA and glycine by  a-amino-(5-ketobutyrate lyase (Fig. 21-14, Reaction 7). The glycine can be  converted, through serine, to pyruvate as described above. Thus, threonine is  both glucogenic and ketogenic.   Serine Hydroxymethyltransferase Catalyzes PLP-Dependent C a —C p Bond  Formation and Cleavage. Threonine can also be converted directly to glycine  and acetaldehyde (which is subsequently oxidized to acetyl-CoA) via Reaction  5 of Fig. 21-14, which breaks threonines C a —C p bond. This PLP-dependent  reaction is catalyzed by serine hydroxymethyltransferase, the same enzyme that  adds a hydroxymethyl group to glycine to produce serine (Fig. 21-14, Reaction 4).  In the glycine —> serine reaction, the amino acids C a —H bond is cleaved (as  occurs in transamination; Fig. 21-8) and a C a —C p bond is formed. In contrast,    FIG. 21-16 The TT-orbital framework of a  PLP-amino acid Schiff base. The bond from X  to C a is in a plane perpendicular to the plane of  the PLP TT-orbital system (top) and is therefore  labile. The broken bond’s electron pair ( bottom )  is delocalized over the conjugated molecule.   the degradation of threonine to glycine by serine hydroxymethyltransferase acts  in reverse, beginning with C a —Cp bond cleavage (at left).   With the cleavage of any of the bonds to C a , the PLP group delocalizes  the electrons of the resulting carbanion. This feature of PLP action is the key  to understanding how the same amino acid-PLP Schiff base can undergo  cleavage of different bonds to C a in different enzymes (bonds a, b, or c in the  upper right of Fig. 21-8). The bond that is cleaved is the one that lies in the  plane perpendicular to that of the TT-orbital system of the PLP (Fig. 21-16).  This arrangement allows the PLP TT-orbital system to overlap the bonding or¬  bital containing the electron pair being delocalized. Any other geometry would  reduce or even eliminate this orbital overlap (the newly formed double bond  would be twisted out of planarity) yielding a higher energy arrangement.  Different bonds to C a can be positioned for cleavage by rotation around the  C a —N bond. Evidently, each enzyme binds its amino acid-PLP Schijf base  adduct with the appropriate geometry for bond cleavage.   Transamination of aspartate leads directly to oxaloacetate. Asparagine is also  converted to oxaloacetate in this manner after its hydrolysis to aspartate by   Interestingly, l- asparaginase is an effective chemotherapeutic agent in the treat- Section 4 Breakdown of Amino Acids   ment of cancers that must obtain asparagine from the blood, particularly acute  lymphoblastic leukemia.    C Arginine, Glutamate, Glutamine, Histidine, and Proline  Are Degraded to a-Ketoglutarate    Arginine, glutamine, histidine, and proline are all degraded by conversion to glu¬  tamate (Fig. 21-17), which in turn is oxidized to a-ketoglutarate by glutamate  dehydrogenase (Section 21 - 2 B). Conversion of glutamine to glutamate involves  only one reaction: hydrolysis by glutaminase. In the kidney, the action of glut-  aminase produces ammonia, which combines with a proton to form the ammo¬  nium ion (NH^) and is excreted. During metabolic acidosis (Box 2-2), kidney  glutaminase helps eliminate excess acid. Although free NH 3 in the blood could  serve the same acid-absorbing purpose, ammonia is toxic. It is therefore converted  to glutamine by glutamine synthetase in the liver (Section 21-5A). Glutamine    FIG. 21-17 The degradation of arginine, glutamate,  glutamine, histidine, and proline to u-ketoglutarate. The enzymes  catalyzing the reactions are (l) glutamate dehydrogenase,    (7) spontaneous, (8) histidine ammonia lyase, (9) urocanate hydratase,  ( 10 ) imidazolone propionase, and ( 11 ) glutamate formiminotransferase.   Chapter 21 Amino Acid Metabolism therefore acts as an ammonia transport system between the liver, where much of   it is synthesized, and the kidneys, where it is hydrolyzed by glutaminase.   Histidines’s conversion to glutamate is more complicated: It is nonoxida-  tively deaminated, then it is hydrated, and its imidazole ring is cleaved to form  A^formiminoglutamate. The formimino group is then transferred to tetrahy-  drofolate, forming glutamate and Af 5 -formiminotetrahydrofolate (Section  21-4D). Both arginine and proline are converted to glutamate through the in¬  termediate formation of glutamate-5-semialdehyde.   Methionine, threonine, isoleucine, and valine have degradative pathways that  all yield propionyl-CoA, which is also a product of odd-chain fatty acid degra¬  dation. Propionyl-CoA is converted to succinyl-CoA by a series of reactions  requiring biotin and coenzyme B 12 (Section 20-2E).   Methionine Breakdown Involves Synthesis of S-Adenosylmethionine and  Cysteine. Methionine degradation (Fig. 21-18) begins with its reaction with  ATP to form S-adenosylmethionine (SAM; alternatively AdoMet). This sul-  fonium ions highly reactive methyl group makes it an important biological methy¬  lating agent. For instance, SAM is the methyl donor in the synthesis of  phosphatidylcholine from phosphatidylethanolamine (Section 20-6A).   Donation of a methyl group from SAM leaves S-adenosylhomocysteine,  which is then hydrolyzed to adenosine and homocysteine. The homocysteine  can be methylated to re-form methionine via a reaction in which N 5 -  methyltetrahydrofolate (see below) is the methyl donor. Alternatively, the ho¬  mocysteine can combine with serine to yield cystathionine, which subsequently  forms cysteine (cysteine biosynthesis) and a-ketobutyrate. The a-ketobutyrate  continues along the degradative pathway to propionyl-CoA and then succinyl-  CoA. High homocysteine levels are associated with disease (Box 21-1).    The cellular level of homocysteine depends on its rate of synthe¬  sis through methylation reactions utilizing SAM (Fig. 21-18, Reac¬  tions 2 and 3) and its rate of utilization through remethylation to  form methionine (Fig. 21-18, Reaction 4) and reaction with ser¬  ine to form cystathionine in the cysteine biosynthetic pathway  (Fig. 21-18, Reaction 5). An increase in homocysteine levels  leads to hyperhomocysteinemia, elevated concentrations of  homocysteine in the blood, which is associated with cardiovascu¬  lar disease. The link was first discovered in individuals with  homocysteinuria, a disorder in which excess homocysteine is  excreted in the urine. These individuals develop atherosclerosis as  children, possibly because homocysteine causes oxidative damage  to the walls of blood vessels even in the absence of elevated  LDL levels (Section 20-7C).   Hyperhomocysteinemia is also associated with neural tube  defects, the cause of a variety of severe birth defects including  spina bifida (defects in the spinal column that often result in  paralysis) and anencephaly (the invariably fatal failure of the brain  to develop, which is the leading cause of infant death due to  congenital anomalies). Hyperhomocysteinemia is readily controlled  by ingesting the vitamin precursors of the coenzymes that partici¬  pate in homocysteine breakdown, namely, B 6 (pyridoxine, the PLP  precursor; Fig. 21-7), B 12 (Fig. 20-17), and folate (Section 21-4D).    Folate, especially, alleviates hyperhomocysteinemia; its administra¬  tion to pregnant women dramatically reduces the incidence  of neural tube defects in newborns. Because neural tube  development is one of the earliest steps of embryogenesis,  women of childbearing age are encouraged to consume adequate  amounts of folate even before they become pregnant. Despite the  foregoing, clinical trials indicate that vitamin therapy in individuals  with hyperhomocysteinemia does not reduce their incidence of  cardiovascular disease.   Around 10% of the population is homozygous for an Ala —>  Val mutation in yV 5 ,W 10 -methylenetetrahydrofolate reductase  (MTHFR), which catalyzes the conversion of A/ 5 ,/V 10 -methylene-  THF to A/ 5 -methyl-THF (Fig. 21-20, top center). This reaction  generates the A/ 5 -methyl-THF required to convert homocysteine  to methionine (Fig. 21-18, Reaction 4). The mutation does not  affect the enzyme’s reaction kinetics but instead increases the  rate at which its essential flavin cofactor dissociates. Folate  derivatives that bind to the enzyme decrease the rate of flavin  loss, thus increasing the enzyme’s overall activity and decreasing  the homocysteine concentration. The prevalence of the MTHFR  mutation in the human population suggests that it has (or once  had) some selective advantage; however, this is as yet a matter  of speculation.   breakdown of methionine yields cysteine and succinyl-CoA. The  enzymes are (l) methionine adenosyltransferase in a reaction that  yields the biological methylating agent S-adenosylmethionine (SAM),  (2) methyltransferase, (3) adenosylhomocysteinase, (4) methionine  synthase (a coenzyme B 12 -dependent enzyme), (5) cystathionine   (9) methylmalonyl-CoA racemase, and (10) methylmalonyl-CoA mutase  (a coenzyme B 12 -dependent enzyme). Reactions 8-10 are discussed  in Section 20-2E. Threonine is converted to a-ketobutyrate by  ( 11 ) serine-threonine dehydratase.    Threonine Is Mainly Degraded to Propionyl-CoA in Humans. The reactions  shown in Fig. 21-14 account for only ~20% of threonine breakdown in hu¬  mans. The remainder, as is indicated in Fig. 21-18, Reaction 11, forms  a-ketobutyrate in a reaction catalyzed by serine—threonine dehydratase  (Fig. 21-15), and the a-ketobutyrate is converted via propionyl-CoA to  succinyl-CoA.   Tetrahydrofolates Are One-Carbon Carriers. Many biosynthetic processes in¬  volve the addition of a C x unit to a metabolic precursor. In most carboxylation   FIG. 21 -19 The two-stage reduction of folate to THF. Both reactions are catalyzed by dihydrofolate reductase.    reactions (e.g., pyruvate carboxylase; Fig. 16-18), the enzyme uses a biotin co¬  factor. In some reactions, S-adenosylmethionine (Fig. 21-18) functions as a  methylating agent. However, tetrahydrofolate (THF) is more versatile than ei¬  ther of those cofactors because it can transfer C x units in several oxidation states.   THF is a 6-methylpterin derivative linked in sequence to a />-aminoben-  zoic acid and a Glu residue:    Up to five additional Glu residues are linked to the first glutamate via isopep¬  tide bonds to form a polyglutamyl tail. THF is derived from the vitamin folic  acid (Latin: folium , leaf), a doubly oxidized form of THF that must be enzy¬  matically reduced before it becomes an active coenzyme (Fig. 21-19). Both  reductions are catalyzed by dihydrofolate reductase (DHFR). Mammals can¬  not synthesize folic acid, so it must be provided in the diet or by intestinal  microorganisms.   Ci units are covalently attached to THF at positions N5, N10, or both  N5 and N10. These C x units, which may be at the oxidation levels of for¬  mate, formaldehyde, or methanol (Table 21-2), are all interconvertible by en¬  zymatic redox reactions (Fig. 21-20).   THF acquires C x units in the conversion of serine to glycine by serine hy-  droxymethyltransferase (the reverse of Reaction 4, Fig. 21-14), in the cleavage  of glycine (Fig. 21-14, Reaction 3), and in histidine breakdown (Fig. 21-17,  Reaction 11). The C x units carried by THF are used in the synthesis of  thymine nucleotides (Section 23-3B) and in the synthesis of methionine from  homocysteine (Fig. 21-18, Reaction 4). By promoting the latter process, sup¬  plemental folate helps prevent diseases associated with abnormally high levels  of homocysteine (Box 21-1).   r Sulfonamides (sulfa drugs) such as sulfanilamide are antibiotics that are  structural analogs of the y?-aminobenzoic acid constituent of THF (at left).  They competitively inhibit bacterial synthesis of THF at the y>-aminobenzoic acid  incorporation step, thereby blocking THF-requiring reactions. The inability of  mammals to synthesize folic acid leaves them unaffected by sulfonamides,  which accounts for the medical utility of these widely used antibacterial agents.   FIG. 21-21 The degradation of the  branched-chain amino acids. Isoleucine (A),  valine (B), and leucine (C) follow an initial  common pathway utilizing three enzymes:   ( 2 ) branched-chain a-keto acid dehydrogenase  (BCKDH), and (3) acyl-CoA dehydrogenase.  Isoleucine degradation then continues (left) to  yield acetyl-CoA and succinyl-CoA; valine degra¬  dation continues ( center ) to yield succinyl-CoA;  and leucine degradation continues (right) to yield  acetyl-CoA and acetoacetate.    (A) Isoleucine: R x = CH 3 —, R 2 = CH 3 —CH 2 -   (B) Valine : R x = CH 3 —, R 2 = CH 3 —   (C) Leucine : R x = H— , R 2 = (CH 3 ) 2 CH —   Degradation of the branched-chain amino acids isoleucine, leucine, and valine  begins with three reactions that employ common enzymes (Fig. 21-21):   The remaining reactions of the isoleucine degradation pathway are analogous  to those of fatty acid oxidation (Section 20-2C), thereby yielding acetyl-CoA  and propionyl-CoA, which is subsequently converted via three reactions to  succinyl-CoA. The degradation of valine, which contains one less carbon than  isoleucine, yields C0 2 and propionyl-CoA, which is then converted to  succinyl-CoA. The further degradation of leucine, which yields acetoacetate  instead of propionyl-CoA, is considered in Section 21-4E.   Branched-chain a-keto acid dehydrogenase (BCKDH), which catalyzes  Reaction 2 of Fig. 21-21, is a multienzyme complex that closely resembles the  pyruvate dehydrogenase and a-ketoglutarate dehydrogenase complexes (Sections  17-2 and 17-3D). Indeed, all three multienzyme complexes share a common  subunit, E3 (dihydrolipoamide dehydrogenase), and employ the coenzymes TPP,  lipoamide, and FAD in addition to their terminal oxidizing agent, NAD + .   T A genetic deficiency in BCKDH causes maple syrup urine disease, so named  because the consequent buildup of branched-chain a-keto acids imparts the   urine with the characteristic odor of maple syrup. Unless promptly treated by a diet  low in branched-chain amino acids, maple syrup urine disease is rapidly fatal.    E Leucine and Lysine Are Degraded Only to Acetyl-CoA  and/or Acetoacetate   Leucine degradation begins in the same manner as isoleucine and valine degra¬  dation (Fig. 21-21), but the dehydrogenated CoA adduct (3-methylcrotonyl-  CoA is converted to acetyl-CoA and acetoacetate, a ketone body.   The predominant pathway for lysine degradation in mammalian liver  produces acetoacetate and 2 C0 2 via the initial formation of the  a-ketoglutarate-lysine adduct saccharopine (Fig. 21-22). This pathway is worth    FIG. 21-22 The pathway of  lysine degradation in  mammalian liver. The enzymes are   ? Identify the reactions that  resemble those of fatty  acid oxidation and  ketogenesis.   Chapter 21 Amino Acid Metabolism examining in detail because we have encountered 7 of its 11 reactions in other   pathways. Reaction 4 is a PLP-dependent transamination. Reaction 5 is the  oxidative decarboxylation of an a-keto acid by a multienzyme complex simi¬  lar to pyruvate dehydrogenase (Section 17-2). Reactions 6, 8, and 9 are stan¬  dard reactions of fatty acyl-CoA oxidation: dehydrogenation by FAD,  hydration, and dehydrogenation by NAD + . Reactions 10 and 11 are standard  reactions in ketone body formation. Two moles of C0 2 are produced at  Reactions 5 and 7 of the pathway.   The saccharopine pathway is thought to predominate in mammals because  a genetic defect in the enzyme that catalyzes Reaction 1 in the sequence re¬  sults in hyperlysinemia and hyperlysinuria (elevated levels of lysine in the  blood and urine, respectively) along with mental and physical retardation. This  is yet another example of how the study of rare inherited disorders has helped  to trace metabolic pathways.   The complexity of the major tryptophan degradation pathway (outlined in  Fig. 21-23) precludes a detailed discussion of all its reactions. However, one  reaction is of particular interest: The fourth reaction is catalyzed by kynuren¬  inase, whose PLP group facilitates cleavage of the Cp—C 7 bond to release  alanine. The kynureninase reaction follows the same initial steps as transami¬  nation (Fig. 21-8), but an enzyme nucleophilic group then attacks C 7 of the  resonance-stabilized intermediate, resulting in Cp—C 7 bond cleavage. The  remainder of the tryptophan skeleton is converted in five reactions to  a-ketoadipate, which is also an intermediate in lysine degradation.  a-Ketoadipate is broken down to 2 C0 2 and acetoacetate in seven reactions,  as shown in Fig. 21-22.   enzymatic reactions shown are (l) tryptophan-2,3-dioxygenase, to a-ketoadipate, which is converted to acetyl-CoA and acetoacetate   (2) formamidase, (3) kynurenine-3-monooxygenase, and (4) kynureninase in seven reactions as shown in Fig. 21-22, Reactions 5-11.   Since the first reaction in phenylalanine degradation is its hydroxylation to ty¬  rosine, a single pathway (Fig. 21-24) is responsible for the breakdown of both  amino acids. The final products of the six-reaction degradation are fumarate,  a citric acid cycle intermediate, and acetoacetate, a ketone body. Defects in  the enzymes that catalyze Reactions 1 and 4 cause disease (Box 21-2).   Pterins Are Redox Cofactors. The hydroxylation of phenylalanine by the  Fe(III)-containing enzyme phenylalanine hydroxylase (Fig. 21-24, Reaction  1 ) requires the cofactor biopterin, a pterin derivative. Pterins are compounds  that contain the pteridine ring (Fig. 21-25). Note the resemblance between   Note the similar structures of pteridine  and the isoalloxazine ring of flavin  coenzymes.    Archibald Garrod realized in the early 1900s that human genetic  diseases result from specific enzyme deficiencies. We have re¬  peatedly seen how this realization has contributed to the elucida¬  tion of metabolic pathways. The first such disease Garrod  recognized was alcaptonuria, which results in the excretion of  large quantities of homogentisic acid (Section 14-4). This condi¬  tion results from deficiency of homogentisate dioxygenase  (Fig. 21-24, Reaction 4). Alcaptonurics suffer no ill effects other  than arthritis later in life (although their urine darkens alarmingly  on exposure to air because of the rapid oxidation of the  homogentisate they excrete).   Individuals suffering from phenylketonuria (PKU) are not so  fortunate. Severe mental retardation occurs within a few months  of birth if the disease is not detected and treated immediately.  PKU is caused by the inability to hydroxylate phenylalanine  (Fig. 21-24, Reaction 1) and therefore results in increased blood  levels of phenylalanine (hyperphenylalaninemia). The excess  phenylalanine is transaminated to phenylpyruvate    with the disease and gave the disease its name. All babies born in  the United States are now screened for PKU immediately after  birth by testing for elevated levels of phenylalanine in the blood.   Classical PKU results from a deficiency in phenylalanine hy¬  droxylase. When this was established in 1947, it was the first  inborn error of metabolism whose basic biochemical defect had  been identified. Since all of the tyrosine breakdown enzymes are  normal, treatment consists in providing the patient with a low-  phenylalanine diet and monitoring the blood level of phenylalanine  to ensure that it remains within normal limits for the first 5 to 10  years of life (the adverse effects of hyperphenylalaninemia seem  to disappear after that age). Aspartame (NutraSweet), an often  used sweetening ingredient in diet soft drinks and many other  dietetic food products, is Asp-Phe-methyl ester (Box 8-2) and is  therefore a source of dietary phenylalanine. Consequently, a warn¬  ing label for phenylketonurics appears on all those products.   Phenylalanine hydroxylase deficiency also accounts for another  common symptom of PKU: Its victims have lighter hair and skin  color than their siblings. This is because elevated phenylalanine  levels inhibit tyrosine hydroxylation (Fig. 21-39), the first reaction  in the formation of the skin pigment melanin.   Other variants of hyperphenylalaninemia have been discovered  since the introduction of infant screening techniques. These are  caused by deficiencies in the enzymes catalyzing the formation or  regeneration of 5,6,7,8-tetrahydrobiopterin, the phenylalanine  hydroxylase cofactor (Fig. 21-26).   • Describe the two general metabolic fates  of the carbon skeletons of amino acids.   • List the seven metabolites that represent  the end products of amino acid catabo¬  lism. Which are glucogenic? Which are  ketogenic?   • Which three amino acids are substrates or  products of serine hydroxymethyltrans-  ferase?   • Describe the roles of the cofactors  pyridoxal-5'-phosphate, tetrahydrofolate,  and tetrahydrobiopterin in the catabolism  of amino acids.    the pteridine ring and the isoalloxazine ring of the flavin coenzymes; the  positions of the nitrogen atoms in pteridine are identical to those of the B  and C rings of isoalloxazine. Folate derivatives also contain the pterin ring  (Section 21-4D).   Pterins, like flavins, participate in biological oxidations. The active form  of biopterin is the fully reduced form, 5,6,7,8-tetrahydrobiopterin. It is pro¬  duced from 7,8-dihydrobiopterin and NADPH, in what may be considered  a priming reaction, by dihydro folate reductase (Fig. 21-26), which also re¬  duces folate to dihydrofolate and then to tetrahydrofolate (Fig. 21-19). In the  phenylalanine hydroxylase reaction, 5,6,7,8-tetrahydrobiopterin is hydroxy-  lated to pterin-4a-carbinolamine (Fig. 21-26), which is then converted to  7 ,8-dihydrobiopterin (quinoid form) by pterin-4a-carbinolamine dehy¬  dratase. This quinoid is subsequently reduced by the NAD(P)H-requiring en¬  zyme dihydropteridine reductase to regenerate the active cofactor. Although  dihydrofolate reductase and dihydropteridine reductase produce the same  product, they use different tautomers of the substrate.    Some amino acids are synthesized in one or a few steps from common metabolites.  The essential amino acids are mostly derived from other amino acids and glucose  metabolites.    Many amino acids are synthesized by pathways that are present only in plants and  microorganisms. Since mammals must obtain these amino acids in their diets,  these substances are known as essential amino acids. The other amino acids,   FIG. 21-26 The formation, utilization, and regeneration of 5,6,7,8-  tetrahydrobiopterin in the phenylalanine hydroxylase reaction.    which can be synthesized by mammals from common intermediates, are  termed nonessential amino acids. Their a-keto acid carbon skeletons are con¬  verted to amino acids by transamination reactions (Section 21-2A) utilizing  the preformed a-amino nitrogen of another amino acid, usually glutamate.  Although it was originally presumed that glutamate can be synthesized from  ammonia and a-ketoglutarate by GDH acting in reverse, it now appears that  the predominant physiological direction of this enzyme is glutamate break¬  down (Section 21-2B). Consequently, preformed a-amino nitrogen should also  be considered to be an essential nutrient. In this context, it is interesting to note  that, in addition to the four well-known taste receptors, those for sweet, sour,  salty, and bitter tastes, a fifth taste receptor has been characterized, that for  the meaty taste of monosodium glutamate (MSG), which is known as  umami (a Japanese name).   The essential and nonessential amino acids for humans are listed in  Table 21-3. Arginine is classified as essential, even though it is synthesized by  the urea cycle (Section 21-3A), because it is required in greater amounts than  can be produced by that route during the normal growth and development of  children (but not adults). The essential amino acids occur in animal and veg¬  etable proteins. Different proteins, however, contain different proportions of  the essential amino acids. Milk proteins, for example, contain them all in the  proportions required for proper human nutrition. Bean protein, on the other  hand, contains an abundance of lysine but is deficient in methionine, whereas  wheat is deficient in lysine but contains ample methionine. A balanced protein    ^Although mammals synthesize arginine, they cleave  most of it to form urea (Section 21-3A).    diet therefore must contain a variety of different protein sources that comple¬  ment each other to supply the proper amounts of all the essential amino acids.   In this section we study the pathways involved in the formation of the  nonessential amino acids. We also briefly consider such pathways for the essen¬  tial amino acids as they occur in plants and microorganisms. Keep in mind that  there is considerable variation in these pathways among different species. In contrasty  the basic pathways of carbohydrate and lipid metabolism are all but universal   All the nonessential amino acids except tyrosine are synthesized by simple  pathways leading from one of four common metabolic intermediates: pyru¬  vate, oxaloacetate, a-ketoglutarate, and 3-phosphoglycerate. Tyrosine, which is  really misclassified as being nonessential, is synthesized by the one-step hy-  droxylation of the essential amino acid phenylalanine (Fig. 21 -24). Indeed, the  dietary requirement for phenylalanine reflects the need for tyrosine as well.  The presence of dietary tyrosine therefore decreases the need for phenylalanine.   Alanine, Asparagine, Aspartate, Glutamate, and Glutamine Are Synthesized  from Pyruvate, Oxaloacetate, and a-Ketoglutarate. Pyruvate, oxaloacetate,  and a-ketoglutarate are the a-keto acids (the so-called carbon skeletons) that  correspond to alanine, aspartate, and glutamate, respectively. Indeed, the  synthesis of each of the amino acids is a one-step transamination reaction  (Fig. 21-27, Reactions 1-3). The ultimate source of the a-amino group in    FIG. 21 -27 The syntheses of alanine, aspartate,  glutamate, asparagine, and glutamine. These  reactions involve, respectively, transamination of  (l) pyruvate, (2) oxaloacetate, and (3) a-ketoglutarate,  and amidation of (4) aspartate and (5) glutamate.    ** How many types of reactions are required to  synthesize these five amino acids?   ganisms, plants, and lower eukaryotes by glutamate synthase (Section 21-7),  an enzyme that is absent in vertebrates.   Asparagine and glutamine are, respectively, synthesized from aspartate and  glutamate by ATP-dependent amidation. In the glutamine synthetase reac¬  tion (Fig. 21-27, Reaction 5), glutamate is first activated by reaction with ATP  to form a 7 -glutamylphosphate intermediate. NH 4 then displaces the phos¬  phate group to produce glutamine. Curiously, aspartate amidation by  asparagine synthetase to form asparagine follows a different route; it uses glu¬  tamine as its amino group donor and cleaves ATP to AMP + PP Z (Fig. 21-27,   Glutamine is the amino group donor in the formation of many biosynthetic  products as well as being a storage form of ammonia. The control of glutamine  synthetase is therefore vital for regulating nitrogen metabolism. Mammalian  glutamine synthetases are activated by a-ketoglutarate, the product of gluta¬  mates oxidative deamination (Section 21 - 2 B). This control presumably helps  prevent the accumulation of the ammonia produced by that reaction.   Bacterial glutamine synthetase, as Earl Stadtman showed, has a much more  elaborate control system. The enzyme, which consists of 12 identical 469-  residue subunits arranged at the corners of a hexagonal prism (Fig. 21-28), is  regulated by several allosteric effectors as well as by covalent modification.  Several aspects of its control system bear note. Nine allosteric feedback in¬  hibitors, each with its own binding site, control the activity of bacterial glutamine  synthetase in a cumulative manner. Six of these effectors—histidine, tryptophan,  carbamoyl phosphate (as synthesized by carbamoyl phosphate synthetase),  glucosamine- 6 -phosphate, AMP, and CTP—are all end products of pathways  leading from glutamine. The other three—alanine, serine, and glycine—reflect  the cells nitrogen level.    FIG. 21 -28 X-Ray structure of glutamine synthetase from  the bacterium Salmonella typhimurium. The enzyme consists of 12  identical subunits, here drawn in worm form, arranged with D 6  symmetry (the symmetry of a hexagonal prism), (a) View down the  sixfold axis of symmetry showing only the six subunits of the upper  ring in alternating blue and green. The subunits of the lower ring are  roughly directly below those of the upper ring. A pair of Mn 2+ ions  (magenta spheres) that occupy the positions of the Mg 2+ ions    required for enzymatic activity are bound in each active site. Each  adenylylation site, Tyr 397 (orange), lies between two subunits. Also  drawn in one active site are ADP (cyan) and phosphinothricin (red), a  competitive inhibitor of glutamate, (b) Side view along one of the  twofold axes showing only the eight nearest subunits. The sixfold axis  is vertical in this view. [Based on an X-ray structure by David  Eisenberg, UCLA. PDBid 1 FPYJ   dition of an AMP group) of a specific Tyr residue (Fig. 21-29). The enzymes  susceptibility to cumulative feedback inhibition increases, and its activity  therefore decreases, with its degree of adenylylation. The level of adenylylation  is controlled by a complex metabolic cascade that is conceptually similar to  that controlling glycogen phosphorylase (Section 16-3B). Both adenylylation  and deadenylylation of glutamine synthetase are catalyzed by adenylyltrans-  ferase in complex with a tetrameric regulatory protein, P n . This complex  deadenylylates glutamine synthetase when P n is uridylylated (also at a Tyr  residue) and adenylylates glutamine synthetase when P n lacks UMP residues.  The level of P n uridylylation, in turn, depends on the relative levels of two  enzymatic activities located on the same protein: a uridylyltransferase that   FIG. 21-29 The regulation of bacterial glutamine synthetase. relative activities of uridylyltransferase, which is sensitive to the levels  The adenylylation/deadenylylation of a specific Tyr residue is of a variety of nitrogen metabolites, and uridylyl-removing enzyme,   controlled by the level of uridylylation of a specific adenylyltransferase • whose activity is independent of those metabolite levels.   Pn Tyr residue. This uridylylation level, in turn, is controlled by the    uridylylates P n and a uridylyl-removing enzyme that hydrolytically excises  the attached UMP groups of P n . The uridylyltransferase is activated by a-  ketoglutarate and ATP and inhibited by glutamine and P z , whereas uridylyl-  removing enzyme is insensitive to those metabolites. This intricate metabolic  cascade therefore renders the activity of E. coli glutamine synthetase extremely  responsive to the cells nitrogen requirements.   Glutamate Is the Precursor of Proline, Ornithine, and Arginine. Conversion  of glutamate to proline (Fig. 21-30, Reactions 1-4) involves the reduction of  the y-carboxyl group to an aldehyde followed by the formation of an internal  Schiff base whose further reduction yields proline. Reduction of the glutamate  y-carboxyl group to an aldehyde is an endergonic process that is facilitated by  first phosphorylating the carboxyl group in a reaction catalyzed by y-glutamyl  kinase. The unstable product, glutamate-5-phosphate, has not been isolated  from reaction mixtures but is presumed to be the substrate for the reduction  that follows. The resulting glutamate-5-semialdehyde (which is also a prod¬  uct of arginine and proline degradation; Fig. 21-17) cyclizes spontaneously to    FIG. 21 -30 The biosynthesis of the glutamate family of  amino acids: arginine, ornithine, and proline. The catalysts for  proline biosynthesis are (l) 'y-glutamyl kinase, (2) dehydrogenase,  (3) nonenzymatic, and (4) pyrroline-5-carboxylate reductase. In  mammals, ornithine is produced from glutamate-5-semialdehyde by    the action of ornithine-8-aminotransferase (5). Ornithine is converted  to arginine via the urea cycle (Section 21-3A).    What are the sources of free energy for these biosynthetic  reactions?    FIG. 21-31 The conversion of 3-phosphoglycerate to serine. The pathway  enzymes are (l) 3-phosphoglycerate dehydrogenase, (2) a PLP-dependent aminotransferase,  and (3) phosphoserine phosphatase.    form the internal Schiff base A 1 -pyrroline-5-carboxylate. The final reduction  to proline is catalyzed by pyrroline-5-carboxylate reductase. Whether the en¬  zyme requires NADH or NADPH is unclear.   In humans, a three-step pathway leads from glutamate to ornithine via a  branch from proline biosynthesis after Step 2 (Fig. 21-30). Glutamate-5-  semialdehyde, which is in equilibrium with A 1 -pyrroline- 5 -carboxylate, is directly  transaminated to yield ornithine in a reaction catalyzed by ornithine-8-amino  transferase (Fig. 21-30, Reaction 5). Ornithine is converted to arginine by  the reactions of the urea cycle (Fig. 21-9).   Serine is formed from the glycolytic intermediate 3 -phosphoglycerate in a  three-reaction pathway (Fig. 21-31):   1. Conversion of 3-phosphoglycerate s 2 -OH group to a ketone, yielding  3 -phosphohydroxypyruvate, serine’s phosphorylated keto acid analog.   1. Direct conversion of serine to glycine by serine hydroxymethyltrans-  ferase in a reaction that also yields W 5 ,W 10 -methylene-THF (Fig. 21-14,  Reaction 4 in reverse).   2 . Condensation of the 7V 5 ,W 10 -methylene-THF with C0 2 and NH 4  by glycine synthase (Fig. 21-14, Reaction 3 in reverse).   In animals, cysteine is synthesized from serine and homocysteine, a break¬  down product of methionine (Fig. 21-18, Reactions 5 and 6 ). Homocysteine  combines with serine to yield cystathionine, which subsequently forms cys¬  teine and a-ketobutyrate. Since cysteines sulflhydryl group is derived from the  essential amino acid methionine, cysteine can be considered to be an essential  amino acid.   Essential amino acids, like nonessential amino acids, are synthesized from  familiar metabolic precursors. Their synthetic pathways are present only in mi¬  croorganisms and plants, however, and usually involve more steps than those  of the nonessential amino acids. The enzymes that synthesize essential amino  acids were apparently lost early in animal evolution, possibly because of the  ready availability of these amino acids in the diet. We will focus on only a few  of the many reactions in the biosynthesis of essential amino acids.   bacteria, aspartate is the common precursor of lysine, methionine, and threo¬  nine (Fig. 21-32). The biosyntheses of these essential amino acids all begin    FIG. 21 -32 The biosynthesis of the aspartate family of amino acids: lysine,  methionine, and threonine. The pathway enzymes shown are (l) aspartokinase, (2) p-  aspartate semialdehyde dehydrogenase, (3) homoserine dehydrogenase, and (4) methionine  synthase (a coenzyme B 12 -dependent enzyme).    with the aspartokinase-catalyzed phosphorylation of aspartate to yield  aspartyl-P-phosphate. We have seen that the control of metabolic pathways  commonly occurs at the first committed step of the pathway. One might there¬  fore expect lysine, methionine, and threonine biosynthesis to be controlled as  a group. Each of these pathways is, in fact, independently controlled. E. coli  has three isozymes of aspartokinase that respond differently to the three amino  acids in terms both of feedback inhibition of enzyme activity and repression  of enzyme synthesis. In addition, the pathway direction is controlled by feed¬  back inhibition at the branch points by the amino acid products of the  branches.   catalyzes the methylation of homocysteine to form methionine using 7V 5 -  methyl-THF as its methyl group donor (Reaction 4 in both Figs. 21-18 and  21-32). Methionine synthase is the only coenzyme B 12 —associated enzyme in  mammals besides methylmalonyl-CoA mutase (Section 20-2E). However, the  coenzyme B 12 ’s Co ion in methionine synthase is axially liganded by a methyl    group to form methylcobalamin rather than by a 5 r -adenosyl group as in  methylmalonyl-CoA mutase (Fig. 20-17). In mammals, the primary function  of methionine synthase is not de novo methionine synthesis, as Met is an es¬  sential amino acid. Instead, it functions in the cyclic synthesis of SAM for use  in biological methylations (Fig. 21-18).   Leucine, Isoleucine, and Valine Are Derived from Pyruvate. Valine and  isoleucine follow the same biosynthetic pathway utilizing pyruvate as a starting  reactant, the only difference being in the first step of the series (Fig. 21-33).  In this thiamine pyrophosphate—dependent reaction, which resembles those cat¬  alyzed by pyruvate decarboxylase (Fig. 15-20) and transketolase (Fig. 15-32),  pyruvate forms an adduct with TPP that is decarboxylated to hydroxyethyl-  TPP. This resonance-stabilized carbanion adds either to the keto group of a sec¬  ond pyruvate to form acetolactate on the way to valine, or to the keto group  of a-ketobutyrate to form cy-aceto-cy-hydroxybutyrate on the way to  isoleucine. The leucine biosynthetic pathway branches off from the valine path¬  way. The final step in each of the three pathways, which begin with pyruvate    FIG. 21 -33 The biosynthesis of the pyruvate family of amino valine and leucine, and the other to isoleucine. Note also that valine   acids: isoleucine, leucine, and valine. The first enzyme, acetolactate aminotransferase catalyzes the formation of both valine and   synthase (a TPP enzyme), catalyzes two reactions, one leading to isoleucine from their respective a-keto acids.    rather than an amino acid, is the PLP-dependent transfer of an amino group  from glutamate to form the amino acid.   The Aromatic Amino Acids Phenylalanine, Tyrosine, and Tryptophan Are  Synthesized from Glucose Derivatives. The precursors of the aromatic  amino acids are the glycolytic intermediate phosphoenolpyruvate (PEP) and  erythrose-4-phosphate (an intermediate in the pentose phosphate pathway;  Fig. 15-30). Their condensation forms 2-keto-3-deoxy-D-arabinoheptu-  losonate-7-phosphate (Fig. 21-34). This C 7 compound cyclizes and is ulti¬  mately converted to chorismate, the branch point for tryptophan synthesis.  Chorismate is converted to either anthranilate and then to tryptophan, or to  prephenate and on to tyrosine or phenylalanine. Although mammals synthe¬  size tyrosine by the hydroxylation of phenylalanine (Fig. 21-24), many mi¬  croorganisms synthesize it directly from prephenate. The last step in the  synthesis of tyrosine and phenylalanine is the addition of an amino group  through transamination. In tryptophan synthesis, the amino group is part of  the serine molecule that is added to indole.   FIG. 21 -34 The biosynthesis of phenylalanine, tryptophan, and tyrosine. The   enzymes shown are (l) 2-keto-3-deoxy-D-arabinoheptulosonate-7-phosphate synthase,  (2) anthranilate synthase, (3) tryptophan synthase, a subunit, (4) tryptophan synthase, |3  subunit (a PLP-dependent enzyme), and (5) chorismate mutase.   Chapter 21 Amino Acid Metabolism Indole Is Channeled between Two Active Sites in Tryptophan Synthase.   The final two reactions of tryptophan biosynthesis (Reactions 3 and 4 in  Fig. 21-34) are both catalyzed by tryptophan synthase:   1 . The a subunit (268 residues) of the a 2 (3 2 bifunctional enzyme cleaves  indole-3-glycerol phosphate, yielding indole and glyceraldehyde-3-  phosphate.   2 . The (3 subunit (396 residues) joins indole with serine in a PLP-  dependent reaction to form tryptophan.   Either subunit alone is enzymatically active, but when they are joined in the  a 2 (3 2 tetramer, the rates of both reactions and their substrate affinities increase  by 1 to 2 orders of magnitude. Indole, the intermediate product, does not ap¬  pear free in solution because it is channeled between the two subunits.   The X-ray structure of tryptophan synthase from Salmonella typhimurium .,  determined by Craig Hyde, Edith Miles, and David Davies, reveals that the  protein forms a 150-A-long, twofold symmetric a(3(3a complex in which the  active sites of neighboring a and (3 subunits are separated by ~25 A (Fig. 21-35).  The active sites are joined by a solvent-filled tunnel that is wide enough to permit the  passage of the intermediate substrate , indole. This structure, the first in which a tun¬  nel connecting two active sites was observed, suggests the following series of  events: The indole-3-glycerol phosphate substrate binds to the a subunit  through an opening into its active site, its “front door,” and the glyceraldehyde-  3-phosphate product leaves via the same route. Similarly, the (3 subunit active  site has a “front door” opening to the solvent through which serine enters and  tryptophan leaves. Both active sites also have “back doors” that are connected  by the tunnel. The indole intermediate presumably diffuses between the two ac¬  tive sites via the tunnel and hence does not escape to the solvent. Channeling may  be particularly important for indole since this nonpolar molecule otherwise can  escape the bacterial cell by diffusing through its plasma and outer membranes.   In order for channeling to increase tryptophan synthases catalytic effi¬  ciency, (1) its connected active sites must be coupled such that their catalyzed  reactions occur in phase, and (2) after substrate has bound to the a subunit,  its active site (“front door”) must close off to ensure that the product indole  passes through the tunnel (“back door”) to the (3 subunit rather than escap¬  ing into solution. A variety of experimental evidence indicates that this series  of events is facilitated through allosteric signals derived from covalent trans¬  formations at the (3 subunit s active site. These switch the enzyme between an    FIG. 21-35 Ribbon diagram of the bifunctional enzyme  tryptophan synthase from S. typhimurium. Only one ap protomer of the  bifunctional appa heterotetramer is shown. The a subunit is blue, the p  subunit’s N-terminal domain is orange, its C-terminal domain is red-orange,  and all p sheets are tan. The active site of the a subunit is located by its  bound competitive inhibitor, indolepropanol phosphate (IPP; red ball-and-  stick model), whereas that of the p subunit is marked by its PLP coenzyme  (yellow ball-and-stick model). The solvent-accessible surface of the  ~25-A-long tunnel connecting the a and p active sites is outlined by  yellow dots. Several indole molecules ( green ball-and-stick models) have  been modeled into the tunnel to show that it is wide enough for indole to  pass from one active site to the other. [Courtesy of Craig Hyde, Edith  Miles, and David Davies, NIH.] Q See Interactive Exercise 30.    open, low-activity conformation to which substrates bind, and a closed, high-  activity conformation from which indole cannot escape.   Histidine Biosynthesis Includes an Intermediate in Nucleotide  Biosynthesis. Five of histidines six C atoms are derived from 5-phospho-  ribosyl-a-pyrophosphate (PRPP; Fig. 21-36), a phospho-sugar intermediate  that is also involved in the biosynthesis of purine and pyrimidine nucleotides  (Sections 23-1A and 23-2A). The histidines sixth carbon originates from ATP.  The ATP atoms that are not incorporated into histidine are eliminated as  5-aminoimidazole-4-carboxamide ribonucleotide (Fig. 21-36, Reaction 2),  which is also an intermediate in purine biosynthesis.   The unusual biosynthesis of histidine from a purine (A^-S'-phospho-  ribosyl ATP, the product of Reaction 1 in Fig. 21-36) has been cited as evi¬  dence supporting the hypothesis that life was originally RNA-based. His  residues, as we have seen, are often components of enzyme active sites, where  they act as nucleophiles and/or general acid—base catalysts. The discovery that  RNA can have catalytic properties therefore suggests that the imidazole moi¬  ety of purines plays a similar role in RNA enzymes. This further suggests that  the histidine biosynthetic pathway is a “fossil” of the transition to more effi¬  cient protein-based life-forms.   • What are the metabolic precursors of the  nonessential amino acids?   • Summarize the types of reactions required  to synthesize the nonessential amino  acids.   • List the compounds that are used to syn¬  thesize the essential amino acids in plants  and microorganisms.   • Compare the catabolic and anabolic path¬  ways for one or more of the amino acids.  In which pathways do oxidation reactions  or reduction reactions predominate?    • Heme is synthesized from glycine and succinyl-CoA and is degraded to a variety  of colored compounds for excretion.   • The synthesis of bioactive amines begins with amino acid decarboxylation.   • Arginine gives rise to the hormonally active gas nitric oxide.    Certain amino acids, in addition to their major function as protein build¬  ing blocks, are essential precursors of a variety of important biomolecules,  including nucleotides and nucleotide coenzymes, heme, and various hor¬  mones and neurotransmitters. In this section, we consider the pathways lead¬  ing to some of these substances. The biosynthesis of nucleotides is considered  in Chapter 23.   Heme, as we have seen, is an Fe-containing prosthetic group that is an essen¬  tial component of many proteins, notably hemoglobin, myoglobin, and the  cytochromes. The initial reactions of heme biosynthesis are common to the  formation of other tetrapyrroles including chlorophyll in plants and bacteria  (Fig. 19-2) and coenzyme B 12 in bacteria (Fig. 20-17).   Elucidation of the heme biosynthetic pathway involved some interesting  detective work. David Shemin and David Rittenberg, who were among the  first to use isotopic tracers in the elucidation of metabolic pathways, demon¬  strated, in 1945, that all of hemes C and N atoms can be derived from acetate  and glycine. Heme biosynthesis takes place partly in the mitochondrion and  partly in the cytosol (Fig. 21-37). Mitochondrial acetate is metabolized via  the citric acid cycle to succinyl-CoA, which condenses with glycine in a reac¬  tion that produces C0 2 and 8 -aminolevulinic acid (ALA). ALA is trans¬  ported to the cytosol, where it combines with a second ALA to yield  porphobilinogen (PBG). The reaction is catalyzed by the Zn-requiring en¬  zyme porphobilinogen synthase.   r Inhibition of PBG synthase by lead is one of the major manifestations of  acute lead poisoning. Indeed, it has been suggested that the accumulation,  in the blood, of ALA, which resembles the neurotransmitter y-aminobutyric  acid (Section 21-6B), is responsible for the psychosis that often accompanies  lead poisoning.   The next phase of heme biosynthesis is the condensation of four PBG mol¬  ecules to form uroporphyrinogen III, the porphyrin nucleus, in a series of  reactions catalyzed by porphobilinogen deaminase (also called uropor¬  phyrinogen synthase) and uroporphyrinogen III synthase. The initial  product, hydroxymethylbilane, is a linear tetrapyrrole that cyclizes.   Protoporphyrin IX, to which Fe is added to form heme, is produced from  uroporphyrinogen III in a series of reactions catalyzed by (1) uroporphyrino¬  gen decarboxylase, which decarboxylates all four acetate side chains (A) to  form methyl groups (M); (2) coproporphyrinogen oxidase, which oxidatively  decarboxylates two of the propionate side chains (P) to vinyl groups (V); and  (3) protoporphyrinogen oxidase, which oxidizes the methylene groups link¬  ing the pyrrole rings to methenyl groups. During the coproporphyrinogen ox¬  idase reaction, the porphyrin is transported back into the mitochondrion. In   8-Aminolevulinic acid (ALA) is synthesized in the mitochondrion from  succinyl-CoA and glycine by ALA synthase. ALA is transported to the  cytosol, where two molecules condense to form PBG, four molecules  of which condense to form a porphyrin ring. The next three reactions  involve oxidation of the pyrrole ring substituents, yielding    protoporphyrinogen IX, which is transported back into the  mitochondrion during its formation. After oxidation of the methylene  groups, ferrochelatase catalyzes the insertion of Fe 2+ to yield heme.  A, P, M, and V, respectively, represent acetyl, propionyl, methyl, and  vinyl (—CH=CH 2 ) groups. C atoms originating as the carboxyl  group of acetate are red.    Defects in heme biosynthesis in liver or erythroid cells result in  the accumulation of porphyrin and/or its precursors and are there¬  fore known as porphyrias. Two such defects are known to affect  erythroid cells: uroporphyrinogen III cosynthase deficiency (con¬  genital erythropoietic porphyria) and ferrochelatase deficiency  (erythropoietic protoporphyria). The former results in accumula¬  tion of uroporphyrinogen derivatives. Excretion of those compounds  colors the urine red; their deposition in the teeth turns them  reddish brown; and their accumulation in the skin renders it  extremely photosensitive, so that it ulcerates and forms disfiguring  scars. Increased hair growth is also observed in afflicted individu¬  als; fine hair may cover much of the face and extremities. These  symptoms have prompted speculation that the werewolf legend  has a biochemical basis.    The most common porphyria that primarily affects liver is por¬  phobilinogen deaminase deficiency (acute intermittent porphyria).  This disease is marked by intermittent attacks of abdominal pain  and neurological dysfunction. Excessive amounts of ALA and PBG  are excreted in the urine during and after such attacks. The urine  may become red resulting from the excretion of excess porphyrins  synthesized from PBG in nonhepatic cells, although the skin does  not become unusually photosensitive. King George III, who ruled  England during the American Revolution, and who has been widely  portrayed as being mad, in fact, had attacks characteristic of acute  intermittent porphyria; he was reported to have urine the color of  port wine, and had several descendants who were diagnosed as  having the disease. American history might have been quite differ¬  ent had George III not inherited that metabolic defect.    the final reaction of heme biosynthesis, ferrochelatase inserts Fe(II) into pro¬  toporphyrin IX.   The Liver and Erythroid Cells Regulate Heme Biosynthesis. The two ma¬  jor sites of heme biosynthesis are erythroid cells, which synthesize ~85% of  the body’s heme groups, and the liver, which synthesizes most of the remain¬  der. In liver, the level of heme synthesis must be adjusted according to  metabolic conditions. For example, the synthesis of the heme-containing cy¬  tochromes P450 (Section 12-4D) fluctuates with the need for detoxification.  In contrast, heme synthesis in erythroid cells is a one-time event; heme and  protein synthesis cease when the cell matures, so that the hemoglobin must  last the erythrocytes lifetime (~120 days).   In liver, the main control target in heme biosynthesis is ALA synthase, the  enzyme catalyzing the pathways first committed step. Heme, or its Fe(III) ox¬  idation product hemin, controls this enzymes activity through feedback inhi¬  bition, inhibition of the transport of ALA synthase from its site of synthesis  in the cytosol to its reaction site in the mitochondrion (Fig. 21-37), and re¬  pression of ALA synthase synthesis.   In erythroid cells, heme exerts quite a different effect on its biosynthesis.  Heme induces, rather than represses, protein synthesis in reticulocytes (im¬  mature erythrocytes). Although the vast majority of the protein synthesized by  reticulocytes is globin, heme may also induce these cells to synthesize the en¬  zymes of heme biosynthesis. Moreover, the rate-determining steps of heme  biosynthesis in erythroid cells may be the ferrochelatase and porphobilinogen  deaminase reactions rather than the ALA synthase reaction. This is consistent  with the supposition that when erythroid heme biosynthesis is “switched on,”  all of its steps function at their maximal rates rather than any one step limit¬  ing the flow through the pathway. Heme-stimulated synthesis of globin also  ensures that heme and globin are synthesized in the correct ratio for assembly  into hemoglobin (Section 28-3C). Genetic defects in heme biosynthesis cause  conditions known as porphyrias (Box 21-3).   Heme Degradation Products Are Excreted. At the end of their lifetime, red  cells are removed from the circulation and their components degraded. Heme  catabolism (Fig. 21-38) begins with oxidative cleavage, by heme oxygenase, of  the porphyrin between rings A and B to form biliverdin, a green linear  tetrapyrrole. Biliverdin’s central methenyl bridge (between rings C and D) is  then reduced to form the red-orange bilirubin. The changing colors of a heal¬  ing bruise are a visible manifestation of heme degradation.   In the reaction forming biliverdin, the methenyl bridge carbon between  rings A and B is released as CO, which is a tenacious heme ligand (with 200-  fold greater affinity for hemoglobin than 0 2 ; Section 7-1A). Consequently,  ~ 1 % of hemoglobin s binding sites are blocked by CO even in the absence  of air pollution.   The highly lipophilic bilirubin is insoluble in aqueous solutions. Like  other lipophilic metabolites, such as free fatty acids, it is transported in the   blood in complex with serum albumin. Bilirubin derivatives are secreted in  the bile and for the most part are further degraded by bacterial enzymes in  the large intestine. Some of the resulting urobilinogen is reabsorbed and trans¬  ported via the bloodstream to the kidney, where it is converted to the yellow  urobilin and excreted, thus giving urine its characteristic color. Most uro¬  bilinogen, however, is microbially converted to the deeply red-brown sterco-  bilin, the major pigment of feces.   T When the blood contains excessive amounts of bilirubin, the deposition of  this highly insoluble substance colors the skin and the whites of the eyes  yellow. This condition, called jaundice (French: jaune , yellow), signals either  an abnormally high rate of red cell destruction, liver dysfunction, or bile duct  obstruction. Newborn infants, particularly when premature, often become  jaundiced because they lack an enzyme that degrades bilirubin. Jaundiced  infants are treated by bathing them with light from a fluorescent lamp; this  photochemically converts bilirubin to more soluble isomers that the infant can  degrade and excrete.    X = OH, R = CH 3 Epinephrine (adrenalin)  X = OH, R = H Norepinephrine   Epinephrine (adrenaline), norepinephrine, dopamine, serotonin (5-  hydroxytryptamine), y-aminobutyric acid (GABA), and histamine (at left)   are hormones and/or neurotransmitters derived from amino acids. Epinephrine,  as we have seen, activates muscle adenylate cyclase, thereby stimulating glyco¬  gen breakdown (Section 16-3B); deficiency in dopamine production is associ¬  ated with Parkinson’s disease, a degenerative condition causing “shaking palsy”;  serotonin causes smooth muscle contraction; GABA is one of the brains ma¬  jor inhibitory neurotransmitters; and histamine is involved in allergic responses  (as allergy sufferers who take antihistamines will realize), as well as in the con¬  trol of acid secretion by the stomach.   The biosynthesis of each of these physiologically active amines involves  decarboxylation of the corresponding precursor amino acid. Amino acid de¬  carboxylases are PLP-dependent enzymes that form a PLP—Schiff base with  the substrate so as to stabilize (by delocalization) the C a carbanion formed on  C a —COO bond cleavage (Section 21-2A):   Formation of histamine (from histidine) and formation of GABA (from  glutamate) are one-step processes; the synthesis of serotonin from tryptophan  requires a hydroxylation step as well as decarboxylation. The various  catecholamines —dopamine, norepinephrine, and epinephrine—are related to  catechol (at left) and are sequentially synthesized from tyrosine (Fig. 21-39):   1. Tyrosine is hydroxylated to 3,4-dihydroxyphenylalanine (l-DOPA)  in a reaction that requires 5,6,7,8-tetrahydrobiopterin (Fig. 21-26).   4. Methylation of norepinephrines amino group by S-adenosylmethio-  nine (Fig. 21-18) produces epinephrine.   FIG. 21 -39 The sequential synthesis of l-DOPA, dopamine, norepinephrine, and  epinephrine from tyrosine.    The specific catecholamine that a cell produces depends on which enzymes of  the pathway are present. In adrenal medulla, for example, epinephrine is the  predominant product. In some areas of the brain, norepinephrine is more com¬  mon. In other areas, the pathway stops at dopamine. In melanocytes, l-DOPA  is the precursor of red and black melanins, which are irregular cross-linked  polymers that give hair and skin much of their color.   Arginine is the precursor of a substance that was originally called endothelium-  derived relaxing factor (EDRF) because it was synthesized by vascular  endothelial cells and caused the underlying smooth muscle to relax. The sig¬  nal for vasodilation was not a peptide, as expected, but the stable free radical  gas nitric oxide, NO. The identification of NO as a vasodilator came in part  from studies that identified NO as the decomposition product that mediates  the vasodilating effects of compounds such as nitroglycerin (at right).  Nitroglycerin is often administered to individuals suffering from angina pec¬  toris, a disease caused by insufficient blood flow to the heart muscle, to rap¬  idly but temporarily relieve their chest pain.   The reaction that converts arginine to NO and citrulline is catalyzed by  nitric oxide synthase (NOS):   • How does the regulation of heme synthe¬  sis differ between the liver and reticulo¬  cytes?   • Identify the amino acids that give rise to  catecholamines, serotonin, GABA, and  histamine.   • What are the substrates and products of  the nitric oxide synthase reaction?    The reaction proceeds via an enzyme-bound hydroxyarginine intermediate and  requires an array of redox coenzymes. NOS is a homodimeric protein of 125-  to 160-kD subunits, and each subunit contains one FMN, one FAD, one  tetrahydrobiopterin (Fig. 21-26), and one Fe(III)-heme. These cofactors facil¬  itate the five-electron oxidation of arginine to produce NO.   Because NO is a gas, it rapidly diffuses across cell membranes, although  its high reactivity (half-life ~5 s) prevents it from acting much farther than  ~1 mm from its site of synthesis. NO is produced by endothelial cells in re¬  sponse to a wide variety of agents and physiological conditions. Neuronal cells  also synthesize NO (neuronal NOS is ~55% homologous to endothelial  NOS). This endothelium-independent NO synthesis induces the dilation of  cerebral and other arteries and is responsible for penile erection. The brain  contains more NOS than any other tissue in the body, suggesting that NO is  essential for the function of the central nervous system. A third type of NOS  is found in leukocytes (white blood cells). These cells produce NO as part of  their cytotoxic arsenal. NO combines with superoxide (Section 18-4B) to pro¬  duce peroxynitrite (ONOO ), which combines with a proton and decays to  nitrogen dioxide (N0 2 ) and a hydroxyl radical (*OH):   The highly reactive *OH kills invading bacteria. The sustained release of NO  has been implicated in endotoxic shock (an often fatal immune system over¬  reaction to bacterial infection), in inflammation-related tissue damage, and in  the damage to neurons in the vicinity of but not directly killed by a stroke  (which often does greater harm than the stroke itself).    • The reduction of N 2 to NH 3 by nitrogenase is an energetically costly process.   • Ammonia is incorporated into amino acids by the action of glutamate synthase.   FIG. 21 -40 Root nodules of a soybean  plant. Bacteria of the genus Rhizobium, which  carry out nitrogen fixation, live symbiotically  within the root nodules of such legumes.    The most prominent chemical elements in living systems are O, H, C, N, and  P. The elements O, H, and P occur widely in metabolically available forms  (H 2 0, 0 2 , and P z ). However, the major forms of C and N, C0 2 and N 2 , are  extremely stable (unreactive); for example, the N=N triple bond has a bond  energy of 945 kj • mol 1 (versus 351 kj • mol 1 for a C—O single bond).  C0 2 , with minor exceptions, is metabolized (fixed) only by photosynthetic or¬  ganisms (Chapter 19). Nitrogen fixation is even less common; N 2 is converted  to metabolically useful forms by only a few strains of bacteria, called diazotrophs.  These organisms include certain marine cyanobacteria and bacteria that colo¬  nize the root nodules of legumes (plants belonging to the pea family, includ¬  ing beans, clover, and alfalfa; Fig. 21-40). Once N 2 is fixed, the nitrogen is  assimilated (incorporated) into biological molecules as amino groups that can  then be transferred to other molecules. In this section we examine the processes  of nitrogen fixation and nitrogen assimilation.   Diazotrophs produce the enzyme nitrogenase, which catalyzes the reduction  of N 2 to NH 3 :   N 2 + 8 H + + 8 e~ + 16 ATP + 16 H 2 0 ->   In legumes, this nitrogen-fixing system produces more metabolically useful  nitrogen than the legume needs; the excess is excreted into the soil, enriching  it. It is therefore common agricultural practice to plant a field with alfalfa    every few years to build up the supply of usable nitrogen in the soil for later  use in growing other crops.   Nitrogenase Contains Novel Redox Centers. Nitrogenase, which catalyzes  the reduction of N 2 to NH 3 , is a complex of two proteins:   1 . The Fe-protein, a homodimer that contains one [4Fe-4S] cluster and  two ATP-binding sites.   2 . The MoFe-protein, an a 2 (3 2 heterotetramer that contains Fe and Mo.   The X-ray structure of Azotobacter vinelandii nitrogenase in complex with the  inhibitor ADP • AIF 4 (which mimics the transition state in ATP hydrolysis),  determined by Douglas Rees, reveals that each MoFe-protein associates with  two molecules of Fe-protein (Fig. 21-41).   Each Fe-protein dimers single [4Fe-4S] cluster is located in a solvent-  exposed cleft between the two subunits and is symmetrically linked to Cys 97  and Cys 132 from both subunits such that an Fe-protein resembles an “iron  butterfly” with the [4Fe-4S] cluster at its head. Its nucleotide-binding sites are  located at the interface between its two subunits.   The MoFe-protein’s a and (3 subunits assume similar folds and extensively  associate to form a pseudo twofold symmetric a (3 dimer, two of which more  loosely associate to form the twofold symmetric a 2 (3 2 tetramer (Fig. 21-41).  Each a(3 dimer has two bound redox centers:   1. The P-cluster (Fig. 21-42^,6), which consists of two [4Fe-3S] clusters  linked through an additional sulfide ion forming the eighth corner of  both of the clusters to make cubane-like structures, and bridged by two   FIG. 21-41 X-Ray structure of the A. vinelandii nitrogenase in complex with  ADP -AIFT- The enzyme, which is viewed along its molecular twofold axis, is an (<*( 372)2  heterooctamer in which the (3-a-<*-(3 assembly, the MoFe-protein, is flanked by two 72  Fe-proteins whose 289-residue subunits are related by local twofold symmetry. The  homologous a subunits (cyan and red; 491 residues) and (3 subunits (light red and light blue;  522 residues) are related by pseudotwofold symmetry. The two 7 subunits forming each  Fe-protein (pink and green with their Switch I and Switch II segments red and blue) bind to the  MoFe-protein with the twofold axis relating them coincident with the pseudo-two-fold axis  relating the MoFe-protein’s a and (3 subunits. The ADP • AIF 4 , [4Fe-4S] cluster, Fe Mo-cofactor,  and P-cluster are drawn in space-filling form with C green, N blue, 0 red, S yellow, Fe orange,  Mo pink, and the AIFT ion purple. [Based on an X-ray structure by Douglas Rees, California  Institute of Technology. PDBid 1N2C.] ©■ See Interactive Exercise 31.    FIG. 21 -42 The prosthetic groups of the nitrogenase MoFe-  protein. The molecules are drawn in ball-and-stick form with C green, N  blue, 0 red, S yellow, Fe orange, and Mo pink, (a) The reduced Klebsiella  pneumoniae P-cluster. It consists of two [4Fe-3S] complexes linked by an  additional sulfide ion forming the eighth corner of each cubane-like  structure, and bridged by two Cys thiol ligands, each coordinating one Fe  from each cluster. Four additional Cys thiols coordinate the remaining four  Fe atoms, (b) The 2-electron-oxidized K. pneumoniae P-cluster. In  comparison with the reduced complex in Part a, two of the Fe—S bonds  from the centrally located sulfide ion that bridges the two [4Fe-3S] clusters  have been replaced by ligands from the Cys 87a amide N and the Ser  186(3 side chain 0 yielding a [4Fe-3S] cluster (left) and a [4Fe-4S] cluster  (right) that remain linked by a direct Fe—S bond and two bridging Cys  thiols, (c) The A vinelandii Fe Mo-cofactor. It consists of a [4Fe-3S] cluster  and a [1Mo-3Fe-3S] cluster that are bridged by three sulfide ions. The  FeMo-cofactor is linked to the protein by only two ligands at its opposite  ends, one from His 442a to the Mo atom and the other from Cys 275a to  an Fe atom. The Mo atom is additionally doubly liganded by homocitrate.  What is most likely an N atom (blue sphere) is liganded to the FeMo-  cluster’s six central Fe atoms (dashed black lines). [Parts a and b based on  X-ray structures by David Lawson, John Innes Centre, Norwich, U.K. Part c  based on an X-ray structure by Douglas Rees, California Institute of  Technology. PDBids (a) 1QGU, (b) 1QH1, and (c) 1M1NJ    thiol ligands, each coordinating one Fe from each cluster. Four  additional Cys thiols coordinate the remaining four Fe atoms.  The positions of two of the Fe atoms in one of the [4Fe—3S]  clusters change on oxidation, rupturing the bonds from these Fe  atoms to the linking sulfide ion. These bonds are replaced in the  oxidized state by a Ser oxygen ligand to one of the Fe atoms,  and by a bond to the amide N of a Cys from the other Fe atom.   2 . The FeMo-cofactor (Fig. 21 -42c), which consists of a [4Fe-3S]  cluster and a [lMo-3Fe-3S] cluster bridged by three sulfide  ions. The FeMo-cofactor s Mo atom is approximately octahe-  drally coordinated by three cofactor sulfurs, a His imidazole ni¬  trogen, and two oxygens from a bound homocitrate ion (at left)  (an essential component of the FeMo-cofactor). A central cav¬  ity in the FeMo-cofactor contains what most probably is a ni¬  trogen atom (although a C or an O atom cannot be ruled out).  This putative N atom is liganded to the FeMo-cofactors cen¬  tral six Fe atoms such that it completes the approximate tetra¬  hedral coordination environment of each of the Fe atoms.   Nitrogen fixation by nitrogenase requires a source of electrons. These are gen¬  erated either oxidatively or photosynthetically, depending on the organism.  The electrons are transferred to ferredoxin, a [4Fe-4S]-containing electron  carrier that transfers an electron to the Fe-protein of nitrogenase, beginning  the nitrogen fixation process (Fig. 21-43).   The transfer of electrons during the nitrogenase reaction requires ATP-  dependent protein conformational changes and the dissociation of the Fe-  protein from the MoFe-protein after each electron transfer. During the  reaction cycle, two molecules of ATP bind to the Fe-protein and are hy¬  drolyzed as an electron passes to the MoFe-protein. ATP hydrolysis induces a  conformational change in the Fe-protein that alters its redox potential from  — 0.29 to —0.40 V, making the electron capable of N 2 reduction ( < 8°' for the  reaction N 2 + 6 H + + 6 e \ 2 NH 3 is —0.34 V).    How are events at the ATP-binding site linked to electron transfer? ATP  hydrolysis occurs at a significant rate only when the Fe-protein is in complex  with the MoFe-protein (and, of course, electron transfer can occur only when  the two proteins are associated). Intriguingly, the binding of ADP • AIF 4 to  the Fe-protein induces conformational changes in two regions of the Fe-  protein, designated Switch I and Switch II (Fig. 21-41), that are homologous  to segments of signal-transducing G proteins in which nucleotide hydrolysis  is coupled to protein conformational changes (Section 13-3B). At Switch I,  these conformational changes affect the interactions between the Fe-protein  and the MoFe-protein, whereas at Switch II they affect the environment of  the [4Fe-4S] cluster. X-Ray structures of the Fe-protein alone and with bound  ADP or ATP indicate that ATP hydrolysis triggers a structural change includ¬  ing rotation of the two Fe-protein subunits by about 13° toward each other.  This conformational change brings the [4Fe-4S] cluster closer to the P-  cluster of the adjacent MoFe-protein (from 18 to 14 A apart), thereby pro¬  moting electron transfer from the Fe-protein to the MoFe-protein.   Kinetic studies of nitrogenase indicate that the rate-limiting step of N 2  reduction is the dissociation of the Fe-protein from the MoFe-protein. If the  Fe-protein functions in the same way as a G protein, then nucleotide hydrol¬  ysis might trigger its dissociation from the MoFe-protein. Reassociation could  occur when the ADP exits the active site and is replaced by ATP. ATP hydrol¬  ysis would not occur again until the Fe-protein was docked with the MoFe-  protein and poised to transfer electrons.    N 2 Reduction Is Energetically Costly. The actual reduction of N 2 occurs on  the MoFe-protein in three discrete steps, each involving an electron pair:    An electron transfer must occur six times per N 2 molecule fixed, so a total of  12 ATP are required to fix one N 2 molecule. However, nitrogenase also  reduces H 2 0 to H 2 , which in turn reacts with diimine to re-form N 2 .   This futile cycle is favored when the ATP level is low and/or the reduction of  the Fe-protein is sluggish. Even when ATP is plentiful, however, the cycle can¬  not be suppressed beyond about one H 2 molecule produced per N 2 reduced  and hence appears to be a requirement for the nitrogenase reaction. The  total cost of N 2 reduction is therefore 8 electrons transferred and 16 ATP  hydrolyzed. Under cellular conditions, the cost is closer to 20—30 ATP.  Consequently, nitrogen fixation is an energetically expensive process.    FIG. 21 -43 The flow of electrons in the nitrogenase-catalyzed reduction of N 2 .    Amino Acid Metabolism Although atmospheric N 2 is the ultimate nitrogen source for all living   things, most plants do not support the symbiotic growth of nitrogen-fixing  bacteria. They must therefore depend on a source of “prefixed” nitrogen such  as nitrate or ammonia. These nutrients come from lightning discharges (the  source of — 10 % of naturally fixed N 2 ), decaying organic matter in the soil,  or from fertilizer applied to it (—50% of the nitrogen fixed is now generated  by the Haber-Bosch process, which is used in the industrial production of  ammonia from N 2 and H 2 ). One major long-term goal of genetic engineer¬  ing is to induce agriculturally useful nonleguminous plants to fix their own  nitrogen. This would free farmers, particularly those in developing countries,  from either purchasing fertilizers, periodically letting their fields lie fallow (giv¬  ing legumes the opportunity to grow), or following the slash-and-burn tech¬  niques that are rapidly destroying the worlds tropical forests.   After atmospheric N 2 has been converted to a biologically useful form (e.g.,  ammonia), it must be assimilated into a cells biomolecules. Once nitrogen has  been introduced into an amino acid, the amino group can be transferred to  other compounds by transamination. Because most organisms do not fix  nitrogen and hence must rely on prefixed nitrogen, nitrogen assimilation  reactions are critical for conserving this essential element.   We have already considered the reaction catalyzed by glutamine synthetase  (Section 21-5A), which in microorganisms represents a metabolic entry point  for fixed nitrogen (in animals, this reaction helps “mop up” excess ammonia).  The glutamine synthetase reaction requires the nitrogen-containing compound  glutamate as a substrate. So what is the source of the amino group in gluta¬  mate? In bacteria and plants, but not animals, the enzyme glutamate synthase  converts a-ketoglutarate and glutamine to two molecules of glutamate:   a-Ketoglutarate + glutamine + NADPH + H + —>2 glutamate + NADP +   This reductive amination reaction requires electrons from NADPH and  takes place at three distinct active sites in the a 2 (3 2 heterotetramer (Fig. 21-44).  X-Ray structures of the enzyme reveal that the substrate-binding sites are widely  separated. As a result, the ammonia that is transferred from glutamine to a-  ketoglutarate must travel through a 31-A-long tunnel in the protein. This  channeling probably helps prevent the loss of NH 3 to the cytosol and main¬  tains this intermediate in its more reactive deprotonated state (in the cytosol,  NH 3 would immediately acquire a proton to become NH 4 ). Furthermore,  the tunnel is blocked by the side chains of several residues and opens only  when a-ketoglutarate is bound to the enzyme and NADPH is available. This  mechanism apparently prevents the wasteful hydrolysis of glutamine.   The net result of the glutamine synthetase and glutamate synthase  reactions is   Thus, the combined action of these two enzymes assimilates fixed nitrogen (NH4)  into an organic compound (a-ketoglutarate) to produce an amino acid (gluta¬  mate). Once the nitrogen is assimilated as glutamate, it can be used in the  synthesis of other amino acids by transamination.   The Nitrogen Cycle Describes the Interconversion of Nitrogen in the  Biosphere. The ammonia produced by the nitrogenase reaction and incorpo¬  rated into amino acids is eventually recycled in the biosphere as described by  the nitrogen cycle (Fig. 21-45). Nitrate is produced by certain bacteria that    FIG. 21-44 The glutamate synthase reaction, (l) Electrons are  transferred from NADPH to FAD at active site 1 on the |3 subunit to  yield FADH 2 . ( 2 ) Electrons travel from the FADH 2 to FMN at site 2  on an a subunit to yield FMNH 2 . (3) Glutamine is hydrolyzed to    glutamate and ammonia at site 3. (4) The ammonia produced in Step  3 moves to site 2 by channeling, where it reacts with a-ketoglutarate.  (5) The a-iminoglutarate product is reduced by FMNH 2 to form  glutamate.    FIG. 21-45 The nitrogen cycle. Nitrogen  fixation by nitrogenase converts N 2 to the  biologically useful ammonia. Nitrate can also be  converted to ammonia by the sequential actions of  nitrate reductase and nitrite reductase. Ammonia is  transformed to N 2 by nitrification followed by  denitrification. Ammonia may be assimilated into  nitrogen-containing biomolecules, which may be  decomposed back to ammonia.   • Describe the reactions that assimilate NH 3  into amino acids. Which reactions occur in  mammals?   • List the enzymes of amino acid metabo¬  lism in which channeling occurs.    oxidize NH 3 to N0 2 and then N0 3 , a process called nitrification. Still other  organisms convert nitrate back to N 2 , which is known as denitrification. In  addition, nitrate is reduced to NH 3 by plants, fungi, and many bacteria, a  process called ammonification in which nitrate reductase catalyzes the two-  electron reduction of nitrate to nitrite (N0 2 ):   NO 3 + 2 H + +2^-> NO^ + H 2 0   N0 2 + 7 H + + 6 e~ -> NH 3 + 2 H 2 0   The direct anaerobic oxidation of NH 3 back to N 2 without the intermediacy  of nitrate, the reverse of nitrogen fixation, has recently been discovered in  certain bacteria.   • Intracellular proteins and extracellular proteins taken up by endo-  cytosis are degraded by lysosomal proteases.   • Proteins tagged by the addition of multiple copies of ubiquitin  enter the barrel-shaped proteasome, where they are digested into  ~ 8-residue fragments.   • The degradation of an amino acid almost always begins with the  removal of its amino group in a PLP-facilitated transamination  reaction.   • A nitrogen atom from ammonia (a product of the oxidative deam¬  ination of glutamate) and bicarbonate are incorporated into car¬  bamoyl phosphate for entry into the urea cycle.   • A second nitrogen atom introduced from aspartate enters the  cycle to produce urea for excretion.   • The rate-limiting step of this process is catalyzed by carbamoyl  phosphate synthetase.   • The 20 “standard” amino acids are degraded to compounds that  give rise either to glucose or to ketone bodies or fatty acids: pyru¬  vate, a-ketoglutarate, succinyl-CoA, fumarate, oxaloacetate,  acetyl-CoA, or acetoacetate.   • The cofactors involved in amino acid degradation include PLP,  tetrahydrofolate, and biopterin.   • The nonessential amino acids are synthesized in all organisms us¬  ing simple pathways with the starting materials pyruvate, oxaloac¬  etate, a-ketoglutarate, and 3-phosphoglycerate.   • The essential amino acids, which are made only in plants and mi¬  croorganisms, require more complicated pathways that vary  among organisms.   • Amino acids are the precursors of various biomolecules. Heme  synthesis begins with glycine and succinyl-CoA derived from  acetyl-CoA, and the porphyrin ring is built in a series of reactions  that occur in the mitochondria and the cytosol.   • Various hormones and neurotransmitters are synthesized by the  decarboxylation and hydroxylation of histidine, glutamate, tryp¬  tophan, and tyrosine.   • The five-electron oxidation of arginine yields the bioactive stable  radical nitric oxide.   • Nitrogen fixation in bacteria, which requires 8 electrons and at  least 16 ATP, is catalyzed by nitrogenase, a multisubunit protein  with redox centers containing Fe, S, and Mo.   • Fixed nitrogen is incorporated into amino acids via the glutamine  synthetase and glutamate synthase reactions.    deamination 722  hyperammonemia 722  urea cycle 723  channeling 725  glucogenic amino acid 727  ketogenic amino acid 727  THF 729    SAM 732  biopterin 739  essential amino acid 740  nonessential amino acid 741  adenylylation 744  uridylylation 744  porphyria 754    1. Explain why protein degradation by proteasomes requires ATP  even though proteolysis is an exergonic process.   2 . The compound shown here inhibits an archaebacterial protea-  some, binding to each of its active sites, which (unlike those of  eukaryotic proteasomes) are all identical. What can you conclude  about the substrate specificity of the archaebacterial proteasome?   3. Explain why cellular stressors such as high temperatures or  oxidative damage would trigger the production of proteasomes.   4. Ritonavir, an inhibitor of HIV protease (Box 12-3), also inhibits  the chymotrypsin-like activity of the proteasome. Explain why  this dual inhibitory effect could contribute to the neurotoxic side  effects of long-term ritonavir use.   3. Explain why it was advantageous for human ancestors to evolve  a taste receptor for glutamate.   6 . An L-amino acid oxidase in mammalian peroxisomes requires H 2 0  and 0 2 as substrates and catalyzes amino acid deamination, produc¬  ing H 2 0 2 as a product. Write a balanced equation for the reaction.   7. Explain why the symptoms of a partial deficiency in a urea  cycle enzyme can be attenuated by a low-protein diet.   8 . Production of the enzymes that catalyze the reactions of the urea  cycle can increase or decrease according to the metabolic needs of  the organism. High levels of these enzymes are associated with high-  protein diets as well as starvation. Explain this apparent paradox.   9. (a) How many ATP equivalents are consumed by the reactions  of the urea cycle? (b) Operation of the urea cycle actually gener¬  ates more ATP than it consumes. Explain.   10. Helicobacter pylori, the bacterium responsible for gastric ulcers,  can survive in the stomach (where the pH is as low as 1.3) in  part because it synthesizes large amounts of the enzyme urease.   (a) Write the reaction for urea hydrolysis by urease, (b) Explain  why this reaction could help establish a more hospitable environ¬  ment for H. pylori, which tolerates acid but prefers to grow at  near-neutral pH.   12 . Alanine, cysteine, glycine, serine, and threonine are amino acids  whose breakdown yields pyruvate. Which, if any, of the remain¬  ing 15 amino acids also do so?   13. Unlike the other 19 standard amino acids, lysine does not un¬  dergo transamination. What is the fate of its a- and £-amino  groups when it is catabolized?   14. Compare the ATP yield for the complete oxidation of glutamate  and methionine, both of which contain five carbons atoms, to C0 2 .   15. In the degradation pathway for isoleucine (Fig. 21-21), draw the re¬  actions that convert tiglyl-CoA to acetyl-CoA and propionyl-CoA.   16. Draw the amino acid-Schiff base that forms in the breakdown  of 3 -hydroxykynurenine to yield 3 -hydroxyanthranilate in the  tryptophan degradation pathway (Fig. 21-23, Reaction 4) and in¬  dicate which bond is to be cleaved.   17. Tyrosine and cysteine are listed as nonessential amino acids, but  an inadequate diet may cause tyrosine and/or cysteine insuffi¬  ciency. Explain.    18. Compile a list of all the cofactors involved in adding or remov¬  ing one-carbon groups in carbohydrate, lipid, and amino acid me¬  tabolism. Provide an example of a reaction that uses each cofactor.   19. Which three mammalian enzymes can potentially react with and  thereby decrease the concentration of free NH 4 ?   20. E. coli can convert glutamate to ornithine. Describe the two  reactions that do so.   21 . What are the metabolic consequences of a defective uridylyl-  removing enzyme in E. coli?   22. Many of the most widely used herbicides inhibit the synthesis of  aromatic amino acids. Explain why the compounds are safe to  use near animals.   24. The compound shown here is used to treat trypanosome infection.  What amino acid does the compound resemble?    25. One of the symptoms of kwashiorkor, the dietary protein defi¬  ciency disease in children, is the depigmentation of the skin and  hair. Explain the biochemical basis of this symptom.   27. Some diazotrophs produce a vanadium-containing VFe protein  in addition to the MoFe protein. The vanadium-containing  nitrogenase converts N 2 to NH 3 and also converts CO to com¬  pounds such as ethane and propane. What aspect of the standard  nitrogenase reaction is responsible for the production of alkanes?   28. Nitrosomonas is a photophobic chemoautotrophic bacterium that  converts ammonia into nitrite. Explain how the organism can fix  C0 2 via the Calvin cycle in the absence of sunlight.    How do anammox bacteria par¬  ticipate in the nitrogen cycle? Where do the organisms live and  how do they contain the toxic intermediates of their specialized  metabolism?   Groll, M. and Clausen, M., Molecular shredders: How proteasomes ful¬  fill their roles. Curr. Opin. Struct. Biol. 13, 665-673 (2003).   Navon, A. and Ciechanover, A., The 26S proteosome: from basic mech¬  anism to drug targeting, J. Biol. Chem. 284, 33713-33718 (2009).   Varshavsky, A., Regulated protein degradation, Trends Biochem. Sci. 30,  283-286 (2005).   Withers, P.C., Urea: Diverse functions of a waste product,’ Clin. Exp.  Pharm. Physiol. 25, 722-727 (1998). [Discusses the various roles of  urea, contrasting it with ammonia with respect to toxicity, acid-base  balance, and nitrogen transport.]   Brosnan, J.T., Glutamate, at the interface between amino acid and car¬  bohydrate metabolism, J. Nutr. 130, 988S-990S (2000). [Summa¬  rizes the metabolic roles of glutamate as a fuel and as a participant  in deamination and transamination reactions.]   Dunn, M.F., Niks, D., Ngo, H., Barends, T.R.M., and Schlichting, I.,  Tryptophan synthase: the workings of a channeling nanomachine,  Trends Biochem. Sci. 33, 254-264 (2008).   Eisenberg, D., Gill, H.S., Pfluegl, M.U., and Rotstein, S.H.,  Structure-function relationships of glutamine synthetases, Biochim.  Biophys. Acta 1477, 122-145 (2000).   Eliot, A.C. and Kirsch, J.E, Pyridoxal phosphate enzymes, Annu. Rev.  Biochem. 73, 383-415 (2004).    Huang, X., Holden, H.M., and Raushel, EM., Channeling of substrates  and intermediates in enzyme-catalyzed reactions, Annu. Rev.  Biochem. 70, 149-180 (2001).   Katagiri, M. and Nakamura, M., Animals are dependent on preformed  a-amino nitrogen as an essential nutrient, Life 53, 125-129 (2002).   Maron, B.A. and Loscalzo, J., The treatment of hyperhomocysteinemia,  Annu. Rev. Med. 60, 39-54 (2009).   Medina, M.A., Urdiales, J.L., and Amores-Sanchez, M.I., Roles of homo¬  cysteine in cell metabolism: Old and new functions, Eur. J. Biochem.  268, 3871-3882 (2001).   Smith, T.J. and Stanley, C.A., Untangling the glutamate dehydrogenase  allosteric nightmare, Trends Biochem. Sci. 33, 557-564 (2008).   Valle, D. (Ed.), The Online Metabolic & Molecular Bases of Inherited Dis¬  ease, http://www.ommbid.com/. [Part 8 contains numerous chapters  on defects in amino acid metabolism.]   Howard, J.B. and Rees, D.C., How many metals does it take to fix N 2 ?  A mechanistic overview of biological nitrogen fixation, Proc. Natl.  Acad. Sci. 103, 17088-17093 (2006).   Igarashi, R.Y. and Seefeldt, L.C., Nitrogen fixation: The mechanism of  Mo-dependent nitrogenase, Crit. Rev. Biochem. Mol. Biol. 38,  351-384 (2003).   Seefeldt, L.C., Hoffman, B.M., and Dean, D.R., Mechanisms of Mo-  dependent nitrogenase, Annu. Rev. Biochem. 78, 701-722 (2009).    This human adipocyte, or fat cell, is specialized for the synthesis and storage of  triacylglycerols. The amount of fat deposited here depends on a complex signaling  system that coordinates fuel acquisition, use, and storage throughout the body.  [Science Photo Library/Photo Researchers, Inc.]    In even the simplest prokaryotic cell, metabolic processes must be coordinated  so that opposing pathways do not operate simultaneously and so that the or¬  ganism can respond to changing external conditions such as the availability of  nutrients. In addition, the organisms metabolic activities must meet the de¬  mands set by genetically programmed growth and reproduction. The chal¬  lenges of coordinating energy acquisition and expenditure are markedly more  complex in multicellular organisms, in which individual cells must cooperate.  In animals and plants, this task is simplified by the division of metabolic  labor among tissues.   In animals, the interconnectedness of various tissues is ensured by neu¬  ronal circuits and by hormones. Such regulatory systems do not simply switch  cells on and off but elicit an almost infinite array of responses. The exact re¬  sponse of a cell to a given regulatory signal depends on the cells ability to rec¬  ognize the signal and on the presence of synergistic or antagonistic signals.   Our examination of mammalian carbohydrate, lipid, and amino acid  metabolism (Chapters 15 - 21 ) would be incomplete without a discussion of  how such processes are coordinated at the molecular level and how their mal¬  functions produce disease. In this chapter, we summarize the specialized me¬  tabolism of different organs and the pathways that link them. We also  examine the mechanisms by which extracellular hormones influence intra¬  cellular events. We conclude with a discussion of disruptions in mammalian  fuel metabolism.   A The Brain Requires a Steady Supply of Glucose  B Muscle Utilizes Glucose, Fatty Acids, and  Ketone Bodies   E Kidney Filters Wastes and Maintains Blood pH  F Blood Transports Metabolites in Interorgan  Metabolic Pathways   A Insulin Release Is Triggered by Glucose  B Glucagon and Catecholamines Counter the  Effects of Insulin   3 Metabolic Homeostasis: The Regulation of Energy  Metabolism, Appetite, and Body Weight   B Adipocytes and Other Tissues Help Regulate  Fuel Metabolism and Appetite  C Energy Expenditure Can Be Controlled by  Adaptive Thermogenesis   A Starvation Leads to Metabolic Adjustments  B Diabetes Mellitus Is Characterized by High  Blood Glucose Levels  C Obesity Is Usually Caused by Excessive  Food Intake   • The major metabolic pathways for glucose, fatty acids, and amino acids center on  pyruvate and acetyl-CoA.   • Adipose tissue stores triacylglycerols and releases fatty acids as needed.   • The liver makes all types of fuel available to other tissues.    Many of the metabolic pathways discussed so far have to do with the oxida¬  tion of metabolic fuels for the production of ATP. These pathways, which en¬  compass the synthesis and breakdown of glucose, fatty acids, and amino acids,  are summarized in Fig. 22-1.   1. Glycolysis. The metabolic degradation of glucose begins with its con¬  version to two molecules of pyruvate with the net generation of two  molecules of ATP (Section 15-1).   2. Gluconeogenesis. Mammals can synthesize glucose from a variety of  precursors, such as pyruvate, via a series of reactions that largely re¬  verse the path of glycolysis (Section 16-4).   FIG. 22-1 The major pathways of fuel metabolism in mammals. Proteins,  glycogen, and triacylglycerols are built up from and broken down to smaller units: amino  acids, glucose-6-phosphate, and fatty acids. Oxidation of those fuels yields metabolic  energy in the form of ATP. Pyruvate (a product of glucose and amino acid degradation) and  acetyl-CoA (a product of glucose, amino acid, and fatty acid degradation) occupy central  positions in mammalian fuel metabolism. Compounds that give rise to pyruvate, such as  oxaloacetate, can be used for gluconeogenesis; acetyl-CoA can give rise to ketone bodies  but not glucose. Not all the pathways shown here occur in all cells or occur simultaneously  in a given cell.   alyzed by glycogen phosphorylase and glycogen synthase are recipro¬  cally regulated by hormonally controlled phosphorylation and   4. Fatty acid synthesis and degradation. Fatty acids are broken down  through (3 oxidation to form acetyl-CoA (Section 20-2), which,  through its conversion to malonyl-CoA, is also the substrate for fatty  acid synthesis (Section 20-4).   5. The citric acid cycle. The citric acid cycle (Section 17-1) oxidizes  acetyl-CoA to C0 2 and H 2 0 with the concomitant production of re¬  duced coenzymes whose reoxidation drives ATP synthesis. Many gluco¬  genic amino acids can be oxidized via the citric acid cycle following  their breakdown to one of its intermediates (Section 21-4), which, in  turn, are broken down to pyruvate and then to acetyl-Co A, the cycles  only substrate.   6. Oxidative phosphorylation. This mitochondrial pathway couples the  oxidation of NADH and FADH 2 produced by glycolysis, (3 oxidation,  and the citric acid cycle to the phosphorylation of ADP (Section 18-3).   7. Amino acid synthesis and degradation. Excess amino acids are de¬  graded to metabolic intermediates of glycolysis and the citric acid cy¬  cle (Section 21-4). The amino group is disposed of through urea  synthesis (Section 21-3). Nonessential amino acids are synthesized via  pathways that begin with common metabolites (Section 21-5A).   Two compounds lie at the crossroads of the major metabolic pathways:  acetyl-CoA and pyruvate (Fig. 22-1). Acetyl-CoA is the common degradation  product of glucose, fatty acids, and ketogenic amino acids. Its acetyl group  can be oxidized to C0 2 and H 2 0 via the citric acid cycle and oxidative phos¬  phorylation or used to synthesize ketone bodies or fatty acids. Pyruvate is the  product of glycolysis and the breakdown of glucogenic amino acids. It can be  oxidatively decarboxylated to yield acetyl-CoA, thereby committing its atoms  either to oxidation or to the biosynthesis of fatty acids. Alternatively, pyruvate  can be carboxylated via the pyruvate carboxylase reaction to form oxaloacetate,  which can either replenish citric acid cycle intermediates or give rise to glu¬  cose or certain amino acids.   Only a few tissues , such as liver ; can carry out all the reactions shown in Fig.   22-1 , and in a given cell only a small portion of all possible metabolic reactions  occur at a significant rate. The flux through any sequence of reactions depends  on the presence of the appropriate enzyme catalysts and on the organisms need  for the reaction products.   We will consider the metabolism of five mammalian organs: brain, mus¬  cle, adipose tissue, liver, and kidney. Metabolites flow between these organs in  well-defined pathways in which flux varies with the nutritional state of the an¬  imal (Fig. 22-2). For example, immediately following a meal, glucose, amino  acids, and fatty acids are directly available from the intestine. Later, when those  fuels have been exhausted, the liver supplies other tissues with glucose and ke¬  tone bodies, whereas adipose tissue provides them with fatty acids. All these  organs are connected via the bloodstream. In addition, the metabolic activi¬  ties of numerous microorganisms contribute to mammalian fuel metabolism  (Box 22-1).   Brain tissue has a remarkably high respiration rate. Although the human brain  constitutes only —2% of the adult body mass, it is responsible for —20% of  its resting 0 2 consumption. Most of the brains energy production powers the  plasma membrane (Na + -K + )-ATPase (Section 10-3A), which maintains the  membrane potential required for nerve impulse transmission.    Under usual conditions, glucose is the brains primary fuel (although dur¬  ing an extended fast, the brain gradually switches to ketone bodies; Section  22-4A). Since brain cells store very little glycogen, they require a steady supply  of glucose from the blood. A blood glucose concentration of less than half the  normal value of ~5 mM results in brain dysfunction. Levels much below this  result in coma, irreversible damage, and ultimately death.   Muscles major fuels are glucose (from glycogen), fatty acids, and ketone bod¬  ies. Rested, well-fed muscle synthesizes a glycogen store comprising 1 to 2%  of its mass. Although triacylglycerols are a more efficient form of energy stor¬  age (Section 9-IB), the metabolic effort of synthesizing glycogen is cost-  effective because glycogen can be mobilized more rapidly than fat and because  glucose can be metabolized anaerobically, whereas fatty acids cannot.    The human body contains about ten trillion (10 13 ) cells, and there  are probably ten times that number of microorganisms living in the  intestine. These organisms, mostly bacteria, form an integrated  community called the microbiome. At one time, the existence of  microbes inside the human host was believed to be a form of  commensalism, a relationship in which neither party has much to  gain or lose in the partnership. It is now clear, however, that the  microbiome plays an active role in providing nutrients (including  some vitamins), regulating fuel use and storage, and preventing  disease.   DNA sequencing studies have revealed that an individual may  host several hundred to a thousand different species. The mixture  remains fairly constant throughout the person’s lifetime but can  vary markedly between individuals, even in the same household. It  is likely that a few species are common to all humans, constituting  a sort of “core” microbiome. The Human Microbiome Project  (https://commonfund.nih.gov/hmp/) aims to better characterize the    species that inhabit the human body and assess their contribution  to human health and disease.   Bacteria and fungi in the small intestine ferment undigested  carbohydrates, mainly polysaccharides that cannot be broken down  by human digestive enzymes, and produce acetate, propionate, and  butyrate. These short-chain fatty acids are absorbed by the host  and are transformed into triacylglycerols for long-term storage.  There is evidence that thin and obese individuals harbor different  species of gut bacteria, suggesting that certain microbial species  might play a role in releasing more calories from food—calories  that would then be stored as fat. Intestinal bacteria also produce  vitamin K, biotin, and folate, some of which can be taken up and  used by the host. The importance of microbial digestion is illus¬  trated by mice grown in a germ-free environment. Without the  normal bacterial partners, the mice need to consume about 30%  more food than normal animals whose digestive systems have  been colonized by microorganisms.   In muscle, glycogen is converted to glucose-6-phosphate (G6P) for entry into  glycolysis. Muscle cannot export glucose, however, because it lacks glucose-6-  phosphatase. Furthermore, although muscle can synthesize glycogen from glucose,  it does not participate in gluconeogenesis because it lacks the required enzymatic  machinery. Consequently, muscle carbohydrate metabolism serves only muscle.   Muscle contraction is driven by ATP hydrolysis (Section 7-2)  and therefore requires either an aerobic or an anaerobic ATP  regeneration system. Respiration (the citric acid cycle and ox¬  idative phosphorylation) is the body’s major source of ATP  resupply. Skeletal muscle at rest uses ~30% of the 0 2 con¬  sumed by the human body. A muscles respiration rate may  increase in response to a heavy workload by as much as 25-  fold. Yet, its rate of ATP hydrolysis can increase by a much  greater amount. The ATP is initially regenerated by the reac¬  tion of phosphocreatine with ADP (Section 14-2C):    (phosphocreatine is resynthesized in resting muscle by the re¬  versal of this reaction). Under conditions of maximum exer¬  tion, however, such as during a sprint, a muscle has only about  a 4-s supply of phosphocreatine. It must then shift to ATP  production via glycolysis of G6P, a process whose maximum  flux greatly exceeds those of the citric acid cycle and oxidative  phosphorylation. Much of the G6P is therefore degraded  anaerobically to lactate (Section 15-3A). As we will see in  Section 22-IF, export of lactate relieves much of the muscles respiratory bur¬  den. Muscle fatigue, which occurs after ~20 s of maximal exertion, is not  caused by exhaustion of the muscles glycogen supply but by the drop in pH  that results from the buildup of lactate. This phenomenon may be an adapta¬  tion that prevents muscle cells from committing suicide by fully depleting their  ATP supply. Exercise lasting more than a minute or two is fueled primarily by  oxidative phosphorylation, which generates ATP more slowly but much more  efficiently than glycolysis alone. The source of ATP during exercise of varying  duration is summarized in Fig. 22-3.   FIG. 22-3 Source of ATP during exercise  in humans. The supply of endogenous ATP is  extended for a few seconds by phosphocreatine,  after which anaerobic glycolysis generates ATP.  The shift from anaerobic to aerobic metabolism  (oxidative phosphorylation) occurs after about 90  sec, or slightly later in trained athletes. [Adapted  from McArdle, W.D., Katch, F.I., and Katch, V.L,  Exercise Physiology, 2nd ed., Lea & Febiger  (1986), p. 348.]    The Heart Is Largely Aerobic. The heart is a muscular organ that acts con¬  tinuously rather than intermittently. Therefore, heart muscle relies entirely on  aerobic metabolism and is richly endowed with mitochondria; they occupy up  to 40% of its cytoplasmic space. The heart can metabolize fatty acids, ketone  bodies, glucose, pyruvate, and lactate. Fatty acids are the resting heart’s fuel of  choice, but during heavy work, the heart greatly increases its consumption of  glucose, which is derived mostly from its relatively limited glycogen store.   The function of adipose tissue is to store and release fatty acids as needed for  fuel as well as to secrete hormones involved in regulating metabolism. Adipose  tissue is widely distributed throughout the body but occurs most prominently  under the skin, in the abdominal cavity, and in skeletal muscle. The adipose  tissue of a normal 70-kg man contains ~15 kg of fat. This amount represents  some 590,000 kj of energy (141,000 dieters Calories), which is sufficient to  maintain life for ~3 months.   Adipose tissue obtains most of its fatty acids for storage from circulating  lipoproteins as described in Section 20-IB. Fatty acids are activated by the for¬  mation of the corresponding fatty acyl-CoA and then esterified with glycerol-  3-phosphate to form the stored triacylglycerols. The glycerol-3-phosphate  arises from the reduction of dihydroxyacetone phosphate, which must be gly-  colytically generated from glucose.   In times of metabolic need, adipocytes hydrolyze triacylglycerols to fatty acids  and glycerol through the action of hormone-sensitive lipase (Section 20-5).  If glycerol-3-phosphate is abundant, many of the fatty acids so formed are  reesterified to triacylglycerols. If glycerol-3-phosphate is in short supply, the  fatty acids are released into the bloodstream. Thus, fatty acid mobilization  depends in part on the rate of glucose uptake since glucose is the precursor of  glycerol-3-phosphate. Metabolic need is signaled directly by a decrease in [glucose]  as well as by hormonal stimulation.   FIG. 22-4 Relative enzymatic activities of  hexokinase and glucokinase over the  physiological blood glucose range. Glucokinase  has much lower affinity for glucose (K M ~ 5 mM)  than does hexokinase (K M ~ 0.1 mM) and  exhibits sigmoidal rather than hyperbolic variation  with [glucose]. [The glucokinase curve was  generated using the Hill equation (Eq. 7-8) with  K = 10 mM and n = 1.5 as obtained from  Cardenas, M.L, Rabajille, E., and Niemeyer, H.,   The liver maintains the proper levels of circulating fuels for use by  the brain, muscles, and other tissues. It is uniquely situated to carry  out this task because all the nutrients absorbed by the intestines  except fatty acids are released into the portal vein, which drains  directly into the liver.   One of the livers major functions is to act as a blood glucose “buffer. ” It  does so by taking up and releasing glucose in response to hormones  and to the concentration of glucose itself. After a carbohydrate-con¬  taining meal, when the blood glucose concentration reaches ~6 mM,  the liver takes up glucose by converting it to G6P. This reaction is  catalyzed by glucokinase, a liver isozyme of hexokinase. The hexo-  kinases in most cells obey Michaelis—Menten kinetics, have a high glu¬  cose affinity (K M <0.1 mM), and are inhibited by their reaction product (G6P).  Glucokinase, in contrast, has much lower glucose affinity (reaching half-maximal  velocity at ~5 mM) and displays sigmoidal kinetics. Consequently, glucokinase ac¬  tivity increases rapidly with blood [glucose] over the normal physiological range (Fig.  22-4). Glucokinase, moreover, is not inhibited by physiological concentrations of  G6P. Therefore, the higher the blood [glucose], the faster the liver converts glu¬  cose to G6P. At low blood [glucose], the liver does not compete with other tis¬  sues for the available glucose, whereas at high blood [glucose], when the glucose  needs of those tissues are met, the liver can take up the excess glucose at a rate  roughly proportional to the blood glucose concentration.    Glucokinase is a monomeric enzyme, so its sigmoidal kinetic behavior is  somewhat puzzling (models of allosteric interactions do not explain coopera¬  tive behavior in a monomeric protein; Section 7-ID). Glucokinase is subject  to metabolic control, however. Emile Van Schaftingen has isolated a gluco¬  kinase regulatory protein from rat liver, which, in the presence of the gly¬  colytic intermediate fructose-6-phosphate (F6P), is a competitive inhibitor of  glucokinase. Since F6P and the glucokinase product G6P are equilibrated in  liver cells by phosphoglucose isomerase, glucokinase is, in effect, inhibited by  its product. Fructose-1-phosphate (F1P), an intermediate in liver fructose me¬  tabolism (Section 15-5A), overcomes this inhibition. Since fructose is normally  available only from dietary sources, fructose may be the signal that triggers the  uptake of dietary glucose by the liver.   G6P has several alternative fates in the liver, depending on the glucose de¬  mand (Fig. 22-5):   1. G6P can be converted to glucose, by the action of glucose-6-  phosphatase, for transport via the bloodstream to the peripheral  organs. This occurs only when the blood [glucose] drops below ~5 mM.  During exercise or fasting, low concentrations of blood glucose cause  the pancreas to secrete glucagon. Glucagon receptors on the liver cell  surface respond by activating adenylate cyclase. The resulting increase  in intracellular [cAMP] triggers glycogen breakdown (Section 16-3).   2. G6P can be converted to glycogen (Section 16-2) when the body’s de¬  mand for glucose is low.   3. G6P can be converted to acetyl-CoA via glycolysis and the action of  pyruvate dehydrogenase. This glucose-derived acetyl-Co A, if it is not  oxidized via the citric acid cycle and oxidative phosphorylation to gen¬  erate ATP, can be used to synthesize fatty acids (Section 20-4), phos¬  pholipids (Section 20-6), and cholesterol (Section 20-7A).   4. G6P can be degraded via the pentose phosphate pathway (Section 15-6)  to generate the NADPH required for the biosynthesis of fatty acids and  other compounds.    FIG. 22-5 Metabolic fate of glucose-6-  phosphate (G6P) in liver. G6P can be   converted (1) to glucose for export or (2) to  glycogen for storage. Acetyl-CoA derived from  G6P degradation (3) is the starting material for  lipid biosynthesis. It is also consumed in  generating ATP by respiration. Degradation of  G6P via the pentose phosphate pathway  (4) yields NADPH.    The Liver Can Synthesize or Degrade Triacylglycerols. Fatty acids are also  subject to alternative metabolic fates in the liver. When the demand for meta¬  bolic fuels is high, fatty acids are degraded to acetyl-CoA and then to ketone  bodies for export to the peripheral tissues. The liver itself cannot use ketone bod¬  ies as fuel, because liver cells lack 3-ketoacyl-CoA transferase, an enzyme required  to convert ketone bodies back to acetyl-CoA (Section 20-3). Fatty acids rather  than glucose or ketone bodies are therefore the livers major acetyl-CoA source  under conditions of high metabolic demand. The liver generates its ATP from  this acetyl-CoA through the citric acid cycle and oxidative phosphorylation.   When the demand for metabolic fuels is low, fatty acids are incorporated into  triacylglycerols that are secreted into the bloodstream as VLDF for uptake by adi¬  pose tissue. Fatty acids are also incorporated into phospholipids (Section 20-6).  Under these conditions, fatty acids synthesized in the liver are not oxidized to  acetyl-CoA because fatty acid synthesis (in the cytosol) is separated from fatty  acid oxidation (in the mitochondria) and because malonyl-CoA, an intermediate  in fatty acid synthesis, inhibits the transport of fatty acids into the mitochondria.   Amino Acids Are Metabolic Fuels. The liver degrades amino acids to a va¬  riety of metabolic intermediates that can be completely oxidized to C0 2 and  H 2 0 or converted to glucose or ketone bodies (Section 21-4). Oxidation of  amino acids provides a significant fraction of metabolic energy immediately  after a meal, when amino acids are present in relatively high concentrations   Chapter 22 Mammalian Fuel Metabolism: Integration [ n t he blood. During a fast, when other fuels become scarce, glucose is pro-   alanine and glutamine. Thus, proteins , in addition to their structural and func¬  tional roles , are important fuel reserves.   The kidney filters urea and other waste products from the blood while it recov¬  ers important metabolites such as glucose. In addition, the kidney maintains the  bloods pH by regenerating depleted blood buffers such as bicarbonate (lost by  the exhalation of C0 2 ) and by excreting excess H + together with the conjugate  bases of excess metabolic acids such as the ketone bodies acetoacetate and |3-  hydroxybutyrate. Protons are also excreted in the form of NH 4 with the am¬  monia derived from glutamine or glutamate. The remaining amino acid skele¬  ton, a-ketoglutarate, can be converted to glucose by gluconeogenesis (the kidney  is the only tissue besides liver that can undertake glucose synthesis). During  starvation, the kidneys generate as much as 50% of the body’s glucose supply.   The ability of the liver to supply other tissues with glucose or ketone bodies,  or the ability of adipocytes to make fatty acids available to other tissues, de¬  pends, of course, on the circulatory system, which transports metabolic fuels,  intermediates, and waste products among tissues. In addition, several impor¬  tant metabolic pathways are composed of reactions occurring in multiple tis¬  sues. In this section, we describe two well-known interorgan pathways.   Glucose and Lactate Are Transported in the Cori Cycle. The ATP that pow¬  ers muscle contraction is generated through oxidative phosphorylation (in mi¬  tochondrion-rich slow-twitch muscle fibers; Box 15-3) or by rapid catabolism  of glucose to lactate (in fast-twitch muscle fibers). Slow-twitch fibers also pro¬  duce lactate when ATP demand exceeds oxidative flux. The lactate is transferred  via the bloodstream to the liver, where it is reconverted to pyruvate by lactate  dehydrogenase and then to glucose by gluconeogenesis. Thus, liver and muscle  are linked by the bloodstream in a metabolic cycle known as the Cori cycle  (Fig. 22-6) in honor of Carl and Gerty Cori (Box 16-1), who first described it.   The ATP-consuming glycolysis/gluconeogenesis cycle would be a futile cy¬  cle if it occurred within a single cell. In this case, the two halves of the path¬  way occur in different organs. Liver ATP is used to resynthesize glucose from  lactate produced in muscle. The resynthesized glucose returns to the muscle,  where it may be stored as glycogen or catabolized immediately to generate  ATP for muscle contraction.   The ATP consumed by the liver during the operation of the Cori cycle is  regenerated by oxidative phosphorylation. After vigorous exertion, it may take   Lactate produced by muscle  glycolysis is transported by the  bloodstream to the liver, where  it is converted to glucose by  gluconeogenesis. The bloodstream  carries the glucose back to the  muscle, where it may be stored as  glycogen. Q See the Animated  Figures.   FIG. 22-7 The glucose-  alanine cycle. Pyruvate produced  by muscle glycolysis is the  amino-group acceptor for muscle  aminotransferases. The resulting  alanine is transported by the  bloodstream to the liver, where it is  converted back to pyruvate (its  amino group is disposed of via urea  synthesis). The pyruvate is a  substrate for gluconeogenesis, and  the bloodstream carries the resulting  glucose back to the muscles.    at least 30 min for the oxygen consumption rate to decrease to its resting level.  The elevated 0 2 consumption pays off the oxygen debt created by the de¬  mand for ATP to drive gluconeogenesis.   The Glucose-Alanine Cycle Transfers Alanine to the Liver. In a pathway  similar to the Cori cycle, alanine rather than lactate travels from muscle to the  liver. In muscle, certain aminotransferases use pyruvate as their a-keto acid  substrate rather than a-ketoglutarate or oxaloacetate (Section 21-2A):   R—CH —COO - + H 3 C —C—COO - R—C—COO - + H 3 C—CH—COO -   The product amino acid, alanine, is released into the bloodstream and trans¬  ported to the liver, where it undergoes transamination back to pyruvate. This  pyruvate is a substrate for gluconeogenesis, and the resulting glucose can be  returned to the muscles to be glycolytically degraded. This is the glucose-  alanine cycle (Fig. 22-7). The amino group carried by alanine ends up in  either ammonia or aspartate and can be used for urea biosynthesis (which  occurs only in the liver). Thus, the glucose—alanine cycle is a mechanism for  transporting nitrogen from muscle to liver.   During fasting, the glucose formed in the liver by this route is also used  by other tissues, breaking the cycle. Because the pyruvate originates from mus¬  cle protein degradation, muscle supplies glucose to other tissues even though  it does not carry out gluconeogenesis.   • Summarize the importance of pyruvate  and acetyl-CoA for catabolism and  anabolism.   • Without looking at Fig. 22-1, draw a  diagram of the major metabolic pathways  involving proteins, glycogen, and triacyl-  glycerols.   • Summarize the major features of fuel  metabolism in the brain, muscle, adipose  tissue, liver, and kidney.   • Explain why the high K M of glucokinase is  important for the role of the liver in buffer¬  ing blood glucose.   • List all the possible fates for a glucose  molecule in the liver.   • Describe the conditions under which the  Cori cycle and the glucose-alanine cycle  operate.   • Without looking at the text, draw diagrams  of these two metabolic cycles.    Insulin release in response to glucose promotes fuel uptake and storage.  Glucagon and the catecholamines promote fuel mobilization.    Multicellular organisms coordinate their activities at every level of their organ¬  ization through complex signaling systems (Chapter 13). Most cells in a mul¬  ticellular organism cannot take up fuel for growth and metabolism without first  receiving a signal to do so. The human endocrine system responds to the needs  of an organism by secreting a wide variety of hormones that enable the body  to maintain metabolic homeostasis (balance between energy inflow and out¬  put), respond to external stimuli, and follow various developmental programs.   We have already discussed steroid hormones (Section 9-IE), the peptide hor¬  mones insulin and glucagon (Sections 13-1 A, 13-4D, and 16-3C), and the cat¬  echolamines epinephrine and norepinephrine (Sections 13-IB, 16-3C, and    21-6B). With the exception of steroids, which can diffuse through cell mem¬  branes and interact directly with intracellular components (and which we discuss  further in Section 28-3B), these extracellular signals must first bind to a cell-  surface receptor. In this section, we review the action of hormones synthesized  by the pancreas and adrenal glands, since these play the largest roles in regulating  the metabolism of fuels in various mammalian tissues. We will also review the  receptors and pathways by which these hormones exert their effects on cells.   The pancreas responds to increases in the concentration of blood glucose by  secreting insulin, which therefore serves as a signal for plentiful metabolic fuel.  Pancreatic (3 cells are most sensitive to glucose at concentrations of 5.5 to 6.0 mM  (normal blood glucose concentrations range from 3.6 to 5.8 mM). There is  no evidence for a cell-surface glucose “receptor” that might relay a signal to  the secretory machinery in the (3 cell. In fact, glucose enters (3 cells via pas¬  sive transport, and its metabolism generates the signal for insulin secretion.   The rate-limiting step of glucose metabolism in (3 cells is the reaction cat¬  alyzed by glucokinase (the same enzyme that occurs in hepatocytes).  Consequently, glucokinase is considered the (3 cells glucose “sensor.”  Glucokinases G6P product is not used to synthesize glycogen, and the activ¬  ity of the pentose phosphate pathway is minor. Furthermore, lactate dehydro¬  genase activity is low. As a result, essentially all the G6P produced in (3 cells  is degraded to pyruvate and then converted to acetyl-CoA for oxidation by  the citric acid cycle. This one-way, linear catabolic pathway for glucose directly  links the (3 cells rate of oxidative phosphorylation to the amount of available  glucose. By mechanisms that are not entirely understood, the overall level of  the (3 cells respiratory activity regulates insulin synthesis and secretion.   Insulin Promotes Fuel Storage in Muscle and Adipose Tissue. Insulin sig¬  naling is very complex (Fig. 13-31). It acts as the primary regulator of blood  glucose concentration by promoting glucose uptake in muscles and adipose  tissue and by inhibiting hepatic glucose production. Insulin also stimulates cell  growth and differentiation by increasing the synthesis of glycogen, proteins,  and triacylglycerols.   Muscle cells and adipocytes express an insulin-sensitive glucose trans¬  porter known as GLUT4. Insulin stimulates GLUT4 activity. Such an  increase can occur through an increase in the intrinsic activity of the  transporter molecules, but in the case of GLUT4, the increase is ac¬  complished through the appearance of additional transporter mol¬  ecules in the plasma membrane (Fig. 22-8). In the absence of  insulin, GLUT4 is localized in intracellular vesicles and tubular  structures known as GLUT4 storage vesicles. Insulin promotes  the fusion of these vesicles to the plasma membrane in a process  that is mediated by SNAREs (Section 9-4F). GLUT4 appears  on the cell surface only a few minutes after insulin stimulation.  GLUT4 has a relatively low K M for glucose (2—5 mM), so cells  containing this transporter can rapidly take up glucose from the  blood. On insulin withdrawal, the glucose transporters are grad¬  ually sequestered through endocytosis.   The insulin-dependent GLUT4 transport system allows muscle  and adipose tissue to quickly stockpile metabolic fuel immediately    FIG. 22-8 GLUT4 activity. Glucose uptake in muscle and fat cells is  regulated by the insulin-stimulated exocytosis (the opposite of endocytosis)  Plasma of specialized membranous vesicles containing GLUT4 (left). On insulin   after a meal. Tissues such as the brain, which uses glucose almost exclusively as Section 2 Hormonal Control of Fuel Metabolism  a fuel, constitutively express an insulin-insensitive glucose transporter.   Consequently, the central nervous system does not experience large fluctuations  in glucose absorption. Significantly, the liver also lacks GLUT4 and therefore does  not respond to increases in insulin levels by increasing its rate of glucose uptake.   Once glucose enters cells, it can be used to synthesize glycogen (in mus¬  cle) and triacylglycerols (in adipocytes; the glucose must first be metabolized  to acetyl-CoA for fatty acid synthesis). Insulin specifically promotes those  metabolic activities. As discussed in Section 16-3, insulin activates glycogen  synthase by promoting its dephosphorylation. In adipocytes, insulin activates  the pyruvate dehydrogenase complex (by activating the associated phosphatase;   Section 17-4A), and it activates acetyl-CoA carboxylase and increases levels of  fatty acid synthase (Section 20-5). At the same time, insulin inhibits lipolysis  by inhibiting hormone-sensitive lipase.   Insulin Blocks Liver Gluconeogenesis and Glycogenolysis. Although the  liver does not respond to insulin by increasing its rate of uptake of glucose, in¬  sulin binding to its receptor on hepatocytes has several consequences. The inac¬  tivation of phosphorylase kinase decreases the rate of glycogenolysis, and the  activation of glycogen synthase promotes glycogen synthesis. Insulin also inhibits  transcription of the genes encoding the gluconeogenic enzymes phospho-  enolpyruvate carboxykinase, fructose-1,6-bisphosphatase, and glucose-6-phos-  phatase (Section 16-4) and stimulates transcription of the genes for the glycolytic  enzymes glucokinase and pyruvate kinase. The expression of lipogenic enzymes  such as acetyl-CoA carboxylase and fatty acid synthase also increases. The result  of these regulatory changes is that the liver stores glucose (as glycogen and as tri-  acylglycerols) rather than producing glucose by glycogenolysis or gluconeogenesis. The  major metabolic effects of insulin are summarized in Table 22-1.   As we discussed in Section 16-3C, the peptide hormone glucagon activates a  series of intracellular events that lead to glycogenolysis in the liver. This con¬  trol mechanism helps make glucose available to other tissues when the con¬  centration of circulating glucose drops. Muscle cells, which lack a glucagon  receptor and cannot respond directly to the hormone, benefit indirectly from  the glucose released by the liver. Glucagon also stimulates fatty acid mobiliza¬  tion from adipose tissue by activating hormone-sensitive lipase.   The catecholamines elicit responses similar to glucagons. Epinephrine and  norepinephrine, which are released during times of stress, bind to two different  types of receptors (Section 13-IB): the (3-adrenergic receptor, which is linked to  the adenylate cyclase system (Fig. 13-23), and the a-adrenergic receptor,  whose second messenger, inositol-1,4,5-trisphosphate (IP3; Section 13-4A),  causes intracellular Ca 2+ concentrations to increase.    Liver cells respond to epinephrine directly and indirectly. Epinephrine pro¬  motes the release of glucagon from the pancreas, and glucagon binding to its  receptor on liver cells stimulates glycogen breakdown. Epinephrine also binds  directly to both a- and (3-adrenergic receptors on the surfaces of liver cells  (Fig. 16-14, righi). Binding to the (3-adrenergic receptor results in increased  intracellular cAMP, which leads to glycogen breakdown and gluconeogenesis.  Epinephrine binding to the a-adrenergic receptor stimulates an increase in in¬  tracellular [Ca 2+ ], which reinforces the cells’ response to cAMP (recall that  phosphorylase kinase, which activates glycogen phosphorylase and inactivates  glycogen synthase, is fully active only when phosphorylated and in the pres¬  ence of increased [Ca 2+ ]; Section 16-3B). In addition, glycogen synthase is  inactivated through phosphorylation catalyzed by several Ca 2+ -dependent  protein kinases.   Epinephrine binding to the (3-adrenergic receptor on muscle cells simi¬  larly promotes glycogen degradation, thereby mobilizing glucose that can be  metabolized by glycolysis to produce ATP. In adipose tissue, epinephrine bind¬  ing to several a- and (3-type adrenergic receptors leads to activation of  hormone-sensitive lipase, which results in the mobilization of fatty acids that   (a) Immediately after a meal, when glucose and fatty acids are  abundant, insulin signals tissues to store fuel as glycogen and  triacylglycerols. Insulin also stimulates tissues other than liver to take    up glucose via the GLUT4 transporter, (b) When dietary fuels are not  available, glucagon stimulates the liver to release glucose and adipose  tissue to release fatty acids. During stress, epinephrine elicits similar  responses.   can be used as fuels by other tissues. In addition, epinephrine stimulates smooth Section 2 Hormonal Control of Fuel Metabolism   muscle relaxation in the bronchi and blood vessels supplying skeletal muscle,   while it stimulates constriction of the blood vessels that supply skin and other   sudden action by mobilizing energy reserves and directing them where they are   needed. The major responses to glucagon and epinephrine are summarized in   Table 22 - 1 , and the metabolic effects of insulin, glucagon, and epinephrine   Several Types of Receptors and Biochemical Signaling Pathways  Participate in Metabolic Regulation. As described in Chapter 13, every sig¬  naling pathway consists of a receptor, a mechanism for transmitting the  ligand-binding event to the cell interior, and a series of intracellular responses  that may involve a second messenger and/or chemical changes catalyzed by ki¬  nases and phosphatases. There are three major signal transduction pathways:  those involving receptor tyrosine kinases associated with the Ras signaling cas¬  cade (Section 13-2), those in which adenylate cyclase generates cAMP (3',5'-  cyclic AMP) as a second messenger (Section 13-3), and those involving  phosphoinositide cleavage by phospholipase C (PKC), with inositol-1,4,5-  trisphosphate (IP 3 ) and 1 , 2 -diacylglycerol (DAG), as well as Ca 2+ , as second  messengers (Section 13-4). Both second messenger systems involve G pro¬  tein-coupled receptors (Section 13-3). The major G protein activating aden¬  ylate cyclase is G s , and the major G protein activating phospholipase C is G q .   Second messenger cAMP Second messengers IP 3 , DAG, Ca 2+    FIG. 22-10 Overview of the major signal transduction  pathways, (a) The binding of a ligand to a receptor tyrosine kinase  initiates the Ras signaling cascade of phosphorylations, which leads  to changes in gene expression. ( b ) The binding of a ligand to its  corresponding G protein-coupled receptor stimulates (or in some  cases inhibits) adenylate cyclase (AC) to synthesize the second  messenger cAMR The cAMP, in turn, activates protein kinase A  (PKA) to phosphorylate its target proteins, leading to a cellular    response, (c) The binding of a ligand to its corresponding G q  protein-coupled receptor initiates the phosphoinositide pathway, which  activates phospholipase C (PLC) to hydrolyze the membrane lipid PIP 2  to the second messengers IP 3 and DAG. IP 3 causes the release of  Ca 2+ from mitochondria, DAG activates protein kinase C (PKC), and  Ca 2+ activates both PKC and Ca 2+ -calmodulin dependent protein  kinase (CaMK). The phosphorylation of target proteins by PKC and  CaMK leads to cellular responses.   • Summarize the role of insulin in regulating  fuel metabolism in the liver, muscle, and  adipose tissue.   • How do glucagon and epinephrine affect  fuel metabolism in liver, muscle, and  adipose tissue?   • Describe the three main types of signaling  pathways that participate in metabolic  regulation.    The hormones described above and in Section 13-1, as well as many other  hormones, growth factors, and other signaling molecules, use signal-transduction  pathways to set in motion a series of biochemical reactions that produce bio¬  logical responses such as altered metabolism, cell differentiation, and cell  growth and division. The exact nature of the response depends on many fac¬  tors. Cells respond to a hormone, generally referred to as a ligand, only if they  display the appropriate receptor. Specific intracellular responses are modulated  by the number, type, and cellular locations of the elements of the signaling  systems. Furthermore, a given cell typically contains receptors for many differ¬  ent ligands, so the response to one particular ligand may depend on the level of  engagement of the other ligands with the cells signal-transduction machinery.  As a result, cells can react to discrete signals or combinations of signals with varia¬  tions in the magnitude and duration of the cellular response.    3 Metabolic Homeostasis: The Regolatioo of  Energy Metabolism, Appetite, and Body Weight    • AMP-dependent protein kinase activates ATP-generating processes and inhibits  ATP-consuming processes.   • Fuel use and appetite are regulated by the adipose tissue hormones adiponectin  and leptin as well as by hormones produced by the hypothalamus, stomach, and  intestine.    The complex mechanisms that regulate mammalian fuel metabolism permit  the body to respond efficiently to changing energy demands and to accom¬  modate changes in the availability of various fuels, maintaining reasonable  metabolic homeostasis. This balance depends on the interactions of many fac¬  tors. In this section we discuss some of the participants in this complex net¬  work regulating energy expenditure, appetite, and body weight.   All of the metabolic pathways discussed above are affected in one way or an¬  other by the need for ATP, as is indicated by the cells AMP-to-ATP ratio  (Section 15-4A). Several enzymes are either activated or inhibited allosterically  by AMP, and several others are phosphorylated by AMP-dependent protein  kinase (AMPK), a major regulator of metabolic homeostasis. AMPK activates  metabolic breakdown pathways that generate ATP while inhibiting biosynthetic  pathways so as to conserve ATP for more vital processes. AMPK is an a(3y het-  erotrimer found in all eukaryotic organisms from yeast to humans. The a sub¬  unit contains a Ser/Thr protein kinase domain, and the y subunit contains  sites for allosteric activation by AMP and inhibition by ATP. Like other pro¬  tein kinases, AMPKs kinase domain must be phosphorylated for activity.  Binding of AMP to the y subunit causes a conformational change that ex¬  poses Thr 172 in the activation loop of the a subunit, promoting its phos¬  phorylation and increasing its activity at least 100-fold. AMP can activate the  phosphorylated enzyme up to 5-fold more. The major kinase that phos-  phorylates AMPK is named LKB1. The specific knockout of LKB1 in mouse  liver results in the loss of the phosphorylated form of AMPK.   AMPK Activates Glycolysis in Ischemic Cardiac Muscle. AMPKs targets in¬  clude the heart isozyme of the bifunctional enzyme PFK-2/FBPase-2, which con¬  trols the fructose-2,6-bisphosphate (F2,6P) concentration (Section 16-4C). The  phosphorylation of this isozyme activates the PFK-2 activity, increasing [F2,6P],  which in turn activates PFK-1 and glycolysis. Consequently, in ischemic (blood-   starved) heart muscle cells, when there is insufficient oxygen for oxidative phos- Section 3 Metabolic Homeostasis: The Regulation  phorylation to maintain adequate concentrations of ATP, the resulting AMP of Energy Metabolism, Appetite, and Body Weight  buildup causes the cells to switch to anaerobic glycolysis for ATP production.    AMPK Inhibits Lipogenesis and Gluconeogenesis in Liver. AMPK-medi-  ated phosphorylation also inhibits acetyl-CoA carboxylase (ACC, which cat¬  alyzes the first committed step of fatty acid synthesis; Section 20-4B),  hydroxymethylglutaryl-CoA reductase (HMG-CoA reductase, which catalyzes  the rate-determining step in cholesterol biosynthesis; Section 20-7B), and  glycogen synthase (which catalyzes the rate-limiting reaction in glycogen syn¬  thesis; Section 16-2B). Consequently, when the rate of ATP production is in¬  adequate, these biosynthetic pathways are turned off, thereby conserving ATP  for the most vital cellular functions.    AMPK Promotes Fatty Acid Oxidation and Glucose Uptake in Skeletal  Muscle. The inhibition of ACC results in a decrease in the concentration of  malonyl-CoA, the starting material for fatty acid biosynthesis. Malonyl-CoA  has an additional role, however. It is an inhibitor of carnitine palmitoyl trans¬  ferase I (Section 20-5), which is required to transfer cytosolic palmitoyl-CoA  into the mitochondria for oxidation. The decrease in malonyl-CoA concentra¬  tion therefore allows more palmitoyl-CoA to be oxidized. AMPK also increases  the expression of GLUT4 in muscle cells, as well as its recruitment to the  muscle cell plasma membrane (Section 22-2F), thus facilitating the insulin-  independent entry of glucose into the cells.   AMPK Inhibits Lipolysis in Adipocytes. AMPK phosphorylates hormone-  sensitive triacylglycerol lipase in adipose tissue (Section 20-5). This phos¬  phorylation inhibits rather than activates the enzyme, in part by preventing  the relocation of the enzyme to the lipid droplet, the cellular location of lipoly¬  sis. As a result, fewer triacylglycerols are broken down so that fewer fatty acids  are exported to the bloodstream. This latter process seems paradoxical (fatty  acid oxidation would help relieve an ATP deficit), although it has been spec¬  ulated that it prevents the cellular buildup of fatty acids to toxic levels. The  major effects of AMPK activation on glucose and lipid metabolism in liver,  skeletal muscle, heart muscle, and adipose tissue are diagrammed in Fig. 22-11.    FIG. 22-11 Major effects of AMP-activated  protein kinase (AMPK) on glucose and lipid  metabolism in liver, muscle, and adipose  tissue. In skeletal muscle, AMPK stimulates  glucose uptake and glucose and fatty acid  oxidation while inhibiting glycogen synthesis. In  heart muscle, AMPK stimulates glycolysis. In  liver, AMPK inhibits lipid biosynthesis and  gluconeogenesis while activating fatty acid  oxidation. In adipose tissue, AMPK inhibits fatty  acid biosynthesis and lipolysis. [After M.C. Towler  and D.G. Hardie, Circulation Res. 100, 328 (2007).]   FIG. 22-1 2 Adiponectin trimers,  hexamers, and multimers. These complexes  are referred to as low molecular weight (LMW),  medium molecular weight (MMW), and high  molecular weight (HMW) forms. [After Kadowaki,  T. and Yamauchi, T., Endocrine Rev. 26, 439  (2005).]   B Adipocytes and Other Tissues Help Regulate Fuel  Metabolism and Appetite   In addition to insulin, glucagon, and epinephrine, a variety of other hormones  help regulate fuel acquisition, use, and storage. Some of these, such as  adiponectin, are secreted exclusively by adipocytes, a cell type whose en¬  docrine activity was not recognized until the early 2000s.   Adiponectin Regulates AMPK Activity. Adiponectin is a 247-residue protein  hormone that helps regulate energy homeostasis and glucose and lipid me¬  tabolism by controlling AMPK activity. Its monomers consist of an N-  terminal collagenlike domain and a C-terminal globular domain. Adiponectin  occurs in the bloodstream in several forms: a low molecular weight (LMW)  trimer formed by the coiling of its collagenlike domains into a triple helix  (Section 6-1C) as well as hexamers (MMW) and multimers (HMW) that  form disulfide-cross-linked bouquets (Fig. 22-12). In addition, globular  adiponectin, formed by the cleavage of the collagenlike domain to release glob¬  ular monomers, occurs in lower concentrations.   The binding of adiponectin to adiponectin receptors, which occur on  the surfaces of both liver and muscle cells, acts to increase the phosphoryla¬  tion and activity of AMPK. This, as we have seen (Section 22-3A), inhibits  gluconeogenesis and stimulates fatty acid oxidation in liver and stimulates glu¬  cose uptake and glucose and fatty acid oxidation in muscle. All of these ef¬  fects act to increase insulin sensitivity, in part because adiponectin and insulin  elicit similar responses in tissues such as liver. Decreased adiponectin is asso¬  ciated with insulin resistance (Section 22-4B). Paradoxically, the blood con¬  centration of adiponectin, which is secreted by adipocytes, decreases with  increased amounts of adipose tissue. This may be because increased adipose  tissue is also associated with increased production of tumor necrosis factor¬  ed (TNFt*), a protein growth factor that decreases both the expression and se¬  cretion of adiponectin from adipose tissue.   Leptin Is a Satiety Hormone. Leptin (Greek: leptos, thin) is a 146-residue  polypeptide that is normally produced by adipocytes (Fig. 22-13). It was dis¬  covered by studying genetically obese mice. Most animals, including humans,  tend to have stable weights; that is, if they are given free access to food, they  eat just enough to maintain their “set-point” weight. However, a strain of mice  that are homozygous for defects in the obese gene (known as oblob mice) are  over twice the weight of normal {OBIOB) mice (Fig. 22-14) and overeat when  given access to unlimited quantities of food. The OB gene encodes leptin.  When leptin is injected into oblob mice, they eat less and lose weight. Leptin  has therefore been considered a “satiety” signal that affects the appetite con¬  trol system of the brain. Leptin also causes higher energy expenditure.   The simple genetic basis for obesity in ob/ob mice does not appear to ap¬  ply to most obese humans. Leptin levels in humans increase with the percent¬  age of body fat, consistent with the synthesis of leptin by adipocytes. Thus,  obesity in humans is apparently the result not of faulty leptin production but  of leptin resistance, perhaps due to a decrease in the level of a leptin receptor  in the brain or the saturation of the receptor that transports leptin across the  blood—brain barrier into the central nervous system.    FIG. 22-13 X-Ray structure of human leptin-EIOO. This mutant form of leptin (Trp  100 Glu) has comparable biological activity to the wild-type protein but crystallizes more  readily. The protein, which is colored in rainbow order from its N-terminus {blue) to its  C-terminus {red), forms a four-helix bundle as do many protein growth factors (e.g., human  growth hormone; Fig. 13-3). Residues 25 to 38 are not visible in the X-ray structure.  [Based on an X-ray structure by Faming Zhang, Eli Lilly & Co., Indianapolis, Indiana. PDBid  1AX8.] O See Interactive Exercise 32.   T A small minority of obese individuals are leptin deficient in a manner sim- Section 3 Metabolic Homeostasis: The Regulation  ilar to ob/ob mice. For example, two grossly obese children who are mem- of Energy Metabolism, Appetite, and Body Weight  bers of the same highly consanguineous (descended from the same ancestors)  family (they are cousins and both sets of parents are cousins) have been shown  to be homozygous for a defective OB gene. The children, at the ages of 8 and  2 years old, respectively, weighed 86 and 29 kg and were noted to have re¬  markably large appetites. Their OB genes have a deletion of a single guanine  nucleotide in codon 133 that causes all succeeding codons to be misread (a  frameshift mutation; Section 27-1 A), which evidently renders the mutant lep¬  tin biologically inactive. In fact, their leptin serum levels were only —10% of  normal. Leptin injections have relieved their symptoms.   A diminished response to leptin leads to high concentrations of neuropep¬  tide Y,    a 36-residue peptide released by the hypothalamus, a part of the brain that  controls many physiological functions. Neuropeptide Y stimulates appetite,  which in turn leads to fat accumulation. Both leptin and insulin, when bound  to their respective receptors in the hypothalamus, inhibit neuropeptide Y  secretion.   Ghrelin and PYY 3 _ 36 Act as Short-Term Regulators of Appetite. It is not   surprising that fuel metabolism, body weight, and appetite are linked. In ad¬  dition to leptin, insulin, and neuropeptide Y, other peptide hormones are  known to be involved in appetite control. For example, ghrelin is an appetite-  stimulating peptide secreted by the empty stomach.   Ghrelin appears to boost levels of neuropeptide Y, and most likely is part of  a short-term appetite-control system, since ghrelin levels fluctuate on the  order of hours, increasing before meals and decreasing immediately afterward.  The gastrointestinal tract secretes an appetite-suppressing hormone called   FIG. 22-14 Normal (OB/OB, left) and  obese (ob/ob, right ) mice. [Courtesy of  Richard D. Palmiter, University of Washington.]    In animals and humans, an infusion of this peptide decreases food intake by  inhibiting neuropeptide Y secretion. In summary, ghrelin stimulates appetite  while leptin, insulin, and PYY 3 _ 36 all suppress it by regulating the secretion  of neuropeptide Y from the hypothalamus.   The nutritional calories contained in food are utilized by an organism either  in the performance of work or in the release of heat. Excess fuel is stored as  glycogen or fat for future use. In well-balanced individuals, the storage of ex¬  cess fuel remains constant over many years. However, when fuel consumed  consistently outpaces energy expenditures, obesity results.   The body has several mechanisms for guarding against obesity. One of  them, discussed above, is appetite control. Another is diet-induced thermo¬  genesis, a form of adaptive thermogenesis (heat production in response to en¬  vironmental stress). We have already discussed adaptive thermogenesis in   • Compare the roles of leptin, ghrelin,  insulin, and PYY in controlling appetite.    response to cold, which is accomplished by the uncoupling of oxidative phos¬  phorylation in brown adipose tissue (Box 18-4). The mechanism of this ther¬  mogenesis involves release of norepinephrine from the brain in response to cold  and its binding to (3-adrenergic receptors on brown adipose tissue, which is  followed by an increase in cAMP that triggers an enzymatic phosphorylation  cascade leading to the activation of hormone-sensitive lipase. The increase in  the concentration of free fatty acids provides fuel for oxidation and leads to  the opening of a proton channel called uncoupling protein 1 (UCP1; alter¬  natively, thermogenin) in the inner mitochondrial membrane. The opening  of UCP1 dissipates the electrochemical gradient generated by electron trans¬  port (Section 18-2) and thereby uncouples electron transport from oxidative  phosphorylation. The consequent continuation of oxidation without ATP syn¬  thesis generates heat.   While it is clear from metabolic measurements that a high energy intake  increases thermogenesis, the cause of this increase in adult humans is unclear  and is under active investigation. Adult humans have little brown adipose tis¬  sue, where UCP1-mediated thermogenesis occurs. However, homologs of the  UCP1 are known (Box 18-4): UCP2 is present in many tissues, including  white adipose tissue, and its synthesis is induced by leptin. UCP3 occurs in  muscle (as well as in brown adipose tissue) and could be significant because  skeletal muscle represents up to 40% of total body weight and has high mi¬  tochondrial capacity. However, it has yet to be demonstrated that UCP3 in  muscle is involved in diet-induced thermogenesis.    • During starvation, the body makes metabolic changes to maintain blood glucose  levels.   • Diabetes may be caused by either insufficient production of insulin (type 1) or an  insensitivity to its presence (type 2).   • Obesity may result from improper regulation of appetite or energy expenditure.    The complex systems that regulate fuel metabolism can malfunction, produc¬  ing acute or chronic diseases of variable severity. Considerable effort has been  directed at elucidating the molecular basis of conditions such as diabetes and  obesity, both of which are essentially disorders of fuel metabolism. In this sec¬  tion, we examine the metabolic changes that occur in starvation, diabetes, and  obesity.   Because humans do not eat continuously, the disposition of dietary fuels and  the mobilization of fuel stores shifts dramatically during the few hours be¬  tween meals. Yet humans can survive fasts of up to a few months by adjust¬  ing their fuel metabolism. Such metabolic flexibility certainly evolved before  modern humans became accustomed to thrice-daily meals.   Absorbed Fuels Are Allocated Immediately. When a meal is digested, nu¬  trients are broken down to small, usually monomeric, units for absorption by  the intestinal mucosa. From there, the products of digestion pass through the  circulation to the rest of the body. Dietary proteins, for example, are broken  down to amino acids for absorption. The small intestine uses amino acids for  fuel, but most travel to the liver via the portal vein. There, they are used for  protein synthesis or, if present in excess, oxidized to produce energy or con¬  verted to glycogen for storage. They may also be converted to glucose or   triacylglycerols for export, depending on conditions. There is no dedicated Section 4 Disturbances in Fuel Metabolism   storage depot for amino acids; whatever the liver does not metabolize circu¬  lates to peripheral tissues to be catabolized or used for protein synthesis.   Dietary carbohydrates, like proteins, are degraded in the intestine, and the  absorbed monomeric products (e.g., glucose derived from dietary starch) are  delivered to the liver via the portal vein. As much as one-third of the dietary  glucose is immediately converted to glycogen in the liver; at least half of the  remainder is converted to glycogen in muscle cells, and the rest is oxidized by  these and other tissues for immediate energy needs. Excess glucose is converted  to triacylglycerol in the liver and exported for storage in adipose tissue. Both  glucose uptake and glycogen and fatty acid biosynthesis are stimulated by in¬  sulin, whose concentration in the blood increases in response to high blood  [glucose].   Dietary fatty acids are packaged as triacylglycerols in chylomicrons  (Section 20-1), which circulate first in the lymph and then in the bloodstream  and therefore are not delivered directly to the liver as are absorbed amino acids  and carbohydrates. Instead, a significant portion of the dietary fatty acids are  taken up by adipose tissue. Lipoprotein lipase first hydrolyzes the triacylglyc¬  erols, and the released fatty acids are absorbed and reesterified in the  adipocytes.   Blood Glucose Remains Nearly Constant. As tissues take up and metabo¬  lize glucose, blood [glucose] drops, thereby causing the pancreatic a cells to  release glucagon. This hormone stimulates glycogen breakdown and the release  of glucose from the liver. It also promotes gluconeogenesis from amino acids  and lactate. The reciprocal effects of insulin and glucagon ., which both respond to  and regulate blood [glucose], ensure that the concentration of glucose available to  extrahepatic tissues remains relatively constant.   However, the body stores less than a day’s supply of carbohydrate  (Table 22-2). After an overnight fast, the combination of increased glucagon  secretion and decreased insulin secretion promotes the mobilization of fatty  acids from adipose tissue (Section 20-5). The diminished insulin also decreases  glucose uptake by muscle tissue. Muscles therefore switch from glucose to fatty  acid metabolism for energy production. This adaptation spares glucose for use  by tissues, such as the brain, that cannot utilize fatty acids.   Source: Cahill, G.E., Jr., New Engl. J. Med. 282, 669 (1970).    FIG. 22-1 5 Liver glycogen depletion  during fasting. The liver glycogen content in  seven subjects was measured using 13 C NMR  over the course of a 64-hour fast. [After  Rothman, D.L, Magnusson, L, Katz, L.D., Shulman,  R.G., and Shulman, G.I., Science 254, 575  (1991).]   Gluconeogenesis Supplies Glucose during Starvation. After a lengthy fast,  the livers store of glycogen becomes depleted (Fig. 22-15). Under these con¬  ditions, the rate of gluconeogenesis increases. Gluconeogenesis supplies ~96%  of the glucose produced by the liver after 40 hours of fasting. The kidney also  is active in gluconeogenesis under these conditions. In animals, glucose can¬  not be synthesized from fatty acids. This is because neither pyruvate nor ox-  aloacetate, the precursors of glucose in gluconeogenesis, can be synthesized in  a net manner from acetyl-CoA. During starvation, glucose must therefore be  synthesized from the glycerol product of triacylglycerol breakdown and, more  importantly, from the amino acids derived from the proteolytic degradation  of proteins, the major source of which is muscle. The breakdown of muscle  cannot continue indefinitely, however, since loss of muscle mass would even¬  tually prevent an animal from moving about in search of food. The organism  must therefore make alternate metabolic arrangements.   Ketone Bodies Become a Major Energy Source during Starvation. After sev¬  eral days of starvation, the liver directs acetyl-CoA, which is derived from fatty  acid (3 oxidation, to the synthesis of ketone bodies (Section 20-3). These fuels  are then released into the blood. The brain gradually adapts to using ketone  bodies as fuel through the synthesis of the appropriate enzymes: After a 3-day  fast, only about one-third of the brains energy requirements are satisfied by ke¬  tone bodies, and after 40 days of starvation, ~70% of its energy needs are so  met. The rate of muscle breakdown during prolonged starvation consequently  decreases to ~25% of its rate after a several-day fast. The survival time of a starv¬  ing individual therefore depends much more on the size of fat reserves than on mus¬  cle mass. Indeed, highly obese individuals can survive a year or more without  eating (and have done so in clinically supervised weight-reduction programs).   Caloric Restriction May Increase Longevity. Caloric restriction is a modified  form of starvation whereby energy intake is reduced 30-40%, while micronu¬  trient (vitamin and mineral) levels are maintained. Rodents kept on such a  diet live up to 50% longer than rodents on normal diets. The lifespans of a  large range of organisms from yeast to primates are similarly extended.  Considerable research effort is being expended to determine the biochemical  basis of these observations.    T in the disease diabetes mellitus (often called just diabetes), which is the  third leading cause of death in the United States after heart disease and can¬  cer, insulin either is not secreted in sufficient amounts or does not efficiently    stimulate its target cells. As a consequence, blood glucose levels become so ele¬  vated that the glucose “spills over” into the urine, providing a convenient diag¬  nostic test for the disease. Yet, despite the high blood glucose levels, cells “starve”  since insulin-stimulated glucose entry into cells is impaired. Triacylglycerol hy¬  drolysis, fatty acid oxidation, gluconeogenesis, and ketone body formation are  accelerated and, in a condition known as ketosis, ketone body levels in the blood  become abnormally high. Since ketone bodies are acids, their high concentra¬  tion puts a strain on the buffering capacity of the blood and on the kidney,  which controls blood pH by excreting the excess H into the urine. This H  excretion is accompanied by NH4 , Na + , K + , P z , and H 2 0 excretion, causing  severe dehydration (excessive thirst is a classic symptom of diabetes) and a de¬  crease in blood volume—ultimately life-threatening situations.   1. Insulin-dependent or juvenile-onset diabetes mellitus, which most  often strikes suddenly in childhood.   2. Non-insulin-dependent or maturity-onset diabetes mellitus, which  usually develops gradually after the age of 40, but is becoming more  common in younger adults.   The accumulation of glucose in the lens leads to  swelling and precipitation of lens proteins. The  resulting opacification causes blurred vision and  ultimately complete loss of sight [© Sue  Ford/Photo Researchers.]    Insulin-Dependent Diabetes Is Caused by a Deficiency of Pancreatic p  Cells. In insulin-dependent (type 1) diabetes mellitus, insulin is absent or  nearly so because the pancreas lacks or has defective P cells. This condition  usually results from an autoimmune response that selectively destroys pancre¬  atic p cells. Individuals with insulin-dependent diabetes, as Frederick Banting  and Charles Best first demonstrated in 1921 (Box 22-2), require daily insulin  injections to survive and must follow carefully balanced diet and exercise regi¬  mens. Their lifespans are, nevertheless, reduced by up to one-third as a result of  degenerative complications such as kidney malfunction, nerve impairment, and  cardiovascular disease that apparently arise from the imprecise metabolic control  provided by periodic insulin injections. The hyperglycemia (high blood [glu¬  cose]) of diabetes mellitus also leads to blindness through retinal degeneration  and the glucosylation of lens proteins, which causes cataracts (Fig. 22-16).   The usually rapid onset of the symptoms of insulin-dependent diabetes  had suggested that the autoimmune attack on the pancreatic P cells is of short  duration. Typically, however, the disease develops over several years as the im¬  mune system slowly destroys the (3 cells. Only when >80% of the cells have  been eliminated do the classic symptoms of diabetes suddenly emerge.  Consequently, one of the most successful treatments for insulin-dependent di¬  abetes is a (3-cell transplant, a procedure that became possible with the devel¬  opment of relatively benign immunosuppressive drugs.    Non-Insulin-Dependent Diabetes May Be Caused by a  Deficiency of Insulin Receptors or Insulin Signal Transduction.   Non-insulin-dependent (type 2) diabetes mellitus, which accounts  for over 90% of the diagnosed cases of diabetes and affects 18%  of the population over 65 years of age, usually occurs in obese in¬  dividuals with a genetic predisposition for the condition. These in¬  dividuals have normal or even greatly elevated insulin levels, but  their cells are not responsive to insulin and are therefore said to be  insulin resistant. As a result, blood glucose concentrations are much  higher than normal, particularly after a meal (Fig. 22-17).    FIG. 22-1 7 Twenty-four-hour plasma glucose profiles in normal and  non-insulin-dependent diabetic subjects. The basal level of glucose and  the peaks following meals are higher in the diabetic individuals. [After Bell, G.I.,  Pilkis, S.J., Weber, I.T., and Polonsky, K.S., Annu. Rev. Physiol. 58, 178 (1996).]   One of the most celebrated clinical research  stories is the discovery and therapeutic use of  insulin, the hormone missing in type 1 diabetes.  Since the end of the nineteenth century, clinical  scientists understood that there was a  connection between diabetes and secretions  from the pancreas, specifically, from the clumps  of cells known as the islets of Langerhans.  Several unsuccessful attempts were made to  extract the secretions. In 1920, Frederick  Banting, reading an account of such work,  wondered whether an active extract could be  purified if the pancreatic ducts, which collect  potentially destructive digestive enzymes, were  first ligated (closed off).   Banting, a young surgeon whose medical practice had not yet  gotten off the ground, was working as a demonstrator at the  University of Western Ontario. He took his idea to John J.R.  Macleod, the head of the Department of Physiology at the  University of Toronto. A skeptical Macleod gave Banting the use  of laboratory space for the summer of 1921, access to  experimental dogs, and the help of assistant Charles Best.   Banting, despite his medical training and experience with the  Canadian Army Medical Service, was not much of an  experimentalist. Best, however, had just completed a degree in  physiology and biochemistry and had already worked as an  assistant to Macleod. Best quickly learned surgical techniques  from Banting, as Banting picked up analytical techniques, such as  measuring sugar levels in dogs’ blood and urine, from Best.   Banting’s and Best’s experimental protocol required  pancreatectomized dogs, but the surgery was difficult, and many  dogs died of infection (it was rumored that strays from the streets of  Toronto were gathered as replacements). Successfully  pancreatectomized dogs, who developed the fatal symptoms of  diabetes, were injected with a pancreatic extract that Banting and  Best named insulin, after the Latin word insula (island). Finally, at the  end of July, Banting and Best achieved their desired result, when the  insulin injection dramatically reduced the dog’s blood sugar level. The  experiment was repeated, with similar results, in other dogs.   Although Banting’s temporary position was to have been  terminated at the end of the summer, his results were promising    enough to secure him additional funding, the supervision of a  more enthusiastic Macleod, and the continued assistance of Best,  who had originally intended to work with Banting for only 2  months. By fall, Banting’s and Best’s success at prolonging the  lives of diabetic dogs was putting enormous pressure on their  ability to produce insulin in large amounts and of consistent purity.  Assistance was offered by James Collip, a biochemist on  sabbatical leave from the University of Alberta. Collip’s insulin  preparations, which were made without any clue as to the  proteinaceous nature of the hormone, proved good enough for  experimental use in humans. At that time, a diagnosis of diabetes  was a virtual death sentence, with the only treatment being a  severely limited diet that probably extended life for only a few  months (many patients died of malnutrition).   In January 1923, Banting injected Collip’s extract into Leonard  Thompson, a 14-year-old diabetic who was near death. Thompson’s  blood sugar level immediately returned to normal, and his strength  increased. Such a dramatic clinical outcome did not go unnoticed,  and within months, Banting opened a clinic to begin treating  diabetics desperate for a treatment that restored their hopes for a  near-normal life. He enlisted the help of the Connaught Antitoxin  Laboratories at the University of Toronto and the Eli Lilly  Pharmaceutical Company to produce insulin on a scale suitable  for clinical testing. Best, just 23, was put in charge of insulin  production for all of Canada. It eventually became clear that  Banting’s original assumption-that the presence of digestive  enzymes compromised the purification of insulin—was mistaken,  and insulin could be successfully isolated from an intact pancreas.   Amid some controversy, the 1923 Nobel Prize for Physiology  or Medicine was awarded to Banting and Macleod. Banting,  annoyed by the omission of Best, split his share of the prize with  his assistant. Macleod, who had not offered much support during  the initial research, split his share with Collip. Banting  subsequently held a variety of administrative posts and pursued  research in silicosis, but he never accomplished much. He was  killed in a plane crash en route to Britain for a wartime mission in  1941. Best, who replaced Macleod as head of physiology at the  age of 29, continued his research into the biological action of  insulin. He also supervised efforts to purify the anticoagulant  glycosaminoglycan heparin and to produce dried human serum for  medical use by the military.   Banting, F.G. and Best, C.H., The internal secretion of the pancreas, J. Lab. Clin. Med.  7, 251-266 (1922).    The hyperglycemia that accompanies insulin resistance induces the pan¬  creatic (3 cells to increase their production of insulin. Yet the high basal level  of insulin secretion diminishes the ability of the (3 cells to respond to further  increases in blood glucose. Consequently, the hyperglycemia and its attendant  complications tend to worsen over time.   A small percentage of cases of type 2 diabetes result from mutations in  the insulin receptor that affect its insulin-binding ability or tyrosine kinase ac¬  tivity. However, a clear genetic cause has not been identified in the vast ma¬  jority of cases. It is therefore likely that many factors play a role in the  development of the disease. For example, the increased insulin production  resulting from overeating may eventually suppress the synthesis of insulin    receptors. This hypothesis accounts for the observation that diet alone often  decreases the severity of the disease.   Another hypothesis, put forward by Gerald Shulman, is that the elevated  concentration of free fatty acids in the blood caused by obesity decreases in¬  sulin signal transduction. This high concentration ultimately results in an ac¬  tivation of a PKC isoform that phosphorylates insulin receptor substrates  (IRSs) on Ser and Thr, thereby inhibiting the Tyr phosphorylation that acti¬  vates them. The failure to activate IRSs decreases the cell’s response to insulin  (Fig. 13-31).   Other treatments for non-insulin-dependent diabetes are drugs such as  metformin and the thiazolidinediones (TZDs) (at right), which decrease in¬  sulin resistance by either suppressing glucose release by the liver (metformin)  or promoting insulin-stimulated glucose disposal in muscle (TZDs). The drugs  act by increasing AMPK activity but by different mechanisms. TZDs cause a  large increase in the AMP-to-ATP ratio in muscle cells, with the expected con¬  comitant increase in AMPK phosphorylation and activity. Metformin, how¬  ever, increases AMPK phosphorylation and activity without changing the  AMP-to-ATP ratio. In both cases, the increase in AMPK activity decreases glu-  coneogenesis in liver and increases glucose utilization in muscle (Fig. 22-11).  In addition, the TZDs decrease insulin resistance by binding to and activat¬  ing a transcription factor known as a peroxisome proliferator-activated  receptor -7 (PPAR- 7 ), primarily in adipose tissue. Among other things, PPAR-  7 activation induces the synthesis of adiponectin (Section 22-3B), which leads  to an increase in AMPK activity. In adipose tissue, AMPK action leads to a  decrease in lipolysis and fatty acid export, decreasing the concentration of free  fatty acids in the blood and therefore decreasing insulin resistance (see above).  The TZDs are not without side effects, however. For example, rosiglitazone  (Avandia)    appears to increase the risk of heart attack and stroke by as much as 43%.  Consequently, it has been banned in Europe, and its use in the United States  is limited to patients for whom no other drug is effective.   Rodent adipocytes secrete a 108-residue polypeptide hormone called  resistin. The hormone is named for its ability to block the action of insulin  on adipocytes. In fact, resistin production is decreased by TZDs, a feature that  led to its discovery. Overproduction of resistin was proposed to contribute to  the development of non-insulin-dependent diabetes. However, an interesting  divergence between rodents and humans is that in humans, resistin is pro¬  duced by macrophages and its role in human metabolic regulation is therefore  unclear.   The human body regulates glycogen and protein levels within relatively nar¬  row limits, but fat reserves, which are much larger, can become enormous.  The accumulation of fatty acids as triacylglycerols in adipose tissue is largely  a result of excess fat or carbohydrate intake compared to energy expenditure.  Fat synthesis from carbohydrates occurs when the carbohydrate intake is high  enough that glycogen stores, to which excess carbohydrate is normally directed,  approach their maximum capacity.   Chapter 22 Mammalian Fuel Metabolism: Integration A chronic imbalance between fat and carbohydrate consumption and uti-   and Regulation lization increases the mass of adipose tissue through an increase in the num¬   ber of adipocytes or their size (once formed, adipocytes are not lost, although  their size may increase or decrease). The increase in adipose tissue mass in¬  creases the pool of fatty acids that can be mobilized to generate metabolic en¬  ergy. Eventually, a steady state is achieved in which the mass of adipose tissue  no longer increases and fat storage is balanced by fat mobilization. This phe¬  nomenon explains in part the high incidence of obesity in affluent societies,  where fat- and carbohydrate-rich foods are plentiful and physical activity is  not a requirement for survival. Considerable evidence suggests that behavior  (e.g., eating habits and levels of physical activity) influences an individuals  body composition. Yet, as we have seen (Section 22-3), some cases of obesity  are also the result of innate disturbances in an individuals capacity to metab¬  olize fuels.   The hormonal mechanisms that contribute to obesity and its attendant  health problems, such as diabetes, may have conferred a selective advantage in  premodern times. For example, the ability to gain weight easily would have  protected against famine. One theory proposes that as humans evolved, the  regulation of fuel metabolism was geared toward overeating large quantities of  food when available, thereby obtaining essential nutrients while storing excess  nonessential energy as fat for use during famine. In modern cultures in which  famines are rare and nutritious food is readily available, human genetic her¬  itage has apparently contributed to an epidemic of obesity: An estimated 30%  of U.S. adults are obese and another 35% are overweight.   • Describe the metabolic changes that  occur during early and late starvation.   • How does insulin resistance or the lack of  insulin contribute to the typical symptoms  of diabetes?    T Obesity Is a Contributing Factor in Metabolic Syndrome. Metabolic syn¬  drome is a disturbance in metabolism characterized by insulin resistance,  inflammation, and a predisposition to several disorders including type 2 dia¬  betes, hypertension, and atherosclerosis. Those disorders are accompanied by  an increase in coronary heart disease. Obesity, physical inactivity, and possi¬  bly genetic determinants have been implicated in its occurrence, which affects  up to 25% of the population in the United States. Metabolic syndrome is par¬  ticularly prevalent among members of ethnic groups that have adopted mod¬  ern diets and lifestyles in recent generations.   Individuals with metabolic syndrome tend to have a relatively high pro¬  portion of visceral (abdominal) fat. This type of fat exhibits a different hor¬  mone profile than subcutaneous fat. For example, visceral fat produces less  leptin and adiponectin (hormones that increase insulin sensitivity). Visceral fat  also produces more TNFa, which is a powerful mediator of inflammation and  a normal part of the body’s immune defenses. Chronic inflammation triggered  by excess visceral fat may be responsible for some of the symptoms, such as  atherosclerosis, that characterize metabolic syndrome. The TNFa signaling  pathway in cells may lead to phosphorylation of IRS-1, a modification that  prevents its activation by the insulin receptor kinase. This would explain the  insulin resistance of metabolic syndrome. Exercise, calorie/weight reduction,  adiponectin, leptin, metformin, and TZDs have all been successfully used to  treat metabolic syndrome. All of these increase AMPK activity, making AMPK  itself a promising target for drug development.   • The pathways for the synthesis and degradation of the major  metabolic fuels (glucose, fatty acids, and amino acids) converge  on acetyl-CoA and pyruvate. In mammals, flux through the path¬  ways is tissue specific.   • The brain uses glucose as its primary metabolic fuel. Muscle can  oxidize a variety of fuels but depends on anaerobic glycolysis for  maximum exertion. Adipose tissue stores excess fatty acids as tri-  acylglycerols and mobilizes them as needed.   • The liver maintains the concentrations of circulating fuels. The  action of glucokinase allows liver to take up excess glucose, which  can then be directed to several metabolic fates. The liver also con¬  verts fatty acids to ketone bodies and metabolizes amino acids de¬  rived from the diet or from protein breakdown. Both the liver and  kidney carry out gluconeogenesis.   • The Cori cycle and the glucose-alanine cycle are multiorgan path¬  ways through which the liver and muscle exchange metabolic in¬  termediates.   • Hormones such as insulin, glucagon, and epinephrine transmit  regulatory signals to target tissues by binding to receptors that  transduce the signal to responses in the interior of the cell. Insulin  promotes fuel storage, and glucagon and epinephrine mobilize  stored fuels.   • AMP-dependent protein kinase (AMPK), the cell’s fuel gauge,  senses the cell’s need for ATP and activates metabolic breakdown  pathways while inhibiting biosynthetic pathways.   • Adiponectin, an adipocyte hormone that increases insulin sensi¬  tivity, acts by activating AMPK.   • Appetite is controlled in the hypothalamus by the hormones lep-  tin, insulin, ghrelin, and PYY.   • During starvation, when dietary fuels are unavailable, the liver re¬  leases glucose first by glycogen breakdown and then by gluconeo¬  genesis from amino acid precursors. Eventually, ketone bodies  supplied by fatty acid breakdown meet most of the body’s energy  needs.   • Diabetes mellitus causes hyperglycemia and other physiological  difficulties resulting from destruction of insulin-producing pancre¬  atic |3 cells or from insulin resistance (from loss of receptors or  their insensitivity to insulin).   • Obesity, an imbalance between food intake and energy expendi¬  ture, may result from abnormal regulation by peptide hormones  produced in different tissues.   • Metabolic syndrome is caused by obesity, physical inactivity, and  possibly genetic determinants.   1. Why does oxidative metabolism, which generates ATP, cease  when a cell’s ATP supply is exhausted?   2. Individuals with a deficiency of medium chain acyl-CoA dehy¬  drogenase (MCAD) tend to develop nonketotic hypoglycemia.  Explain why this defect makes them unable to undertake (a) ke-  togenesis and (b) gluconeogenesis.   3. Many cancer cells generate ATP primarily by glycolysis, an anaer¬  obic pathway, even when 0 2 is available (this phenomenon was  first noted by Otto Warburg in 1924). Explain how a high flux  of carbon through the glycolytic pathway could help support a  high rate of growth of the cancer cells.   4. Growth factor stimulation of cells leads to inhibition of pyruvate  kinase. As a result, glycolytic intermediates are redirected to the  pentose phosphate pathway. Explain why this would help pro¬  mote cell growth and division.   3. Adaptation to high altitude includes increases in GLUTl and  phosphofructokinase in muscle. Explain why this would be ad¬  vantageous.    6 . The heart cannot convert lactate back to glucose, as the liver  does, but instead uses it as a fuel. What is the ATP yield from  the complete catabolism of one mole of lactate?   7. The passive glucose transporter, called GLUTl (Fig. 10-13), is  present in the membranes of many cells, but not in the liver.  Instead, liver cells express the GLUT 2 transporter, which exhibits  different transport kinetics. Given what you know about the role  of the liver in buffering blood glucose, compare the K M values of  GLUTl and GLUT 2 .   8 . Use the results of Problem 7 to explain why a deficiency of  GLUT 2 produces symptoms resembling those of Type 1 glyco¬  gen storage disease (Box 16-2).   9. The small intestine catabolizes amino acids, producing significant  amounts of NH 4 . Describe the metabolic fate of the NH 4 .   10. (a) Identify the two reactions that allow the kidney to produce  NH 4 . (b) Which gluconeogenic precursor is thereby generated?  Describe the pathway by which it can be converted to glucose.   11. Explain why insulin is required for adipocytes to synthesize tri-  acylglycerols from fatty acids.   12. Predict the effect of an overdose of insulin on brain function in  a normal person.   13. Pancreatic |3 cells express a receptor for fatty acids. Fatty acid  binding to the protein appears to stimulate insulin secretion.  Does this phenomenon make metabolic sense?   14. Fatty acids appear to stimulate insulin secretion to a much greater  extent when glucose is also present. Why is this significant?   13. Would you expect insulin to increase or decrease the activity of  the enzyme ATP-citrate lyase?   16. Stimulation of a certain G s protein-coupled receptor activates  protein kinase A (PKA). Predict the effect of PKA activation  on the following substrates of PKA: (a) acetyl-CoA carboxylase,  (b) glycogen synthase, (c) hormone-sensitive lipase, and (d) phos-  phorylase kinase.   17. Why can adipose tissue be considered to be an endocrine organ?   18. If the circulatory system of an obiob mouse is surgically joined  to that of a normal mouse, what will be the effect on the ap¬  petite and weight of the ob/ob mouse?   19. In experiments to test the appetite-suppressing effects of PYY 3 _ 36 ,  why must the hormone be administered intravenously rather than  orally?   20. Extremely obese recipients of gastric bypass surgery, which closes  off most of the stomach and reroutes a portion of the small in¬  testine, frequently become more insulin sensitive immediately af¬  ter the surgery, even before they have lost any weight. Explain.   21. Experienced runners know that it is poor practice to ingest large  amounts of glucose immediately before running a marathon.  What is the metabolic basis for this apparent paradox?   22. After several days of starvation, the capacity of the liver to me¬  tabolize acetyl-CoA via the citric acid cycle is greatly diminished.  Explain.   23. Explain why type 1 diabetics require insulin injections, whereas in¬  sulin injections are effective in only a portion of type 2 diabetics.   24. High concentrations of free fatty acids in the blood are known to  cause insulin resistance in muscle, but only after 3 hours. This sug¬  gests that a metabolite of the fatty acids may be responsible for this  phenomenon. It is also known that an isoform of protein kinase  C is activated during the process and that high concentrations of  free fatty acids result in intramuscular accumulation of triacylglyc-  erols. With this information, review the mechanism of activation    of PKC and the pathway of triacylglycerol biosynthesis and suggest  a metabolite that may be responsible for PKC activation.   25. AMPK activates phosphofructokinase-2. Explain how this would  contribute to the antidiabetic effect of stimulating AMPK activ¬  ity.   26. Discuss, in molecular terms, how physical inactivity might lead  to insulin resistance.   Case 25 Glycogen Storage Diseases  Focus concept: Disturbances in glycogen utilization result if an en¬  zyme involved in glycogen synthesis or degradation is missing.   • The link between glycogen metabolism and fat metabolism  Case 28 The Bacterium Helicobacter pylori and  Peptic Ulcers   Focus concept: The entire genome of Helicobacter pylori has been  sequenced. This allows biochemists to examine the organism’s pro¬  teins in detail. In this case, mechanisms employed by Helicobacter  pylori that permit it to survive in the acidic environment of the stom¬  ach are examined.   Focus concept: The characteristics of phenylalanine hydroxylase,  the enzyme missing in persons afflicted with the genetic disorder  phenylketonuria (PKU), are examined.   • Integration of amino acid metabolic pathways with carbohy¬  drate metabolic pathways    Anecdotal evidence suggests  that sleep and obesity are related: less sleep tends to promote  weight gain, and vice versa. What are the hormonal links between  the basic drives for food and rest? Is there evidence that increas¬  ing the time spent sleeping can lead to weight loss?    Bollen, M., Keppens, S., and Stalmans, W., Specific features of glycogen  metabolism in the liver, Biochem. J. 336, 19-31 (1998).   Coll, A.P., Farooqi, I.S., and O’Rahilly, S., The hormonal control of food  intake, Cell 129, 251-262 (2007).   Huang, H. and Czech, M.P., The GLUT4 glucose transporter, Cell Me¬  tabolism 5 , 237-252 (2007).   Kadowaki, T. and Yamauchi, T., Adiponectin and adiponectin receptors,  Endocrine Rev. 26, 439-451 (2005).   Kahn, S.E., Hull, R.L., and Utzschneider, K.M., Mechanisms linking  obesity to insulin resistance and type 2 diabetes, Nature 444,  840-856 (2006).    Nakar, V.A., et al., AMPK and PPARy are exercise mimics, Cell 134,  405-415 (2008).   Obesity, Science 299, 845-860 (2003). [A series of informative reports  on the origins and treatments of obesity.]   Spiegelman, B.M. and Flier, J.S., Obesity and the regulation of energy  balance, Cell 104, 531-543 (2001).   Towler, M.C. and Hardie, D.G., AMP-Activated protein kinase in meta¬  bolic control and insulin signaling, Circulation Res. 100, 328-341  (2007).   A Purine Synthesis Yields Inosine Monophosphate  B IMP Is Converted to Adenine and Guanine  Ribonucleotides   C Purine Nucleotide Biosynthesis Is Regulated  at Several Steps  D Purines Can Be Salvaged   A UMP Is Synthesized in Six Steps  B UMP Is Converted to UTP and CTP  C Pyrimidine Nucleotide Biosynthesis Is Regulated  at ATCase or Carbamoyl Phosphate Synthetase II   A Ribonucleotide Reductase Converts  Ribonucleotides to Deoxyribonucleotides  B dUMP Is Methylated to Form Thymine   A Purine Catabolism Yields Uric Acid  B Some Animals Degrade Uric Acid  C Pyrimidines Are Broken Down to Malonyl-CoA  and Methylmalonyl-CoA    When mammalian cells are damaged, they release urate, a product of nucleotide  metabolism. White blood cells, such as the one shown here, respond to the urate by  activating other immune defenses and by increasing their phagocytic activity in order  to engulf and eliminate the damaged cell. [David Scharf/Photo Researchers, Inc.]    Nucleotides are phosphate esters of a pentose (ribose or deoxyribose) in which  a purine or pyrimidine base is linked to Cl' of the sugar (Section 3-1).  Nucleoside triphosphates are the monomeric units that act as precursors of  nucleic acids; nucleotides also perform a wide range of other biochemical func¬  tions. For example, we have seen how the cleavage of “high-energy” com¬  pounds such as ATP provides the free energy that makes various reactions  thermodynamically favorable. We have also seen that nucleotides are compo¬  nents of some of the central cofactors of metabolism, including FAD, NAD + ,  and coenzyme A. The importance of nucleotides in cellular metabolism is in¬  dicated by the observation that nearly all cells can synthesize them both  de novo (anew) and from the degradation products of nucleic acids. However,  unlike carbohydrates, amino acids, and fatty acids, nucleotides do not provide  a significant source of metabolic energy.   In this chapter, we consider the nature of the nucleotide biosynthetic path¬  ways. In doing so, we will examine how they are regulated and the conse¬  quences of their blockade, both by genetic defects and through the  administration of chemotherapeutic agents. We then discuss how nucleotides  are degraded. In following the general chemical themes of nucleotide metab¬  olism, we will break our discussion into sections on purines, pyrimidines, and  deoxynucleotides (including thymidylate). The structures and nomenclature of  the major purines and pyrimidines are given in Table 3-1.    • IMP is synthesized through the assembly of a purine base on ribose-5-phosphate.   • Kinases convert IMP-derived AMP and GMP to ATP and GTP   • Purine nucleotide synthesis is regulated by feedback inhibition and feedforward  activation.    In 1948, John Buchanan obtained the first clues to the de novo synthesis of  purine nucleotides by feeding a variety of isotopically labeled compounds to  pigeons and chemically determining the positions of the labeled atoms in their  excreted uric acid (a purine; at left).   The results of his studies demonstrated that N1 of purines arises from the  amino group of aspartate; C2 and C8 originate from formate; N3 and N9 are  contributed by the amide group of glutamine; C4, C5, and N7 are derived  from glycine (strongly suggesting that this molecule is wholly incorporated  into the purine ring); and C6 comes from HC0 3 .    The actual pathway by which the precursors are incorporated into the  purine ring was elucidated in subsequent investigations performed largely by  Buchanan and G. Robert Greenberg. The initially synthesized purine deriva¬  tive is inosine monophosphate (IMP; at left), the nucleotide of the base  hypoxanthine. IMP is the precursor of both AMP and GMP Thus, contrary  to expectation, purines are initially formed as ribonucleotides rather than as free  bases. Additional studies have demonstrated that such widely divergent organ¬  isms as E. coli, yeast, pigeons, and humans have virtually identical pathways  for the biosynthesis of purine nucleotides, thereby further demonstrating the  biochemical unity of life.   IMP is synthesized in a pathway composed of 11 reactions (Fig. 23-1):   1. Activation of ribose-5-phosphate. The starting material for purine  biosynthesis is a-D-ribose-5-phosphate, a product of the pentose phos¬  phate pathway (Section 15-6). In the first step of purine biosynthesis,  ribose phosphate pyrophosphokinase activates the ribose by reacting  it with ATP to form 5-phosphoribosyl-a-pyrophosphate (PRPP).  This compound is also a precursor in the biosynthesis of pyrimidine  nucleotides (Section 23-2A) and the amino acids histidine and trypto¬  phan (Section 21-5B). As is expected for an enzyme at such an impor¬  tant biosynthetic crossroads, the activity of ribose phosphate  pyrophosphokinase is precisely regulated.   2. Acquisition of purine atom N9. In the first reaction unique to purine  biosynthesis, amidophosphoribosyl transferase catalyzes the displace¬  ment of PRPP s pyrophosphate group by glutamines amide nitrogen.  The reaction occurs with inversion of the a configuration at Cl of  PRPP, thereby forming (5-5-phosphoribosylamine and establishing  the anomeric form of the future nucleotide. The reaction, which is  driven to completion by the subsequent hydrolysis of the released PP Z ,  is the pathways flux-controlling step.   of IMP. Here the purine residue is built up on a ribose ring in 11  enzyme-catalyzed reactions. The X-ray structures for all the enzymes  are shown to the outside of the corresponding reaction arrow. The  peptide chains of monomeric enzymes are colored in rainbow order  from N-terminus (blue) to C-terminus (red). The oligomeric enzymes,  all of which consist of identical polypeptide chains, are viewed along a  rotation axis with their various chains differently colored. Bound    ligands are shown in space-filling form with C green, N blue, 0 red,  and P orange. [PDBids: enzyme 1, 1DKU; enzyme 2, 1AOO; enzyme  3, 1GS0; enzyme 4, 1CDE; enzyme 5, 1VK3; enzyme 6, 1CLI; enzyme  7, 1D7A (PurE) and 1 B6S (PurK); enzyme 8, 1A48; enzyme 9, 1C3U;  enzymes 10 and 11, 1G8M.] Q See the Animated Figures.   FIG. 23-2 X-Ray structure of E. coli GAR  transformylase in complex with GAR and  5dTHF. The monomeric protein is colored in  rainbow order from its N-terminus (blue) to its  C-terminus (red). GAR (upper middle ) and  5dTHF (lower left) are drawn in stick form with  GAR C green, 5dTHF C magenta, N blue, 0 red,  and P orange. Note the close (3.3-A) approach  of the GAR amino group to N10 of 5dTHF.  [Based on an X-ray structure by Robert Almassy,  Agouron Pharmaceuticals, San Diego, California.  PDBid 1CDE.1    3. Acquisition of purine atoms C4, C5, and N7. Glycine s carboxyl  group forms an amide with the amino group of phosphoribosylamine,  yielding glycinamide ribotide (GAR). This reaction is reversible, de¬  spite its concomitant hydrolysis of ATP to ADP + P z . It is the only  step of the purine biosynthetic pathway in which more than one purine  ring atom is acquired.   formyl donor in the reaction is 7V 10 -formyltetrahydrofolate (N 10 -  formyl-THF), a coenzyme that transfers C x units (THF cofac¬  tors are described in Section 21-4D). The X-ray structure of the  enzyme catalyzing the reaction, GAR transformylase, in com¬  plex with GAR and the THF analog 5-deazatetrahydrofolate  (5dTHF) was determined by Robert Almassy (Fig. 23-2). Note  the proximity of the GAR amino group to N10 of 5dTHF. This  supports enzymatic studies suggesting that the GAR transformylase  reaction proceeds via the nucleophilic attack of the GAR amine group  on the formyl carbon of A^ 10 -formyl-THF to yield a tetrahedral inter¬  mediate.   5. Acquisition of purine atom N3. The amide amino group of a second  glutamine is transferred to the growing purine ring to form formyl-  glycinamidine ribotide (FGAM). This reaction is driven by the cou¬  pled hydrolysis of ATP to ADP + P z .   6. Formation of the purine imidazole ring. The purine imidazole ring  is closed in an ATP-requiring intramolecular condensation that yields  5-aminoimidazole ribotide (AIR). The aromatization of the imid¬  azole ring is facilitated by the tautomeric shift of the reactant from its  imine to its enamine form.   7. Acquisition of C6. Purine C6 is introduced as HC0 3 (C0 2 ) in a re¬  action catalyzed by AIR carboxylase that yields carboxyaminoim-  idazole ribotide (CAIR). In yeast, plants, and most prokaryotes  (including E. coli), AIR carboxylase consists of two proteins called  PurE and PurK. Although PurE alone can catalyze the carboxylation  reaction, its K M for HC0 3 is —110 mM, so the reaction would re¬  quire an unphysiologically high HC0 3 concentration (—100 mM) to  proceed. PurK decreases the HC0 3 concentration required for the  PurE reaction by > 1000-fold but at the expense of ATP hydrolysis.   8. Acquisition of Nl. Purine atom N1 is contributed by aspartate in an  amide-forming condensation reaction yielding 5-aminoimidazole-4-  (Af-succinylocarboxamide) ribotide (SACAIR). This reaction, which  is driven by the hydrolysis of ATP, chemically resembles Reaction 3.    9. Elimination of fumarate. SACAIR is cleaved with the release of fu-  marate, yielding 5-aminoimidazole-4-carboxamide ribotide (AICAR).  Reactions 8 and 9 chemically resemble the reactions in the urea cycle in  which citrulline is aminated to form arginine (Section 21-3A). In both  pathways, aspartates amino group is transferred to an acceptor through  an ATP-driven coupling reaction followed by the elimination of the  aspartate carbon skeleton as fumarate.   10. Acquisition of C2. The final purine ring atom is acquired through  formylation by 7V 10 -formyl-THF, yielding 5-formaminoimidazole-4-  carboxamide ribotide (FAICAR). This reaction and Reaction 4 of  purine biosynthesis are inhibited indirectly by sulfonamides, structural  analogs of the y?-aminobenzoic acid constituent of THF (Section 21-4D).   11. Cyclization to form IMP. The final reaction in the purine biosynthetic  pathway, ring closure to form IMP, occurs through the elimination of  water. In contrast to Reaction 6, the cyclization that forms the imid¬  azole ring, this reaction does not require ATP hydrolysis.   In animals, Reactions 10 and 11 are catalyzed by a bifunctional enzyme, as are Section 1 Synthesis of Purine Ribonucleotides   Reactions 7 and 8. Reactions 3, 4, and 6 also take place on a single protein.   The intermediate products of these multifunctional enzymes are not readily released to  the medium but are channeled to the succeeding enzymatic activities  of the pathway. As in the reactions catalyzed by the pyruvate dehydrogenase com¬  plex (Section 17-2), fatty acid synthase (Section 20-4C), bacterial glutamate syn¬  thase (Section 21-7), and tryptophan synthase (Section 21-5B), channeling in  the nucleotide synthetic pathways increases the overall rate of these multistep  processes and protects intermediates from degradation by other cellular enzymes.   IMP does not accumulate in the cell but is rapidly converted to AMP and GMP.   AMP, which differs from IMP only in the replacement of its 6-keto group by  an amino group, is synthesized in a two-reaction pathway (Fig. 23-3, left). In  the first reaction, aspartates amino group is linked to IMP in a reaction pow¬  ered by the hydrolysis of GTP to GDP + P z to yield adenylosuccinate. In the  second reaction, adenylosuccinate lyase eliminates fumarate from adenylosuc¬  cinate to form AMP. The same enzyme catalyzes Reaction 9 of the IMP path¬  way (Fig. 23-1). Both reactions add a nitrogen with the elimination of fumarate.   GMP is also synthesized from IMP in a two-reaction pathway (Fig. 23-3,  right). In the first reaction, IMP is dehydrogenated via the reduction of NAD +  to form xanthosine monophosphate (XMP; the ribonucleotide of the base  xanthine). XMP is then converted to GMP by the transfer of the glutamine    FIG. 23-3 Conversion of IMP to AMP or GMP in separate  two-reaction pathways. The X-ray structures of the homo-oligomeric  enzymes catalyzing the reactions are shown as is described in the    legend for Fig. 23-1. [PDBids: adenylosuccinate synthetase, 1GIM;  adenylosuccinate lyase, 1C3U; IMP dehydrogenase, 1JR1; GMP  synthetase, 1GPM.]    amide nitrogen in a reaction driven by the hydrolysis of ATP to AMP + PP Z  (and subsequently to 2 P z ). In B and T lymphocytes, which mediate the im¬  mune response, IMP dehydrogenase activity is high in order to supply the  guanosine the cells need for proliferation. The fungal compound mycophe¬  nolic acid ( at left ) inhibits the enzyme and is used as an immunosuppressant  following kidney transplants.   Nucleoside Diphosphates and Triphosphates Are Synthesized by the  Phosphorylation of Nucleoside Monophosphates. In order to participate in  nucleic acid synthesis, nucleoside monophosphates must first be converted to the  corresponding nucleoside triphosphates. First, nucleoside diphosphates are syn¬  thesized from the corresponding nucleoside monophosphates by base-specific  nucleoside monophosphate kinases. For example, adenylate kinase (Section  14-2C) catalyzes the phosphorylation of AMP to ADP:    These nucleoside monophosphate kinases do not discriminate between ribose  and deoxyribose in the substrate.   Although the reaction is written with ATP as the phosphoryl donor, this en¬  zyme exhibits no preference for the bases of its substrates or for ribose over  deoxyribose. Furthermore, the nucleoside diphosphate kinase reaction, as  might be expected from the nearly identical structures of its substrates and  products, normally operates close to equilibrium (AG ~ 0). ADP is, of course,  also converted to ATP by a variety of energy-releasing reactions such as those  of glycolysis and oxidative phosphorylation. Indeed, it is those reactions that  ultimately drive the foregoing kinase reactions.   The pathways synthesizing IMP, ATP, and GTP are individually regulated in  most cells so as to control the total amounts of purine nucleotides available  for nucleic acid synthesis, as well as the relative amounts of ATP and GTP.  This control network is diagrammed in Fig. 23-4.   The IMP pathway is regulated at its first two reactions: those catalyzing the  synthesis of PRPP and 5-phosphoribosylamine. Ribose phosphate pyrophospho-  kinase, the enzyme catalyzing Reaction 1 of the IMP pathway (Fig. 23-1), is in¬  hibited by both ADP and GDP. Amidophosphoribosyl transferase, the enzyme  catalyzing the first committed step of the IMP pathway (Reaction 2), is likewise  subject to feedback inhibition. In this case, the enzyme binds ATP, ADP, and  AMP at one inhibitory site and GTP, GDP, and GMP at another. The rate of  IMP production is therefore independently but synergistically controlled by the levels  of adenine nucleotides and guanine nucleotides. Moreover, amidophosphoribosyl  transferase is allosterically stimulated by PRPP (feedforward activation).   A second level of regulation occurs immediately below the branch point  leading from IMP to AMP and GMP. AMP and GMP are each competitive  inhibitors of IMP in their own synthesis, which prevents excessive buildup of  the pathway products. In addition, the rates of adenine and guanine nucleotide  synthesis are coordinated. Recall that GTP powers the synthesis of AMP from  IMP, whereas ATP powers the synthesis of GMP from IMP (Fig. 23-3). This  reciprocity balances the production of AMP and GMP (which are required in  roughly equal amounts in nucleic acid biosynthesis): The rate of synthesis of  GMP increases with [ATP], whereas that of AMP increases with [GTP].    FIG. 23-4 Control of the purine  biosynthesis pathway. Red octagons and  green circles indicate control points. Feedback  inhibition is indicated by dashed red arrows, and  feedforward activation is represented by a  dashed green arrow. Q See the Animated  Figures.   In most cells, the turnover of nucleic acids, particularly some types of RNA,  releases adenine, guanine, and hypoxanthine (Section 23-4A). These free  purines are reconverted to their corresponding nucleotides through salvage  pathways. In contrast to the de novo purine nucleotide synthetic pathway,  which is virtually identical in all cells, salvage pathways are diverse in charac¬  ter and distribution. In mammals, purines are mostly salvaged by two differ¬  ent enzymes. Adenine phosphoribosyltransferase (APRT) mediates AMP  formation using PRPP:   Hypoxanthine-guanine phosphoribosyltransferase (HGPRT) catalyzes the  analogous reaction for both hypoxanthine and guanine:   Hypoxanthine + PRPP v IMP + PP Z  Guanine + PRPP v GMP + PP Z   Lesch-Nyhan Syndrome Results from HGPRT Deficiency. The symp¬  toms of Lesch-Nyhan syndrome, which is caused by a severe HGPRT  deficiency, indicate that purine salvage reactions have functions other than  conservation of the energy required for de novo purine biosynthesis. This sex-  linked congenital defect (it affects mostly males) results in excessive uric acid  production (uric acid is a purine degradation product; Section 23-4A) and  neurological abnormalities such as spasticity, mental retardation, and highly  aggressive and destructive behavior, including a bizarre compulsion toward  self-mutilation. For example, many children with Lesch-Nyhan syndrome  have such an irresistible urge to bite their lips and fingers that they must be  restrained. If the restraints are removed, communicative patients will plead that  the restraints be replaced, even as they attempt to injure themselves.   The excessive uric acid production in patients with Lesch-Nyhan syn¬  drome is readily explained. The lack of HGPRT activity leads to an accumu¬  lation of the PRPP that would normally be used to salvage hypoxanthine and    • List the starting materials and cofactors  required for IMP biosynthesis.   • Describe the reactions that convert IMP  to ATP and GTP.    • How do guanine and adenine nucleotides  inhibit their own synthesis? How do they  promote synthesis of each other?   guanine. The excess PRPP activates amidophosphoribosyl transferase (which  catalyzes Reaction 2 of the IMP biosynthetic pathway), thereby greatly  accelerating the synthesis of purine nucleotides and thus the formation of their  degradation product, uric acid. Yet the physiological basis of the associated  neurological abnormalities remains obscure. That a defect in a single enzyme  can cause such profound but well-defined behavioral changes nevertheless has  important neurophysiological implications.   • UMP is synthesized as a pyrimidine base to which ribose-5-phosphate is added.   • The early steps of pyrimidine nucleotide synthesis are the pathway’s major control  points.    The biosynthesis of pyrimidines is simpler than that of purines. Isotopic la¬  beling experiments have shown that atoms Nl, C4, C5, and C6 of the pyrim¬  idine ring are all derived from aspartic acid, C2 arises from HC0 3 , and N3  is contributed by glutamine.   UMP, which is also the precursor of CMP, is synthesized in a six-reaction path¬  way (Fig. 23-5). In contrast to purine nucleotide synthesis, the pyrimidine ring  is coupled to the ribose-5-phosphate moiety after the ring has been synthesized.   1. Synthesis of carbamoyl phosphate. The first reaction of pyrimidine  biosynthesis is the synthesis of carbamoyl phosphate from HC0 3 and  the amide nitrogen of glutamine by the cytosolic enzyme carbamoyl  phosphate synthetase II. This reaction consumes two molecules of ATP:  One provides a phosphate group and the other energizes the reaction.  Carbamoyl phosphate is also synthesized in the urea cycle (Section 21-3A).  In that reaction, catalyzed by the mitochondrial enzyme carbamoyl phos¬  phate synthetase I, ammonia is the nitrogen source.   2. Synthesis of carbamoyl aspartate. Condensation of carbamoyl phos¬  phate with aspartate to form carbamoyl aspartate is catalyzed by  aspartate transcarbamoylase (ATCase). This reaction proceeds with¬  out ATP hydrolysis because carbamoyl phosphate is already “activated.”  The structure and regulation of E. coli ATCase are discussed in Section  12-3.   3. Ring closure to form dihydroorotate. The third reaction of the path¬  way is an intramolecular condensation catalyzed by dihydroorotase to  yield dihydroorotate.   4. Oxidation of dihydroorotate. Dihydroorotate is irreversibly oxidized  to orotate by dihydroorotate dehydrogenase. The eukaryotic enzyme,  which contains FMN and nonheme Fe, is located on the outer surface  of the inner mitochondrial membrane, where quinones supply its ox¬  idizing power. The other five enzymes of pyrimidine nucleotide biosyn¬  thesis are cytosolic in animal cells. Inhibition of dihydroorotate  dehydrogenase blocks pyrimidine synthesis in T lymphocytes, thereby  attenuating the autoimmune disease rheumatoid arthritis.    FIG. 23-5 The de novo synthesis of UMP. The X-ray structures of the six enzymes  catalyzing the reactions are shown as is described in the legend for Fig. 23-1. [PDBids:  enzyme 1, 1 BXR; enzyme 2, 5AT1; enzyme 3, 1J79; enzyme 4, 1D3H; enzyme 5, 10PR;  enzyme 6, 1 DBT.] Q See the Animated Figures.    5. Acquisition of the ribose phosphate moiety. Orotate reacts with  PRPP to yield orotidine-5'-monophosphate (OMP) in a reaction cat¬  alyzed by orotate phosphoribosyl transferase. The reaction, which is  driven by the hydrolysis of the eliminated PP Z , fixes the anomeric form  of pyrimidine nucleotides in the p configuration. Orotate phosphori¬  bosyl transferase also salvages other pyrimidine bases, such as uracil and  cytosine, by converting them to their corresponding nucleotides.   6. Decarboxylation to form UMP. The final reaction of the pathway is  the decarboxylation of OMP by OMP decarboxylase (ODCase) to  form UMP. ODCase enhances the rate (k c J by a factor of 2 X 10 23  over that of the uncatalyzed reaction, making it the most catalytically  proficient enzyme known. Nevertheless, the reaction requires no co¬  factors to help stabilize its putative carbanion intermediate. Although  the mechanism of the ODCase reaction is not fully understood, the  removal of OMPs phosphate group, which is quite distant from the    C6 carboxyl group, deceases the reaction rate by a factor of 7 X 10 7 ,  thus providing a striking example of how binding energy can be  applied to catalysis (preferential transition state binding).   In bacteria, the six enzymes of UMP biosynthesis occur as independent  proteins. In animals, however, as Mary Ellen Jones demonstrated, the first  three enzymatic activities of the pathway—carbamoyl phosphate synthetase II,  ATCase, and dihydroorotase—occur on a single 210-kD polypeptide chain.   r The pyrimidine biosynthetic pathway is a target for antiparasitic drugs. For  example, the parasite Toxoplasma gondii (Fig. 23-6), which infects most mam¬  mals, causes toxoplasmosis, a disease whose complications include blindness,  neurological dysfunction, and death in immunocompromised individuals (e.g.,  those with AIDS). Most parasites have evolved to take advantage of nutrients  supplied by their hosts, but T. gondii is unable to meet its needs exclusively  through nucleotide salvage pathways and retains the ability to synthesize uracil  de novo. Drugs that target the parasites carbamoyl phosphate synthetase II (an  enzyme whose structure and kinetics distinguish it from its mammalian counter¬  part) could therefore prevent T. gondii growth. Moreover, there is evidence that  77 gondii strains that have been engineered to lack carbamoyl phosphate syn¬  thetase II are avirulent and could be useful as vaccines in humans and livestock.   The synthesis of UTP from UMP is analogous to the synthesis of purine nu¬  cleoside triphosphates (Section 23-IB). The process occurs by the sequential ac¬  tions of a nucleoside monophosphate kinase and nucleoside diphosphate kinase:    UMP + ATP UDP + ADP  UDP + ATP UTP + ADP    CTP is formed by the amination of UTP by CTP synthetase (Fig. 23-7). In  animals, the amino group is donated by glutamine, whereas in bacteria it is  supplied directly by ammonia.    C Pyrimidine Nucleotide Biosynthesis Is Regulated at ATCase or  Carbamoyl Phosphate Synthetase II   In bacteria, the pyrimidine biosynthetic pathway is primarily regulated at  Reaction 2, the ATCase reaction (Fig. 23-8 d). In E. coli, control is exerted  through the allosteric stimulation of ATCase by ATP and its inhibition by CTP  (Section 12-3). In many bacteria, however, UTP is the major ATCase inhibitor.   In animals, ATCase is not a regulatory enzyme. Rather, pyrimidine biosyn¬  thesis is controlled by the activity of carbamoyl phosphate synthetase II, which  is inhibited by UDP and UTP and activated by ATP and PRPP (Fig. 23-8 £).  A second level of control in the mammalian pathway occurs at OMP decar¬  boxylase, for which UMP and to a lesser extent CMP are competitive in-    hibitors. In all organisms, the rate of OMP production varies with the avail¬  ability of its precursor, PRPP. Recall that the PRPP level depends on the ac¬  tivity of ribose phosphate pyrophosphokinase (Fig. 23-1, Reaction 1), which  is inhibited by ADP and GDP (Section 23-1C).   T Orotic Aciduria Results from an Inherited Enzyme Deficiency. Orotic  aciduria, an inherited human disease, is characterized by the urinary ex¬  cretion of large amounts of orotic acid, retarded growth, and severe anemia.  It results from a deficiency in the bifunctional enzyme catalyzing Reactions 5  and 6 of pyrimidine nucleotide biosynthesis. Consideration of the biochem¬  istry of this situation led to its effective treatment: the administration of uri¬  dine and/or cytidine. The UMP formed through the phosphorylation of the  nucleosides, besides replacing that normally synthesized, inhibits carbamoyl  phosphate synthetase II so as to attenuate the rate of orotic acid synthesis. No  other genetic deficiency in pyrimidine nucleotide biosynthesis is known in hu¬  mans, presumably because such defects are lethal in utero.    FIG. 23-8 Regulation of pyrimidine  biosynthesis. The control networks are shown  for (a) E. coli and ( b ) animals. Red octagons and  green circles indicate control points. Feedback  inhibition is represented by dashed red arrows,  and activation is indicated by dashed green  arrows. Q See the Animated Figures.   • Compare the pathways of purine and  pyrimidine nucleotide synthesis with re¬  spect to (a) precursors, (b) energy cost,   • How does regulation of pyrimidine synthesis  differ in bacteria and animals?    • Ribonucleotide reductase uses a free radical mechanism to convert  ribonucleotides to deoxyribonucleotides.   • Thymidylate synthase transfers a methyl group to dUMP to form thymine.    DNA differs chemically from RNA in two major respects: (1) Its nucleotides  contain 2'-deoxyribose residues rather than ribose residues, and (2) it contains    the base thymine (5-methyluracil) rather than uracil. In this section, we con¬  sider the biosynthesis of these DNA components.   Deoxyribonucleotides are synthesized from their corresponding ribonucleotides by  the reduction of their C2' position rather than by their de novo synthesis from  deoxyribose-containing precursors.    Enzymes that catalyze the formation of deoxyribonucleotides by the re¬  duction of the corresponding ribonucleotides are named ribonucleotide re¬  ductases (RNRs). There are three classes of RNRs, which differ in their  prosthetic groups, although they all replace the 2'-OH group of ribose with  H via a free-radical mechanism. We will discuss the mechanism of the so-   (a) Schematic diagram of the quaternary structure. The enzyme  consists of two identical pairs of subunits, R1 2 and R2 2 . Each R2  subunit contains a binuclear Fe(lll) complex that generates a phenoxy  radical at Tyr 122. The R1 subunits each contain three different  allosteric effector sites and five catalytically important Cys residues.  The enzyme’s two active sites occur at the interface between the R1  and R2 subunits. ( b ) A ribbon diagram of R2 2 viewed perpendicularly  to its twofold axis with the subunits drawn in blue and yellow. The Fe(lll)  ions are represented by orange spheres, and the radical-harboring Tyr  122 side chains are shown in space-filling representation with their C  and 0 atoms green and red. (c) The binuclear Fe(lll) complex of R2.  Each Fe(lll) ion is octahedrally coordinated by a His N atom and five  0 atoms, including those of the O 2- ion and the Glu carboxyl group   that bridges the two Fe(lll) ions. [Part b based on an X-ray structure by  Hans Eklund, Swedish University of Agricultural Sciences, Uppsala,  Sweden. PDBid 1 RIB.] 0 See Interactive Exercise 33.   called Class I RNRs, which contain an Fe or Mn prosthetic group and which Section 3 Formation of Deoxyribonucleotides   Class I RNRs reduce ribonucleoside diphosphates (NDPs) to the corre¬  sponding deoxyribonucleoside diphosphates (dNDPs). E. coli ribonucleotide  reductase, as Peter Reichard demonstrated, is mainly present in vitro as a het-  erotetramer that can be decomposed to two catalytically inactive homodimers,   Rl 2 and R2 2 (Fig. 23-9 a\ opposite). Each R1 subunit contains a substrate¬  binding site that includes several redox-active thiol groups. The R1 subunits  also contain three effector-binding sites that control the enzymes catalytic ac¬  tivity as well as its substrate specificity (see below).   The X-ray structure of R2 2 (Fig. 23-9 b), determined by Hans Eklund, re¬  veals that the subunits are bundles of eight unusually long helices. Each sub¬  unit contains a novel binuclear Fe(III) prosthetic group whose Fe(III) ions are  liganded by a variety of groups including an O 2 ion (Fig. 23-9 c). The Fe(III)  complex interacts with Tyr 122 to form an unusual tyrosyl free radical. Recall  that tyrosine radicals also take part in the reactions catalyzed by cytochrome c  oxidase (Section 18-2F) and plant Photosystem II (Section 19-2C).   JoAnne Stubbe has proposed the following catalytic mechanism for E. coli  ribonucleotide reductase (Fig. 23-10):   1. Ribonucleotide reductases free radical (X*) abstracts an H atom from  Cy of the substrate in the reactions rate-determining step.   2 and 3. Acid-catalyzed cleavage of the C2'—OH bond releases H 2 0 to  yield a radical-cation intermediate. The C3'-OH groups unshared    FIG. 23-10 Enzymatic  mechanism of ribonucleotide  reductase. The reaction occurs via  a free radical-mediated process in  which reducing equivalents are  supplied by the formation of an  enzyme disulfide bond. [After  Stubbe, J.A., J. Biol. Chem. 265,  5330 (1990).]   4. The radical-cation intermediate is reduced by the enzymes redox-  active sulfhydryl pair to yield a 3'-deoxynucleotide radical and a protein  disulfide group (this group must eventually be reduced to regenerate  the enzymes activity).   5. The 3' radical abstracts an H atom from the protein to yield the product  deoxynucleoside diphosphate and restore the enzyme to its radical state.   The Tyr 122 radical in R2 is buried 10 A beneath the surface of the protein, too  far for the enzymes catalytic site to abstract an electron directly from the sub¬  strate. Evidently, the protein mediates electron transfer from this tyrosyl radical  to another group that is closer to the substrate, probably the thiyl radical (—S*)  form of Cys 439 in R1 (represented as X* in Fig. 23-10). Two other R1 Cys  residues probably form the redox-active sulfhydryl pair that directly reduces the  substrate. The resulting disulfide bond is reduced via disulfide interchange with  yet two other Cys residues, which are positioned to accept electrons from exter¬  nal reducing agents to regenerate the active enzyme. Thus, each R1 subunit con¬  tains at least five Cys residues that chemically participate in nucleotide reduction.    The Inability of Oxidized Ribonucleotide Reductase to Bind Substrate  Serves an Essential Protective Function. Comparison of the X-ray struc¬  tures of reduced R1 (in which the redox-active Cys 225 and Cys 462 residues  are in their SH forms) and oxidized R1 (in which Cys 225 and Cys 462 are  disulfide-linked) reveals that Cys 462 in reduced R1 has rotated away from its  position in oxidized R1 to become buried in a hydrophobic pocket, whereas  Cys 225 moves into the region formerly occupied by Cys 462. The distance  between the formerly disulfide-linked S atoms thereby increases from 2.0 A  to 5.7 A. These movements are accompanied by small shifts of the surround¬  ing polypeptide chain. R1 Cys 225 in oxidized ribonucleotide reductase pre¬  vents the binding of substrate through steric interference of its S atom with  the substrate NDP s 02' atom.   The inability of oxidized ribonucleotide reductase to bind substrate has  functional significance. In the absence of substrate, the enzymes free radical  is stored in the interior of the R2 subunit, close to its dinuclear iron center.  When substrate is bound, the radical is presumably transferred to it via a se¬  ries of protein side chains in both R2 and Rl. If the substrate is unable to  properly react after accepting this free radical, as would be the case if the en¬  zyme were in its oxidized state, the free radical could potentially destroy both  the substrate and the enzyme. Thus, an important role of the enzyme is to con¬  trol the release of the radical's powerful oxidizing capability. It does so in part by  preventing the binding of substrate while the enzyme is in its oxidized form.    Thioredoxin Reduces Ribonucleotide Reductase. The final step in the ribonu¬  cleotide reductase catalytic cycle is reduction of the enzymes newly formed disulfide  bond to re-form its redox-active sulfhydryl pair. One of the enzymes physiological  reducing agents is thioredoxin, a ubiquitous monomeric protein with a pair of  neighboring Cys residues (and which also participates in regulating the Calvin  cycle; Section 19-3C). Thioredoxin reduces oxidized ribonucleotide reductase via  disulfide interchange.   FIG. 23-1 1 X-Ray structure of human  thioredoxin in its reduced (sulfhydryl) state.   The backbone of the 105-residue monomer is  colored in rainbow order from its N-terminus  (blue) to its C-terminus (red). The side chains of  the redox-active Cys residues are shown in  space-filling form with C green and S yellow.  [Based on an X-ray structure by William  Montfort, University of Arizona. PDBid 1ERT.]    The X-ray structure of thioredoxin (Fig. 23-11) reveals that its redox-active  disulfide group is located on a molecular protrusion, making the protein the only  known example of a “male” enzyme.   Oxidized thioredoxin is, in turn, reduced in a reaction mediated by thiore¬  doxin reductase, which contains redox-active thiol groups and an FAD pros¬  thetic group. This enzyme, a homolog of glutathione reductase (Box 15-4) and  lipoamide dehydrogenase (Section 17-2B), catalyzes a similar reaction, the  NADPH-mediated reduction of a substrate disulfide bond. NAD PH therefore  serves as the terminal reducing agent in the ribonucleotide reductase—catalyzed  reduction of NDPs to dNDPs (Fig. 23-12).   The synthesis of the four dNTPs in the amounts required for DNA synthesis  is accomplished through feedback control. Maintaining the proper intracellu¬  lar ratios of dNTPs is essential for normal growth. Indeed, a deficiency of any  dNTP is lethal, whereas an excess is mutagenic because the probability that a given  dNTP will be erroneously incorporated into a growing DNA strand increases with  its concentration relative to those of the other dNTPs.   The activities of both E. coli and mammalian ribonucleotide reductases  are remarkably responsive to the levels of nucleotides in the cell. For example,  ATP induces the reduction of CDP and UDP; dTTP induces the reduction  of GDP and inhibits the reduction of CDP and UDP; and dATP inhibits the  reduction of all NDPs. Barry Cooperman has shown that the catalytic activ¬  ity of mouse ribonucleotide reductase varies with its state of oligomerization,  which in turn is governed by the binding of nucleotide effectors to three in¬  dependent allosteric sites on R1 (Fig. 23-9 a): (1) the specificity site, which  binds ATP, dATP, dGTP, and dTTP; (2) the activity site, which binds ATP  and dATP; and (3) the hexamerization site, which binds only ATP.  Coopermans model for the allosteric regulation of Class I ribonucleotide re-    FIG. 23-12 An electron-transfer pathway for nucleoside diphosphate (NDP)  reduction. NADPH provides the reducing equivalents for the process through the  intermediacy of thioredoxin reductase, thioredoxin, and ribonucleotide reductase.    FIG. 23-13 Regulation of  ribonucleotide reductase. Allosteric  regulators alter the state of oligomerization.  States shown in green have high activity,  and states shown in red have little or no  activity. R2 has been omitted for simplicity.  [After Kashlan, O.B., Scott, C.P., Lear, J.D.,  and Cooperman, B.S., Biochemistry 41 , 461  ( 2002 ).]    ductase, which quantitatively accounts for the enzymes regulatory properties,  has the following features (Fig. 23-13):   1. The binding of ATP, dATP, dGTP, or dTTP to the specificity site in¬  duces the catalytically inactive R1 monomers to form a catalytically ac¬  tive dimer, Rl 2 .   2. The binding of dATP or ATP to the activity site causes the dimers to  form tetramers, Rl 4 a , that slowly but reversibly change conformation  to a catalytically inactive state,   3. The binding of ATP to the hexamerization site induces the tetramers  to further aggregate to form catalytically active hexamers, Rl^, the en¬  zymes major active form.   The concentration of ATP in a cell is such that, in vivo , R1 is almost entirely  in its tetrameric or hexameric forms. As a consequence, ATP couples the over¬  all rate of DNA synthesis to the cells energy state.   dNTPs Are Produced by Phosphorylation of dNDPs. The final step in the  production of all dNTPs is the phosphorylation of the corresponding dNDPs:   This reaction is catalyzed by nucleoside diphosphate kinase, the same enzyme  that phosphorylates NDPs (Section 23-IB). As before, the reaction is written  with ATP as the phosphoryl donor, although any NTP or dNTP can func¬  tion in that capacity.   The dTTP substrate for DNA synthesis is derived from dUTP, which is hy¬  drolyzed to dUMP by dUTP diphosphohydrolase (dUTPase):   The dUMP is then methylated to generate dTMP, and the dTMP is phospho-  rylated to form dTTP. The apparent reason for the energetically wasteful process  of dephosphorylating dUTP and rephosphorylating dTMP is that cells must  minimize their concentration of dUTP in order to prevent incorporation of  uracil into their DNA (the enzyme system that synthesizes DNA from dNTPs  does not efficiently discriminate between dUTP and dTTP; Section 25-2A).   Human dUTPase is a homotrimer of 141-residue subunits. Its X-ray struc¬  ture, determined by John Tainer, reveals the basis for the enzymes exquisite  specificity for dUTP. Each subunit binds dUTP in a snug-fitting cavity that  sterically excludes thymines C5 methyl group via the side chains of conserved   FIG. 23-14 X-Ray structure of human dUTPase. (a) The active  site region of dUTPase in complex with dUTP. The protein is  represented by its molecular surface colored according to its  electrostatic potential (negative, red; positive, blue; and near neutral,  white). The dUTP is shown in ball-and-stick form with its N, 0, and P  atoms blue, red, and yellow. Mg 2+ ions that have been modeled into  the structure are represented by green spheres. Note the  complementary fit of the uracil ring into its binding pocket, particularly  the close contacts that discriminate against a methyl group on C5 of  the pyrimidine ring and a 2'-0H group on the ribose ring, (b) The   binding site of dUMP, showing the hydrogen bonding system  responsible for the enzyme’s specific binding of a uracil ring. The  dUMP and the polypeptide backbone binding it are shown in ball-and-  stick form with atoms colored as in Part a; hydrogen bonds are  indicated by white dotted lines; and a conserved water molecule is  represented by a pink sphere. The side chain of a conserved Tyr is  tightly packed against the ribose ring so as to discriminate against the  presence of a 2'-0H group. [Courtesy of John Tainer and Clifford Mol,  The Scripps Research Institute, La Jolla, California.]    residues (Fig. 23-14^). The enzyme differentiates uracil from the similarly  shaped cytosine via a set of hydrogen bonds that in part mimic adenines base  pairing interactions (Fig. 23-l4£). The 2'-OH group of ribose is likewise ster-  ically excluded by the side chain of a conserved Tyr.    Thymidylate Synthase Transfers a Methyl Group to dUMP. Thymidylate  (dTMP) is synthesized from dUMP by thymidylate synthase with TV 5 ,TV 10 -  methylenetetrahydrofolate (7V 5 ,A^ 10 -methylene-THF) as the methyl donor:   FIG. 23-1 5 Catalytic mechanism of thymidylate synthase. The methyl group is  supplied by A/ 5 ,A/ 10 -methylene-THF, which is concomitantly oxidized to dihydrofolate.    Note that the transferred methylene group (in which the carbon has the oxi¬  dation state of formaldehyde) is reduced to a methyl group (which has the ox¬  idation state of methanol) at the expense of the oxidation of the THF cofactor   Thymidylate synthase, a highly conserved 65-kD homodimeric protein,  follows a mechanistic scheme proposed by Daniel Sand (Fig. 23-15):   1. An enzyme nucleophile, identified as the thiolate group of Cys 146,  attacks C6 of dUMP to form a covalent adduct.   3. An enzyme base abstracts the acidic proton at the C5 position of the  enzyme-bound dUMP, forming an exocyclic methylene group and  eliminating the THF cofactor. The abstracted proton subsequently ex¬  changes with solvent.    FIG. 23-16 Regeneration of A/ 5 ,A/ 10 -methylenetetrahydrofolate. The DHF product  of the thymidylate synthase reaction is converted back to A/ 5 , A/ 10 -methylene-THF by the  sequential actions of ( 1 ) dihydrofolate reductase and ( 2 ) serine hydroxymethyltransferase.  The sites of action of some inhibitors are indicated by red octagons. Thymidylate synthase  is inhibited by FdUMP, whereas dihydrofolate reductase is inhibited by the antifolates  methotrexate, aminopterin, and trimethoprim (Box 23-1).    4. The redox change occurs via the migration of the C6-H atom of THF  as a hydride ion to the exocyclic methylene group, converting it to a  methyl group and yielding DHF. This reduction promotes the displace¬  ment of the Cys thiolate group from the intermediate to release prod¬  uct, dTMP, and re-form the active enzyme.    Tetrahydrofolate Is Regenerated in Two Reactions. The thymidylate synthase  reaction is biochemically unique in that it oxidizes THF to DHF; no other enzy¬  matic reaction employing a THF cofactor alters this coenzyme's net oxidation state.  The DHF product of the thymidylate synthase reaction is recycled back to  A^,A^°-methylene-THF through two sequential reactions (Fig. 23-16):   1. DHF is reduced to THF by NADPH as catalyzed by dihydrofolate  reductase (DHFR; Fig. 23-17). Although in most organisms DHFR  is a monomeric, monofunctional enzyme, in protozoa and some plants   FIG. 23-1 7 X-Ray structure of human  dihydrofolate reductase in complex with folic  acid. The polypeptide is colored in rainbow order  from its N-terminus (blue) to its C-terminus  (red). The folic acid is drawn in space-filling form  with C green, N blue, and 0 red. [Based on an  X-ray structure by Joseph Kraut, University of  California at San Diego. PDBid 1DHF.] © See  Interactive Exercise 34.   • What is the role of thioredoxin and  NADPH in the formation of deoxyribonu-  cleotides?    Describe the roles of dUTPase, thymi-  dylate synthase, and dihydrofolate  reductase in the synthesis of dTMP  What is the role of NADPH in the  formation of thymidylate?    DHFR and thymidylate synthase occur on the same polypeptide chain  to form a bifunctional enzyme that has been shown to channel DHF  from its thymidylate synthase to its DHFR active sites.   2. Serine hydroxymethyltransferase (Section 21-4A) transfers the hydroxy¬  methyl group of serine to THF yielding A/ 5 ,A^ 10 -methylene-THF and  glycine.   Inhibition of thymidylate synthase or DHFR blocks dTMP synthesis and is  therefore the basis of cancer chemotherapies (Box 23-1).    Most foodstuffs, being of cellular origin, contain nucleic acids. Dietary nucleic  acids survive the acidic medium of the stomach; they are degraded to their com¬  ponent nucleotides, mainly in the intestine, by pancreatic nucleases and intestinal  phosphodiesterases. The ionic nucleotides, which cannot pass through cell mem¬  branes, are then hydrolyzed to nucleosides by a variety of group-specific nucleoti¬  dases and nonspecific phosphatases. Nucleosides may be directly absorbed by the  intestinal mucosa or further degraded to free bases and ribose or ribose-1 -phos¬  phate through the action of nucleosidases and nucleoside phosphorylases:    Radioactive labeling experiments have demonstrated that only a small frac¬  tion of the bases of ingested nucleic acids are incorporated into tissue nucleic  acids. Evidently, the de novo pathways of nucleotide biosynthesis largely satisfy an  organisms need for nucleotides. Consequently, ingested bases are mostly degraded  and excreted. Cellular nucleic acids are also subject to degradation as part of the  continual turnover of nearly all cellular components. In this section, we outline  these catabolic pathways and discuss the consequences of several of their inher¬  ited defects. A summary of nucleotide metabolism is shown in Fig. 23-18.    FIG. 23-1 8 Summary of nucleotide  metabolism. Nucleotides are synthesized from  amino acids and ribose-5-phosphate by  pathways in which the base is built onto the  sugar (purine synthesis) or the sugar is added to  the base (pyrimidine synthesis). Nucleotide  catabolism yields a derivative of the base and  releases the sugar as ribose-1-phosphate.    dTMP synthesis is a critical process for rapidly proliferating cells,  such as cancer cells, which require a steady supply of dTMP for  DNA synthesis. Interruption of dTMP synthesis can therefore kill  those cells. Most normal mammalian cells, which grow slowly if  at all, require less dTMP and so are less sensitive to agents that  inhibit thymidylate synthase or dihydrofolate reductase (notable  exceptions are the bone marrow cells that constitute the blood-  forming tissue and much of the immune system, the intestinal  mucosa, and hair follicles).   is an irreversible inhibitor of thymidylate synthase. This substance,  like dUMP, binds to the enzyme (an F atom is not much larger  than an H atom) and undergoes the first two steps of the normal  enzymatic reaction (Fig. 23-15). In Step 3, however, the enzyme  cannot abstract the F atom as F + (F is the most electronegative  element) so that the enzyme is frozen in an enzyme-FdUMP-THF  ternary covalent complex.   The X-ray structure of the covalent thymidylate synthase-  FdUMP-THF ternary complex has been determined.   The enzyme’s active site region is shown with the polypeptide  chain colored in rainbow order from its N-terminus (blue) to its  C-terminus (red). The FdUMP, THF, and Cys 146 side chain are  drawn in stick form colored according to atom type (FdUMP C  cyan, THF and Cys C green, N blue, 0 red, F magenta, P orange,  and S yellow). The C5 and C6 atoms of FdUMP form covalent  bonds with the CH 2 group substituent to N5 of THF and the S  atom of Cys 146.   Enzyme inhibitors, such as FdUMP, that inactivate an enzyme  only after undergoing part or all of the normal catalytic reaction  are called mechanism-based inhibitors (alternatively, suicide  substrates because they cause the enzyme to “commit suicide”).  Because of their extremely high specificity, mechanism-based  inhibitors are among the most useful therapeutic agents.   Inhibition of DHFR blocks dTMP synthesis as well as all other  THF-dependent biological reactions, because the THF converted  to DHF by the thymidylate synthase reaction cannot be regener¬  ated. Methotrexate (amethopterin), aminopterin, and  trimethoprim   are DHF analogs that competitively although nearly irreversibly  bind to DHFR with an —1000-fold greater affinity than does DHF.  These antifolates (substances that interfere with the action of  folate cofactors) are effective anticancer agents, particularly  against childhood leukemias. In fact, a successful chemotherapeu¬  tic strategy is to treat a cancer victim with a lethal dose of  methotrexate and some hours later “rescue” the patient (but hope¬  fully not the cancer) by administering massive doses of 5-formyl-THF  and/or thymidine. Trimethoprim binds much more tightly to bacterial  DHFRs than to those of mammals and is therefore a clinically  useful antibacterial agent. A variety of compounds that inhibit  protozoan DHFR are used to treat (and prevent) malaria and  other parasitic infections.   [Based on an X-ray structure by William Montfort, University of  Arizona. PDBid 1TSNJ    Compare the activity of PNP to that of glycogen phosphorylase  (Section 16-1 A).   The major pathways of purine nucleotide and deoxynucleotide catabolism in  animals are diagrammed in Fig. 23-19. The pathways in other organisms dif¬  fer somewhat, but all the pathways lead to uric acid. Of course, the pathway  intermediates may be directed to purine nucleotide synthesis via salvage reac¬  tions. In addition, ribose-1-phosphate, a product of the reaction catalyzed by  purine nucleoside phosphorylase (PNP), is a precursor of PRPP.   Rather, adenine nucleosides and nucleotides are deaminated by adenosine  deaminase (ADA) and AMP deaminase to their corresponding inosine de¬  rivatives, which can then be further degraded.   ADA is an eight-stranded al (3 barrel (Fig. 23-20) with its active site  in a pocket at the C-terminal end of the (3 barrel, as in nearly all known   polypeptide is drawn in ribbon form colored according to its secondary  structure (helices cyan, |3 strands magenta, and loops pink) and viewed  approximately down the axis of the enzyme’s a/p barrel from the N-terminal  ends of its p strands. The transition state analog 6-hydroxy-1,6-dihydropurine  ribonucleoside (HDPR) is shown in stick form with its C, N, and 0 atoms  green, blue, and red. The enzyme-bound Zn 2+ ion, which is coordinated by  HDPR’s 6-hydroxyl group, is represented by a silver sphere. [Based on an  X-ray structure by Florante Quiocho, Baylor College of Medicine. PDBid  1 ADA.] O See Interactive Exercise 35.    a/|3 barrel enzymes. A catalytically essential zinc ion is bound  in the deepest part of the active site pocket. Mutations that af¬  fect the active site of ADA selectively kill lymphocytes, causing  severe combined immunodeficiency disease (SCID). Without  special protective measures, the disease is invariably fatal in in¬  fancy because of overwhelming infection.   Biochemical considerations provide a plausible explanation  of SCID’s etiology (causes). In the absence of active ADA, de-  oxyadenosine is phosphorylated to yield levels of dATP that are  50-fold greater than normal. This high concentration of dATP  inhibits ribonucleotide reductase (Section 23-3A), thereby pre¬  venting the synthesis of the other dNTPs, choking off DNA syn¬  thesis and thus cell proliferation. The tissue-specific effect of  ADA deficiency on the immune system can be explained by the  observation that lymphoid tissue is particularly active in de-  oxyadenosine phosphorylation. ADA deficiency was one of the  first genetic diseases to be successfully treated by gene therapy  (Section 3-5D).   This pathway functions in muscle to prime the  citric acid cycle by generating fumarate.    The Purine Nucleotide Cycle Generates Fumarate. The deamination of  AMP to IMP, when combined with the synthesis of AMP from IMP (Fig.  23-3, left), has the net effect of deaminating aspartate to yield fumarate  (Fig. 23-21). John Lowenstein demonstrated that this purine nucleotide cycle  has an important metabolic role in skeletal muscle. An increase in muscle  activity requires an increase in the activity of the citric acid cycle. This process  usually occurs through the generation of additional citric acid cycle interme¬  diates (Section 17-5B). Muscles, however, lack most of the enzymes that cat¬  alyze these anaplerotic (filling up) reactions in other tissues. Instead, muscle  replenishes its citric acid cycle intermediates with fumarate generated in the  purine nucleotide cycle.   The importance of the purine nucleotide cycle in muscle metabolism is  indicated by the observation that the activities of the three enzymes involved  are all severalfold higher in muscle than in other tissues. In fact, individuals  with an inherited deficiency in muscle AMP deaminase (myoadenylate  deaminase deficiency) are easily fatigued and usually suffer from cramps  after exercise.   (XO) converts hypoxanthine (the base of IMP) to xanthine, and xanthine to  uric acid (Fig. 23-19, bottom). The reaction product is an enol (which has a  p K of 5.4; hence the name uric acid). The enol tautomerizes to the more  stable keto form:    In mammals, xanthine oxidase occurs almost exclusively in the liver and  the small intestinal mucosa. It is a dimeric protein of identical 130-kD sub¬  units, each of which contains an entire “zoo” of electron-transfer agents: an  FAD, a Mo complex that cycles between its Mo(VI) and Mo(IV) oxidation  states, and two different Fe-S clusters. The final electron acceptor is 0 2 , which  is converted to H 2 0 2 , a potentially harmful oxidizing agent (Section 18-4B)  that is subsequently converted to H 2 0 and 0 2 by catalase.   In humans and other primates, the final product of purine degradation is uric  acid, which is excreted in the urine. The same is true in birds, terrestrial rep¬  tiles, and many insects, but those organisms, which do not excrete urea, also  catabolize their excess amino acid nitrogen to uric acid via purine biosynthe¬  sis. This complicated system of nitrogen excretion has a straightforward func¬  tion: It conserves water. Uric acid is only sparingly soluble in water, so its  excretion as a paste of uric acid crystals is accompanied by very little water.  In contrast, the excretion of an equivalent amount of the much more water-  soluble urea osmotically sequesters a significant amount of water.   In all other organisms, uric acid is further processed before excretion (Fig.  23-22). Mammals other than primates oxidize it to their excretory product,  allantoin, in a reaction catalyzed by the Cu-containing enzyme urate oxidase.  A further degradation product, allantoic acid, is excreted by teleost (bony)  fish. Cartilaginous fish and amphibia further degrade allantoic acid to urea  prior to excretion. Finally, marine invertebrates decompose urea to NH 4 .    T Gout Is Caused by an Excess of Uric Acid. Gout is a disease character¬  ized by elevated levels of uric acid in body fluids. Its most common man¬  ifestation is excruciatingly painful arthritic joint inflammation of sudden onset,  most often of the big toe (Fig. 23-23), caused by deposition of nearly insol¬  uble crystals of sodium urate. Sodium urate and/or uric acid may also precip¬  itate in the kidneys and ureters as stones, resulting in renal damage and urinary  tract obstruction.   Gout, which affects ~3 per 1000 persons, predominantly males, has been  traditionally, although inaccurately, associated with overindulgent eating and  drinking. The probable origin of this association is that in previous centuries,  when wine was often contaminated with lead during its manufacture and stor¬  age, heavy drinking resulted in chronic lead poisoning that, among other  things, decreases the kidneys ability to excrete uric acid.   The most prevalent cause of gout is impaired uric acid excretion (although  usually for reasons other than lead poisoning). Gout may also result from a  number of metabolic insufficiencies, most of which are not well characterized.    FIG. 23-22 Degradation of uric acid to  ammonia. The process is arrested at different  stages in the indicated species, and the resulting  nitrogen-containing product is excreted.    FIG. 23-23 The Gout, a cartoon by James  Gillray (1799). [Science Source/Photo Researchers,  Inc.]    Gertrude Elion (1918-1999) Gertrude  Elion’s choice of career was made at age 15  when she graduated from high school in New  York City. As she watched her grandfather  suffer and die from stomach cancer, she  decided to dedicate herself to finding a cure for  cancer. When she enrolled at Hunter College  the next fall, she began to study chemistry.  Despite her academic accomplishments, she  was unable to obtain a scholarship for graduate  school and spent the next two years as a teacher and lab assistant.  She eventually obtained a Master of Science degree in 1941.   After discovering that working in an industrial chemistry  laboratory was boring, Elion obtained a position with George  Hitchings at the pharmaceutical company Burroughs Wellcome.  There, she found an intellectually stimulating environment where  she was able to expand her understanding of chemistry as well as  pharmacology, immunology, and other fields. Although she  ultimately received numerous honorary degrees, Elion never  formally earned a doctoral degree; after several years of attending  classes at night, she was forced to choose between her work with  Hitchings, which she loved, and full-time study. Fortunately for  science and medicine, Elion kept her job and significantly  advanced the development of drugs for cancer, gout, organ  transplants, and infectious diseases.   Hitchings asked Elion to investigate purine metabolism, with the  idea that disrupting a cell’s supply of nucleotides could interfere  with its ability to synthesize DNA. Ideally, compounds that blocked  DNA synthesis would disable rapidly growing cancer cells, bacteria,  and viruses, without affecting normal cells. Without knowing the  structures of the relevant enzymes, or even of DNA, Elion and her  colleagues used a simple bacterial growth assay to investigate the  ability of various purine derivatives to act as “antimetabolites.”   Elion’s first clinical success was 6-mercaptopurine, which was  used to treat leukemia in children. Many  patients showed a dramatic recovery, but to  Elion’s dismay they often relapsed and later  died. Their initial improvement, however,  confirmed Elion’s belief that interfering with  nucleotide metabolism was a sound  therapeutic strategy.    A related compound, azathioprine, is  converted to 6-mercaptopurine  intracellularly. It turned out to be effective  not as an anticancer drug but as an  inhibitor of the immune response. This  drug helped solve the problem of  rejection in organ transplants and was  used for the first successful human  kidney transplant in 1961.   Studies aimed at improving the  effectiveness of purine derivatives by  preventing their degradation by xanthine oxidase led to the  discovery of allopurinol (Section 23-4B), which is still used to treat  gout and some parasitic diseases. Elion also helped develop the  antibacterial agent trimethoprim (Box 23-1) and the widely used  antiviral drug acyclovir (Zovir).   After Hitchings’ retirement, Elion  became head of the Department of  Experimental Therapy in 1967. Although  she formally retired in 1983, Elion’s  line of research continued to bear fruit  with the development of azidothymidine  (AZT; Box 12-3), a nucleoside analog  that was the sole drug that was  effective for treating AIDS until 1991  and is still in use.   Elion, together with Hitchings and  James Black [who discovered cimetidine  (Tagamet), the first drug to inhibit stomach acid secretion, and  propranolol (Inderol), which is widely used in the treatment of  high blood pressure], was awarded the Nobel Prize for  Physiology or Medicine in 1988, a rare accomplishment for a  scientist in the pharmaceutical industry. It was her research,  which was based on an understanding of nucleotide metabolism,  rather than her ability to simply synthesize novel compounds,  that merited recognition.   Elion, G.B., Hitchings, G.H., and Vanderwerff, H., Antagonists of nucleic acid  derivatives. VI. Purines, J. Biol. Chem. 192 , 505-518 (1951).   Elion, G.B., Kovensky, A., and Hitchings, G.H., Metabolic studies of allopurinol, an  inhibitor of xanthine oxidase, Biochem. Pharmacol. 15 , 863-880 (1966).   One well-understood cause is HGPRT deficiency (Lesch-Nyhan syndrome in  severe cases), which leads to excessive uric acid production through PRPP ac¬  cumulation (Section 23-ID).   Gout can be treated by administering the xanthine oxidase inhibitor  allopurinol {at right), a hypoxanthine analog with interchanged N7 and C8 po¬  sitions. Xanthine oxidase hydroxylates allopurinol, as it does hypoxanthine, yield¬  ing alloxanthine {at right), which remains tightly bound to the reduced form of  the enzyme, thereby inactivating it. Allopurinol consequently alleviates the symp¬  toms of gout by decreasing the rate of uric acid production while increasing the  levels of the more soluble hypoxanthine and xanthine. Although allopurinol con¬  trols the gouty symptoms of Lesch-Nyhan syndrome, it has no effect on its neu¬  rological symptoms. Allopurinol, along with several other notable purine  derivatives, was developed by the chemist Gertrude Elion (Box 23-2).    FIG. 23-24 The major pathways of pyrimidine catabolism in animals. The amino  acid products of these reactions are taken up in other metabolic processes. UMP and  dTMP are degraded by the same enzymes; the pathway for dTMP degradation is given in  parentheses.   • What compounds are produced by the  degradation of purines and pyrimidines?   • Describe the reactions catalyzed by nucle¬  oside phosphorylase, adenosine deami¬  nase, and xanthine oxidase.   • What are the physiological implications of  excreting waste nitrogen in the form or  urate, urea, or ammonia?   • Describe how purine catabolism is related  to SCID, muscle function, and gout.   Animal cells degrade pyrimidine nucleotides to their component bases (Fig.  23-24, top). The reactions, like those of purine nucleotides, occur through de¬  phosphorylation, deamination, and glycosidic bond cleavages. The resulting  uracil and thymine are then broken down in the liver through reduction (Fig.  23-24, middle) rather than by oxidation as occurs in purine catabolism. The  end products of pyrimidine catabolism, (3-alanine and (3-aminoisobutyrate,  are amino acids and are metabolized as such. They are converted, through  transamination and activation reactions, to malonyl-CoA and methylmalonyl-  CoA (Fig. 23-24, bottom left). Malonyl-CoA is a precursor of fatty acid syn¬  thesis (Fig. 20-26), and methylmalonyl-CoA is converted to the citric acid  cycle intermediate succinyl-CoA (Fig. 20-16). Thus, to a limited extent , catab¬  olism of pyrimidine nucleotides contributes to the energy metabolism of the cell.   • The purine nucleotide IMP is synthesized in 11 steps from ribose-  5-phosphate, aspartate, fumarate, glutamine, glycine, and HC0 3 .  Purine nucleotide synthesis is regulated at its first and second  steps.   • IMP is the precursor of AMP and GMP, which are phosphory-  lated to produce the corresponding di- and triphosphates.   • The pyrimidine nucleotide UMP is synthesized from 5-phospho-  ribosyl pyrophosphate, aspartate, glutamine, and HC0 3 in six re¬  actions. UMP is converted to UTP and CTP by phosphorylation  and amination.   • Pyrimidine nucleotide synthesis is regulated in bacteria at the  ATCase step and in animals at the step catalyzed by carbamoyl  phosphate synthetase II.   • Deoxyribonucleoside diphosphates are synthesized from the  corresponding NDPs by the action of ribonucleotide reductase,  which contains a binuclear Fe(III) prosthetic group, a tyrosyl  radical, and several redox-active sulfhydryl groups. Enzyme   • Ribonucleotide reductase is regulated by allosteric effectors, which  ensure that deoxynucleotides are synthesized in the amounts re¬  quired for DNA synthesis.   • dTMP is synthesized from dUMP by thymidylate synthase. The  dihydrofolate produced in the reaction is converted back to  tetrahydrofolate by dihydrofolate reductase (DHFR).   • Purine nucleotides are degraded by nucleosidases and purine nu¬  cleoside phosphorylase (PNP). Adenine nucleotides are deami-  nated by adenosine deaminase and AMP deaminase. The synthesis  and degradation of AMP in the purine nucleotide cycle yield the  citric acid cycle intermediate fumarate in muscles. Xanthine oxi¬  dase catalyzes the oxidation of hypoxanthine to xanthine and of  xanthine to uric acid.   • In primates, birds, reptiles, and insects, the final product of purine  degradation is uric acid, which is excreted. Other organisms de¬  grade urate further.   • Pyrimidines are broken down to intermediates of fatty acid me¬  tabolism.   3. List all the enzymes of nucleotide biosynthesis that use glutamine  as an amino group donor.   4. Certain glutamine analogs irreversibly inactivate enzymes that  bind glutamine. Identify the nucleotide biosynthetic intermedi¬  ates that accumulate in the presence of those compounds.   3. Calculate the cost, in ATP equivalents, of synthesizing de novo  (a) IMP, (b) AMP, and (c) CTP. Assume all substrates (e.g.,  ribose-3-phosphate and glutamine) and cofactors are available.    6. Rats are given cytidine that is l4 C-labeled at both its base and  ribose components. Their DNA is then extracted and degraded  with nucleases. Describe the labeling pattern of the recovered de-  oxycytidylate residues if deoxycytidylate production in the cell  followed a pathway in which (a) intact CDP is reduced to dCDP,  and (b) CDP is broken down to cytosine and ribose before  reduction.   7. Why does it make metabolic sense for UTP to inhibit carbamoyl  phosphate synthetase II, whereas ATP activates the enzyme?   8. Why does it make metabolic sense for ADP and GDP to inhibit  ribose phosphate pyrophosphokinase?   9. Explain why hydroxyurea, which destroys tyrosyl radicals, is use¬  ful as an antitumor agent.   11. Mouse embryonic stem cells divide extremely rapidly, about once  every 5 hours. These cells require large amounts of threonine in  their medium. Explain how threonine catabolism (Fig. 21-14)  helps meet the cells’ energy needs and their high rate of DNA  synthesis.   12. The purine and pyrimidine rings are built from the amino acids  aspartate, glutamine, and glycine. Why is serine required for the  synthesis of deoxyribonucleotides?   13. Why do individuals who are undergoing chemotherapy with  FdUMP or methotrexate often temporarily go bald?   14. Normal cells die in a nutrient medium containing thymidine and  methotrexate, whereas mutant cells defective in thymidylate syn¬  thase survive and grow. Explain.   13. Explain why methotrexate inhibits the synthesis of histidine and  methionine.   16. Some microorganisms lack DHFR activity, but their thymidylate  synthase has an FAD cofactor. What is the function of the FAD?   17. Is trimethoprim a mechanism-based inhibitor of bacterial dihy¬  drofolate reductase?   19. Why is it important that muscle cells have low levels of gluta¬  mate dehydrogenase?   21. Describe how the fumarate produced by the purine nucleotide  cycle could be catabolized to C0 2 .   22. Calculate the ATP yield of converting the carbons of thymine to   23. Individuals with a complete deficiency of dihydropyrimidine de¬  hydrogenase exhibit a variety of symptoms. What compounds are  likely to be present at high levels in the urine of these patients?   24. Individuals with a partial deficiency of dihydropyrimidine dehy¬  drogenase experience severe toxic effects when given high doses  of the anticancer drug 5-fluoro uracil. Explain.    25. In animals, one pathway for NAD + synthesis begins with nico¬  tinamide. Draw the structures generated by the reactions shown.    26. In the synthesis of FAD, riboflavin is phosphorylated by a kinase  to form FMN (Fig. 18-10). The FMN then reacts with ATP.  How many “high-energy” bonds are broken in this process?    One long-standing shortcom¬  ing of the RNA world hypothesis (Section 3-2C) is the lack of a  plausible explanation for the origin of ribonucleotides. Although a  molecule such as UMP has a modular structure (base + ribose +  phosphate), why is it unlikely that these components spontaneously  assembled to form the first nucleotides? Why is it more likely that  such compound arose from the combination of two molecules that  were half-base, half-ribose?    Carreras, C.W. and Santi, D.V., The catalytic mechanism and structure  of thymidylate synthase, Annu. Rev. Biochem. 64, 721-762 (1995).   Cotruvo, J.A., Jr. and Stubbe, J., Class I ribonucleotide reductases:  metallocofactor assembly and repair in vitro and in vivo, Annu. Rev.  Biochem. 80, 733-767 (2011).   Finer-Moore, J.S., Santi, D.V., and Stroud, R.M., Lessons and conclu¬  sions from dissecting the mechanism of a bisubstrate enzyme:  thymidylate synthase mutagenesis, function, and structure, Biochem¬  istry 42, 248-256 (2003).   Greasley, S.E., Horton, P., Ramcharan, J., Beardsley, G.P., Benkovic, S.J.,  and Wilson, I.A., Crystal structure of a bifunctional transformylase  and cyclohydrolase enzyme in purine biosynthesis, Nature Struct.  Biol. 8, 402-406 (2001).    Kappock, T.J., Ealick, S.E., and Stubbe, J., Modular evolution of the  purine biosynthetic pathway, Curr. Opin. Chem. Biol. 4, 567-572  ( 2000 ).   Liu, S., Neidhardt, E.A., Grossman, T.H., Ocain, T., and Clardy, J.,  Structures of human dihydroorotate dehydrogenase in complex with  antiproliferative agents, Structure 8, 25-33 (1999).   Nordlund, P. and Reichard, P., Ribonucleotide reductases, Annu. Rev.  Biochem. 75, 681-706 (2006).   Valle, D. (Ed.), The Online Metabolic & Molecular Bases of Inherited Dis¬  ease, http://www.ommbid.com/. [Part 11 contains chapters on de¬  fects in purine and pyrimidine metabolism.]   DNA molecules, such as that spilling out of the osmotically lysed bacteriophage T2  shown here, are enormous. A cell must be able to efficiently package and safely store  its DNA. However, despite DNA’s highly condensed structure, a cell must be able to  read and interpret the meaning of the encoded genetic information. [From Kleinschmidt,  A.K., Lang, D., Jacherts, D., and Zahn, R.K., Biochimica et Biophysics Acta, 61, 861  (1962). Reproduced with permission from Elsevier.]   A Nucleic Acids Are Stabilized by Base Pairing,  Stacking, and Ionic Interactions   C Eukaryotic Transcription Factors May Include  Zinc Fingers or Leucine Zippers    In every organism, the ultimate source of biological information is nucleic acid.  The shapes and activities of individual cells are, to a large extent, determined  by genetic instructions contained in DNA (or RNA, in some viruses).  According to the central dogma of molecular biology (Section 3-3B), sequences  of nucleotide bases in DNA encode the amino acid sequences of proteins. Many  of the cells proteins are enzymes that carry out the metabolic processes we have  discussed in Chapters 15—23. Other proteins have a structural or regulatory  role or participate in maintaining and transmitting genetic information.   Two kinds of nucleic acids, DNA and RNA, store information and make  it available to the cell. The structures of these molecules are consistent with  the following:   1. Genetic information must be stored in a form that is manageable in  size and stable over a long period.    2. Genetic information must be decoded—often many times—in order to  be used. Transcription is the process by which nucleotide sequences in  DNA are copied onto RNA so that they can direct protein synthesis, a  process known as translation.   3. Information contained in DNA or RNA must be accessible to proteins  and other nucleic acids. These agents must recognize nucleic acids (in  many cases, in a sequence-specific fashion) and bind to them in a way  that alters their function.   4. The progeny of an organism must be equipped with the same set of  instructions as in the parent. Thus, DNA is replicated (an exact copy  made) so that each daughter cell receives the same information.   As we will see, many cellular components are required to execute all the  functions of nucleic acids. Yet nucleic acids are hardly inert “read-only” enti¬  ties. RNA in particular, owing to its single-stranded nature, is a dynamic mol¬  ecule that provides structural scaffolding as well as catalytic proficiency in a  number of processes that decode genetic information. In this chapter, we  focus on the structural properties of nucleic acids, including their interactions  with proteins, that allow them to carry out their duties. In subsequent chap¬  ters, we will examine the processes of replication (Chapter 25), transcription  (Chapter 26), and translation (Chapter 27).    • A DNA helix can have the A, B, or Z conformation.   • The conformational freedom of the glycosidic bond, the ribose ring, and the  sugar-phosphate backbone is limited.   • Supercoiled DNA can be described in terms of linking number, twist, and  writhing number.   • Topoisomerases cut one (type I) or both (type II) strands of DNA to add or  remove supercoils.   We begin our discussion of nucleic acid structure by examining the various  forms of DNA, with an eye toward understanding how the molecule safe¬  guards genetic information while leaving it accessible for replication and tran¬  scription.   DNA is a two-stranded polymer of deoxynucleotides linked by phosphodiester  bonds (Figs. 3-3 and 3-6). The biologically most common form of DNA is  known as B-DNA, which has the structural features first noted by James  Watson and Francis Crick together with Rosalind Franklin and others (Section  3-2B and Box 24-1):   1. The two antiparallel polynucleotide strands wind in a right-handed  manner around a common axis to produce an ~20-A-diameter dou¬  ble helix.   2. The planes of the nucleotide bases, which form hydrogen-bonded  pairs, are nearly perpendicular to the helix axis. In B-DNA, the bases  occupy the core of the helix while the sugar—phosphate backbones  wind around the outside, forming the major and minor grooves. Only  the edges of the base pairs are exposed to solvent.    Rosalind Franklin (1920-1958) James  Watson and Francis Crick were the first to  publish an accurate model of the structure  of DNA. This seminal discovery was not  only based on their own insights but, like  all scientific discoveries, built on the work  of others. One of the major contributors to  this process was Rosalind Franklin, who  was probably never fully aware of her role  in this discovery and did not share the  1962 Nobel Prize in Physiology or Medicine awarded to Watson,  Crick, and Maurice Wilkins.   Franklin was born in England in an intellectually oriented and  well-to-do family. She excelled in mathematics, and although  career opportunities for women with her talents were few, she  obtained a doctorate in physical chemistry at the University of  Cambridge in 1945. From 1947 to 1950 she worked in a French  government laboratory, where she became an authority in applying  X-ray diffraction techniques to imperfectly crystalline substances  such as coal; her numerous publications on the structures of coal  and other forms of carbon changed the way that these  substances were viewed.   In 1951, she returned to England at the invitation of John  Randall, the head of the Medical Research Council’s (MRC’s)  Biophysics Research Unit at King’s College London, to investigate  the structure of DNA. Randall had indicated to Franklin that she  would be given an independent position, whereas Wilkins, who had  been working on DNA at King’s for some time, was given the  impression that Franklin would be working under his direction.   That misunderstanding, together with their sharply contrasting  personalities (Franklin was quick, assertive, and confrontational,  whereas Wilkins was deliberate, shy, and indirect), led to a falling  out between the two such that they were barely on speaking  terms. Hence, they worked independently. Some popular accounts  of Franklin’s life and work have implied that the atmosphere at  King’s College was inhospitable to women, but Franklin’s  correspondence as well as firsthand accounts indicate that the  atmosphere was in fact congenial. Still, she was unhappy at  King’s, apparently for personal reasons, and in the spring of 1953,  departed for Birkbeck College in London. There, until her untimely  death of ovarian cancer in 1958, she carried out groundbreaking  investigations on the structures of viruses.   Early in her tenure at King’s, Franklin discovered, through the  analysis of X-ray fiber diffraction patterns, that DNA (which she  obtained from Wilkins, who had gotten it from Rudolf Signer in  Switzerland) exists in two distinct conformations, which she called  the A and B forms. Prior to this discovery, the X-ray diffraction  patterns of DNA that had been obtained were confusing because  they were of mixtures of the A and B forms. Through careful  control of the humidity, Franklin obtained an X-ray fiber diffraction  photograph of B-DNA of unprecedented clarity (Fig. 3-5) that  strikingly indicated the DNA’s helical character. Analysis of the  photograph permitted Franklin to determine that B-DNA is double  helical, it has a diameter of 20 A, and each turn of the helix is  34 A long and contains 10 base pairs, each separated by 3.4 A.  Further analysis suggested that the hydrophilic sugar-phosphate  chains were on the outside of the helix and the relatively   hydrophobic bases were on the inside. Although Franklin was aware  of Chargaff’s rules (Section 3-2A) and Jerry Donohue’s work  concerning the tautomeric forms of the bases (Section 3-2B), she  did not deduce the existence of base pairs in double-stranded DNA.   In January 1953, Wilkins showed Franklin’s X-ray photograph  of B-DNA to Watson, when he visited King’s College. Moreover, in  February 1953, Max Perutz (Box 7-2), Crick’s thesis advisor at  Cambridge University, showed Watson and Crick his copy of the  1952 Report of the MRC, which summarized the work of all of its  principal investigators, including that of Franklin. Within a week  (and after 13 months of inactivity on the project), Watson and  Crick began building a model of DNA with a backbone structure  compatible with Franklin’s data [in earlier modeling attempts, they  had placed the bases on the outside of the helix (as did a model  published by Linus Pauling; Box 6-1) because they assumed that  the bases could transmit genetic information only if they were  externally accessible]. On several occasions Crick acknowledged  that Franklin’s findings were crucial to this enterprise.   Watson and Crick published their model of B-DNA in Nature in  April 1953. The paper was followed, back-to-back, by papers by  Wilkins and by Franklin on their structural studies of DNA.  Franklin’s manuscript had been written in March 1953, before she  knew about Watson and Crick’s work. The only change that  Franklin made to her manuscript when she became aware of  Watson and Crick’s model was the addition of a single sentence,  “Thus our general ideas are not inconsistent with the model  proposed by Watson and Crick in the preceding communication.”  She was apparently unaware that the Watson-Crick model was, to  a significant extent, based on her work. Since Watson and Crick  did not acknowledge Franklin in their 1953 Nature paper, her  paper was widely taken as data that supported the Watson-Crick  model rather than being an important element in its formulation.  Only after her death did Watson and Crick indicate the crucial role  of Franklin’s contributions.   Interestingly, Watson, Crick, and Franklin developed a close  friendship. Starting in 1954 they maintained a correspondence  and commented on each other’s work. In the summer of 1954,  Watson offered to drive Franklin across the United States from  Woods Hole, Massachusetts, to her destination at Caltech, where  he was also going. In the spring of 1956, she toured Spain with  Crick and his wife Odile and, later, stayed with them at their  house in Cambridge when she was recovering from her  treatments for ovarian cancer.   If Watson and Crick had not formulated their model of B-DNA,  would Franklin, who had a distaste for speculative modeling, have  eventually done so? We, of course, will never know. And, had she  lived, would Franklin have received the Nobel prize together with  Watson, Crick, and Wilkins? (The prize cannot be awarded  posthumously.) Those who argue that she was an indispensable  participant in the discovery process would say yes. However, the  Nobel committee typically awards those who initiate the research  (i.e., Wilkins), and the prize cannot be shared by more than three  individuals.    Franklin, R.E. and Gosling, R.G., Molecular configuration in sodium thymonucleate,  Nature 171, 740-741 (1953).   Maddox, B., Rosalind Franklin: The Dark Lady of DNA, HarperCollins (2002).    FIG. 24-1 The Watson-Crick base pairs. The line joining the Cl' atoms is the same  length in both A*T and G*C base pairs and makes equal angles with the glycosidic bonds  to the bases. This gives DNA a series of pseudo-twofold symmetry axes that pass through  the center of each base pair (red line ) and are perpendicular to the helix axis. [After Arnott,  S., Dover, S.D., and Wonacott, A.J., Acta Cryst. B25, 2196 (1969).] Q See Kinemage  Exercise 17-2.    3. Each base pair has approximately the same width (Fig. 24-1), which  accounts for the near-perfect symmetry of the DNA molecule, regard¬  less of base composition. A • T and G • C base pairs are interchange¬  able: They can replace each other in the double helix without altering the  positions of the sugar-phosphate backbones Cl' atoms. Likewise, the part¬  ners of a Watson-Crick base pair can be switched (i.e., by changing a  G • C to a C • G or an A • T to a T • A). In contrast, any other com¬  bination of bases would significantly distort the double helix.   4. The canonical (ideal) B-DNA helix has 10 base pairs (bp) per turn (a  helical twist of 36° per bp) and, since the aromatic bases have van der  Waals thicknesses of 3.4 A and are partially stacked on each other,  the helix has a pitch (rise per turn) of 34 A.   Double-helical DNA can assume several distinct structures depending on the  solvent composition and base sequence. The major structural variants of DNA  are A-DNA and Z-DNA. The geometries of the molecules are summarized in   A-DNA’s Base Pairs Are Inclined to the Helix Axis. Under dehydrating con¬  ditions, B-DNA undergoes a reversible conformational change to A-DNA,  which forms a wider and flatter right-handed helix than does B-DNA. A-DNA    Source: Mainly Arnott, S., in Neidle, S. (Ed.), Oxford Handbook of Nucleic Structure , p. 35, Oxford University Press (1999).    has 11.6 bp per turn and a pitch of 34 A, which gives it an axial hole (Fig.  24-2 c, left). A-DNA’s most striking feature, however, is that the planes of its  base pairs are tilted 20° with respect to the helix axis. Since the axis does not  pass through its base pairs, A-DNA has a deep major groove and a very shal¬  low minor groove; it can be described as a flat ribbon wound around a 6-A-  diameter cylindrical hole.   Z-DNA Forms a Left-Handed Helix. Occasionally, a seemingly familiar sys¬  tem exhibits quite unexpected properties. Over 25 years after the discovery  of the Watson-Crick DNA structure, the crystal structure determination of  d(CGCGCG) by Andrew Wang and Alexander Rich revealed, quite surpris¬  ingly, that this self-complementary sequence formed a left-handed double  helix. This helix, which was dubbed Z-DNA, has 12 Watson-Crick base  pairs per turn, a pitch of 44 A, a deep minor groove, and no discernible major  groove. Z-DNA therefore resembles a left-handed drill bit in appearance  (Fig. 24-2, right).   Fiber diffraction and NMR studies have shown that complementary  polynucleotides with alternating purines and pyrimidines, such as poly d(GC) •  poly d(GC) or poly d(AC) • poly d(GT), assume the Z conformation at high  salt concentrations. The salt stabilizes Z-DNA relative to B-DNA by reducing  the electrostatic repulsions between closest approaching phosphate groups on  opposite strands (which are 8 A apart in Z-DNA and 12 A apart in B-DNA).   Does Z-DNA have a biological function? The discovery of Z-DNA—binding  proteins strongly suggests that Z-DNA does exist in vivo. Rich has determined  the X-ray structure of a Z-DNA—binding domain named Z a in complex with  DNA. One Za domain binds to each strand of Z-DNA via hydrogen bonds  and ionic interactions between polar and basic side chains and the sugar-  phosphate backbone of the DNA; none of the DNAs bases participate in these  associations (Fig. 24-3). The proteins DNA-binding surface is complemen¬  tary in shape to the Z-DNA and is positively charged, as is expected for a pro¬  tein that interacts with closely spaced anionic phosphate groups. Sequences  capable of forming Z-DNA frequently occur near the start of genes, and the  reversible conversion of B-DNA to Z-DNA at these sites may play a role in  the control of transcription.   FIG. 24-2 Structures of A-, B-, and Z-DNA. (a) View  perpendicular to the helix axis. The sugar-phosphate backbones,  which wind about the periphery of each molecule, are outlined by a  green ribbon and the bases, which occupy its core, are red. Note that  the two sugar-phosphate chains in each helix run in opposite  directions so as to form right-handed double helices in A- and B-DNAs  and a left-handed double helix in Z-DNA. (b) Space-filling models  colored according to atom type with C white, N blue, 0 red, and P  orange, respectively. H atoms have been omitted for clarity, (c) View  along the helix axis. The ribose ring 0 atoms are red and the nearest  nucleotide pair is white. Note that the helix axis passes far “above”  the major groove of A-DNA, through the base pairs of B-DNA, and  through the edge of the minor groove of Z-DNA. Consequently,  A-DNA has a hollow core whereas B- and Z-DNAs have solid cores.  Also note that the deoxyribose residues in A- and B-DNAs have the    same conformation in each helix, but those in Z-DNA have two  different conformations so that alternate ribose residues lie at  different radii. [Based on structures by Olga Kennard, Dov Rabinovitch,  Zippora Shakked, and Mysore Viswamitra, Cambridge University, U.K.,  Nucleic Acid Database ID ADH010 (A-DNA); Richard Dickerson and  Horace Drew, Caltech, PDBid 1BNA (B-DNA); and Andrew Wang and  Alexander Rich, MIT, PDBid 2DCG (Z-DNA). Illustrations in Parts a  and c, Irving Geis, Images from the Irving Geis Collection, Howard  Hughes Medical Institute. Reprinted with permission. Model  coordinates for Part b generated by Helen Berman, Rutgers  University.] Q See Kinemage Exercises 17-1, 17-4, 17-5, and 17-6.    Would a protein that binds to a segment of B-DNA be able  to bind to A-DNA or Z-DNA?    RNA Can Form an A Helix. Double-stranded RNA is the genetic material of  certain viruses, but it is synthesized only as a single strand. Nevertheless, single-  stranded RNA can fold back on itself so that complementary sequences base-  pair to form double-stranded stems with single-stranded loops (Fig. 3-9).  Moreover, short segments of double-stranded RNA have been implicated in  the control of gene expression (Section 28-3C). Double-stranded RNA is  unable to assume a B-DNA-like conformation because of steric clashes involv¬  ing its 2'-OH groups. Rather, it usually assumes a conformation resembling  A-DNA (Fig. 24-2) that ideally has 11.0 bp per helical turn, a pitch of 30.9 A,  and base pairs that, on average, are inclined to the helix axis by 16.7°.   FIG. 24-3 X-Ray structure of Zol in complex with  Z-DNA. The DNA sequence d(CGCGCG) is shown in stick  form with its backbone red and its base pairs pink. The  Za domains are drawn with a helices blue and |3 sheets  light green. Note that each Za domain contacts only one  strand of Z-DNA and that none of the bases of the  Z-DNA interact directly with the protein. [Courtesy of  Alexander Rich, MIT. PDBid 1QBJJ   FIG. 24-4 X-Ray structure of a 10-bp  RNA-DNA hybrid helix. The complex consists  of the RNA UUCGGGCGCC that is base paired  to its DNA complement. The structure is shown  in stick form with RNA C atoms cyan, DNA C  atoms green, N blue, 0 red except for RNA 02'  atoms, which are magenta, and P gold. [Based  on an X-ray structure by Nancy Horton and  Barry Finzel, Pharmacia & Upjohn, Inc.,  Kalamazoo, Michigan. PDBid 1 FIX.] Q See  Interactive Exercise 36.    Hybrid double helices, which consist of one strand each of RNA and  DNA, also have an A-DNA-like conformation (Fig. 24-4), although hybrid  helices also have B-DNA-like qualities in that their overall conformation is  intermediate to those of A-DNA and B-DNA. Short stretches of RNA-DNA  hybrid helices occur during the initiation of DNA replication by small seg¬  ments of RNA (Section 25-1) and during the transcription of RNA on DNA  templates (Section 26-1C).   The structurally distinct A, B, and Z forms of DNA are not thought to freely  interconvert in vivo. Rather, the transition from one form to another requires  unusual physical conditions (e.g., dehydration) or the influence of DNA-binding  proteins. In addition, real DNA molecules deviate from the ideal structures  described in the preceding section. X-Ray structures of B-DNA segments re¬  veal that individual residues significantly depart from the average conformation  in a sequence-dependent manner. For example, the helical twist per base pair  may range from 26° to 43°. Each base pair can also deviate from its ideal con¬  formation by rolling or twisting like the blade of a propeller. Such conforma¬  tional variation appears to be important for the sequence-specific recognition  of DNA by the proteins that process genetic information.   B-DNA molecules, which are 20 A thick and many times as long, are  not perfectly rigid rods. In fact, it is imperative that the molecules be somewhat  flexible so that they can be packaged in cells. DNA helices can adopt differ¬  ent degrees of curvature ranging from gentle arcs to sharp bends. The more  severe distortions from linearity generally occur in response to the binding of  specific proteins.   Bond Rotation Is Hindered. The conformation of a nucleotide unit, as  Fig. 24-5 indicates, is specified by the six torsion angles of the sugar—phos¬  phate backbone and the torsion angle describing the orientation of the base  around the glycosidic bond (the bond joining CT to the base). It would seem  that these seven degrees of freedom per nucleotide would render polynu¬  cleotides highly flexible. Yet these torsion angles are subject to a variety of in¬  ternal constraints that greatly restrict their rotational freedom.   The rotation of a base around its glycosidic bond (angle x) is greatly hin¬  dered. Purine residues have two sterically permissible orientations known as   FIG. 24-6 The sterically allowed orientations of purine and pyrimidine bases  with respect to their attached ribose units. In B-DNA, the nucleotide residues all have  the anti conformation.    the syn (Greek: with) and anti (Greek: against) conformations (Fig. 24-6).  Only the anti conformation of pyrimidines is stable, because, in the syn con¬  formation, the sugar residue sterically interferes with the pyrimidines C2 sub¬  stituent. In most double-helical nucleic acids , all bases arc in the anti  conformation. The exception is Z-DNA (Section 24-1 A), in which the alter¬  nating pyrimidine and purine residues are anti and syn, respectively (this is  one reason why the repeating unit of Z-DNA is a dinucleotide).   The flexibility of the ribose ring itself is also limited. The vertex angles of  a regular pentagon are 108°, a value quite close to the tetrahedral angle  (109.5°), so one might expect the ribofuranose ring to be nearly flat. However,  the ring substituents are eclipsed when the ring is planar. To relieve this crowd¬  ing, which occurs even between hydrogen atoms, the ring puckers; that is, it  becomes slightly nonplanar. In the great majority of known nucleoside and  nucleotide X-ray structures, four of the ring atoms are coplanar to within a  few hundredths of an angstrom and the remaining atom is out of the plane  by several tenths of an angstrom. The out-of-plane atom is almost always C2'  or C3' (Fig. 24-7). The two most common ribose conformations are known  as C3 '-endo and Cl'-endo; “ endo ” (Greek: endon, , within) indicates that the  displaced atom is on the same side of the ring as C5', whereas “ exo (Greek: exo,  out of) indicates displacement toward the opposite side of the ring from C5'.   The ribose pucker is conformationally important in nucleic acids because  it governs the relative orientations of the phosphate substituents to each ribose  residue. In fact, B-DNA has the C2 '-endo conformation, whereas A-DNA is    FIG. 24-7 Nucleotide sugar conformations, (a) The C3 '-endo sugar-phosphate backbone are indicated. [After Saenger, W.,   conformation (C3' is displaced to the same side of the ring as C5'), Principles of Nucleic Acid Structure, p. 237, Springer-Verlag   which occurs in A-DNA. ( b ) The C2 '-endo conformation, which occurs (1983).] Q See Kinemage Exercise 17-3.   C3 '-endo. In Z-DNA, the purine nucleotides are all C3 ’-endo and the pyrimi¬  dine nucleotides are Cl’-endo.   Finally, if the torsion angles of the sugar—phosphate chain (angles a to £ in  Fig. 24-5) were completely free to rotate, there could probably be no stable  nucleic acid structure. However, these angles are actually quite restricted. This  is because of noncovalent interactions between the ribose ring and the phos¬  phate groups and, in polynucleotides, steric interference between residues. The  overall result is that the sugar—phosphate chains of the double helix are stiff  although the sugar—phosphate conformational angles are reasonably strain-free.   The chromosomes of many viruses and bacteria are circular molecules of du¬  plex DNA. In electron micrographs (e.g., Fig. 24-8), some of the molecules  have a peculiar twisted appearance, a phenomenon known as supercoiling or  superhelicity. Supercoiled DNA molecules are more compact than “relaxed”  molecules with the same number of nucleotides. This has important conse¬  quences for packaging DNA in cells (Section 24-5) and for the unwinding  events that occur as part of DNA replication and RNA transcription.   Superhelix Topology Can Be Simply Described. Consider a double-helical  DNA molecule in which each of its polynucleotide strands forms a covalently  closed circle, thus forming a circular duplex molecule. A geometric property  of such an assembly is that its number of coils cannot be altered without first  cleaving at least one of its strands. You can easily demonstrate this with a buck¬  led belt in which each edge of the belt represents a strand of DNA. The num¬  ber of times the belt is twisted before it is buckled cannot be changed without  unbuckling the belt (cutting a polynucleotide strand).   1. Z, the linking number, is the number of times that one DNA strand  winds around the other. This integer quantity is most easily counted  when the molecule is made to lie flat on a plane. The linking number  cannot be changed by twisting or distorting the molecule, as long as  both its polynucleotide strands remain covalently intact.   FIG. 24-8 Electron micrographs of circular duplex DNAs. Their conformations vary  from no supercoiling (left) to tightly supercoiled (right). [Electron micrographs by Laurien  Polder. From Kornberg, A. and Baker, T.A., DNA Replication (2nd ed.), p. 36, W.H. Freeman  (1992). Permission provided courtesy of Roger Kornberg.]    2. T, the twist, is the number of complete revolutions that one polynu¬  cleotide strand makes around the duplex axis. By convention, T is pos¬  itive for right-handed duplex turns so that, for B-DNA, the twist is  normally the number of base pairs divided by 10.5 (the observed num¬  ber of base pairs per turn of the B-DNA double helix in aqueous  solution).   3. W, the writhing number, is the number of turns that the duplex axis  makes around the superhelix axis. It is a measure of the DNA’s super-  helicity. The difference between writhing and twisting is illustrated in  Fig. 24-9. When a circular DNA is constrained to lie in a plane, W — 0.   The two DNA conformations diagrammed on the right of Fig. 24-10 are  topologically equivalent; that is, they have the same linking number, Z, but  differ in their twists and writhing numbers (topology is the study of the  geometric properties of objects that are unaltered by deformation but not by  cutting).   Since L is a constant in an intact duplex DNA circle , for every new double¬  helical twisty A Ty there must be an equal and opposite superhelical twist; that is ,  A W = —A T. For example, a closed circular DNA without supercoils (Fig.  24-10, upper righi) can be converted to a negatively supercoiled conformation  (Fig. 24-10, lower righi) by winding the duplex helix the same number of pos¬  itive (right-handed) turns.   Supercoiled DNA Is Relaxed by Nicking One Strand. Supercoiled DNA  may be converted to relaxed circles (as appears in the leftmost panel of Fig.  24-8) by treatment with pancreatic DNase I, an endonuclease (an enzyme  that cleaves phosphodiester bonds within a polynucleotide strand), which  cleaves only one strand of a duplex DNA. One single-strand nick is sufficient  to relax a supercoiled DNA. This is because the sugar-phosphate chain oppo¬  site the nick is free to swivel about its backbone bonds (Fig. 24-5) so as to  change the molecules linking number and thereby alter its superhelicity.  Supercoiling builds up elastic strain in a DNA circle, much as it does in a  rubber band. This is why the relaxed state of a DNA circle is not supercoiled.   Naturally Occurring DNA Circles Are Underwound. The linking numbers of  natural DNA circles are less than those of their corresponding relaxed circles;   FIG. 24-1 O Two ways of introducing one supercoil into a DNA that has 10  duplex turns. The two closed circular forms shown (right) are topologically equivalent; that  is, they are interconvertible without breaking any covalent bonds. The linking number L ,  twist T, and writhing number 1/1/ are indicated for each form. Strictly speaking, the linking  number is defined only for a covalently closed circle.   relaxed state (left), the spring or cord is in a  helical form that has a large writhing number  and a small twist. As the coil is pulled out  (center) until it is nearly straight (right), its  writhing number becomes small and its twist  becomes large.    FIG. 24-1 1 Progressive unwinding of a  negatively supercoiled DNA molecule. As the   double helix in a negatively supercoiled circle  (1/1/ < 0) is unwound without breaking covalent  bonds (T decreases), 1/1/ increases until it  reaches 0. Further unwinding of the double helix  then causes the DNA to supercoil in the  opposite direction, yielding a positively coiled  superhelix (1/1/ > 0).    that is, they are underwound. However, because DNA tends to adopt an over¬  all conformation that maintains its normal twist of 1 turn/10.5 bp, the mol¬  ecule is negatively supercoiled (W < 0; Fig. 24-11, left). If the duplex is  unwound (if T decreases), then ^increases (Z, must remain constant). At first,  this reduces the superhelicity of an underwound circle. However, with continued  unwinding, the value of W passes through zero (a relaxed circle; Fig. 24-11,  center) and then becomes positive, yielding a positively coiled superhelix  (Fig. 24-11, right).    DNA functions normally only if it is in the proper topological state. In such  basic biological processes as replication and transcription, complementary  polynucleotide strands must separate. The negative supercoiling of naturally  occurring DNAs in both prokaryotes and eukaryotes promotes such separa¬  tions since it tends to unwind the duplex helix (an increase in W must be  accompanied by a decrease in T). If DNA lacks the proper superhelical tension ,  the above vital processes cannot occur.   The supercoiling of DNA is controlled by a remarkable group of enzymes  known as topoisomerases. They are so named because they alter the topolog¬  ical state (linking number) of circular DNA but not its covalent structure.  There are two classes of topoisomerases in prokaryotes and eukaryotes:   1. Type I topoisomerases act by creating transient single-strand breaks  in DNA. Type I enzymes are further classified as type IA and type IB  topoisomerases, which differ in sequence and reaction mechanism.   2. Type II topoisomerases act by making transient double-strand breaks  in DNA.   Type IA Topoisomerases Relax Negatively Supercoiled DNA. Type I topo¬  isomerases catalyze the relaxation of supercoiled DNA by changing the linking num¬  ber in increments of one. Type IA enzymes, which are present in all cells, relax  only negatively supercoiled DNA. A clue to the mechanism of type IA  topoisomerases was provided by the observation that they reversibly catenate  (interlink) single-stranded circles (Fig. 24-12#). Apparently the enzyme oper¬  ates by cutting a single strand, passing a single-strand loop through the result¬  ing gap, and then resealing the break (Fig. 24-12/?). For negatively supercoiled  DNA, this increases the linking number and makes the writhing number more    FIG. 24-1 2 Type IA topoisomerase action. By cutting a single-stranded DNA,  passing a loop of a second strand through the break, and then resealing the break, a type  IA topoisomerase can (a) catenate two single-stranded circles or ( b ) unwind duplex DNA  by one turn.   positive (fewer negative supercoils). The process can be repeated until all of  the supercoils have been removed (W = 0). This strand passage mechanism  is supported by the observation that the denaturation of type IA topoisomerase  that has been incubated with single-stranded circular DNA yields a linear  DNA that has its 5'-terminal phosphoryl group linked to the enzyme via a  phospho-Tyr diester linkage:    Formation of the covalent enzyme—DNA intermediate conserves the free energy of  the cleaved phosphodiester bond so that no free energy input is needed to later  reseal the nick in the DNA.   E. coli topoisomerase III, a type IA enzyme, is a 659-residue monomer  (type I topoisomerases are denoted by odd Roman numerals; e.g., topoisom¬  erase I, III, etc.). The X-ray structure of its catalytically inactive Y328F  mutant in complex with the single-stranded octanucleotide d(CGCAACTT)  was determined by Alfonso Mondragon. The enzyme folds into four domains  that enclose an ~20 by 28 A hole which is large enough to contain duplex  DNA and which is lined with numerous Arg and Lys residues (Fig. 24-13).  The octanucleotide binds in a groove that is also lined with Arg and Lys side  chains with its sugar—phosphate backbone in contact with the protein and with  most of its bases exposed for possible base pairing. This single-stranded DNA  assumes a B-DNA-like conformation even though its complementary strand  would be sterically excluded from the groove. The DNA strand is oriented    FIG. 24-1 3 X-Ray structure of the Y328F  mutant of E. coli topoisomerase III in  complex with the single-stranded  octanucleotide d(CGCAACTT). The protein’s  four domains are drawn in different colors and  the two views shown are related by a 90°  rotation about a vertical axis. The DNA is drawn  in space-filling form with C white, N blue, 0 red,  and P yellow. This type IA topoisomerase’s active  site is marked by the side chain of Phe 328,  which is shown in space-filling form in light  green. [Based on an X-ray structure by Alfonso  Mondragon, Northwestern University. PDBid 117D.]   Chapter 24 Nucleic Acid Structure with its 3' end near the active site where, if the mutant Phe 328 were the   wild-type Tyr, its side chain would be properly positioned to nucleophilically  attack the phosphate group bridging the DNAs C-6 and T-7 nucleotides to  form a 5 r -phospho-Tyr linkage with T-7 and release C-6 with a free 3 r -OH.  This structure suggests the mechanism for the type IA topoisomerase-catalyzed  strand passage reaction that is diagrammed in Fig. 24-14.   Type IB topoisomerases can relax both negative and positive supercoils. In  doing so, they transiently cleave one strand of a duplex DNA through the  nucleophilic attack of an active site Tyr on a DNA P atom to yield a 3Hinked  phospho-Tyr intermediate and a free 5 r -OH group on the succeeding nu¬  cleotide (in contrast to the 5'-linked phospho-Tyr and free 3'-OH group  formed by type IA topoisomerases). The X-ray structure of the catalytically  inactive Y723F mutant of human topoisomerase I, a type IB topoisomerase,  in complex with a 22-bp duplex DNA was determined by Wim Hoi. The core  domain of the bilobal protein is wrapped around the DNA in a tight embrace  (Fig. 24-15). If the mutant Phe 723 were the wild-type Tyr, its OH group  would be ideally positioned to nucleophilically attack the P on the scissile  P—05 ' bond so as to form a covalent linkage with the 3’ end of the cleaved  strand. The protein interacts to a much greater extent with the five base pairs  of the DNAs “upstream” segment (which would contain the cleaved strands  newly formed 5' end) than it does with the base pairs of the DNAs “down¬  stream” segment (to which Tyr 723 would be covalently linked), and in both  cases it does so in a largely sequence-independent manner.   Topoisomerase I does not appear sterically capable of unwinding super-  coiled DNA via the strand passage mechanism that type IA topoisomerases    The gap between  domains I and III  closes, returning  the enzyme to its  initial state.   III and the ssDNA is cleaved. The new  5' end becomes covalently linked to  the active site Tyr while the new 3'  end remains tightly but noncovalently  bound in the binding groove.   The gap between domains I and I  reopens to permit the escape of the  green strand, yielding the reaction  product in which the green strand has  been passed through a transient break  in the red strand.   The unbroken {green)  strand is passed  through the opening  formed by the  cleaved {red) strand  to enter the protein’s  central hole.   The unbroken strand is  trapped by the partial  closing of the gap.    FIG. 24-1 4 Proposed mechanism for type IA  topoisomerases. The enzyme is shown in blue with the yellow patch  representing the binding groove for single-stranded (ss) DNA. The two  DNA strands, which are drawn in red and green, could represent the  two strands of a covalently closed circular duplex or two ss circles. If  the two strands form a negatively supercoiled duplex DNA, the    reaction results in its linking number, L, increasing by 1; if they are  separate ss circles, they become catenated or decatenated. For  negatively supercoiled duplex DNA, this process can be repeated until  all of its supercoils have been removed (1/1/ = 0). [After a drawing by  Alfonso Mondragon, Northwestern University.]    FIG. 24-1 5 X-Ray structure of the Y723F mutant of human topoisomerase I in  complex with a 22-bp duplex DNA. This type IB topoisomerase consists of several  domains and subdomains that are drawn here in different colors. Tyr 723 is on the  C-terminal domain (green). The DNA’s uncleaved strand is cyan, and the upstream and  downstream portions of the scissile strand are magenta and pink, respectively. [Courtesy of  Wim Hoi, University of Washington. PDBid 1A36.]    follow (Fig. 24-14). Rather, it is likely that topoisomerase I relaxes DNA  supercoils by permitting the cleaved duplex DNAs loosely held downstream  segment to rotate relative to the tightly held upstream segment. This rotation  can only occur about the sugar-phosphate bonds in the uncleaved strand (a,  (3, y, £, and £ in Fig. 24-5) that are opposite the cleavage site because the  cleavage frees those bonds to rotate. In support of this mechanism, the pro¬  tein region surrounding the downstream segment contains 16 conserved, pos¬  itively charged residues that form a ring about the duplex DNA, which  apparently holds the DNA but not in any specific orientation. Nevertheless,  the downstream segment is unlikely to rotate freely because the cavity con¬  taining it is shaped so as to interact with the downstream segment during some  portions of its rotation. Hence, type IB topoisomerases are said to mediate a  controlled rotation mechanism in relaxing supercoiled DNA. The unwind¬  ing is driven by the superhelical tension in the DNA and hence requires no  other energy input. Eventually, the DNA is religated by a reversal of the cleav¬  age reaction and the now less supercoiled DNA is released.   Type II Topoisomerases Function Via an ATP-Dependent Strand Passage  Mechanism. Type II topoisomerases are multimeric enzymes that require ATP  hydrolysis to complete a reaction cycle in which two DNA strands are cleaved,  duplex DNA is passed through the break, and the break is resealed. Type II  topoisomerases therefore change the linking number in increments of two  rather than one, as do type I topoisomerases. Both prokaryotic and eukary¬  otic type II enzymes relax negative and positive supercoils. However, only the  prokaryotic enzyme (also known as DNA gyrase or just gyrase) can introduce  negative supercoils [in contrast, the negative supercoils in eukaryotic chromo¬  somes result primarily from its packaging in nucleosomes (Section 24-5 B)  rather than from topoisomerase action]. Gyrases are A 2 B 2 heterotetramers,  whereas eukaryotic type II topoisomerases are homologous to gyrases but with  their A and B subunits fused so that they are homodimers.    FIG. 24-1 6 X-Ray structures of yeast  topoisomerase II. (a) Structure of a homodimer  of the N-terminal ATPase domain (residues  7-406 of the 1428-residue subunits) in complex  with AMPPNP as viewed with its twofold axis  vertical. The protein is displayed in ribbon form  with one subunit gray and the other colored in  rainbow order from its N-terminus (blue) to its  C-terminus (red). The bound AMPPNP  molecules are shown in space-filling form with C  green, N blue, 0 red, and P orange, (b) Structure  of a homodimer of the DNA breakage/reunion  domain (residues 419-1177) in complex with a  doubly nicked 34-bp DNA. The structure is  viewed with its twofold axis vertical. The DNA,  which describes a 150° arc, is drawn in stick  form with C green, N blue, 0 red, and P orange  embedded in its semitransparent molecular  surface (orange). The protein is displayed in  ribbon form with one subunit gray and the other  colored in rainbow order from its N-terminus  (blue) to its C-terminus (red). The side chains of  the two active site Tyr residues (Y782) are  shown in space-filling form with C magenta and  0 red. The polypeptides in Parts a and b, which  in the intact protein are joined by a 12-residue  linker (represented by the dotted lines),  presumably share the same twofold axis as  drawn, although their relative orientation about  the axis is unknown. [Based on X-ray structures  by James Berger, University of California at  Berkeley. PDBids 1 PVG and 2RGR.] Q See  Interactive Exercise 37.   Type II topoisomerases superficially resemble the type I enzymes in that  they have a pair of catalytic Tyr residues, which form transient covalent  intermediates with the 5' ends of duplex DNA. They thereby mediate the  cleavage of the two DNA strands at staggered sites to produce 4-nucleotide  “sticky ends.”   The X-ray structure of the homodimeric ATPase fragment of yeast topo¬  isomerase II (type II topoisomerases are denoted by even Roman numerals;  e.g., topoisomerase II, IV, etc.) in complex with the nonhydrolyzable ATP  analog AMPPNP (Section 13-2A), determined by James Berger, consists of  two domains (Fig. 24-16^). The N-terminal domain binds AMPPNP and the  C-terminal domains form the walls of a large hole through the dimer, which  in the X-ray structure of the structurally similar E. coli gyrase is 20 A across,  the same width as the B-DNA double helix.    G-segments are  resealed, lower  gate is opened,  and T-segment is  released.    Double stranded DNA  G-segment binding to  the enzyme induces a  conformational change,    ATP is hydrolyzed and released  and the lower gate is closed to  regenerate the G-segment •  enzyme complex ready to again  bind ATP and the T-segment.    changes induce ATPase  domain dimerization,  G-segment cleavage,  and upper gate opening  to form a hypothetical  intermediate (in square  brackets).    FIG. 24-1 7 Model for the enzymatic mechanism of type II  topoisomerases. The protein’s ATPase domain and the upper and  lower portions of the breakage/reunion domain are colored yellow,    red, and purple, respectively, and the DNA’s G- and T-segments (G for  gate; T for transported) are colored gray and green. [Courtesy of  James Wang, Harvard University.]    Berger also determined the X-ray structure of the yeast topoisomerase II  breakage/reunion domain in complex a 15-bp DNA that has a self-comple¬  mentary 4-nucleotide overhang on the 5' end of one its strands. The DNA  thereby forms a twofold symmetric 34-bp duplex with nicks on opposite  strands separated by 4 bp (Fig. 24-16£). These are precisely the sites at which  the enzyme would cleave an intact DNA by linking its newly formed 5'-ending  strands to the active site Tyr residues. The protein binds the DNA in a  positively charged groove that spans the width of the dimer and, in doing so,  bends it through an arc of 150° (as we will see, DNA-binding proteins often  deform their bound DNA, although such an extreme deformation is unusual).  Interestingly, the DNA between the two cleavage sites is essentially in the  A form. There are almost no direct contacts between the protein and the DNA  bases, as is expected for a protein with little sequence specificity. Note also  that the C-terminal portions of the protein come together to enclose a large  centrally located empty space.   Consideration of the foregoing two structures and those of the correspon¬  ding portions of E. coli gyrase suggests a model for the mechanism of type II  topoisomerases (Fig. 24-17). The DNA to be cleaved first binds to the  enzyme and is clamped in place. In the presence of ATP, the bound DNA is  cleaved and a second duplex DNA is passed through the opening into the cen¬  tral hole of the protein. The cleaved DNA is then resealed and the transported  DNA exits the complex at a point opposite its point of entry. ATP hydroly¬  sis to ADP + P z prepares the enzyme for an additional catalytic cycle. The  importance of topoisomerases in maintaining DNA in its proper topological  state is indicated by the fact that many antibiotics and chemotherapeutic  agents are inhibitors of topoisomerases (Box 24-2).   • Describe the structural differences  between A-, B-, and Z-DNA, including  handedness, diameter, and presence of  grooves.   • In what nucleic acid conformation(s) does  the ribose pucker with C2' endo ?   • Summarize the relationship between  linking number, twist, and writhing number.   • Why is it necessary for naturally occurring  DNA molecules to be negatively  supercoiled?   • How do type I A, type IB, and type II  topoisomerases alter DNA topology?  Which processes require the input of  free energy?    Type II topoisomerases are inhibited by a variety of compounds. For  example, ciprofloxacin and novobiocin (below) specifically inhibit  DNA gyrase but not eukaryotic topoisomerase II and are therefore  antibiotics. In fact, ciprofloxacin is the most efficacious oral antibiotic  presently in clinical use. In contrast, novobiocin’s adverse side effects  and the rapid generation of bacterial resistance to its presence have  resulted in the discontinuation of its use in the treatment of human  infections. A number of substances, including doxorubicin and  etoposide (below), inhibit eukaryotic type II topoisomerases and are  therefore widely used in cancer chemotherapy.   Different type II topoisomerase inhibitors act in one of two  ways. Many of these agents, including novobiocin, inhibit their  target enzyme’s ATPase activity. They therefore kill cells by  blocking topoisomerase activity, which results in the arrest of DNA  replication and RNA transcription since the cell can no longer  control the level of supercoiling in its DNA. However, other  substances, including ciprofloxacin, doxorubicin, and etoposide,  enhance the rate at which their target type II topoisomerases  cleave double-stranded DNA and/or reduce the rate at which the  breaks are resealed. Consequently, these substances induce    higher than normal levels of transient protein-bridged breaks in  the DNA of treated cells. The protein bridges are easily ruptured  by the passage of the replication and transcription machinery,  thereby rendering the breaks permanent. Although all cells have  extensive enzymatic machinery for repairing damaged DNA  (Section 25-5), a sufficiently high level of DNA damage can  overwhelm the repair mechanisms and trigger cell death. Conse¬  quently, since rapidly replicating cells such as cancer cells have  elevated levels of type II topoisomerases, they are far more likely  to incur lethal DNA damage through the inhibition of their type II  topoisomerases than are slow-growing or quiescent cells.   Camptothecin (below) and its derivatives, the only known  inhibitors of type IB topoisomerases, act by prolonging the lifetime  of the covalent enzyme-DNA intermediate. These substances bind  to the complex between the ends of the severed DNA strand,  positioning the 5'-OH groups over 4.5 A away from the  phosphate groups that must be reattached in the religation  reaction. Because the nicked DNA cannot be replicated,  camptothecin prevents the proliferation of rapidly growing cells  such as cancer cells and hence is a potent anticancer agent.    • A double-stranded nucleic acid structure is stabilized by hydrogen bonding  between base pairs, by stacking interactions, and by ionic interactions.   • DNA can be denatured by heating and renatured through annealing.   • RNA assumes more varied shapes than DNA and in some cases has  catalytic activity.    DNA does not exhibit the structural complexity of proteins because it has only  a limited repertoire of secondary structures and no comparable tertiary or   residues of proteins have a far greater range of chemical and physical proper- Acid Structures   ties than do the four DNA bases. Nevertheless, many RNAs have well-defined  tertiary structures. In this section, we examine the forces that determine the  structures of nucleic acids.   A Nucleic Acids Are Stabilized by Base Pairing, Stacking,  and Ionic Interactions   Base pairing is apparently a “glue” that holds together double-stranded nucleic  acids, although other intramolecular forces help shape nucleic acid structures.   Only Watson-Crick base pairs normally occur in DNA structures, but other  hydrogen-bonded base pairs are known, particularly in RNA (Fig. 24-18). For  example, in some A • T base pairs, adenine N7 is the hydrogen bond  acceptor (Hoogsteen geometry; Fig. 24-18/2) rather than N1 (Watson-Crick  geometry; Fig. 24-1).   Observations of DNA structures indicate that Watson-Crick geometry is the  most stable mode of base pairing in the double helix , even though non-Watson—Crick  base pairs are theoretically possible. Initially, it was believed that the geomet¬  rical constraints of the double helix precluded other types of base pairing.   Recall that because A • T, T • A, G • C, and C • G base pairs are geometrically  similar, they can be interchanged without altering the conformation of the  sugar-phosphate chains. However, experimental measurements show that a  major reason that other base pairs do not appear in the double helix is that  Watson-Crick base pairs have an intrinsic stability that non-Watson-Crick base  pairs lack; that is, the bases in a Watson-Crick pair have a higher mutual affin¬  ity than those in a non-Watson—Crick pair. Nevertheless, as we will see, the  double-helical segments of many RNAs contain unusual base pairs, such as  G • U, that help stabilize their tertiary structures.   Hydrogen Bonds Only Weakly Stabilize Nucleic Acid Structures. It is clear  that hydrogen bonding is required for the specificity of base pairing in DNA.   Yet, as is also true for proteins (Section 6-4A), hydrogen bonding contributes  little to the stability of nucleic acid structures. This is because, on denaturation,  the hydrogen bonds between the base pairs of a native nucleic acid are re¬  placed by energetically similar hydrogen bonds between the bases and water.   Other types of forces must therefore play an important role in stabilizing nu¬  cleic acid structures.   Stacking Interactions Result from Hydrophobic Forces. Purines and pyrim¬  idines tend to form extended stacks of planar parallel molecules. This has been   FIG. 24-1 8 Some non-Watson-Crick base pairs, (a) Hoogsteen pairing between  adenine and thymine residues. ( b ) An A* A base pair, (c) A U • C base pair. (R represents  ribose-5-phosphate).    FIG. 24-1 9 The stacking of adenine  rings in the X-ray structure of  9-methyladenine. The partial overlap of the  rings is typical of the association between bases  in crystal structures and in double-helical nucleic  acids. [After Stewart, R.F. and Jensen, L.H.,   Source: Ornstein, R.L., Rein, R., Breen, D.L., and  MacElroy, R.D., Biopolymers 17, 2356 (1978).   observed in the structures of nucleic acids (Fig. 24-2 e) and in the structures  of crystallized nucleic acid bases (e.g., Fig. 24-19). These stacking interac¬  tions are a form of van der Waals interaction (Section 2-1A). Interactions  between stacked G and C bases are greater than those between stacked A and  T bases (Table 24-2), which largely accounts for the greater thermal stability  of DNAs with a high G + C content. Note also that differing sets of base  pairs in a stack have different stacking energies. Thus, the stacking energy of a  double helix is sequence-dependent.   One might assume that hydrophobic interactions in nucleic acids are qual¬  itatively similar to those that stabilize protein structures. This is not the case.  Recall that folded proteins are stabilized primarily by the increase in entropy of  the solvent water molecules (the hydrophobic effect; Section 6-4A); in other  words, protein folding is enthalpically opposed and entropically driven. In con¬  trast, thermodynamic measurements reveal that the base stacking in nucleic acids  is enthalpically driven and entropically opposed, although the theoretical basis  for this observation is not well understood. The difference between hydropho¬  bic forces in proteins and in nucleic acids may reflect the fact that nucleic acid  bases are much more polar than most protein side chains (e.g., adenine versus  the side chain of Phe or Leu). Whatever their origin, hydrophobic forces are of  central importance in determining nucleic acid structures, as is clear from the  denaturing effect of adding nonpolar solvents to aqueous solutions of DNA.   Cations Shield the Negative Charges of Nucleic Acids. Any theory of the  stability of nucleic acid structures must take into account the electrostatic in¬  teractions of their charged phosphate groups. For example, DNA is stabilized  by Na + ions because the ions electrostatically shield the anionic phosphate  groups from each other. Other monovalent cations such as Li + and K + have  similar nonspecific interactions with phosphate groups. Divalent cations, such  as Mg 2+ , Mn 2+ , and Co 2+ , in contrast, specifically bind to phosphate groups,  so they are far more effective shielding agents for nucleic acids than are mono¬  valent cations. For example, an Mg 2+ ion has an influence on the DNA double  helix comparable to that of 100 to 1000 Na + ions. Indeed, enzymes that me¬  diate reactions with nucleic acids or nucleotides almost always require an  Mg 2+ ion cofactor for activity. Mg 2+ ions also play an essential role in stabi¬  lizing the complex structures assumed by many RNAs.   When a solution of duplex DNA is heated above a characteristic temperature,  its native structure collapses and its two complementary strands separate and  assume random conformations (Fig. 24-20). The denaturation process is  accompanied by a qualitative change in the DNAs physical properties. For    example, the characteristic high viscosity of native DNA solutions, which arises  from the resistance to deformation of its rigid and rodlike duplex molecules,  drastically decreases when the DNA decomposes to the conformationally  flexible single chains. Likewise, DNAs ultraviolet absorbance, which is almost  entirely due to its aromatic bases, increases by —40% on denaturation  (Fig. 24-21) as a consequence of the disruption of the electronic interactions  among neighboring bases. The increase in absorbance is known as the hyper-  chromic effect.   Monitoring the changes in absorbance at a single wavelength (usually 260  nm) as the temperature increases reveals that the increase in absorbance oc¬  curs over a narrow temperature range (Fig. 24-22). This indicates that the de¬  naturation of DNA is a cooperative phenomenon in which the collapse of one part  of the structure destabilizes the remainder. In analogy with the melting of a solid,  Fig. 24-22 is referred to as a melting curve, and the temperature at its mid¬  point is defined as its melting temperature, 7^.   FIG. 24-21 The UV absorbance spectra  of native and heat-denatured E. coli DNA.   Note that denaturation does not change the  general shape of the absorbance curve but only  increases its intensity. [After Voet, D., Gratzer,    FIG. 24-22 An example of a DNA melting curve. The relative absorbance is the  ratio of the absorbance (customarily measured at 260 nm) at the indicated temperature  to that at 25°C. The melting temperature, T m , is the temperature at which half of the  maximum absorbance increase is attained. O See the Animated Figures.    FIG. 24-23 Partially renatured DNA. This  schematic representation shows the imperfectly  base-paired structures assumed by DNA that  has been heat-denatured and then rapidly  cooled. Note that both intramolecular and  intermolecular aggregation can occur.   The stability of the DNA double helix, and hence its T m , depends on sev¬  eral factors, including the nature of the solvent, the identities and concentra¬  tions of the ions in solution, and the pH. T m also increases linearly with the  mole fraction of G • C base pairs, due to their greater stacking energy and not,  as one might naively assume, because G • C base pairs contain one more  hydrogen bond than A • T base pairs.   DNA Can Be Renatured by Cooling. If a solution of denatured DNA is rap¬  idly cooled below its T m , the resulting DNA will be only partially base-paired  (Fig. 24-23), because the complementary strands will not have had sufficient  time to find each other before the randomly base-paired structure becomes  effectively “frozen in.” If, however, the temperature is maintained ~25°C below  the T m , enough thermal energy is available for short base-paired regions to  rearrange by melting and re-forming. Under such annealing conditions, as Julius  Marmur discovered in 1960, denatured DNA eventually completely renatures.  Likewise, complementary strands of RNA and DNA, in a process known as hy¬  bridization, form RNA-DNA hybrid double helices (e.g., Fig. 24-4) that are  only slightly less stable than the corresponding DNA double helices.   The structure of RNA is stabilized by the same forces that stabilize DNA, and  its conformational flexibility is limited by many of the same features that limit  DNA conformation. In fact, RNA may be even more rigid than DNA owing  to the presence of a greater number of water molecules that form hydrogen  bonds to the 2'-OH groups of RNA. Even so, RNA comes in a greater vari¬  ety of shapes and sizes than DNA and, in some cases, has catalytic activity.   RNAs May Contain Double-Stranded Segments. Bacterial ribosomes, which  are two-thirds RNA and one-third protein, contain three highly conserved  RNA molecules (Section 27-3). The smallest of these is the 120-nucleotide 5S  RNA [so named because it sediments in the ultracentrifuge at the rate of  5 Svedbergs (S); Section 5-2E]. 5S RNA has a secondary structure consisting of  several base-paired regions connected by loops of various kinds (Fig. 24-24#).  These structural elements can be discerned in the X-ray structure of this RNA  molecule, in which the base-paired segments form mostly A-type helices   II | | | o o | o o o o | | | | | o o   FIG. 24-24 Secondary and tertiary structure of 5S  RNA from Haloarcula marismortui. (a) Secondary structure  diagram. This single-stranded RNA molecule contains four  double-stranded base-paired helices, as well as hairpin loops,  internal loops, and bulges. The dashes indicate Watson-Crick  base pairing, and circles indicate other modes of base pairing.  ( b ) The X-ray structure in stick form colored according to its  secondary structure as in Part a. [Based on an X-ray structure  by Peter Moore and Thomas Steitz, Yale University. PDBid 1JJ2.]   The 76-nucleotide RNA is shown in stick form  with C green, N blue, 0 red, and P orange and  with successive P atoms linked by orange rods.  Nearly all the bases are stacked, thereby  stabilizing the compact structure of the tRNA.  [Based on an X-ray structure by Alexander Rich  and Sung-Hou Kim, MIT. PDBid 6TNA.]    (Fig. 24-24 b). Approximately two-thirds of the bases in 5S rRNA participate  in base-pairing interactions; the remaining nucleotides, which are located in  loops and at the 3' and 5' ends, are free to interact with ribosomal proteins  or with unpaired nucleotides in other rRNA molecules.   Large RNA molecules presumably fold in stages, as do multidomain  proteins (Section 6-5). RNA folding is almost certainly a cooperative  process, with the rapid formation of short duplex regions preceding the  collapse of the structure into its mature conformation. The complete fold¬  ing process may take several minutes, it may involve relatively stable in¬  termediates, and it may require the assistance of proteins.    RNA Molecules Are Stabilized by Stacking Interactions. The three-  dimensional structure of the yeast transfer RNA that forms a covalent com¬  plex with Phe (tRNA Phe ) was elucidated in 1974 by Alexander Rich in  collaboration with Sung-Hou Kim and, in a different crystal form, by Aaron  Klug. The molecule is compact and L-shaped, with each leg of the L ~60 A  long (Fig. 24-25). The structural complexity of yeast tRNA Phe is reminis¬  cent of that of a protein. Although only 42 of its 76 bases occur in double¬  helical stems, 71 of them participate in stacking associations.   tRNA structures are characterized by the presence of covalently modified  bases and unusual base pairs, including hydrogen-bonding associations involv¬  ing three bases. These tertiary interactions contribute to the compact structure  of the tRNAs (tRNA structure is discussed in greater detail in Section 27-2A).    Some RNAs Are Catalysts. Although the discovery of catalytic RNA in 1982  was initially greeted with skepticism, subsequent studies have demonstrated that  the catalytic potential of RNA is virtually unlimited. At least nine naturally oc¬  curring types of catalytic RNAs have been described, and many more ribozymes  have been developed in the laboratory Most naturally occurring RNA catalysts    The observations that nucleic acids but not proteins can direct  their own synthesis, that cells contain batteries of protein-based  enzymes for manipulating DNA (Chapter 25) but few for process¬  ing RNA, and that many coenzymes in biosynthetic processes and  energy-harvesting pathways are ribonucleotides (e.g., ATP, NAD + ,  FAD, and coenzyme A) led to the hypothesis that RNAs were the  original biological catalysts in precellular times—the so-called  RNA world. Indeed, RNA remains the carrier of genetic  information in viruses with single- or double-stranded RNA  genomes. The fact that RNA-based functions are distributed  across all three domains of life provides additional evidence that  the role of RNA in modern cells reflects its establishment very  early in evolution.   Assuming that an RNA world did once exist, we must ask two  questions: Why did protein catalysts take the place of most  ribozymes, and why did DNA become the dominant molecule of  heredity? The first question is readily answered by observing that  the 20 amino acid residues of proteins contain functional groups—  hydroxyl, sulfhydryl, amide, and carboxylate groups-that are  lacking in RNA, whose four nucleotide residues are more uniform  and less reactive. Thus, proteins probably replaced RNA as cellular  workhorses by virtue of their greater chemical virtuosity.  Nevertheless, RNA continues to play the dominant role in the  synthesis of proteins: Ribosomes are predominantly RNA,  which is now known to catalyze peptide bond formation  (Section 27-4B).   As for DNA, it is more stable than RNA because RNA is  highly susceptible to base-catalyzed hydrolysis by the reaction  mechanism at the right. The base-induced deprotonation of the  2'-OH group facilitates its nucleophilic attack on the adjacent  phosphorus atom, thereby cleaving the RNA backbone. The  resulting 2',3'-cyclic phosphate group subsequently hydrolyzes to  produce a 2'- or 3'-nucleotide product (the RNase A-catalyzed  hydrolysis of RNA follows a nearly identical reaction sequence but  generates only 3'-nucleotides; Section 1 1-3A). DNA is not  susceptible to such degradation because it lacks a 2'-OH group.  This greater chemical stability of DNA makes it more suitable than  RNA for the long-term storage of genetic information.    participate in aspects of RNA metabolism such as mRNA splicing (Section   26- 3B) and hydrolytic RNA processing (Section 26-3C). The formation of pep¬  tide bonds during protein synthesis is catalyzed by ribosomal RNA (Section   27- 4). In vitro, RNA molecules can catalyze some of the reactions required for  their own production, such as the synthesis of glycosidic bonds and nucleotide  polymerization. This functional versatility has led to the hypothesis that RNA  once carried out many of the basic activities of early life, before DNA or pro¬  teins had evolved. This scenario has been dubbed the RNA world (Box 24-3).   One of the best-characterized ribozymes is the hammerhead ribozyme, a  minimally ~40-nt molecule that participates in the replication of certain virus¬  like RNAs that infect plants and also occurs in schistosomes (species of para¬  sitic flatworms). The hammerhead ribozyme catalyzes the site-specific cleavage  of one of its own phosphodiester bonds. The secondary structures of the 63-  nt hammerhead ribozyme from Schistosoma mansoni, which cleaves itself be¬  tween its C-17 and C-l.l nucleotides, has three duplex stems and a catalytic  core of two nonhelical segments (Fig. 24-26#). The X-ray structure of this    FIG. 24-26 Structure of a hammerhead ribozyme. (a) The sequence  and schematic structural representation of the ribozyme oriented and colored  to match the X-ray structure drawn in Part b. Base pairs are represented by  colored lines, and dotted lines represent tertiary interactions. Nucleotides are  labeled according to the universal numbering system for hammerhead  ribozymes. (b) X-Ray structure of the ribozyme drawn in paddle form with C  atoms the same color as in Part a except that those of G-12 are light green  and those of C-1.1 are light blue, N blue, 0 red, and P orange. Adjacent P  atoms in the same strand are connected by thin orange rods. The P atom of  the scissile phosphate group (bridging 017 and C-1.1) and the nucleophile  (02' of C-17) are represented by small spheres, (c) The ribozyme’s active  site residues are drawn in stick form with C green, 0 red, N blue, and P  orange. Hydrogen bonds are represented by dashed black lines. The  proposed reaction mechanism is indicated by the curved arrows with the  dotted black line marking the in-line trajectory taken by atom 02' of C-17 in  nucleophilically attacking the P atom of the scissile phosphate group. [Part a  courtesy of and Parts b and c based on an X-ray structure by William Scott,  University of California at Santa Cruz. PDBid 2G0Z.]    hammerhead ribozyme (Fig. 24-26 b), determined by William Scott, reveals that  it forms three A-type helices. The nucleotides in the helical stems mainly form  normal Watson—Crick base pairs, whereas the various loops and bulges inter¬  act through a variety of hydrogen bonding and base stacking interactions.   The bases in the highly conserved catalytic core of the hammerhead  ribozyme participate in an extensive hydrogen bonded network (Fig. 24-26 c).  This helps position C-17 such that its 02' atom is properly oriented for nu¬  cleophilic attack on the P atom linking atom 03' of C-17 to atom 05' of  C-1.1, so as to yield a cyclic 2',3'-phosphodiester on C-17 together with a free  5'-OH on C-1.1. The 2'-OH of the invariant G-8 and N1 of the invariant  G-12 when deprotonated appear to be properly positioned to act as acid and  base catalysts in the reaction. Interestingly, no metal ions are present in the  ribozymes active site region.   • Describe the forces that stabilize nucleic  acid structure. Which is most important?   • Explain the molecular events of nucleic  acid denaturation and renaturation.    The hammerhead ribozyme is not a true catalyst, since it is its own sub¬  strate and hence cannot return to its original state. However, other ribozymes  catalyze multiple turnovers without themselves undergoing alteration. Ribozymes  are comparable to protein enzymes in their rate enhancement (~10 7 -fold for  the hammerhead ribozyme), in their ability to use cofactors such as metal ions  or imidazole groups, and in their regulation by small allosteric effectors.  Although RNA molecules lack the repertoire of functional groups possessed by  protein catalysts, RNA molecules can assume conformations that specifically  bind substrates, orient them for reaction, and stabilize the transition state—all  hallmarks of enzyme catalysts.    Nucleic acids can be fractionated on the basis of size, composition, and  sequence by chromatography and electrophoresis.    In Section 5-2 we considered the most common procedures for isolating and  characterizing proteins. Most of these methods, often with some modifications,  are also used to fractionate nucleic acids according to size, composition, and se¬  quence. There are also many techniques that apply only to nucleic acids. In this  section, we outline some of the most useful nucleic acid fractionation procedures.   Nucleic acids in cells are invariably associated with proteins. Once cells have  been broken open, their nucleic acids are usually deproteinized. This can be ac¬  complished by shaking the protein-nucleic acid mixture with a phenol solution  so that the protein precipitates and can be removed by centrifugation.  Alternatively, the protein can be dissociated from the nucleic acids by detergents,  guanidinium chloride, or high salt concentration, or it can be enzymatically de¬  graded by proteases. In all cases, the nucleic acids, a mixture of RNA and DNA,  can then be isolated by precipitation with ethanol. The RNA can be recovered  from such precipitates by treating them with pancreatic DNase to eliminate the  DNA. Conversely, the DNA can be freed of RNA by treatment with RNase.   In all these and subsequent manipulations, the nucleic acids must be  protected from degradation by nucleases that occur both in the experimental  material and on human hands. Nucleases can be inhibited by chelating agents  such as EDTA, which sequester the divalent metal ions they require for  activity. Laboratory glassware can also be autoclaved to heat-denature the nu¬  cleases. Nevertheless, nucleic acids are generally easier to handle than proteins  because most lack a complex tertiary structure and are therefore relatively  tolerant of extreme conditions.   Many of the chromatographic techniques that are used to separate proteins  (Section 5-2C) also apply to nucleic acids. However, hydroxyapatite, a form  of calcium phosphate [Ca 5 (P04) 3 0H], is particularly useful in the chromato¬  graphic purification and fractionation of DNA. Double-stranded DNA binds  to hydroxyapatite more tightly than do most other molecules. Consequently,  DNA can be rapidly isolated by passing a cell lysate through a hydroxyapatite  column, washing the column with a phosphate buffer of concentration low  enough to release only the RNA and protein, and then eluting the DNA with  a concentrated phosphate solution.   Affinity chromatography is used to isolate specific nucleic acids. For  example, most eukaryotic messenger RNAs (mRNAs) have a poly(A) sequence  at their 3' ends (Section 26-3A). They can be isolated on agarose or cellulose  to which poly(U) is covalently attached. The poly(A) sequences specifically    bind to the complementary poly(U) in high salt and at low temperature and  can later be released by altering those conditions.   Nucleic acids of a given type can be separated by polyacrylamide gel elec¬  trophoresis (Section 3-4B) because their electrophoretic mobilities in such gels  vary inversely with their molecular masses. However, DNAs of more than a  few thousand base pairs cannot penetrate even a weakly cross-linked polyacryl¬  amide gel and so must be separated in agarose gels. Yet conventional gel elec¬  trophoresis is limited to DNAs of <100,000 bp, because larger DNA  molecules tend to worm their way through the agarose at a rate independent  of their size. Charles Cantor and Cassandra Smith overcame this limitation by  developing pulsed-field gel electrophoresis (PFGE), which can resolve  DNAs of up to 10 million bp (6.6 million kD). In the simplest type of PFGE  apparatus, the polarity of the electrodes is periodically reversed, with the du¬  ration of each pulse varying from 0.1 to 1000 s, depending on the sizes of the  DNAs being separated. With each change in polarity, the migrating DNA  must reorient to the new electrical field before it can resume movement. A  DNA molecule may actually migrate backward for part of the time. Because  small molecules reorient more quickly than large molecules, different-sized  DNA molecules gradually separate (Fig. 24-27).   Intercalation Agents Stain Duplex DNA. The various DNA bands in a gel  can be stained by planar aromatic cations such as ethidium ion, acridine  orange, or proflavin:    FIG. 24-27 Pulsed-field gel electrophoresis  (PFGE) of a set of yeast chromosomes. The   yeast chromosomes, which were run as identical  samples in the 13 inner lanes, have sizes 260,  290, 370, 460, 580/600, 700, 780, 820, and  850 kb. The two outer lanes, which provide  molecular mass standards, contained twenty  successively larger multimers of a —43.5-kb  bacteriophage X DNA (top to bottom) to an  observed limit of —850 kb. [Electrophoretogram  by Margit Burmeister, University of Michigan in  Wilson, K. and Walker, J., Principles and  Techniques of Biochemistry and Molecular Biology  (6th ed.), p. 477, Cambridge University Press  (2005).]   The dyes bind to double-stranded DNA by intercalation (slipping in between  the stacked bases; Fig. 24-28), where they exhibit a fluorescence under UV   FIG. 24-28 X-Ray structure of a complex  of ethidium with 5-iodo-UpA. Ethidium (red)  intercalates between the base pairs of the  double-helically paired dinucleotide and thereby  provides a model of the binding of ethidium to  duplex DNA. [After Tsai, C.-C., Jain, S.C., and  Sobell, H.M., Proc. Natl. Acad Sci. 72, 629 (1975).]   After electrophoresis, the gel was soaked in a  solution of ethidium bromide, washed, and  photographed under UV light. The fluorescence  of the ethidium cation is strongly enhanced by  binding to DNA, so each fluorescent band marks  a different sized DNA fragment. [Klaus  Guldbrandsen/Photo Researchers.]   light that is far more intense than that of the free dye (Fig. 24-29). As little  as 50 ng of DNA can be detected in a gel by staining it with ethidium bro¬  mide. Single-stranded DNA and RNA also stimulate the fluorescence of ethid¬  ium but to a lesser extent than does duplex DNA.   Southern Blotting Identifies DNAs with Specific Sequences. DNA with a  specific base sequence can be identified through a procedure developed by  Edwin Southern known as Southern blotting (Fig. 24-30). This procedure  takes advantage of the ability of nitrocellulose to tenaciously bind single-  stranded but not duplex DNA. Following gel electrophoresis of double-  stranded DNA, the gel is soaked in 0.5 M NaOH to convert the DNA to its  single-stranded form. The gel is then overlaid by a sheet of nitrocellulose  paper. Molecules in the gel are forced through the nitrocellulose by drawing  out the liquid with a stack of absorbent towels compressed against the far side  of the nitrocellulose, or by using an electrophoretic process (electrobiotting).  The single-stranded DNA binds to the nitrocellulose at the same position it  had in the gel. After drying at 80°C, which permanently fixes the DNA in  place, the nitrocellulose sheet is moistened with a minimal quantity of solu¬  tion containing a single-stranded DNA or RNA probe that is complementary  in sequence to the DNA of interest. The probe is typically tagged with a    Incubate the  nitrocellulose-  bound DNA with  32 P-labeled  DNA or RNA of a  specific sequence.    FIG. 24-30 The detection of DNAs containing specific base sequences by Southern blotting.   fluorescent group or a radioactive isotope such as 32 P. The moistened nitro¬  cellulose is held at a suitable renaturation temperature for several hours to per¬  mit the probe to hybridize to its target sequence(s), washed to remove the  unbound probe, dried, and then analyzed. If the probe is radioactive, an au¬  toradiogram is made by placing the nitrocellulose for a time over a sheet of  X-ray film. The positions of the molecules that are complementary to the ra¬  dioactive probe are indicated by a blackening of the developed film (alterna¬  tively, a position-sensitive radiation detector can be used). If the probe is  fluorescent, an imaging instrument is used to produce a digital photograph.   Specific RNA sequences can be detected through a variation of the  Southern blot, punningly named a Northern blot, in which the RNA is  immobilized on nitrocellulose paper and probed with a complementary RNA  or DNA probe. A specific protein can be analogously detected in an  immunoblot or Western blot through the use of antibodies directed against  the protein in a procedure similar to that used in an ELISA (Fig. 5-3).   • Which fractionation procedures separate  nucleic acids according to their binding  affinity? According to their size?    Restriction endonucleases make base-specific contacts with palindromic DNA.  Prokaryotic repressors interact with DNA by inserting a helix or a p strand into  the major groove.   Many eukaryotic DNA-binding proteins contain zinc fingers or leucine zipper  structural motifs.    The accessibility of genetic information depends on the ability of proteins to  recognize and interact with DNA in a manner that allows the encoded infor¬  mation to be copied as DNA (in replication) or as RNA (in transcription).  Even the most basic steps of these processes require many proteins that inter¬  act with each other and with the nucleic acids. In addition, organisms regu¬  late the expression of most genes, which requires yet other proteins that act  as repressors or activators of transcription.   Many proteins bind DNA nonspecifically, that is, without regard to the se¬  quence of nucleotides. For example, histones (which are involved in packaging  DNA; Section 24-5A) and certain DNA replication proteins (which must po¬  tentially interact with all the sequences of an organisms genome) bind to DNA  primarily through interactions between protein functional groups and the  sugar-phosphate backbone of DNA. Proteins that recognize specific DNA se¬  quences presumably also bind nonspecifically but loosely to DNA so that they  can scan the polynucleotide chain for their target sequences to which they then  bind specifically and tightly. Most DNA-protein complexes have dissociation  constants ranging from 10 9 to 10 12 M; these values represent binding that  is 10 3 to 10 7 times stronger than nonspecific binding. Sequence-specific DNA-  protein interactions must be extremely precise so that the proteins can exert their  effects at sites selected from among—in the human genome—billions of base  pairs. How do such proteins interact with their target sites on DNA?   Sequence-specific DNA-binding proteins generally do not disrupt the base  pairs of the duplex DNA to which they bind. They do, however, discriminate  among the four base pairs (A* T, T* A, G* C, and C* G) according to the func¬  tional groups of the base pairs that project into DNA’s major and minor grooves.  As can be seen in Fig. 24-2, these groups are more exposed in the major groove  of B-DNA than in its narrower minor groove. Moreover, the major groove  contains more sequence-specific functional groups than does the minor groove  (Fig. 24-1). However, as revealed in a systematic survey of 129 protein-DNA  complexes, only about one-third of the interactions between the binding pro¬  tein and the DNA involve hydrogen bonds, either directly or via intervening  water molecules. Approximately two-thirds of all interactions are van der Waals    FIG. 24-31 X-Ray structure of EcoRI endonuclease in  complex with DNA. The segment of duplex DNA has the  self-complementary sequence TCGC GAATTC GCG (12 bp with an  overhanging T at both 5' ends; the enzyme’s 6-bp target sequence  is underlined) and is drawn as a space-filling model with its  sugar-phosphate chains yellow, its recognition sequence bases cyan,  and its other bases white. The protein is drawn in ribbon form with its    two identical subunits red and blue. The complex is shown (a) with its  DNA helix axis vertical and (b) in end view. The complex’s twofold  axis is horizontal and the DNA’s major groove faces right (toward the  protein) in both views. This was the first known X-ray structure of a  protein-DNA complex. [Based on an X-ray structure by John  Rosenberg, University of Pittsburgh. PDBid 1 ERI.j Q See Kinemage  Exercise 18-1.    contacts. Ionic interactions with backbone phosphate groups also occur. The  complementarity of the binding partners in many cases is augmented by an  “induced fit” phenomenon in which the protein and the nucleic acid change  their conformations for greater stability. In some cases, these changes allow the  binding proteins to interact with other proteins or alter the accessibility of the  DNA to other molecules. Below, we examine some examples of specific  DNA—protein binding. We will encounter many more examples of nucleic  acid—protein interactions in subsequent chapters.   Type II restriction endonucleases rid bacterial cells of foreign DNA by cleaving the  DNA at specific sites that have not yet been methylated by the host s modifica¬  tion methylase (Section 3-4A). Restriction enzymes recognize palindromic DNA  sequences of ~4 to 8 base pairs with such remarkable specificity that a single  base change can reduce their activity by a millionfold. This degree of specificity  is necessary to prevent accidental cleavage of other sites in a DNA sequence.   The X-ray structure of EcoRI endonuclease in complex with a segment of  B-DNA containing the enzyme’s recognition sequence was determined by John  Rosenberg. The DNA binds in the symmetric cleft between the two identical  276-residue subunits of the dimeric enzyme (Fig. 24-31), thereby accounting  for the DNAs palindromic recognition sequence. Protein binding causes the  dihedral angle between the recognition sequences central two base pairs to  open up by ~50° toward the minor groove. These base pairs thereby become  unstacked, but the DNA remains nearly straight due to compensating bends  at the adjacent base pairs. Nevertheless, this unwinds the DNA by 28° and  widens the major groove by 3.5 A at the recognition site. The N-terminal ends  of a pair of parallel helices from each protein subunit are inserted into the    FIG. 24-32 X-Ray structure of EcoRV endonuclease in  complex with DNA. The 10-bp segment of DNA has the  self-complementary sequence GG GATAT CCC (the enzyme’s 6-bp  target sequence is underlined). The DNA and protein are colored as  in Fig. 24-31. (a) The complex as viewed along its twofold axis, facing  the DNA’s major groove. The two symmetry-related protein loops that  overlie the major groove (composed of residues 182-186) are the   only parts of the enzyme that make base-specific contacts with the  DNA. (b) The complex as viewed from the right in Part a (the DNA’s  major groove faces left). Note how the protein kinks the DNA toward  its major groove. [Based on an X-ray structure by Fritz Winkler,  Hoffman-LaRoche Ltd., Basel, Switzerland. PDBid 4RVE.] Q See  Kinemage Exercise 18-2.    widened major groove, where they participate in a hydrogen-bonded network  with the bases of the recognition sequence. The phosphodiester cleavage points  are located two bases from the center of the palindrome (Table 3-2).   Certain other restriction endonucleases, including EcoRV (Fig. 24-32),  also induce kinks in the DNA, in some cases by opening up the minor groove  and compressing the major groove. However, complementary hydrogen bonding  between the nucleotide bases and protein side chain and backbone groups , rather  than the DNA distortion per se, is the primary prerequisite for the formation of  a sequence-specific endonuclease—DNA complex. For example, in the BamHI—  DNA complex, every potential hydrogen bond donor and acceptor in the  major groove of the recognition site takes part in direct or water-mediated  hydrogen bonds with the protein. No other DNA sequence could support this  degree of complementarity with BamHI.   In prokaryotes, the expression of many genes is governed at least in part by re¬  pressors, proteins that bind at or near the gene so as to prevent its transcrip¬  tion (Section 28-2). These repressors often contain ~20-residue polypeptide  segments that form a helix-turn-helix (HTH) motif containing two a helices  that cross at an angle of ~120°. HTH motifs, which are apparently evolution¬  ary related, occur as components of domains that otherwise have widely vary¬  ing structures, although all of them bind DNA. Note that HTH motifs are  structurally stable only when they are components of larger proteins.   Like restriction endonuclease recognition sites, the sequences to which  repressors bind (called operators) exhibit palindromic symmetry or nearly so.  Typically, the repressors are dimeric, although their interactions with DNA may    FIG. 24-33 X-Ray structure of a portion of the 434 phage repressor in complex  with its target DNA. One strand of the 20-bp DNA has the sequence  d(TATACAAGAAAGTTTGTACT). (a) The complex as viewed with the homodimeric protein’s  twofold axis horizontal. The protein is drawn in ribbon form with one of its subunits blue  and the other pink and with their helix-turn-helix (HTH) motifs in darker shades. The DNA  is drawn in stick form with C white, N blue, 0 red, and P orange, and with successive P  atoms in the same chain connected by orange rods, (b) A schematic drawing indicating  how the helix-turn-helix motif, which encompasses helices a2 and a3, interacts with the  DNA. Residues are identified by single-letter codes. Short bars emanating from the  polypeptide chain represent peptide NH groups, dashed lines represent hydrogen bonds,  and numbered circles represent DNA phosphates. The small circle is a water molecule.   [Part a based on an X-ray structure by and Part b courtesy of Aneel Aggarwal, John  Anderson, and Stephen Harrison, Harvard University. PDBid 20R1.] ©See Interactive  Exercise 39 and Kinemage Exercise 19.    not be perfectly symmetrical. Binding involves interactions between the amino  acid side chains extending from the second helix of the HTH motif (the “recog¬  nition” helix) and the bases and sugar-phosphate chains of the DNA. The X-  ray structure of a dimeric repressor from bacteriophage 434 bound to its 20-bp  recognition sequence was determined by Stephen Harrison. The 434 repressor  associates with the DNA in a twofold symmetric manner with a recognition  helix from each subunit bound in successive turns of the DNAs major groove  (Fig. 24-33). The repressor closely conforms to the DNA surface and interacts  with its paired bases and sugar-phosphate chains through elaborate networks  of hydrogen bonds, salt bridges, and van der Waals contacts. In the  repressor-DNA complex, the DNA bends around the protein in an arc of ra¬  dius ~65 A, which compresses the minor groove by ~ 2 . 5 A near its center (be¬  tween the two protein monomers) and widens it by ~2.5 A toward its ends.   The E. coli trp Repressor Binds DNA Indirectly. The E. coli trp repressor reg¬  ulates the transcription of genes required for tryptophan biosynthesis (Section  28-2C). Paul Sigler determined the X-ray structure of the protein in complex  with a DNA containing an 18-bp palindrome (of single-strand sequence TG-  T ACTAGT TA ACTAGTA C, where the trp repressors target sequence is under¬  lined) that closely resembles the trp operator. The homodimeric repressor protein  also has HTH motifs whose recognition helices bind, as expected, in successive  major grooves of the DNA, each in contact with half of the operator sequence  (ACTAGT ; Fig. 24-34). There are numerous hydrogen-bonding contacts be¬  tween the trp repressor and the DNAs nonesterified phosphate oxygens.  Astoundingly, however, there are no direct hydrogen bonds or nonpolar contacts  that can explain the repressors specificity for its operator. Rather, all but one of  the side chain-base hydrogen-bonding interactions are mediated by bridging wa¬  ter molecules. In addition, the operator contains several base pairs that are not  in contact with the repressor but whose mutation nevertheless greatly decreases  repressor binding affinity. This suggests that the operator assumes a sequence-spe¬  cific conformation that makes favorable contacts with the repressor. Other DNA se¬  quences could conceivably assume the same conformation but at too high an  energy cost to form a stable complex with the repressor. This phenomenon, in  which a protein senses the base sequence of DNA through the DNAs backbone  conformation and/or flexibility, is referred to as indirect readout. This finding  puts to rest the notion that proteins recognize nucleic acid sequences exclusively  via particular sets of pairings between amino acid side chains and nucleotide  bases analogous to Watson-Crick base pairing.    The met Repressor Binds DNA via a Two-Stranded p Sheet. Simon  Phillips first determined the X-ray structure of the E. coli met repressor in  the absence of DNA (the met repressor regulates the transcription of genes  involved in methionine biosynthesis). The homodimeric protein lacks an    FIG. 24-34 X-Ray structure of an E. coli trp repressor-operator complex.   The complex is viewed with its molecular twofold axis horizontal. The protein’s two  identical subunits are drawn in ribbon form colored pink and blue with their HTH  motifs more deeply colored. The 18-bp self-complementary DNA is shown in stick  form with C white, N blue, 0 red, P orange, and with successive P atoms in the same  chain connected by orange rods. The trp repressor binds its operator only when  L-tryptophan, drawn in space-filling form with C green, is simultaneously bound. Note  that the protein’s recognition helices bind, as expected, in successive major grooves  of the DNA but extend approximately perpendicular to the DNA helix axis, whereas  those of 434 phage repressor are nearly parallel to the major grooves of its bound  DNA (Fig. 24-33). [Based on an X-ray structure by Paul Sigler, Yale University. PDBid  1TRO.] Q See Interactive Exercise 40.    HTH motif, but model-building studies suggested that the repressor might  bind to its palindromic target DNA via a symmetry-related pair of protrud¬  ing a helices, reminiscent of the way the recognition helices of HTH motifs  interact with DNA. However, the subsequently determined X-ray structure of  the met repressor—operator complex showed that the protein actually binds its  target DNA sequence through a pair of symmetrically related (3 strands  (located on the opposite side of the protein from the protruding a helices)  that form a two-stranded antiparallel (3 sheet that inserts into the DNAs  major groove (Fig. 24-35). The (3 strands make sequence-specific contacts  with the DNA via hydrogen bonding and, probably, indirect readout. This  result indicates that the conclusions of even what appear to be straightforward  model-building studies should be viewed with skepticism. In the case of the met  repressor—operator complex, the model-building study favored the incorrect  model because it could not take into account the small conformational adjust¬  ments that both the protein and the DNA make on binding one another.   FIG. 24-35 X-Ray structure of the E. coli met repressor-SAM-  operator complex. The complex is viewed along its twofold axis of  symmetry. The self-complementary 18-bp DNA is drawn in stick form, and  S-adenosylmethionine (SAM), which must be bound to the repressor for it  to also bind DNA, is shown in space-filling form. DNA C atoms are white,  SAM C green, N blue, 0 red, P orange, and S yellow. Note that the DNA  has four bound repressor subunits: Pairs of subunits (light cyan and  lavender) form symmetric dimers in which each subunit donates one strand  of the two-stranded |3 sheet that is inserted in the DNA’s major groove  (upper left and lower right). Two such dimers pair across the complex’s  twofold axis via their antiparallel N-terminal helices, which contact each  other over the DNA’s minor groove. [Based on an X-ray structure by Simon  Phillips, University of Leeds, U.K. PDBid 1CMA.] Q See Interactive  Exercise 41.   In eukaryotes, genes are selectively expressed in different cell types; this requires  more complicated regulatory machinery than in prokaryotes. Prokaryotic re¬  pressors of known structure either contain an HTH motif or resemble the met  repressor. However, eukaryotic DNA-binding proteins employ a much wider  variety of structural motifs to bind DNA. A number of proteins known as tran¬  scription factors (Section 26-2C) promote the transcription of genes by bind¬  ing to DNA sequences at or near those genes. In this section, we describe a  variety of the DNA-binding motifs in eukaryotic transcription factors.   Zinc Fingers Form Compact DNA-Binding Structures. The first of the pre¬  dominantly eukaryotic DNA-binding motifs, the zinc finger, was discovered  by Klug in transcription factor IIIA (TFIIIA) from Xenopus laevis (an African  clawed toad). The 344-residue TFIIIA contains nine similar, tandemly re¬  peated, ~30-residue modules, each of which contains two invariant Cys  residues and two invariant His residues. Each of these units binds a Zn 2+ ion,  which is tetrahedrally liganded by the Cys and His residues (a Cys 2 —His 2 zinc  finger; Fig. 6-37). In some zinc fingers, the two Zn 2+ -liganding His residues  are replaced by two additional Cys residues, whereas others have six Cys  residues liganding two Zn 2+ ions. Indeed, structural diversity is a hallmark of  zinc finger proteins. In all cases, however, the Zn 2+ ions appear to knit  together relatively small globular domains, thereby eliminating the need for  much larger hydrophobic protein cores (Section 6-4A).   The Cys 2 —His 2 zinc finger contains a two-stranded antiparallel (3 sheet and  one a helix. Three of these motifs are incorporated into a 72-residue segment  of the mouse protein Zif268, whose X-ray structure in complex with a target  DNA was elucidated by Carl Pabo (Fig. 24-36). The three zinc fingers are  arranged as separate domains in a C-shaped structure that fits snugly into the  DNAs major groove. Each zinc finger interacts in a conformationally identical  manner with successive 3-bp segments of the DNA, predominantly through  hydrogen bonds between the zinc fingers a helix and one strand of the DNA.  Each zinc finger specifically hydrogen-bonds to two bases in the major groove.  Interestingly, five of the six associations involve Arg—guanine pairs. In addition to  these sequence-specific interactions, each zinc finger hydrogen-bonds with the  DNAs phosphate groups via conserved Arg and His residues.    FIG. 24-36 X-Ray structure of the  three-zinc finger segment of Zif268 in  complex with DNA. The 10-bp DNA has a  single nucleotide overhang at each end. The  DNA is drawn in paddle form with C magenta,   N blue, 0 red, and P orange and with successive  P atoms connected by orange rods. The protein  is drawn in cartoon form with finger 1 lavender,  finger 2 colored in rainbow order from its  N-terminus (blue) to its C-terminus (red), and  finger 3 dark yellow. The Zn 2+ ions are  represented by cyan spheres, and their liganding  His and Cys side chains are drawn in stick form  with C green, N blue, and S yellow. Note how  the N-terminal end of each zinc finger’s helix  extends into the DNAs major groove to contact  three base pairs. [Based on an X-ray structure  by Carl Pabo, MIT. PDBid 1ZAA.] © See  Interactive Exercise 42.   FIG. 24-37 X-Ray structure of the GAL4 DNA-binding domain  in complex with DNA. (a) A ribbon diagram of the protein’s zinc  finger domain (residues 8-40) colored in rainbow order from its  N-terminus (blue) to its C-terminus (red). The six Cys side chains are  shown in stick form with C green and S yellow, and the two Zn 2+  ions are represented by cyan spheres, (b) The complex of the dimeric  protein with the 19-bp DNA. The DNA is drawn in paddle form with  C green, N blue, 0 red, and P orange and with successive P atoms  connected by orange rods. The protein is shown in ribbon form with    one subunit pink and the other lavender. The views are along the  complex’s twofold axis (left) and turned 90° with the twofold axis  horizontal (right). Note how the C-terminal end of each subunit’s  N-terminal helix extends into the DNA’s major groove. [Based on an  X-ray structure by Ronen Marmorstein and Stephen Harrison, Harvard  University. PDBid 1 D66J © See Interactive Exercise 43.    The Cys 2 —His 2 zinc finger broadly resembles the prokaryotic HTH motif as  well as most other DNA-binding motifs we will encounter (including other types  of zinc fingers). All these DNA-binding motifs provide a platform for inserting  an a helix into the major groove of B-DNA. However, the Cys 2 —His 2 zinc fin¬  gers, unlike other DNA-binding motifs, occur as modules that each contact suc¬  cessive DNA segments (some transcription factors contain over 60 zinc  fingers). Such a modular system can recognize extended asymmetric base sequences.   A binuclear Cys^ zinc finger mediates the DNA binding of the yeast  protein GAL4, a transcriptional activator of several genes that encode galac¬  tose-metabolizing enzymes. Residues 1 to 65 of this 881-residue protein in¬  clude six Cys residues that collectively bind two Zn 2+ ions (Fig. 24-37^).  Each Zn 2+ ion is tetrahedrally coordinated by four Cys residues, with two of  the residues ligating both metal ions. GAL4 binds to its 17-bp target DNA  as a symmetric dimer (Fig. 24-3 7b), although in the absence of DNA it is a  monomer. Each subunit includes a compact zinc finger (residues 8-40) that  binds DNA, an extended linker (residues 41-49), and an a helix (residues  50—64) that assists in GAL4 dimerization. The N-terminal helix of the zinc  finger inserts into the DNAs major groove, making sequence-specific contacts  with a highly conserved CCG sequence at each end of the recognition  sequence. The bound DNA retains its B conformation.   The dimerization helices of GAL4 (center of Fig. 24-37 b) are positioned  over the minor groove of the DNA. The linkers connecting those helices to  the zinc fingers wrap around the DNA, largely following its minor groove.  The two symmetrically related DNA-binding zinc fingers thereby approach   Chapter 24 Nucleic Acid Structure the ma jor groove from opposite sides of the DNA, ~1.5 helical turns apart,   rather than from the same side of the DNA, ~1 turn apart, as do HTH  motifs. The resulting relatively open structure could permit other proteins to  bind simultaneously to the DNA.   Some Transcription Factors Include Leucine Zippers. Segments of certain  eukaryotic transcription factors, such as the yeast protein GCN4, contain a Leu  at every seventh position. We have already seen that a helices with the seven-  residue pseudorepeating sequence ( a-b-c-d-e-f-g) n , in which the a and ^/residues  are hydrophobic, have a hydrophobic strip along one side that allows them to  dimerize as a coiled coil (e.g., a-keratin; Section 6-1C). Steven McKnight sug¬  gested that DNA-binding proteins containing such heptad repeats also form  coiled coils in which the Leu side chains might interdigitate, much like the teeth  of a zipper (Fig. 24-38). In fact, these leucine zippers mediate the dimerization  of certain DNA-binding proteins but are not themselves DNA-binding motifs.   The X-ray structure of the 33-residue polypeptide corresponding to the  leucine zipper of GCN4 was determined by Peter Kim and Thomas Alber.  The first 30 residues, which contain ~3.6 heptad repeats (Fig. 24-38/2), coil  into an ~8-turn a helix that dimerizes as McKnight predicted to form —1/4  turn of a parallel left-handed coiled coil (Fig. 24-38 £). The dimer can be en¬  visioned as a twisted ladder whose sides consist of the helix backbones and  whose rungs are formed by the interacting hydrophobic side chains. The con¬  served Leu residues in the heptad position d, which corresponds to every  second rung, are not interdigitated as McKnight originally suggested, but  instead make side-to-side contacts. The alternate rungs are likewise formed by  the a residues of the heptad repeat (which are mostly Val). These contacts form  an extensive hydrophobic interface between the two helices.   In many leucine zipper—containing proteins, a DNA-binding region that is  rich in basic residues is immediately N-terminal to the leucine zipper, and hence  these proteins are known as basic region leucine zipper (bZIP) proteins. For  example, in GCN4, the C-terminal 56 residues form an extended a helix. The  last 25 residues of two such helices associate as a leucine zipper. The N-termi-  nal portions of the helices smoothly diverge to bind in the major grooves on  opposite sides of the DNA, thereby clamping the DNA in a sort of scissors   FIG. 24-38 The GCN4 leucine zipper motif, (a) A helical wheel  representation of the motif’s two helices as viewed from their  N-termini. The sequences of residues at each position are indicated  by the adjacent column of one-letter codes. Residues that form ion  pairs in the crystal structure are connected by dashed lines. Note that  all residues at positions d and d' are Leu (L), those at positions a and  a' are mostly Val (V), and those at other positions are mostly polar.    [After O’Shea, E.K., Klemm, J.D., Kim, P.S., and Alber, I, Science 254,  540 (1991).] (b) The X-ray structure, in side view, in which the helices  are shown in ribbon form. Side chains are shown in stick form with the  contacting Leu residues at positions d and d' yellow and side chains at  positions a and a' with C green, N blue, and 0 red. [Based on an X-ray  structure by Peter Kim, MIT, and Tom Alber, University of Utah School  of Medicine. PDBid 2ZTA.] Q See Kinemage Exercise 20.   FIG. 24-39 X-Ray structure of the GCN4 bZIP region in  complex with its target DNA viewed with its twofold axis vertical.   The DNA consists of a 19-bp segment with a single nucleotide  overhang at each end and contains the protein’s palindromic (except  for the central base pair) 7-bp target sequence. The DNA is drawn in  paddle form with C magenta, N blue, 0 red, and P orange and with  successive P atoms connected by orange rods. The two identical  GCN4 subunits, shown in ribbon form, each contain a continuous  52-residue a helix. At their C-terminal ends (yellow), the two subunits  associate in a parallel coiled coil (a leucine zipper), and at their basic  regions (green), they smoothly diverge to each engage the DNA in its  major groove at the target sequence. [Based on an X-ray structure by  Stephen Harrison, Harvard University. PDBid 1YSA.] © See  Interactive Exercise 44.   FIG. 24-40 X-Ray structure of Max binding to DNA viewed  with its twofold axis vertical. Residues 22-113 of the 160-residue  transcription factor Max form a complex with a 22-bp DNA containing  the protein’s palindromic 6-bp target sequence. The DNA is drawn in  paddle form with C magenta, N blue, 0 red, and P orange and with  successive P atoms connected by orange rods. The homodimeric  protein is shown in ribbon form. Its N-terminal basic region (green)  forms an a helix that engages its target sequence in the DNA’s major  groove and then merges smoothly with the HI helix (yellow) of the  basic helix-loop-helix (bHLH) motif. Following the loop (red), the  protein’s two H2 helices (purple) of the bHLH motif form a short  parallel four-helix bundle with the two HI helices. Each H2 helix then  merges smoothly with the leucine zipper (Z) motif (cyan) to form a  parallel coiled coil. The protein’s C-terminal ends are pink. [Based on  an X-ray structure by Edward Ziff and Stephen Burley, The Rockefeller  University. PDBid 1AN2.] © See Interactive Exercise 45.    grip (Fig. 24-39). The basic region residues that are conserved in bZIP pro¬  teins make numerous contacts with both the bases and the phosphate oxygens  of their DNA target sequence without distorting its conformation.   Many eukaryotic transcription factors contain a conserved DNA-binding  basic region that is immediately followed by two amphipathic helices con¬  nected by a loop to form a so-called basic helix-loop-helix (bHLH) motif.  The basic region, together with the N-terminal portion of the first (HI)  helix of the bHLH motif, binds in the major groove of its target DNA. The  C-terminal (H2) helix of the bHLH motif mediates the dimerization of the  protein via the formation of a coiled coil. In many proteins, the bHLH  motif is continuous with a leucine zipper (Z) that presumably augments the  dimerization. Thus, as is shown in Fig. 24-40 for the protein Max, the dimeric  bHLH/Z protein grips the DNA in a manner reminiscent of a pair of forceps.  Each basic region contacts specific bases of the DNA as well as phosphate  groups. Side chains of both the loop and the N-terminal end of the H2 helix  also contact DNA phosphate groups.   • Describe the types of interactions  between nucleic acids and proteins.   • Why do restriction endonucleases make  both site-specific and nonspecific contacts  with DNA?   • How does the DNA binding of a zinc  finger-containing transcription factor differ  from that of a prokaryotic repressor?   • Explain why dimeric or multidomain proteins  can bind to DNA with very high specificity.   consists of two sister (identical) chromatids  joined at their centromere (the constricted region  near the center of a chromosome through which  it attaches to the mitotic spindle). Chromosomes  constitute a cell’s largest molecular entities.  [Andrew Syred/Photo Researchers, Inc.]   • Eukaryotic DNA is negatively supercoiled around a core of histone proteins to  form a nucleosome.   Although each base pair of B-DNA contributes only ~3.4 A to its contour  length (the end-to-end length of a stretched-out native molecule), DNA mol¬  ecules are generally enormous (as shown on page 821). Indeed, the 23 chro¬  mosomes of the 3 billion-bp human genome have a total contour length of  almost 1 m. One of the enduring questions of molecular biology is how such  vast quantities of genetic information can be scanned and decoded in a rea¬  sonable time while stored in a small portion of the cells volume.   The elongated shape of duplex DNA (its diameter is only 20 A) and its  relative stiffness make it susceptible to mechanical damage when outside the  protective environment of the cell. For example, a Drosophila chromosome, if  expanded by a factor of 500,000, would have the shape and some of the me¬  chanical properties of a 6-km-long strand of uncooked spaghetti. In fact, the  shear degradation of DNA by stirring, shaking, or pipetting a DNA solution  is a standard laboratory method for preparing DNA fragments.   Prokaryotic genomes typically comprise a single circular DNA molecule.  However, most eukaryotes condense and package their genome in several chro¬  mosomes. Each chromosome is a complex of DNA and protein, a material  known as chromatin, and is a dynamic entity whose appearance varies dra¬  matically with the stage of the cell cycle (the general sequence of events that  occur in the lifetime of a eukaryotic cell; Section 28-4A). For example, chro¬  mosomes assume their most condensed forms only during the metaphase stage  of cell division (Fig. 24-41). During the remainder of the cell cycle, when the  DNA is transcribed and replicated, the chromosomes of most cells become  so highly dispersed that they cannot be distinguished. Yet the DNA of the  chromosomes is still compacted relative to its free B-helix form. Human chro¬  mosomes have contour lengths between 1.5 and 8.4 cm but in their most  condensed state are only 1.3 to 10 (xm long. In this section, we examine how  DNA is packaged in cells to achieve that degree of condensation.   Chromatin is about one-half protein by mass, and most of this protein con¬  sists of histones. To understand how DNA is packaged, we must first exam¬  ine the histone proteins. The five major classes of histones, HI, H2A, H2B,  H3, and H4, all have a large proportion of positively charged residues (Arg  and Lys; Table 24-3). These proteins can therefore bind DNA’s negatively  charged phosphate groups through electrostatic interactions.   Histones Are Highly Conserved. The amino acid sequences of histones H2A,  H2B, H3, and H4 are remarkably conserved. For example, histones H4 from cows  and peas, species that diverged 1.2 billion years ago, differ by only two conserva¬  tive residue changes, which makes this protein among the most evolutionarily con¬  served proteins known (Section 5-4A). Such evolutionary stability implies that the  histones have critical functions to which their structures are so well tuned that they are  all but intolerant to change. The fifth histone, HI, is more variable than the other  histones; we will see below that it also has a somewhat different role.   Histones are subject to posttranslational modifications that include meth-  ylation, acetylation, and phosphorylation of specific Arg, Glu, His, Lys, Ser, Thr,  and Tyr residues. These modifications , many of which are reversible , all decrease the  histones positive charges , thereby significantly altering histone-DNA interactions.  Despite the histones’ great evolutionary stability, their degree of modification  varies enormously with the species, the tissue, and the stage of the cell cycle.  As we will see (Section 28-3A), modification of histones has been linked to    transcriptional activation. The modifications probably create new protein bind¬  ing sites on the histones themselves or on the DNA with which they associate.   The Nucleosome Core Particle Contains a Histone Octamer and 146 bp   of DNA. The first level of chromatin organization was pointed out by Roger  Kornberg in 1974 from several lines of evidence:   1. Chromatin contains roughly equal numbers of histones H2A, H2B,  H3, and H4, and no more than half that number of histone HI.   2. Electron micrographs of chromatin preparations at low ionic strength  reveal ~ 100-A-diameter particles connected by thin strands of appar¬  ently naked DNA, rather like beads on a string (Fig. 24-42).   3. Brief digestion of chromatin by micrococcal nuclease (which  hydrolyzes double-stranded DNA) cleaves the DNA only between the  above particles; apparently the particles protect the DNA closely asso¬  ciated with them from nuclease digestion. Gel electrophoresis indicates  that each particle contains ~200 bp of DNA.   Kornberg proposed that the chromatin particles, which are called nucleo-  somes, consist of the octamer (H2A) 2 (H2B) 2 (H3)2(H4) 2 in association with  ~200 bp of DNA. The fifth histone, HI, was postulated to be associated in  some other manner with the nucleosome (see below).   Micrococcal nuclease initially degrades chromatin to single nucleosomes  in complex with histone HI. Further digestion trims away additional DNA,  releasing histone HI. This leaves the so-called nucleosome core particle,  which consists of a 146-bp strand of DNA associated with the histone oc¬  tamer. A segment of linker DNA, the DNA that is removed by the nucle¬  ase, joins neighboring nucleosomes. Its average length varies between 10 and  50 bp among organisms and tissues.   The X-ray structure of the nucleosome core particle, independently deter¬  mined by Timothy Richmond and Gerard Bunick, reveals a nearly twofold  symmetric complex in which B-DNA is wrapped around the outside of the  histone octamer in 1.65 turns of a left-handed superhelix (Fig. 24-43). This  is the origin of sup ere oiling in eukaryotic DNA. Despite having only weak   FIG. 24-42 Electron micrograph of D.  melanogaster chromatin showing strings of  closely spaced nucleosomes. [Courtesy of  Oscar L Miller, Jr., and Steven McKnight,  University of Virginia.]   FIG. 24-43 X-Ray structure of the chicken nucleosome core  particle, (a) The entire core particle as viewed along its superhelical  axis (left) and rotated 90° about the vertical axis (right). The proteins  of the histone octamer are drawn in ribbon form with H3 blue, H4  green, H2A yellow, and H2B red. The 146-bp DNA is drawn in  paddle form with one strand cyan and the other strand orange. In  both views, the pseudo-twofold axis is vertical and passes through the  DNA center at the top. (b) The half of the nucleosome core particle  that is closest to the viewer in Part a, left. The numbers 0 through 7    arranged about the inside of the DNA superhelix mark the positions  of sequential of double-helical turns. The four histones that are drawn  in their entirety are primarily associated with this DNA segment,  whereas only fragments of the H3 from the other half of the particle  are shown. The two 4-helix bundles shown are labeled H3' H3 and  H4 H2B. [Based on an X-ray structure by Gerard Bunick, University of  Tennessee and Oak Ridge National Laboratory, Oak Ridge, Tennessee.  PDBid 1EQZJ    FIG. 24-44 X-Ray structure of half of a histone octamer within the nucleosome  core particle. Those portions of H2A, H2B, H3, and H4 that form the histone folds are  yellow, red, blue, and green, respectively, with their N- and C-terminal tails colored in lighter  shades. [Based on an X-ray structure by Gerard Bunick, University of Tennessee and Oak  Ridge National Laboratory, Oak Ridge, Tennessee. PDBid 1 EQZJ    sequence similarity, all four types of histones share a similar ~70-residue  fold in which a long central helix is flanked on each side by a loop and  a shorter helix (Fig. 24-44). Pairs of histones interdigitate in a sort  of “molecular handshake” to form the crescent-shaped heterodimers  H2A-H2B and H3-H4, each of which binds 2.5 turns of duplex  DNA that curves around it in a 140° bend. The H3-H4 pairs  interact, via a bundle of four helices from the two H3 histones,  to form an (H3-H4) 2 tetramer with which each H2A-H2B  pair interacts, via a similar four-helix bundle between H2B and  H4, to form the histone octamer (Fig. 24-43 £).   The histones bind exclusively to the inner face of the DNA,  primarily via its sugar-phosphate backbones, through hydrogen  bonds, salt bridges, and helix dipoles (their positive N-terminal  ends), all interacting with phosphate oxygens, as well as through  hydrophobic interactions with the deoxyribose rings. There are few contacts be¬  tween the histones and the bases, in accord with the nucleosomes lack of se¬  quence specificity. However, an Arg side chain is inserted into the DNAs minor  groove at each of the 14 positions at which it faces the histone octamer. The  DNA superhelix has a radius of 42 A and a pitch (rise per turn) of 26 A. The  DNA does not follow a uniform superhelical path but, rather, is bent fairly  sharply at several locations due to outward bulges of the histone core. Moreover,  the DNA double helix exhibits considerable conformational variation along its  length such that its twist, for example, varies from 7.5 to 15.2 bp/turn with  an average value of 10.4 bp/turn (versus 10.5 bp/turn for DNA in solution).  Approximately 75% of the DNA surface is accessible to solvent and hence ap¬  pears to be available for interactions with DNA-binding proteins.   Linker Histones Bring Nucleosomes Together. In the micrococcal nuclease  digestion of chromatin fibers, the ~200-bp DNA is first degraded to 166 bp.  Then there is a pause before histone HI is released and the DNA is further  shortened to 146 bp. Since the 146-bp DNA of the core particle makes 1.65  superhelical turns, the 166-bp intermediate should make nearly two full  superhelical turns, which would bring its two ends close together. Klug has pro¬  posed that histone HI binds to nucleosomal DNA at this point, where the  DNA segments enter and leave the core particle (Fig. 24-45 tf). Chromatin    FIGURE 24-45 Binding of histone  HI to the nucleosome. (a) Model of  histone HI binding to the DNA of the  166-bp nucleosome. The two complete  superhelical turns of the DNA (blue)  enable HI (orange cylinder) to bind to the  DNAs two ends and its middle. The  histone octamer is represented by the  yellow central spheroid, (b) Electron  micrograph of HI-containing chromatin.  [Courtesy of Fritz Thoma, Eidgenossiche  Technische Hochschule, Zurich, Switzerland.]    fibers containing HI have closely spaced nucleosomes (Fig. 24-45 £), consistent  with DNA entering and exiting the nucleosome on the same side. In HI-de¬  pleted chromatin (e.g., Fig. 24-42), the nucleosomes are more dispersed, sug¬  gesting that linker histones play a role in condensing chromatin fibers and  regulating the access of other proteins to the DNA.   Winding the DNA helix around a nucleosome reduces its contour length  sevenfold (the 560-A length of 166 bp is compressed to an ~80-A-high  supercoil). Under physiological conditions, chromatin condenses further  by folding in a zigzag fashion to form a helical filament with a diameter of  ~30 nm. A model of this 30-nm fiber based on the X-ray structure of a  tetranucleosome indicates that sequential nucleosomes are rotated ~71° to each  other. The nucleosomes follow a zigzag path so that all the odd-numbered  nucleosomes, and all the even-numbered nucleosomes, stack next to each other  (Fig. 24-46). The ~250-A-diameter structure, in which the two stacks wind  around each other in two left-handed helices, has approximately 18.9 nucleo¬  somes per turn, covering a distance of about 316 A along the fiber axis.   The 30-nm fiber is stabilized by interactions involving H2A-H2B dimers  and histone H4. As the X-ray structure of the nucleosome shows (Fig. 24-  43a), H4s positively charged N-terminal tail can bind to an intensely nega¬  tively charged region on an exposed face of the H2A—H2B dimer in a  neighboring nucleosome. Acetylation of H4s N-terminal tail reduces its pos¬  itive charge and would therefore weaken this interaction. The highly basic tails  of H2B and H3 also extend out from the nucleosome core (Fig. 24-43 a). Since  histone tails contain numerous acetylation sites, higher-order chromatin struc¬  ture is likely to be controlled, at least in part, by the acetylation of certain  histone residues but not others.   FIG. 24-46 Model of the 30-nm chromatin  fiber consisting of 22 nucleosomes in which  the fiber axis is vertical. This model is based  on the X-ray structure of a four-nucleosome  segment in which each nucleosome consists of  147 bp of DNA plus a histone octamer.  Successive pairs of nucleosomes are colored  cyan and pink and the DNA is represented by  wider ribbons than those of the polypeptide  chains. The first (N1), second (N2), third (N3),  and seventh (N7) nucleosome in the linear DNA  sequence are indicated. The inset is a schematic  diagram of the DNA’s connectivity. Note how this  model consists of a left-handed two-start helix  of stacked nucleosomes. [Courtesy of Daniela  Rhodes, MRC Laboratory of Molecular Biology,  Cambridge, U.K. The X-ray structure on which  this model is based was determined by Timothy  Richmond, ETH, Zurich, Switzerland. PDBid  1ZBBJ   FIG. 24-47 Electron micrographs of a histone-depleted magnification reveals that the DNA is attached to the scaffold in   metaphase human chromosome, (a) The fibrous central protein loops. [Courtesy of Ulrich Laemmli, University of Geneva, Switzerland.]    extensive halo of DNA (Fig. 24-47 a). The strands of DNA that can be followed  form loops that enter and exit the scaffold at nearly the same point (Fig. 24-47£).  Most of the loops are 15 to 30 [im long (which corresponds to 45-90 kb), so  when condensed as 30-nm filaments, they would be ~0.6 fxm long. Electron  micrographs of metaphase chromosomes in cross section, such as Fig. 24-48,  strongly suggest that the loops of chromatin are radially arranged and would each  contribute 0.3 |Jim to the diameter of the chromosome (a fiber must double back  on itself to form a loop). These loops, along with the 0.4-pim width of the  scaffold, would account for the 1.0-fxm diameter of the metaphase chromosome.   Note the mass of chromatin fibers  radially projecting from the central  scaffold. [Courtesy of Ulrich Laemmli,  University of Geneva, Switzerland.]   FIG. 24-49 Model diagramming the  various levels of metaphase chromatin  organization. B-DNA winds around histone  octamers to form nucleosomes; these fold into a  30-nm fiber. Loops of chromatin are attached to  a protein scaffold to form the metaphase  chromosome.    A typical human chromosome, which contains — 140 million bp and is  about 6 \xm long, would therefore have —2000 radial loops. Presumably,  nonhistone proteins, which constitute —10% of the chromosomal proteins,  must be involved in organizing the DNA in these higher-order structures  (Fig. 24-49) and managing the interconversion of highly condensed  metaphase chromosomes with the more dispersed DNA present during the  rest of the cell cycle.   In prokaryotes, DNA is also packaged through its association with highly  basic proteins that functionally resemble histones. Nucleosome-like particles  condense to form large loops that are attached to a protein scaffold, yielding  a relatively compact chromosome.   • How do histones differ from the  DNA-binding proteins described in the  preceding section?   • Describe the levels of DNA packaging in  eukaryotic cells, from B-DNA to a  metaphase chromosome.   • The most common form of DNA is B-DNA, which is a right-  handed double helix containing A • T and G • C base pairs of  similar geometry. The A-DNA helix, which also occurs in double-  stranded RNA, is wider and flatter than the B-DNA helix. The  left-handed Z-DNA helix may occur in sequences of alternating  purines and pyrimidines.   • The flexibility of nucleotides in nucleic acids is constrained by the  allowed rotation angles around the glycosidic bond, the puckering    of the ribose ring, and the torsion angles of the sugar-phosphate  backbone.   • The linking number (Z) of a covalently closed circular DNA is  topologically invariant. Consequently, any change in the twist ( T)  of a circular duplex must be balanced by an equal and opposite  change in its writhing number (W), which indicates its degree of  supercoiling.   • Naturally occurring DNA is negatively supercoiled (underwound).  Topoisomerases relax supercoils by cleaving one or both strands of   the DNA, passing the DNA through the break (type IA and II  topoisomerases) or allowing controlled rotation of the uncleaved  strand (type IB topoisomerase), and resealing the broken strand(s).   • The structures of nucleic acids are stabilized by Watson-Crick base  pairing, by hydrophobic interactions between stacked base pairs,  and by divalent cations that shield adjacent phosphate groups.   • Nucleic acids can be denatured by increasing the temperature  above their T m and renatured by lowering the temperature to  ~25°C below their T m .   • RNA molecules assume a variety of structures consisting of  double-stranded stems and single-stranded loops. Some RNAs  have catalytic activity.   • Nucleic acids are fractionated by methods similar to those used to  fractionate proteins, including solubilization, affinity chromatog¬  raphy, and electrophoresis.   • Sequence-specific DNA-binding proteins interact primarily with  bases in the major groove and with phosphate groups through  direct and indirect hydrogen bonds, van der Waals interactions,  and ionic interactions. The conformations of both the protein and  the DNA may change on binding.   • Common structural motifs in DNA-binding proteins include the  helix—turn—helix (HTH) motif in prokaryotic repressors, and zinc  fingers, leucine zippers, and basic helix—loop—helix (bHLH)  motifs in eukaryotic transcription factors.   • The DNA of eukaryotic chromatin winds around histone oc-  tamers to form nucleosome core particles that further condense in  the presence of linker histones. Additional condensation is accom¬  plished by folding chromatin into 30-nm-diameter filaments,  which are then attached in loops to a fibrous protein scaffold to  form a condensed (metaphase) chromosome.    Southern blotting 848  repressor 851  HTH motif 851  operator 851  indirect readout 852  transcription factor 854  zinc finger 854  heptad repeat 856  leucine zipper 856  bHLH motif 857    contour length 858  shear degradation 858  chromosome 858  chromatin 858  histone 858  nucleosome 859  nucleosome core particle  linker DNA 859  30-nm fiber 861    1. Unusual bases and non-Watson-Crick base pairs frequently  appear in tRNA molecules. Which base is most likely to pair with  hypoxanthine (Section 23-1)? Draw this base pair.   2. Draw the structure of a G • U base pair.   3. Amino acid residues in proteins are each specified by three  contiguous bases. What is the contour length of a segment of  B-DNA that encodes a 50-kD protein?   4. Calculate the contour length for the gene in in Problem 3 if it  assumed an A-DNA conformation.   3. A standard A*T base pair is shown in Fig. 24-1. Under certain  conditions, an adenine residue in B-DNA can transiently adopt  the syn conformation. Draw the resulting A • T base pair. (Hint:  It has two hydrogen bonds.) Is the helix diameter larger or  smaller in this alternate arrangement?   6. A standard G • C base pair is shown in Fig. 24-1. Under certain  conditions, the guanine residue can transiently adopt the syn  conformation. Draw the resulting G • C base pair, which has two  hydrogen bonds.   7. The ends of eukaryotic chromosomes terminate in a G-rich sin¬  gle-stranded overhang that can fold up on itself to form a four-  stranded structure. In this structure, four guanine residues assume    a hydrogen-bonded planar arrangement with an overall geome¬  try that can be represented as    Draw the complete structure of this “G quartet,” including the  hydrogen bonds between the purine bases.   8. Show schematically how a single strand of four repeating  TTAGGG sequences can fold to generate a structure with three  stacked G quartets linked by TTA loops.    9. Compounds known as peptide nucleic acids (PNAs) have been  developed as nucleic acid-binding probes. A PNA molecule has  a polypeptide-like backbone with purine and pyrimidine bases  attached as side chains. Draw the PNA backbone resulting from  amide bond formation between two molecules of N-(2-  aminoethyl) glycine.   10. The asterisk in the structure shown in Problem 9 indicates the  site where the base is attached via a two-carbon linker. Compare  the overall backbone length for a PNA and a DNA molecule and  explain why the two molecules can form base-paired duplexes.   11. Which of the DNAs shown in Figure 24-8 would move fastest  during agarose gel electrophoresis?   12. You have discovered an enzyme secreted by a particularly viru¬  lent bacterium that cleaves the C2'-C3' bond in the deoxyribose  residues of duplex DNA. What is the effect of this enzyme on  supercoiled DNA?   13. When the helix axis of a closed circular duplex DNA of 2310 bp  is constrained to lie in a plane, the DNA has a twist ( T) of 207.  When released, the DNA takes up its normal twist of 10.3 bp  per turn. Indicate the values of the linking number (Z), writhing  number ( W ), and twist for both the constrained and uncon¬  strained conformational states of this DNA circle. What is the  superhelix density, cr = Wl T, of both the constrained and  unconstrained DNA circles?   14. A closed circular duplex DNA has a 100-bp segment of alternat¬  ing C and G residues. On transfer to a high salt solution, the  segment undergoes a transition from the B conformation to the  Z conformation. What is the change in its linking number,  writhing number, and twist?   13. How is the melting curve of duplex DNA affected by decreasing  the ionic strength of the solution?   16. How is the melting curve of duplex DNA affected by adding a  small amount of ethanol?   17. Compare the melting temperature of a 1-kb segment of DNA  containing 20% A residues to that of a 1-kb segment containing  30% A residues under the same conditions.   18. The melting curve for the polyribonucleotide poly(A) is shown  below, (a) Explain why absorbance increases with increasing tem¬  perature. (b) Why does the shape of the curve differ from the  one shown in Fig. 24-22?   19. For the RNA sequence AUUGGCAUCCGAUAA, draw the  secondary structure that maximizes its base pairing.   20. In addition to the standard base-paired helical structures (e.g.,  Fig. 24-2), DNA can form X-shaped hairpin structures called  cruciforms in which most bases are involved in Watson-Crick  pairs. Such structures tend to occur at sequences with inverted  repeats. Draw the cruciform structure formed by the DNA  sequence TCAAGTCCACGGTGGACTTGC.   21. E. coli ribosomes contain three RNA molecules named for their  sedimentation behavior: 5S, 16S, and 23S. Draw a diagram  showing the approximate positions of the three RNA species  following electrophoresis.   22. Mouse genomic DNA is treated with a restriction endonuclease  and electrophoresed in an agarose gel. A radioactive probe made  from the human gene rxr-1 is used to perform a Southern blot.  The experiment was repeated three times. Explain the results of  these repeated experiments:   Experiment 1. The autoradiogram shows a large smudge at a  position corresponding to the top of the lane in which the mouse  DNA was electrophoresed.   Experiment 2. The autoradiogram shows a smudge over the  entire lane containing the mouse DNA.   23. What is the probability that the palindromic symmetry of the trp  repressor target DNA sequence (Section 24-4B) is merely acci¬  dental?   24. The E. coli genome contains approximately 4639 kb. (a) How  many copies of the 6-bp recognition sequence for the trp re¬  pressor would be expected to occur in the E. coli chromosome?  (b) Explain why it is advantageous for the trp repressor to be a  dimer that recognizes two adjacent 6-bp sequences.   25. Protamines are small proteins that take the place of histones dur¬  ing spermatogenesis in vertebrates. What amino acids are likely  to be abundant in protamines?   26. The enzyme ornithine decarboxylase generates a product that has  been proposed to play a role in stabilizing compacted DNA.  Draw the structure of the ornithine decarboxylase reaction prod¬  uct and explain how it interacts with DNA.   27. For a linear B-DNA molecule of 50,000 kb, calculate (a) the con¬  tour length and (b) the length of the DNA as packaged in nu-  cleosomes with linker histones present.   28. Calculate the length of the 50,000-bp DNA in a 30-nm fiber.   29. Most protein-coding genes are present in just one copy per  genome. Explain why eukaryotic genomes typically contain  dozens of copies of histone genes.   30. Eukaryotic DNA is typically covalently modified by methylation  of certain nucleotide bases. How would the addition of methyl  groups affect the packaging of DNA in nucleosomes?    How many histone variants oc¬  cur in human cells? How do they differ from each other? How does  the histone variant CENP-A relate to chromosomal structure? Which  histone variant is associated with X-chromosome inactivation?   Focus concept: Understanding the mechanism of action of an  enzyme can lead to the construction of hyperactive variant enzymes  with a greater catalytic efficiency than the wild-type enzyme.   Arnott, S., DNA polymorphism and the early history of the double  helix, Trends Biol. Sci. 31, 349-354 (2006).   Doudna, J.A. and Cech, T.R., The chemical repertoire of natural  ribozymes, Nature 418, 222-228 (2002). [Describes some of the  major ribozymes and how they relate to the RNA world.]   Rich, A., The double helix: a tale of two puckers, Nature Struct. Biol. 10,  247-249 (2003). [A brief history of DNA and RNA structural  studies.]   Snustad, D.P. and Simmons, M.J., Principles of Genetics, 5th ed., Wiley  (2009). [This and other genetics textbooks review DNA structure  and function.]   The double helix—50 years, Nature 421, 395-453 (2003). [A supple¬  ment containing a series of articles on the historical, cultural, and  scientific influences of the DNA double helix on the fiftieth anniver¬  sary of its discovery.]   Deweese, J.E., Osheroff, M.A., and Osheroff, N., DNA topology and  topoisomerases, Biochem. Mol. Biol. Educ. 37, 2-10 (2009).   Dong, K.C. and Berger, J.M., Structure and function of DNA topo¬  isomerases, in Rice, P.A. and Correll, C.C. (Eds.), Protein—Nucleic  Acid Interactions, RSC Publishing (2008).   Wang, J.C., Cellular roles of DNA topoisomerases: a molecular perspec¬  tive, Nature Rev. Mol. Cell Biol. 3, 430-440 (2002).   Aggarwal, A.K., Structure and function of restriction endonucleases, Curr.  Opin. Struct. Biol. 5, 11-19 (1995).   Berg, J.M. and Shi, Y., The galvanization of biology: a growing appreci¬  ation of the roles of zinc, Science 271, 1081-1085 (1996). [Summa¬  rizes different types of zinc fingers and discusses why zinc is suitable  for stabilizing small protein domains.]   Ellenberger, T.E., Getting a grip on DNA recognition: structures of the  basic region leucine zipper, and the basic region helix-loop-helix  DNA-binding domains, Curr. Opin. Struct. Biol. 4, 12-21 (1994).    Luscombe, N.M., Austin, S.E., Berman, H.M., and Thornton, J.M., An  overview of the structures of protein-DNA complexes, Genome Biol. 1,  reviews 001.1-001.37 (2000). [This review, which contains many  molecular models, is available online at http://genomebiology.com/  2000/1 /1 / reviews/001.]   Marmorstein, R., Carey, M., Ptashne, M., and Harrison, S.C., DNA  recognition by GAL4: structure of a protein-DNA complex, Nature  356, 408-414 (1992).   Sarai, A. and Kono, H., Protein-DNA recognition patterns and predic¬  tions, Annu. Rev. Biophys. Biomol. Struct. 34, 379-398 (2005).   Somers, W.S. and Phillips, S.E.V., Crystal structure of the met  repressor-operator complex at 2.8 A resolution reveals DNA recog¬  nition by (3-strands, Nature 359 , 387-393 (1992).   Andrews, A.J. and Luger, K., Nucleosome structure(s) and stability:  variations on a theme, Annu. Rev. Biophys. 40, 99-117 (2011).   Gruss, C. and Knippers, R., Structure of replicating chromatin, Prog.  Nucleic Acid Res. Mol. Biol. 52, 337-365 (1996).   Kornberg, R.D. and Lorch, Y., Chromatin rules, Nature Struct. Mol. Biol.  14, 986-988 (2007).   Luger, K., Mader, A.W., Richmond, R.K., Sargent, D.E, and Richmond,  T.J., Crystal structure of the nucleosome core particle at 2.8 A res¬  olution, Nature 389, 251-260 (1997); and Harp, J.M., Hanson,  B.L., Timm, D.E., and Bunick, G.J., Asymmetries in the nucleo¬  some core particle at 2.5 A resolution, Acta Cryst. D56, 1513-1534  ( 2000 ).   Schalch, T., Duda, S., Sargent, D.E, and Richmond, T.J., X-Ray struc¬  ture of a tetranucleosome and its implications for the chromatin  fibre, Nature 436, 138-141 (2005).   Widom, J., Structure, dynamics, and function of chromatin in vitro,  Annu. Rev. Biophys. Biomol. Struct. 27, 285-327 (1998).    In these human nuclei, fluorescent spots mark the locations of DNA polymerase, many  copies of which are required to replicate the enormous chromosomes quickly and  efficiently. [Courtesy Catherine Green, University of Cambridge, Cambridge, UK.]    Watson and Cricks seminal paper describing the DNA double helix ended  with the statement: “It has not escaped our notice that the specific pairing we  have postulated immediately suggests a possible copying mechanism for the  genetic material.” As they predicted, when DNA replicates, each polynu¬  cleotide strand acts as a template for the formation of a complementary strand  through base pairing interactions. The two strands of the parent molecule must  therefore separate so that a complementary daughter strand can be enzymati¬  cally synthesized on the surface of each parent strand. This results in two mol¬  ecules of duplex DNA, each consisting of one polynucleotide strand from the  parent molecule and a newly synthesized complementary strand (Fig. 3-11).  Such a mode of replication is termed semiconservative (in conservative repli¬  cation, the parental DNA would remain intact and both strands of the daugh¬  ter duplex would be newly synthesized).   The semiconservative nature of DNA replication was elegantly demon¬  strated in 1958 by Matthew Meselson and Franklin Stahl. The density of DNA  was increased by labeling it with 15 N, a heavy isotope of nitrogen ( l4 N is  the naturally abundant isotope). This was accomplished by growing E. coli  in a medium that contained 15 NH 4 C1 as its only nitrogen source. The  labeled bacteria were then abruptly transferred to a l4 N-containing medium  and the density of their DNA was monitored over several generations by  ultracentrifugation.   Meselson and Stahl found that after one generation (one doubling of the  cell population), all the DNA had a density exactly halfway between the den¬  sities of fully 15 N-labeled DNA and unlabeled DNA. This DNA must there¬  fore contain equal amounts of l4 N and 15 N as is expected after one generation   C The Leading and Lagging Strands Are  Synthesized Simultaneously  D Replication Terminates at Specific Sites  E DNA Is Replicated with High Fidelity   A Eukaryotes Use Several DNA Polymerases  B Eukaryotic DNA Is Replicated from  Multiple Origins   A Some Damage Can Be Directly Reversed  B Base Excision Repair Requires a Glycosylase  C Nucleotide Excision Repair Removes a Segment  of a DNA Strand   D Mismatch Repair Corrects Replication Errors  E Some DNA Repair Mechanisms  Introduce Errors   B DNA Can Be Repaired by Recombination  C Transposition Rearranges Segments of DNA    FIG. 25-1 The Meselson and Stahl experiment. Parental ( 15 N-labeled or “heavy”)  DNA is replicated semiconservatively so that in the first generation, DNA molecules contain  one parental (blue) strand and one newly synthesized (red) strand. In succeeding generations,  the proportion of 14 N-labeled (“light”) strands increases, but hybrid molecules containing  one heavy and one light strand persist. Q See the Animated Figures.    Draw a diagram of the DNA molecules that would be produced by conservative  replication.   of semiconservative replication (Fig. 25-1). Conservative DNA replication, in  contrast, would preserve the fully 15 N-labeled parental strand and generate  an equal amount of unlabeled DNA. After two generations, half of the DNA  molecules were unlabeled and the remainder were l4 N— 15 N hybrids. In suc¬  ceeding generations, the amount of unlabeled DNA increased relative to the  amount of hybrid DNA although the hybrid never totally disappeared. This  is in accord with semiconservative replication but at odds with conservative  replication, in which hybrid DNA never forms.   The details of DNA replication, including the unwinding of the parental  strands and the assembly of complementary strands from nucleoside triphos¬  phates, have emerged gradually since 1958. DNA replication is far more com¬  plex than the overall chemistry of the process might suggest, in large part  because replication must be extremely accurate in order to preserve the integrity  of the genome from generation to generation. In this chapter, we examine the  protein assemblies that mediate DNA replication in prokaryotes and eukary¬  otes. We also discuss the mechanisms for ensuring fidelity during replication  and for correcting polymerization errors and other types of DNA damage.    • DNA polymerase requires a template and primers to synthesize DNA.    DNA is replicated by enzymes known as DNA-directed DNA polymerases  or simply DNA polymerases. These enzymes use single-stranded DNA  (ssDNA) as templates on which to catalyze the synthesis of the complemen¬  tary strand from the appropriate deoxynucleoside triphosphates (Fig. 25-2).  The reaction occurs through the nucleophilic attack of the growing DNA  chains 3 r -OH group on the a-phosphoryl of an incoming nucleoside triphos¬  phate. The otherwise reversible reaction is driven by the subsequent hydrolysis    FIG. 25-2 Action of DNA  polymerases. These enzymes assemble  incoming deoxynucleoside triphosphates  on single-stranded DNA templates. The  OH group at the 3' end of a growing  polynucleotide strand is a nucleophile that  attacks the a-phosphate group of an  incoming dNTP that base-pairs with the  template DNA strand, and hence the  growing strand is elongated in its 5' —> 3'  direction. Although formation of a new  phosphodiester bond is reversible, the  subsequent hydrolysis of the PP ) product  makes the overall reaction irreversible.   By B 2 ' B 3 ' B 4 ' B 5 '   FIG. 25-3 An autoradiogram and its interpretive drawing of a replicating E. coli  chromosome. The bacterium had been grown in a medium containing [ 3 H]thymidine,  thereby labeling the subsequently synthesized DNA so that it appears as a line of dark  grains in the photographic emulsion (red lines in the drawing). The size of the replication  eye indicates that this circular chromosome is about one-eighth duplicated. The arrows  indicate the direction of movement of the replication forks. [Courtesy of John Cairns, Cold  Spring Harbor Laboratory.]    of the eliminated PP Z . The incoming nucleotides are selected by their ability  to form Watson—Crick base pairs with the template DNA so that the newly  synthesized DNA strand forms a double helix with the template strand. Nearly  all known DNA polymerases can add a nucleotide only to the free 3 r -OH  group of a base-paired polynucleotide so that DNA chains are extended only in  the 5' —> 3' direction.    DNA Replication Occurs at Replication Forks. John Cairns observed DNA  replication through the autoradiography of chromosomes from E. coli grown  in a medium containing [ 3 H] thymidine. These circular chromosomes contain  replication “eyes” or “bubbles” (Fig. 25-3), called 0 structures (after their  resemblance to the Greek letter theta), that form when the two parental strands  of DNA separate to allow the synthesis of their complementary daughter  strands. DNA replication involving 0 structures is known as 0 replication.   The branch points in a replication eye, at which DNA synthesis occurs,  are called replication forks. Autoradiographic studies have demonstrated that  0 replication is almost always bidirectional. In other words, DNA synthesis  proceeds in both directions from the point where replication is initiated, so that  the replication forks move apart.    Replication Is Semidiscontinuous. The low-resolution images provided by  autoradiograms such as Fig. 25-3 suggest that duplex DNAs two antiparallel  strands are simultaneously replicated at an advancing replication fork. Yet since  DNA polymerases extend DNA strands only in the 5' —> 3' direction, how can  they copy the parent strand that extends in the 5' —> 3' direction  past the replication fork? This question was answered in 1968 by  Reiji Okazaki through the following experiment. If a growing E.  coli culture is pulse labeled for 30 s with [ 3 H]thymidine before  the DNA is isolated, much of the radioactive and hence newly  synthesized DNA consists of 1000- to 2000-nucleotide (nt) frag¬  ments (in eukaryotes, these so-called Okazaki fragments are  100-200 nt long). When the cells are transferred to an unlabeled  medium after the [ 3 H] thymidine pulse, the size of the labeled frag¬  ments increases over time. The Okazaki fragments must therefore  become covalently incorporated into larger DNA molecules.   Okazaki interpreted his experimental results in terms of the semidiscon¬  tinuous replication model (Fig. 25-4). The two parent strands are replicated  in different ways. The newly synthesized DNA strand that extends 5' —> 3' in  the direction of replication fork movement, the leading strand, is continu¬  ously synthesized in its 5' —> 3' direction as the replication fork advances. The  other new strand, the lagging strand, is also synthesized in its 5' —> 3' direc¬  tion. However, it can only be made discontinuously, as Okazaki fragments, as   FIG. 25-4 Semidiscontinuous DNA  replication. Both daughter strands (leading strand  red, lagging strand blue) are synthesized in their  5' —> 3' direction. The leading strand is  synthesized continuously, whereas the lagging  strand is synthesized discontinuously. The lagging  strand segments are known as Okazaki fragments.    FIG. 25-5 Priming of DNA synthesis by  short RNA segments. Each Okazaki fragment  consists of an RNA primer (green) that has been  extended by DNA polymerase. The RNA primer  is later removed.   single-stranded parental DNA becomes newly exposed at the replication fork.  The Okazaki fragments are later covalently joined together by the enzyme   • Explain why DNA polymerase requires a  template and a primer.    DNA Synthesis Extends RNA Primers. Given that DNA polymerases require  a free 3'-OH group to extend a DNA chain, how is DNA synthesis initiated?  Careful analysis of Okazaki fragments revealed that their 5' ends consist of  RNA segments of 1 to 60 nt (the length is species dependent) that are com¬  plementary to the template DNA chain (Fig. 25-5). In E. coli , these RNA  primers are synthesized by the enzyme primase. Multiple primers are required  for lagging strand synthesis, but only one primer is required to initiate syn¬  thesis of the leading strand. Mature DNA, however, does not contain RNA.  The RNA primers are eventually replaced with DNA.    • DNA polymerases include 3' —> 5' exonuclease activity and, in the case of Pol I,   • DNA replication requires additional proteins to separate DNA strands, unwind the  helix, bind to single-stranded DNA, and synthesize RNA primers.   • A sliding clamp is loaded onto the template DNA to enhance the processivity of  DNA polymerase.   • Complete replication of DNA requires DNA ligase, a termination factor, and  topoisomerases.   • Balanced nucleotide levels, the polymerase mechanism, its proofreading mecha¬  nism, and the activities of DNA repair enzymes contribute to the accuracy of DNA  replication.    DNA replication involves a great variety of enzymes in addition to those men¬  tioned above. Many of the required enzymes were first isolated from prokary¬  otes and are therefore better understood than their eukaryotic counterparts.  Accordingly, we begin with a detailed consideration of prokaryotic DNA  replication. Replication in eukaryotes is discussed in Section 25-3.   In 1957, Arthur Kornberg discovered an E. coli enzyme that catalyzes the syn¬  thesis of DNA, based on its ability to incorporate the radioactive label from    Kornberg (1918-2007) Like a number  of his contemporaries during the Great  Depression, Arthur Kornberg entered  medical school because of the lack of  jobs in teaching or industry. He  subsequently served in the U.S. Coast  Guard as a ship’s doctor but quickly  realized that he was better suited to the  laboratory than the sea. When he began  his work at the National Institutes of Health in 1942, the classic  studies of nutrition and vitamins were giving way to the new  science of enzymology. After purifying aconitase from rat hearts,  Kornberg stated that he found enzymes intoxicating. He likened  purifying an enzyme to ascending an uncharted mountain, with the  reward being the view from the top and the satisfaction of being  the first one there.   Kornberg spent 6 months at the Cori laboratory (Box 16-1)  and in 1948 began working on enzymes involved in the synthesis  of nucleotide cofactors. He found that the enzyme nucleotide  pyrophosphorylase catalyzed a reaction in which a nucleotide was  incorporated into a coenzyme:   Next, Kornberg began to search for an enzyme that could  assemble many nucleotides to make a nucleic acid chain. Some  people have mistakenly assumed that Kornberg was inspired to  look for DNA polymerase by the publication of the Watson-Crick  model of DNA in 1953. In fact, Kornberg was following his  instincts as an enzymologist, and his curiosity stemmed from his  familiarity with other enzymes, including glycogen phosphorylase,  which was capable of synthesizing the polymer glycogen in vitro.  Kornberg had also experimented with enzymes involved in the  synthesis of phospholipids, which—like DNA—contain  phosphodiester bonds (he abandoned this line of investigation  because he didn’t like working with “greasy” molecules).   To purify a DNA-synthesizing enzyme, Kornberg started with  extracts of fast-growing E coli cells, which had replaced slower  growing yeast cells as a laboratory subject. He added radioactive  thymidine to the extracts and measured the production of  radioactive DNA. Disappointingly, the incorporation of thymidine    was extremely low, but thymidine phosphate (TMP) worked better,  and thymidine triphosphate (TTP) better still. Kornberg also  discovered that the amount of newly synthesized DNA increased  when a small amount of DNA was included in the reaction mixture.  This was not entirely unexpected, as Kornberg already knew that a  small amount of glycogen could serve as a primer for additional  glycogen synthesis. At first, however, Kornberg believed that the  DNA added to his reaction mixture acted as substrates for  nucleases that were present in cell extracts and thereby protected  the newly synthesized radioactive DNA from degradation. Later, he  realized that the added DNA functioned as a template for the  synthesis of a new strand and through its partial degradation also  supplied the other, nonradioactive nucleotides required for  polymerization. The idea of a template was, at the time, foreign to  most enzymologists and other biochemists, but biologists seemed  more receptive to the role of a template in DNA synthesis. For his  discovery and characterization of DNA polymerase (later known as  DNA polymerase I or Pol I), Kornberg was awarded the 1959  Nobel Prize in Physiology or Medicine.   Even with purified Pol I in hand, Kornberg was faced with the  need to prove that the reaction product was biologically active. He  therefore used Pol I to synthesize the 5386-bp DNA of  bacteriophage 4>X174 on viral DNA templates and then used DNA  ligase to close up the synthetic DNA molecule to yield a circular  DNA that was infectious. To Kornberg’s chagrin, the popular press  misunderstood this work, hailing it as the creation of life in a  test tube.   Kornberg later confessed that he was amazed by the virtuosity  of the DNA polymerase he had isolated from E. coli: The enzyme  could synthesize a chain of thousands of nucleotides with an  accuracy that exceeded chemical predictions. However, during the  1970s, genetic studies and other evidence clearly indicated that  other proteins were responsible for DNA replication in E coli, and  DNA polymerases II and III were soon discovered and  characterized. For the next two decades, Kornberg led the effort  to determine the mechanism of DNA replication. Indeed, many of  the leading researchers in the field were trained in his laboratory.   Kornberg, A., Active center of DNA polymerase, Science 163 , 1410-1418 (1969).  Kornberg, A, For Love of Enzymes: The Odyssey of a Biochemist, Harvard University  Press (1989). [A scientific autobiography.]    [ l4 C]thymidine triphosphate into DNA (Box 25-1). This enzyme, which is  now known as DNA polymerase I or Pol I, consists of a single 928-residue  polypeptide. Pol I is said to be a processive enzyme because it catalyzes a  series of successive nucleotide polymerization steps, typically 20 or more, without  releasing the single-stranded template.   Pol I Has Exonuclease Activity. In addition to its polymerase activity, Pol I  has two independent hydrolytic activities that occupy separate active sites: a  y —> 5' exonuclease and a 5' — > 3' exonuclease. The 3' —> 5' exonuclease  activity allows Pol I to edit its mistakes. If Pol I erroneously incorporates a  mispaired nucleotide at the end of a growing DNA chain, the polymerase  activity is inhibited and the 3' —> 5' exonuclease hydrolytically excises the    FIG. 25-6 The 3' —» 5' exonuclease  function of DNA polymerase I. This enzymatic  activity excises mispaired nucleotides from the 3'  end of the growing DNA strand (blue).   FIG. 25-7 The 5' —» 3' exonuclease  function of DNA polymerase I. This enzymatic  activity excises up to 10 nucleotides from the 5'  end of a single-strand nick. The nucleotide  immediately past the nick (X) may or may not be  paired.    offending nucleotide (Fig. 25-6). The polymerase activity then resumes DNA  replication. This proofreading mechanism explains the high fidelity of DNA  replication by Pol I.   The Pol I 5' —> 3' exonuclease binds to duplex DNA at single-strand nicks  (breaks). It cleaves the nicked DNA strand in a base-paired region beyond the  nick to excise the DNA as either mononucleotides or oligonu¬  cleotides of up to 10 residues (Fig. 25-7).   Although Pol I was the first of the E. coli DNA polymerases  to be discovered, it is not E. colts primary replicase. Rather, its  most important (and only essential) function is in lagging strand  synthesis, in which it removes the RNA primers and replaces  them with DNA. This process involves the 5' —> 3' exonuclease  and polymerase activities of Pol I working in concert to excise the  ribonucleotides on the 5' end of the single-strand nick between  the new and old (previously synthesized) Okazaki fragments and  to replace them with deoxynucleotides that are appended to the  y end of the new Okazaki fragment (Fig. 25-8). The nick is thereby trans¬  lated (moved) toward the DNA strands 3 ' end, a process known as nick  translation. When the RNA has been entirely excised, the nick is sealed by  the action of DNA ligase (Section 25-2C), thereby linking the new and old  Okazaki fragments.    FIG. 25-8 The replacement of RNA  primers by DNA in lagging strand synthesis.   (1) The RNA primer at the 5' end of a  previously synthesized Okazaki fragment  (j Fragment 1) is excised through the action of Pol  I’s 5' —» 3' exonuclease function and replaced  through its polymerase function, which adds  deoxynucleotides to the 3' end of the newly  synthesized Okazaki fragment (Fragment 2). This,  in effect, translates the nick originally at the 5'  end of the RNA to the position that was  occupied by its 3' end (nick translation). (2) The  nick is sealed by the action of DNA ligase.   Biochemists use nick translation to prepare radioactive DNA. Double- Section 2 Prokaryotic DNA Replication   stranded DNA (dsDNA) is nicked in only a few places by treating it with  small amounts of pancreatic DNase I. Radioactively labeled dNTPs are then  added and Pol I translates the nicks, thereby replacing unlabeled deoxynu-  cleotides with labeled deoxynucleotides.   Pol I also functions in the repair of damaged DNA. As we discuss in  Section 25-5, damaged DNA is detected by a variety of DNA repair systems,  many of which endonucleolytically cleave the damaged DNA on the 5 ' side  of the lesion. Pol Is 5' —> 3' exonuclease activity then excises the damaged  DNA while its polymerase activity fills in the resulting single-strand gap in  the same way it replaces the RNA primers of Okazaki fragments. Thus, Pol I  has indispensable roles in E. coli DNA replication and repair although it is  not, as was first supposed, responsible for the bulk of DNA synthesis.   The Klenow Fragment Is a Model DNA Polymerase. E. coli DNA polymerase  I, a protein whose three enzymatic activities occupy three separate active sites,  can be proteolytically cleaved to a large or “Klenow” fragment (residues  324-928), which contains both the polymerase and the 3' —> 5' exonuclease  activities, and a small fragment (residues 1-323), which contains the 5' —> 3'  exonuclease activity. The X-ray structure of the Klenow fragment, determined  by Thomas Steitz, reveals that it consists of two domains (Fig. 25-9). The smaller  domain (residues 324-517; the lower portion of the structure shown in Fig.   25-9) contains the 3' —> 5' exonuclease site. The larger domain (residues  521-928) contains the polymerase active site at the bottom of a prominent  cleft, a surprisingly large distance (—25 A) from the 3' —» 5' exonuclease site.   The cleft, which is lined with positively charged residues, has the appropriate  size and shape (—22 A X —30 A) to bind a B-DNA molecule in a manner  resembling a right hand grasping a rod (note the thumb, fingers, and palm  structures in Fig. 25-9). The active sites of nearly all DNA and RNA poly¬  merases of known structure are located at the bottoms of similarly shaped clefts.    FIG. 25-9 X-Ray structure of E. coli DNA  polymerase I Klenow fragment in complex  with a double-helical DNA. The protein is  drawn in ribbon form colored in rainbow order  from its N-terminus (blue) to its C-terminus (red)  and embedded in its semitransparent molecular  surface. The DNA is shown in stick form with  the C atoms of its 10-nt template strand cyan,  the C atoms of its 13-nt primer strand magenta,  N blue, 0 red, and P orange, and with successive  P atoms in each polynucleotide strand connected  by orange rods. A Zn 2+ ion (purple) marks the  3' —» 5' exonuclease active site. Note that the 3'  end of the primer strand occupies the enzyme’s  3' —> 5' exonuclease site and hence this structure  is a so-called editing complex. [Based on an  X-ray structure by Thomas Steitz, Yale University.  PDBid 1 KLN.] Q See Interactive Exercise 46.   Chapter 25 DNA Replication, Repair, and phosphate backbone of the duplex DNA, consistent with Pol Is lack of   Recombination sequence specificity in binding DNA. The separation of the polymerase and   y —> 5' exonuclease active sites suggests that the bound DNA undergoes a  large conformational shift in shuttling between the sites.   DNA Polymerase Senses Watson-Crick Base Pairs via Sequence-  Independent Interactions. The C-terminal domain of the thermostable  Thermus aquaticus (Taq) DNA polymerase I (Klentaql) is 50% identical in  sequence and closely similar in structure to the large domain of Klenow  fragment (Klentaql lacks a functional 3' —> 5' exonuclease site). Gabriel  Waksman crystallized Klentaql in complex with a double-stranded DNA mol¬  ecule with a GGAAA overhang at its 5' end (representing a template with a  complementary DNA primer). The crystals were then incubated with 2', 3'-  dideoxy-CTP (ddCTP, which lacks a 3'-OH group). The X-ray structure of  the crystals (Fig. 25-10^) reveals that a ddC residue is covalently linked to the  y end of the primer, where it forms a Watson-Crick pair with the G residue  at the y end of the overhang. Moreover, a separate ddCTP molecule (to which  the primers new 3'-terminal ddC residue is incapable of forming a covalent  bond) occupies the enzymes active site, where it forms a Watson-Crick pair  with the templates next G. Clearly, Klentaql retains its catalytic activity in the  crystals.   A DNA polymerase must distinguish correctly paired bases from mis¬  matches and yet do so via sequence-independent interactions with the incom¬  ing dNTP. The Klentaql structure reveals that this occurs in an active site  pocket whose shape is complementary to Watson-Crick base pairs. In addition,   FIG. 25-10 X-Ray structure of Klentaql in complex with DNA  in the closed and open conformations, (a) The closed conformation  (with ddCTP). (b) The open conformation (ddCTP depleted). The  protein, which is viewed as in Fig. 25-9, is represented in worm form  with its N-terminal, palm, fingers, and thumb domains colored yellow,  magenta, green, and blue, respectively. The DNA is shown in stick  form, whereas the primer strand’s 3'-terminal ddC residue is drawn in    space-filling form as is the ddCTP in Part a that is base-paired with a  template G residue in the enzyme’s active site pocket. The atoms are  colored according to type with template strand C cyan, primer strand  C green, ddCTP C yellow, N blue, 0 red, and P orange. [Based on  X-ray structures by Gabriel Waksman, Washington University School of  Medicine. PDBids 3KTQ and 2KTQ.]   although the bound dsDNA is mainly in the B conformation, the 3 base Section 2 Prokaryotic DNA Replication   pairs nearest the active site assume the A conformation, as has also been observed   in the X-ray structures of several other DNA polymerases in their complexes with   DNA. The resulting wider and shallower minor groove (Section 24-1 A) permits   protein side chains to form hydrogen bonds with the otherwise inaccessible   N3 atoms of the purine bases and 02 atoms of the pyrimidine bases. The   be seen from an inspection of Fig. 24-1 (in contrast, the positions of the   hydrogen bonding acceptors in the major groove vary with the sequence).   However, with a non-Watson-Crick pairing—that is, with a mismatched  dNTP in the active site—these hydrogen bonds would be greatly distorted if  not completely disrupted. The polymerase also makes extensive sequence-  independent hydrogen bonding and van der Waals interactions with the DNAs  sugar-phosphate backbone.   After the Klentaql • DNA • ddCTP crystals described above were depleted  of ddCTP by soaking them in a ddCTP-free solution, the enzymes fingers  domain assumed a so-called open conformation (Fig. 25-10^), which differs  significantly from the closed conformation shown in Fig. 25-10/2. Evidently,  the fingers helices in the open conformation move toward the active site when  ddCTP binds, thereby burying the nucleotide in the catalytically active closed  complex. These observations are consistent with kinetic measurements indi¬  cating that the binding of the correct dNTP to Pol I induces a rate-limiting  conformational change that yields a tight ternary complex. It therefore appears  that the enzyme rapidly samples the available dNTPs in its open conforma¬  tion, but only when it binds the correct dNTP in a Watson-Crick pairing  with the template base does it form the catalytically competent closed confor¬  mation. After formation of the phosphodiester bond, a second conformational  change releases the PP Z product, and the DNA is translocated in the active site  so as to position it for the next reaction cycle.   X-ray structures of a variety of DNA polymerases suggest that they share a  common catalytic mechanism for nucleotidyl transfer. Their active sites all  contain two metal ions, usually Mg 2+ , that are liganded by two invariant Asp  side chains in the palm domain (Fig. 25-11). Metal ion B is liganded by all  three phosphate groups of the bound dNTP, whereas metal ion A bridges the  a-phosphate group of this dNTP and the primers 3'-OH group. Metal ion  A presumably activates the primers 3'-OH group for a nucleophilic attack on  the a-phosphate group, whereas metal ion B functions to orient its bound  triphosphate group and to electrostatically shield its negative charges as well  as the additional negative charge on the transition state leading to the release  of the PP Z ion.   DNA Polymerase III Is E. coli’s DNA Replicase. The discovery of normally  growing E. coli mutants that have very little (but not entirely absent) Pol I  activity stimulated the search for additional DNA polymerizing activities. This  effort was rewarded by the discovery of two more enzymes, designated, in the  order they were discovered, DNA polymerase II (Pol II) and DNA poly¬  merase III (Pol III) . The properties of these enzymes are compared with those    FIG. 25-1 1 Role of metal ions in nucleotidyl transfer. A and B represent  enzyme-bound metal ions, usually Mg 2+ , in the DNA polymerase active site. Atoms are  colored according to type (C gray, N blue, 0 red, and P yellow), and metal ion coordination  is represented by green dotted lines. Metal ion A activates the primer’s 3'-0H group for  nucleophilic attack (arrow) on the a-phosphate group of the incoming nucleotide (here,  ddCTP). Metal ion B orients and electrostatically stabilizes the negatively charged  triphosphate group. [Courtesy of Tom Ellenberger, Harvard Medical School.]   Source: Kornberg, A. and Baker, T.A., DNA Replication (2nd ed.), p. 167, Freeman (1992).    of Pol I in Table 25-1. Pol II and Pol III had not previously been detected  because their combined activities in the assays used are normally <5% that  of Pol I. Pol II participates in DNA repair; mutant cells lacking Pol II can  therefore grow normally. The absence of Pol III, however, is lethal, demon¬  strating that it is E. colls DNA replicase.   The catalytic core of Pol III consists of three subunits: a, which contains  the complex’s DNA polymerase activity (Fig. 25-12); 8 , its 3' —> 5' exonucle¬  ase; and 0. However, at least seven other subunits (t, y, 8, 8', T, and (3)  combine with them to form a labile multisubunit enzyme known as the Pol  III holoenzyme. The catalytic properties of the Pol III core resemble those of  Pol I except that Pol III lacks 5' —> 3' exonuclease activity on dsDNA. Thus,  Pol III can synthesize a DNA strand complementary to a single-stranded template  and can edit the polymerization reaction to increase replication fidelity, but it  cannot catalyze nick translation.   The E. coli chromosome is a supercoiled DNA molecule of 4.6 X 10 6 bp.  Since DNA polymerase requires a single-stranded template, other proteins par¬  ticipate in DNA replication by locating the replication initiation site, unwind¬  ing the DNA, and preventing the single strands from reannealing. Replication  in E. coli begins at a 245-bp region known as oriC. Elements of this sequence   Pol III ol subunit. The protein is drawn in ribbon  form with its thumb, PHP, palm, and fingers  domains green, blue, magenta, and orange,  respectively. Note the handlike shape of the  protein but its entirely different fold from that of  Klenow fragment and Klentaql (Figs. 25-9 and  25-10), with which it has no significant  sequence similarity. [Based on an X-ray structure  by Mike O’Donnell, The Rockefeller University,  and John Kuriyan, University of California at  Berkeley. PDBid 2HQA.]    are highly conserved among gram-negative bacteria. Multiple copies of a  52-kD protein known as DnaA bind to oriC and cause ~45 bp of an AT-rich  segment of the DNA to separate into single strands. This melting requires the  free energy of ATP hydrolysis and is probably also facilitated by both the  AT-rich nature of the DNA segment and the negative supercoiling (under¬  winding) of the circular DNA chromosome [the latter being generated by  DNA gyrase, a type II topoisomerase (Section 24-ID) whose activity is  required for prokaryotic DNA replication].    Helicases Unwind DNA. DnaA bound to oriC recruits two hexameric com¬  plexes of DnaB, one to each end of the melted region. DnaB is a helicase  that further separates the DNA strands. Helicases are a diverse group of  enzymes that unwind DNA during replication, transcription, and a variety of  other processes. DnaB is one of 12 helicases expressed by E. coli. Helicases  function by translocating along one strand of a double-helical nucleic acid so  as to mechanically unwind the helix in their path, a process that is driven by  the free energy of NTP hydrolysis.   E. coli DnaB, a hexamer of identical 471-residue subunits, separates the  two strands of the parental DNA by translocating along the lagging strand  template in the 5' —> 3' direction, while hydrolyzing ATP (it can also use GTP  and CTP but not UTP). Some helicases move in the 3' —> 5' direction, and  some are dimers rather than hexamers.   The El protein of bovine papillomavirus, a 605-residue hexameric heli¬  case, translocates along ssDNA in the 3' —> 5' direction (the opposite direc¬  tion of DnaB). The X-ray structure of its C-terminal 274 residues in complex  with a 13-nt poly(dT) and ADP (Fig. 25-13) was determined by Leemor  Joshua-Tor. Each protein subunit consists of two domains: a 74-residue N-  terminal oligomerization domain and a 200-residue C-terminal AAA+ domain  (AAA+ for ATPases /Associated with cellular /Activities; a functionally diverse  protein family). The protein forms a two-layered hexagonal ring in which the  oligomerization domains form a rigid collar with nearly perfect sixfold symmetry.   FIG. 25-13 X-Ray structure of bovine papillomavirus El  helicase in complex with poly(dT) and ADP. (a) The protein is  drawn in ribbon form viewed along the homohexamer’s pseudo-sixfold  axis with each protein subunit differently colored and with the  oligomerization domain of each subunit more lightly shaded than its  AAA+ domain. Protein loops ( purple ) extend radially inward from each  subunit to interact with the DNA’s phosphate groups. The poly(dT), 6  nt of which are visible, is drawn in stick form with C gray, N blue, 0    red, and P orange and with successive P atoms joined by orange  rods. Its 5' end is closest to the viewer. The ADP is shown in  space-filling form with C green, N blue, 0 red, and P orange, (b) Side  view of the protein related to that in Part a by a 90° rotation about  the horizontal axis. The blue and green AAA+ domains in Part a have  been deleted to expose the DNA, which is drawn in space-filling form.  [Based on an X-ray structure by Leemor Joshua-Tor, Cold Spring  Harbor Laboratory, New York. PDBid 2GXA.]   FIG. 25-14 X-Ray structure of SSB in  complex with dC(pC) 34 . The homotetramer,  which has D 2 symmetry, is viewed along one of  its twofold axes with its other twofold axes  horizontal and vertical. Each of its subunits  (which include the N-terminal 134 residues of  the 177-residue polypeptide) are differently  colored. Its two bound ssDNA molecules are  drawn in space-filling form colored according to  atom type with the upper strand C cyan, lower  strand C green, N blue, 0 red, and P orange.  (The lower strand is partially disordered and  hence appears to consist of two fragments).  [Based on an X-ray structure by Timothy  Lohman and Gabriel Waksman, Washington  University School of Medicine. PDBid 1 EYG.]    In contrast, the AAA+ domains deviate significantly from this symmetry  (Fig. 25-13 a). An ADP is bound at a radially peripheral site between each  neighboring pair of AAA+ domains. The poly(dT) forms a right-handed  helix that binds in the minimally ~13-A-diameter central channel of the  AAA+ domain hexamer (which is too narrow to admit dsDNA) with its 5'  end toward the top of the hexamer in Fig. 25-13. The DNAs phosphate  groups each interact with a positively charged loop (residues 505-508) that  extends radially inward from each AAA+ domain such that these loops form  an arrangement that resembles a right-handed spiral staircase that tracks the  ssDNAs sugar-phosphate backbone. Apparently, the protein steps through a  series of ATP-driven conformational changes that, via interactions with the  loops, pushes the ssDNA through the channel from bottom to top in Fig.  25-13^. During this process, each loop maintains its grip on the same phos¬  phate group. ATP hydrolysis occurs toward the bottom of the spiral staircase  and ADP release occurs between subunits located toward its top. A new ATP  then binds to this site, which causes the topmost loop to drop to the bottom  of the staircase, where it binds the next available phosphate group and repeats  the catalytic cycle. Thus the El helicase mechanically separates the strands of  dsDNA by pulling itself along the groove of one strand in its 3' —> 5' direc¬  tion but without turning relative to the DNA.   separated DNA strands behind an advancing helicase do not reanneal  to form dsDNA because they become coated with single-strand bind¬  ing protein (SSB). The SSB coat also prevents ssDNA from forming  secondary structures (such as stem-loops) and protects it from nucle¬  ases. Evidently, DNA polymerase displaces SSB from the template  strand as replication proceeds.   E. coli SSB is a homotetramer of 177-residue subunits that can  bind to DNA in several different ways. In the major binding mode  (Fig. 25-14), each U-shaped strand of ssDNA is draped across two  of SSB s four subunits. This would permit an unlimited series of SSB  tetramers to interact end-to-end along the length of a ssDNA. The  DNA-binding cleft of SSB, which is contained in its N-terminal 115  residues, is positively charged so that the protein can interact electro¬  statically with DNA phosphate groups. The cleft is too narrow to ac¬  commodate dsDNA.    The Primosome Synthesizes RNA Primers. All DNA synthesis, both of  leading and lagging strands, requires the prior synthesis of an RNA primer.  Primer synthesis in E. coli is mediated by an ~600-kD protein assembly  known as a primosome, which includes the DnaB helicase and an RNA-  synthesizing primase called DnaG, as well as five other types of subunits. E. coli  DnaG is a monomeric protein whose catalytic domain does not resemble any  of the other DNA and RNA polymerases of known structure. Nevertheless, it  catalyzes the same polymerization reaction (Fig. 25-2 using NTPs rather than  dNTPs) to produce an RNA segment of ~ 11 nucleotides.   The primosome is propelled in the 5' —> 3' direction along the DNA tem¬  plate for the lagging strand (i.e., toward the replication fork) in part by DnaB-  catalyzed ATP hydrolysis. This motion, which displaces the SSB in its path,  is opposite in direction to that of template reading during DNA chain syn¬  thesis. Consequently, the primosome reverses its migration momentarily to  allow primase to synthesize an RNA primer in the 5' —> 3' direction (Fig. 25-5).   The primosome is required to initiate each Okazaki fragment. The single  RNA segment that primes the synthesis of the leading strand can be synthe¬  sized, at least in vitro , by either primase or RNA polymerase (the enzyme that  synthesizes RNA transcripts from a DNA template; Section 26-1), but its rate  of synthesis is greatly enhanced when both enzymes are present.   In E. coli, the Pol III holoenzyme catalyzes the synthesis of both the leading  and lagging strands. This occurs in a single multiprotein particle, the repli-  some, which contains two Pol III enzymes. In some other prokaryotes and in  eukaryotes, two different polymerases synthesize the leading and lagging  strands, but like Pol III, they are part of a multiprotein replisome. In order for  the replisome to move as a single unit in the 5' —> 3' direction along the leading  strand, the lagging strand template must loop around (Fig. 25-15). After com¬  pleting the synthesis of an Okazaki fragment, the lagging strand holoenzyme  relocates to a new primer near the replication fork and resumes synthesis. The  result of this process is a continuous leading strand and a series of RNA-primed  Okazaki fragments separated by single-strand nicks. The RNA primers are replaced  with DNA through Pol I—catalyzed nick translation, and the nicks in the  lagging strand are then sealed through the action of DNA ligase (see below).    RNA primer to be replaced  with DNA by Pol I;  nick sealed by DNA ligase    FIG. 25-1 5 The replication of E. coli DNA. (a) The replisome,  which contains two DNA polymerase III holoenzymes, synthesizes both  the leading and the lagging strands. The lagging strand template must  loop around to permit the holoenzyme to extend the primed lagging  strand. ( b ) The holoenzyme releases the lagging strand template when  it encounters the previously synthesized Okazaki fragment. This may    signal the primosome to initiate synthesis of a lagging strand RNA  primer, (c) The holoenzyme rebinds the lagging strand template and  extends the RNA primer to form a new Okazaki fragment. Note that  in this model, leading strand synthesis is always ahead of lagging  strand synthesis.   Chapter 25 DNA Replication, Repair, and a Sliding Clamp Promotes Pol III Processivity. The Pol III core enzyme   is, it has a processivity of — 12 residues. However, the Pol III holoenzyme has  a processivity of >5000 residues, due to the presence of its P subunit. A P  subunit bound to a cut circular DNA slides to the break and falls off. This  suggests that the P subunit forms a ring around the DNA that functions as a  sliding clamp (alternatively, /J clamp) that can move along it, thereby keeping  the Pol III holoenzyme from diffusing away. The p clamp also increases the rate  of nucleotide polymerization.   The X-ray structure of the p clamp in complex with DNA, determined  by John Kuriyan and Mike O’Donnell, reveals that the protein is a dimer of  C-shaped monomers that form an — 80-A-diameter donut-shaped structure  (Fig. 25-16). Each P subunit contains three domains of similar structure (al¬  though with <20% sequence identity) so that the dimeric ring is a pseudo-  symmetrical six-pointed star. The interior surface of the ring is positively  charged, whereas its outer surface is negatively charged.   The central —35-A-diameter hole of the P clamp is larger than the 20-  and 26-A diameters of B- and A-DNAs (the hybrid helices containing RNA  primers and DNA have an A-DNA-like conformation; Section 24-1 A). The  X-ray structure of the P clamp-DNA complex indicates that the proteins  a helices span the major and minor grooves of the DNA rather than entering  into them as do, for example, the recognition helices of helix—turn—helix  motifs (Section 24-4B). It appears that the p subunit is designed to minimize  its associations with double-stranded DNA. This presumably permits the  protein to freely slide along the DNA helix.   E. coli replicates its DNA at a rate of —1000 nt/s. Thus, in lagging strand  synthesis, the DNA polymerase holoenzyme must be reloaded onto the  template strand every second or so (Okazaki fragments are —1000 nt long).  Consequently, a new P clamp, which promotes Pol Ills processivity, must be  installed around the lagging strand template about once every second. The y  complex of the Pol III holoenzyme (subunit composition yT 2 88'x^) is also    FIG. 25-1 6 X-Ray structure of the p clamp of E. coli Pol III  holoenzyme in complex with DNA. (a) The homodimeric sliding clamp is  drawn in ribbon form embedded in its semitransparent surface diagram and  viewed along its twofold axis with one subunit magenta and the other  colored in rainbow order from its N-terminus (blue) to its C-terminus (red).  The DNA, which consists of a 10-bp double-stranded segment with a 4-nt  single-stranded extension at the 5' end of one of the strands, is drawn in  stick form with the C atoms of the template strand cyan, those of the  primer strand green, N blue, 0 red, and P orange, and with an orange rod  connecting successive P atoms in each strand, (b) Cutaway diagram of the  structure in Part a rotated 90° about the horizontal axis. The DNA, which is  shown in space-filling form, is inclined by —22° to the protein’s twofold  axis, which is vertical in this diagram. [Based on an X-ray structure by John  Kuriyan, University of California at Berkeley, and Mike O’Donnell, The  Rockefeller University. PDBid 3BEP.]    FIG. 25-1 7 Architecture of the E. coli replisome. See the text for details. Compare  this to Fig. 25-15 in which only the t 2 component of the clamp loader is drawn. [Courtesy  of Charles Richardson, Harvard Medical School.]    known as the clamp loader because it opens the dimeric (3 clamp to load it  onto the DNA template in an ATP-dependent manner. The y complex bridges  the replisomes two Pol III cores (aS0) via the C-terminal segments (t c ) of its  two t subunits (Fig. 25-17), which also bind the DnaB helicase. This appar¬  ently allows the helicase to match the pace of the two polymerases.   Once the (3 clamp has been loaded onto the DNA, the Pol III core binds  to the (3 clamp more tightly than does the 7 complex, thereby displacing it  and permitting processive DNA replication to occur. When the polymerase  encounters the previously synthesized Okazaki fragment, that is, when the gap  between the two successively synthesized Okazaki fragments has been reduced  to a nick, the Pol III core releases the DNA and loses its affinity for the  (3 clamp. However, the components of the replisome don’t go far; the 7 subunit  and the Pol III core are held in the vicinity of the lagging strand template  through their linkage to the Pol III core engaged in leading strand synthesis  (which remains tethered to the DNA by its associated (3 clamp; Fig. 25-17).  Consequently, the 7 complex can quickly load a new (3 clamp around the  template DNA at the next primer so that Pol III can begin synthesizing a new  Okazaki fragment. The leading-strand Pol III pauses only momentarily while  the lagging-strand polymerase is repositioned.   The sliding clamp, which remains around the completed Okazaki frag¬  ment, probably recruits Pol I and DNA ligase so as to replace the RNA primer  on the previously synthesized Okazaki fragment with DNA and seal the  remaining nick. However, the sliding clamp must eventually be recycled. It was  initially assumed that this was the job of the clamp loader. However, it is now  clear that the release of the sliding clamp from its associated DNA is carried  out by free 8 subunit (the “wrench” in the clamp loader that cracks apart the  (3 subunits forming the sliding clamp), which is synthesized in fivefold excess  over that required to populate the cells few clamp loaders.   DNA Ligase Is Activated by NAD + or ATP. The free energy for the DNA   ligase reaction is obtained, in a species-dependent manner, through the  coupled hydrolysis of either NAD + to nicotinamide mononucleotide   E. coli DNA ligase. In eukaryotic and T4 ligases,  NAD + is replaced by ATP so that PP/ rather  than NMN + is eliminated in the first reaction  step. Here A, R, and N represent the adenine,  ribose, and nicotinamide residues, respectively.    Compare the free energy requirements  of the reactions catalyzed by DNA  polymerase and DNA ligase.    (NMN + ) + AMP, or ATP to PP, + AMP E. coli DNA ligase,  a 77-kD monomer that uses NAD + , catalyzes a three-step reac-  tion (Fig. 25-18):   1 . The adenylyl group of NAD + is transferred to the 8-amino  group of an enzyme Lys residue to form an unusual phospho-  amide adduct.   2 . The adenylyl group of this activated enzyme is transferred to  the 5 r -phosphoryl terminus of the nick to form an adenyly-  lated DNA. Here, AMP is linked to the 5'-nucleotide via a  pyrophosphate rather than the usual phosphodiester bond.   3. DNA ligase catalyzes the formation of a phosphodiester bond  by attack of the 3'-OH on the 5'-phosphoryl group, thereby  sealing the nick and releasing AMP.   ATP-requiring DNA ligases, such as those of eukaryotes, release  PP Z in the first step of the reaction rather than NMN + . The  DNA ligase from the bacteriophage T4 is notable because it can  link together two duplex DNAs that lack complementary single-  stranded ends (blunt end ligation) in a reaction that is a boon to  genetic engineering (Section 3-5).   The E. coli replication terminus is a large (350-kb) region flanked by ten nearly  identical nonpalindromic ~23-bp terminator sites, TerH y Terl y TerE y TerD y  and TerA on one side and TerJ y TerG y TerF y TerB y and TerC on the other  (Fig. 25-19; note that oriC is directly opposite the termination region on the  E. coli chromosome). A replication fork, traveling counterclockwise as drawn  in Fig. 25-19 passes through TerJ, TerGy TerFy TerB y and TerC but stops on  encountering either TerA } TerD y TerEy Terl y or TerH ( TerD , TerEy Terly and  TerH are presumably backup sites for TerA). Similarly, a clockwise-traveling  replication fork passes TerHy Terly TerEy TerDy and TerA but halts at TerC or,  failing that, TerBy TerFy TerGy or TerJ. Thus, these termination sites are polar;  they act as one-way valves that allow replication forks to enter the terminus  region but not to leave it. This arrangement guarantees that the two replication  forks generated by bidirectional initiation at oriC will meet in the replication  terminus even if one of them arrives there well ahead of its counterpart.   The arrest of replication fork motion at Ter sites requires the action of  Tus protein, a 309-residue monomer that is the product of the tus gene (for  terminator utilization .substance). Tus protein specifically binds to a Ter site,  where it prevents strand displacement by DnaB helicase, thereby arresting  replication fork advancement.   The X-ray structure of Tus in complex with a 15-bp Ter fragment  (Fig. 25-20) reveals that the protein forms a deep positively charged cleft in    FIG. 25-1 9 Map of the E. coli  chromosome showing the positions of the  Ter sites. The TerC, TerB, TerF, and TerG  sites, in combination with Tus protein, allow a  counterclockwise-moving replisome to pass  but not a clockwise-moving replisome. The  opposite is true of the TerA, TerD, and TerE  sites. Consequently, two replication forks that  initiate bidirectional DNA replication at oriC  will meet between the oppositely facing Ter  sites.   FIG. 25-20 X-Ray structure of E. coli Tus  in complex with a 15-bp Ter-containing DNA.   The protein is drawn in ribbon form colored in  rainbow order from its N-terminus (blue) to its  C-terminus (red). The DNA is shown in stick  form with C gray, N blue, 0 red, and P orange  and with successive P atoms in the same strand  joined by orange rods. [Based on an X-ray  structure by Kosuke Morikawa, Protein  Engineering Research Institute, Osaka, Japan.    which the DNA binds. A 5-bp segment of the DNA near the side of Tus that  permits the passage of the replication fork is deformed and underwound rela¬  tive to canonical (ideal) DNA: Its major groove is deeper, and its minor  groove is significantly expanded. Protein side chains at the bottom of the cleft  penetrate the DNAs widened major groove to make sequence-specific con¬  tacts such that the protein cannot release the bound DNA without a large  conformational change. Nevertheless, the mechanism through which Tus pre¬  vents replication fork advancement from one side of a Ter site but not the  other is unclear. Curiously, however, this system is not essential for termina¬  tion. When the replication terminus is deleted, replication simply stops, ap¬  parently through the collision of opposing replication forks. Nevertheless, this  termination system is highly conserved in gram-negative bacteria.   As two oppositely moving replication forks collide at the termination site,  the newly synthesized strands become covalently linked to yield two covalently  closed double-stranded chromosomes. The parental DNA strands remain  wound about each other by several turns (presumably because DNA gyrase  cannot gain access to the DNA when the colliding replication forks closely  approach each other) and consequently the product dsDNA strands must be  wound about each other by the same number of turns (Fig. 25-21). The result¬  ing catenated circular dsDNAs must be separated so that each can be passed  to a different daughter cell. This is the job of the type II topoisomerase named  topoisomerase IV (Section 24-ID).   Since a single polypeptide as small as the Pol I Klenow fragment can replicate  DNA by itself, why does E. coli maintain a battery of over 20 intricately  coordinated proteins to replicate its chromosome? The answer apparently is to  ensure the nearly perfect fidelity of DNA replication required to accurately trans¬  mit genetic information.    FIG. 25-21 The formation and separation of catenated dsDNAs at the replication  termination site. The parental strands are red and black and the daughter strands are  green and blue. For clarity, the double-helical character of the newly formed dsDNA  molecules is not shown.   • Summarize the functions of the following  proteins in E. coli DNA replication: DNA  polymerase I, DNA polymerase III, DnaA,  helicase, SSB, primase, the sliding clamp,  clamp loader, DNA ligase, Tus, and topo-  isomerases.   • Why is it important for DNA polymerase I  to have three active sites?   • What is the role of the metal ions in the  polymerase active site?   • How many sliding clamps and clamp¬  loading events are required for synthesis  of the leading and lagging strands?   • What feature of the clamp allows it to  slide along the DNA?   • Explain why Tus is the only replication  protein that is sequence-specific.   • Describe the four factors that contribute  to the high fidelity of DNA replication.    The rates of reversion of mutant E. coli or T4 phages to the wild type  indicates that only one mispairing occurs per 10 8 to 10 10 base pairs replicated.  This corresponds to error per 1000 bacteria per generation. Such high  replication accuracy arises from four sources:   1. Cells maintain balanced levels of dNTPs through the mechanisms  discussed in Sections 23-1C and 23-2C. This is important because a  dNTP present at aberrantly high levels is more likely to be misincor-  porated and, conversely, one present at low levels is more likely to be  replaced by one of the dNTPs present at higher levels.   2. The polymerase reaction itself has extraordinary fidelity because it  occurs in two stages. First, the incoming dNTP base-pairs with the  template while the enzyme is in an open, catalytically inactive confor¬  mation. Polymerization occurs only after the polymerase has closed  around the newly formed base pair, which properly positions the cat¬  alytic residues (induced fit; Fig. 25-10). The protein conformational  change constitutes a double-check for correct Watson-Crick base pairing  between the dNTP and the template.   4. A remarkable set of enzyme systems in all cells repairs residual errors  in the newly synthesized DNA as well as any damage that may occur  after its synthesis through chemical and/or physical insults. We discuss  the DNA repair systems in Section 25-5.   In addition, the inability of a DNA polymerase to initiate chain elonga¬  tion without a primer increases DNA replication fidelity. The first few  nucleotides of a chain are those most likely to be mispaired because of the  cooperative nature of base-pairing interactions (Section 24-2). The use of RNA  primers eliminates this source of error since the RNA is eventually replaced  by DNA under conditions that permit more accurate base pairing.    • Eukaryotic DNA replication requires DNA polymerases with different degrees of  accuracy and processivity.   • Prereplication complexes assemble at multiple origins distributed throughout the  eukaryotic genome.   • Telomerase uses an RNA template to extend the 3' ends of eukaryotic chromo¬  somes.    Eukaryotic and prokaryotic DNA replication mechanisms are remarkably sim¬  ilar, although the eukaryotic system is vastly more complex in terms of the  amount of DNA to be replicated and the number of proteins required (esti¬  mated at >27 in yeast and mammals). Several different modes of DNA repli¬  cation occur in eukaryotic cells, which contain nuclear DNA as well as  mitochondrial and, in plants, chloroplast DNA. In this section, we consider  some of the proteins of eukaryotic DNA replication as well as the challenge  of replicating the ends of linear chromosomes.   Animal cells contain at least 13 distinct DNA polymerases, which have been  named with Greek letters according to their order of discovery. A newer clas¬  sification scheme uses sequence homology to group eukaryotic as well as  prokaryotic polymerases into six families: A, B, C, D, X, and Y. In this    section, we describe the three main enzymes involved in replicating eukary¬  otic nuclear DNA: polymerases c x, 8, and 8, which are all members of the  B-family of polymerases (Table 25-2).   DNA polymerase a (pol a), like all DNA polymerases, replicates DNA  by extending a primer in the 5' 3' direction under the direction of a ssDNA   template. This enzyme has no exonuclease activity and therefore cannot proof¬  read its polymerization product. Pol a is only moderately processive (polymer¬  izing '■"TOO nucleotides at a time) and associates tightly with a primase,  indicating that it is involved in initiating DNA replication. The pol a/primase  complex synthesizes a 7- to 10-nt RNA primer and extends it by an additional  15 or so deoxynucleotides. Its lack of proofreading activity is not problematic,  since the first few residues of newly synthesized DNA are typically removed  and replaced along with the RNA primer.   DNA polymerase 8 (pol 8) does not associate with a primase and contains  a y —> 5' exonuclease active site. In addition, the processivity of pol 8 is  essentially unlimited (it can replicate the entire length of a template DNA), but  only when it is in complex with a sliding-clamp protein named proliferating  cell nuclear antigen (PCNA). The X-ray structure of PCNA (Fig. 25-22), also  determined by Kuriyan, reveals that it forms a trimeric ring with almost iden¬  tical structure (and presumably function) as the E. coli (3 2 sliding clamp (Fig.  25-16). Intriguingly, PCNA and the (3 clamp exhibit no significant sequence  identity, even when their structurally similar portions are aligned.   Pol 8 in complex with PCNA is required for lagging strand DNA synthesis.  During replication, RFC (replication factor C, a clamp loader that is the  eukaryotic counterpart of the E. coli y complex) loads PCNA onto the  template strand near the primer. This displaces pol a, which in a process called   three subunits, which form a threefold symmetric  ring, are drawn in ribbon form embedded in their  semitransparent surface diagram. One of the  subunits is colored in rainbow order from its  N-terminus (blue) to its C-terminus (red), another  is pink, and the third is light green. A space¬  filling model of B-DNA viewed along its helix  axis has been drawn in the center of the PCNA  ring. [Based on an X-ray structure by John  Kuriyan, University of California at Berkeley.  PDBid 1AXCJ O See Interactive Exercise 48.    Compare this structure to that of the p  clamp of the E. coli Pol III holoenzyme  (Fig. 25-16).    Reverse transcriptase (RT) is an essential enzyme of  retroviruses, which are RNA-containing eukaryotic viruses such  as human immunodeficiency virus (HIV, the causative agent of  AIDS; Box 12-3). RT, which was independently discovered in 1970  by Howard Temin and David Baltimore, synthesizes DNA in the  5' —» 3' direction from an RNA template. Although the activity of  the enzyme was initially considered antithetical to the central dogma  of molecular biology (Section 3-3B), there is no thermodynamic  prohibition to the RT reaction (in fact, under certain conditions,   Pol I can copy RNA templates). RT catalyzes the first step in the  conversion of the virus’ single-stranded RNA genome to a  double-stranded DNA.   After the virus enters a cell, its RT uses the viral RNA as a  template to synthesize a complementary DNA strand, yielding an  RNA-DNA hybrid helix. The DNA synthesis is primed by a host  cell tRNA whose 3' end unfolds to base-pair with a complementary  segment of viral RNA. The viral RNA strand is then nucleolytically  degraded by an RNase H (an RNase activity that hydrolyzes the  RNA of an RNA-DNA hybrid helix). Finally, the DNA strand acts  as a template for the synthesis of its complementary DNA, yielding  dsDNA that is then integrated into a host cell chromosome.   RT has been a particularly useful tool in genetic engineering  because it can transcribe mRNAs to complementary strands of  DNA (cDNA; see diagram). mRNA-derived cDNAs can be used,  for example, to express eukaryotic structural genes in E coli  (Section 3-5D). Since E. coli lacks the machinery to splice out  introns (Section 26-3B), the use of genomic DNA to express a  eukaryotic structural gene in E coli would require the prior exci¬  sion of its introns—a technically difficult feat.   HIV-1 reverse transcriptase is a dimeric protein whose subunits  are synthesized as identical 66-kD polypeptides, known as p66  (< opposite, top), that each contain a polymerase domain and an  RNase H domain. However, the RNase H domain of one of the    two subunits is proteolytically excised, thereby yielding a 51-kD  polypeptide named p51 ( opposite, bottom). Thus, RT is a dimer of  p66 and p51.   The X-ray structure of HIV-1 RT shows that the two subunits  have different structures, although each has a fingers, palm, and    template switching allows pol 8 to bind to the lagging strand template strand  and subsequently processively extend the new DNA strand.   Pol 8 , a heterotetrameric nuclear enzyme, is the most enigmatic partici¬  pant in DNA replication. Pol £ is highly processive in the absence of PCNA  and has a 3' —> 5' exonuclease activity that degrades single-stranded DNA to  6- or 7-residue oligonucleotides rather than to mononucleotides, as does that  of pol 8. Although pol £ is necessary for the viability of yeast, its essential  function can be carried out by only the noncatalytic C-terminal half of its  256-kD catalytic subunit, which is unique among B-family DNA polymerases.  This suggests that the C-terminal half of the pol £ catalytic subunit is required  for the assembly of the replication complex. Nevertheless, Thomas Kunkel has  shown that pol £ is probably the leading strand replicase, although it may also  contribute to lagging strand synthesis. Moreover, pol 8 may also participate in  leading strand synthesis.   DNA polymerase y (pol y), an A-family enzyme, occurs exclusively in  the mitochondrion, where it presumably replicates the mitochondrial genome.  Chloroplasts contain a similar enzyme. An additional member of the  polymerase family of proteins is the viral enzyme reverse transcriptase, an  RNA-directed DNA polymerase (Box 25-2).   thumb domain as well as a “connection”  domain. The RNase H domain of the p66  subunit follows the connection domain. The  p66 and p51 subunits are not related by  twofold molecular symmetry (a rare but not  unprecedented phenomenon) but instead  associate in a sort of head-to-tail arrangement.  Consequently, RT has only one polymerase  active site.   Reverse transcriptase lacks a proofreading  exonuclease function and hence is highly  error-prone. Indeed, it is HIV’s capacity to  rapidly evolve, even within a single host, that  presents a major obstacle to the development  of an anti-HIV vaccine. This high rate of  mutation is also the main contributor to the  ability of HIV to rapidly develop resistance to  drugs that inhibit virally encoded enzymes,  including RT. Q See Interactive Exercise 49.   [Based on an X-ray structure by Edward Arnold,  Rutgers University. PD Bid 3JYT.]   prokaryotic DNA replication, a helicase is required to pry apart the two  template strands. In eukaryotes, this function is carried out by the heterohexa-  meric complex known as MCM. Single-stranded DNA becomes coated with  the trimeric replication protein A (RPA), the eukaryotic equivalent of the  bacterial SSB. The eukaryotic replisome also includes additional proteins that  have no prokaryotic counterparts and whose functions are poorly understood.   Eukaryotes lack a nick-translating polymerase like E. coli Pol I. Instead,  the RNA primers of Okazaki fragments are removed through the actions of  two enzymes: RNase HI removes most of the RNA, leaving only a 5'-  ribonucleotide adjacent to the DNA, which is then removed through the ac¬  tion of flap endonuclease-1 (FEN1). Yet, as we have seen, pol a extends  the RNA primer by ~15 nt of DNA before it is displaced by pol 8. Since  pol a lacks proofreading ability, this primer extension is more likely to con¬  tain errors than the DNA synthesized by pol 8. However, FEN1, which is  recruited by PCNA, provides what is, in effect, pol as proofreading func¬  tion: It is also an endonuclease that excises mismatch-containing oligonu¬  cleotides up to 15 nt long from the 5' end of an annealed DNA strand.  Moreover, FEN1 can make several such excisions in succession to remove   FIG. 25-23 Removal of RNA primers in  eukaryotes. (1) RNase HI excises all but the  5'-ribonucleotide of the RNA primer. (2) FEN1,  a 5' —» 3' endonuclease, then removes the  remaining ribonucleotide along with a segment  of adjoining DNA if it contains mismatches.   (3) The excised nucleotides are replaced as  DNA polymerase 8 completes the synthesis of  the next Okazaki fragment (on the left in this  diagram). The nick is eventually sealed by DNA  ligase.    How does this process compare to the  removal of RNA primers in E. co//?    more distant mismatches. The excised segment is later replaced by pol 8 as  it synthesizes the succeeding Okazaki fragment (Fig. 25-23).   Replication fork movement in eukaryotes is —10 times slower than in prokary¬  otes. Since a eukaryotic chromosome typically contains 60 times more DNA  than does a prokaryotic chromosome, its bidirectional replication from a  single origin, as in prokaryotes, would require ~ 1 month. Electron micrographs  such as Fig. 25-24, however, show that eukaryotic chromosomes contain multi¬  ple origins , one every 3 to 300 kb, depending on both the species and the  tissue, so replication usually requires just a few hours to complete.   In yeast, the initiation of DNA replication occurs at autonomously repli¬  cating sequences (ARS), which are conserved 11-bp sequences adjacent to  easily unwound DNA. In mammalian genomes, replication origins do not  exhibit sequence conservation, yet these sites all support the binding of a six-  subunit origin recognition complex (ORC). Additional ATP-hydrolyzing  proteins help assemble the helicase MCM. The resulting prereplication  complex (pre-RC) is not competent to initiate replication until it has been  activated by other factors that control progress through the cell cycle  (Section 28-4A). Presumably, the separate control of pre-RC assembly and  activation allows cells to select replication origins before MCM unwinds the  template DNA and replication commences. Once DNA synthesis is under  way, no new pre-RC complexes can form, thereby ensuring that the DNA is  replicated once and only once per cell cycle.   Initiation sites are uniformly distributed across the genome in early  embryogenesis, when cell division is rapid. However, after cells have differenti¬  ated, the distribution of replication origins changes, possibly reflecting patterns  of gene expression and/or alterations in DNA packaging in different cell types.   Cytological observations indicate that the various chromosomal regions are  not all replicated simultaneously. Rather, clusters of 20 to 80 adjacent repli-  cons (replicating units; DNA segments that are each served by a replication  origin) are activated simultaneously. New sets of replicons are activated until  the entire chromosome has been replicated. DNA replication proceeds in each  direction from the origin of replication until each replication fork collides with  a fork from the adjacent replicon. Eukaryotes appear to lack termination  sequences analogous to the Ter sites in E. coli.   Nucleosomes Reassemble behind the Replication Forks. Unlike prokary¬  otic DNA, eukaryotic DNA is packaged in nucleosomes (Section 24-5A).  Some alteration of this structure is probably necessary for initiation, but once  replication is under way, nucleosomes do not seem to impede the progress of  DNA polymerases. Experiments with labeled histones indicate that nucleo¬  somes just ahead of the replication fork disassemble and the freed histones,  either individually or as dimers or tetramers, immediately reassociate with the  emerging daughter duplexes. The parental histones randomly associate with  the leading and lagging duplexes. DNA replication (which occurs in the    FIG. 25-24 Electron micrograph of a  fragment of replicating Drosophila DNA. The   arrows indicate the multiple replication eyes.  [From Kreigstein, H.J. and Hogness, D.S., Proc.  Natl. Acad Sci. 71, 136 (1974).]    nucleus) is coordinated with histone protein synthesis in the cytosol so that  new histones are available in the required amounts.   The ends of linear chromosomes present a problem for the replication  machinery. Specifically DNA polymerase cannot synthesize the extreme 5' end of  the lagging strand (Fig. 25-25). Even if an RNA primer were paired with the  3' end of the DNA template, it could not be replaced with DNA (recall that  DNA polymerase operates only in the 5' —> 3' direction; it can only extend  an existing primer, and the primer must be bound to its complementary  strand). Consequently, in the absence of a mechanism for completing the lagging  strand, linear chromosomes would be shortened at both ends by at least the length  of an RNA primer with each round of replication.   Telomeres Are Built from an RNA Template. The ends of eukaryotic chro¬  mosomes, the telomeres (Greek: telos, end), have an unusual structure.  Telomeric DNA consists of 1000 or more tandem repeats of a short G-rich  sequence (TTGGGG in the protozoan Tetrahymena and TTAGGG in  humans) on the 3 r -ending strand of each chromosome end.   Elizabeth Blackburn, Carol Greider, and Jack Szostak have shown that  telomeric DNA is synthesized and maintained by an enzyme named telo¬  merase, which is a ribonucleoprotein (a complex of protein and RNA). The  RNA component (451 nt in humans) includes a segment that is complemen¬  tary to the repeating telomeric sequence and acts as a template for a reaction  in which nucleotides are added to the 3' end of the DNA (Fig. 25-26, top).  Telomerase repeatedly translocates to the new 3' end of the DNA strand,  thereby adding multiple telomeric sequences to the DNA (Fig. 25-26, bottom ).  The DNA strand complementary to the telomeric G-rich strand is apparently  synthesized by the normal cellular machinery for lagging strand synthesis,  leaving a 100- to 300-nt single-stranded overhang on the G-rich strand.   Telomerase functions similarly to reverse transcriptase (Box 25-2); in fact, its  highly conserved catalytic subunit, called TERT, is homologous to reverse   FIG. 25-25 Replication of a linear  chromosome. Leading strand synthesis can  proceed to the end of the chromosome (top).  However, DNA polymerase cannot synthesize the  extreme 5' end of the lagging strand because it  can only extend an RNA primer that is paired  with the 3' end of a template strand (bottom).  Removal of the primer and degradation of the  remaining single-stranded extension would cause  the chromosome to shorten with each round of  replication.    FIG. 25-26 Mechanism for the synthesis  of telomeric DNA by Tetrahymena telomerase.   The telomere’s 5'-ending strand is later extended  by normal lagging strand synthesis. [After  Greider, C.W. and Blackburn, E.H., Nature 337,  336 (1989).]    FIG. 25-27 X-Ray structure of TERT. The protein, from the flour  beetle Tribolium castaneum, is shown in ribbon form embedded in its  semitransparent surface diagram with its four domains in different colors.  TRBD, the telomere repeat binding c/omain, binds RNA. [Based on an X-ray  structure by Emmanuel Skordalakes, The Wistar Institute, Philadelphia,  Pennsylvania. PDBid 3DU5.]    Compare the structures of TERT and the reverse transcriptase  subunit p66 shown in Box 25-2.   • Describe how pol a, pol 8, PCNA, RNase  HI, FEN1, and DNA ligase participate in  eukaryotic DNA synthesis.   • In what ways does replication differ  between eukaryotes and prokaryotes?   • What does telomerase do and why is this  enzyme unlike other DNA polymerases?    transcriptase (Fig. 25-27). In addition to the fingers, palm, and thumb domains  typical of other polymerases, TERT includes an N-terminal RNA-binding domain.   The absence of telomerase, which allows the gradual truncation of chro¬  mosomes with each round of DNA replication, contributes to the normal  senescence of cells. Conversely, enhanced telomerase activity permits the un¬  controlled replication and cell growth that occur in cancer (Box 25-3).   Telomeres Form G-Quartets. Guanine-rich polynucleotides are notoriously  difficult to work with. This is because of their propensity to aggregate via  Hoogsteen-type base pairing (Section 24-2A) to form cyclic tetramers known  as G-quartets (Fig. 25-28 a). Indeed, the G-rich overhanging strands of telo¬  meres fold back on themselves to form a hairpin, two of which associate in  an antiparallel fashion to form stable complexes of stacked G-quartets (Fig.  25-28 £). Such structures presumably serve as binding sites for capping pro¬  teins, which may help regulate telomere length and prevent activation of  DNA repair mechanisms that recognize the ends of broken DNA molecules.   base-pairing interactions in the  G-quartet. ( b ) Schematic diagram of the  NMR structure in which the strand  directions are indicated by arrows. The  nucleotides are numbered 1 to 12 in  one strand and 1* to 12* in the  symmetry-related strand. Guanine  residues G1 to G4 are represented by  blue rectangles, G9 to G12 are cyan,  G1* to G4* are red, and G9* to G12*  are pink. [After Schultze, P., Smith, F.W.,  and Feigon, J., Structure 2, 227 (1994).  PDBid 156DJ    Without the action of telomerase, a chromosome would be short¬  ened at both ends by at least the length of an RNA primer with  every cycle of DNA replication and cell division. It was therefore  initially assumed that, in the absence of active telomerase, essen¬  tial genes located near the ends of chromosomes would eventu¬  ally be lost, thereby killing the descendants of the originally  affected cells. However, it is now evident that telomeres serve  another vital chromosomal function that must be compromised  before this can happen. Free DNA ends trigger DNA damage  repair systems that normally function to rejoin the ends of broken  chromosomes (Section 25-5E). Consequently, exposed telomeric  DNA would result in the end-to-end fusion of chromosomes, a  process that leads to chromosomal instability and eventual cell  death (fused chromosomes often break in mitosis; their two  centromeres cause them to be pulled in opposite directions).  However, in a process known as capping, telomeric DNA is  specifically bound by proteins that hide the DNA ends. There is  mounting evidence that capping is a dynamic process in which the  probability of a telomere spontaneously uncapping increases as  telomere length decreases.   The somatic cells of multicellular organisms lack telomerase  activity. This explains why such cells in culture can undergo only a  limited number of doublings (20-60) before they reach senes¬  cence (a stage in which they cease dividing) and eventually die.  Indeed, otherwise immortal Tetrahymena cultures with mutationally  impaired telomerases exhibit characteristics reminiscent of senes¬  cent mammalian cells before dying off. Apparently, the loss of    telomerase function in somatic cells is a basis for aging in multicel¬  lular organisms.   Despite the foregoing, there is only a weak correlation be¬  tween the proliferative capacity of a cultured cell and the age of  its donor. There is, however, a strong correlation between the  initial telomere length in a cell and its proliferative capacity. Cells  that initially have relatively short telomeres undergo significantly  fewer doublings than cells with longer telomeres. Moreover, fibro¬  blasts from individuals with progeria (a rare disease characterized  by rapid and premature aging resulting in childhood death) have  short telomeres, an observation that is consistent with their known  reduced proliferative capacity in culture. In contrast, sperm (which  are essentially immortal) have telomeres that do not vary in length  with donor age, which indicates that telomerase is active during  germ-cell growth. Likewise, those few cells in culture that become  immortal (capable of unlimited proliferation) exhibit an active  telomerase and a telomere of stable length, as do the cells of  unicellular eukaryotes (which are also immortal).   What selective advantage might multicellular organisms gain  by eliminating the telomerase activity in their somatic cells? An  intriguing possibility is that cellular senescence is a mechanism  that protects multicellular organisms from cancer. Indeed, cancer  cells, which are immortal and grow uncontrollably, contain active  telomerase. For example, the enzyme is active in ovarian cancer  cells but not in normal ovarian tissue. This hypothesis makes  telomerase inhibitors an attractive target for antitumor  drug development.    • DNA is susceptible to damage from a variety of sources.   • Mutagenicity, which is related to carcinogenicity, can be tested in living cells.    The fidelity of DNA replication carried out by DNA polymerases and their  attendant proofreading functions is essential for the accurate transmission of  genetic information during cell division. Yet errors in polymerization occasion¬  ally occur and, if not corrected, may alter the nucleotide sequences of genes.  DNA can also be chemically altered by agents that are naturally present in the  cell or in the cells external environment.   In many cases, damaged DNA can be repaired, as discussed in Section 25-5.  Severe lesions, however, may be irreversible, leading to the loss of genetic in¬  formation and, often, cell death. Even when damaged DNA can be mended,  the restoration may be imperfect, producing a mutation, a heritable alteration  of genetic information. In multicellular organisms, genetic changes are usually  notable only when they occur in germline cells so that the change is passed  on to all the cells of the organisms offspring. Damage to the DNA of a so¬  matic cell, in contrast, rarely has an effect beyond that cell unless the muta¬  tion contributes to a malignant transformation (cancer).   Environmental agents such as ultraviolet light, ionizing radiation, and certain  chemical agents can physically damage DNA. For example, UV radiation  (200—300 nm) promotes the formation of a cyclobutyl ring between adjacent   The dimer forms on UV irradiation of two  adjacent thymine residues on a DNA strand. The  ~1.6-A-long covalent bonds joining the thymine  rings (red) are much shorter than the normal  3.4-A spacing between stacked rings in B-DNA,  thereby locally distorting the DNA.    thymine residues on the same DNA strand to form an intrastrand thymine  dimer (Fig. 25-29). Similar cytosine and thymine—cytosine dimers also form  but less frequently. Such pyrimidine dimers locally distort DNAs base-paired  structure, interfering with transcription and replication. Ionizing radiation also  damages DNA either through its direct action on the DNA molecule or  indirectly by inducing the formation of free radicals, particularly the hydroxyl  radical (HO*), in the surrounding aqueous medium. This can lead to strand  breakage.   The DNA damage produced by chemical mutagens, substances that  induce mutations, falls into two major classes:   1. Point mutations, in which one base pair replaces another. These are  subclassified as:   (a) Transitions, in which one purine (or pyrimidine) is replaced  by another.   (b) Transversions, in which a purine is replaced by a pyrimidine or  vice versa.   2. Insertion/deletion mutations, in which one or more nucleotide pairs  are inserted in or deleted from DNA.   Point Mutations Result from Altered Bases. One cause of point mutations  is treatment of DNA with nitrous acid (HN0 2 ), which oxidatively deaminates  aromatic primary amines. Cytosine is thereby converted to uracil and adenine  to the guaninelike hypoxanthine (which forms two of guanines three hydro¬  gen bonds with cytosine; Fig. 25-30). Hence, treatment of DNA with nitrous  acid results in both A • T —» G • C and G • C —> A • T transitions. Despite  its potential mutagenic activity, nitrite (the conjugate base of nitrous acid) is  used as a preservative in prepared meats such as frankfurters because it also  prevents the growth of Clostridium botulinum ., the organism that causes bot¬  ulism. Base deamination reactions also occur spontaneously in the absence of  nitrous acid.   Cellular metabolism itself exposes DNA to the damaging effects of reac¬  tive oxygen species (e.g., the superoxide ion 0 2 % the hydroxyl radical, and  H 2 0 2 ) that are normal by-products of oxidative metabolism (Section 18-4B).   FIG. 25-30 Oxidative deamination by nitrous acid, (a) Cytosine is converted to  uracil, which base-pairs with adenine, (b) Adenine is converted to hypoxanthine, a guanine  derivative (it lacks guanine’s 2-amino group) that base-pairs with cytosine.    Over 100 different oxidative modifications to DNA have been catalogued. For  example, guanine can be oxidized to 8-oxoguanine (oxoG, right). When the  modified DNA strand is replicated, the oxoG can base-pair with either C or  A, causing aG • C^T • A transversion.   Alkylating agents such as dimethyl sulfate, nitrogen mustard, ethylni-  trosourea, and A^methyl-A^-nitro-A^nitrosoguanidine (MNNG)    can also generate transversions. For example, the exposure of DNA to MNNG  yields, among other products, 0 6 -methylguanine residues ( right ), which can  base-pair with either C or T. The metabolic methylating agent S-adenosylmethio-  nine (Section 21-4D) occasionally nonenzymatically methylates a base to form  derivatives such as 3-methyladenine and 7-methylguanine residues. However,  base methylations also have normal physiological functions (Box 25-4).   The alkylation of the N7 position of a purine nucleotide renders its gly-  cosidic bond susceptible to hydrolysis, leading to loss of the base. The result¬  ing gap in the sequence is filled in by an error-prone enzymatic repair system.  Transversions arise when the missing purine is replaced by a pyrimidine. Even  in the absence of alkylating agents, the glycosidic bonds of an estimated  20,000 of the 6.0 billion purine nucleotides in each diploid human cell spon¬  taneously hydrolyze every day.   Insertion/deletion mutations may arise from the treatment of DNA with  intercalating agents such as acridine orange or proflavin (Section 24-3B). The  distance between two consecutive base pairs is roughly doubled by the inter¬  calation of such a molecule between them. The replication of the distorted  DNA occasionally results in the insertion or deletion of one or more  nucleotides in the newly synthesized polynucleotide. (Insertions and deletions of  large segments generally arise from aberrant crossover events; Section 25-6A.)    All Mutations Are Random. The bulk of the scientific data regarding mutage¬  nesis is that mutations, whether the result of polymerase errors, spontaneous  modification, or chemical damage to DNA, occur at random. This paradigm  was challenged by John Cairns, who demonstrated that bacteria unable to di¬  gest lactose preferentially acquired the mutations they needed in order to use  lactose when it was the only nutrient available. This observation, which sug¬  gests that bacteria can “direct” mutations that benefit them, more likely reflects  a nonspecific adaptive response in which the overall rate of mutation—useful  as well as nonuseful—increases when the cells are under metabolic stress. The  hypermutable state appears to reflect the activation of error-prone DNA repair and  recombination systems that are relatively inactive in normally growing cells.   Not all alterations to DNA have phenotypic consequences. For example, muta¬  tions in noncoding segments of DNA are often invisible. Similarly, the redun¬  dancy of the genetic code (more than one trinucleotide may specify a particular  amino acid; Section 27-1C) can mask point mutations. Even when a proteins  amino acid sequence is altered, its function may be preserved if the substitution  is conservative (Section 5-4A) or occurs on a surface loop. Nevertheless, even a  single point mutation, if appropriately located, can irreversibly alter cellular    Not all DNA modifications are detrimental. For example, the A and  C residues of DNA may be methylated, in a species-specific pat¬  tern, to form yv 6 -methyladenine (m 6 A), yv 4 -methylcytosine  (m 4 C), and 5-methylcytosine (m 5 C) residues:   The methyl groups project into B-DNA’s major groove, where they  can interact with DNA-binding proteins. In most cells, only a few  percent of the susceptible bases are methylated, although this  figure rises to >30% of the C residues in some plants.   Bacterial DNAs are methylated at their own particular restric¬  tion sites by modification methylases, thereby preventing the cor¬  responding restriction endonuclease from cleaving the DNA  (Section 3-4A). Other methyltransferases also modify DNA  bases in a sequence-specific manner. For example, in E. coli, the  Dam methyltransferase methylates the A residue in all GATC  sequences and the Dcm methyltransferase methylates both C    residues in CCjGG at their C5 positions. Note that both of these  sequences are palindromic.   In addition to its role in restriction-modification systems, DNA  methylation in prokaryotes functions most conspicuously as a  marker of parental DNA in the repair of mismatched base pairs.  Any replicational mispairing that has eluded the editing functions  of Pol I and Pol III may still be corrected by a process known as  mismatch repair (Section 25-5D). However, if this system is to  correct errors rather than perpetuate them, it must distinguish the  parental DNA, which has the correct base, from the daughter  strand, which has the incorrect although normal base. The obser¬  vation that E. coli cells with a deficient Dam methyltransferase  have higher mutation rates than wild-type bacteria suggests how  this distinction is made. A newly replicated daughter strand is  undermethylated compared to the parental strand because DNA  methylation lags behind DNA synthesis.   5-Methylcytosine is the only methylated base in most eukary¬  otic DNAs, including those of vertebrates. This modification occurs  largely in the CG dinucleotide of various palindromic sequences.  CG is present in the vertebrate genome at only about one-fifth its  randomly expected frequency. The upstream regions of many  genes, however, have normal CG frequencies and are therefore  known as CpG islands.   There is clear evidence that DNA methylation switches off  eukaryotic gene expression, particularly when it occurs in the pro¬  moter regions upstream of a gene’s transcribed sequence (Section  28-3A). For example, globin genes are less methylated in erythroid  cells than they are in nonerythroid cells and, in fact, the specific  methylation of the control region in a recombinant globin gene  inhibits its transcription. Moreover, the methylation pattern of a  parental DNA strand directs the methylation of its daughter strand  (a methylated CG sequence directs a methyltransferase to meth¬  ylate its complementary CG sequence), so that the “inheritance” of  a methylation pattern in a cell line permits all the cells to have the  same differentiated phenotype. Variations in methylation are  responsible for genomic imprinting in mammals, the phenomenon  in which certain maternal and paternal genes are differentially  expressed in the offspring (Section 28-3A).    metabolism, for example, by causing cancer. As many as 80% of human cancers  may be caused by carcinogens that damage DNA or interfere with its replication  or repair. Consequently, many mutagens are also carcinogens.   There are presently over 80,000 man-made chemicals of commercial im¬  portance, and —1000 new ones are introduced every year. The standard ani¬  mal tests for carcinogenesis, exposing rats or mice to high levels of the  suspected carcinogen and checking for cancer, are expensive and require ~3  years to complete. Thus, relatively few substances have been tested in that  manner. Likewise, epidemiological studies in humans are costly, time-consuming,  and often inconclusive.   Bruce Ames devised a rapid and effective bacterial assay for carcinogen¬  icity that is based on the high correlation between carcinogenesis and muta¬  genesis. He constructed special strains of the bacterium Salmonella typhimurium  that are his (cannot synthesize histidine and therefore cannot grow in its ab¬  sence). Mutagenesis in these strains is indicated by their reversion to the his +  phenotype. In the Ames test, ~10 9 test bacteria are spread on a culture plate    FIG. 25-31 The Ames test for mutagenesis. A filter paper disk containing a  mutagen, in this case the alkylating agent ethyl methanesulfonate, is centered on a culture  plate containing his~ strains of Salmonella typhimurium in a medium that lacks histidine. A  dense halo of revertant bacterial colonies appears around the disk from which the mutagen  diffused. The larger colonies distributed around the culture plate are spontaneous  revertants. The bacteria near the disk have been killed by the toxic mutagen’s high  concentration. [Courtesy of Raymond Devoret, Institut Curie, Orsay, France.]    Describe the appearance of a culture plate in which the disk contains a  substance that is not mutagenic.    that lacks histidine. A mutagen placed in the culture medium causes some  of the his bacteria to become his + so that they grow into visible colonies  after 2 days at 37°C (Fig. 25-31). The mutagenicity of a substance is scored as  the number of such colonies minus the few spontaneously revertant colonies  that occur in the absence of the mutagen. Dose-response curves, which are gen¬  erated by testing a given compound at a number of concentrations, are almost  always linear, indicating that there is no threshold concentration for mutagenesis.   About 80% of the compounds determined to be carcinogens in whole-  animal experiments are also mutagenic by the Ames test. Because many non¬  carcinogens are converted to carcinogens in the liver or in other tissues via a  variety of detoxification reactions (e.g., those catalyzed by the cytochromes  P450; Section 12-4D), a small amount of rat liver homogenate is therefore  included in the Ames test medium in order to approximate the effects of mam¬  malian metabolism. Alternatively, carcinogenicity testing may be carried out  using mammalian cell cultures rather than bacteria.   Several compounds to which humans had been extensively exposed that  were found to be mutagenic by the Ames test were later found to be carcino¬  genic in animal tests. These include tris(2,3-dibromopropyl)phosphate, which  was used as a flame retardant in childrens sleepwear in the mid-1970s and can  be absorbed through the skin; and furylfuramide, which was used in Japan in  the 1960s and 1970s as an antibacterial additive in many prepared foods (and  which had passed two animal tests before it was found to be mutagenic).   • Make a list of the environmental and  chemical agents that can alter DNA.   • Why is it practical to test carcinogens by  a mutagenesis assay?    • Some DNA damage can be repaired by the action of a single enzyme.   • Damaged bases can be removed and replaced by base excision repair.   • In nucleotide excision repair, a segment of one damaged DNA strand is removed  and replaced.    DNA damage must be repaired to maintain the integrity of genetic informa¬  tion. The biological importance of DNA repair is indicated by the great vari¬  ety of repair mechanisms in even simple organisms such as E. coli. These  systems include enzymes that simply reverse the chemical modification of nu¬  cleotide bases as well as more complicated multienzyme systems that depend  on the inherent redundancy of the information in duplex DNA to restore the  damaged molecule.   Several enzymes recognize and reverse certain types of DNA damage. For  example, pyrimidine dimers (Fig. 25-29) may be restored to their monomeric  forms by photoreactivation catalyzed by light-absorbing enzymes known as   DNA photolyases. These 55- to 65-kD monomeric enzymes are found in  many prokaryotes and eukaryotes but not in humans. Photolyases contain two  prosthetic groups: a light-absorbing cofactor and FADH . In the E. coli  enzyme, the cofactor 7V 5 ,A^ 10 -methenyltetrahydrofolate (Section 21-4D)  absorbs UV-visible light (300-500 nm) and transfers the excitation energy to  the FADH , which then transfers an electron to the pyrimidine dimer, thereby  splitting it. The resulting pyrimidine anion reduces the FADH* to regenerate  the enzyme.   The X-ray structure of the 474-residue DNA photolyase from the  cyanobacterium Anacystis nidulans in complex with a 9-bp dsDNA containing  a synthetic thymine dimer whose bridging phosphate group was replaced by  a —O—CH 2 —O— group (which does not affect the enzyme’s ability to split  the dimer) reveals how the enzyme splits the thymine dimer. The DNA binds  to a highly positively charged surface on the protein with its thymine dimer  flipped out of the double helix and bound in a deep cavity (Fig. 25-32). This  flip-out is probably facilitated by the relatively weak base-pairing interactions  of the thymine dimer and the distortions it imposes on the double helix. In  the following discussions we will see that this so-called base flipping (really  nucleotide flipping since entire nucleotides flip out of the double helix) is by  no means an unusual process for enzymes that perform chemistry on the bases  of dsDNA. The DNA outside of the thymine dimer assumes the B confor¬  mation but at the thymine dimer is bent by 50° away from the protein, thereby  unstacking the adenine bases complementary to the dimerized thymine bases.  The “hole” in the DNA helix left by the flipped out thymine dimer is par¬  tially occupied by an irregular protein ridge.   In the X-ray structure, the thymine dimers C5—C5 and C6—C6 bonds  are broken. Yet, in the crystal, the enzyme-bound thymine dimer is stable in  the dark for at least a year. Apparently, the X-rays used to generate the dif¬  fraction data mimic the effects of the light that normally ruptures these bonds.  Moreover, the FAD s isoalloxazine ring exhibits a 9° “butterfly” bend about its  N5-N10 axis (the isoalloxazine rings atomic numbering scheme is given in  Fig. 14-14), which indicates that it is in the fully reduced FADH form. The  isoalloxazine ring and adenine ring of the FAD , which has a folded confor¬  mation, are in van der Waals contact with one or the other bases of the  thymine dimer, and the isoalloxazine ring is ~10 A distant from the flavin-  like ring of the MTHF. This permits the observed efficient energy transfer in  the photolyase reaction (which has a quantum yield of ~0.9).   Another type of direct DNA repair is the reversal of base methylation  by alkyltransferases. For example, 0 6 -methylguanine and 0 6 -ethylguanine  lesions of DNA are repaired by 0 6 -alkylguanine-DNA alkyltransferase,  which directly transfers the offending methyl or ethyl group to one of its  own Cys residues. This reaction inactivates the protein, which therefore can¬  not be strictly classified as an enzyme. Apparently, the cost of sacrificing the  alkyltransferase is justified by the highly mutagenic nature of the modified  guanine residue.    FIG. 25-32 X-Ray structure of A. nidulans DNA photolyase in complex with  dsDNA containing a synthetic thymine dimer. The protein is drawn in worm  form embedded in its semitransparent surface diagram. The DNA, in which the  phosphate group bridging the nucleotides of the thymine dimer is replaced by a  —0—CH 2 —0— group, is drawn mainly in stick form but with the bases of the  thymine dimer in space-filling form, all colored according atom type (C green,   N blue, 0 red, and P orange) and with successive P atoms in each  polynucleotide chain connected by orange rods. The FAD and  A/ 5 ,A/ 10 -methenyltetrahydrofolate are drawn in stick form with their flavin and  flavinlike rings in space-filling form and with FAD C magenta and MTHF C  yellow. [Based on an X-ray structure by Thomas Carell, Ludwig Maximilians  University, Munich, Germany, and Lars-Oliver Essen, Philipps University, Marburg,  Germany. PDBid 1TEZ]    Damaged bases that cannot be directly repaired may be removed and replaced  in a process known as base excision repair (BER). This pathway, as its name  implies, begins with removal of the damaged base. Cells contain a variety of  DNA glycosylases that each cleave the glycosidic bond of a corresponding  type of altered nucleotide, leaving a deoxyribose residue with no attached base  (Fig. 25-33). Such apurinic or apyrimidinic sites (AP or abasic sites) also  result from the occasional spontaneous hydrolysis of glycosidic bonds. The  deoxyribose residue is then cleaved on one side by an AP endonuclease, the  deoxyribose and several adjacent residues are removed by the action of a cel¬  lular exonuclease (possibly associated with a DNA polymerase), and the gap  is filled in and sealed by a DNA polymerase and DNA ligase.   The enzymes of BER, which correct the most frequent type of DNA dam¬  age, include a glycosylase that recognizes 8-oxoguanine and the enzyme  uracil-DNA glycosylase (UDG), which excises uracil residues. The latter arise  from cytosine deamination as well as the occasional misincorporation of uracil  instead of thymine into DNA (Box 25-5).   The X-ray structure of human UDG in complex with a 10-bp DNA  containing a U • G mismatch (which forms a doubly hydrogen-bonded base  pair whose shape differs from that of Watson-Crick base pairs; Section 24-1 A),  determined by John Tainer, reveals that the enzyme has bound the DNA with  the U • G base pairs uridine nucleotide flipped out of the double helix  (Fig. 25-34). Moreover, the enzyme has hydrolyzed the uridines glycosidic  bond, yielding the free uracil base and an AP site on the DNA, although both   These enzymes hydrolyze the glycosidic bond of  their corresponding altered base (red) to yield an  AP site.   FIG. 25-34 X-Ray structure of a complex of human uracil-DNA  glycosylase with a 10-bp DNA containing a U • G base pair. The   protein (the C-terminal 223 residues of the 304-residue monomer) is  represented by its ribbon diagram embedded in its transparent molecular  surface. The DNA, viewed looking into its major groove, is drawn in stick  form colored according to atom type (C green, N blue, 0 red, and P  orange). The U • G base pair’s uridine nucleotide has flipped out of the  double helix (to the right of the DNA) and has been hydrolyzed to yield an  AP site (stick form with C cyan) and uracil (space-filling form with C cyan),  which remains bound in the enzyme’s binding pocket. The side chain of Arg  272 (space-filling form with C yellow) has intercalated into the DNA base  stack to fill the space vacated by the flipped-out uracil base. [Based on an  X-ray structure by John Tainer, The Scripps Research Institute, La Jolla,  California. PDBid 4SKN.]   Three of the deoxynucleotide bases in DNA (adenine, guanine,  and cytosine) also occur as ribonucleotide bases in RNA. The  fourth deoxynucleotide base, thymine, is synthesized—at consider¬  able metabolic effort (Section 23-3B)—from uracil, which occurs in  RNA. Since uracil and thymine have identical base-pairing proper¬  ties, why do cells bother to synthesize thymine at all?   This enigma was solved by the discovery of cytosine’s pen¬  chant for conversion to uracil by deamination, either spontaneously  or by reaction with nitrites (Section 25-4A). If U were a normal  DNA base, the deamination of C would be highly mutagenic  because there would be no indication of whether the resulting    mismatched G • U base pair had initially been G • C or A • U. Since  T is DNA’s normal base, however, any U in DNA is almost certainly a  deaminated C and can be removed by uracil-DNA glycosylase.   Uracil-DNA glycosylase also has an important function in DNA  replication. dUTP, an intermediate in dTTP synthesis, is present in  all cells in small amounts. DNA polymerases do not discriminate  well between dUTP and dTTP, both of which can base-pair with  template A’s. Consequently, newly synthesized DNA contains an  occasional U. These U’s are rapidly replaced by T through base  excision repair.   products remain bound to the enzyme. The cavity in the DNAs base stack that  would otherwise be occupied by the flipped-out uracil is filled by the side chain  of Arg 272, which intercalates into the DNA from its minor groove side.   How does UDG detect a base-paired uracil in the center of DNA and  how does it discriminate so acutely between uracil and other bases, particu¬  larly the closely similar thymine? The X-ray structure indicates that the phos¬  phate groups flanking the flipped-out base are 4 A closer together than they  are in B-DNA (8 A versus 12 A), which causes the DNA to kink by ~45° in  the direction parallel to the view in Fig. 25-34. Tainer has postulated that  UDG rapidly scans a DNA molecule for uracil by periodically binding to it  so as to compress and thereby slightly bend the DNA backbone. The DNA  bends more readily at a uracil-containing site (a U • G base pair is smaller  than C • G and hence leaves a space in the base stack, whereas a U • A base  pair is even weaker than T • A), permitting the enzyme to flip out the uracil  by inserting Arg 272 into the minor groove. The exquisite specificity of UDG s  binding pocket for uracil prevents the binding and hydrolysis of any other  base that the enzyme might have induced to flip. Thus the overall shapes of  adenine and guanine exclude them from this pocket, whereas thymines 5-  methyl group is sterically blocked by the rigidly held side chain of Tyr 147.  Cytosine, which has approximately the same shape as uracil, is excluded  through a set of hydrogen bonds emanating from the protein that mimic those  made by adenine in a Watson-Crick A • U base pair (dUTPase discriminates  similarly between uracil and other bases; Fig. 23-14).   AP sites in mammalian DNA are highly cytotoxic because they irreversibly  trap mammalian topoisomerase I in its covalent complex with DNA (Section  24-ID). Moreover, since the ribose at the AP site lacks a glycosidic bond, it  can readily convert to its linear form (Section 8-IB), whose reactive aldehyde  group can cross-link to other cell components. This rationalizes why AP sites  remain tightly bound to UDG in solution. UDG activity is enhanced by AP  endonuclease, the next enzyme in the base excision repair pathway, but the  two enzymes do not interact in the absence of DNA. This suggests that UDG  remains bound to an AP site it generated until it is displaced by the more  tightly binding AP endonuclease, thereby protecting the cell from the AP site’s  cytotoxic effects. Its seems likely that other damage-specific DNA glycosylases  function similarly.   C Nucleotide Excision Repair Removes a Segment of a DNA Strand   All cells have a more elaborate pathway, nucleotide excision repair (NER),  to correct pyrimidine dimers and other DNA lesions in which the bases are  displaced from their normal position or have bulky substituents. The NER  system appears to respond to helix distortions rather than by the recognition  of any particular group. In humans, NER is the major defense against two  important carcinogens, sunlight and tobacco smoke.   In E. coli, NER is carried out in an ATP-dependent process through the  actions of the UvrA, UvrB, and UvrC proteins (the products of the uvrA, uvrB ,  and uvrC genes). This system, which is often referred to as the UvrABC  endonuclease (although there is no complex that contains all three subunits),  cleaves the damaged DNA strand at the seventh and at the third or fourth phos-  phodiester bonds from the lesions 5 ' and 3' sides, respectively (Fig. 25-35). The  excised 11- or 12-nt oligonucleotide is then displaced by the binding of UvrD  (also called helicase II) and is replaced through the actions of Pol I and  DNA ligase.   Genetically Defective NER. In eukaryotes, NER requires at least 16  proteins and removes oligonucleotides of ~30 residues. The proteins are con¬  served from yeast to humans but none of them exhibit any sequence similarity    FIG. 25-35 The mechanism of nucleotide  excision repair (NER) of pyrimidine dimers.    to the prokaryotic proteins, suggesting that the two NER systems arose by  convergent evolution. Many of the enzymes involved in the human pathway  have been identified through mutations that are manifested as two genetic  diseases. The inherited disease xeroderma pigmentosum (XP; Greek: xeros,  dry + derma , skin) is mainly characterized by the inability of skin cells to  repair UV-induced DNA lesions. Individuals suffering from this autosomal  recessive condition are extremely sensitive to sunlight. During infancy they de¬  velop marked skin changes such as dryness, excessive freckling, and keratoses  (a type of skin tumor; the skin of these children is described as resembling  that of farmers with many years of sun exposure), together with eye damage,  such as opacification and ulceration of the cornea. Moreover, they develop  often fatal skin cancers at an ~2000-fold greater rate than normal and inter¬  nal cancers at a 10- to 20-fold increased rate. Curiously, many individuals with  XP also have a bewildering variety of seemingly unrelated symptoms includ¬  ing progressive neurological degeneration and developmental deficits.   Cockayne syndrome (CS), a rare inherited disease that is also associated  with defective NER, arises from defects in three of the same genes that are  defective in XP as well as in two additional genes. Individuals with CS are  hypersensitive to UV radiation and exhibit stunted growth, neurological  dysfunction due to neuron demyelination, and the appearance of premature  aging but, intriguingly, have a normal incidence of skin cancer. The proteins  that are defective in CS normally recognize an RNA polymerase whose  progress has been halted by a damaged or distorted DNA template. In order  for transcription to resume, the stalled RNA polymerase must be removed so  that the DNA damage can be repaired by the NER system. In CS, the DNA  cannot be repaired, which causes the cell to undergo apoptosis (programmed  cell death; Section 28-4C). The death of transcriptionally active cells may  account for the developmental symptoms of Cockayne syndrome.   Any replicational mispairing that has eluded the editing functions of the DNA  polymerases may still be corrected by a process known as mismatch repair    FIG. 25-36 The mechanism of mismatch repair in E. coli. (1) MutS binds.   (2) MutL binds. (3) The mismatch-containing DNA loops out. (4) MutH endonuclease binds.  (5) The unmethylated DNA strand containing the mismatched base is excised.    (MMR). The MMR system can also correct insertions or deletions of up to  four nucleotides (which arise from the slippage of one strand relative to the  other in the active site of DNA polymerase). The importance of mismatch  repair is indicated by the fact that defects in the human mismatch repair  system result in a high incidence of cancer, most notably hereditary nonpoly¬  posis colorectal cancer syndrome, which affects several organs and may be  the most common inherited predisposition to cancer.   Mismatch repair in E. coli is carried out by three proteins (Fig. 25-36).  A homodimer of MutS binds to a mismatched base pair, or to unpaired bases,  and then binds a homodimer of MutL. Then, in an ATP-dependent process,  the resulting MutS 2 MutL 2 complex translocates along the DNA in both  directions, thereby causing the duplex DNA to form a mismatch-containing  loop that is closed by the MutS 2 MutL 2 complex. If an MMR system is to cor¬  rect errors in replication rather than perpetuate them, it must distinguish the  parental DNA, which has the correct base, from the daughter strand, which  has an incorrect although normal base. The mismatch repair system in E. coli  can distinguish the two DNA strands because newly replicated DNA remains  hemimethylated until methyltransferases have had sufficient time to methylate  the daughter strand (Box 25-4). On encountering a hemimethylated GATC  palindrome, the MutS 2 MutL 2 complex recruits MutH and activates this  endonuclease to make a nick on the 5' side of the unmethylated GATC  sequence. This site may be located on either side of the mismatch and over  1000 bp distant from it. The UvrD helicase, which also participates in NER,  separates the parental and daughter strands. An exonuclease completely  removes the defective daughter segment, which is then replaced by the action  of DNA polymerase III.   Eukaryotic cells contain several homologs of MutS and MutL but not of  MutH and must therefore use some other cue than methylation status to  differentiate the daughter and parental strands. One possibility is that a newly  synthesized daughter strand is identified by its as yet unsealed nicks between  Okazaki fragments. DNA resynthesis is probably mediated by pol 8.   Individuals with a variant form of XP exhibit increased rates of skin cancer  even though their NER proteins are normal. The defect in this disorder has  been pinpointed to an enzyme known as DNA polymerase r\ (pol T|). When  functioning normally, pol T| can bypass DNA lesions such as UV-induced  thymine dimers, incorporating two adenine bases in the new strand. Although  it is useful as a translesion polymerase, pol T| is relatively inaccurate and has  no proofreading exonuclease activity: It inserts an incorrect base on average  every 30 nucleotides.   The existence of error-prone polymerases provides a fail-safe mechanism  for replicating stretches of DNA that cannot be navigated by the standard  replication machinery. In fact, such alternative DNA polymerases typically out¬  number the polymerases devoted to normal replication. For example, eukary¬  otes contain at least five such “bypass” polymerases that act with different  degrees of accuracy. However, the errors generated by bypass polymerases may  still be corrected by the mismatch repair system.    End-Joining Repairs Double-Strand Breaks. Both the base excision repair  and nucleotide excision repair pathways act when a lesion affects only one   strand of DNA. However, DNA is susceptible to double-strand  breaks (DSBs) generated by ionizing radiation or the free radical  by-products of oxidative metabolism, as well as when the replica¬  tion fork encounters a nick. In fact, around 5 to 10% of divid¬  ing cells in culture exhibit at least one chromosome break at any  given time. Unrepaired DSBs can be lethal to cells or cause chro¬  mosomal aberrations that may lead to cancer. Hence the efficient  repair of DSBs is essential for cell viability and genomic integrity.   Cells have two general mechanisms to repair DSBs: recombi¬  nation repair and nonhomologous end-joining (NHEJ). Here we  discuss NHEJ, a process which, as its name implies, directly rejoins  DSBs. The recombination repair of DSBs is discussed in Section 25-6B.   In NHEJ, the ends of the broken DNA must be aligned, frayed ends  trimmed off or filled in, and the strands ligated. The core of the end-join¬  ing machinery in eukaryotes includes the protein Ku (a heterodimer of ho¬  mologous 70- and 83-kD subunits, Ku70 and Ku80), which appears to be  the cells broken-DNA sensor. The X-ray structure of Ku in complex with a  14-bp DNA, determined by Jonathan Goldberg, reveals that the protein cra¬  dles the dsDNA segment along its entire length and encircles its central —3  bp segment (Fig. 25-37). Ku makes no specific contacts with the DNAs bases  and few with its backbone but instead fits snugly into the major and minor  grooves. Ku-DNA complexes dimerize so as to align both halves of a double¬  strand break while leaving the strand ends accessible to nucleases, polymerases,  and ligases. Nucleotide trimming, of course, generates mutations, but an  unrepaired double-strand break would be even more detrimental to the cell.   The reason that the mutations generated by NHEJ are usually not unac¬  ceptably deleterious is that only a small fraction of the mammalian genome is  expressed (Section 28-1 A). In fact, the genome in a somatic cell of a 70-year-  old human typically contains —2000 “scars” caused by NHEJ.    FIG. 25-37 X-Ray structure of human Ku  protein in complex with a 14-bp DNA. The   subunits of Ku70 (red helices and yellow  strands) and Ku80 (blue helices and green (3  strands) are viewed along the pseudotwofold axis  relating them. The DNA is drawn in space-filling  form with its sugar-phosphate backbone dark  gray and its base pairs light gray. Note that the  DNA is surrounded by a ring of protein.   [Courtesy of John Tainer, The Scripps Research  Institute, La Jolla, California. Based on an X-ray  structure by Jonathan Goldberg, Memorial Sloan-  Kettering Cancer Center, New York, New York.  PDBid 1JEYJ    The E. coli SOS Response Is Mutagenic. In E. coli , agents that damage  DNA induce a complex system of cellular changes known as the SOS response.  Cells undergoing the SOS response cease dividing and increase their capacity  to repair damaged DNA. LexA, a repressor, and RecA, a DNA-binding pro¬  tein, regulate the activity of this system. (RecA is also a key player in homol¬  ogous recombination, discussed in the following section, which offers a means  for repairing damaged DNA after it has been replicated.) During normal  growth, LexA represses SOS gene expression. However, when DNA is dam¬  aged (and cannot fully replicate), the resulting single strands bind to RecA to  form a complex that activates LexA to cleave and thereby inactivate itself. The  SOS genes, which include recA and lexA as well as the NER genes uvrA and  uvrB (Section 25-5C), are thereby expressed. On the repair of the DNA, the  DNA • RecA complex is no longer present, so the newly synthesized LexA  again represses the expression of the SOS genes.   Among the 43 genes controlled by the SOS system are DNA polymerases  IV and V, both of which replicate DNA with low fidelity and lack proofread¬  ing exonuclease activity. Because these translesion polymerases can synthesize  DNA even when there is no information as to which bases were originally  present, the SOS repair system is error-prone and consequently mutagenic. It is  therefore a process of last resort that is initiated only —50 min after SOS  induction if the DNA has not already been repaired by other means. Hence,  the SOS repair system is a testimonial to the proposition that survival with a  chance of loss of function (and the possible gain of a new one) is advanta¬  geous, in the Darwinian sense, over death, although only a small fraction of  cells survive this process. It has therefore been suggested that, under condi¬  tions of environmental stress, the SOS system functions to increase the rate  of mutation so as to increase the rate at which the E. coli adapt to the new  conditions.   • Describe the enzymatic activities required  for base excision repair, nucleotide  excision repair, and mismatch repair.   • What are the symptoms of defective  nucleotide excision repair and mismatch  repair?   • Describe why cells require error-prone  repair systems such as pol r|, nonhomolo¬  gous end-joining, and the SOS response.    • In homologous recombination, DNA strands cross over to exchange places.   • Transposons move themselves and sometimes additional genes to new chromoso¬  mal locations.    FIG. 25-38 Crossing-over, (a) An electron  micrograph and (b) its interpretive drawing of  two homologous pairs of chromatids during  meiosis in the grasshopper Chorthippus parallels.  Nonsister chromatids (different colors ) may  recombine at any of the points where they cross  over. [© Science VU/B. John/Visuals Unlimited/  © Corbis.]    FIG. 25-39 The Holliday model of  general recombination between homologous  DNA duplexes. Q See the Animated Figures.    Over the years, genetic studies have shown that genes are not immutable. In  higher organisms, pairs of genes may exchange by crossing-over when homol¬  ogous chromosomes are aligned (Fig. 25-38). Bacteria, which do not contain  duplicate chromosomes, also have an elaborate mechanism for recombining  genetic information. In addition, foreign DNA can be installed in a hosts  chromosome through recombination. In this section, we examine the molec¬  ular events of recombination and discuss the biochemistry of transposons,  which are mobile genetic elements.   Homologous recombination (also called general recombination) occurs be¬  tween DNA segments with extensive homology; site-specific recombination  occurs between two short, specific DNA sequences. The prototypical model  for homologous recombination (Fig. 25-39) was proposed by Robin Holliday  in 1964. The corresponding strands of two aligned homologous DNA duplexes   secondary structure of the four-stranded Holliday junction formed by  the palindromic sequence d(CCGGTACCGG) in which the four strands,  A, B, C, and D, are individually colored, their nucleotides are numbered  1 to 10 from their 5' to 3' termini, and Watson-Crick base-pairing  interactions are represented by black dashes. The twofold axis relating  the two helices is represented by the black lenticular symbol, (b) The  three-dimensional structure of the Holliday junction, as viewed along    its twofold axis, in which the oligonucleotides are represented in stick  form with their backbones traced by ribbons, all colored as in Part a.  With the exception of the backbones of strands B and D at the  crossovers, the two arms of this structure each form an undistorted  B-DNA helix, including the stacking of the base pairs flanking the  crossovers. Note that Fig. 25-39 g is a schematic representation of  this structure as viewed from the side in this drawing. [Courtesy of  Shing Ho, Oregon State University. PDBid 1 DCW.]    are nicked, and the nicked strands cross over to pair with the nearly comple¬  mentary strands on the homologous duplex, thereby forming a segment of  heterologous DNA, after which the nicks are sealed (Fig. 25-39 a—e). The  crossover point is a four-stranded structure known as a Holliday junction. A  Holliday junction has, in fact, been observed in the X-ray structure of the  self-complementary d(CCGGTACCGG), determined by Shing Ho (Fig. 25-40).  The crossover point can move in either direction, often thousands of  nucleotides, in a process known as branch migration (Fig. 25-39 e, f).   1. Cleavage of the strands that did not cross over exchanges the ends of  the original duplexes to form, after nick sealing, the traditional recom¬  binant DNA molecule (right branch of Fig. 25-39 ]—l).   2. Cleavage of the strands that crossed over exchanges a pair of homolo¬  gous single-stranded segments (left branch of Fig. 25-39 j—l).   RecA Promotes Recombination in E. coli. The observation that recA~ E. coli  have a 10 4 -fold lower recombination rate than the wild-type indicates that  RecA protein has an important function in recombination. Indeed, RecA greatly  increases the rate at which complementary strands renature in vitro. This ver¬  satile protein (recall it also stimulates the autoproteolysis of LexA to trigger  the SOS response; Section 25-5E) polymerizes cooperatively without regard to  base sequence on ssDNA or on dsDNA that has a single-stranded gap. The  resulting filaments, which may contain up to several thousand RecA  monomers, specifically bind the homologous dsDNA, and, in an ATP-dependent  reaction, catalyze strand exchange.   EM studies by Edward Egelman revealed that RecA filaments bound to  ssDNA or dsDNA form a right-handed helix with ~6.2 RecA monomers per  turn and a pitch (rise per turn) of 95 A. The DNA in these filaments binds  to the protein with 3 nt (or bp) per RecA monomer and hence is underwound  with —18.5 nt (or bp) per turn (versus 10 bp per turn for canonical B-DNA).   Chapter 25 DNA Replication, Repair, and The formation of RecA-DNA filaments is highly cooperative; it requires   Recombination five or six RecA protomers to form a stable assembly. Consequently, attempts   to crystallize RecA-DNA filaments over many years were unsuccessful. Nikola  Pavletich ingeniously solved this conundrum by linking five E. coli RecA genes  (each corresponding to residues 1-335 of this 353 -residue protein) in tandem  via 14-residue linkers and mutating the first and last RecA so as to prevent  them from forming longer filaments. This fusion protein, which has DNA-  dependent ATPase and strand-exchange activities comparable to that of  monomeric RecA, formed crystals containing both ssDNA and dsDNA.   The X-ray structure of the RecA 5 -(ADP-AlF 4 ) 5 -(dT) 15 -(dA ) 12 complex  (Fig. 25-41; ADP-AIF 4 is a nonhydrolyzable ATP analog) exhibits a straight  filament axis with overall helical parameters that are closely similar to those  derived from EM studies. Each RecA unit consists of a largely helical 30-residue  N-terminal segment, a 240-residue a/(3 ATPase core, and a 64-residue C-  terminal globular domain. The linkers connecting adjacent RecA units are dis¬  ordered. Each RecA unit makes extensive contacts with its nearest neighbors  so as to form a filament with a deep helical groove that exposes the DNA  bound in its interior (Fig. 25-41 a is viewed looking into this groove).   The two DNA strands, which lie close to the filament axis, form a complete  set of Watson-Crick base pairs (Fig. 25-41 £). However, rather than being  smoothly stretched out, as had been expected, the dsDNA assumes an irreg¬  ular conformation in which each 3-bp segment that is bound to a RecA unit  closely resembles B-DNA with the steps between successive base pairs in this  triplet having an axial rise of ~ 3 .4 A and a helical twist of ~30° (versus 3.4  A and 36° for canonical B-DNA; Table 24-1). In contrast, the step between  successive base pair triplets has an axial rise of 8.4 A and a helical twist of  —4°, thereby forming a 5 -A-high gap between successive triplets that is partially   FIG. 25-41 X-ray structure of the  RecA 5 -(ADP-AIFT) 5 -(dT) 15 -(dA) 12 complex viewed with its  filament axis vertical, (a) The RecA units RecA 1 (the N-terminal unit)  through RecA 4 are colored green, cyan, magenta, and gray,  respectively, with the C-terminal unit, RecA 5 , colored in rainbow order  from its N-terminus (blue) to its C-terminus (red). The DNA and  ADP-AIFT are drawn in space-filling form with DNA C gray, ADP C    green, N blue, 0 red, P orange, F light blue, and Al purple, (b) The  DNA in the complex, viewed and colored as in Part a, but with C  atoms of the (dT) 15 green. [Based on an X-ray structure by Nikola  Pavletich, Memorial Sloan-Kettering Cancer Center, New York, New  York. PDBid 3CMX.]   FIG. 25-42 A model for RecA-mediated pairing. (1) A ssDNA (3) RecA rotates the bases of the aligned homologous strands to  binds to RecA to form an initiation complex. (2) dsDNA binds to the effect strand exchange in an ATP-driven process. [After West, S.C.,   initiation complex so as to transiently form a three-stranded helix. Annu. Rev. Biochem. 61, 618 (1992).]    filled by the side chain of the conserved lie 199. The sugar—phosphate  backbone of the (dT ) 15 (the DNA strand furthest from the viewer in Fig.  25-41) makes extensive contacts with RecA. In contrast, the other strand [the  (dA) 12 ] makes few contacts with the protein; it is held in place almost entirely  by base pairing with the first strand. The ADP-A1F 4 _ is sandwiched between  adjacent a/(3 ATPase cores, where it is completely buried.   The X-ray structure of the ssDNA-containing RecA 6 -(ADP-AlF 4 ) 6 —  (dT ) 18 complex closely resembles that of the foregoing dsDNA-containing  complex but with the absence of the DNA strand closest to the viewer in  Fig. 25-41. Thus, each RecA unit binds a (dT ) 3 segment that is held in a  B-DNA-like conformation with successive (dT ) 3 segments separated by a  7 .8-A axial rise.   How does RecA mediate DNA strand exchange between ssDNA and  dsDNA? On encountering a duplex DNA with a strand that is complemen¬  tary to its bound ssDNA, RecA partially unwinds the duplex and exchanges  the ssDNA with the corresponding strand on the dsDNA, a process that  involves a three-stranded DNA intermediate (Fig. 25-42). ATP binding to RecA  is required for strand exchange, but ATP hydrolysis does not occur until the  dissociation step. The strand-exchange process tolerates only a limited degree  of mispairing and therefore occurs only between homologous DNA segments.  As the RecA filament rotates about its axis, the duplex DNA is spooled in  (Fig. 25-43). Presumably, by keeping most of its bound ssDNA in the B con¬  formation, RecA prevents the formation of non-Watson—Crick base pairings.   Homologous DNA molecules are paired in advance of strand  exchange in a three-stranded helix. The ATP-driven rotation of the    RecA filament ( purple ) around its helix axis causes duplex DNA to be  “spooled in” to the filament, left to right as drawn. [After West, S.C.,  Annu. Rev. Biochem. 61, 617 (1992).]    Two such strand-exchange processes must occur simultaneously in a Holliday  junction. In eukaryotes, the protein Rad51, which is homologous to E. coli  RecA, apparently functions in a similar ATP-dependent manner to mediate  DNA recombination.   E. coli, the single-strand segments to which RecA binds are made by the  RecBCD protein, the 330-kD heterotrimeric product of the SOS genes recB,  recC, and recD, which has both helicase and nuclease activities (Fig. 25-44).  The process begins with RecBCD binding to the end of a dsDNA and then  unwinding it via its ATP-driven helicase function. As it does so, it nucleo-  lytically degrades the unwound single strands behind it, with the 3'-ending strand  being cleaved more often and hence broken down to smaller fragments than  the 5'-ending strand. However, on encountering the sequence GCTGGTGG  from its 3' end (the so-called Chi sequence, which occurs about every 5 kb  in the E. coli genome), it stops cleaving the 3'-ending strand and increases the  rate at which it cleaves the 5'-ending strand, thereby yielding the 3'-ending  single-strand segment to which RecA binds. This explains the observation that  regions containing Chi sequences have elevated rates of recombination. RecB  also functions to recruit RecA and nucleate its binding to the 3'-ending strand.   Dale Wigley determined the X-ray structure of E. coli RecBCD in com¬  plex with a 51-nt DNA that could form a hairpin loop containing an up to  21-bp dsDNA stem (Fig. 25-45). The structure shows that RecB (1180  residues) and RecC (1122 residues) are intimately intertwined, with RecBs C-  terminal nuclease domain connected to the rest of the subunit by an extended  21-residue polypeptide tether. A 15-bp segment of dsDNA enters the protein  through a tunnel between RecB and RecC. There it encounters a loop from  RecC that appears to wedge the two strands apart, with the 6-nt 3'-ending  single strand of the DNA binding to RecB and the 10-nt 5'-ending single  strand binding to RecD (608 residues; the 5-nt loop connecting the two  strands of the dsDNA at the top of Fig. 25-45 is disordered).   The structure explains the different rates of cleavage of the two DNA  strands. The 3'-ending strand emerges from a tunnel through RecC in the  vicinity of the RecB nuclease domain, which is positioned to processively  cleave it. The 5'-ending strand competes with the 3'-ending strand for the  nuclease site, but since the 5'-ending strand is less favorably located, it is  cleaved less frequently. However, after RecC has bound a Chi sequence, the  3 '-ending strand is no longer available for cleavage, which permits the nucle¬  ase to cleave the 5'-ending strand more frequently.    FIG. 25-44 The generation of a 3'-ending  single-strand DNA segment by RecBCD to  initiate recombination. (1) RecBCD binds to a  free end of a dsDNA and, in an ATP-driven  process, advances along the helix, unwinding the  DNA and degrading the resulting single strands  behind it, with the 3'-ending strand cleaved more  often than the 5'-ending strand. (2) When  RecBCD encounters a properly oriented Chi  sequence, it binds it and thus stops cleaving the  3'-ending strand but increases the frequency at  which it cleaves the 5'-ending strand. This  generates the potentially invasive 3'-ending  strand segment to which RecA binds.    FIG. 25-45 X-Ray structure of E. coli RecBCD in complex with a 51-nt DNA  capable of forming a 21-bp hairpin loop. The protein is drawn in semitransparent ribbon  form with RecB yellow, RecC cyan, and RecD magenta. Note how the RecB nuclease  domain is linked to the rest of the subunit by an extended polypeptide tether. The DNA is  shown in space-filling form with C gray, N blue, 0 red, and P orange. A loop from RecC,  which is drawn in space-filling form in green, is situated so as to wedge apart the two  strands of the incoming dsDNA with the 3'-ending strand binding to the 3' —» 5' helicase  of RecB and the 5'-ending strand passing through RecC to bind to the 5' —» 3' helicase of  RecD. [Based on an X-ray structure by Dale Wigley, The London Research Institute, Herts,  U.K. PDBid 3K70.]    RecBCD can commence unwinding DNA only at a free duplex end. Such  ends are not normally present in E. coli, which has a circular genome, but  become available during recombinational processes.   RuvABC Mediates Branch Migration and the Resolution of the Holliday  Junction. Branch migration requires the proteins RuvA and RuvB. The X-ray  structure of Mycobacterium leprae (the cause of leprosy) RuvA in complex with  a Holliday junction, determined by Kosuke Morikawa, indicates that RuvA  forms a homotetramer with the appearance of a four-petaled flower (C 4 sym¬  metry) and is relatively flat (80 X 80 X 45 A) with one face concave and the  other convex (Fig. 25-46). The concave face (facing the viewer in Fig. 25-46),  which is highly positively charged and is studded with numerous conserved  residues, has four symmetry-related grooves that bind the Holliday junctions  four arms. Four negatively charged projections or “pins” are centrally located.  The repulsive forces between the pins and the Holliday junctions anionic phos¬  phate groups probably facilitate the separation of the ssDNA segments and  guide them from one double helix to another. However, this is not the entire  story. The X-ray structure of a RuvA-Holliday junction complex crystallized  under different conditions than that in Fig. 25-46 resembles the complex in  Fig. 25-46 but with a second RuvA tetramer in face-to-face contact with the  first. Here the Holliday junction is contained in two intersecting tunnels run¬  ning through the resulting ^-symmetric RuvA octamer.   In the presence of dsDNA, RuvB, an AAA+ family ATPase, oligomerizes  to form a pseudohexameric ring with a 30-A-diameter hole through which a  dsDNA can be threaded, much like a hexameric helicase (Section 25-2B). A  model for the RuvA • RuvB • Holliday junction complex, based on X-ray   FIG. 25-46 X-Ray structure of a  RuvA-Holliday junction complex. The protein  tetramer, which is viewed along its fourfold axis,  is represented by its molecular surface with its  subunits alternately colored pink and light blue.  The DNA is drawn in space-filling form colored  according to atom type with C atoms in different  chains in different colors, N blue, 0 red, and P  orange. [Based on an X-ray structure by Kosuke  Morikawa, Biomolecular Engineering Research  Institute, Osaka, Japan. PDBid 1C7Y.]   The model is based on X-ray structures and on electron micrographs  such as that in the inset. The proteins are represented by their sur¬  face diagrams with the RuvA tetramer green and the two oppositely  oriented RuvB hexamers white. The DNA of the Holliday junction is  drawn in space-filling form with its homologous blue and magenta  strands complementary to its red and gray strands. A second RuvA  tetramer, which overlies the RuvA tetramer shown and thus    sandwiches the DNA in two perpendicular tunnels, has been removed  to show the path of the DNA. The complex is postulated to drive  branch migration via the ATP-driven movement of loops projecting  from the center of the RuvB hexamers along the DNA. This pumps  the horizontal dsDNAs through the RuvB hexamers to the center of  the Holliday junction, where the strands separate and then base-pair  with their homologs to form new dsDNAs, which are pumped out  vertically. [Courtesy of Peter Artymiuk, University of Sheffield, U.K.]    structures and electron micrographs, is shown in Fig. 25-47. In the complex,  two RuvB hexamers encircle the DNA on opposite sides of a RuvA octamer.  This arrangement suggests a mechanism for branch migration in which RuvBs  DNA-contacting loops unidirectionally “walk” along the horizontal dsDNAs  grooves in an ATP-dependent process resembling that proposed for the El  helicase (Section 25-2B. This pulls the dsDNAs through the center of each  RuvB hexamer toward the RuvA octamer. The DNA strands of the Holliday  junction are thereby pushed through the center of RuvA, where they rejoin to  form the vertical dsDNAs and exit the complex. (An alternative model, in  which the two RuvB hexamers counterrotate so as to screw the dsDNA into  the RuvA octamer, seems unlikely due to the large contact area between RuvA  and RuvB.) The direction of branch migration depends on which pair of op¬  posing arms of the Holliday junction the RuvB hexamers are loaded onto.   The final stage of homologous recombination is the resolution of the  Holliday junction into its two homologous dsDNAs. This process is carried  out by RuvC, a homodimeric nuclease. One of the RuvA tetramers must dis¬  sociate from the octameric RuvA complex to expose the Holliday junction  DNA. RuvC presumably sits down on the open face of the resulting  RuvA-Holliday junction complex (the side visible in Fig. 25-46) to cleave  oppositely located strands at the Holliday junction (Figs. 25-39/, j). The re¬  sulting single-strand nicks are then sealed by DNA ligase (Figs. 25-39£, /).   In haploid organisms such as bacteria, homologous recombination between  chromosomal DNA and exogenously supplied DNA as occurs, for example,  in transformation (Section 3-3A) is such a rare event that the vast majority of  such cells never participate in this process. Similarly, in multicellular organ¬  isms, the only time that gene shuffling through homologous recombination  occurs is during meiosis (Fig. 25-38), which takes place only in germline cells.  Why then do nearly all cells have elaborate systems for mediating homologous  recombination? This is because damaged replication forks occur at least once  per bacterial cell generation and perhaps ten times per eukaryotic cell cycle.  The underlying DNA lesions can be corrected via homologous recombination  in a process named recombination repair [error-prone repair (Section 25-5E),    which is highly mutagenic, is a process of last resort]. Indeed, the rates of syn¬  thesis of RuvA and RuvB are greatly enhanced by the SOS response. Thus, as  Michael Cox pointed out, the primary function of homologous recombination is  to repair damaged replication forks.   1. When a replication fork encounters an unrepaired single-strand nick  or gap, the fork collapses (falls apart).    FIG. 25-48 The recombination repair of a replication fork  that has encountered a single-strand nick. Thick lines indicate  parental DNA, thin lines indicate newly synthesized DNA, and the    arrows point in the 5' —> 3' direction. [After Cox, M.M., Annu. Rev.  Genet. 35, 53 (2001).]   The repair process begins via the RecBCD + RecA—mediated invasion  of the newly synthesized and undamaged 3 r -ending strand into the ho¬  mologous dsDNA starting at its broken end.   Branch migration, as mediated by RuvAB, then yields a Holliday junc¬  tion, which exchanges the replication forks 3'-ending strands.   RuvC then resolves the Holliday junction yielding a reconstituted repli¬  cation fork ready for replication restart.   Thus, the 5 r -ending strand of the nick has, in effect, become the 5' end of an  Okazaki fragment.   The final step in the recombination repair process is the restart of DNA  replication. This process is, of necessity, distinct from the replication initiation  that occurs at oriC (Section 25-2B). Origin-independent replication restart  is mediated by a specialized seven-subunit primosome, which has therefore  been named the restart primosome.   Recombination Repair Reconstitutes Double-Strand Breaks. We have seen  that double-strand breaks (DSBs) in DNA can be rejoined, often mutagenically,  by nonhomologo us end-joining (NHEJ; Section 25-5E). DSBs may also be non-  mutagenically repaired through a recombination repair process known as ho¬  mologous end-joining, which occurs via two Holliday junctions (Fig. 25-49):   1 . The DSBs double-stranded ends are resected to produce single-  stranded ends. One of the 3 r -ending strands invades the corresponding   FIG. 25-49 The repair of a double-strand break in DNA by  homologous end-joining. Thick lines indicate parental DNA, thin    lines indicate newly synthesized DNA, and the arrows point in the  5' —¥ 3' direction. [After Haber, J.E., Trends Genet. 16, 259 (2000).]    sequence of a homologous chromosome to form a Holliday junction,  a process mediated by Rad51 (the eukaryotic RecA homolog). The  other 3'-ending strand pairs with the displaced strand segment on the  homologous chromosome to form a second Holliday junction.   2 . DNA polymerase fills in the gaps, and ligase seals the joints.   3. Both Holliday junctions are resolved to yield two intact double-  stranded DNAs.   Thus, the sequences that may have been lost in the formation of the DSB are  copied from the homologous chromosome. Of course, a limitation of homol¬  ogous end-joining, particularly in haploid cells, is that a homologous chromo¬  somal segment may not be available.   r The importance of recombination repair in humans is demonstrated by the  observation that defects in the proteins BRCA1 (1863 residues) and  BRCA2 (3418 residues), both of which interact with Rad51, are associated  with a greatly increased incidence of breast, ovarian, prostate, and pancreatic  cancers. Indeed, individuals with mutant BRCA1 or BRCA2 genes have up to  an 80% lifetime risk of developing cancer.   In the early 1950s, Barbara McClintock reported that the variegated pigmenta¬  tion pattern of maize (Indian corn) kernels results from the action of genetic el¬  ements that can move within the maize genome. This proposal was resoundingly  ignored because it was contrary to the then-held orthodoxy that chromosomes  consist of genes linked in fixed order. Another 20 years were to pass before ev¬  idence of mobile genetic elements was found in another organism, E. coli.   Transposons Move Genes between Unrelated Sites. It is now known that  transposable elements, or transposons, are common in both prokaryotes and  eukaryotes, where they influence the variation of phenotypic expression over  the short term and evolutionary development over the long term. Each trans-  poson codes for the enzymes that insert it into the recipient DNA. This  process differs from homologous recombination in that it requires no homol¬  ogy between donor and recipient DNA and occurs at a rate of only one event  in every 10 4 to 10 7 cell divisions.   1 . The simplest transposons are named insertion sequences or IS ele¬  ments. They are normal constituents of bacterial chromosomes and  plasmids (autonomously replicating circular DNA molecules that usu¬  ally consist of several thousand base pairs; Section 3-5A). For example,  a common E. coli strain has eight copies of IS1 and five copies of IS2.  IS elements generally consist of <2000 bp, comprising a so-called  transposase gene and, in some cases, a regulatory gene, flanked by short  inverted (having opposite orientation) terminal repeats. On each side of  an inserted IS element is a directly (having the same orientation)  repeated segment of host DNA (Fig. 25-50). This suggests that an IS    ■ ■ ■ ABCD 12345 . 5'4'3'2'r ABCD - - -   ■ ■ ■ A'B'C'D' 1'2'3'4'5'. 5 4 3 2 1 A'B'C'D' - - -    FIG. 25-50 Structure of IS elements. These and other transposons have inverted  terminal repeats ( numerals ) and are flanked by direct repeats of host DNA sequences  (letters).    FIG. 25-51 A model for transposon insertion. A staggered cut that is later filled in generates direct repeats of the target sequence.    element is inserted in the host DNA at a staggered cut that is later filled  in (Fig. 25-51). The length of the target sequence (most commonly  5-9 bp), but not its sequence, is characteristic of the IS element.   2. More complex transposons carry genes not involved in the transposition  process , such as antibiotic-resistance genes. For example, Tn3 consists  of 4957 bp and has inverted terminal repeats of 38 bp each.    The central region of Tn3 codes for three proteins: (1) a 1015-residue  transposase named TnpA; (2) a 185-residue protein known as TnpR,  which represses the expression of both tnpA and tnpR and mediates  the site-specific recombination reaction necessary for transposition (see  below); and (3) a (5-lactamase (encoded by amp) that inactivates the  antibiotic ampicillin (Box 8-3). The site-specific recombination occurs  in an AT-rich region, the internal resolution site, between tnpA and  tnpR.   3. The so-called composite transposons (Fig. 25-52) consist of a gene-  containing central region flanked by two identical or nearly identical    FIG. 25-52 A composite transposon. This element consists of two identical or  nearly identical IS-like modules (green) flanking a central region carrying various genes.  The IS-like modules may have either (a) direct or (b) inverted relative orientations.    IS-like modules that have either the same or an inverted relative  orientation. Composite transposons apparently arose by the association  of two originally independent IS elements. Experiments demonstrate  that composite transposons can transpose any sequence of DNA in their  central region.   The Mechanisms of Certain Transposons Require Replication. The trans¬  posons known as replicative transposons do not simply jump from point to  point within a genome. Instead, their mechanism of transposition involves the  replication of the transposon. A model for the movement of such a transposon  between two plasmids consists of the following steps (Fig. 25-53):   1. A pair of staggered single-strand cuts (such as in Fig. 25-51) is made  at the target sequence of the recipient plasmid. Similarly, single-strand  cuts are made on opposite strands on either side of the transposon.   2 . Each of the transposons free ends is ligated to a protruding single  strand at the insertion site. This forms a replication fork at each end  of the transposon.   3. The transposon is replicated, thereby yielding a cointegrate (the fusion  of the two plasmids). Such cointegrates have been isolated.   4. Through a site-specific crossover between the internal resolution sites  of the two transposons, the cointegrate is resolved into two separate  plasmids, each of which contains the transposon. This recombination  process is catalyzed by a transposon-coded resolvase (TnpR in Tn3)  rather than by RecA.   Transposition Is Responsible for Much Genetic Rearrangement. In addi¬  tion to mediating their own insertion into DNA, transposons promote inver¬  sions y deletionSy and rearrangements of the host DNA. Inversions can occur when  the host DNA contains two copies of a transposon in inverted orientation.  The recombination of the transposons inverts the region between them  (Fig. 25-54^). If, instead, the two transposons have the same orientation,  recombination deletes the segment between them (Fig. 25-54 £). The deletion  of a chromosomal segment in this manner, followed by its integration into the  chromosome at a different site by a separate recombination event, results in  chromosomal rearrangement.   Transposons can be considered natures genetic engineering “tools.” For  example, the rapid evolution, since antibiotics came into common use, of plas¬  mids that confer resistance to several antibiotics (Section 3-5A) has resulted  from the accumulation of the corresponding antibiotic-resistance transposons  in these plasmids. Transposon-mediated rearrangements may also have been  responsible for forming new proteins by linking two formerly independent    newly synthesized DNA. [After Shapiro, J.A., Proc. Natl. Acad. Sci. 76,  1934 (1979).]    FIG. 25-54 Chromosomal rearrangement via recombination. transposons with inverted orientations, (b) The deletion of a DNA  (a) The inversion of a DNA segment between two identical segment between two identical transposons with the same orientation.    gene segments. Moreover, transposons can apparently mediate the transfer of  genetic information between unrelated species.   Most Eukaryotic Transposons Resemble Retroviruses. Transposons occur  in such distantly related eukaryotes as yeast, maize, fruit flies, and humans. In  fact, ~3% of the human genome consists of DNA-based transposons  although, in most cases, their sequences have mutated so as to render them  inactive; that is, these transposons are evolutionary fossils.   Most eukaryotic transposons exhibit little similarity to those of prokary¬  otes. Often, their base sequences resemble those of retroviruses (Box 25-2),  which suggests that these transposons are degenerate retroviruses. The trans¬  position of these so-called retrotransposons occurs via a pathway that resem¬  bles the replication of retroviral DNA: (1) their transcription to RNA, (2) the  reverse transcriptase-mediated copying of the RNA to cDNA, and (3) the  largely random insertion of the DNA into the host organisms genome as  mediated by enzymes known as integrases (which resemble DNA transposases  in structure and mechanism). Other retrotransposons replicate via a different  mechanism. Collectively, retrotransposons account for ~42% of the human  genome.   • Describe the processes of strand invasion,  branch migration, and resolution during  homologous recombination.   • What is the role of recombination in re¬  pairing damaged DNA?   • Can recombination occur in haploid as  well as diploid cells?   • What percentage of the human genome  consists of transposons and  retrotransposons?   • DNA is replicated semiconservatively through the action of DNA  polymerases that use the separated parental strands as templates  for the synthesis of complementary daughter strands.   • Replication is semidiscontinuous: The leading strand is synthe¬  sized continuously while the lagging strand is synthesized as RNA-  primed Okazaki fragments that are later joined.   • E. coli DNA polymerase I (Pol I), which has 3' —> 5' and 5' —> 3'  exonuclease activities in addition to its 5' —> 3' polymerase  activity, excises RNA primers and replaces them with DNA.  Pol III is the primary polymerase in E. coli .   • To initiate replication, parental strands are first melted apart at a  specific site named oriC and further unwound by a helicase.   Single-strand binding protein (SSB) prevents the resulting single  strands from reannealing. A primase-containing primosome syn¬  thesizes an RNA primer.   • Because DNA polymerases operate only in the 5' —> 3' direction,  the lagging strand template must loop back to the replisome,  which contains two polymerases. A sliding clamp increases Pol Ill’s  processivity. DNA ligase seals the nicks between Okazaki frag¬  ments. Replication proceeds until the two replication forks meet  between oppositely facing Ter sequences.   • The high fidelity of DNA replication is achieved by the regula¬  tion of dNTP levels, by the low error rate of the polymerase  reaction, by the requirement for RNA primers, by 3' —> 3' proof¬  reading, and by DNA repair mechanisms.   • Eukaryotes contain a number of DNA polymerases. Pol a extends  primers, the highly processive pol 8 is the lagging strand polymerase,  and pol £ appears to be the leading strand polymerase. Eukaryotic  replication has multiple origins and proceeds through nucleosomes.   • In order to replicate the 5' end of the lagging strand, eukaryotic  chromosomes end with repeated telomeric sequences appended by  the ribonucleoprotein telomerase. The 3' extension at the end of  each chromosome serves as a template for primer synthesis.  Somatic cells lack telomerase activity, which may prevent them  from transforming to cancer cells.   • Mutations in nucleotide sequences arise spontaneously from repli¬  cation errors and from alterations triggered by agents such as UV  light, radiation, and chemical mutagens. Many compounds that  are mutagenic in the Ames test are also carcinogenic.   • Some forms of DNA damage (e.g., alkylated bases and pyrimidine  dimers) may be reversed in a single step. In base excision repair,  a glycosylase removes damaged bases. In nucleotide excision repair,  an oligonucleotide containing the lesion is removed and replaced.  Repair pathways that use error-prone polymerases, nonhomolo-  gous end-joining, and the E. coli SOS response are mutagenic.   • Homologous recombination, in which strands of homologous  DNA segments are exchanged, involves a crossover structure  (Holliday junction). Recombination requires proteins to unwind  DNA, mediate strand exchange, drive branch migration, and re¬  solve the Holliday junction. Damaged DNA can be repaired  through recombination.   • Transposons are genetic elements that move within a genome,  often by mechanisms involving their replication. Transposons  mediate rearrangement of the host DNA.    1. Approximately how many Okazaki fragments are synthesized in  the replication of the E. coli chromosome?   2. Approximately how many Okazaki fragments are synthesized in  the replication of the human genome?   3. Explain why a DNA polymerase that could synthesize DNA in  the y — » 5' direction would have a selective disadvantage even  if it had 5' —> 3' proofreading activity.   4. You have discovered a drug that inhibits the activity of inorganic  pyrophosphatase. What effect would this drug have on DNA  synthesis?    3. Why are there no Pol I mutants that completely lack 5' —> 3' exo¬  nuclease activity?   6. Why is it necessary to use only the Klenow fragment, rather  than intact E. coli Pol I, in DNA sequencing reactions (Section   7. The three DNA base pairs that fit within the active site of DNA  polymerase have a conformation similar to that of A-DNA rather  than the usual B-DNA. Why is it important for DNA polymerase  to accommodate an A-type helix?    8. A reaction mixture contains DNA polymerase, the four dNTPs,  and one of the DNA molecules whose structure is represented  below. Which reaction mixtures generate PP Z ?   9. E. coli oriC is rich in A • T base pairs. Why is this advantageous?   10. Explain why DNA gyrase is required for efficient unwinding of  DNA by helicase at the replication fork.   11. In some experimental systems, a newly synthesized RNA primer  is transferred from primase to the clamp or clamp loader rather  than directly to DNA polymerase. Explain why this would max¬  imize the efficiency of DNA replication.   12. Why is the observed mutation rate of E. coli 10 8 to 10 10 per  base pair replicated, even though the error rates of Pol I and  Pol III are 10 6 to 10 7 per base pair replicated?   13. Why can’t a linear duplex DNA, such as that of bacteriophage  T7, be fully replicated by just E. coli-z ncoded proteins?   14. How does a eukaryotic cell distinguish the end of a broken  chromosome from the end of an intact linear chromosome?   13. Telomerase adds six deoxynucleotides to the end of a DNA strand  using a built-in RNA template. Explain why the templating  region of the RNA includes one and a half repeats of the telo-  meric sequence (~9 nucleotides).   16. Give the name of the enzyme that catalyzes each of the follow¬  ing reactions:   tautomerization from its keto form to its enol form than does  thymine. 5BU can be incorporated into newly synthesized DNA  when it pairs with adenine on the template strand. However, the  enol form of 5BU can pair with guanine rather than adenine, (a)  Draw the 5BU • G base pair, (b) What type of mutation results?   Hydroxylamine (NH 2 OH) converts cytosine to the compound  shown below.   With which base does this modified cytosine pair? Does this gen¬  erate a transition or a transversion mutation?   19. Oxidative deamination of adenine produces hypoxanthine (the  base of inosine), which can base pair with cytosine, (a) If no  repair takes place, describe the makeup of the DNA in the two  daughter cells following cell division, (b) Describe the makeup of  the DNA in the four daughter cells following a second round of  cell division.   20. A certain mutant DNA polymerase is error-prone, tending to  incorporate C opposite a template A. When such a DNA poly¬  merase replicates a segment of DNA containing an A • T base  pair, what will be the DNA composition in the daughter cells af¬  ter (a) one and (b) two rounds of cell division? Assume that DNA  repair does not occur.   21. Use the data in Table 11-1 to calculate the half-life in years of a  phosphodiester bond in the absence of a nuclease. ( Hint : See  Section 12-1 A.)   22. According to the text, approximately 20,000 of 6.0 billion gly-  cosidic bonds in the DNA of a diploid human cell spontaneously  hydrolyze each day. What is the half-life in years of such a bond?  {Hint See Section 12-1 A.)   23. In eukaryotic cells, a specific triphosphatase cleaves deoxy-8-  oxoguanosine triphosphate (oxo-dGTP) to oxo-dGMP + PP Z .  What is the advantage of this reaction?   24. Yeast and some other organisms produce proteins that closely  resemble 0 6 -alkylguanine-DNA alkyltransferase but lack a Cys  at the active site, (a) Explain why these proteins cannot remove  an alkyl group attached to guanine, (b) These proteins do protect  cells against DNA alkylation damage. In which type of repair  process are these proteins most likely to participate?   25. Certain sites in the E. coli chromosome are known as hot spots  because they have unusually high rates of point mutations. Many  of these sites contain a 5-methylcytosine residue. Explain the  existence of such hot spots.   26. Since mammalian DNA contains roughly 25% thymine residues,  why do mammalian cells need a thymine-DNA glycosylase?   27. What type of repair system would cells most likely use to repair  DNA with a deaminated adenine?   28. Explain why base excision repair, nucleotide excision repair, and  mismatch repair—which all require nucleases to excise damaged  DNA—require DNA ligase.   29. In E. coli, all newly synthesized DNA appears to be fragmented  (an observation that could be interpreted to mean that the lead¬  ing strand as well as the lagging strand is synthesized discontin-  uously). However, in E. coli mutants that are defective in    uracil-DNA glycosylase, only about half the newly synthesized  DNA is fragmented. Explain.   30. Broken DNA can be repaired by DNA ligase (single-strand  breaks) or nonhomologo us end-joining (double-strand breaks).  Explain why the cell’s set of repair enzymes also includes  tyrosyl-DNA phosphodiesterases.   3 E A DNA template strand containing a thymine dimer can be repli¬  cated by DNA polymerase r\. Does pol r\ fix the damaged DNA?   32. E. coli DNA polymerase V has the ability to bypass thymine  dimers. However, Pol V tends to incorporate G rather than A op¬  posite the damaged T bases. Would you expect Pol V to be more  or less processive than Pol III? Explain.   33. Predict whether loss of the following E. coli genes would be lethal  or not: (a) dnaB (which encodes DnaB), (b) polA (which encodes  Pol I), (c) ssb, (d) recA.   34. During bacterial conjugation, two cells of the same or similar  species make close contact and transfer genetic information. In  the donor cell, an endonuclease nicks the DNA so that a single  strand can enter the recipient cell. Explain why the stable incor¬  poration of the transferred DNA requires RecBCD activity in the  recipient cell.    Following DNA replication in  eukaryotes, the two DNA molecules (sister chromatids) remain  associated through the binding of a protein complex called cohesin.  Describe cohesin’s structure and explain how it binds to DNA and  how it eventually releases the DNA. What role does cohesin play  in recombination?   Focus concept: The activity of the pentose phosphate pathway  enzyme glucose-6-phosphate dehydrogenase has been found to be  important in the regulation of cell growth.   Autexier, C. and Lue, N.E, The structure and function of telomerase  reverse transcriptase, Annu. Rev. Biochem. 75, 493-517 (2006).   Blackburn, E.H., Telomere states and cell fates, Nature 408, 53-56 (2000).   Bowman, G.D., Goedken, E.R., Kazmirski, S.L., O’Donnell, M., and  Kuriyan, J., DNA polymerase clamp loaders and DNA replication,  FEBS Lett. 579, 863-867 (2005).   Enemark, E.J. and Joshua-Tor, L., Mechanism of DNA translocation in  a replicative hexameric helicase, Nature 442, 270-275 (2006).   Georgescu, R.E., Kim, S.-S., Yurieva, O., Kuriyan, J., Kong, X.-P., and  O’Donnell, M., Structure of a sliding clamp on DNA, Cell 132,  43-54 (2008).   Gillis, A.J., Schuller, A.P., and Skordalakes, E., Structure of the Tribolium cas-  taneum telomerase catalytic subunit TERT, Nature 455, 633-637 (2008).   Groth, A., Rocha, W., Verreault, A., and Almouzni, G., Chromatin challenges  during DNA replication and repair, Cell 128, 721-733 (2007).   Hamdan, S.M. and Richardson, C.C., Motors, switches, and contacts in  the replisome, Annu. Rev. Biochem. 78, 205-243 (2009).   Johansson, E. and MacNeill, S.A., The eukaryotic replicative DNA poly¬  merase takes shape, Trends Biochem. Sci. 35, 339-347 (2010).   McHenry, C.S., DNA replicases from a bacterial perspective, Annu. Rev.  Biochem. 80, 403-436 (2011).   O’Donnell, M., Replisome architecture and dynamics in E. coli, J. Biol.  Chem. 281, 10653-10656 (2006); and Johnson, A. and O’Donnell,  M., Cellular DNA replicases: Components and dynamics at the  replication fork, Annu. Rev. Biochem. 74, 283-315 (2005).   O’Sullivan, R.J. and Karlseder, J., Telomeres: protecting chromosomes against  genome instability, Nature Rev. Mol. Cell Biol. 11, 171-181 (2010).   Rothwell, P.J. and Waksman, G., Structure and mechanism of DNA poly¬  merases, Adv. Trot. Chem. 71, 401-440 (2005).   Singleton, M.R., Dillingham, M.S., and Wigley, D.B., Structure and  mechanism of helicases and nucleic acid translocases, Annu. Rev.  Biochem. 76, 23-50 (2007).   Friedberg, E.C., Walker, G.C., Siede, W., Wood, R.D., Schultz, R.A., and  Ellenberger, T., DNA Repair and Mutagenesis (2nd ed.), ASM Press  (2006).    Jackson, S.P. and Bartek, J., The DNA-damage response in human  biology and disease, Nature 461, 1071-1078 (2009). [Summarizes  current understanding of how signaling pathways triggered by DNA  damage lead to DNA repair.]   Lieber, M.R., The mechanism of double-strand DNA break repair by the  nonhomologo us DNA end-joining pathway, Annu. Rev. Biochem.  79, 181-211 (2010).   Sancar, A., Lindsey-Bolz, L.A., Unsal-Ka^maz, K., and Linn, S., Molec¬  ular mechanisms of mammalian DNA repair and the DNA damage  checkpoints, Annu. Rev. Biochem. 73, 39-85 (2004).   Valle, D. (Ed.), The Online Metabolic & Molecular Bases of Inherited  Disease, http://www.ommbid.com/. [Chapters 28 and 32 contain  discussions of xeroderma pigmentosum, Cockayne syndrome, and  hereditary nonpolyposis colorectal cancer.]   Chen, A., Yang, H., and Pavletich, N.P., Mechanism of homologous  recombination from the RecA-ssDNA/dsDNA structures, Nature  453, 489-494 (2008).   Cox, M.M., Motoring along with the bacterial RecA protein, Nature  Rev. Mol. Cell Biol. 8, 127-138 (2007); and Regulation of bacte¬  rial RecA protein function, Crit. Rev. Biochem. Mol. Biol. 42,  41-63 (2007).   Craig, N.L., Craigie, R., Gellert, M., and Lambowitz, A.M. (Eds.),  Mobile DNA II, ASM Press (2002). [A compendium of authoritative  articles.]   Kowalczykowski, S.C., Initiation of genetic recombination and  recombination-dependent replication, Trends Biochem. Sci. 25,  156-165 (2000). [Presents models for and describes the proteins  involved in recombination/replication events in bacteria.]   West, S.C., Molecular views of recombination proteins and their control,  Nature Rev. Mol. Cell. Biol. 4, 435-445 (2003).   Yamada, K., Ariyoshi, M., and Morikawa, K., Three-dimensional struc¬  tural views of branch migration and resolution in DNA homologous  recombination, Curr. Opin. Struct. Biol. 14, 130-137 (2004).    Vaults, large ribonucleoprotein particles in animal cells, are constructed mainly from  78 protein subunits. The hollow structure includes about a dozen small RNA molecules  that are likely regulatory rather than structural. Various studies suggest that vaults may   C The RNA Chain Grows from the 5' to 3' End   A Messenger RNAs Undergo 5' Capping and  Addition of a 3' Tail   D Transfer RNAs Are Processed by Nucleotide  Removal, Addition, and Modification    play a role in cell signaling, drug metabolism, or pathogen clearance. [Courtesy of  Tomitake Tsukihara, University of Osaka, Japan.]    DNA is confined almost exclusively to the nucleus of eukaryotic cells, as  shown by microscopists in the 1930s. By the 1950s, the site of protein syn¬  thesis was identified by showing that radioactively labeled amino acids that  had been incorporated into proteins were associated with cytosolic RNA—  protein complexes called ribosomes. Thus, protein synthesis is not immediately  directed by DNA because, at least in eukaryotes, DNA and ribosomes are never  in contact. The intermediary between DNA and the protein-biosynthesis  machinery, as outlined in Francis Cricks central dogma of molecular biology  (Section 3-3B), is RNA.   Cells contain three major types of RNA: ribosomal RNA (rRNA), which  constitutes two-thirds of the ribosomal mass; transfer RNA (tRNA), a set of  small, compact molecules that deliver amino acids to the ribosomes for assem¬  bly into proteins; and messenger RNA (mRNA), whose nucleotide sequences  direct protein synthesis. In addition, a host of other noncoding RNA species  play various roles in the regulation of gene expression and the processing of  newly transcribed RNA molecules (Table 26-1). All types of RNA can be  shown to hybridize with complementary sequences on DNA from the same  organism. Thus, all cellular RNAs are transcribed from DNA templates.    The transcription of DNA to RNA is carried out by RNA polymerases  (RNAPs) that operate as multisubunit complexes, as do the DNA polymerases  that catalyze DNA replication. In this chapter, we examine the catalytic prop¬  erties of RNAPs and discuss how these proteins—unlike DNA polymerases—  are targeted to specific genes. We will also see how newly synthesized RNA is  processed to become fully functional.   FIG. 26-1 An electron micrograph of  E. coli RNA polymerase holoenzyme. This  soluble enzyme, one of the largest known, is  attached to various promoter sites on  bacteriophage T7 DNA. [From Williams, R.C.,  Proc. Natl. Acad. Sci. 74, 2313 (1977).]   • Bacterial transcription begins with the RNAP holoenzyme binding to a promoter  to melt apart the DNA.   • As the polymerase processively extends an RNA chain, another polymerase can  initiate transcription.   • Transcription termination in prokaryotes depends on intrinsic terminator sequences  or Rho factor.   RNAP, the enzyme responsible for the DNA-directed synthesis of RNA, was  discovered independently in I960 by Samuel Weiss and Jerard Hurwitz. The  enzyme couples together the ribonucleoside triphosphates (NTPs) ATP, CTP,  GTP, and UTP on DNA templates in a reaction that is driven by the release  and subsequent hydrolysis of PP Z :   All cells contain RNAP. In bacteria, one enzyme synthesizes all of the  cells RNA except the short RNA primers employed in DNA replication  (Section 25-2B). Eukaryotic cells contain four or five RNAPs that each syn¬  thesize a different class of RNA. We will first consider the bacterial enzyme  because it is smaller and simpler than the eukaryotic enzymes.   The E. coli RNAP holoenzyme is an ~449-kD protein with subunit compo¬  sition a 2 (3P'u)a. Once RNA synthesis has been initiated, however, the cr sub¬  unit (also called the cr factor) dissociates from the core enzyme a 2 |3(Ta),  which carries out the actual polymerization process. RNAP is large enough to  be clearly visible in electron micrographs (Fig. 26-1).   FIG. 26-2 X-Ray structure of the Tth core RNAP in complex with a 23-nt template  DNA, a 14-nt nontemplate DNA, and a 16-nt RNA. The protein is drawn in ribbon form with its  two a subunits yellow and green, its |3 subunit cyan, its |3' subunit pink, and its w subunit gray.  Its bound Mg 2+ and Zn 2+ ions are represented by red and orange spheres, respectively.   The DNA and RNA are shown in ladder form with template DNA green, nontemplate  DNA blue, and RNA red. [Based on an X-ray structure by Dmitry Vassylyev,    The X-ray structure of the related RNAP core enzymes from  Thermus aquaticus (Taq) and Thermus thermophilus (Tth) were in¬  dependently determined by Seth Darst and Dmitry Vassylyev.   The structure of the Tth core enzyme in complex with DNA and  RNA has the overall shape of a crab claw whose two pincers are  formed by the p and (T subunits (Fig. 26-2). In this so-called open  complex, the ~27-A-high space between the pincers, the main chan¬  nel, is occupied by the dsDNA. The template DNA continues to the ac¬  tive site at the end of the main channel where it base-pairs with the  incoming NTP (not present in the structure) at the so-called i + \ site near  a bound Mg 2+ ion. The 3' end of the RNA forms a 9-bp hybrid helix with  the 5' end of the template DNA strand and then exits the protein through a  channel between the p and p' subunits, the RNA exit channel, in which it  adopts a conformation similar to that of a single strand within an RNA dou¬  ble helix. However, the paths taken by the template and nontemplate DNA  strands to rejoin at the end of the so-called transcription bubble are unclear.   In the X-ray structure of the Taq holoenzyme in complex with dsDNA  containing a promoter, the cr subunit extends across the top of the core  enzyme, which causes the pincers to come together so as to narrow the channel  between them by ~10 A (Fig. 26-3). In this structure, the DNA binds to the  cr subunit via its promoter elements (see Section 26-IB) to form the so-called  closed complex in which the DNA has not yet entered the main channel to  form the transcription bubble (via an unknown mechanism). The outer sur¬  face of the holoenzyme is almost uniformly negatively charged, whereas those  surfaces that interact with nucleic acids, particularly the inner walls of the  main channel, are positively charged.    Only One Strand of DNA Is Copied. RNA synthesis is normally initiated only  at specific sites on the DNA template. In contrast to replication, which requires   FIG. 26-3 Low (6.5 A) resolution X-ray structure of Taq RNAP  holoenzyme in complex with a promoter-containing dsDNA. The   subunits of the core enzyme, which is viewed and colored as in Fig. 26-2,  are represented by their partially transparent molecular surface (which  appears striated due to the structure’s low resolution, which permits only  the polypeptide backbones to be traced). The cr subunit is drawn in worm  form colored in rainbow order from its N-terminus (blue) to its C-terminus  (red). The DNA’s sugar-phosphate backbone is drawn in cartoon form with  the template DNA strand green, the nontemplate DNA’s -10 and -35  elements (Section 26-1 B) red and magenta, and its remaining portions  blue. [Based on an X-ray structure by Seth Darst, The Rockefeller  University. PDBid 1 L9Z.]    FIG. 26-4 Sense and antisense DNA strands. The template strand of duplex DNA  is known as its antisense or noncoding strand. Its complementary sense or coding strand  has the same nucleotide sequence and orientation as the transcribed RNA.    that both strands of the chromosome be entirely copied, the regulated expres¬  sion of genetic information involves much smaller, single-strand portions of  the genome. The DNA strand that serves as a template during transcription  is known as the antisense or noncoding strand since its sequence is comple¬  mentary to that of the RNA. The other DNA strand, which has the same  sequence as the transcribed RNA (except for the replacement of U with T),  is known as the sense or coding strand (Fig. 26-4). The two strands of DNA  in an organisms chromosome can therefore contain different sets of genes.   Keep in mind that “gene” is a relatively loose term that refers to sequences  that encode polypeptides, as well as those that correspond to the sequences of  rRNA, tRNA, and other RNA species. Furthermore, a gene typically includes  sequences that participate in initiating and terminating transcription (and  translation) that are not actually transcribed (or translated). The expression of  many genes also depends on regulatory sequences that do not directly flank  the coding regions but may be located a considerable distance away.   Most protein-coding genes (called structural genes) in eukaryotes are  transcribed individually. In prokaryotic genomes, however, genes are frequently  arranged in tandem along a single DNA strand so that they can be transcribed  together. These genetic units, called operons, typically contain genes with re¬  lated functions. For example, the three different rRNA genes of E. coli occur  in single operons (Section 26-3C). The E. coli lac operon, whose expression  is described in detail in Section 28-2A, contains three genes encoding pro¬  teins involved in lactose metabolism as well as sequences that control their  transcription (Fig. 26-5). Other operons contain genes encoding proteins re¬  quired for biosynthetic pathways, for example, the trp operon, whose six gene  products (proteins are often referred to as gene products) catalyze tryptophan  synthesis. An operon is transcribed as a single unit, giving rise to a polycistronic  mRNA that directs the more-or-less simultaneous synthesis of each of the en¬  coded polypeptides (the term cistron is a somewhat archaic synonym for gene).  In contrast, eukaryotic structural genes, which are not part of operons, give  rise to monocistronic mRNAs.    FIG. 26-5 The E. coli lac operon. This DNA includes genes encoding the proteins  mediating lactose metabolism and the genetic sites that control their expression. The Z, Y,  and A genes, respectively, specify the proteins p-galactosidase (Box 8-1), galactoside  permease (Section 10-3D), and thiogalactoside transacetylase. The closely linked  regulatory gene, /, which is not part of the lac operon, encodes a repressor that inhibits  transcription of the lac operon.   How does RNAP recognize the correct DNA strand and initiate RNA synthe¬  sis at the beginning of a gene (or operon)? RNAP binds to its initiation sites  through base sequences known as promoters that are recognized by the correspon¬  ding a factor. The existence of promoters was first revealed through mutations  that enhanced or diminished the transcription rates of certain genes. Promoters  consist of ~ 40-bp sequences that are located on the 5' side of the transcrip¬  tion start site. By convention, the sequence of this DNA is represented by its  sense (nontemplate) strand so that it will have the same sequence and direc¬  tionality as the transcribed RNA. A base pair in a promoter region is assigned  a negative or positive number that indicates its position, upstream or down¬  stream in the direction of RNAP travel, from the first nucleotide that is tran¬  scribed to RNA; this start site is +1 and there is no 0. Because RNA is  synthesized in the 5' —> 3' direction (see below), the promoter is said to lie  upstream of the RNAs starting nucleotide.   The holoenzyme forms tight complexes with promoters (dissociation  constant A"— 10 14 M). This tight binding can be demonstrated by showing  that the holoenzyme protects the bound DNA segments from digestion  in vitro by the endonuclease DNase I. Sequence determinations of the protected  regions from numerous E. coli genes have identified the “consensus” sequence  of E. coli promoters (Fig. 26-6). Their most conserved sequence is a hexamer  centered at about the — 10 position (sometimes called the Pribnow box, after  David Pribnow, who described it in 1975). It has a consensus sequence of  TATAAT in which the leading TA and the final T are highly conserved.   Upstream sequences around position —35 also have a region of sequence sim¬  ilarity, TTGACA. The initiating (+1) nucleotide, which is nearly always A or  G, is centered in a poorly conserved CAT or CGT sequence. Most promoter  sequences vary considerably from the consensus sequence (Fig. 26-6).   Nevertheless, a mutation in one of the partially conserved regions can greatly   rrnDl CAAAAAAATAC T TGTGCAAAAAAT T GGGAT C C C TATAATGC GC C T C CGT T GAGAC GA   FIG. 26-6 The sense (coding) strand sequences of selected  E. coli promoters. A 6-bp region centered around the -10 position  (red shading) and a 6-bp sequence around the -35 region (blue  shading) are both conserved. The transcription initiation sites (+1),  which in most promoters occur at a single purine nucleotide, are  shaded in green. The bottom row shows the consensus sequence of  298 E. coli promoters with the number below each base indicating its    percentage occurrence. [After Rosenberg, M. and Court, D., Annu. Rev.  Genet. 13, 321-323 (1979). Consensus sequence from Lisser, D. and  Margalit, H., Nucleic Acids Res. 21, 1512 (1993).]    Approximately how many helical turns separate the two  blocks of conserved sequence?   Chapter 26 Transcription and RNA Processing increase or decrease a promoters initiation efficiency. This is because the rates   at which E. coli genes are transcribed vary directly with the rates at which their  promoters form stable initiation complexes with the RNAP holoenzyme.   Initiation Requires the Formation of a Transcription Bubble. The promoter  regions in contact with the RNAP holoenzyme have been identified by a pro¬  cedure named footprinting. In this procedure, DNA is incubated with a pro¬  tein to which it binds and is then treated with an alkylation agent such as  dimethyl sulfate (DMS). This results in alkylation of the DNAs bases followed  by backbone cleavage at the alkylated positions. However, those portions of  the DNA that bind proteins are protected from alkylation and hence cleav¬  age. The resulting pattern of protection is called the proteins footprint. The  footprint of RNAP holoenzyme indicates that it contacts the promoter prima¬  rily at its —10 and — 35 regions. In some genes, additional upstream sequences  may also influence RNAP binding to DNA.   DMS methylates G residues at N7 and A residues at N3 in both double-  and single-stranded DNA. DMS also methylates N1 of A and N3 of C, but  only if the latter positions are not involved in base-pairing interactions. The  pattern of DMS methylation therefore reveals whether the DNA is single or  double stranded. Footprinting studies indicate that holoenzyme binding  “melts” (separates) ~11 bp of DNA (from —9 to +2). The resulting tran¬  scription bubble (e.g., Fig. 26-4) resembles the region of unwound DNA at  the replication origin (Section 25-2B).   The core enzyme , which does not specifically bind promoters , tightly binds du¬  plex DNA (the complex’s dissociation constant is if — 5 X 10 12 M, and its  half-life is ~60 min). The holoenzyme , in contrast , binds to nonpromoter DNA  comparatively loosely (AT ~ 10 7 M and a half-life of >1 s). Evidently, the cr  subunit allows the holoenzyme to move rapidly along a DNA strand in search  of the cr subunit’s corresponding promoter. Once transcription has been ini¬  tiated and the <r subunit jettisoned, the tight binding of the core enzyme to  DNA apparently stabilizes the ternary enzyme—DNA—RNA complex.   Gene Expression Is Controlled by Different tr Factors. Because different cr  factors recognize different promoters, a cell's complement of a factors determines  which genes are transcribed. Development and differentiation, which involve the  temporally ordered expression of sets of genes, can be orchestrated through a  “cascade” of cr factors. For example, infection of Bacillus subtilis by bac¬  teriophage SP01 requires the expression of different sets of phage genes at dif¬  ferent times. The first set, known as the early genes, are transcribed using the  bacterial cr factor. One of the early phage gene products is a a subunit known as  cr 8 ? 28 , which displaces the host cr factor and thereby permits the RNAP to rec¬  ognize only the phage middle gene promoters. The phage middle genes, in turn,  specify q-sp 33/34 5 which promotes transcription of only phage late genes.   Several bacteria, including E. coli and Bacillus subtilis , likewise have sev¬  eral different cr factors. These are not necessarily used in a sequential manner.  For example, cr factors in E. coli that differ from its primary cr factor (which  is named cr 70 because its molecular mass is 70 kD) control the transcription  of coordinately expressed groups of special-purpose genes whose promoters  are quite different from those recognized by a 70 .   C The RNA Chain Grows from the 5' to 3' End   Because RNA synthesis, like DNA synthesis, proceeds in the 5' —> 3' direc¬  tion (Fig. 26-7), the growing RNA molecule has a 5'-triphosphate group.  Mature RNA molecules, as we will see, may also be chemically modified at  one or both ends.   A portion of the double-stranded DNA template remains opened up at  the point of RNA synthesis (Fig. 26-4). This allows the antisense strand to   to the 3' end of the growing RNA chain via attack of the 3'-OH  group on the incoming nucleoside triphosphate. A radioactive label at   the 7 position of an NTP (indicated by an asterisk) is retained in the  initial nucleotide of the RNA (the 5' end) but is lost as PP/ during the  polymerization of subsequent nucleotides.    direct the synthesis of its complementary RNA strand. The RNA chain  transiently forms a short length of RNA-DNA hybrid duplex. The unpaired  “bubble” of DNA in the open initiation complex apparently travels along the  DNA with the RNAP (Fig. 26-8).   As DNAs helical turns are pushed ahead of the advancing transcription  bubble, they become more tightly wound (more positively supercoiled) while  the DNA behind the bubble becomes equivalently unwound (more negatively  supercoiled). This scenario is supported by the observation that the transcrip¬  tion of plasmids in E. coli induces their positive supercoiling in gyrase mu¬  tants (which cannot relax positive supercoils; Section 24-ID) and their  negative supercoiling in topoisomerase I mutants (which cannot relax negative  supercoils). Inappropriate superhelicity in the DNA being transcribed halts  transcription. Quite possibly, the torsional tension in the DNA generated by  negative superhelicity behind the transcription bubble is required to help drive  the transcriptional process, whereas too much such tension prevents the open¬  ing and maintenance of the transcription bubble.   Strain that builds up in the DNA template is apparently responsible for  a curious property of RNAP: It frequently releases its newly synthesized RNA  after only ~9 to 11 nt have been polymerized, a process known as abortive  initiation. When RNAP begins transcribing, it keeps its grip on the promoter  (which is on the DNAs nontemplate strand). Consequently, conformational  tension builds up as the template strand is pulled through the RNAP s active  site, a process called scrunching because the resulting increased size of the  transcription bubble in the downstream direction must somehow be accom¬  modated within the RNAP. In successful initiation, the strain eventually pro¬  vides enough energy to strip the promoter from the RNAP, which then  continues its progress along the template. In abortive initiation, the RNAP   FIG. 26-8 DNA supercoiling during transcription. In the region  being transcribed, the DNA double helix is unwound by about a turn  to permit the DNAs antisense strand to form a short segment of  DNA-RNA hybrid double helix. As the RNA polymerase advances  along the DNA template (here to the right), the DNA unwinds ahead  of the RNA’s growing 3' end and rewinds behind it, thereby stripping  the newly synthesized RNA from the template strand. Because the    ends of the DNA as well as the RNA polymerase are apparently  prevented from rotating by attachments within the cell (black bars),  the DNA becomes overwound (positively supercoiled) ahead of the  advancing transcription bubble and underwound (negatively  supercoiled) behind it (consider the consequences of placing your  finger between the twisted DNA strands in this model and pushing  toward the right). [After Futcher, B., Trends Genet 4, 272 (1988).]   In rapidly proliferating bacterial cells, DNA synthesis is likely to  occur even as genes are being transcribed. The DNA replica¬  tion machinery moves along the circular chromosome at a rate  many times faster than the movement of the transcription  machinery. Collisions between DNA polymerase and RNA  polymerase seem unavoidable. What happens when the two  enzyme complexes collide? Using in vitro model systems, Bruce  Alberts has shown that when both enzymes are moving in the  same direction, the replication fork passes the RNA polymerase  without displacing it, leaving it fully competent to resume RNA  chain elongation.   When the replication fork collides head-on with a transcription  complex, however, the replisome pauses briefly before moving past  the RNA polymerase. Surprisingly, this causes the RNA    polymerase to switch its template strand. The growing RNA chain  dissociates from the original template DNA strand and hybridizes  with the newly synthesized daughter DNA strand of the same  sequence before RNA elongation resumes.   Head-on collisions are disadvantageous because (1) replication  slows when DNA polymerase pauses, and (2) dissociation of the  RNA polymerase during the jump from one template strand to the  other could abort the transcription process. Indeed, in many  bacterial and phage genomes, the most heavily transcribed genes  are oriented so that replication and transcription complexes move  in the same direction. It remains to be seen whether a similar  arrangement holds in eukaryotic genomes, which contain multiple  replication origins and genes that are much larger than are  prokaryotic genes.   fails to escape the promoter and instead relieves the conformational tension  by releasing the newly synthesized RNA fragment, thereby letting the tran¬  scription bubble relax to its normal size. The RNAP then reinitiates transcrip¬  tion from the +1 position.   RNA Polymerase Is Processive. Once the open complex has been formed,  transcription proceeds without dissociation of the enzyme from the template.  Processivity is accomplished without an obvious clamplike structure (e.g., the  p clamp of E. coli DNA polymerase III; Fig. 25-16), although the RNAP it¬  self apparently functions as a sliding clamp by binding tightly but flexibly to  the DNA—RNA complex. In experiments in which the RNAP was immobi¬  lized and a magnetic bead was attached to the DNA, up to 180 rotations (rep¬  resenting nearly 1900 base pairs at 10.5 bp per turn) were observed before the  polymerase slipped. Such processivity is essential, since genes are often thou¬  sands (in eukaryotes, sometimes millions) of nucleotides in length.   The tight association between RNAP and DNA and their multiple attach¬  ment sites may explain why a transcription complex does not completely dissoci¬  ate from a DNA template even when transcription is interrupted by the DNA  replication machinery proceeding along the same strand of DNA (see Box 26-1).   Transcription Is Rapid. In E. coli , the in vivo rate of transcription is 20 to  50 nt/s at 37°C (but still many times slower than the DNA replication rate  of —1000 nt/s; Section 25-2C). The error frequency in RNA synthesis is one  wrong base incorporated for every —10 4 transcribed. This frequency, which is  10 4 to 10 6 times higher than that for DNA synthesis, is tolerable because most  genes are repeatedly transcribed, because the genetic code contains numerous  synonyms (Section 27-1C), and because amino acid substitutions in proteins  are often functionally innocuous.   Once an RNAP molecule has initiated transcription and moved away from  the promoter, another RNAP can follow suit. The synthesis of RNAs that are  needed in large quantities, rRNAs, for example, is initiated as often as is ster-  ically possible, about once per second. This gives rise to an arrowhead appear¬  ance of the transcribed DNA (Fig. 26-9). mRNAs encoding proteins are  generally synthesized at less frequent intervals, and there is enormous varia¬  tion in the amounts of different polypeptides produced. For example, an  E. coli cell may contain 10,000 copies of a ribosomal protein, whereas a reg¬  ulatory protein may be present in only a few copies per cell. Many enzymes,  particularly those involved in basic cellular “housekeeping” functions, are syn¬  thesized at a more or less constant rate; they are called constitutive enzymes.   FIG. 26-9 An electron micrograph of three contiguous  ribosomal genes undergoing transcription. The “arrowhead”  structures result from the increasing lengths of the nascent RNA  chains as the RNA polymerases synthesizing them move from the    initiation site on the DNA to the termination site. [Courtesy of Ulrich  Scheer, University of Wurzburg, Germany.]    Other enzymes, termed inducible enzymes, are synthesized at rates that vary  with the cells circumstances. To a large extent, the regulation of gene expres¬  sion relies on mechanisms that govern the rate of transcription, as we will see  in Section 28-2. The products of transcription also vary in their stabilities.  Ribosomal RNA turns over much more slowly than mRNA, which is rapidly  synthesized and rapidly degraded (sometimes so fast that the 5' end of an  mRNA is degraded before its 3' end has been synthesized).   Electron micrographs such as Fig. 26-9 suggest that DNA contains specific sites  at which transcription is terminated. The transcription termination sequences  of about half of E. coli genes share two common features (Fig. 26-10):   1. A series of 4 to 10 consecutive A • T base pairs, with the As on the  template strand. The transcribed RNA is terminated in or just past this  sequence.   2. A G + C—rich region with a palindromic sequence that immediately  precedes the series of A • T s.   The RNA transcript of this region, called an intrinsic terminator, can there¬  fore form a self-complementary “hairpin” structure that is terminated by sev¬  eral U residues.   The structural stability of an RNA transcript at its terminators  G + C-rich hairpin and the weak base pairing of its oligo(U) tail to template  DNA appear to be important factors in ensuring proper chain termination.  The formation of the G + C—rich hairpin causes RNAP to pause for several  seconds at the termination site. This probably induces a conformational  change in the RNAP that permits the nontemplate DNA strand to displace  the weakly bound oligo(U) tail from the template strand, thereby sponta¬  neously terminating transcription.   Despite the foregoing, experiments by Michael Chamberlin indicate that  the RNA-terminator hairpin and U-rich 3' tail do not function independently  of their upstream and downstream flanking regions. Indeed, terminators that   Its transcription yields an RNA (red) with a  self-complementary G + C-rich segment that  forms a base-paired hairpin immediately followed  by a sequence of 4 to 10 consecutive U’s that  base-pair with the template A’s (black) in the  transcription bubble. The oval symbol represents  the binding site for an incoming NTP. [After a  drawing by Joo-Soep Park and Jeffrey Roberts,  Cornell University.]   Chapter 26 Transcription and RNA Processing lack a U-rich segment can be highly efficient when joined to the appropriate   sequence immediately downstream from the termination site. Termination ef¬  ficiency also varies with the concentrations of nucleoside triphosphates, with  the level of supercoiling in the DNA template, with changes in the salt con¬  centration, and with the sequence of the terminator. These results suggest that  termination is a complex multistep process.   Termination Often Requires Rho Factor. Approximately half the termination  sites in E. coli lack any obvious similarities to the intrinsic terminator described  above and are unable to form strong hairpins. Instead, they require the action  of a protein known as Rho factor to terminate transcription. Rho factor, a hexa-  mer of identical 419-residue subunits, enhances the termination efficiency of  spontaneously terminating transcripts and induces termination of nonsponta-  neously terminating transcripts.   Several key observations have led to a model of Rho-dependent termination:   1. Rho is a helicase that unwinds RNA-DNA and RNA-RNA double  helices by translocating along a single strand of RNA in its 5' —> 3'  direction. This process is powered by the hydrolysis of NTPs to  NDPs + P z with little preference for the identity of the base.   2. Genetic manipulations indicate that Rho-dependent termination re¬  quires the presence of a specific recognition sequence on the newly  transcribed RNA upstream of the termination site. The recognition se¬  quence must be on the nascent RNA rather than the DNA as is  demonstrated by Rho s inability to terminate transcription in the pres¬  ence of pancreatic RNase A. The essential features of this termination  site have not been fully elucidated; the construction of synthetic ter¬  mination sites indicates that it consists of 80 to 100 nucleotides that  lack a stable secondary structure and contain multiple regions that are  rich in C and poor in G.   These observations suggest that Rho factor attaches to the RNA at its recogni¬  tion sequence and then migrates along the RNA in the 5' —> 3' direction until  it encounters an RNAP paused at the termination site (without the pause, Rho  might not be able to overtake the RNAP). There, as Jeffrey Roberts has shown,  Rho pushes the RNAP forward in a way that partially rewinds its dsDNA he¬  lix at the transcription bubble while unwinding the RNA-DNA hybrid helix,  thus releasing the RNA. Rho-terminated transcripts have 3' ends that typically  vary over a range of ~50 nt. This suggests that Rho gradually pries the RNA  away from its template DNA rather than liberating the RNA at a specific point.   Each Rho subunit consists of two domains: Its N-terminal domain binds  single-stranded polynucleotides and its C-terminal domain, which is homolo¬  gous to the a and (3 subunits of the F 1 -ATPase (Section 18-3B), binds an  NTP. The X-ray structure of Rho in complex with AMPPNP and an 8-nt  RNA (Fig. 26-11^), determined by James Berger, reveals that Rho forms a  hexameric lock washer-shaped helix that is 120 A in diameter with an ~30-  A-diameter central hole and whose first and sixth subunits are separated by a  12-A gap and a rise of 45 A along the helix axis. The RNA binds along the  interior of the helix to the so-called primary RNA binding sites on the N-  terminal domains and to the so-called secondary RNA binding sites on the  C-terminal domains. This structure represents an open state that is poised to  bind mRNA that has entered its central cavity through the notch.   In the X-ray structure of Rho in complex with rU 12 and the ATP mimic  ADP • BeF 3 (Fig. 26-11 h), the proteins six subunits have formed a closed ring  in which each subunit has a different conformation. Protein loops that extend  from the walls of the central channel to interact with the RNA are helically  arranged like the steps of a right-handed spiral staircase such that they track  the RNAs sugar-phosphate backbone, much like the central loops of El  protein (a hexagonal helicase) tracks its centrally bound ssDNA (Section 25-2B).   FIG. 26-1 1 X-Ray structures of Rho factor, (a) Rho in complex  with r(UC) 4 (only one UC unit of which is visible) and AMPPNR Each  of the protein’s six subunits are drawn in tube-and-arrow form in  different colors with the upper right subunit colored in rainbow order  from its N-terminus (blue) to its C-terminus (red). The UC units and  the AMPPNP are shown in space-filling form with UC C green,  AMPPNP C gray, N blue, 0 red, and P orange. The hexamer has a  lock washer-like shape with the blue subunit —45 A closer to the  viewer than the pink subunit, (b) Rho in complex with rU 12 (only 6 nt  of which are visible) and ADP • BeF 3 . The protein is drawn and   colored as in Part a. The RNA and ADP • BeF 3 are shown in stick  and space-filling form, respectively, with RNA C green, ADP C gray, N  blue, 0 red, P orange, Be light green, and F light blue. Note that each  of the Rho subunits has a different conformation. [Based on X-ray  structures by James Berger, University of California at Berkeley.    The different conformations of Rho s six subunits indicate that they sequentially  undergo a series of six NTP-driven conformational changes and that the  changes are allosterically coupled so that they progress around the hexamer in  a wavelike manner. Since each of the foregoing loops maintains its grip on the  same nucleotide during the process, the helicase translocates along its bound  RNA, in much the same way that El protein translocates along its bound  ssDNA. Why, then, do Rho and El protein move in opposite directions?  Comparison of the structures of Rho and El protein indicates that the rela¬  tive order of the conformational states around the hexamer for Rho is oppo¬  site that for El protein. Evidently, the “firing order” of Rhos NTPase sites  around the hexamer is the reverse of that of El protein, thus accounting for  their differing directions of translocation.    • Eukaryotes use three RNA polymerases to carry out DNA-directed RNA synthesis.   • The three RNA polymerases recognize different and sometimes highly variable  promoter sequences.   • A set of general transcription factors are required to initiate transcription  in eukaryotes.    Although the fundamental principles of transcription are similar in prokary¬  otes and eukaryotes, eukaryotic transcription is distinguished by having multiple   • Compare DNA and RNA polymerases with  respect to structure, substrates, products,  mechanism, error rate, and template  specificity.   • What is the significance of the different  DNA-binding properties of prokaryotic RNA  polymerase core enzyme and holoenzyme?   • Why is it important for the transcription  bubble to remain a constant size?   • Explain why a single gene can quickly give  rise to many copies of its gene products  (RNA or protein).   • Describe how transcription is terminated  with and without Rho factor.    A wide variety of compounds inhibit transcription in prokaryotes  and eukaryotes. These agents are therefore toxic to susceptible  organisms; that is, they function as antibiotics. Such compounds  are also useful research tools since they arrest the transcription  process at well-defined points.   Two related antibiotics, rifamycin B, which is produced by  Streptomyces mediterranei, and its semisynthetic derivative  rifampicin,   Rifamycin B R 1 = CH 2 COO ; R 2 = H  Rifampicin R x = H; R 2 = CH = N—N N—CH 3   specifically inhibit transcription by prokaryotic but not eukaryotic  RNA polymerases. The selectivity and high potency of rifampicin  (2 X 10 -8 M results in 50% inhibition of bacterial RNA poly¬  merase) make it a medically useful bactericidal agent. Rifamycins  inhibit neither the binding of RNA polymerase to the promoter nor  the formation of the first phosphodiester bonds, but they prevent  further chain elongation. The inactivated RNA polymerase remains  bound to the promoter, thereby blocking initiation by uninhibited  enzyme. Once RNA chain initiation has occurred, however,  rifamycins have no effect on the subsequent elongation process.  The rifamycins can therefore be used in the laboratory to dissect  transcriptional initiation and elongation.   Actinomycin D (right), a useful anti neoplastic (anticancer)  agent produced by Streptomyces antibioticus, tightly binds to duplex    DNA and, in doing so, strongly inhibits both transcription and DNA  replication, presumably by interfering with the passage of RNA  and DNA polymerases. The NMR structure of actinomycin D in  complex with an 8-bp DNA is shown opposite with the drug in  space-filling form (with C green, H white, N blue, and 0 red) and  the DNA in stick form (with C cyan, H white, N blue, 0 red, and    RNAPs and by much more complicated control sequences. Moreover, the eukary¬  otic transcription machinery, as we will see, is far more complex than that of  prokaryotes, requiring well over 100 polypeptides that form assemblies with  molecular masses of several million daltons to recognize the control sequences  and initiate transcription.   Eukaryotic nuclei contain three distinct types of RNA polymerase that differ  in the RNAs they synthesize:   1. RNA polymerase I (RNAP I), which is located in the nucleoli (dark-  staining nuclear bodies where ribosomes are assembled; Fig. 1-8), syn¬  thesizes the precursors of most rRNAs.    P orange, and with orange rods connecting successive P atoms).  The actinomycin’s phenoxazone ring system intercalates between  the DNA’s base pairs, thereby unwinding the DNA helix by 23°  and separating the neighboring base pairs by 7.0 A. Actinomycin’s  chemically identical cyclic depsipeptides (depsipeptides have both  peptide bonds and ester linkages) extend in opposite directions  from the intercalation site along the minor groove of the DNA.  Other intercalation agents, including ethidium and proflavin (Sec¬  tion 24-3B), also inhibit nucleic acid synthesis, presumably by  similar mechanisms.   The poisonous mushroom Amanita phalloides (death cap), which  is responsible for the majority of fatal mushroom poisonings in Eu¬  rope, contains several types of toxic substances, including a series of    which is representative of the amatoxins, forms a tight 1:1 complex  with RNAP II (K = 10“ 8 M) and a looser one with RNAP III  (,K = 10 -6 M) so as to specifically block their elongation steps.   The X-ray structure of RNAP II in complex with a-amanitin reveals  that a-amanitin binds in the funnel beneath the protein’s bridge  helix (Fig. 26-13b) such that it interacts almost exclusively with the  residues of the bridge helix and the adjacent part of Rpbl. The  a-amanitin binding site is too far away from the enzyme active site  to directly interfere with NTP entry or RNA synthesis, consistent  with the observation that a-amanitin does not influence the affinity  of RNAP II for NTPs. Most probably, a-amanitin binding impedes  the conformational change of the bridge helix postulated to motivate  the RNAP translocation step (Section 26-2A), which further sup¬  ports this mechanism. RNAP I as well as mitochondrial, chloroplast,  and prokaryotic RNA polymerases are insensitive to a-amanitin.   Despite the amatoxins’ high toxicity (5-6 mg, contained in  —40 g of fresh mushrooms, is sufficient to kill a human adult),  they act slowly. Death, usually from liver dysfunction, occurs no  earlier than several days after mushroom ingestion (and after  recovery from the effects of other mushroom toxins). This, in part,  reflects the slow turnover rate of eukaryotic mRNAs and proteins.    [Structure of actinomycin D-DNA complex based on an NMR struc¬  ture by Andrew Wang, University of Illinois. PDBid 1 DSC.]    2. RNA polymerase II (RNAP II), which occurs in the nucleoplasm,  synthesizes the mRNA precursors.   3. RNA polymerase III (RNAP III), which also occurs in the nucleo¬  plasm, synthesizes the precursors of 5S rRNA, the tRNAs, and a vari¬  ety of other small nuclear and cytosolic RNAs.   In addition to these nuclear enzymes, eukaryotic cells contain separate mitochon¬  drial and (in plants) chloroplast RNAPs. The essential function of RNAPs in all  cells makes them attractive targets for antibiotics and other drugs (Box 26-2).   Eukaryotic RNAPs, which have molecular masses of as much as 600 kD,  have considerably greater subunit complexity than the prokaryotic enzymes.  Each eukaryotic RNAP contains two nonidentical “large” (>120 kD) sub¬  units, which are homologs of the prokaryotic (3 and (3' subunits, and an  array of up to 12 different “small” (<50 kD) subunits, two of which are   Chapter 26 Transcription and RNA Processing homologs of the prokaryotic a subunit and one of which is a homolog of the   0) subunit. Five of the small subunits, including the homolog, are identical  in all three eukaryotic enzymes, and the a homologs are identical in RNAPs  I and III. Moreover, the sequences of these subunits are highly conserved  (~50% identical) across species from yeast to humans. In fact, in all ten cases  tested, a human RNAP II subunit could replace its counterpart in yeast with¬  out loss of cell viability.   In a crystallographic tour de force, Roger Kornberg determined the X-ray  structure of yeast RNAP II (Fig. 26-12). This enzyme, as expected, resembles  Tth and Taq RNAP (Figs. 26-2 and 26-3) in its overall crab claw-like shape  and in the positions and core folds of their homologous subunits, although  RNAP II is somewhat larger and has several subunits that have no counter¬  part in the bacterial RNAPs. RNAP II binds two Mg 2+ ions at its active site  in the vicinity of five conserved acidic residues, which suggests that RNAPs  catalyze RNA elongation via a two-metal ion mechanism similar to that em¬  ployed by DNA polymerases (Section 25-2A). As is the case with the bacter¬  ial RNAPs, the surface of RNAP II is almost entirely negatively charged except  for the DNA-binding cleft and the region about the active site, which are pos¬  itively charged.   RNAP IPs Rpbl subunit (pink in Fig. 26-12), the homolog of the (T sub¬  unit in prokaryotic RNAPs, has an extraordinary C-terminal domain (CTD). In  mammals, the CTD contains 52 highly conserved repeats with the consensus  sequence Pro-Thr-Ser-Pro-Ser-Tyr-Ser (26 repeats in yeast, with other eukary¬  otes having intermediate values). As many as 50 Ser residues in this hydroxyl-  rich protein segment are subject to reversible phosphorylation by CTD kinases  and CTD phosphatases. RNAP II initiates transcription only when the CTD  is unphosphorylated but commences elongation only after the CTD has been  phosphorylated, which suggests that this process triggers the conversion of  RNAP IPs initiation complex to its elongation complex. Charge—charge repul¬  sions between nearby phosphate groups probably cause a highly phosphorylated  CTD to project as far as 500 A from the globular portion of the polymerase.   FIG. 26-12 X-Ray structure of yeast RNA polymerase II. and the C-terminus of the |3' homolog are not visible in this structure,   (a) The enzyme is oriented similarly to the Tth RNAP in Fig. 26-2, and ( b ) View of the enzyme from the right of Part a, showing the   subunits that are homologous to those in Tth RNA polymerase are DNA-binding cleft. The circle has the approximate diameter of B-DNA.   given the same colors as in Fig. 26-2. The position of an active site [Based on an X-ray structure by Roger Kornberg, Stanford University.   Mg 2+ ion is marked by a red sphere, and eight Zn 2+ ions are PDBid 1150.]   As we will see, the phosphorylated CTD provides the binding sites for numer- Section 2 Transcription in Eukaryotes   The RNA Polymerase Structure Explains Its Function. To produce a snap¬  shot of RNAP II in action, Kornberg incubated the enzyme with a dsDNA  molecule bearing a 3' single-stranded tail (the template strand) together with  all the NTP substrates except UTP. As a consequence, the polymerase synthe¬  sized a short (14-nt) RNA strand before pausing at the first template A residue  (when the crystals of this complex were soaked in UTP, transcription resumed,  demonstrating that the complex was active). The X-ray structure and a cut¬  away diagram of this paused RNAP II are shown in Fig. 26-13.   In the RNAP-DNA-RNA complex, a massive (~50-kD) portion of the  Rpb2 subunit (the |3 homolog; cyan in Fig. 26-12), named the “clamp,” has  swung downward over the DNA to trap it in the cleft, in large part account¬  ing for the enzymes essentially infinite processivity.   The DNA unwinds by three nucleotides before entering the active site  (which is contained on Rpbl). Past this point, however, a portion of Rpb2  dubbed the “wall” directs the template strand out of the cleft in an ~90° turn.   As a consequence, the template base at the active (i + 1) site points toward  the floor of the active site where it can be read out by the polymerase. This  base is paired with the ribonucleotide at the 3' end of the RNA, which is po¬  sitioned above a 12-A-diameter pore at the end of a funnel (also called the  secondary channel) to the protein exterior through which NTPs presumably  gain access to the otherwise sealed-off active site.   The hybrid helix adopts a nonstandard conformation intermediate be¬  tween those of A- and B-DNAs, which is underwound relative to that in the   (a) X-Ray structure of the complex as viewed from the bottom of Fig.  26-12a (portions of Rpb2 that form the near side of the cleft have  been removed to expose the bound RNA-DNA complex). The protein  is represented by its backbone in which the clamp, which is closed  over the downstream DNA duplex, is yellow, the bridge helix is green,  and the remaining portions of the protein are gray. The template DNA  strand (cyan), the nontemplate DNA strand (green), and the newly  synthesized RNA (red) are drawn with their well-ordered portions in  ladder form and their less ordered portions in backbone form. An   active site Mg 2+ ion is represented by a magenta sphere, (b) Cutaway  diagram of the transcribing complex in Part a in which the cut sur¬  faces of the protein are light gray, its remaining surfaces are darker  gray, and several of its functionally important structural features are  labeled. The DNA, RNA, and active site Mg 2+ ion are colored as in  Part a with portions of the DNA and RNA that are not visible in the  X-ray structure represented by dashed lines. a-Amanitin (orange ball)  is discussed in Box 26-2. [Modified from diagrams by Roger  Kornberg, Stanford University. PDBid 1I6H.] Q See Interactive  Exercise 50.   Chapter 26 Transcription and RNA Processing X-ray structure of an RNA-DNA hybrid helix alone (Fig. 24-4). Nearly all   contacts that the protein makes with the RNA and DNA are to their  sugar-phosphate backbones; none are with the edges of their bases. The speci¬  ficity of the enzyme for a ribonucleotide rather than a deoxyribonucleotide ap¬  pears due to the enzyme’s recognition of both the incoming ribose sugar and  the RNA-DNA hybrid helix. After about one turn of hybrid helix, a loop ex¬  tending from the clamp known as the “rudder” separates the RNA and tem¬  plate DNA strands, thereby permitting the DNA double helix to re-form as  it exits the enzyme (although the unpaired 5' tail of the nontemplate strand  and the 3' tail of the template strand are disordered in the X-ray structure).   How does RNAP translocate its bound DNA-RNA assembly at the end of  each catalytic cycle? A highly conserved helical segment of Rpbl called the  “bridge” (because it bridges the two pincers forming the enzymes cleft) non-  specifically contacts the template DNA base at the +1 position. The bridge is  straight in the X-ray structures of RNAP II (Figs. 26-12 a and 26-13/2) but bent  in the X-ray structure of the bacterial RNAPs (Fig. 26-2). If the bridge helix, in  fact, alternates between its straight and bent conformations, it would move by  3 to 4 A. Kornberg has therefore speculated that translocation occurs through  the bending of the bridge helix so as to push the paired nucleotides at position  i + 1 to position i — 1. The recovery of the bridge helix to its straight confor¬  mation would then yield an empty site at position i + 1 for entry of the next  NTP, thereby preparing the enzyme for a new round of nucleotide addition. The  reversal of this process is presumably prevented by the binding of the next sub¬  strate NTP, and hence this mechanism is that of a Brownian ratchet [in which  otherwise random thermal (Brownian) back-and-forth fluctuations are converted  to coherent forward motion by inhibiting the backward motion].   RNAP II selects its substrate ribonucleotide through a two-stage process.  The incoming NTP first binds to the so-called E (for entry) site (Fig. 26-14),  which exhibits no selectivity for the identity of its base. The NTP then piv¬  ots to enter the A (for addition) site, which accepts only an NTP that forms  a Watson-Crick base pair with the template base in the / + 1 position. This  process is mediated by the RNAPs so-called trigger loop, which swings in be¬  neath the correctly base-paired NTP in the A site to form an extensive hydro¬  gen-bonded network involving both the NTP and other portions of the RNAP,  interactions that acutely discriminate against dNTPs.   FIG. 26-1 4 The A and E sites and the trigger loop in RNA  polymerase II. A cutaway view of the transcribing complex viewed  as in Fig. 26-13. The template DNA is cyan, the nontemplate DNA  green, newly transcribed RNA red, GTP in the A site orange, and ATP    in the E site blue. The trigger loop is magenta, the bridge helix is  green, and the two Mg 2+ ions at the active site are represented by  magenta spheres. The RNAP II surface is gray. [Courtesy of Dong  Wang and Roger Kornberg, Stanford University. PDBid 2E2H.]   RNAPs Can Correct Their Mistakes. RNAPs cannot read through a dam- Section 2 Transcription in Eukaryotes   aged template strand and consequently stall at the damage site. Moreover, if   into RNA, the DNA-RNA hybrid helix becomes distorted, which also causes   the RNAP to stall. How, then, do RNAPs avoid accumulating at damaged or   mispaired sites, which, if it occurred on an essential gene, would be lethal?   Instead, they frequently backtrack such that the RNAs penultimate nucleotide,  which was in the i — \ position, has re-entered the i + 1 position and the  3'-nucleotide, now in the i + 2 position, enters the secondary channel where  it binds in the so-called P (for proofreading) site. If the forward movement of  the RNA is impeded by damage to the template or by mispairing, further  backtracking becomes favored so that several more ribonucleotides enter the  secondary channel. The backtracking of only one or a few nucleotides is re¬  versible. Otherwise, transcription is arrested until the RNA is hydrolytically  cleaved at the active site. In E. coli, this requires the assistance of the homol¬  ogous proteins GreA and/or GreB, whereas with RNAP II, this function is  carried out by the unrelated protein TFIIS. These proteins induce the RNAP  active site to hydrolyze the phosphodiester bond between the ribonucleotides  in the / + 1 and i — l positions (a reaction that is not the reverse of the  polymerase reaction since this would be pyrophosphorolysis). In this way,   RNAP can correct its mistakes and resume RNA synthesis. RNAP I and  RNAP III also efficiently correct their mistakes.   In eukaryotes, RNAP does not include a removable cr factor. Instead, a number  of accessory proteins identify promoters and recruit RNAP to the transcription  start site (as described more fully in Section 26-2C). As expected, eukaryotic  promoters are more complex and diverse than prokaryotic promoters. In addi¬  tion, the three eukaryotic RNAPs recognize different types of promoters.   Mammalian RNA Polymerase I Has a Bipartite Promoter. Both prokaryotic  and eukaryotic genomes contain multiple copies of their rRNA genes in or¬  der to meet the enormous demand for these rRNAs (which comprise, e.g.,   80% of an E. coli cells RNA content). Since the numerous rRNA genes in a  given eukaryotic cell have essentially identical sequences, its RNAP I recog¬  nizes only one promoter. Yet, in contrast to RNAP II and III promoters,   RNAP I promoters are species specific; that is, an RNAP I recognizes only its  own promoter and those of closely related species.   RNAP I promoters were identified by determining how the transcription  rate of an rRNA gene is affected by a series of increasingly longer deletions  approaching its transcription start site from either its upstream or its down¬  stream side. Such studies have indicated, for example, that mammalian RNAP  I requires a so-called core promoter element, which spans positions — 31 to  + 6 and hence overlaps the transcribed region. However, efficient transcription  also requires an upstream promoter element, which is located between  residues —187 and —107. These elements, which are G + C-rich and ~85%  identical, are bound by specific transcription factors which then recruit RNAP I  to the transcription start site.   RNA Polymerase II Promoters Are Complex and Diverse. The promoters  recognized by RNAP II are considerably longer and more diverse than those  of prokaryotic genes. The structural genes expressed in all tissues (the house¬  keeping genes, which are thought to be constitutively transcribed) have GC-  rich stretches of DNA located upstream from their transcription start sites.   These CpG islands (Box 25-4) function analogously to prokaryotic promoters. On  the other hand, structural genes that are selectively expressed in one or a few   Chapter 26 Transcription and RNA Processing ce JJ types often lack these GC-rich sequences. Instead, they contain a core pro¬   moter of 40 to 50 nt that includes the transcription start site. Within the core  promoter for a given gene are one or more conserved elements that act—for  the most part in a cooperative fashion—to direct transcription initiation.   Among the best-known core promoter elements is the TATA box, an AT-  rich sequence located 25 to 31 bp upstream from the transcription start site.  The TATA box (consensus sequence TATAjAx) resembles the —10 region  of a prokaryotic promoter (TATAAT), although it differs in its location rela¬  tive to the transcription start site ( — 27 versus —10). Around two-thirds of  protein-encoding genes lack a TATA box, but approximately half have a con¬  served 7-nt Inr (initiator) element that includes the initiating (+1) nucleotide.  The consensus sequences and relative positions of several core promoter ele¬  ments are shown in Fig. 26-15. Most of these elements have been identified  from their conservation in many genes from different eukaryotic species, and  their participation in RNAP II-directed transcription has been verified exper¬  imentally by mutating their sequences and/or positions and by introducing  them into genes whose original core promoter elements had been deleted.  None of the core promoter elements occurs in all promoters, although some  general patterns are apparent; for example, core promoters that lack a TATA  box use MTE or DPE, and vice versa. All of the core promoter elements,  either singly or as sets, interact with their corresponding protein factors that  in turn recruit RNAP II to the initiation site (Section 26-2C).   The gene region extending between about — 50 and —110 also contains  promoter elements. For instance, many eukaryotic structural genes have a con¬  served consensus sequence of CCAAT (the CCAAT box) located between  about —70 and —90 whose alteration greatly reduces the genes transcription  rate. Evidently, the sequences upstream of the core promoter constitute additional  DNA-binding sites for proteins involved in transcription initiation.   Enhancers Are Transcriptional Activators That Can Have Variable Positions  and Orientations. Perhaps the most surprising aspect of eukaryotic transcriptional  control elements is that some of them need not have fixed positions and orientations  relative to their corresponding transcribed sequences. For example, the genome of  simian virus 40 (SV40), in which such elements were first discovered, con¬  tains two repeated sequences of 72 bp each that are located upstream from the  promoter for early gene expression. Transcription is unaffected if one of the re¬  peats is deleted but is nearly eliminated when both are absent. The analysis of  a series of SV40 mutants containing only one of the repeats demonstrated that  its ability to stimulate transcription from its corresponding promoter is all but  independent of its position and orientation. Indeed, transcription is unimpaired  when this segment is several thousand base pairs upstream or downstream from  the transcription start site. Gene segments with such properties are named en¬  hancers to indicate that they differ from promoters, which are site-specific and  strand-specific regarding transcription initiation.    FIG. 26-1 5 Some core promoter sequences for RNA  polymerase II. The consensus sequence of each element is given  (some positions can accommodate two or more nucleotides; N  represents any nucleotide). The approximate positions of the various  motifs are given relative to the transcription initiation (+1) site. Note  that a specific core promoter may contain all, some, or none of these    motifs. The BRE (TFIIB recognition element) is an upstream  extension of the TATA box. The DPE (downstream core promoter  element) and the MTE (motif fen element) must be accompanied  by an Inr.   Compare these sequences to the E. coli promoter in Fig. 26-6.   Enhancers occur in cellular genes as well as eukaryotic viruses and are re- Section 2 Transcription in Eukaryotes   quired for the full activities of their cognate promoters. Rather than interacting  with RNAP II directly, enhancers are recognized by specific transcription factors that  stimulate RNAP II to bind to the corresponding but distant promoter. This requires  that the DNA between the enhancer and promoter loop around so that the tran¬  scription factor can simultaneously contact the enhancer and RNAP II at the  promoter. Most cellular enhancers are associated with genes that are selectively  expressed in specific tissues. It therefore seems, as we discuss in Section 28-3B,  that enhancers mediate much of the selective gene expression in eukaryotes.   RNA Polymerase III Promoters Can Be Located Downstream from Their  Transcription Start Sites. The promoters of some genes transcribed by RNAP  III are located entirely within the genes transcribed regions. In a gene for the 5S  RNA of Xenopus borealis , deletions of base sequences that start from outside  one or the other end of the transcribed portion of the gene prevent transcrip¬  tion only if they extend into the segment between nucleotides +40 and +80.   This portion of the gene is effective as a promoter because it contains the  binding site for a transcription factor that stimulates upstream binding of  RNAP III. Further studies have shown, however, that the promoters of other  RNAP Ill-transcribed genes lie entirely upstream of their start sites. These up¬  stream sites also bind transcription factors that recruit RNAP III.   Differentiated eukaryotic cells possess a remarkable capacity for the selective ex¬  pression of specific genes. The synthesis rates of a particular protein in two cells  of the same organism may differ by as much as a factor of 10 9 . Thus, for ex¬  ample, reticulocytes (immature red blood cells) synthesize large amounts of he¬  moglobin but no detectable insulin, whereas the pancreatic (3 cells produce large  quantities of insulin but no hemoglobin. In contrast, prokaryotic systems gen¬  erally exhibit no more than a thousandfold range in their transcription rates so  that at least a few copies of all the proteins they encode are present in any cell.   Nevertheless, the basic mechanism for initiating transcription of structural genes is  the same in eukaryotes and prokaryotes: Protein factors bind selectively to the pro¬  moter regions of DNA. With class II promoters (those transcribed by RNAP II),  a complex of at least six general transcription factors (GTFs; Table 26-2) op¬  erates as a formal equivalent of a prokaryotic a factor. The structures of several  eukaryotic transcription factors are described in Section 24-4C.    TABLE 26-2 Properties and Functions of the Eukaryotic General Transcription Factors   Chapter 26 Transcription and RNA Processing The six GTFs, which are highly conserved from yeast to humans, are re¬   quired for the synthesis of all mRNAs, even those with strong promoters. The  GTFs allow a low (basal) level of transcription that can be augmented by the  participation of other gene-specific factors. We will examine the action of some  of those other proteins in Section 28-3. The GTFs, whose names begin with  TF (for transcription factor) followed by the Roman numeral II to indicate that  they are involved in transcription by RNAP II, combine with the enzyme and  promoter DNA in an ordered pathway to form a preinitiation complex (PIC).   PIC Formation Often Begins with TATA-Binding Protein Binding to the  TATA Box. As indicated in Section 26-2B, the promoters of many eukaryotic  structural genes contain a TATA box at around position —27. The GTFs are  targeted to this and other sequences that make up the core promoter. The first  transcription factor to bind to TATA box-containing promoters is the  TATA-binding protein (TBP), which as its name indicates, binds to the  TATA box and thereby helps identify the transcription start site. TBP is sub¬  sequently joined on the promoter by additional subunits to form, in humans,  the ~1122-kD, 17-subunit complex TFIID.   The highly conserved C-terminal domain of TBP contains two ~40%  identical direct repeats of 66 residues separated by a highly basic segment. Its  X-ray structure, which was independently determined by Roger Kornberg and  Stephen Burley, reveals a saddle-shaped molecule that consists of two struc¬  turally similar domains arranged with pseudo twofold symmetry (Fig. 26-16^).  TBP s structure suggests that it could fit snugly astride an undistorted B-DNA  helix. However, the X-ray structures of TBP-DNA complexes, independently  determined by Burley and Paul Sigler, reveal a quite different interaction. The  DNA indeed binds to the concave surface of TBP but with its duplex axis  nearly perpendicular rather than parallel to the saddles cylindrical axis  (Fig. 26-1 6b). The bound DNA is kinked by —45° between the first two base  pairs of the 7-bp TATA box and between its last base pair and the succeeding   (a) A ribbon diagram of yeast TBP in the absence of DNA, in which a  helices are red, p strands are yellow, and the remainder of the  polypeptide backbone is cyan. The protein’s pseudotwofold axis of  symmetry is vertical. [Based on an X-ray structure by Roger Kornberg,  Stanford University. PDBid 1TBPJ (b) The structure of human TBP in  complex with a 16-bp TATA box-containing duplex DNA. The DNA,  which is largely in the B form, is drawn in stick form and colored  according to atom type with sense strand C green, antisense strand C  cyan, N blue, 0 red, and P orange. It enters its binding site with the 5'  end of the sense strand on the right and exits on the left with its helix  axis nearly perpendicular to the page. The protein is drawn in worm form  colored in rainbow order from its N-terminus (blue) to its C-terminus  (red). The side chains of Phe residues 193, 210, 284, and 301, which  induce sharp kinks in the DNA, are drawn in stick form (magenta).  Between the kinks, which are located at each end of the TATA box, the  DNA is partially unwound with the central eight strands of the protein’s  ten-stranded p sheet inserted into the DNA’s greatly widened minor  groove. The TBP does not contact the DNA’s major groove. [Based on an  X-ray structure by Stephen Burley, Structural GenomiX, Inc., San Diego,  California. PDBid 1CDW.] ©See Interactive Exercise 51.   tie conformational change on binding DNA, does so via hydrogen bonding  and van der Waals interactions. The kinked and partially unwound DNA is  stabilized by a wedge of two Phe side chains on each side of the saddle struc¬  ture that pry apart the two base pairs flanking each kink from their minor  groove sides. The bent conformation of DNA creates a stage for the assembly  of other proteins to form the PIC.   TFIIA, TFIIB, and TAFs Interact with TBP and RNAP II. The other GTFs re¬  quired for basal transcription assemble as is diagrammed in Fig. 26-17. The PIC  requires, at a minimum, TBP, TFIIB, TFIIE, TFIIF, and TFIIH. TFIIB con¬  sists of two domains, an N-terminal domain (TFIIB N ), which interacts with  RNAP II, and a C-terminal domain (TFIIB C ), which binds DNA and interacts  with TBP. The X-ray structures of TFIIA-TBP-DNA and TFIIB C -TBP-DNA  complexes have permitted the generation of a plausible model for the   £ The TBP component of TFIID binds  /to the TATA box of the promoter.    FIG. 26-1 7 The assembly of the preinitiation complex (PIC) D., Curr. Opin. Cell Biol. 4, 490 (1992).] O See Interactive   on a TATA box-containing promoter. [After Zawel, L. and Reinberg, Exercise 51.   (a) Model of the TFIIA-TBP-TFIIB c -promoter complex based on the  independently determined X-ray crystal structures of the  TFIIA-TBP-promoter and TFIIB c -TBP-promoter complexes. The  DNA (white sticks ) has been extended in both directions beyond the  TATA box. The pseudosymmetrical TBP (cyan and purple ), viewed from  above, induces two sharp kinks in the DNA, giving it a cranklike  shape. Both TFIIA (yellow and green) and TFIIB C (magenta and red)  interact with TBP, but each interacts with independent sites on the  DNA. The transcription start site (+1) is at the left. [Courtesy of  Stephen Burley, Structural GenomiX, Inc., San Diego, California. Based  on PDBids 1YTF and 1VOL] (b) Model of the TFIIB-TBP-RNAP    ll-promoter open complex based on the X-ray structures of the  TFIIB n -RNAP II and TFIIB c -TBP-promoter complexes. The view of  the RNAP II is approximately from the top of Fig. 26-13 after it has  been rotated 90° about the horizontal axis. The proteins are  represented by their solvent-accessible surfaces with TFIIB red, TBP  purple, and the various portions of RNAP II in other colors. The DNA  is drawn in cartoon form with its template and nontemplate strands  cyan and green and with the dotted segments representing its  transcription bubble. Its upstream end is at the upper right. Note how  the leftmost portion of TFIIB, its B finger, extends into the active  center of RNAP II to contact the DNA. [Courtesy of Roger Kornberg,  Stanford University. Based on PDBids 3K7A, 1 R5U, and 1VOL]    TFIIA-TFIIB C -TBP-DNA complex (Fig. 26-18^). The three proteins bind to  the DNA just upstream from the transcription start site, leaving ample room  for additional proteins and RNAP II to bind. Since the pseudosymmetric TBP  has been shown to bind to the TATA box in either orientation, it appears that  base-specific interactions between TFIIB and the promoter function to position  TFIIB to properly orient the TBP on the promoter.   The X-ray structure of the TFIIB n -RNAP II complex, also determined  by Kornberg, reveals that TFIIB N contacts the “dock” domain of RNAP II  near its RNA exit channel and that TFIIB N inserts its “finger” domain into  the active center of RNAP II. A model of the TFIIB-TBP-RNAP II-DNA  complex based on this X-ray structure together with that of the  TFIIB C -TBP-DNA complex is shown in Fig. 26-18 A.   The remaining components of TFIID, which are known as TBP-associ-  ated factors (TAFs), form a horseshoe-shaped complex to which TFIIA and  TFIIB are bound (Fig. 26-19). TBP is located at the top of the cavity, and  the promoter DNA passes through the cavity. Portions of 9 of the 14 known    FIG. 26-19 Electron microscopy-based image of the human  TFIID-TFIIA-TFIIB complex at 35 A resolution. The complex consists of three  domains, A, B, and C, arranged in a horseshoe shape roughly 200 A wide, 135 A high,  and 110 A thick. The red and green meshes indicate the positions of TFIIA and TFIIB  as determined by comparison with the EM-based images of the TFIID-TFIIA complex  and TFIID alone. The yellow mesh indicates the binding position of an anti-TBP  antibody. [Courtesy of Eva Nogales, University of California at Berkeley.]   TAFs, which are highly conserved from yeast to humans, are homologous to Section 2 Transcription in Eukaryotes   nonlinker histones (Section 24-5A). Indeed, X-ray and other studies suggest  that four of the TAFs associate to form a nucleosome-like heterooctamer.   Nevertheless, it is unlikely that the DNA wraps around the TAFs as it does  in a nucleosome because many of the histone residues that make critical con¬  tacts with the DNA in the nucleosome have not been conserved in the  histonelike TAFs.   In the final steps of PIC formation (Fig. 26-17), TFIIF recruits RNAP II  to the promoter in a manner reminiscent of the way that cr factor interacts  with bacterial RNAP. In fact, the second largest of TFIIF s three subunits is  homologous to cr 70 , the predominant bacterial cr factor, and, moreover, can  specifically interact with bacterial RNAPs (although it does not participate in  promoter recognition). Finally, TFIIE and TFIIH join the assembly. Once  this complex has been assembled, the ATP-dependent helicase activity of  TFIIH induces the formation of the open complex so that RNA synthesis  can commence. Note that the human PIC, exclusive of the ~ 12-subunit,   ~600-kD RNAP II, contains 37 subunits with an aggregate mass of —2000 kD  (Table 26-2). Many of the proteins in the PIC are the targets of transcrip¬  tional regulators.   Promoters That Lack a TATA Box Also Bind TBP. Since the TATA-binding  protein is a component of TFIID, a general transcription factor for RNAP II,  how does the preinitiation complex form properly at TATA-less promoters? In  many cases, the presence of the Inr element is sufficient to direct RNAP II to  the correct start site. These systems require the participation of many of the  same GTFs that initiate transcription from TATA box—containing promoters.   Surprisingly, they also require TBP. This suggests that with TATA-less promot¬  ers, Inr recruits TFIID such that its component TBP binds to the —30 region  in a sequence-nonspecific manner. Indeed, in Inr-containing promoters that  also contain a TATA box, the two elements act synergistically to promote tran¬  scriptional initiation. Nevertheless, a mutant TBP that is defective in TATA  box binding will support efficient transcription from some TATA-less promot¬  ers although not from others. This suggests that the former promoters do not  require a stable interaction with TBP. Consequently, the scheme outlined in  Fig. 26-17 should be taken as a flexible framework for RNAP II transcription  initiation in eukaryotes, with the exact protein requirements depending on the  nature of the promoter and the presence of additional protein factors.   TBP Is a Universal Transcription Factor. RNAP I and RNAP III require dif¬  ferent sets of GTFs from each other and from RNAP II to initiate transcrip¬  tion at their respective promoters. This is not unexpected considering the very  different organizations of these three classes of promoters (Section 26-2B).   Indeed, the promoters recognized by RNAP I (class I promoters) and nearly  all those recognized by RNAP III (class III promoters) lack TATA boxes. Thus,  it came as a surprise when it was demonstrated that TBP is required for ini¬  tiation by both RNAP I and RNAP III. TBP participates by combining with  different sets of TAFs to form the GTFs SLI (with class I promoters) and  TFIIIB (with class III promoters). As with certain class II TATA-less promot¬  ers, a TBP mutant that is defective for TATA box binding can still support in  vitro transcriptional initiation by both RNAP I and RNAP III. Clearly, TBP,  the only known universal transcription factor, is an unusually versatile protein.   Elongation Requires Different Transcription Factors. After RNAP II initi¬  ates RNA synthesis and successfully produces a short transcript, the transcrip¬  tion machinery undergoes a transition to the elongation mode. The switch  appears to involve displacement of the finger domain of TFIIB, which would  otherwise clash with the growing RNA chain in the active site, as well as phos¬  phorylation of the C-terminal domain (CTD) of RNAP IPs Rpbl subunit.   • What are the functions of the three  eukaryotic RNA polymerases?   • Explain the functions of RNAP M’s CTD,  clamp, wall, funnel, rudder, bridge, and  trigger.   • What are the advantages of having multi¬  ple types of promoters and enhancers?   • Summarize the role of TBP in transcription  initiation in eukaryotes.   • Explain why both the sequence and struc¬  ture of DNA help recruit RNA polymerase.   • Why is TBP considered to be a universal  transcription factor?    Phosphorylated RNAP II releases some of the transcription-initiating factors  and advances beyond the promoter region. In fact, when RNAP II moves away  from (“clears”) the promoter, it leaves behind some GTFs, including TFIID.  These proteins can reinitiate transcription by recruiting another RNAP II to  the promoter. Consequently, the first RNAP to transcribe a gene may act as a  “pioneer” polymerase that helps pave the way for additional rounds of transcription.   During elongation, a six-protein complex called Elongator binds to the  phosphorylated CTD of Rpbl, taking the place of the jettisoned transcription  factors. Although Elongator is not essential for transcription by RNAP II  in vitro , its presence accelerates transcription. Interestingly, TFIIF and TFIIH  remain associated with the polymerase during elongation. Well over a dozen  other proteins are known to associate with an elongating RNAP II. Some of  the proteins are involved in processing the nascent RNA, modifying the pack¬  aging of the DNA template, and terminating transcription.   Eukaryotes Lack Precise Transcription Termination Sites. The sequences  signaling transcriptional termination in eukaryotes have not been identified.  This is largely because the termination process is imprecise; that is, the pri¬  mary transcripts of a given structural gene have heterogeneous 3' sequences.  However ; a precise termination site is not required because the transcript under¬  goes processing that includes endonucleolytic cleavage at a specific site (see below).  The endonuclease may act even while the polymerase is still transcribing, so  RNA cleavage itself may signal the polymerase to stop.    • Eukaryotic mRNAs are modified by a 5' cap and a 3' poly(A) tail.   • Eukaryotic genes include introns that must be spliced out by the action of  snRNPs in the spliceosome.   • A single gene can generate several protein products through alternative  mRNA splicing.   • Prokaryotic and eukaryotic rRNA and tRNA precursors are variously processed by  endonucleolytic cleavage, covalent modification, splicing, and nucleotide addition.    The immediate products of transcription, the primary transcripts, are not  necessarily functional. In order to acquire biological activity, many of them  must be specifically altered: (1) by the exo- and endonucleolytic removal of  polynucleotide segments; (2) by appending nucleotide sequences to their 3'  and 5' ends; and/or (3) by the modification of specific nucleotide residues.  The three major classes of RNA—mRNA, rRNA, and tRNA—are altered in  different ways in prokaryotes and in eukaryotes. In this section, we outline  these posttranscriptional modification processes.    A Messenger RNAs Undergo 5' Capping and Addition of a 3' Tail   In prokaryotes, most primary mRNA transcripts are translated without fur¬  ther modification. Indeed, protein synthesis usually begins before transcription  is complete (Section 27-4). In eukaryotes, however, mRNAs are synthesized  in the cell nucleus, whereas translation occurs in the cytosol. Eukaryotic  mRNA transcripts can therefore undergo extensive posttranscriptional process¬  ing while still in the nucleus.   Eukaryotic mRNAs Have 5' Caps. Eukaryotic mRNAs have a cap structure  consisting of a 7-methylguanosine (m 7 G) residue joined to the transcript's ini¬  tial (5j nucleotide via a 5'-5' triphosphate bridge (Fig. 26 - 20 ). The cap, which  adds to the growing transcript when it is ~30 nt long, identifies the eukaryotic   translation start site (Section 27-4A). Capping involves several enzymatic  reactions: (1) the removal of the leading phosphate group from the mRNAs  5' terminal triphosphate group by an RNA triphosphatase; (2) the guanyly-  lation of the mRNA by capping enzyme, which requires GTP and yields the  5'-5' triphosphate bridge and PP Z ; and (3) the methylation of guanine by  guanine-7-methyltransferase, in which the methyl group is supplied by S-  adenosylmethionine (SAM). In addition, the cap may be O 2 -methylated at  the first and second nucleotides of the transcript by a SAM-requiring 2'-  O-methyltransferase. Capping enzyme binds to RNAP IPs phosphorylated  CTD (Section 26-2A), and hence it appears that capping marks the comple¬  tion of RNAP IPs switch from transcription initiation to elongation. Capped  mRNAs are resistant to 5'-exonucleolytic degradation.   Eukaryotic mRNAs Have Poly(A) Tails. Eukaryotic RNA transcripts have  heterogeneous 3' sequences because the transcription termination process is  imprecise. Mature eukaryotic mRNAs, however, have well-defined 3' ends termi¬  nating in poly (A) tails of —250 nt (—80 nt in yeast). The poly(A) tails are  enzymatically appended to the primary transcripts in two reactions:   1. A transcript is cleaved 15 to 25 nt past a highly conserved AAUAAA  sequence and less than 50 nt before a less-conserved U-rich or  G + U-rich sequence. The precision of this cleavage reaction has ap¬  parently eliminated the need for accurate transcription termination.  Nevertheless, the identity of the endonuclease that cleaves the RNA is  uncertain although cleavage factors I and II (CFI and CFII) are re¬  quired for this process.   2. The poly (A) tail is subsequently generated from ATP through the step¬  wise action of poly (A) polymerase (PAP), a template-independent  RNA polymerase that elongates an mRNA primer with a free 3'-OH  group. PAP is activated by cleavage and polyadenylation specificity  factor (CPSF) when the latter protein recognizes the AAUAAA se¬  quence. Once the poly(A) tail has grown to — 10 residues, the AAUAAA  sequence is no longer required for further chain elongation. This sug¬  gests that CPSF becomes disengaged from its recognition site in a man¬  ner reminiscent of the way cr factor is released from the transcription  initiation site once the elongation of prokaryotic RNA is underway.   CPSF binds to the phosphorylated RNAP II CTD; deleting the CTD inhibits  polyadenylation. Evidently, the CTD couples polyadenylation to transcription.   PAP is part of a 500- to 1000-kD complex that also contains the proteins  required for mRNA cleavage. Consequently, the cleaved transcript is polyadeny-  lated before it can dissociate and be digested by cellular nucleases (see below).  The maximum length of the poly(A) tail may be determined by the stoichio¬  metric binding of multiple copies of poly (A) binding protein II (PAB II).   In vitro studies indicate that a poly(A) tail is not required for mRNA trans¬  lation. Rather, the observation that an mRNAs poly (A) tail shortens as it ages  in the cytosol suggests that poly(A) tails have a protective role. In fact, the only  mature mRNAs that lack poly(A) tails, those encoding histones (which are re¬  quired in large quantities only during the relatively short period during the cell  cycle when DNA is being replicated), have cytosolic lifetimes of <30 min ver¬  sus hours or days for most other mRNAs. The poly(A) tails are specifically com-  plexed in the cytosol by poly (A)-binding protein (PABP; not related to PAB  II), which organizes poly(A)-bearing mRNAs into ribonucleoprotein particles.  PABP is thought to protect mRNA from degradation as is suggested, for ex¬  ample, by the observation that the addition of PABP to a cell-free system con¬  taining mRNA and mRNA-degrading nucleases greatly reduces the rate at which  the mRNAs are degraded and the rate at which their poly(A) tails are shortened.  However, as we will see (Section 28-3C), the degradation of mRNAs is regulated  by multiple signals, not just the lengths of their poly(A) tails.    FIG. 26-20 Structure of the 5' cap of  eukaryotic mRNAs. A cap may be   CF-methylated at the transcript’s leading  nucleoside (the predominant cap in multicellular  organisms), at its first two nucleosides, or at  neither of those positions (the predominant cap  in unicellular eukaryotes). If the first nucleoside  is adenosine (it is usually a purine), it may also  be /V 6 -methylated.   Chapter 26 Transcription and RNA Processing The cleavage of a transcript past its AAUAAA sequence does not, in it¬   self, terminate transcription. However, in yeast, the protein Rttl03, which  binds to the phosphorylated CTD of RNAP II, recognizes the AAUAAA se¬  quence and recruits the 5 r —> 3' exonuclease known as Rati (Xrn2 in hu¬  mans). Then, in what is termed the torpedo model, the highly processive  Ratl/Xrn2 loads onto the newly liberated 5' end of the still nascent RNA and  rapidly degrades it until it intercepts the RNAP and induces it to terminate  RNA synthesis. It has been hypothesized that this occurs in much the same  way as Rho factor terminates bacterial transcription (Section 26-ID). This  frees the RNAP to initiate a new round of transcription.   The most striking difference between eukaryotic and prokaryotic structural  genes is that the coding sequences of most eukaryotic genes are interspersed with  unexpressed regions . The primary transcripts, also called pre-mRNAs or het¬  erogeneous nuclear RNAs (hnRNAs), are variable in length and are much  larger (~2000 to >20,000 nt) than expected from the known sizes of eukary¬  otic proteins. Rapid labeling experiments demonstrated that little of the  hnRNA is ever transported to the cytosol; most of it is quickly degraded in  the nucleus. Yet the hnRNAs 5' caps and 3' tails eventually appear in cytoso¬  lic mRNAs. The straightforward explanation of these observations, that pre-  mRNAs are processed by the excision of internal sequences, seemed so bizarre  that it came as a great surprise in 1977 when Phillip Sharp and Richard  Roberts independently demonstrated that this is actually the case (Box 26-3).  Thus, pre-mRNAs are processed by the excision of nonexpressed intervening se¬  quences (introns) y following which the flanking expressed sequences (exons)  are spliced (joined) together.   A pre-mRNA typically contains eight introns whose aggregate length av¬  erages 4 to 10 times that of its exons. This situation is graphically illustrated  in Fig. 26-21, which is an electron micrograph of chicken ovalbumin mRNA   FIG. 26-21 The chicken ovalbumin gene  and its mRNA. The electron micrograph and its  interpretive drawing show the hybridization of the  antisense (template) strand of the chicken  ovalbumin gene and its corresponding mRNA.  The complementary segments of the DNA (blue  line in drawing) and mRNA (green dashed line)  have annealed to reveal the exon positions  (/_, 1-7). The looped-out segments (I—VII), which  have no complementary sequences in the  mRNA, are the introns. [From Chambon,   For several decades following the discovery  that DNA is the genetic material in all  organisms, it was believed that genes were  continuous sequences of nucleic acid. A  messenger RNA molecule was thought to be  a faithful copy of the gene, aligning exactly  with the DNA sequence. In fact, this is largely  true for simple genetic systems such as  bacteria and bacteriophages, which were widely  used in early studies of molecular biology.  However, Richard Roberts and Phillip Sharp,  working independently, showed in 1977 that  genes could be discontinuous. We now  understand that the mRNA segments  corresponding to these “split genes” must be  spliced together in order for the gene to be  properly expressed.   Richard Roberts began his scientific career  as an organic chemist. While working on his thesis, he read a  book by John Kendrew (who had determined the X-ray structure  of myoglobin; Section 7-1 A) and became hooked on the new field  of molecular biology. His first project in the world of nucleic acids  was to determine the sequence of nucleotides in a tRNA  molecule. He was able to take advantage of new sequencing  techniques devised by Frederick Sanger (who subsequently  developed the dideoxy sequencing method; Section 3-4C). Roberts  next turned his attention to restriction endonucleases. He  recognized that the enzymes could be invaluable tools for cutting  large DNA molecules down to size, including the adenovirus-2  genome that he began to characterize.   During roughly the same period, Phillip Sharp completed a  thesis describing his use of statistical and physical theories to  describe the DNA polymer. By his own admission, he was not a  skilled experimenter. His outlook changed, however, when he  joined a molecular biology laboratory that made extensive use of  electron microscopy to examine DNA-RNA heteroduplexes. When  Sharp subsequently turned to the study of eukaryotic gene  expression, the only practical experimental systems were animal  viruses with DNA genomes, such as adenovirus-2. Because that    virus, which is a cause of the common cold, infects mammalian  cells, its genes were thought to resemble those of its host.   Thus, in the mid-1970s, both Roberts (at the Cold Spring  Harbor Laboratory on Long Island, New York) and Sharp (at the  Massachusetts Institute of Technology) came to be mapping the  adenovirus genome. They located viral genes by obtaining  expressed mRNA molecules and hybridizing them to segments of  the viral DNA. Sharp observed that the nuclei of virus-infected  cells accumulated viral mRNAs that were not transported to the  cytoplasm, and he speculated that the nuclear mRNAs were  processed in order to generate the cytoplasmic mRNAs.   Meanwhile, Roberts observed that “late” (mature) mRNAs began  with an oligonucleotide that was not encoded in the DNA next to  the main body of the mRNA.   Both scientists prepared samples in which the mRNA was  allowed to hybridize with the DNA, and then visualized the hybrid by  electron microscopy. The results were as exciting as they were  unexpected: A single mRNA molecule hybridized with as many as  four well-separated segments of the DNA molecule, so that the  unpaired DNA sequences between the hybrid segments looped out  (as in Fig. 26-21). The inescapable conclusion was that viral genetic  information was organized discontinuously, a notion that contradicted  commonly held views about the nature of genes. Nevertheless,  other researchers were eager to see whether split genes occurred  in other viruses and animal cells. Within a year, similar results were  confirmed for a handful of other genes. Subsequent research  showed that most animal genes are discontinuous. The sequences  that are ultimately expressed were termed exons, and the  intervening sequences that did not appear in the mature mRNA  were named introns. For their discoveries of split genes, Roberts  and Sharp shared the 1993 Nobel Prize in Physiology or Medicine.   The existence of split genes solved some biochemical puzzles  but, as is always the case with important discoveries, introduced  new ones: How are introns cut out and the remaining exons  joined together? Are the same exons spliced together in all cells?  Could evolution occur more rapidly through exon shuffling?  Answers to these questions are still being refined.   Chow, L.T., Gelinas, R.E., Broker, T.R., and Roberts, R.J., An amazing sequence  arrangement at the 5' ends of adenovirus 2 messenger RNA, Cell 12 , 1-8 (1977).  Berget, S.M., Moore, C., and Sharp, P.A., Spliced segments at the 5' terminus of  adenovirus 2 late mRNA, Proc. Natl. Acad. Sci. 74 , 3171-3175 (1977).    hybridized to the antisense (template) strand of the ovalbumin gene (ovalbu¬  min is the major protein component of egg white). In humans, the number  of introns in a gene varies from none to 364 (in the ~2400-kb gene encod¬  ing the 34,350-residue muscle protein titin, the largest known gene and  single-chain protein; Section 7-2A), with intron lengths ranging from ~65 to  ~800,000 nt (in the gene encoding the muscle protein dystrophin; Section 7-2A)  and averaging ~3500 nt (exons, in contrast, have lengths that average — 150 nt  and range up to 17,106 nt in the gene encoding titin). The introns from  corresponding genes in two vertebrate species rarely vary in number and po¬  sition, but often differ extensively in length and sequence so as to bear little  resemblance to one another.    ii 11 hi | | iv | | v | | vi | | vn   Cap— | | I | | II11 III | | IV | | V | | VI | | VII    FIG. 26-22 The sequence of steps in the production of  mature eukaryotic mRNA. This example shows the chicken  ovalbumin gene. Following transcription, the primary transcript is  capped and polyadenylated. The introns are excised and the exons    spliced together to form the mature mRNA. Note that capping and  splicing may occur cotranscriptionally, so that introns nearer the  capped 5' end may be excised before transcription of the 3' end is  complete.    The production of a translation-competent eukaryotic mRNA begins with  the transcription of the entire gene, including its introns (Fig. 26-22). Capping  occurs soon after initiation, and splicing commences during the elongation  ph ase of transcription. The mature mRNA emerges only after splicing is com¬  plete and the RNA has been polyadenylated. The mRNA is then transported  to the cytosol, where the ribosomes are located, for translation into protein.   Exons Are Spliced in a Two-Stage Reaction. Sequence comparisons of  exon—intron junctions from a diverse group of eukaryotes indicate that they  have a high degree of homology (Fig. 26-23), with most of them having an  invariant GU at the introns 5' boundary and an invariant AG at its 3' bound¬  ary. These sequences are necessary and sufficient to define a splice junction. The  splicing reaction occurs via two transesterification reactions (Fig. 26-24):   1. A 2',5 r -phosphodiester bond forms between an intron adenosine  residue and the introns 5 r -terminal phosphate group. The 5' exon is  thereby released and the intron assumes a novel lariat structure (so  called because of its shape). The adenosine at the lariat branch point  is typically located in a conserved sequence 20 to 50 residues upstream    FIG. 26-23 The consensus sequences at the exon-intron junctions of eukaryotic  pre-mRNAs. The subscripts indicate the percentage of pre-mRNAs in which the specified  base(s) occurs. Note that the 3' splice site is preceded by a tract of 11 predominantly  pyrimidine nucleotides. [Based on data from Padgett, R.A., Grabowski, P.J., Konarska, M.M.,  Seiler, S.S., and Sharp, P.A., Annu. Rev. Biochem. 55, 1123 (1986).]   transesterification reactions splice together the  exons of eukaryotic pre-mRNAs. The exons and  introns are drawn in blue and orange, and R and  Y represent purine and pyrimidine residues.   (1) The 2'-OH group of a specific intron A  residue nucleophilically attacks the 5'-phosphate  at the 5' intron boundary to displace the 3'  end of the 5' exon, thereby yielding a  2',5'-phosphodiester bond and forming a lariat  structure. ( 2 ) The liberated 3'-OH group attacks  the 5'-phosphate of the 5'-terminal residue of  the 3' exon, forming a 3',5'-phosphodiester bond,  thereby displacing the intron in lariat form and  splicing the two exons together.    How many bonds are broken and how  many are formed during splicing?    of the y splice site. Mutations that change this branch point A residue  abolish splicing at that site.   2. The y exons free 3'-OH group displaces the 3' end of the intron,  forming a phosphodiester bond with the 5'-terminal phosphate of the  y exon and yielding the spliced product. The intron is thereby elimi¬  nated in its lariat form with a free 3'-OH group. Mutations that alter  the conserved AG at the 3' splice junction block this second step,  although they do not interfere with lariat formation. The lariat is even¬  tually debranched (linearized) and, in vivo , is rapidly degraded.   Note that the splicing process proceeds without free energy input; its transes¬  terification reactions preserve the free energy of each cleaved phosphodiester  bond through the concomitant formation of a new one. The sequences required  for splicing are the short consensus sequences at the 3' and 5' splice sites and  at the branch site. Nevertheless, these sequences are poorly conserved, and even  highly sophisticated computer programs are only ~50% successful in predict¬  ing actual splice sites over apparently equally good candidates that are not.   A simplistic interpretation of Fig. 26-24 suggests that any 5' splice site  could be joined with any following 3' splice site, thereby eliminating all the  intervening exons together with the introns joining them. However, such exon  skipping does not normally take place (but see below). Rather, all of a  pre-mRNAs introns are individually excised in what appears to be a largely  fixed order that more or less proceeds in the 5' —> 3' direction. This occurs,  at least in part, because splicing occurs cotranscriptionally. Thus, as a newly  synthesized exon emerges from an RNAP II, it is bound by splicing factors (see  below) that are also bound to the RNAP IPs highly phosphorylated CTD. This  tethers the exon and its associated splicing machinery to the CTD so as to en¬  sure that splicing occurs when the next exon emerges from the RNAP II.   Splicing Is Mediated by snRNPs in the Spliceosome. How are splice junc¬  tions recognized and how are the two exons to be joined brought together in   FIG. 26-25 X-Ray structure of an Sm  dimer. The D3 protein is gold and the B protein  is blue. The p5 strand of D3 associates with the  p4 strand of B to form a continuous antiparallel  |3 sheet Note that their corresponding loops  extend in similar directions. [Courtesy of Kiyoshi  Nagai, MRC Laboratory of Molecular Biology,  Cambridge, U.K. PDBid 1 D3BJ    the splicing process? Part of the answer to these questions was established by  Joan Steitz going on the assumption that one nucleic acid is best recognized  by another. The eukaryotic nucleus contains numerous copies of highly con¬  served 60- to 300-nt RNAs called small nuclear RNAs (snRNAs), which  form protein complexes termed small nuclear ribonucleoproteins (snRNPs;  pronounced “snurps”). Steitz noted that the 5' end of one of these snRNAs,  Ul-snRNA (which is so named because of its high uridine content), is par¬  tially complementary to the consensus sequence of 5' splice junctions.  Apparently, Ul-snRNP recognizes the 5' splice junction. Other snRNPs that par¬  ticipate in splicing are U2-snRNP, U4-U6-snRNP (in which the U4- and  U6-snRNAs associate via base pairing), and U5-snRNP.   Splicing takes place in an —60S particle dubbed the spliceosome. The  spliceosome brings together a pre-mRNA, the snRNPs, and a variety of pre-  mRNA binding proteins. The spliceosome, which consists of 5 RNAs and —150  polypeptides, is comparable in size and complexity to the ribosome (Section 27-  3). It is a highly dynamic entity, with its various components associating and dis¬  sociating during specific stages of the splicing process. This includes rearrangement  of base-paired stems among the snRNAs, as the spliceosome carries out the two  transesterification reactions. The U6-snRNA coordinates a catalytically essential  metal ion that enhances the nucleophilicity of the attacking OH group and sta¬  bilizes the leaving group. The existence of self-splicing introns (see below) and the  results of experiments with protein-free snRNAs strongly suggest that the RNA  components of the spliceosome, rather than the proteins, catalyze splicing.   All four snRNPs involved in mRNA splicing contain the same so-called  snRNP core protein, which consists of seven Sm proteins, named B, D l5  D 2 , D 3 , E, F, and G proteins. These proteins collectively bind to a conserved  AAUUUGUGG sequence known as the Sm RNA motif, which occurs in  U1-, U2-, U4-, and U5-snRNAs. The X-ray structures of the D 3 B and D X D 2  heterodimers reveal that these four proteins share a common fold, which con¬  sists of an N-terminal helix followed by a five-stranded antiparallel (3 sheet  that is strongly bent so as to form a hydrophobic core (Fig. 26-25). Model  building based on the X-ray structures together with biochemical and mutagenic  evidence suggest that the seven Sm proteins form a heptameric ring in which  the fourth (3 strand of one Sm protein interacts with the fifth (3 strand of an  adjacent Sm protein through main-chain hydrogen bonding. The funnel-  shaped central hole of the Sm ring, which is lined with positive charges, is  large enough to accommodate snRNA. An Sm-like protein from the hyper-  thermophilic archaebacterium Pyrobaculum aerophilum forms a homohep-  tameric ring, suggesting that the seven eukaryotic Sm proteins arose through  a series of duplications of an archaeal Sm-like protein gene.   Mammalian Ul-snRNP consists of Ul-snRNA and ten proteins, namely,  the seven Sm proteins that are common to all U-snRNPs as well as three that  are specific to Ul-snRNP: U1-70K, Ul-A, and Ul-C. The predicted sec¬  ondary structure of the 165-nt Ul-snRNA (Fig. 26-26 a) contains five  double-helical stems, four of which come together at a four-way junc¬  tion. U1-70K and Ul-A bind directly to RNA stem-loops (SL) 1  and 2, respectively, whereas Ul-C is bound by other proteins.   Kiyoshi Nagai determined the X-ray structure of human U1  snRNP at 5.5 A resolution (Fig. 26-2 6b,c). At this low resolution  the major and minor grooves of the double-stranded RNA stems are visible.  Moreover, the helices and sheets of the proteins are apparent, which allows the  placement of protein folds of known structure. SL2 of the Ul-snRNA had  been altered and shortened to promote crystallization, which eliminated the  binding site for Ul-A. However, U1 snRNP in which Ul-A is depleted is ac¬  tive in a splicing assay. The Sm proteins form the predicted heteroheptameric  ring, which is ~70 A in diameter. The Sm RNA motif and SL4 are threaded  through the ring’s funnel-shaped central hole such that the Sm RNA motif in¬  teracts with the Sm proteins. Ul-C associates with the Sm ring via an inter¬  action between its zinc finger domain and the D3 subunit.   C ■ G  G ■ C  G * U  A G  G A  C C  G-C  G ■ C   U G  G-C  C-G  G-C  U U  C U  A-U  G ■ C  G - C-iw>  -G ■ C  G-C   S'-mgGpppA U|C*9> VAC| U G G ■ C A U A AU U U G UG G U A G U G * U Ggh-3 1    Fortuitously, the 5' ends of two neighboring Ul-snRNAs in the  crystal form a double-helical segment, which serves as a model for  how the 5' end of Ul-snRNA base-pairs with the 5' splice site of the pre-mRNA  in the spliceosome. The zinc finger domain of Ul-C interacts with this double  helix (left side of Fig. 26-26c) and presumably stabilizes it. This is consistent  with the observation that mutants of Ul-C in this region cannot initiate spliceo¬  some formation.    FIG. 26-26 X-Ray structure of human UI-snRNP. (a) The predicted  secondary structure of Ul-snRNA with the RNA segments at which the  proteins U1-70K and U1-A bind indicated. ^ is the symbol for pseudouridine  residues (Section 26-3C). (b) The seven Sm proteins, each shown in ribbon  form and differently colored, form a ring. Ul-C and U1-70K are shown in  shades of orange (U1-A is not visible in this model). The RNA is shown in  paddle form in gray. The yellow spheres represent the selenium atoms in  selenomethionine residues (Met with its S atom replaced by Se), which were  mutagenically inserted into the proteins to aid in solving the structure, (c) The  structure as viewed from above in Part a. The Zn 2+ ion bound to the zinc  finger motif of Ul-C is represented by a green sphere. In the crystal, the 5'  terminal portion of the Ul-snRNA forms a double-helical segment with a  neighboring Ul-snRNA (purple). In vivo, the 5' splice site of a pre-mRNA  would bind here. [Courtesy of Kiyoshi Nagai, MRC Laboratory of Molecular  Biology, Cambridge, U.K. PDBid 3CW1J    Gene Splicing Offers Evolutionary Advantages. Analysis of the large body  of known DNA sequences reveals that introns are rare in prokaryotic struc¬  tural genes, uncommon in lower eukaryotes such as yeast (which has a total  of 239 introns in its —6600 genes and, with two exceptions, only one intron  per polypeptide), and abundant in higher eukaryotes (the only known verte¬  brate structural genes lacking introns are those encoding histones and the an¬  tiviral proteins known as interferons). Pre-mRNA introns, as we have seen,  can be quite long and many genes contain large numbers of them.  Consequently, unexpressed sequences constitute —80% of a typical vertebrate  structural gene and >99% of a few of them.   The argument that introns are only molecular parasites (junk DNA)  seems untenable since it would then be difficult to rationalize why the evolu¬  tion of complex splicing machinery offered any selective advantage over the  elimination of the split genes. What then is the function of gene splicing?   Chapter 26 Transcription and RNA Processing Although, since its discovery, the significance of gene splicing has been de¬   1. Gene splicing is an agent for rapid protein evolution. Many eukary¬  otic proteins consist of modules that also occur in other proteins  (Section 5-4B). For example, SH2 and SH3 domains occur in many  of the proteins involved in signal transduction (Section 13-2B). It  therefore appears that the genes encoding these modular proteins arose by  the stepwise collection of exons that were assembled by (aberrant) recom¬  bination between their neighboring introns.   2. Through alternative splicing, which we discuss below, gene splicing  permits a single gene to encode several (sometimes many) proteins that  may have significantly different functions.   The expression of numerous cellular genes is modulated by the selection of alter¬  native splice sites. Thus, genes containing multiple exons may give rise to tran¬  scripts containing mutually exclusive exons. In effect, certain exons in one type  of cell may be introns in another. For example, a single rat gene encodes seven  tissue-specific variants of the muscle protein a-tropomyosin (Section 7-2A)  through the selection of alternative splice sites (Fig. 26-27).   Alternative splicing occurs in all multicellular organisms and is especially  prevalent in vertebrates. In fact, microarray-based comparisons of the cDNAs  obtained from various tissues indicate that —95% of human structural genes  are subject to at least one alternative splicing event. This helps explain the  discrepancy between the —23,000 genes identified in the human genome and  earlier estimates that it contains over 100,000 structural genes. The variations  in spliced mRNA sequences can take several different forms: Exons can be re¬  tained or skipped; introns may be excised or retained; and the positions of 5'   5TJT1-38 39-80 39-80 81-125 126-164 188 213 234 257 284 3' UT 284 3' UT   FIG. 26-27 Alternative splicing in the rat ct-tropomyosin  gene. Seven alternative splicing pathways give rise to cell-specific  a-tropomyosin variants. The thin kinked lines indicate the positions  occupied by the introns before they are spliced out to form the  mature mRNAs. Tissue-specific exons are indicated together with the  amino acid (aa) residues they encode: “constitutive” exons (those  expressed in all tissues) are green; those expressed only in smooth    muscle (SM) are brown; those expressed only in striated muscle  (STR) are purple; and those variably expressed are yellow. Note that  the smooth and striated muscle exons encoding amino acid residues  39 to 80 are mutually exclusive and, likewise, there are alternative 3'  untranslated (UT) exons. [After Breitbart, R.E., Andreadis, A., and  Nadal-Ginard, B., Annu. Rev Biochem. 56, 481 (1987).]   and 3' splice sites can be shifted to make exons longer or shorter. Alterations Section 3 Posttranscriptional Processing   in the transcriptional start site and/or the polyadenylation site can further con¬  tribute to the diversity of the mRNAs that are transcribed from a single gene.   A particularly striking example of alternative splicing occurs in the  Drosophila DSCAM protein, which functions in neuronal development. The  protein is encoded by 24 exons of which there are 12 mutually exclusive vari¬  ants of exon 4, 48 of exon 6, 33 of exon 9, and 2 of exon 17 (which are there¬  fore known as cassette exons) for a total of 38,016 possible variants of the  protein (compared to ~ 14,000 identified genes in the Drosophila genome).   Although it is unknown whether all possible DSCAM variants are produced,  experimental evidence suggests that the Dscam gene expresses many thousands  of them. [Dscam is a membrane-anchored cell-surface protein of the im¬  munoglobulin superfamily. The specific isoform expressed in a given neuron  binds to itself but rarely to other isoforms. This permits the neuron to distin¬  guish its own processes (axons and dendrites) from those of other neurons and  thereby plays an essential role in neural patterning. However, the precise iden¬  tity of a given isoform appears to be unimportant.] Clearly, the number of genes  in an organisms genome does not by itself provide an adequate assessment of its  protein diversity. Indeed, it has been estimated that, on average, each human  structural gene encodes three different proteins.   The types of changes that alternative splicing confers on expressed proteins  span the entire spectrum of protein properties and functions. Entire functional  domains or even single amino acid residues may be inserted into or deleted  from a protein, and the insertion of a Stop codon may truncate the expressed  polypeptide [a codon is a 3-nt sequence that specifies an amino acid in a  polypeptide; a Stop codon is a 3-nt sequence that instructs the ribosome to ter¬  minate polypeptide synthesis (Section 27-1C)]. Splice variations may, for ex¬  ample, control whether a protein is soluble or membrane bound, whether it is  phosphorylated by a specific kinase, the subcellular location to which it is tar¬  geted, whether an enzyme binds a particular allosteric effector, or the affinity  with which a receptor binds a ligand. Changes in an mRNA, particularly in its  noncoding regions, may also influence the rate at which it is transcribed and  its susceptibility to degradation. Since the selection of alternative splice sites is  both tissue- and developmental stage-specific, splice site choice must be tightly  regulated in both space and time. In fact, ~15% of human genetic diseases are  caused by point mutations that result in pre-mRNA splicing defects. Some of  these mutations delete functional splice sites, thereby activating nearby preex¬  isting cryptic splice sites. Others generate new splice sites that are used in¬  stead of the normal ones. In addition, tumor progression is correlated with  changes in levels of proteins implicated in alternative splice site selection.   How are alternative splice sites selected? One example of this process oc¬  curs in the transformer (tra) gene during sex determination in Drosophila. Exon  2 of tra pre-mRNA contains two alternative 3' splice sites, with the proximal  (close; to exon 1) site used in males and the distal (far) site used in females  (Fig. 26-28). The region between the two sites contains a Stop codon with  the sequence UAG. In males, the splicing factor U2-snRNP auxiliary factor   FIG. 26-28 Alternative splice site selection in the Drosophila  tra gene. Exons of the pre-mRNA are represented by colored  rectangles, and introns are shown as gray bands with those to be  excised flanked by kinked lines. UAG is a Stop codon. In females, the    protein SXL directs the spliceosome to the second of two possible  splice sites, to generate mRNA for a full-length TRA protein. [After a  drawing by Maniatis, T. and Tasic, B., Nature 418, 236 (2002).]   Chapter 26 Transcription and RNA Processing (U2AF) binds to the proximal 3' splice site to yield an mRNA containing the   premature Stop codon, which thereby directs the synthesis of a truncated and  hence nonfunctional TRA protein. However, in females, the proximal 3' splice  site is bound by the female-specific SXL protein, the product of the sex-lethal  (sxl) gene (which is only expressed in females), so as to block the binding of  U2AF, which then binds to the distal 3' splice site, thereby inducing the ex¬  pression of functional TRA protein. In females, the TRA protein directs the  splicing of mRNA from another sex-determination gene, leading to produc¬  tion of a repressor of male-specific genes.   Similar mechanisms for repressing or activating a splice site occur in ver¬  tebrates. Some of the proteins involved in splice site selection—both constitu¬  tive and alternative—include SR proteins and several members of the  heterogeneous nuclear ribonucleoprotein (hnRNP) family. SR proteins each  have one or more RNA recognition motifs (RRMs), which bind RNA se¬  quences, near their N-terminus and a distinctive C-terminal RS domain, which  contains numerous Ser-Arg (SR) repeats and which participates in  protein-protein interactions (U2AF and TRA both contain an SR domain, but  neither has an RRM and so are not considered to be SR proteins). SR pro¬  teins, when appropriately phosphorylated on their RS domains, specifically  bind to corresponding small (5-10 nt) exonic splicing enhancers (ESEs) via  their RRMs and thereby recruit the splicing machinery to the flanking 5' and  3' splice sites. The hnRNPs, which are highly abundant RNA-binding proteins,  lack RS domains and hence cannot recruit the splicing machinery. Instead, they  bind to their corresponding exonic and intronic splicing silencers (ESSs and  ISSs) so as to block the binding of the splicing machinery at the flanking splice  sites. Such interactions help the splicing machinery distinguish true exons  (which are relatively short) from the more numerous sequences within introns  (which are relatively long) that happen to exhibit the consensus splice-site se¬  quences. Indeed, extensive analysis of the sequences of numerous alternative  splice sites has revealed the existence of a “splicing code” that uses combina¬  tions of over 200 RNA features that are present in both introns and exons and  which are recognized by the foregoing regulators. Tissue-specific expression of  hnRNPs and SR proteins, together with the latter’s reversible phosphorylation,  undoubtedly contribute to the complex regulation of mRNA splicing.   mRNA May Be Edited. Additional posttranscriptional modifications of  eukaryotic mRNAs include the methylation of certain A residues. In a few  cases, mRNAs undergo RNA editing, which involves base changes, deletions,  or insertions. In the most extreme examples of this phenomenon, which oc¬  cur in the trypanosomes and related protozoa, several hundred U residues may  be added and removed to produce a translatable mRNA. Not surprisingly, the  mRNA editing machinery includes RNAs known as guide RNAs (gRNAs)  that base-pair with the immature mRNA to direct its alteration.   In a few mRNAs, specific bases are chemically altered, for example, by  enzyme-catalyzed deamination of C to produce U and/or deamination of A  to produce inosine (which is read as G by the ribosome), a process called sub¬  stitutional editing. For example, humans express two forms of apolipopro-  tein B (apoB): apoB-48, which is made only in the small intestine and  functions in chylomicrons to transport triacylglycerols from the intestine to  the liver and peripheral tissues; and apoB-100, which is made only in the liver  and functions with VLDL, IDL, and LDL to transport cholesterol from the  liver to the peripheral tissues (Table 20-1). ApoB-100 is an enormous 4536-  residue protein, whereas apoB-48 consists of apoB-100’s N-terminal 2152  residues and therefore lacks the C-terminal domain of apoB-100 that medi¬  ates LDL receptor binding. Although apoB-48 and apoB-100 are expressed  from the same gene, the mRNAs encoding the two proteins differ by a single  C —^ U change: The codon for Gin 2153 (CAA) in apoB-100 mRNA is, in  apoB-48 mRNA, a UAA Stop codon. The enzyme that catalyzes this conver¬  sion is site-specific cytidine deaminase.   Substitutional editing may contribute to protein diversity. For example, Section 3 Posttranscriptional Processing   Drosophila cacophony pre-mRNA, which encodes a voltage-gated Ca 2+ chan¬  nel subunit, contains 10 different substitutional editing sites and hence has  the potential of generating 2 10 ~ 1000 different isoforms in the absence of  alternative splicing. Substitutional editing can also generate alternative splice  sites. For example, the enzyme ADAR2 (ADAR for adenosine deaminase  acting on 7?NA) edits its own pre-mRNA by converting an intronic AA din¬  ucleotide to AI, which mimics the AG normally found at 3' splice sites (Fig.   26-24). The consequent new splice site adds 47 nucleotides near the 5' end  of the ADAR2 mRNA so as to generate a new translational initiation site. The  resulting ADAR2 isozyme is catalytically active but is produced in smaller  amounts than that from unedited transcripts, perhaps due to a less efficient  translational initiation site. Thus, ADAR2 appears to regulate its own rate of  expression.   Mature Eukaryotic mRNAs Are Actively Transported from the Nucleus to  the Cytoplasm. The translation of prokaryotic mRNAs is often initiated be¬  fore their synthesis is complete. This cannot occur in eukaryotes because the  transcription and posttranscriptional processing of eukaryotic mRNAs occur  in the nucleus but their translation takes place in the cytosol. Consequently,  mature mRNAs must be transported from the nucleus to the cytoplasm. This  is a highly selective process because mature mRNAs comprise only a small  fraction of the RNAs present in the nucleus, the remainder being pre-mRNAs,  excised introns (which are usually much larger than the exons from which they  were liberated), rRNAs, tRNAs, snRNAs, and a variety of RNAs that partic¬  ipate in the processing of rRNAs and tRNAs (Section 26-3C). Indeed, only  ~5% of the RNA that is synthesized ever leaves the nucleus.   How are mature mRNAs recognized and transported? As we have seen,  throughout their residency in the nucleus, pre-mRNAs are continually associ¬  ated with numerous proteins, including those that participate in synthesizing  their m 7 G caps and poly(A) tails, and in splicing out their introns. In addi¬  tion, the exon junction complex (EJC), which consists of four core proteins  and several transiently associating proteins, is deposited onto mRNA during  the splicing process at a site that is 20 to 24 nt upstream of the splice junc¬  tion without regard to its sequence. The population of proteins bound to an  mRNA changes as the mRNA is processed but some of the proteins, includ¬  ing SR proteins, hnRNPs, and EJCs, remain associated with mature mRNAs  in the nucleus. However, it appears that it is its entire collection of bound  proteins rather than any individual protein that serves to identify an mRNA  to the nuclear export machinery.   The eukaryotic nucleus (Fig. 1-8) is a double membrane-enveloped or¬  ganelle that in animals is penetrated by an average of —3000 pores. These are  formed by nuclear pore complexes (NPCs), which are massive (— 120,000 kD),  eightfold symmetric assemblies of —30 different proteins known as  nucleoporins. NPCs, which have inner diameters of —90 A (although this may  be expandable to as much as 260 A), allow the free diffusion of molecules of  up to —40 kD, but most macromolecules, including mRNAs in their complexes  with proteins, require an active transport process to pass through an NPC. Some  of the proteins associated with mature mRNAs bear nuclear export signals that  are recognized by an ATP-driven RNA helicase that in yeast is named Dbp5.   Dbp5 also binds to the NPC, which permits it to pull the mRNA out into the  cytosol while simultaneously stripping away many of its bound proteins. The  proteins are later recycled by returning them to the nucleus through the NPCs.   E. coli has three types of rRNAs, the 5S, 16S, and 23S rRNAs. These are  specified by seven operons, each of which contains one nearly identical copy  of each of the three rRNA genes. The polycistronic primary transcripts of these    FIG. 26-29 The posttranscriptional processing of E. coli  rRNA. The transcriptional map is shown approximately to scale. The  labeled arrows indicate the positions of the various nucleolytic cuts  and the nucleases that generate them. [After Apiron, D., Ghora, B.K.,    Plantz, G., Misra, T.K., and Gegenheimer, P., in Soil, D., Abelson, J.N.,  and Schimmel, P.R. (Eds.), Transfer RNA: Biological Aspects, p. 148,  Cold Spring Harbor Laboratory (1980).]    operons are >5500 nt long and contain, in addition to the rRNAs, transcripts  for as many as four tRNAs (Fig. 26-29). The steps in processing the primary  transcripts to mature rRNAs were elucidated with the aid of mutants defec¬  tive in one or more of the processing enzymes.   The initial processing, which yields products known as pre-rRNAs, com¬  mences while the primary transcript is still being synthesized. It consists of  specific endonucleolytic cleavages by RNase III, RNase P, RNase E, and  RNase F at the sites indicated in Fig. 26-29. The cleavage sites are probably  recognized on the basis of their secondary structures.   The 5' and 3' ends of the pre-rRNAs are trimmed away in secondary pro¬  cessing steps through the action of RNases D, Ml6, M23, and M5 to pro¬  duce the mature rRNAs. These final cleavages occur only after the pre-rRNAs  become associated with ribosomal proteins. During ribosomal assembly, spe¬  cific rRNA residues are methylated, which may help protect them from inap¬  propriate nuclease digestion.   snoRNAs Direct the Methylation of Eukaryotic rRNAs. The eukaryotic  genome typically has several hundred tandemly repeated copies of rRNA genes.  These genes are transcribed and processed in the nucleolus (Fig. 1-8), a small  dark-staining body in the nucleus that is not membrane enveloped. The pri¬  mary eukaryotic rRNA transcript is an ~7500-nt 45S RNA that contains the  18S, 5.8S, and 28S rRNAs separated by spacer sequences:    In the first stage of its processing, 45S RNA is specifically methylated at  numerous sites (106 in humans) that occur mostly in its rRNA sequences.  About 80% of these modifications yield O 2 -methylribose residues, and the re¬  mainder yield methylated bases such as Af 6 ,Af^-dimethyladenine and 2-  methylguanine. The subsequent cleavage and trimming of the 45S RNA  superficially resemble those of prokaryotic rRNAs. In fact, enzymes exhibiting  RNase III- and RNase P-like activities occur in eukaryotes. Eukaryotic ribo¬  somes contain four different rRNAs (Section 27-3). The fourth type, 5S    eukaryotic rRNA, is separately processed in a manner resembling that of tRNA  (Section 26-3D).   The methylation sites in yeast and vertebrate rRNAs generally occur in in¬  variant sequences, although the methylation sites do not appear to have a con¬  sensus structure that might be recognized by a single methyltransferase. How  then are the methylation sites targeted? Pre-rRNAs interact with mem¬  bers of a large family of small nucleolar RNAs (snoRNAs). Mammals  have —200 snoRNAs, most of which are encoded by the introns of  structural genes (and hence not all excised introns are discarded). The  snoRNAs, whose lengths vary from 70 to 100 nt, contain segments of  10 to 21 nt that are precisely complementary to segments of the mature  rRNAs that contain O 2 -methylation sites. The snoRNAs appear to di¬  rect a protein complex containing a methyltransferase to the methyla¬  tion site. Presumably, without the snoRNAs, the cell would need to  synthesize a different methyltransferase to recognize each methylation se¬  quence. Complexes of small RNAs and proteins also catalyze the con¬  version of certain rRNA uridine residues to pseudouridine (iji):   Some Eukaryotic rRNAs Are Self-Splicing. A few eukaryotic rRNA  genes contain introns. In fact, it was Thomas Cecils study of how these  introns are spliced out that led to the astonishing discovery, in 1982,  that RNA can act as an enzyme (a ribozyme; Section 24-2C). Cech  showed that when the isolated pre-rRNA of the ciliated protozoan  Tetrahymena thermophila is incubated with guanosine or a free guanine  nucleotide (GMP, GDP, or GTP), but in the absence of protein, its sin¬  gle 413-nt intron excises itself and splices together its flanking exons;  that is, the pre-rRNA is self-splicing. The three-step reaction sequence of  this process (Fig. 26-30) resembles that of mRNA splicing:   1. The 3'-OH group of the guanosine attacks the introns 5' end,  displacing the 3'-OH group of the 5' exon and thereby form¬  ing a new phosphodiester linkage with the 5' end of the intron.   2. The 3'-terminal OH group of the newly liberated 5' exon at¬  tacks the 5'-phosphate of the 3' exon to form a new phospho¬  diester bond, thereby splicing together the two exons and  displacing the intron.   3. The 3'-terminal OH group of the intron attacks a phosphate of  the nucleotide 15 residues from the introns end, displacing the  5' terminal fragment and yielding the 3' terminal fragment in  cyclic form.    FIG. 26-30 The sequence of reactions in the self-splicing of Tetrahymena  pre-rRNA. (1) The 3'-OH group of a guanine nucleotide attacks the intron’s  5'-terminal phosphate to form a phosphodiester bond, displacing the 5' exon. (2) The  newly generated 3'-0H group of the 5' exon attacks the 5'-terminal phosphate of  the 3' exon, thereby splicing the two exons and displacing the intron. (3) The 3'-OH  group of the intron attacks the phosphate of the nucleotide that is 15 residues from  the 5' end so as to cyclize the intron and displace its 5'-terminal fragment. Throughout  this process, the RNA maintains a folded, internally hydrogen-bonded conformation that  permits the precise excision of the intron.   Chapter 26 Transcription and RNA Processing The self-splicing process consists of a series of transesterifications and there¬   Self-splicing RNAs that react as shown in Fig. 26-30 are known as group I  introns and occur in the nuclei, mitochondria, and chloroplasts of diverse  eukaryotes (but not vertebrates), and in some bacteria. Group II introns,  which occur in the mitochondria and chloroplasts of fungi and plants, em¬  ploy an internal A residue (instead of an external G) to form a lariat interme¬  diate. The chemical similarities of the pre-mRNA and group II intron splicing  reactions therefore suggest that spliceosomes are ribozymal systems whose RNA  components have evolved from primordial self-splicing RNAs and that their pro¬  tein components serve mainly to fine-tune ribozymal structure and function. This,  of course, is consistent with the hypothesis that RNAs were the original bio¬  logical catalysts in precellular times (the RNA world; Box 24-3).   Group I introns are the most abundant self-splicing introns with >2000 such  sequences known. The sequence of the 413-nt Tetrahymena group I intron, to¬  gether with phylogenetic comparisons, indicates that it contains nine double¬  helical segments that are designated PI through P9 (Fig. 26-31 tf; P for base  paired segment). Such analysis further indicates that the conserved catalytic  core of group I introns consists of sets of coaxially stacked helices interspersed  with internal loops that are organized into two domains, the P4-P5-P6 do¬  main (also called P4-P6) and the P3-P7-P8-P9 domain (also called P3-P9).   Cech designed a 247-nt RNA (Fig. 26-31/2) that encompasses both the  P4-P6 and P3-P9 domains of the Tetrahymena group I intron (it lacks the Pl-  P2 domain and the attached exons), with the addition of a 3' G (o>G), which  functions as an internal guanosine nucleophile. This RNA is catalytically ac¬  tive; it binds the P1-P2 domain via tertiary interactions and, with the assis¬  tance of (uG, cleaves PI in a manner similar to the intact intron.   The X-ray structure of this RNA (Fig. 26-31 b,c) reveals that it is largely  composed of three coaxially stacked sets of A-RNA-like helices with P4-P6  consisting of two pseudocontinuous and straight parallel helices connected  by a sharp bend and P3-P9 consisting of a curved helix that wraps around  one side of P4-P6 through extensive interdomain interactions that form the  ribozyme s active site. Of particular note are its so-called A-rich bulge, a 7-  nt sequence about halfway along the short arm of the U-shaped P4-P6, and  the 6-nt sequence at the tip of the short arm of the U, whose central GAAA  assumes a characteristic conformation known as a tetraloop. In both of these  substructures, the bases are splayed outward so as to stack on each other and  to associate in the minor groove of specific segments of the long arm of the  U via hydrogen bonding interactions involving ribose residues as well as  bases. In many such interactions, the close packing of phosphate groups is  mediated by hydrated Mg 2+ ions. Throughout this structure, the defining  characteristic of RNA, its 2'-OH group, is both a donor and an acceptor of  hydrogen bonds to phosphates, bases, and other 2'-OH groups. Interestingly,  although this overall fold is highly conserved among group I introns, their  sequences are poorly conserved with the exception of a few crucial active site  residues.   Divalent metal ions, usually Mg 2+ , are often required for both the struc¬  tural stability and the catalytic activity of ribozymes. In particular, an Mg 2+  ion is in contact with the 2'-OH group of cnG as well as being liganded by  the phosphate groups of three surrounding nucleotides. This both orients the  ribose group of o>G and nucleophilically activates its 3'-OH group. A second  Mg 2+ ion, which accompanies the phosphate group of the RNA substrate,  also participates in the catalytic reaction. Note that two Mg 2+ ions similarly  participate in the phosphoryl-transfer reactions catalyzed by protein enzymes  such as DNA polymerase (Section 25-2A).    FIG. 26-31 The group I intron from Tetrahymena thermophila. (a) The   secondary structure of the 414-nt ribozyme. Its P4-P6 and P3-P9 domains are  shaded in blue and green, respectively, with the catalytically active 3' wG residue  shaded in yellow, the base triples of the P7 domain shaded in red, and the A-rich  bulge, the tetraloop, and the tetraloop receptor on the P4-P6 domain shaded in  orange, pink, and cyan, respectively. Watson-Crick and non-Watson-Crick base  pairing interactions are represented by short horizontal lines and small filled circles,  whereas interdomain interactions are indicated by magenta lines. Every tenth  residue is marked by an outwardly pointing dash. The positions of five residues that  have been mutated to stabilize the ribozyme structure are circled and those of the  seven mutations that facilitated crystallization are bracketed (this mutant form  retains its catalytic activity), (b) The X-ray structure of the ribozyme, drawn in  cartoon form with its bases shown as paddles, and colored as in Part a. The  inferred positions of Mg 2+ ions are represented by orange spheres, (c) As in Part  b but rotated 140° about the vertical axis to better show the A-rich bulge and the  interaction between the tetraloop and the tetraloop receptor. [Part a modified from  a drawing by and Parts b and c based on an X-ray structure by Thomas Cech,  University of Colorado. PDBid 1X8W.] O See Interactive Exercise 52.   D Transfer RNAs Are Processed by Nucleotide Removal, Addition,  and Modification    A tRNA molecule, as we discussed in Secdon 24-2C, consists of around 76 nu¬  cleotides, many of which are chemically modified, that assume a cloverleaf-shaped   FIG. 26-32 Schematic diagram of the  tRNA cloverleaf secondary structure. Each  red dot indicates a base pair in the hydrogen-  bonded stems. The positions of the anticodon  (the 3-nt sequence that binds to mRNA during  translation) and the 3'-terminal —CCA are  indicated.    secondary structure with four base-paired stems (Fig. 26-32). The E. coli  chromosome contains ~60 tRNA genes. Some of them are components of  rRNA operons; the others are distributed, often in clusters, throughout the  chromosome. The primary tRNA transcripts, which contain as many as five  identical tRNA species, have extra nucleotides at the 3' and 5' ends of each  tRNA sequence. The excision and trimming of these tRNA sequences resem¬  ble E. coli rRNA processing (Fig. 26-29) in that the two processes employ  some of the same nucleases.   E. coli RNase P, which processes rRNA and generates the 5' ends of  tRNAs, is a particularly interesting enzyme because it is a ribozyme with a cat-  alytically essential RNA and one protein subunit. At first, the RNA was be¬  lieved to recognize the substrate through base pairing and thereby guide the  protein subunit, which was presumed to be the nuclease, to the cleavage site  on the substrate pre-tRNA. However, Sidney Altman showed that the RNA  component of RNase P is, in fact, the enzyme's catalytic subunit by demonstrating  that free RNase P RNA catalyzes the cleavage of substrate RNA at high salt con¬  centrations. The RNase P protein, which is basic, increases the reaction rate by  two to three orders of magnitude by stabilizing the P RNA structure, binding  the 5' “leader” sequence of the pre-tRNA, and assisting in product release.  RNase P activity also occurs in eukaryotes, although the enzyme includes 9 or  10 protein subunits. RNase P, unlike the self-splicing introns, is a true enzyme  capable of multiple turnovers. Indeed, RNase P mediates one of the two ri-  bozymal activities that occur in all cellular life, the other being ribosomally  catalyzed peptide bond formation (Section 27-4B).   The RNase P holoenzyme from Thermotoga maritima contains a 117-residue  (14-kD) protein and a 338-residue (110-kD) RNA whose secondary structure in¬  cludes numerous base-paired stems. The X-ray structure of this RNase P in com-  plex with tRNA phe [Fig. 26-33; tRNA phe conveys Phe to the ribosome (Section  27-4)], a ribozyme-product complex, was determined by Alfonso Mondragon. It  reveals that RNase P has an overall compact structure typical of protein enzymes  and binds the tRNA Phe through shape complementarity, base stacking, base pair¬  ing, and specific RNA-RNA contacts. The enzymes active site includes a uni¬  versally conserved U residue and two metal ions, probably Mg 2+ , that are  postulated to participate directly in catalysis. A portion of the RNA is present in  all known RNase P s and hence is known as the universal minimum consensus  structure. This structure was presumably present in the primordial RNase P.    FIG. 26-33 X-Ray structure of T.  maritima RNase P in complex with tRNA Phe .   The RNAs are drawn in ladder form with the  tRNA Phe pink and the P RNA colored in rainbow  order from its 5' end (blue) to its 3' end (red).  The protein component of RNase P, which is  positioned to bind a substrate pre-tRNA’s 5'  extension, is drawn in ribbon form colored in  rainbow order from its N-terminus (blue) to its  C-terminus (red). The tRNA Phe lacks the 5'  extension present in pre-tRNAs and hence this  is a ribozyme-product complex. Gray segments  of the RNAs represent additional sequences that  were required to crystallize the complex. [Based  on an X-ray structure by Alfonso Mondragon,  Northwestern University. PDBid 3Q1R.]   A 14-nt intervening sequence  (red) and a 19-nt 5'-terminal  sequence (green) are excised  from the primary transcript, a  —CCA (blue) is appended to  its 3' end, and several of the  bases are modified to form  the mature tRNA  (m 2 G, A/ 2 -methylguanosine;   Gm, 2'-methylguanosine;  m|G, A/ 2 ,A/ 2 -dimethylguanosine;  M* pseudouridine;  i 6 A, A/ 6 -isopentenyladenosine;  m 5 C, 5-methylcytosine;  rrdA, 1-methyladenosine;  see Fig. 27-4).    Many Eukaryotic Pre-tRNAs Have Introns. Eukaryotic genomes each contain  from several hundred to several thousand tRNA genes. Many eukaryotic pri¬  mary tRNA transcripts, for example, that of yeast tRNA Tyr (Fig. 26-34), con¬  tain a small intron as well as extra nucleotides at their 5' and 3' ends. tRNA  processing therefore includes nucleolytic removal of the extra nucleotides. The  three nucleotides, CCA, at the 3' termini of all tRNAs, the sites at which  amino acids are attached (Section 27-2B), are lacking in the immature tRNA  transcripts. This trinucleotide is appended by the enzyme CCA-adding poly¬  merase, which sequentially adds two C’s and an A to tRNA using CTP and  ATP as substrates. This enzyme also occurs in prokaryotes, although, at least  in E. coli, all tRNA genes encode a —CCA terminus. The E. coli CCA-adding  polymerase is therefore likely to function in the repair of degraded tRNAs.    What is the advantage of starting RNA  processing before transcription is  complete?   Discuss the advantages, in terms of pro¬  tein structure and evolution, that result  from alternative mRNA splicing.   What does the existence of self-splicing  RNAs reveal about the evolution of  spliceosomes?   Describe the modifications that occur  during processing of prokaryotic and  eukaryotic rRNAs and tRNAs.   • RNA polymerase synthesizes a polynucleotide chain from ribonu- * The a factor of E. coli RNA polymerase holoenzyme recognizes  cleoside triphosphates using a single strand (the antisense or non- and binds to a promoter sequence to position the enzyme to ini-   • RNA synthesis requires the formation of an open complex. The  transcription bubble travels along the DNA as the RNA chain is  elongated by the processive activity of RNA polymerase.   • In E. coli, RNA synthesis is terminated in response to specific sec¬  ondary structural elements in the transcript and may require the  action of Rho factor.   • Eukaryotes contain three nuclear RNA polymerases that synthe¬  size its different types of RNAs.   • Eukaryotic promoters are diverse: They vary in position relative to  the transcription start site and may consist of multiple sequences.  Enhancers form the binding sites for regulatory proteins that func¬  tion as activators of transcription.   • Basal transcription of eukaryotic genes requires six general tran¬  scription factors, including TBP, that form a preinitiation complex    with RNA polymerase at the promoter. Some of these proteins re¬  main with the polymerase when its C-terminal domain (CTD) is  phosphorylated and shifts to elongation.   • The primary transcripts of most eukaryotic structural genes are  posttranscriptionally modified by the addition of a 5' cap and a  y poly(A) tail. Pre-mRNAs that contain introns undergo splicing,  in which the introns are excised and the flanking exons are joined  together via two transesterification reactions mediated by an  snRNA-containing spliceosome.   • The processing of pre-rRNAs includes nucleolytic cleavage and  snoRNA-assisted methylation. Some eukaryotic rRNA transcripts  undergo splicing catalyzed by the intron itself.   • tRNA transcripts may be processed by the addition, removal, and  modification of nucleotides.   (c) PNPase includes a binding site for long ribonucleotides, which  may promote the enzymes processivity. Why would this be an  advantage for the primary activity of PNPase in vivo}   5' C A AG GTA AC AC T T TAG AG C GG CGCGTC AT T TGATATGATGCGC CCCGCTTCCC GAT A 3*  3' GTTGCAT TGTG AA ATGTCG C C GCGC AGT AA AC TAT AC TAC GCG GGGC G A AG GGC TAT 5*   (a) The enzyme catalyzes a phosphorolysis reaction, as does glyco¬  gen phosphorylase (Section 16-1), rather than hydrolysis.  Write an equation for the mRNA phosphorolysis reaction.   (b) In vitro , PNPase also catalyzes the reverse of the phosphoroly¬  sis reaction. What does this reaction accomplish and how does  it differ from the reaction carried out by RNA polymerase?    4. Design a six-residue nucleic acid probe that would hybridize with  the greatest number of E. coli gene promoters.   5. Why does promoter efficiency tend to decrease with the number  of G • C base pairs in the —10 region of a prokaryotic gene?   6. Explain why inserting 3 bp of DNA at the — 50 position of a eu¬  karyotic gene decreases the rate of RNA polymerase II transcription  initiation to a greater extent than inserting 10 bp at the same site.   7. Many bacterial genes with related functions are arranged in oper-  ons, sets of contiguous genes that are under the control of a sin¬  gle promoter and are transcribed together, (a) What is the  advantage of this arrangement? (b) How might eukaryotic cells,  which do not contain operons, ensure the simultaneous transcrip¬  tion of different genes?   and ~82 different proteins. Why does a cell contain many more  copies of the rRNA genes than the ribosomal protein genes?   9. Replication of many RNA viruses depends on RNA polymerase.  The antiviral drug ribavirin does not inhibit the polymerase but  instead increases its error rate. Explain how this affects the virus.   10. Collisions between DNA polymerase and a slower-moving RNA  polymerase using the same template strand may be responsible for  the observation that leading strand synthesis in bacteria is discon¬  tinuous in vivo. Draw a diagram to show how DNA synthesis that  has been halted after the DNA polymerase-RNA polymerase col¬  lision can resume using the RNA transcript as a primer.   11. Predict the effect of the antibiotic bicyclomycin, an antibiotic  that inhibits Rho, on gene expression in E. coli.   12. Certain E. coli bacteriophages encode a protein called Q, which  binds to RNAP shortly after transcription initiation, before a 70  is released. Q increases the rate of transcription and renders  RNAP resistant to Rho-dependent termination, (a) Explain how  Q could enhance expression of bacteriophage genes, (b) Explain  why the effect of Q depends on its ability to interact with spe¬  cific DNA sequences, not just RNAP.   13. The RNA polymerase from bacteriophage T7 differs structurally  from prokaryotic and eukaryotic RNAPs and is extremely specific  for its own promoter. Why do these properties make T7 RNAP  useful in experiments with recombinant DNA?   14. TFIIB appears to interact with proteins that bind to sequences  at the y end of genes. Explain how this interaction could en¬  hance transcription of a gene.   13. Design an oligonucleotide-based affinity chromatography system  for purifying mature mRNAs from eukaryotic cell lysates.   16. Human cells contain a protein that binds to the 5' triphosphate  groups of RNA. Explain why this protein would be part of the  defense against viral infection.   17. A eukaryotic cell carrying out transcription and RNA processing  is incubated with 32 P-labeled ATP. Where will the radioactive  isotope appear in mature mRNA if the ATP is labeled at the  (a) a position, (b) (3 position, and (c) y position?   18. Compare DNA polymerase, RNA polymerase, poly(A) poly¬  merase, and CCA-adding polymerase with respect to requirement  for a primer, template, and substrates.   19. Explain why the active site of poly(A) polymerase is much nar¬  rower than that of DNA and RNA polymerases.   20. Explain why the O 2 -methylation of ribose residues protects  rRNA from RNases.   21. Would you expect spliceosome-catalyzed intron removal to be re¬  versible in a highly purified in vitro system and in vivo? Explain.   22. Introns in eukaryotic protein-coding genes may be quite large,  but almost none are smaller than about 63 bp. What is the rea¬  son for this minimum intron size?   23. Infection with certain viruses inhibits snRNA processing in eu¬  karyotic cells. Explain why this favors the expression of viral  genes in the host cell.   24. Draw a diagram, including exons and introns, of a gene that encodes  both membrane-bound and soluble forms of a protein. Explain how  one gene can code for two different forms of a protein.    A so-called gene trap vector is  engineered to include a reporter gene flanked by a splice accep¬  tor sequence and a polyadenylation signal. How is this vector used  to reveal patterns of gene expression in mammalian cells? Why  does the gene trapping method depend on the size of a gene’s in¬  trons? How is the trapped gene identified?   Cramer, P. and Arnold, A., Proteins: how RNA polymerases work, Curr.  Opin. Struct. Biol. 19, 680-682 (2009). [The editorial overview of  a series of authoritative reviews on numerous aspects of RNA poly¬  merases. See, in particular, the reviews beginning on pages 691, 701,  708, and 732.]   Kadonaga, K.T., The RNA polymerase II core promoter, Annu. Rev.  Biochem. 72, 449-479 (2003).   Marakami, K.S. and Darst, S.A., Bacterial RNA polymerases: the whole  story. Curr. Opin. Struct. Biol. 13, 31-39 (2003).   Nudler, E., RNA polymerase active center: the molecular engine of tran¬  scription, Annu. Rev. Biochem. 78, 335-361 (2009).   Shilatifard, A., Conway, R.C., and Conway, J.W., The RNA polymerase  II elongation complex, Annu. Rev. Biochem. 72, 693-715 (2003).   Skordalakes, E. and Berger, J.M., Structural insights into RNA-dependent  ring closure and ATPase activation by the Rho termination factor,  Cell 127, 553-564 (2006).   Wang, D., Bushnell, D.A., Westover, K.D., Kaplan, C.D., and Kornberg,  R.D., Structural basis of transcription: Role of the trigger loop in  substrate specificity and catalysis, Cell 127, 941-954 (2006).   Black, D.L., Mechanism of alternative pre-messenger RNA splicing,  Annu. Rev. Biochem. 72, 291-336 (2003).   Blencoe, B.J., Alternative splicing: New insights from global analysis, Cell  126, 37-47 (2006).    Darnell, J., RNA. Life’s Indispensable Molecule, Cold Spring Harbor  Laboratory Press (2011).   Evans, D., Marquez, S.M., and Pace, N.R., RNase P: interface in the  RNA and protein worlds, Trends Biochem. Sci. 31, 333-341 (2006).   Krummel, D.A.P., Oubridge, C., Leung, A.K.W., Li, J., and Nagai, K.,  Crystal structure of human spliceosomal U1 snRNP at 5.5 A reso¬  lution, Nature 458, 475-480 (2009).   Maniatis, T. and Reed, R., An extensive network of coupling among gene  expression machines, Nature 416, 499-506 (2002). [Describes why  efficient gene expression requires coupling between transcription and  RNA processing.]   Nilsen, T.W. and Graveley, B.R., Expansion of the eukaryotic proteome  by alternative splicing, Nature 463, 457-463 (2010). [A review of  different types of alternative splicing and its regulatory mechanisms.]   Nishikura, K., Functions and regulation of RNA editing by ADAR deam¬  inases, Annu. Rev. Biochem. 79, 321-349 (2010).   Proudfoot, N.J., Furger, A., and Dye, M.J., Integrating mRNA process¬  ing with transcription, Cell 108, 501-512 (2002).   Reiter, N.J., Osterman, A., Torres-Larios, A., Swinger, K.K., Pan, T., and  Mondragon, A., Structure of a bacterial ribonuclease P holoenzyme  in complex with tRNA, Nature 468, 784-789 (2010).   Stark, H. and Luhrmann, R., Cryo-electron microscopy of spliceosomal  components, Annu. Rev. Biophys. Biomol. Struct. 35, 435-457 (2006).   Vincens, Q. and Cech, T.R., Atomic level architecture of group I introns  revealed, Trends Biochem. Sci. 31, 41-51 (2005).   Warf, M.B. and Berglund, J.A., Role of RNA structure in regulating pre-  mRNA splicing, Trends Biochem. Sci. 35, 169-178 (2010).   A Codons Are Triplets That Are Read Sequentially  B The Genetic Code Was Systematically  Deciphered   A All tRNAs Have a Similar Structure  B Aminoacyl-tRNA Synthetases Attach Amino  Acids to tRNAs   Several types of organisms, including Acetabularia (an enormous single-celled alga  whose cap is on the order of 1 cm in diameter), use variations of the standard  genetic code, prompting biologists to devise new explanations for how such a  fundamental aspect of molecular biology can permit evolutionary change.   B The Ribosome Decodes the mRNA, Catalyzes  Peptide Bond Formation, Then Moves to the  Next Codon    How is the genetic information encoded in DNA decoded? In the preceding  chapter, we saw how the base sequence of DNA is transcribed into that of  RNA. In this chapter, we consider the remainder of the decoding process by  examining how the base sequences of RNAs are translated into the amino acid  sequences of proteins. This second part of the central dogma of molecular bi¬  ology (DNA —» RNA —» protein) shares a number of features with both DNA  replication and transcription, the other major events of nucleic acid metabo¬  lism. First, all three processes are carried out by large, complicated protein-  containing macromolecular machines whose proper functioning depends on a  variety of specific and nonspecific protein—nucleic acid interactions. Accessory  factors are also required for the initiation, elongation, and termination phases  of these processes. Furthermore, translation, like replication and transcription,  must be executed with accuracy. Although translation involves base pairing be¬  tween complementary nucleotides, it is amino acids, rather than nucleotides,  that are ultimately joined to generate a polymeric product. This process, like  replication and transcription, is endergonic and must be driven by the cleav¬  age of “high-energy” phosphoanhydride bonds.   An understanding of translation requires not only a knowledge of the  macromolecules that participate in polypeptide synthesis, but an appreciation  for the mechanisms that produce a chain of linked amino acids in the exact  order specified by mRNA. Accordingly, we begin this chapter by examining  the genetic code, the correspondence between nucleic acid sequences and  polypeptide sequences. We then consider in turn the structures and properties  of tRNAs and ribosomes. Next, we examine how the translation machinery  operates as a coordinated whole. Finally, we take a brief look at some post¬  translational steps of protein synthesis.   • The genetic code is based on sets of three-nucleotide codons that are read  sequentially.   • Each codon represents one amino acid or a Stop signal.   One of the most fascinating puzzles in molecular biology is how a sequence  of nucleotides composed of only four types of residues can specify the sequence  of up to 20 types of amino acids in a polypeptide chain. Clearly a one-to-  one correspondence between nucleotides and amino acids is not possible. A  group of several bases, termed a codon , is necessary to specify a single amino acid.  A triplet code, that is, one with 3 bases per codon, is more than sufficient to  specify all the amino acids since there are 4 3 = 64 different triplets of bases.  A doublet code with 2 bases per codon (4 2 = 16 possible doublets) would be  inadequate. The triplet code allows many amino acids to be specified by more  than one codon. Such a code, in a term borrowed from mathematics, is said  to be degenerate.   How does the polypeptide-synthesizing apparatus group DNAs continu¬  ous sequences of bases into codons? For example, the code might be overlap¬  ping; that is, in the sequence   ABC might code for one amino acid, BCD for a second, CDE for a third,  etc. Alternatively, the code might be nonoverlapping, so that ABC specifies  one amino acid, DEF a second, GHI a third, etc. In fact, the genetic code is  a nonoverlapping, degenerate, triplet code. A number of elegant experiments,  some of which are outlined below, revealed the workings of the genetic code.   In genetic experiments on bacteriophage T4, Francis Crick and Sydney  Brenner found that a mutation that resulted in the deletion of a nucleotide  could abolish the function of a specific gene. However, a second mutation, in  which a nucleotide was inserted at a different but nearby position, could re¬  store gene function. The two mutations are said to be suppressors of one an¬  other; that is, they cancel each others mutant properties. On the basis of these  experiments, Crick and Brenner concluded that the genetic code is read in a se¬  quential manner starting from a fixed point in the gene. The insertion or dele¬  tion of a nucleotide shifts the reading frame (grouping) in which succeeding  nucleotides are read as codons. Insertions or deletions of nucleotides are there¬  fore known as frameshift mutations.   In further experiments, Crick and Brenner found that whereas two closely  spaced deletions or two closely spaced insertions could not suppress each other  (restore gene function), three closely spaced deletions or insertions could do  so. These observations clearly established that the genetic code is a triplet code.   The foregoing principles are illustrated by the following analogy. Consider  a sentence (gene) in which the words (codons) each consist of three letters  (bases):   Here the spaces separating the words have no physical significance; they are  present only to indicate the reading frame. The deletion of the fourth letter,  which shifts the reading frame, changes the sentence to   so that all words past the point of deletion are unintelligible (specify the wrong    amino acids). An insertion of any letter, however, say an X in the ninth  position,   restores the original reading frame. Consequently, only the words between the  two changes (mutations) are altered. As in this example, such a sentence might  still be intelligible (the gene could still specify a functional protein), particu¬  larly if the changes are close together. Two deletions or two insertions, no mat¬  ter how close together, would not suppress each other but just shift the reading  frame. However, three insertions, say X, Y, and Z in the fifth, eighth, and  twelfth positions, respectively, would change the sentence to   which, after the third insertion, restores the original reading frame. The same  would be true of three deletions. As before, if all three changes were close to¬  gether, the sentence might still retain its meaning. Like this textual analogy,  the genetic code has no internal punctuation to indicate the reading frame; in¬  stead\ the nucleotide sequence is read sequentially, triplet by triplet.   Since any nucleotide sequence may have three reading frames, it is possi¬  ble, at least in principle, for a polynucleotide to encode two or even three dif¬  ferent polypeptides. In fact, some single-stranded DNA bacteriophages (which  presumably must make maximal use of their small complement of DNA) con¬  tain completely overlapping genes that have different reading frames. A simi¬  lar form of coding economy is exhibited by bacteria, in which the ribosomal  initiation sequence of one gene in a polycistronic mRNA often overlaps the  end of the preceding gene.   In order to understand how the genetic code dictionary was elucidated, we  must first review how proteins are synthesized. An mRNA does not directly  recognize amino acids. Rather, it specifically binds molecules oftRNA that each  carry a corresponding amino acid (Fig. 27-1). Each tRNA contains a trinu¬  cleotide sequence, its anticodon, that is complementary to an mRNA codon  specifying the tRNAs amino acid. During translation, amino acids carried by    FIG. 27-1 Transfer RNA in its “cloverleaf”  form. Its covalently linked amino acid residue  is at the top, and its anticodon (X-Y-Z, a  trinucleotide segment that base-pairs with the  complementary mRNA codon during translation)  is at the bottom.   tRNAs are joined together according to the order in which the tRNA anti- Section 1 The Genetic Code   codons bind to the mRNA codons at the ribosome (Fig. 3-13).   The genetic code could, in principle, be determined by simply compar¬  ing the base sequence of an mRNA with the amino acid sequence of the  polypeptide it specifies. In the 1960s, however, techniques for isolating and  sequencing mRNAs had not yet been developed. Moreover, techniques for syn¬  thesizing RNAs were quite rudimentary. They utilized polynucleotide phos-  phorylase, an enzyme from Azotobacter vinelandii that links together  nucleotides without the use of a template.    Thus, the NDPs are linked together at random so that the base composition  of the product RNA reflects that of the reactant NDP mixture. The elucida¬  tion of the genetic code therefore proved to be a difficult task, even with the  development of cell-free translation systems.   E. coli cells that have been gently broken open and centrifuged to remove  cell walls and membranes yield an extract containing DNA, mRNA, ribo¬  somes, enzymes, and other cell constituents necessary for protein synthesis.  When fortified with ATP, GTP, and amino acids, this system synthesizes small  amounts of protein. A cell-free translation system, of course, produces pro¬  teins specified by the cells DNA. Adding DNase halts protein synthesis after  a few minutes because the system can no longer synthesize mRNA, and the  mRNA originally present is rapidly degraded. At this point, purified mRNA  or synthetic mRNA can be added to the system and the resulting polypeptide  products can be subsequently recovered.   In 1961, Marshall Nirenberg and Heinrich Matthaei added the synthetic  polyribonucleotide poly(U) to a cell-free translation system containing isotopi-  cally labeled amino acids and recovered a labeled poly(Phe) polypeptide. They  concluded that UUU must be the codon that specifies Phe. Similar experiments  with poly(A) and poly(C) yielded poly(Lys) and poly (Pro), respectively, thereby  identifying AAA as a codon for Lys and CCC as a codon for Pro.   In another series of experiments, different trinucleotides were tested for their  ability to promote tRNA binding to ribosomes. The ribosomes, with their bound  tRNAs, are retained by a nitrocellulose filter, but free tRNAs are not. The bound  tRNA can be identified by the radioactive amino acid attached to it. This sim¬  ple binding assay revealed that, for instance, UUU stimulates the ribosomal bind¬  ing of only Phe tRNA. Likewise, UUG, UGU, and GUU stimulate the binding  of Leu, Cys, and Val tRNAs, respectively. Hence UUG, UGU, and GUU must  be codons that specify Leu, Cys, and Val, respectively. In this way, the amino  acids specified by some 50 codons were identified. For the remaining codons, the  binding assay was either negative (no tRNA bound) or ambiguous.   The genetic code dictionary was completed and previous results confirmed  through H. Gobind Khoranas chemical synthesis of polynucleotides with spec¬  ified repeating sequences (at the time, an extremely laborious process). In a  cell-free translation system, UCUCUCUC * • • , for example, is read   so that it specifies a polypeptide chain of two alternating amino acid residues.  This particular mRNA stimulated the production of   Alternating sequences of three nucleotides, such as poly(UAC), specify  three different homopolypeptides because ribosomes may initiate polypeptide  synthesis on these synthetic mRNAs in any of the three possible reading frames  (Fig. 27-2). Analyses of the polypeptides specified by various alternating se¬  quences of two and three nucleotides confirmed the identify of many codons  and filled out missing portions of the genetic code.    FIG. 27-2 The three potential reading  frames of an mRNA. Each reading frame  would yield a different polypeptide.   The genetic code, presented in Table 27-1, has several remarkable features:   1. The code is highly degenerate. Three amino acids—Arg, Leu, and Ser—are  each specified by six different codons, and most of the rest are specified  by either four, three, or two codons. Only Met and Trp, two of the least  common amino acids in proteins (Table 4-1), are represented by a single  codon. Codons that specify the same amino acid are termed synonyms.   2. The arrangement of the code table is nonrandom. Most synonyms occupy  the same box in Table 27-1; that is, they differ only in their third nu¬  cleotide. XYU and XYC always specify the same amino acid; XYA and  XYG do so in all but two cases. Moreover, changes in the first codon  position tend to specify similar (if not the same) amino acids, whereas  codons with second position pyrimidines encode mostly hydrophobic  amino acids (gold in Table 27-1), and those with second position  purines encode mostly polar amino acids (blue, red, and purple in  Table 27-1). These observations suggest a nonrandom origin of the ge¬  netic code and indicate that the code evolved so as to minimize the  deleterious effects of mutations (see Box 27-1).   3. UAG, UAA , and UGA are Stop codons . These three codons (also  known as nonsense codons) do not specify amino acids but signal the  ribosome to terminate polypeptide chain elongation.   4. AUG and GUG are Start codons. The codons AUG and, less fre¬  quently, GUG specify the starting point for polypeptide chain synthesis.    Because of the degeneracy of the genetic code, a point mutation  in the third position of a codon seldom alters the specified amino  acid. For example, a GUU —» GUA transversion still codes for Val  and is therefore said to be phenotypically silent. Other point muta¬  tions, even at the first or second codon positions, producing AUU  (lie) or GCU (Ala), for instance, result in the substitution of a  chemically similar amino acid and may have minimal impact on the  encoded protein’s overall structure or function. This built-in protec¬  tion against mutation may be more than accidental.   In the 1960s, the perceived universality of the genetic code  led Francis Crick to propose the “frozen accident” theory, which  holds that codons were allocated to different amino acids entirely  by chance. Once assigned, the meaning of a codon could not  change because of the high probability of disrupting the structure  of the encoded protein. Thus, once established, the genetic code  was thought to have ceased evolving.   However, the distribution of codons as presented in Table 27-1  suggests an alternative evolutionary history of the genetic code, in  which a few simple codons corresponding to a handful of amino  acids gradually became more complex. One scenario begins with  an RNA-based world containing only A and U nucleotides. Uracil  is almost certainly a primordial base since the pyrimidine biosyn¬  thetic pathway yields uracil nucleotides before cytosine or thymine  nucleotides (Section 23-2). Adenine would have been required as  uracil’s complement.   Assuming that a triplet-based genetic code was established at  the outset (and it is difficult to envision how any other arrange¬  ment could have given rise to the present-day triplet code), the  two bases could have coded for 2 3 = 8 amino acids. In fact, the    contemporary genetic code assigns these all-U/A codons to six  amino acids and a stop signal:   The AUA codon may well have originally specified the initiating Met  (now encoded by AUG), bringing the total to seven amino acids.   When G and C appeared in evolving life-forms, these  nucleotides were incorporated into RNA. Codons containing three  or four types of bases could specify additional amino acids, but  because of selective pressure against introducing disruptive muta¬  tions into proteins, the level of codon redundancy increased. An  inspection of Table 27-1 shows that triplet codons made entirely  of G and C specify only four different amino acids, which is half  the theoretical maximum of eight:   GGG, GGC = Gly  GCG, GCC = Ala  CGG, CGC = Arg  CCC, CCG = Pro   This nonrandom allocation of codons to amino acids argues  against a completely random origin for the genetic code. The  gradual introduction of two new bases (G and C) to a primitive  genetic code based on U and A must have allowed greater infor¬  mation capacity (i.e., coding for 20 amino acids) while minimizing  the rate of deleterious substitutions.    However, they also specify the amino acids Met and Val, respectively,  at internal positions in polypeptide chains. We will see in Section 27-4A  how ribosomes differentiate the two types of codons.    ^Nonpolar amino acid residues are gold, basic residues are blue, acidic residues are red, and polar uncharged residues are purple.  ^AUG forms part of the initiation signal as well as coding for internal Met residues.    ? What are the consensus nucleotides in the codons for each colored grouping of amino acids?   • Why do frameshift mutations occur with  the insertion or deletion of one or two,  but not three, nucleotides?   • Why must a polypeptide sequence begin  with a designated Start codon?   • Explain why codons must consist of at  least three nucleotides.    The “Standard” Genetic Code Is Not Universal. For many years, it was  thought that the “standard” genetic code (that given in Table 27-1) was uni¬  versal. This assumption was based in part on the observation that one kind of  organism (e.g., E. coli) can accurately translate the genes for quite different  organisms (e.g., humans). Indeed, this phenomenon is the basis of genetic en¬  gineering. DNA studies in 1981 nevertheless revealed that the genetic codes of  certain mitochondria are variants of the “standard” genetic code. For example, in  mammalian mitochondria, AUA, as well as the standard AUG, is a Met /ini¬  tiation codon; UGA specifies Trp rather than “Stop”; and AGA and AGG are  “Stop” rather than Arg. Apparently, mitochondria, which contain their own  genes and protein-synthesizing systems, are not subject to the same evolution¬  ary constraints as are nuclear genomes. An alternate genetic code also appears  to have evolved in ciliated protozoa, which branched off very early in eukary¬  otic evolution. Thus, the “standard” genetic code, although very widely uti¬  lized, is not universal.   • An aminoacyl-tRNA synthetase charges each tRNA with the appropriate amino  acid.   • The wobble hypothesis explains the ability of a tRNA to recognize more than one  codon.    Cells must translate the language of RNA base sequences into the language  of polypeptides. In 1955, Crick hypothesized that translation occurs  through the mediation of “adaptor” molecules, which we now know are  tRNAs that carry a specific amino acid and recognize the corresponding  mRNA codon.   In 1965, after a 7-year effort, Robert Holley reported the first known base  sequence of a biologically significant nucleic acid, that of the 76-residue yeast  alanine tRNA (tRNA^). Currently, the base sequences of many thousands  of tRNAs from nearly 800 organisms and organelles are known (most from their  DNA sequences). They vary in length from 54 to 100 nucleotides (18-28 kD)  although most have ~76 nucleotides.   Almost all known tRNAs can be schematically arranged in the so-called  cloverleaf secondary structure (Fig. 27-3). Starting from the 5' end, they have  the following common features:   2. A 7-bp stem that includes the 5'-terminal nucleotide and that may con¬  tain non-Watson-Crick base pairs such as G • U. This assembly is known  as the acceptor or amino acid stem because the amino acid residue car¬  ried by the tRNA is covalently attached to its 3'-terminal OH group.   3. A 3- or 4-bp stem ending in a 5- to 7-nt loop that frequently contains  the modified base dihydrouridine (D). This stem and loop are there¬  fore collectively termed the D arm.   4. A 5-bp stem ending in a loop that contains the anticodon. These fea¬  tures are known as the anticodon arm.   5. A 5-bp stem ending in a loop that usually contains the sequence TiJjC  (where vji is the symbol for pseudo uridine; Section 26-3C). This as¬  sembly is called the Ti|jC or T arm.    FIG. 27-3 The cloverleaf secondary structure of tRNA. Filled circles connected by  dots represent Watson-Crick base pairs, and open circles indicate bases involved in  non-Watson-Crick base pairing. Invariant positions are indicated: R and Y represent  invariant purines and pyrimidines, and i|/ represents pseudouridine. The starred nucleotides  are often modified. The D and variable arms contain different numbers of nucleotides in the  various tRNAs.    6. A y CCA sequence with a free 3'-OH group. The —CCA may be  genetically specified or enzymatically added to immature tRNA,  depending on the species (Section 26-3D).    tRNAs have 15 invariant positions (always have the same base) and  8 semi-invariant positions (only a purine or only a pyrimidine) that  occur mostly in the loop regions. The purine on the 3' side of the an¬  ticodon is invariably modified. The site of greatest variability among  the known tRNAs occurs in the so-called variable arm. It has from  to 21 nucleotides and may have a stem consisting of up to 7 bp.    tRNAs Have Numerous Modified Bases. One of the most striking charac¬  teristics of tRNAs is their large proportion, up to 25%, of posttranscription-  ally modified bases. Nearly 80 such bases, found at >60 different tRNA  positions, have been characterized. A few of them, together with their stan¬  dard abbreviations, are indicated in Fig. 27-4. None of these modifications  are essential for maintaining a tRNAs structural integrity or for its proper  binding to the ribosome. However, base modifications may help promote at¬  tachment of the proper amino acid to the acceptor stem or strengthen  codon—anticodon interactions.    tRNA Has a Complex Tertiary Structure. As described in Section 24-2C,  tRNA molecules have an L-shape in which the acceptor and T stems form one    iV 7 -Methylguanosine (m 7 G) AT 2 ,iV 2 -Dimethylguanosine (m 2 G)    FIG. 27-4 A few of the modified nucleosides that occur in may also be methylated at their ribose 2' positions to form residues  tRNAs. Note that although inosine chemically resembles guanosine, it symbolized, for instance, by Cm, Gm, and Urn. © See Kinemage  is biochemically derived by the deamination of adenosine. Nucleosides Exercise 21-2.    FIG. 27-5 The structure of yeast tRNA Phe . (a) The base  sequence drawn in cloverleaf form. Tertiary base-pairing interactions  are represented by thin red lines connecting the participating bases.  Bases that are conserved or semiconserved in all tRNAs are circled  by solid and dashed lines, respectively. The 5' terminus is colored  bright green, the acceptor stem is yellow, the D arm is white, the  anticodon arm is light green, the variable arm is orange, the Ti|/C arm  is cyan, and the 3' terminus is red. (b) The X-ray structure drawn to   show how its base-paired stems form an L-shaped molecule. The  tRNA is drawn in stick form with C atoms colored as in Part a, N  blue, and 0 red. Adjacent P atoms are connected by rods colored as  in Part a. [Based on an X-ray structure by Sung-Hou Kim, PDBid  6TRNA.] Q See Kinemage Exercise 21-1.   Why is it important for the anticodon bases not to form  intramolecular base pairs?    leg and the D and anticodon stems form the other (Fig. 27-5). Each leg of  the L is ~60 A long and the anticodon and amino acid acceptor sites are at  opposite ends of the molecule, some 76 A apart. The narrow 20- to 25-A  width of tRNA is essential to its biological function: During protein synthe¬  sis, three tRNA molecules must simultaneously bind in close proximity at ad¬  jacent codons on mRNA (Section 27-4B).   tRNA’s complex tertiary structure is maintained by extensive stacking interac¬  tions and base pairing within and between the helical stems. Many of the ter¬  tiary base-pairing interactions are non-Watson—Crick associations. Moreover,  most of the bases involved in these interactions are either invariant or semi¬  invariant, consistent with the notion that all tRNAs have similar conformations.  The compact structure of yeast tRNA Phe renders most of its bases inaccessible  to solvent. The most notable exceptions are the anticodon bases and those of  the amino acid—bearing —CCA terminus. Both of these groupings must be  accessible in order to carry out their biological functions.   1. The correct amino acid must be selected for covalent attachment to a  tRNA by an aminoacyl-tRNA synthetase (discussed below).   2. The correct aminoacyl-tRNA (aa-tRNA) must pair with an mRNA  codon at the ribosome (discussed in Section 27-4B).    An amino acid—specific aminoacyl—tRNA synthetase (aaRS) appends an amino  acid to the 3'-terminal ribose residue of its cognate tRNA to form an aa-tRNA  (Fig. 27-6). Aminoacylation occurs in two sequential reactions that are cat¬  alyzed by a single enzyme.   which, with all but three aaRSs, can occur in the absence of tRNA.  Indeed, this intermediate can be isolated, although it normally remains  tightly bound to the enzyme.   2. The mixed anhydride then reacts with tRNA to form the aa-tRNA:   is driven to completion by the hydrolysis of the PP Z generated in the first reaction  step. The aa-tRNA product is a “high-energy” compound (Section 14-2A); for  this reason, the amino acid is said to be “activated” and the tRNA is  said to be “charged.” Amino acid activation resembles fatty acid activation  (Section 20-2A); the major difference is that tRNA is the acyl acceptor in amino  acid activation whereas CoA performs the function in fatty acid activation.    There Are Two Classes of Aminoacyl-tRNA Synthetases. Most cells have  at least one aaRS for each of the 20 amino acids. The similarity of the reac¬  tion catalyzed by these enzymes and the structural similarities among tRNAs  suggest that all aaRSs evolved from a common ancestor and should therefore  be structurally related. This is not the case. In fact, the aaRSs form a diverse  group of enzymes with different sizes and quaternary structures and little sequence  similarity. Nevertheless, these enzymes can be grouped into two classes that  each have the same 10 members in nearly all organisms (Table 27-2). Class I  and Class II aminoacyl-tRNA synthetases differ in several ways:   1. Structural motifs. The Class I enzymes share two homologous  polypeptide segments that are components of a dinucleotide-binding  fold (Rossmann fold, which is also present in many NAD + - and ATP-  binding proteins; Section 6-2C). The Class II synthetases lack the fore¬  going sequences but have three other sequences in common.   2. Anticodon recognition. Many Class I aaRSs must recognize the anti¬  codon to aminoacylate their cognate tRNAs. In contrast, several Class II  enzymes do not interact with their bound tRNAs anticodon.   3. Site of aminoacylation. All Class I enzymes aminoacylate their bound  tRNAs 3'-terminal 2'-OH group, whereas Class II enzymes charge the  3'-OH group. Nevertheless, an aminoacyl group attached at the 2' po¬  sition rapidly equilibrates between the 2' and 3' positions (it must be  at the y position to take part in protein synthesis).   4. Amino acid specificity. The amino acids for which the Class I syn¬  thetases are specific tend to be larger and more hydrophobic than those  for Class II synthetases.   amino acid residue is esterified to the tRNA’s  3'-terminal nucleotide at either its 3'-0H group,  as shown here, or its 2'-0H group.   FIG. 27-7 Experimentally observed  identity elements of tRNAs. The tRNA  backbone is cyan, and each of its nucleotides is  represented by a yellow circle whose diameter is  proportional to the fraction of the various tRNA  acceptor types for which the nucleotide is an  identity element for aaRS recognition. [Courtesy  of William McClain, University of Wisconsin.]   Aminoacyl-tRNA Synthetases Recognize Unique Structural Features of  tRNA. How does an aaRS recognize a tRNA so that it can be charged with  the proper amino acid? First, all tRNAs have similar structures, so the features  that differentiate them must be subtle variations in sequence or local struc¬  ture. On the other hand, since the genetic code is degenerate, more than one  tRNA may carry a given amino acid. The members of each set of these so-  called isoaccepting tRNAs must all be recognized by their cognate aaRS.  Finally, the tRNA must be charged with only the amino acid that corresponds  to its anticodon, and not any of the 19 other amino acids.   Clues to the specificity of synthetase-tRNA interactions have been gleaned  from studies using tRNA fragments, mutationally altered tRNAs, chemical cross-  linking agents, computerized sequence comparisons, and X-ray crystallography.  When the experimentally determined synthetase contact sites, or identity ele¬  ments, for a variety of tRNAs are mapped onto a three-dimensional model of a  tRNA molecule, they cluster in the acceptor stem, the anticodon loop, and other  points on the inner (concave) face of the L (Fig. 27-7). However, there appears  to be little regularity in how the synthetases recognize their cognate tRNAs, and  some enzyme—tRNA interactions do not involve the anticodon at all.   Synthetases that do interact with both the anticodon and the acceptor  stem must have a size and structure adequate to bind both legs of the L-shaped  tRNA. This is evident in the complex of E. coli glutaminyl-tRNA synthetase  (GlnRS) with tRNA Gln and ATP (Fig. 27-8), determined by Thomas Steitz,  the first such structure to be elucidated. GlnRS, a 553-residue monomeric  Class I enzyme, has an elongated shape so that it binds the anticodon near  one end of the protein and the acceptor stem near the other. Genetic and bio¬  chemical data indicate that the identity elements for tRNA Gln include all seven  bases of the anticodon loop. The bases of the anticodon itself are unstacked  and splay outward so as to bind in separate recognition pockets of GlnRS.  The 3' end of tRNA Gln plunges deeply into a protein pocket that also binds  the enzymes ATP and glutamine substrates.   Yeast AspRS, a Class II enzyme, is a homodimer of 557-residue subunits.  Its X-ray structure in complex with tRNA^, determined by Dino Moras, re¬  veals that the protein symmetrically binds two tRNA molecules by contacting  them principally at their acceptor stem and anticodon regions (Fig. 27-9). The  anticodon arm of tRNA Asp is bent by as much as 20 A toward the inside of  the L relative to that in the X-ray structure of uncomplexed tRNA Asp , and its  anticodon bases are unstacked. The hinge point for the bend is a G30 • U40  base pair in the anticodon stem (nearly all other species of tRNA contain a  Watson-Crick base pair at that point). The anticodon bases of tRNA Gln are  also unstacked in contacting GlnRS but with a backbone conformation that  differs from that in tRNA Asp . Evidently, the conformation of a tRNA in com¬  plex with its cognate synthetase is dictated more by its interactions with the  protein (induced fit) than by its sequence. This is perhaps one reason why the  members of each set of isoaccepting tRNAs in a cell are recognized by a single aaRS.   The different modes of tRNA binding by GlnRS and AspRS can be seen  by comparing Figs. 27-8 and 27-9. Although both tRNAs approach their syn¬  thetases along the inside of the L shapes, tRNA Gln does so from the direction  of the minor groove of its acceptor stem, whereas tRNA Asp does so from the   FIG. 27-8 X-Ray structure of E. coli GlnRS • tRNA Gln • ATP. The ATP bound in the  protein’s active site is drawn in space-filling form with C green, N blue, 0 red, and P  orange. The tRNA is drawn in stick form colored as the ATP but with the C atoms of the  anticodon (UCG) and the 3'-CCA end yellow. An orange rod links its successive P atoms.  The protein is represented by a semitransparent lavender surface diagram that reveals the  buried portions of the tRNA and ATP. Note that both the 3' end of the tRNA (top right)  and its anticodon bases ( bottom ) are inserted into deep pockets in the protein. [Based on  an X-ray structure by Thomas Steitz, Yale University. PDBid 1GTR.] ©• See Kinemage  Exercise 22.   FIG. 27-9 X-Ray structure of yeast AspRS • tRNA Asp . The   homodimeric enzyme with its two symmetrically bound tRNAs is viewed  with its twofold axis approximately vertical. The tRNAs are drawn in  skeletal form colored according to atom type with the C atoms of the  anticodon (GUC) and the 3'-CCA end yellow, the remaining C atoms  green, N blue, 0 red, and P orange. An orange rod connects successive  P atoms. The two protein subunits are represented by semitransparent  pink and lavender surface diagrams that reveal the buried portions of  the tRNAs. [Based on an X-ray structure by Dino Moras, CNRS/  INSERM/ULP, lllkirch Cedex, France. PDBid 1ASYJ    direction of its major groove. The 3' end of tRNA Asp thereby continues its  helical track as it plunges into AspRS s catalytic site, whereas the 3' end of  tRNA Gln bends backward into a hairpin turn as it enters its active site. These  structural differences account for the observation that Class I and Class II en¬  zymes aminoacylate different OH groups on the 3' terminal ribose of tRNA.   The charging of a tRNA with its cognate amino acid is a remarkably accurate  process. Experimental measurements indicate, for example, that at equal con¬  centrations of isoleucine and valine, IleRS transfers ~40,000 isoleucines to  tRNA Ile for every valine it so transfers. This high degree of accuracy is surpris¬  ing because valine, which differs from isoleucine only by the lack of a single  methylene group, should fit easily into the isoleucine-binding site of IleRS. The  binding free energy of a methylene group is estimated to be ~12 kj • mol l .  Equation 1-17 indicates that the ratio f of the equilibrium constants  K 2 , with which two substances bind to a particular site is given by   where AAG°' = A Gf — A G 2 is the difference between the free  energies of binding of the two substances. It is therefore esti¬  mated that isoleucyl-tRNA synthetase could discriminate be¬  tween isoleucine and valine by no more than a factor of —TOO.   Paul Berg resolved this apparent paradox by demonstrating that,  in the presence of tRNA Ile , IleRS catalyzes the quantitative hydrol¬  ysis of valine-adenylate, the intermediate of the aminoacylation re¬  action, to valine + AMP rather than forming Val—tRNA Ile . Thus,  isoleucyl-tRNA synthetase subjects aminoacyl—adenylates to a proofreading  or editing step, a process reminiscent of that carried out by DNA  polymerase I (Section 25-2A).   IleRS has two active sites that operate as a double sieve. The first site ac¬  tivates isoleucine and the chemically similar valine. The second active site,  which hydrolyzes aminoacylated tRNA Ile , admits only aminoacyl groups that  are smaller than isoleucine (i.e., Val-tRNA Ile ). The X-ray structure of a bacte¬  rial IleRS, a Class I enzyme, determined by Steitz, reveals that the protein con¬  tains an additional editing domain inserted into its dinucleotide-binding fold  domain (Fig. 27-10). The 3' terminus of tRNA Ile appears to shuttle between  the aminoacylation and editing sites by a conformational change. The combined  selectivities of the aminoacylation and editing steps are responsible for the high fi¬  delity of translation, a phenomenon that occurs at the expense of ATP hydrolysis.   FIG. 27-10 X-Ray structure of  Staphylococcus aureus isoleucyl-tRNA  synthetase in complex with tRNA lle . The   tRNA is white, the protein is colored by domain,  and mupirocin, an antibiotic that binds to IleRS,  is drawn in stick form in pink. The four terminal  residues of the tRNA are labeled. The 3'  terminus of the tRNA is positioned near the  editing active site. [Courtesy of Thomas Steitz,  Yale University. PDBid 1QU2J   amino acids through a variety of noncovalent interactions. In ValRS, the side  chain of Asp 279 protrudes into the editing active site, where it hydrogen-bonds  with the hydroxyl group of threonine. The isosteric (having the same shape)  valine lacks this hydroxyl group and is thereby excluded from the editing  pocket. ThrRS has the opposite problem: It must synthesize Thr-tRNA Thr but  not Val-tRNA Thr . Specificity is conferred by the aminoacylation site, which  contains a Zn 2+ ion that is coordinated by the side chain OH group of  threonine. Valine cannot coordinate the Zn 2+ in this way and hence does not  undergo adenylylation by ThrRS. A separate editing site deals with misacylated  Ser-tRNA Tt ir . TyrRS distinguishes between tyrosine and phenylalanine through  hydrogen bonding with tyrosines OH group. Since no other amino acid re¬  sembles tyrosine, the enzyme can do without an editing function.   Some Organisms Lack Certain aaRSs. Many bacteria lack the expected  complement of 20 aminoacyl-tRNA synthetases. For example, GlnRS is ab¬  sent in gram-positive bacteria, archaebacteria, cyanobacteria, mitochondria,  and chloroplasts. Instead, glutamate is linked to tRNA Gln by the same GluRS  that synthesizes Glu-tRNA Glu . The resulting Glu-tRNA Gln is then converted  to Gln-tRNA Gln by Glu-tRNA Gln amidotransferase in an ATP-requiring re¬  action in which glutamine is the amide donor. Some microorganisms that lack  AsnRS use a similar transamidation pathway for the synthesis of Asn-tRNA Asn  from Asp-tRNA Asn .   Certain archaebacteria lack a gene for CysRS. In these cells, Cys-tRNA Cys  is produced by a ProRS (called ProCysRS). Despite its unusual dual speci¬  ficity, ProCysRS does not generate either Cys-tRNA Pro or Pro-tRNA Cys . This  is because adenylylation of Pro does not require the presence of tRNA Pro , but  the activation of Cys requires tRNA Cys . Presumably, the binding of Pro-AMP  or tRNA Cys elicits conformational changes that are mutually exclusive.   In protein synthesis, the proper tRNA is selected only through codon—anticodon inter¬  actions; the aminoacyl group does not participate in this process (this is one reason  why accurate aminoacylation is critical for protein synthesis). The three nu¬  cleotides of an mRNA codon pair with the three nucleotides of a complemen¬  tary tRNA anticodon in an antiparallel fashion. One might naively guess that  each of the 61 codons specifying an amino acid would be read by a different  tRNA. Yet even though most cells contain numerous groups of isoaccepting  tRNAs, many tRNAs bind to two or three of the codons specifying their cognate amino  acids. For example, yeast tRNA Phe , which has the anticodon GmAA (where Gm  indicates G with a 2'-methyl group), recognizes the codons UUC and UUU,   and yeast tRNA Ala , which has the anticodon IGC (where I is inosine), recog¬  nizes the codons GCU, GCC, and GCA:   5' : : : 3' 5' : : : 3' 5' : : : 3'   Evidently, non-Watson—Crick base pairing can occur at the third codon-  anticodon position (the anticodons first position is defined as its 3' nu¬  cleotide), the site of most codon degeneracy (Table 27-1). Note that the third  (50 anticodon position commonly contains a modified base such as Gm or I.    FIG. 27-1 1 U • G and I • A wobble pairs. Both have been observed in X-ray structures.    TABLE 27-3 Allowed Wobble Pairing Combinations in  the Third Codon-Anticodon Position    The Wobble Hypothesis Accounts for Codon Degeneracy. By combining  structural insight with logical deduction, Crick proposed the wobble hypoth¬  esis to explain how a tRNA can recognize several degenerate codons. He as¬  sumed that the first two codon—anticodon pairings have normal Watson-Crick  geometry and that there could be a small amount of play or “ wobble " in the third  anticodon position to allow limited conformational adjustments in its pairing  geometry. This permits the formation of several non-Watson-Crick pairs such  as U • G and I • A (Fig. 27-11). The allowed pairings for the third codon-  anticodon position are listed in Table 27-3. An anticodon with C or A in its  third position can potentially pair only with its Watson-Crick complementary  codon (although, in fact, there is no known instance of a tRNA with an A in  its third anticodon position). If U or G occupies the third anticodon position,  two codons can potentially be recognized. I at the third anticodon position  can pair with U, C, or A.   A consideration of the various wobble pairings indicates that at least 31  tRNAs are required to translate all 61 coding triplets of the genetic code (there  are 32 tRNAs in the minimal set because translation initiation requires a sep¬  arate tRNA; Section 27-4A). Most cells have >32 tRNAs, some of which have  identical anticodons. In fact, mammalian cells have >150 tRNAs. Some or¬  ganisms contain unique tRNAs that are charged with unusual amino acids  (Box 27-2).   Frequently Used Codons Are Complementary to the Most Abundant tRNA  Species. The analysis of the base sequences of several highly expressed struc¬  tural genes of baker s yeast, Saccharomyces cerevisiae , has revealed a remarkable  bias in their codon usage. Only 25 of the 61 coding triplets are commonly used.  The preferred codons are those that are most nearly complementary\ in the  Watson-Crick sense , to the anticodons in the most abundant species in each set of  isoaccepting tRNAs. A similar phenomenon occurs in E. coli y although several of  its 22 preferred codons differ from those in yeast. The degree with which the  preferred codons occur in a given gene is strongly correlated, in both organisms,  with the genes level of expression. This probably permits the mRNAs of pro¬  teins that are required in high abundance to be rapidly and smoothly translated.   • Why is it important for all tRNAs to have  similar structures?   • What features of tRNA structure are in¬  volved in recognition and aminoacylation  by an aaRS?   • Describe how the double-sieve mechanism  of lleRS promotes accurate tRNA amino¬  acylation.   • Which base pairings account for wobble  at the third codon-anticodon position?   It is widely stated that proteins are synthesized from 20 “standard”  amino acids, but two other amino acids are known to be incorpo¬  rated into proteins during translation (other nonstandard amino  acid residues in proteins are the result of posttranslational modifi¬  cations). In both cases, the amino acid is attached to a unique  tRNA that recognizes a Stop codon.   Several enzymes in eukaryotes and prokaryotes contain the  nonstandard amino acid selenocysteine (Sec):   The Sec residues of selenoproteins are thought to participate in  redox reactions such as those catalyzed by mammalian glutathione  peroxidase (Box 15-4) and thioredoxin reductase (Section 23-3A).  For this reason, selenium is an essential trace element. The  human genome contains an estimated 25 selenoproteins.   Selenocysteine, sometimes called the “twenty-first amino  acid,” is incorporated into proteins with the aid of a tRNA that  interprets the UGA Stop codon as a Sec codon. tRNA Sec is  initially charged with serine in a reaction catalyzed by the same  SerRS that charges tRNA Ser . The resulting Ser-tRNA Sec is enzy¬  matically selenylated to produce selenocysteinyl-tRNA Sec .  Although tRNA Sec must resemble tRNA Ser enough to interact  with the same SerRS, its acceptor stem has 8 bp (rather than  7), its D arm has a 6-bp stem and a 4-base loop (rather than a  4-bp stem and a 7-8-base loop), its Ti|iC stem has 4 bp rather  than 5, and its anticodon, UCA, recognizes a UGA Stop codon  rather than a Ser codon. In addition, several of the invariant    residues of other tRNAs are altered in tRNA Sec . These changes  explain why tRNA Sec is not recognized by EF-Tu • GTP, which  escorts other aminoacyl-tRNAs to the ribosome (Section 27-  4B). Instead, a dedicated protein (a special elongation factor)  named SELB in complex with GTP is required to deliver  Sec-tRNA Sec to the ribosome. SELB • GTP • Sec-tRNA Sec  reads the UGA codon as Sec rather than “Stop,” provided that  the ribosomally bound mRNA has a hairpin loop on the 3' side  of the UGA specifying Sec.   includes the amino acid pyrrolysine (Pyl), a lysine with its e-  nitrogen in amide linkage to a pyrroline group:   Pyl is specified by the codon UAG (normally a Stop codon). Pyl is  carried to the ribosome by tRNA Pyl , which contains a CUA anti¬  codon and differs from typical tRNAs in having a D loop with  5 rather than 8 residues, an anticodon stem with 6 rather than  5 bp, and a TvJiC loop that lacks the sequence Ti|iC. A specific  aminoacyl-tRNA synthetase, PylRS, that differs from known  LysRSs, charges tRNA Pyl with pyrrolysine in an ATP-dependent  reaction, the first known example in nature of the direct aminoacy-  lation of a tRNA with a “nonstandard” amino acid. Unlike the case  for Ser-tRNA Sec , Pyl-tRNA^ 1 is delivered to the ribosome by  EF-Tu. This suggests that the mRNA contains a signal that causes  UAG to be read as a Pyl codon rather than as a Stop codon. A  conserved stem-loop structure located on the 3' side of UAG  codons specifying Pyl may comprise this signal.    • The ribosome consists of a large and a small subunit, both composed of RNA  and a large number of small proteins.   • The complex structure of the ribosome allows it to bind mRNA and three tRNA  molecules and to carry out protein synthesis.    Ribosomes, small organelles that were once thought to be artifacts of cell dis¬  ruption, were identified as the site of protein synthesis in 1955 by Paul  Zamecnik, who demonstrated that l4 C-labeled amino acids are transiently as¬  sociated with ribosomes before they appear in free proteins. The ribosome is  both enormous (~2.5 X 10 6 D in bacteria and 3.9 to 4.5 X 10 6 D in eu¬  karyotes) and complex (ribosomes contain several large RNA molecules and  dozens of different proteins). This complexity is necessary for the ribosome to  carry out the following vital functions:   1. The ribosome binds mRNA such that its codons can be read with high  fidelity.   3. The ribosome mediates the interactions of nonribosomal protein factors  that promote polypeptide chain initiation, elongation, and termination.   5. The ribosome undergoes movement so that it can translate sequential  codons.   In this section, we discuss the structure of the ribosome, beginning with  the smaller and simpler prokaryotic ribosome and ending with the larger and  more complicated eukaryotic ribosome.   Ribosomal components are traditionally described in terms of their rate of sed¬  imentation in an ultracentrifuge, which correlates roughly with their size  (Section 5-2E). Thus, the intact E. coli ribosome has a sedimentation coeffi¬  cient of 70S. As James Watson discovered, the ribosome can be dissociated  into two unequal subunits (Table 27-4). The small (30S) subunit consists of  a 16S rRNA molecule and 21 different proteins, whereas the large (50S) sub¬  unit contains a 5S and a 23S rRNA together with 31 different proteins. By  convention, ribosomal proteins from the small and large subunits are desig¬  nated with the prefixes S and L, respectively, followed by a number that  roughly increases from the largest to the smallest. These proteins, which range  in size from 46 to 557 residues, occur in only one copy per ribosome with  the exception of L12, which is present in four copies. Most of these proteins,  which exhibit little sequence similarity with one another, are rich in the basic  amino acids Lys and Arg and contain few aromatic residues, as is expected for  proteins that are closely associated with polyanionic RNA molecules. The up  to 20,000 ribosomes in an E. coli cell account for ~80% of its RNA content  and 10% of its protein.   Ribosomal RNAs Have Complicated Secondary Structures. The E. coli  16S rRNA, which was sequenced by Harry Noller, consists of 1542 nu¬  cleotides. A computerized search of this sequence for stable double-helical seg¬  ments yielded many plausible but often mutually exclusive secondary  structures. However, the comparison of the sequences of 16S rRNAs from sev¬  eral prokaryotes, under the assumption that their structures have been evolu-  tionarily conserved, led to the flowerlike secondary structure for 16S rRNA    FIG. 27-1 2 Secondary structures of the E. coli ribosomal  RNAs. (a) 16S rRNA and (b) 23S and 5S rRNAs. The rRNAs are  colored by domain. The stems, which contain Watson-Crick base pairs,  G • U base pairs, and other non-Watson-Crick base pairs, form    flowerlike structures within each domain. [Courtesy of Venki  Ramakrishnan, MRC Laboratory of Molecular Biology, Cambridge,  U.K., and Peter Moore, Yale University.]    seen in Fig. 27-12#. In this four-domain structure, which is 54% base paired,  the double-helical stems tend to be short (<8 bp) and many of them are im¬  perfect. The large ribosomal subunits 5S and 23S rRNAs, which consist of  120 and 2904 nucleotides, respectively, also exhibit extensive secondary struc¬  tures (Fig. 27-121?).   The Ribosome Has a Complex Three-Dimensional Structure. The struc¬  ture of the ribosome began to come into focus through electron microscopy  (EM; Fig. 27-13#) and later through cryoelectron microscopy (cryo-EM;   Fig. 27-13^). In the latter technique, the sample is cooled to near liquid N 2  temperatures (— 196°C) so rapidly (in a few milliseconds) that the water in   (b) Cryoelectron microscopy-based image of the E. coli ribosome. The  30S subunit ( yellow ) is on the left and the 50S subunit (blue) is on  the right. The tRNAs that occupy the A, P, and E sites (described  below) are colored magenta, green, and brown. A portion of the 50S  subunit has been cut away to reveal the polypeptide exit tunnel. A  segment of mRNA (5' end brown and 3' end lavender) and the nascent  polypeptide chain (yellow) have been modeled into the structure.  [Courtesy of Joachim Frank, State University of New York at Albany.]    the sample does not have time to crystallize but, rather, assumes a vitreous  (glasslike) state. Consequently, the sample remains hydrated and hence retains  its native shape to a greater extent than in conventional electron microscopy.  Cryo-EM studies, carried out largely by Joachim Frank, revealed that the ri¬  bosome has an irregular shape, about 250 A across, with numerous lobes and  bulges as well as channels and tunnels.   The fine structure of the ribosome was determined by X-ray crystallogra¬  phy, a landmark achievement owing to the enormous size of the particle.  Ribosomal subunits were first crystallized by Ada Yonath in 1980, but it took  another 20 years for improvements in crystal quality and technical advances  in crystallography to meet the challenge of determining the X-ray structures  of these gargantuan molecular complexes. In 2000, Peter Moore and Steitz re¬  ported the X-ray structure of the ~ 100,000-atom 50S ribosomal subunit of  the halophilic (salt-loving) bacterium Haloarcula marismortui at atomic (2.4 A)  resolution, and shortly thereafter Venki Ramakrishnan and Yonath inde¬  pendently reported the X-ray structure of the 30S subunit of 77 thermophilus  at ~3 A resolution. Since then, higher-resolution structures of the entire 70S  ribosomes from 77 thermophilus and E. coli have been obtained.   1. Both the 16S and 23S rRNAs are assemblies of helical elements con¬  nected by loops, most of which are irregular extensions of helices  (Fig. 27-14). These structures, which are in close accord with previous  secondary structure predictions (Fig. 27-12), are stabilized by interac¬  tions between helices such as minor groove to minor groove packing  (recall that A-form RNA has a very shallow minor groove); the inser¬  tion of a phosphate ridge into a minor groove; and the insertion of con¬  served adenines into minor grooves. Similar intramolecular interactions  are observed in other complex RNA structures (e.g., Section 26-3C).   2. Each of the 16S rRNAs four domains, which extend out from a cen¬  tral junction (Fig. 27-12 a), forms a morphologically distinct portion  of the 30S subunit (Fig. 27-14 a). In contrast, the 23S rRNAs six do¬  mains (Fig. 27-12 b) are intricately intertwined in the 5OS subunit (Fig.  27-14£). Since the ribosomal proteins are embedded in the RNA (see  below), this suggests that the domains of the 30S subunit can move  relative to one another during protein synthesis, whereas the 5OS sub¬  unit appears to be rigid.   16S rRNA of T. thermophilus. (b ) The 23S rRNA of H. marismortui.  The rRNAs are colored according to domain as in Fig. 27-12. The  interface view of a ribosomal subunit (left) shows the surface that  associates with the other subunit in the whole ribosome and the back  view (right) shows the opposite (solvent-exposed) side. Note that the    secondary structure domains of the 16S rRNA fold as separate  tertiary structure domains, whereas in the 23S rRNA, the secondary  structure domains are more intertwined. The LI protein, part of the  large subunit, is shown for purposes of orientation. [Courtesy of Venki  Ramakrishnan, MRC Laboratory of Molecular Biology, Cambridge, U.K.,  and Peter Moore, Yale University. PDBids 1J5E and 1JJ2.]    FIG. 27-1 5 Distribution of protein and RNA in the ribosomal  subunits, (a) The 30S subunit of T. thermophilus. (b) The 50S  subunit of H. marismortui. The subunits are drawn in space-filling form  with their RNAs gray and their proteins in various colors. Note that  the interface side of each subunit is largely free of protein. The  globular portions of proteins are exposed on the surface of their    associated subunit (Fig. 27-16), whereas their extended segments are  largely buried in the RNA. [Part a based on an X-ray structure by  Venki Ramakrishnan, MRC Laboratory of Molecular Biology,  Cambridge, U.K. Part b based on an X-ray structure by Peter Moore  and Thomas Steitz, Yale University. PDBids 1J5E and 1JJ2.]    3. The distribution of the proteins in the two ribosomal subunits is not  uniform (Fig. 27-15). The vast majority of the ribosomal proteins are  located on the back and sides of their subunits. In contrast, the face  of each subunit that forms the interface between the two subunits, par¬  ticularly those regions that bind the tRNAs and mRNA (see below),  is largely devoid of proteins.   4. Most ribosomal proteins consist of a globular domain and a tail. The  globular domain, when present, is located on a subunit surface (Fig.  27-15). The tail segment, which is largely devoid of secondary struc¬  ture and unusually rich in basic residues, infiltrates between the RNA  helices into the subunit interior (Fig. 27-16). These protein tails make  far fewer base-specific interactions than do other known RNA-binding  proteins. They tend to interact with the RNA through salt bridges be¬  tween their positively charged side chains and the RNAs’ negatively  charged phosphate oxygen atoms, thereby neutralizing the repulsive  charge-charge interactions between nearby RNA segments. Moreover,  the sequences of these proteins’ tails are more conserved than their at¬  tached globular domains. This is consistent with the hypothesis that  the primordial ribosome consisted entirely of RNA (the RNA world)  and that the proteins that were eventually acquired stabilized its struc¬  ture and fine-tuned its function. Indeed, a comparison of Figs. 27-14   FIG. 27-16 Backbone structures of  selected ribosomal proteins. Globular  portions of the proteins are green, and  extended segments are red. [Courtesy of  Venki Ramakrishnan, MRC Laboratory of  Molecular Biology, Cambridge, U.K. PDBid 1J5E.]    FIG. 27-1 7 X-Ray structure of the T. thermophilus ribosome in  complex with tRNA and mRNA. (a) The RNA components of the  complex are drawn in cartoon form except for the 11-residue mRNA,  which is shown in space-filling form. The E site binds tRNA Phe and the A  and P sites bind Phe-tRNA Phe . Proteins have been omitted for clarity. The  16S RNA is cyan, the 23S RNA is yellow, the 5S RNA is blue, the tRNAs  in the A, P, and E sites are green, magenta, and red, and the mRNA, which  is largely occluded by the 16S RNA, is colored according to atom type  with C pink, N blue, 0 red, and P orange, (b) The 16S RNA in interface  view with its bound tRNAs and mRNA, all represented as in Part a.   (c) The 23S RNA in interface view (rotated 180° about the vertical  direction relative to Part b) with its bound tRNAs all represented as in Part  a. [Based on an X-ray structure by Venki Ramakrishnan, MRC Laboratory  of Molecular Biology, Cambridge, U.K. PDBids 2WDK and 2WDL] Q See  Interactive Exercise 53.    and 27-15 indicates that the structures of the large and  small ribosomal subunits are determined largely by their  rRNA components, not their proteins.    In the intact ribosome, the large and small subunits maintain  the overall shapes of the isolated subunits and contact each other at  12 positions via RNA—RNA, protein—protein, and RNA—protein  bridges (Fig. 27-17). Mg 2+ ions mediate many of these interac¬  tions. Although the bridges are essential for the stable association of the two  ribosomal subunits, some of the contacts must be broken and re-formed in or¬  der to permit the ribosome to carry out the various activities of translation.   In Section 27-4, we will see that the large subunit is mainly involved in  mediating biochemical tasks such as catalyzing the reactions of polypeptide elon¬  gation , whereas the small subunit is the major actor in ribosomal recognition  processes such as mRNA and tRNA binding (although the large subunit also par¬  ticipates in tRNA binding). We will also see that rRNA has the major func¬  tional role in ribosomal processes (recall that RNA has demonstrated catalytic  properties; Sections 24-2C and 26-3B).    Ribosomes Have Three tRNA-Binding Sites. Ribosomes have three function¬  ally distinct tRNA-binding sites: the A or aminoacyl site (it accommodates the  incoming aminoacyl-tRNA), the P or peptidyl site (it accommodates the    FIG. 27-1 8 The interactions of tRNAs  with mRNA. These RNA structures are part of  the ribosomal complex shown in Fig. 27-17. The  RNAs are drawn in cartoon form with A-site C  green, P-site C magenta, E-site C red, mRNA C  pink, N blue, and 0 red and with successive P  atoms connected by orange rods. The Phe  residues appended to the A- and P-site tRNAs  are represented in space-filling form. Note their  close approach. [Based on an X-ray structure by  Venki Ramakrishnan, MRC Laboratory of  Molecular Bioloqy, Cambridqe, U.K. PDBid  2WDK.]   peptidyl-tRNA, the tRNA to which the growing peptide chain is attached),  and the E or exit site (it accommodates a deacylated tRNA that is about to exit  the ribosome). The ribosome binds all three tRNAs in a similar manner with  their anticodon arms bound to the 3OS subunit and their remaining portions  bound to the 5 OS subunit (Fig. 27-17). These interactions, which mainly con¬  sist of RNA—RNA contacts, are made to the tRNAs universally conserved seg¬  ments, thereby permitting the ribosome to bind different species of tRNAs in  the same way. Nevertheless, the three bound tRNAs have slightly different con¬  formations, with the A-site tRNA most closely resembling the X-ray structure  of tRNA Phe (Fig. 27-5 £) and the E-site tRNA most distorted.   Only the A-site and P-site tRNAs interact closely with the mRNA through  base pairing (Fig. 27-18). In the ribosomal complex, the acceptor ends of these  two tRNAs are close together, a necessity for the peptidyl transferase reaction  that takes place there.   The growing polypeptide fits into a tunnel on the 5OS subunit that ex¬  tends from the P site to the outer ribosomal surface (the tunnel is visible in  Fig. 27-13^). The ~100-A-long tunnel is lined with mostly hydrophilic  residues and has an average diameter of ~15 A. This is barely large enough  for an a helix, so significant protein folding probably cannot occur until the  polypeptide exits the ribosome.   Although eukaryotic and prokaryotic ribosomes resemble each other in both  structure and function, they differ in nearly all details. Eukaryotic ribosomes  have particle masses in the range 3.9 to 4.5 X 10 6 D, about 40% larger than  bacterial ribosomes, and have a nominal sedimentation coefficient of 80S.  They dissociate into two unequal subunits with compositions that are dis¬  tinctly different from those of prokaryotes (Table 27-5; compare with Table  27-4). The small (40S) subunit of the rat liver cytoplasmic ribosome, a well-  characterized eukaryotic ribosome, consists of 33 unique polypeptides and an  18S rRNA. Its large (60S) subunit contains 49 different polypeptides and  three rRNAs of 28S, 5.8S, and 5S. The additional complexity of the eukary¬  otic ribosome relative to its prokaryotic counterpart is presumably due to the  eukaryotic ribosomes additional functions: Its mechanism of translational ini¬  tiation is more complex (Section 27-4A); it must be transported from the nu¬  cleus, where it is formed, to the cytoplasm, where translation occurs; and the  machinery with which it participates in the secretory pathway is more com¬  plicated (Section 9-4D).    Sequence comparisons of the corresponding rRNAs from various species  indicate that evolution has conserved their secondary structures rather than  their base sequences (Fig. 27-12). For example, a G • C in a base-paired stem  of E. coli 16S rRNA has been replaced by an A • U in the analogous stem of  yeast 18S rRNA. The 5.8S rRNA, which occurs in the large eukaryotic sub¬  unit in base-paired complex with the 28S rRNA, is homologous in sequence  to the 5' end of prokaryotic 23S rRNA. Apparently 5.8S RNA arose through  mutations that altered rRNAs posttranscriptional processing to produce a  fourth rRNA.   An X-ray structure of the yeast 80S ribosome (Fig. 27-19) was determined  at 4.15 A resolution by Marat Yusupov, taking advantage of the known struc¬  tures of the H. marismortui 5OS subunit and the 77 thermophilus 30S subunit.   FIG. 27-1 9 X-Ray structure of the yeast  80S ribosome. Proteins and rRNAs are drawn  in ribbon form with the 60S proteins orange,   60S RNA yellow, 40S proteins blue, and 40S  RNA cyan. Expansion segments, RNA sequences  that are not present in bacterial ribosomes, are  shown in red. [Courtesy of Marat Yusupov,   Institut de Genetique et de Biologie Moleculaire  et Cellulaire, Strasbourg, France.] © See  Interactive Exercise 53.   • Summarize the relationship between rRNA  and the overall structure of the ribosome.   • Describe the positions and functions of  the three tRNA-binding sites in the  ribosome.   • Describe the similarities and differences  between prokaryotic and eukaryotic  ribosomes.    The shape of the yeast ribosome reveals that there is a high degree of struc¬  tural conservation between eukaryotic and prokaryotic ribosomes (compare  Fig. 27-19 with Fig. 27-17a). The greater size and functional complexity of  the eukaryotic ribosome are due to an additional 25 proteins that have no  counterparts in bacteria, and additional segments among the conserved pro¬  teins, as well as additional rRNA sequences (“expansion segments”) inserted  into the conserved core rRNAs. The expansion segments cluster mostly on the  solvent-exposed surface of the ribosome rather than in the subunit interface.  The yeast ribosome exhibits 16 intersubunit bridges, 12 of which match the  12 that were observed in the X-ray structure of the 77 thermophilus ribosome,  a remarkable evolutionary conservation that indicates the importance of these  bridges. A similar X-ray structure of the 40S subunit from the protozoan  Tetrahymena thermophila has been determined by Nenad Ban.    • Initiation factors help to assemble the ribosomal subunits, deliver the initiator  tRNA, and in eukaryotes, locate the initiation codon.   • The ribosome selects the correct aminoacyl-tRNA, catalyzes the transpeptidation  reaction, and then translocates along the mRNA during the elongation phase of  protein synthesis.    In order to appreciate the manner in which the ribosome orchestrates the  translation of mRNA to synthesize polypeptides, it is helpful to assimilate the  following points:   1. Polypeptide synthesis proceeds from the N-terminus to the C-terminus; that  is, a peptidyl transferase activity appends an incoming amino acid to  a growing polypeptides C-terminus. This was shown to be the case in  1961 by Howard Dintzis, who exposed reticulocytes (immature red  blood cells) that were actively synthesizing hemoglobin to 3 H-labeled  leucine for less time than it takes to synthesize an entire polypeptide.  The extent to which the tryptic peptides from the soluble (completed)  hemoglobin molecules were labeled increased with their proximity to  the C-terminus (Fig. 27-20), thereby indicating that incoming amino  acids are appended to the growing polypeptides C-terminus.    FIG. 27-20 Demonstration that  polypeptide synthesis proceeds from the  N- to the C-terminus. Rabbit reticulocytes  were incubated with [ 3 H]leucine for the indicated  times. The curves show the distribution of  [ 3 H]Leu among the tryptic peptides from the p  subunit of soluble rabbit hemoglobin. The  numbers on the horizontal axis are peptide  identifiers, arranged from the N-terminus to the  C-terminus. [After Dintzis, H.M., Proc. Natl. Acad.  Sci. 47, 255 (1961).]    FIG. 27-21 The ribosomal peptidyl transferase reaction forming a peptide bond.   The amino group of the aminoacyl-tRNA in the A site nucleophilically displaces the tRNA  of the peptidyl-tRNA ester in the P site, thereby forming a new peptide bond and  transferring the nascent polypeptide to the A-site tRNA.    What is the approximate free energy change for the transfer reaction?    2. Chain elongation occurs by linking the growing polypeptide to the incoming  tRNA’s amino acid residue . If the growing polypeptide is released from  the ribosome by treatment with high salt concentrations, its C-terminal  residue is esterified to a tRNA molecule as a peptidyl-tRNA ( at right) -  The nascent (growing) polypeptide must therefore grow by being trans¬  ferred from the peptidyl-tRNA in the P site to the incoming aa-tRNA  in the A site to form a peptidyl-tRNA with one more residue (Fig.  27-21). After the peptide bond has formed, the new peptidyl-tRNA,  which now occupies the A site, is translocated to the P site so that a  new aa-tRNA can enter the A site. The uncharged tRNA in the P site  moves to the E site before it dissociates from the ribosome (we discuss  the details of chain elongation in Section 27-4B).   3. Ribosomes read mRNA in the 5' 3' direction. This was shown through   the use of a cell-free protein-synthesizing system in which the mRNA  was poly(A) with a 3 r -terminal C:   Such a system synthesizes a poly(Lys) that has a C-terminal Asn:   Together with the knowledge that AAA and AAC code for Lys and Asn  (Table 27-1) and the polarity of peptide synthesis, this established that  the mRNA is read in the 5' —> 3' direction. Because mRNA is also  synthesized in the 5' —> 3' direction, prokaryotic ribosomes can com¬  mence translation as soon as a nascent mRNA emerges from RNA  polymerase. This, however, is not possible in eukaryotes because the  nuclear membrane separates the site of transcription (the nucleus) from  the site of translation (the cytosol).   FIG. 27-22 Polysomes, (a) Electron micrograph of polysomes  from silk gland cells of the silkworm Bombyx mori. The 3' end of the  mRNA is on the left. Arrows point to the silk fibroin polypeptides. The  bar represents 0.1 [xm. [Courtesy of Oscar L. Miller, Jr., and Steven  L. McKnight, University of Virginia.] (b) Cryo-EM-based image of an  E. coli polysome. The mRNA (which is mostly occluded) is represented  by a red line, the small ribosomal subunits are yellow, the large  subunits are cyan, and the red cones point to the polypeptide exit    tunnel on each large subunit. This assembly has a pseudohelical  arrangement of ribosomes in which the center-to-center distance  between adjacent ribosomes averages —230 A. Polysomes with  somewhat different although equally densely packed arrangements of  ribosomes have also been observed. [Courtesy of Ulrich Hartl and  Wolfgang Baumeister, Max Planck Institute of Biochemistry,  Martinsreid, Germany.]   4. Active translation occurs on polysomes. In both prokaryotes and eukaryotes,  multiple ribosomes can bind to a single mRNA transcript, giving rise to  a beads-on-a-string structure called a polyribosome (polysome; Fig.  27-22). Individual ribosomes are arranged in a helical fashion so that they  have a maximum density on the mRNA of ~1 ribosome per 80 nt.  Polysomes arise because once an active ribosome has cleared its initiation  site on mRNA, a second ribosome can initiate translation at that site.   The first indication of how ribosomes initiate polypeptide synthesis was the  observation that almost half of the E. coli proteins begin with the otherwise  uncommon amino acid residue Met. In fact, the tRNA that initiates transla¬  tion is a peculiar form of Met—tRNA Met in which the Met residue is 7V-formy-  lated {at left). Because the AAformylmethionine residue (fMet) already has an  amide bond, it can only be the N-terminal residue of a polypeptide. E. coli  proteins are posttranslationally modified by deformylation of their fMet  residue and, in many proteins, by the subsequent removal of the resulting N-  terminal Met. This processing usually occurs on the nascent polypeptide,  which accounts for the observation that mature E. coli proteins all lack fMet.   The tRNA that recognizes the initiation codon, tRNAf 1 **, differs from  the tRNA that carries internal Met residues, tRNA^ et , although they both  recognize the same AUG codon. Presumably, the conformations of these  tRNAs are different enough to permit them to be distinguished in the reactions  of chain initiation and elongation.   In E. coli , uncharged tRNAf 1 ** is aminoacylated with Met by the same  MetRS that charges tRNA^ et . The resulting Met-tRNAf 1 ** is specifically N-  formylated to yield fMet-tRNAf 1 ** by a transformylase that employs N 10 -  formyltetrahydrofolate (Section 21-4D) as its formyl donor. This transformylase  does not recognize Met-tRNA^**.   Base Pairing between mRNA and the 16S rRNA Helps Select the  Translation Initiation Site. AUG codes for internal Met residues as well as   QP phage replicase  4>X174 phage A protein  R17 phage coat protein  Ribosomal S12  Ribosomal L10  trpE   A G C  C A A  UU C  U A A  A AU  U C A  AAA  C U A  C A A  GU A    GG AUG  C U A AU  UU C A G  A C A C A  C U A AG    C UU  ACC  ACC  C C AGG  A A U U A  A A A AG   A A C  G A A  C UU  UUG  C U A  AAA  A AU  AU C    A AC G  A A U U  G A U U  A G C U  AUG C   A A G C  UUU A  G C U A  A AC A  G AC A    FIG. 27-23 Some translation initiation sequences recognized by E. coli  ribosomes. The RNAs are aligned according to their initiation codons ( blue shading). Their  Shine-Dalgarno sequences (red shading) are complementary, counting G • U pairs, to a  portion of the 16S rRNA’s 3' end (below). [After Steitz, J.A., in Chambliss, G., Craven,   G. R., Davies, J., Davis, K., Kahan, L, and Nomura, M. (Eds.), Ribosomes. Structure, Function  and Genetics, pp. 481-482, University Park Press (1979).]   the initiating Met residue of a polypeptide. Moreover, mRNAs usually con¬  tain many AUGs (and GUGs) in different reading frames. Clearly, a transla¬  tion initiation site must be specified by more than just an initiation codon.   In E. coli, the 16S rRNA contains a pyrimidine-rich sequence at its 3'  end. This sequence, as John Shine and Lynn Dalgarno pointed out in 1974,  is partially complementary to a purine-rich tract of 3 to 10 nucleotides, the  Shine-Dalgarno sequence, that is centered ~10 nucleotides upstream from  the start codon of nearly all known prokaryotic mRNAs (Fig. 27-23). Base¬  pairing interactions between an mRNAs Shine-Dalgarno sequence and the 16S  rRNA apparently permit the ribosome to select the proper initiation codon.   The X-ray structure of the 70S ribosome reveals, in agreement with Fig.  27-13 by that an ~30-nt segment of the mRNA is wrapped in a groove that  encircles the neck of the 30S subunit (Fig. 27-24). The mRNA codons in the  A and P sites are exposed on the interface side of the 30S subunit, whereas  its 5' and 3 ' ends are bound in tunnels composed of RNA and protein. The  mRNAs Shine-Dalgarno sequence, which is located near its 5' end, is base-  paired, as expected, with the 16S rRNAs anti-Shine—Dalgarno sequence,  which is situated close to the E site. The proteins that in part form the  tunnel through which the mRNA enters the ribosome (green in Fig. 27-  24) probably function as a helicase to remove secondary structures from the  mRNA that would otherwise interfere with tRNA binding.    Initiation Requires Soluble Protein Factors. Translation initiation in  E. coli is a complex process in which the two ribosomal subunits and  fMet-tRNAf^ et assemble on a properly aligned mRNA to form a    FIG. 27-24 Path of mRNA through the T. thermophilus 30S  subunit. The ribosomal subunit is viewed from its interface side. The 16S  rRNA is cyan, the mRNA is represented in worm form with its A- and P-site  codons orange and red, and the Shine- Dalgarno helix (which includes a  segment of 16S rRNA) magenta, and its remaining segments yellow. The  ribosomal proteins that in part form the entry and exit tunnels for the  mRNA are drawn in green and purple. The remaining ribosomal proteins  have been omitted for clarity [Courtesy of Gloria Culver, Iowa State  University. Based on an X-ray structure by Harry Noller, University of  California at Santa Cruz. PDBid 1JGO.]    complex that can commence chain elongation. This process also requires  initiation factors that are not permanently associated with the ribosome,  designated IF-1, IF-2, and IF-3 in E. coli (Table 27-6).   Translation initiation in E. coli occurs in three stages (Fig. 27-25):   1. On completing a cycle of polypeptide synthesis, the 30S and 5OS sub¬  units are separated (Section 27-4C). IF-3 binds to the 30S subunit so  as to prevent the reassociation of the 5OS complex. The X-ray struc¬  ture of the 30S subunit in complex with the C-terminal domain of  IF-3 (which by itself prevents the association of the 30S and 5OS sub¬  units), determined by Yonath and Francois Franceschi, indicates that  IF-3 binds to its solvent-exposed (back) side. Apparently, IF-3 does not  function by physically blocking the binding of the 5OS subunit.   2. mRNA and IF-2 in a ternary complex with GTP and fMet-tRNAf 1611 ,  along with IF-1, subsequently bind to the 30S subunit in either order.  Since the ternary complex containing fMet-tRNA^ et can bind to the  ribosome before mRNA, fMet—tRNAf 161 binding must not be medi¬  ated by a codon-anticodon interaction; it is the only tRNA-ribosome  association that does not require one, although this interaction helps  bind fMet-tRNA^ et to the ribosome. IF-1 binds in the A site, where  it may prevent the inappropriate binding of a tRNA. IF-3 also func¬  tions at this stage of the initiation process by preventing the binding  of tRNAs other than tRNA t Met .   3. Last, in a process that is preceded by IF-1 and IF-3 release, the 50S  subunit joins the 30S initiation complex in a manner that stimulates  IF-2 to hydrolyze its bound GTP to GDP + P z . This irreversible   Releases mRNA and tRNA from  recycled 30S subunit and aids  new mRNA binding   Together with EF-G, induces  ribosomal dissociation to small  and large subunits    reaction conformationally rearranges the 30S subunit and releases  IF-2 for participation in further initiation reactions.   Initiation results in the formation of an fMet—tRNAj 4 ^ • mRNA • ribosome  complex in which the fMet—tRNA^ et occupies the ribosomes P site while its A site  is poised to accept an incoming aa—tRNA (an arrangement analogous to that at    IF-3 binds to 30S subunit, which  (prevents the reassociation of  the 50S subunit.   IF-2 - GTP • f Met - tRNAf et  + IF-1   mRNA, the IF-2 • GTP • fMet-  tRNA^ et ternary complex, and IF-1  2 > bind to the 30S subunit. The  Shine-Dalgarno sequence  positions the mRNA.   IF-1 and IF-3 are released and the  50S subunit binds. This causes IF-2  to hydrolyze its bound GTP to GDP +  P j, all of which are released, which  prepares the ribosome to accept an  aminoacyl-tRNA in its A site.    FIG. 27-25 Translation initiation pathway in E. coli. The E site, which is unoccupied during this process, has been omitted for clarity.    the conclusion of a round of elongation; Section 27-4B). Note that tRNAf 1 ^  is the only tRNA that directly enters the P site. All other tRNAs must first  enter the A site during chain elongation.   Ribosomal initiation in eukaryotes requires the assistance of at least 12 initi¬  ation factors (designated clFn; “e” for eukaryotic) that consist of 26 polypep¬  tide chains. Nevertheless, Steps 1 and 3 of the prokaryotic process (Fig. 27-25)  are superficially similar in eukaryotes. However, the way in which the mRNAs  initiating codon is identified in eukaryotes is fundamentally different from that  in prokaryotes.   The pairing of the initiator tRNA with the initiating AUG in the riboso¬  mal P site begins with eIF2, a heterotrimer in complex with GTP, escorting  the initiator tRNA to the complex of the 40S subunit with several other ini¬  tiation factors, thereby forming the 43S preinitiation complex (a process that  resembles Step 2 in prokaryotic initiation but without the mRNA). The ini¬  tiator tRNA is tRNAf* 6 * (“i” for initiator). Its appended Met residue is not  formylated as in prokaryotes. Nevertheless, both species of initiator tRNAs are  readily interchangeable in vitro.   Eukaryotic mRNAs lack the complementary sequences to bind to the 18S  rRNA in the Shine—Dalgarno manner. Rather, they have an entirely different  mechanism for recognizing the mRNAs initiating AUG codon. Eukaryotic  mRNAs , nearly all of which have a 7-methylguanosine (m 7 G) cap and a poly (A)  tail (Section 26-3A), are invariably monocistronic and almost always initiate trans¬  lation at their leading AUG. This AUG, which occurs at the end of a 5'-untrans¬  lated region of 50 to 70 nt, is embedded in the consensus sequence  GCCRCCAUGG. Changes in the purine (R) 3 nt before the AUG and in the  G immediately following it each reduce translational efficiency by ~ 10-fold. The  recognition of the initiation site begins by the binding of eIF4F to the mRNAs  m 7 G cap. eIF4F is a heterotrimeric complex of eIF4E, eIF4G, and eIF4A (all  monomers), in which eIF4E (also known as cap-binding protein) recognizes  the mRNAs m 7 G cap and eIF4G serves as a scaffold to join eIF4E with eIF4A.   The structure of eIF4E in complex with m 7 GDP reveals that the protein  binds the m 7 G base by intercalating it between two highly conserved Trp residues  in a region that is adjacent to a positively charged cleft that presumably forms the  mRNA-binding site (Fig. 27-26). The m 7 G base is specifically recognized by  hydrogen bonding to protein side chains in a manner reminiscent of G • C base  pairing. eIF4G also binds poly (A)-binding protein (PABP; Section  26-3A), which coats the mRNAs poly(A) tail, thereby circulariz¬  ing the mRNA. Although this explains the synergism between an  mRNAs m 7 G cap and its poly(A) tail in stimulating translational  initiation, the function of the circle is unclear. However, an at¬  tractive hypothesis is that it enables a ribosome that has finished  translating the mRNA to reinitiate translation without having to  disassemble and then reassemble. Another possibility is that it pre¬  vents the translation of incomplete (broken) mRNAs.   The eIF4F-mRNA complex is subsequently joined by several  additional initiation factors. The resulting complex joins the 43S    FIG. 27-26 X-Ray structure of murine elF4E in complex with the  m 7 G cap analog m 7 GDP. The protein is shown as its solvent-accessible  surface colored according to its electrostatic potential (red negative, blue  positive, and white neutral). The m 7 GDP is drawn in stick form with C  green, N blue, 0 red, and P yellow. The two Trp residues that bind the  m 7 G base are indicated, as are a Lys and Ser residue that flank the  putative mRNA-binding cleft (yellow arrow). [Courtesy of Nahum  Sonenberg, McGill University, Montreal, Quebec, Canada. PDBid 1EJ1.]    preinitiation complex, which then scans down the mRNA in an ATP-driven  process until it encounters the mRNAs initiating AUG codon, thereby form¬  ing the 48S initiation complex. The recognition of the AUG occurs mainly  through base pairing with the CUA anticodon on the bound Met-tRNAf 4 ^,  as was demonstrated by the observation that mutating this anticodon results  in the recognition of the new cognate codon instead of AUG.   In the analog of Step 3 of prokaryotic initiation (Fig. 27-25), the eIF2-  catalyzed hydrolysis of its bound GTP induces the release of all initiation fac¬  tors from the 48S initiation complex. The resulting 40S subunit-tRNAf 4 ^  complex is joined by the 60S subunit in a GTP-dependent reaction mediated  by eIF5B (a monomer and homolog of prokaryotic IF-2), thereby forming  the 80S ribosomal initiation complex.    B The Ribosome Decodes the mRNA, Catalyzes Peptide Bond  Formation, Then Moves to the Next Codon   Ribosomes elongate polypeptide chains in a three-stage reaction cycle (Fig. 27-27):   1. Decoding, in which the ribosome selects and binds an aminoacyl-tRNA  whose anticodon is complementary to the mRNA codon in the A site.   FIG. 27-27 Elongation cycle in E. coli ribosomes. The E site and EF-Ts are replaced by a single multisubunit protein, eEFI, and   is not shown. Eukaryotic elongation follows a similar cycle, but EF-Tu EF-G is replaced by eEF2.   2. Transpeptidation, or peptide bond formation, in which the peptidyl  group in the P-site tRNA is transferred to the aminoacyl group in the  A site.   3. Translocation, in which the A-site and P-site tRNAs are respectively  transferred to the P and E sites, accompanied by their bound mRNA;  that is, the mRNA, together with its base-paired tRNAs, is ratcheted  through the ribosome by one codon.   This process, which occurs at a rate of 10 to 20 amino acid residues per sec¬  ond, requires several nonribosomal proteins known as elongation factors  (Table 27-6).   Decoding: An Aminoacyl-tRNA Binds to the Ribosomal A Site. In the first  stage of the E. coli elongation cycle, a binary complex of GTP and the elon¬  gation factor EF-Tu combines with an aa-tRNA. The resulting ternary com¬  plex binds to the ribosome. Binding of the aa-tRNA in a codon-anticodon  complex to the ribosomal A site is accompanied by the hydrolysis of GTP to  GDP so that EF-Tu • GDP and P z are released. The EF-Tu • GTP complex  is regenerated when GDP is displaced from EF-Tu • GDP by the elongation  factor EF-Ts, which in turn is displaced by GTP.   Aminoacyl-tRNAs can bind to the ribosomal A site without EF-Tu but  at a rate too slow to support cell growth. The importance of EF-Tu is indi¬  cated by the fact that it is the most abundant E. coli protein; it is present in  — 100,000 copies per cell (>5% of the cells protein), which is approximately  the number of tRNA molecules in the cell. Consequently, the cells entire com¬  plement of aa-tRNAs is essentially sequestered by EF-Tu.   The X-ray structure of EF-Tu in complex with Phe-tRNA Phe and the non-  hydrolyzable GTP analog guanosine-5 r -(P,y-imido)triphosphate (GMPPNP  at left, alternatively GDPNP), was determined by Brian Clark and Jens Nyborg  (Fig. 27-28). This compl ex has a corkscrew-shaped structure in which the EF-  Tu and the tRNAs acceptor stem form a knoblike handle and the tRNAs an¬  ticodon helix forms the screw. The macromolecules appear to associate rather  tenuously via three major regions: (1) The 3'-CCA-Phe segment of the  Phe-tRNA phe binds in the cleft between domains 1 and 2 of EF-Tu •  GMPPNP (the blue and green mainly helical domain and the yellow (3 sheet  domain in Fig. 27-28); (2) the 5'-phosphate of the tRNA binds in a depres¬  sion at the junction of EF-Tus three domains; and (3) one side of the TvJiC  stem of the tRNA makes contacts with exposed main chain and side chains  of EF-Tu domain 3 (the orange (3 barrel-containing domain in Fig. 27-28).   EF-Tu binds all aminoacylated tRNAs, but not uncharged tRNAs or ini¬  tiator tRNAs. Evidently, the tight association of the aminoacyl group with EF-  Tu greatly increases the affinity of EF-Tu for the otherwise loosely bound  tRNA. tRNA^ et does not bind to EF-Tu because it has a 3' overhang of 5 nt  versus 4 nt in an elongator tRNA. This, together with the formyl group at¬  tached to the fMet residue, apparently prevents fMet-tRNA^ et from binding  to EF-Tu, thereby explaining why the initiator tRNA never translates internal  AUG codons.   EF-Tu is a G protein that undergoes a large conformational change on hy¬  drolyzing GTP. Its N-terminal domain 1 (blue, cyan, and green in Fig. 27-28)    FIG. 27-28 X-Ray structure of the ternary complex of yeast Phe-tRNA Phe , Thermus  aquaticus EF-Tu, and GMPPNP. The EF-Tu is drawn in ribbon form colored in rainbow order from  its N-terminus (blue) to its C-terminus (red). The tRNA is shown in stick form colored according to  atom type with C green, N blue, 0 red, and P orange and with orange rods linking successive P  atoms. The tRNA’s appended aminoacyl-Phe residue and the GMPPNP that is bound to the EF-Tu  are drawn in space-filling form with C atoms cyan and yellow, respectively. Two bound Mg 2+ ions  are represented by magenta spheres. [Based on an X-ray structure by Brian Clark and Jens Nyborg,  University of Aarhus, Arhus, Denmark. PDBid 1TTT.]   FIG. 27-29 Comparison of the X-ray  structures of ribosomal elongation factor EF-  Tu in its complexes with GDP and GMPPNP.   The protein is drawn in ribbon form with domain 1,  its GTP-binding domain, magenta in the GDP  complex and yellow in the GMPPNP complex.  Domains 2 and 3, which have the same orientation  in both complexes, are green and cyan. The bound  GDP and GMPPNP are shown in space-filling  form with C green, N blue, 0 red, and P orange.  [Based on X-ray structures by Jens Nyborg,  Aarhus University, Arhus, Denmark. PDBids 1EFT  and 1TUI.] © See Interactive Exercise 54.    Does this structure resemble the G  proteins involved in signal transduction  (see Fig. 13-19)?    resembles other G proteins (Sections 13-2B and 13-3B), with Switch I and  Switch II regions signaling the state of the bound nucleotide (GTP or GDP).  GTP hydrolysis also causes domain 1 to reorient with respect to domains 2  and 3 by a dramatic 91° rotation (Fig. 27-29), a conformational change that  eliminates the tRNA-binding site. Since EF-Tu is a G protein, the ribosome  can be classified as its GTPase-activating protein (GAP) and EF-Ts functions  as its guanine nucleotide exchange factor (GEF).   The X-ray structure of the 77 thermophilus ribosome in complex with  tRNAs and EF-Tu • GDP is shown in Fig. 27-30. The complex was stabi¬  lized by the addition of the antibiotic kirromycin, which prevents EF-Tu from  undergoing conformational changes and dissociating from the ribosome after  GTP hydrolysis. The incoming aminoacyl-tRNA has made contact with its  codon but, because it is bound to EF-Tu, is distorted relative to its conforma¬  tion after it has been fully installed in the A site (as in Fig. 27-17).   The Ribosome Monitors Correct Codon-Anticodon Pairing. When EF-Tu •  GTP delivers an aa-tRNA to the ribosome, it first inserts the anticodon end  of the tRNA into the ribosome. The tRNAs aminoacyl end moves fully into  the A site only after GTP is hydrolyzed and EF-Tu dissociates. This multistep  process allows the ribosome to verify that the aa-tRNA has correctly  paired with an mRNA codon. How does the ribosome monitor  codon-anticodon pairing?   The X-ray structure of the 77 thermophilus 30S subunit in  complex with a U<s hexanucleotide “mRNA” and a 17-nt RNA  consisting of the tRNA Phe anticodon arm (Fig. 27-5, but with  its nucleotides unmodified), determined by Ramakrishnan, re¬  veals how an mRNA-specified tRNA initially binds to the ribo¬  some. The codon-anticodon association is stabilized by its   FIG. 27-30 Model of the ribosome with bound EF-Tu. The T.   thermophilus ribosome is drawn in ribbon form with its 23S RNA orange,  its 50S subunit proteins brown, its 16S RNA cyan, and its 30S proteins  blue. The other RNAs are shown as surface models with mRNA black,  tRNA Phe in the E site yellow, tRNA Phe in the P site green, and  Thr-tRNA Thr , which is bound to EF-Tu (red surface model), magenta. DC  represents the decoding center of the ribosome, and PTC is the peptidyl  transferase center. [Courtesy of Martin Schmeing and Venki Ramakrishnan,   The (a) first, (b) second, and (c) third codon-anticodon base pairs as  seen in the X-ray structure of the T. thermophilus 30S subunit in  complex with U 6 (a model mRNA) and the 17-nt anticodon arm of  tRNA Phe (whose anticodon is GAA). The structures are drawn in  ball-and-stick form embedded in their semitransparent van der Waals    surfaces. Codons are purple, anticodons are yellow, and rRNA is  brown or gray with non-C atoms colored according to type (N blue, 0  red, and P green). Protein segments are gray and Mg 2+ ions are  represented by magenta spheres. [Courtesy of Venki Ramakrishnan,  MRC Laboratory of Molecular Biology, Cambridge, U.K. PDBid 11BM.]   interactions with three universally conserved ribosomal bases, A1492, A1493,  and G530 (Fig. 27-31):   1. The first codon-anticodon base pair, that between mRNA U1 and  tRNA A36, is stabilized by the binding of the rRNA A1493 base in  the base pairs minor groove (Fig. 27-31/2).   2. The second codon—anticodon base pair, that between U2 and A35, is  bolstered by A1492 and G530, which both bind in this base pairs mi¬  nor groove (Fig. 27-31£).   3. The third codon—anticodon base pair (the wobble pair; Section 27-2C),  that between U3 and G34, is reinforced through minor groove bind¬  ing by G530 (Fig. 27-31 c). The latter interaction appears to be less  stringent than those in the first and second codon—anticodon positions,  which is consistent with the need for the third codon—anticodon pair¬  ing to tolerate non-Watson-Crick base pairs (Section 27-2C).   Comparison of this structure with that of the 30S subunit alone reveals that  the foregoing rRNA nucleotides undergo conformational changes on the for¬  mation of a codon-anticodon complex (Fig. 27-32): In the absence of tRNA,  the bases of A1492 and A1493 stack in the interior of an RNA loop, whereas  in the codon—anticodon complex, these bases have flipped out of the loop and  the G530 base has switched from the syn to the anti conformation (Section  24-IB). These interactions enable the ribosome to monitor whether an incom¬  ing tRNA is cognate to the codon in the A site; a non-Watson—Crick base  pair could not bind these ribosomal bases in the same way. Indeed, any mu¬  tation of A1492 or A1493 is lethal because pyrimidines in these positions  could not reach far enough to interact with the codon—anticodon complex or  G530, and because a G in either position would be unable to form the re¬  quired hydrogen bonds and its N2 would be subjected to steric collisions. An  incorrect codon—anticodon interaction provides insufficient free energy to bind  the tRNA to the ribosome and it therefore dissociates from the ribosome, still  in its ternary complex with EF-Tu and GTP.    FIG. 27-32 Ribosomal decoding site. The X-ray structures of T. thermophilus 30S  subunit (a) alone and (b) in its complex with U 6 and the anticodon stem-loop of tRNA Phe .  The RNAs are drawn as ribbons with their nucleotides in paddle form with tRNA gold,  A-site mRNA purple, P-site mRNA green, rRNA gray, and nucleotides that undergo  conformational changes red. Protein SI 2 is tan and Mg 2+ ions are represented by red  spheres. [Courtesy of Venki Ramakrishnan, MRC Laboratory of Molecular Biology,  Cambridge, U.K. PDBids (a) 1 FJF and (b) 11BM.]   GTP Hydrolysis by EF-Tu Is a Thermodynamic Prerequisite to Ribosomal Section 4 Translation   Proofreading. An aa-tRNA is selected by the ribosome only according to its  anticodon, a process that has a measured error rate of only ~10 4 per residue.   Yet the binding energy loss arising from a single base mismatch in a codon-  anticodon interaction is estimated to be ~12 kj • mol \ which, according to  Eq. 27-1, cannot account for a ribosomal decoding accuracy of less than  — 10 2 errors per codon. Evidently, the ribosome has some sort of proofread¬  ing mechanism that increases its overall decoding accuracy.   A proofreading step must be entirely independent of the initial selection step.   Only then can the overall probability of error be equal to the product of the  probabilities of error of the individual selection steps. We have seen that DNA  polymerases and aminoacyl-tRNA synthetases maintain the independence of  their two selection steps by carrying them out at separate active sites (Sections  25-2A and 27-2B). Yet the ribosome recognizes the incoming aa—tRNA only  according to its anticodons complementarity to the codon in the A site.   Consequently, the ribosome must somehow examine this codon-anticodon in¬  teraction in two separate ways. How does this occur?   Cryo-EM studies by Frank indicate that in the ribosome • aa-tRNA • EF-  Tu • GTP complex, the aa-tRNA assumes an otherwise energetically unfavorable  conformation in which the anticodon arm has moved via a hinge motion toward  the inside of the tRNAs L. The formation of a correct codon—anticodon com¬  plex triggers EF-Tu to hydrolyze its bound GTP and thereby dissociate from the  aa-tRNA, which then assumes its energetically more favorable conformation  while maintaining its codon—anticodon interaction. This movement swings the  acceptor stem with its attached aminoacyl group into the peptidyl transferase cen¬  ter on the 5OS subunit (see below). A noncognate (mispaired) aa-tRNA would  presumably have insufficient strength in its codon—anticodon interaction to hold  it in place during this conformational change and would therefore dissociate from  the ribosome. EF-Tus irreversible GTPase reaction must precede this proofread¬  ing step because otherwise the dissociation of a noncognate aa-tRNA (the release  of its anticodon from the codon) would simply be the reverse of the initial bind¬  ing step; that is, it would be part of the initial selection step rather than proof¬  reading. GTP hydrolysis therefore provides the second context necessary for  proofreading; it is the entropic price the system must pay for accurate tRNA selection.   Transpeptidation: The Ribosome Is a Ribozyme. In the second (transpep-  tidation) stage of the elongation cycle (Fig. 27-27), the peptide bond is formed  through the nucleophilic displacement of the P site tRNA by the amino group  of the 3'-linked aa-tRNA in the A site (Fig. 27-21). The nascent polypeptide  chain is thereby lengthened at its C-terminus by one residue and transferred  to the A-site tRNA. The reaction occurs without the need of activating cofac¬  tors such as ATP because the ester linkage between the nascent polypeptide  and the P-site tRNA is a “high-energy” bond relative to the peptide bond that  forms during transpeptidation.   The peptidyl transferase center that catalyzes peptide bond formation is lo¬  cated on the large subunit and consists entirely ofrRNA. Numerous observations  had suggested that the ribosome is indeed a ribozyme rather than a conven¬  tional protein enzyme. For example, up to 95% of the protein can be removed  from the large subunit without abolishing its peptidyl transferase activity,  rRNAs are more highly conserved throughout evolution than are ribosomal  proteins, and most mutations that confer resistance to antibiotics that inhibit  protein synthesis occur in genes encoding rRNAs rather than ribosomal pro¬  teins. However, it was the X-ray structure of the large ribosomal subunit that  unambiguously demonstrated that rRNA functions catalytically in protein syn¬  thesis. In hindsight, this seems obvious on evolutionary grounds: Since all pro¬  teins, including those associated with ribosomes, are ribosomally synthesized,  the primordial ribosome must have preceded the primordial proteins and  hence consisted entirely of RNA.   FIG. 27-33 Model of the substrate  complex of the 50S ribosomal subunit.   Atoms are colored according to type with the  A-site substrate (Tyr-tRNA Tyr ) C purple, P-site  substrate C green, 23S rRNA C orange, N blue,  and 0 red. The attacking amino group of the  A-site aminoacyl residue is held in position for  nucleophilic attack (cyan arrow) on the  carbonyl C of the P-site aminoacyl ester through  hydrogen bonds (dashed black lines) to  A2486-N3 and the 2'-0 of the P-site A76.  [Courtesy of Peter Moore and Thomas Steitz,  Yale University.]    The ribosomes active site lies in a highly conserved region in domain V  of the 23S rRNA (Figs. 27-12 b, 27-17, and 27-30). The nearest protein side  chain is ~18 A away from the newly formed peptide bond, and the nearest  Mg 2+ ion is ~8.5 A away—both too far to be involved in catalysis. Clearly,  the ribosomal transpeptidase reaction is catalyzed solely by RNA.   The ribosomal transpeptidase reaction occurs ~10 7 -fold faster than the  uncatalyzed reaction. How does the ribosome catalyze this reaction?  Ribosomally mediated peptide bond formation proceeds via the nucleophilic  attack of the amino group on the carbonyl group of an ester to form a tetra¬  hedral intermediate that collapses to an amide and an alcohol (Fig. 27-21).  However, in the physiological pH range, the attacking amino group is pre¬  dominantly in its ammonium form (RNH^), which lacks the lone pair nec¬  essary to undertake a nucleophilic attack. This suggests that the transpeptidase  reaction is catalyzed in part by a general base that abstracts a proton from the  ammonium group to generate the required free amino group (RNH 2 ).   Inspection of the peptidyl transferase active site in H. marismortui reveals  that the only basic group within 5 A of the inferred position of the attacking  amino group is atom N3 of the invariant rRNA base A2486 (A2451 in  E. coli). It is ~3 A from and hence hydrogen bonded to the attacking amino  group (Fig. 27-33). This further suggests that the protonated A2486-N3 elec¬  trostatically stabilizes the oxyanion of the tetrahedral reaction intermediate and  then donates the proton to the leaving group of the P-site tRNA to yield a  3'-OH group (general acid catalysis). However, in order for A2486-N3 to act  as a general base in abstracting the proton from an ammonium group (whose  pZfis ~10), it must have a pK of at least 7 (recall that proton transfers be¬  tween hydrogen-bonded groups occur at physiologically significant rates only  when the p K of the proton donor is no more than 2 or 3 pH units greater  than that of the proton acceptor; Section 11-5C). Yet, the pZfof N3 in AMP  is <3.5. Moreover, the model displayed in Fig. 27-33 indicates that the tetra¬  hedral intermediates oxyanion would point away from and  hence could not be stabilized by protonated A2486-N3.    The Ribosome Is an Entropy Trap. The resolution to this co¬  nundrum was provided by Marina Rodnina and Richard  Wolfenden, who noted that the uncatalyzed reaction of esters  with amines to form amides occurs quite facilely in aqueous so¬  lution. They therefore measured the rates of both uncatalyzed  peptide bond formation by model compounds and peptidyl  transfer by the ribosome at several different temperatures. This  provided values of AA//* at and AAS* at , the reactions change in  the enthalpy and entropy of activation by the ribosome relative  to the uncatalyzed reaction. Here A A H\„ x — TAASf^ t =    change in the reactions free energy of activation by the ribo¬  some relative to the uncatalyzed reaction, and AGquncat) and  AG*(cat) are the free energies of activation of the uncatalyzed  and catalyzed (ribosomal) reactions (Sections 1-3D and 11-2). The measured  value of AAZ/* at is —19 kj • mol \ a quantity that would be positive, not  negative, if the ribosomal reaction had a significant component of chemical  catalysis such as acid-base catalysis and/or the formation of new hydrogen  bonds. In contrast, the value of 7AAS* at is +52 kj • mol 1 , which indicates  that the Michaelis complex (E • S; Section 12-IB) in the ribosomal reaction  is significantly more ordered relative to the transition state than is the uncat¬  alyzed reaction. This value of 7AAS* at largely accounts for the observed ~10 7 -  fold rate enhancement of the ribosomal reaction relative to the uncatalyzed  reaction (the rate enhancement by the ribosome is given by ^ AAGcat/i?7 ; Section  11-2). Evidently, the ribosome enhances the rate of peptide bond formation by  properly positioning and orienting its substrates and/or excluding water from the   FIG. 27-34 The mechanism of ribosome-catalyzed peptidyl  transfer. The nucleophilic attack of the a-amino group of the  aminoacyl-tRNA (red) on the carbonyl C of the peptidyl-tRNA (blue)  occurs in concert with a proton shuttle that involves the 03' and  2'-0H of the P-site A76 together with the a-amino group of the    aminoacyl-tRNA. The reaction proceeds through a transition state  (center; enclosed by square brackets) that contains a six-membered  ring of partially bonded atoms and which collapses to the reaction  products drawn on the right.    preorganized electrostatic environment of the active site (a form of reactant order¬  ing) rather than by chemical catalysis .   The X-ray structures of the large ribosomal subunit in complex with  aminoacyl-tRNA and peptidyl-tRNA indicate that the ribosome uses an in¬  duced fit mechanism, as occurs in enzymes such as hexokinase (Section 15-2A).  Conformational changes in the 23S rRNA, presumably triggered by proper  binding of the aminoacyl-tRNA in the A site, orient the ester group of the  peptidyl—tRNA for nucleophilic attack. The hydrogen bond between the 2'-  OH of the P-site A76 and the attacking amino group (Fig. 27-33) is crucial  in doing so, as is indicated by the observations that replacing this 2'-OH group  with H or F reduces the reaction rate by at least 10 6 . This suggests that the  peptidyl transferase reaction occurs via the substrate-assisted proton shuttle  mechanism diagrammed in Fig. 27-34.   In the absence of a tRNA in the A site, the peptidyl-tRNA is held in a  position that prevents nucleophilic attack by water (hydrolysis), which would  otherwise release the peptidyl group from the ribosome.   Translocation: The Ribosome Moves to the Next Codon. Following transpep-  tidation, the now uncharged P-site tRNA moves to the E site (not shown in  Fig. 27-27), and the new peptidyl-tRNA in the A site, together with its bound  mRNA, moves to the P site. These events, which leave the A site vacant, pre¬  pare the ribosome for the next elongation cycle. During translocation, the pep¬  tidyl—tRNAs interaction with the codon, which is no longer necessary for amino  acid specification, acts as a placekeeper so that the ribosome can advance by ex¬  actly three nucleotides along the mRNA, as required to preserve the reading  frame. An Mg 2+ -stabilized kink in the mRNA between the A and P codons  may help prevent slipping. Indeed, certain mutant tRNA molecules that cause  frameshifting induce the ribosome to translocate by four nucleotides, thereby  demonstrating that mRNA movement is directly coupled to tRNA movement.   The translocation process requires an elongation factor, EF-G in E. coli,  that binds to the ribosome together with GTP and is released only on hydrol¬  ysis of the GTP to GDP + P z (Fig. 27-27). EF-G release is prerequisite for  beginning the next elongation cycle because the ribosomal binding sites of  EF-G and EF-Tu partially or completely overlap and hence their ribosomal  binding is mutually exclusive. GTP hydrolysis, which precedes translocation,  provides free energy for tRNA movement.   FIG. 27-35 X-Ray structure of EF-G from T. thermophilus in complex  with GMPPNP. The protein is drawn in ribbon form colored in rainbow order from its  N-terminus (blue) to its C-terminus (red). The GMPPNP is drawn in space-filling form  colored according to atom type with C yellow, N blue, 0 red, and P orange. An Mg 2+ ion  that is bound to the GMPPNP is represented by a magenta sphere. Portions of the  structure are not visible. Note the remarkable resemblance in shape between this structure  and that of Phe-tRNA Phe • EF-Tu • GMPPNP (Fig. 27-28). [Based on an X-ray structure  by Anders Liljas and Derek Logan, Lund University, Lund, Sweden. PDBid 2BV3.]    EF-G Structurally Mimics the EF-Tu • tRNA Complex. The X-ray structure  of EF-G (Fig. 27-35) reveals that the protein has an elongated shape com¬  prising five domains, the first two of which (blue through green in Fig.  27-35) are arranged similarly to the first two domains of the EF-Tu •  GMPPNP complex (Fig. 27-28). Most intriguingly, the remaining three do¬  mains of EF-G (yellow, orange, and red in Fig. 27-35) have a conformation  reminiscent of the shape of tRNA bound to EF-Tu. Such molecular mimicry  allows EF-G to drive translocation not just by inducing a conformational  change in the ribosome but by actively displacing the peptidyl—tRNA from  the A site. The X-ray structure of the ribosome with bound tRNAs and  EF-G supports this model (Fig. 27-36). In this complex, EF-G has hydrolyzed  its GTP and translocation has occurred, but EF-G cannot dissociate from the  ribosome due to the presence of the antibiotic fusidic acid. The EF-G occupies  a position similar to that of the EF-Tu • tRNA complex bound to the ribosome  (Fig. 27-30). The structural similarity between EF-G domains 3 to 5 and tRNA  also suggests that in the earliest cells, whose functions were based on RNA,  proteins evolved by mimicking shapes already used successfully by RNA.    Translocation Occurs Via Intermediate States. Variations in chemical foot¬  printing patterns during the elongation cycle, together with X-ray and cryo-  EM studies, indicate that the translocation of tRNA through the ribosome  occurs in several discrete steps (Fig. 27-37):   1. In the posttranslocational state, a deacylated tRNA occupies the E  site of both the 30S and 50S subunits (the E/E state), a peptidyl-  tRNA occupies both P sites (the P/P state), and the A site is vacant.  An aa-tRNA in complex with EF-Tu • GTP binds to the ribosome.  This yields a complex in which the incoming aa—tRNA is bound in  the 30S subunits A site via a codon—anticodon interaction (recall  that the mRNA is bound to the 30S subunit) but with the  EF-Tu • GTP preventing the entry of the tRNAs aminoacyl  end into the 5OS subunits A site, an arrangement termed the  A/T state (T for EF-7u). This structure is also pictured in  Fig. 27-30.   2. As we discussed above, EF-Tu hydrolyzes its bound GTP to  GDP + P z - and is released from the ribosome, thereby permit¬  ting the aa-tRNA to fully bind to the A site (the A/A state)  and releasing the E-site tRNA.    FIG. 27-36 X-Ray structure of the T. thermophilus ribosome with  bound EF-G. The structure is drawn and labeled as in Fig. 27-30 with  EF-G shown as a red surface model. [Courtesy of Martin Schmeing and  Venki Ramakrishnan, MRC Laboratory of Molecular Biology, Cambridge,  U.K. PDBids 2WRI, 2WRJ, 2WRK, and 2WRL]    4. The acceptor end of the new peptidyl-tRNA shifts to the P site of the  5OS subunit, while the tRNAs anticodon end remains associated with  the A site of the 30S subunit (yielding the A/P state). The acceptor  end of the newly deacylated tRNA simultaneously moves to the E site  of the 5OS subunit while its anticodon end remains associated with the  P site of the 30S subunit (the P/E state).   5. EF-G • GTP binds to the ribosome and the resulting GTP hydrolysis  impels the anticodon ends of the two tRNAs, together with their  bound mRNA, to move relative to the small ribosomal subunit such  that the peptidyl-tRNA assumes the P/P state and the deacylated  tRNA assumes the E/E state (the posttranslocational state), thereby  completing the elongation cycle.    FIG. 27-37 Ribosomal binding states in the  elongation cycle. Note how this scheme elaborates  the classical elongation cycle diagrammed in Fig.  27-27. The drawings are accompanied by 17-A-  resolution cryo-EM-based images of the E. coli 70S  ribosome in the corresponding binding states in  which the 30S subunit is transparent yellow, the  50S subunit is transparent blue, and the tRNAs and  elongation factors are colored as in the drawing  they accompany. [Cryo-EM images courtesy of Knud  Nierhaus, Max-Planck-lnstitut fur Molekulare  Genetik, Berlin, Germany, and Joachim Frank, State  University of New York at Albany.]    The majority of known antibiotics, including a great variety of  medically useful substances, block translation. This situation is  presumably a consequence of the translation machinery’s enor¬  mous complexity, which makes it vulnerable to disruption in many  ways. Antibiotics have also been useful in analyzing ribosomal  mechanisms because the blockade of a specific function often  permits its biochemical dissection into its component steps. For  example, the ribosomal elongation cycle was originally character¬  ized through the use of the antibiotic puromycin, which resem¬  bles the 3' end of Tyr-tRNA:    peptidyl-puromycin in which puromycin’s “amino acid residue” is  linked to its “tRNA” via an amide rather than an ester bond. The  ribosome therefore cannot catalyze further transpeptidation, and  polypeptide synthesis is aborted.   Streptomycin is a medically important member of a family of  antibiotics known as aminoglycosides that inhibit prokaryotic  ribosomes in a variety of ways.    Puromycin binds to the ribosomal A site without the need of  elongation factors. The transpeptidation reaction yields a   At low concentrations, streptomycin induces the ribosome to char¬  acteristically misread mRNA: One pyrimidine may be mistaken for    The binding of tRNAs to the A and E sites of the ribosome, as Knud  Nierhaus has shown, exhibits negative allosteric cooperativity. In the pre-  translocational state, the E site binds the newly deacylated tRNA with high  affinity, whereas the now empty A site has low affinity for aa-tRNA. The bind¬  ing of a new aa-tRNA • EF-Tu • GTP compl ex induces the ribosome to un¬  dergo a conformational change that converts the A site to a high-affinity state  and the E site to a low-affinity state that consequently releases the deacylated  tRNA. Thus, the E site is not simply a passive holding site for spent tRNAs  but performs an essential function in the translation process.   The requirement that the GTP-hydrolyzing proteins EF-Tu and EF-G alter¬  nate in their activities ensures that the ribosome cycles unidirectionally through the  transpeptidation and translocation stages of translation. Translocation can occur  in the absence of GTP, which indicates that the free energy of the transpep¬  tidation reaction is sufficient to drive the entire translational process. However,  the GTP hydrolysis catalyzed by EF-Tu and EF-G increases the overall rate of  translation, presumably by reducing the activation barriers between successive  states. Because GTP hydrolysis is irreversible , the accompanying conformational  changes in the ribosome are also irreversible and hence unidirectional    the other in the first and second codon positions, and either  pyrimidine may be mistaken for adenine in the first position. This  inhibits the growth of susceptible cells but does not kill them. At  higher concentrations, however, streptomycin prevents proper chain  initiation and thereby causes cell death.   the first of the “broad-spectrum” antibiotics, inhibits the peptidyl  transferase activity of prokaryotic ribosomes. However, its clinical  uses are limited to severe infections because of its toxic side  effects, which are caused in part by the chloramphenicol sensitivity  of mitochondrial ribosomes. Chloramphenicol binds to the large  subunit near the A site, which explains why it competes for bind¬  ing with puromycin and the 3' end of aminoacyl-tRNA but not  with peptidyl-tRNAs.   and its derivatives are broad-spectrum antibiotics that bind to the  small subunit of prokaryotic ribosomes. Tetracycline binding pre¬  vents the entry of aminoacyl-tRNAs into the A site but allows  EF-Tu to hydrolyze its GTP. As a result, protein synthesis cannot  proceed, and GTP hydrolysis, which occurs every time another  aminoacyl-tRNA attempts to enter the ribosome, presents an  enormous energetic drain on the cell.   Tetracycline-resistant bacterial strains have become quite  common, thereby precipitating a serious clinical problem. Most  often resistance is conferred by a decrease in bacterial cell  membrane permeability to tetracycline rather than any alteration of  ribosomal components that would overcome the inhibitory effect  on translation.   Although it is a protein rather than a small molecule, the en¬  zyme ricin can be considered an antibiotic. Ricin is an A/-glycosi-  dase from castor beans, the source of castor oil. The enzyme  inactivates the large subunit of eukaryotic ribosomes by hydrolyti¬  cally removing the adenine base of a highly conserved residue of  28S rRNA. The modified ribosome is unable to bind elongation  factors, and translation ceases. Because it acts catalytically rather  than stoichiometrically, a single ricin molecule can inactivate tens  of thousands of ribosomes. Its deadliness in minute amounts  makes ricin attractive to bioterrorists, but methods for selectively  targeting ricin to particular cells, such as cancer cells, raise the  possibility of its therapeutic use.   The changes in binding states result in large-scale tRNA movements, in  some instances >50 A. Moreover, cryo-EM studies indicate that on binding  EF-G • GDP(CH 2 )P (like GMPPNP but with a CH 2 group rather than an  NH group bridging its (3 and y phosphates), the 30S subunit rotates with re¬  spect to the 5OS subunit by 6° clockwise when viewed from the 30S subunits  solvent side, which results in a maximum displacement of ~ 1 9 A at the pe¬  riphery of the ribosome. This rotation is accompanied by many smaller con¬  formational changes in both subunits, particularly in the regions about the  entrance and exit to the mRNA channel. Clearly, we are far from fully under¬  standing how the ribosome works at the molecular level. Several of the many  antibiotics that disrupt ribosomal functioning are discussed in Box 27-3.   Elongation in eukaryotes closely resembles that in prokaryotes. In eukaryotes,  the functions of EF-Tu and EF-Ts are assumed by the eukaryotic elongation  factors eEFlA and eEFIB. Likewise, eEF2 functions in a manner analogous  to prokaryotic EF-G. However, the corresponding eukaryotic and prokaryotic  elongation factors are not interchangeable.   EF-G hydrolyzes its bound GTP to GDP + P,-,  which expels the bound tRNAs. RRF, the mRNA, /g  and EF-G • GDP depart, causing the ribosomal  subunits to dissociate, ready for reinitiation.   RF-1 (or RF-2)  recognizes the  corresponding Stop  codon in the A site.    The peptidyl-tRNA is  2 k hydrolyzed and the  polypeptide is released.  RF-3 • GDP binds.   RF-3 exchanges its  bound GDP for GTP  and RF-1 (or RF-2) is  released.    RF-3 hydrolyzes its bound  GTP to GDP + Pj, both of  which are released. RRF  binds in the A site  followed by EF-G • GTP.    FIG. 27-38 The translation termination pathway in E. coli  ribosomes. RF-1 recognizes the Stop codons UAA and UAG,  whereas RF-2 (not shown) recognizes UAA and UGA. Eukaryotic    termination follows an analogous pathway but requires only a single  release factor, eRFI, that recognizes all three Stop codons.    Polypeptide synthesis under the direction of synthetic mRNAs such as poly(U)  results in a peptidyl-tRNA “stuck” in the ribosome. However, the translation  of natural mRNAs, which contain the Stop codons UAA, UGA, or UAG,  yields free polypeptides. In 77. coli, the Stop codons, which normally have no  corresponding tRNAs, are recognized by protein release factors (Table 27-6):  RF-1 recognizes UAA and UAG, whereas the 39% identical RF-2 recognizes  UAA and UGA. In eukaryotes, a single release factor, eRFl, recognizes all  three Stop codons.   Termination, like initiation and elongation, has several stages. The se¬  quence of events in E. coli is diagrammed in Fig. 27-38:   1. RF-1 or RF-2 recognizes a corresponding Stop codon in the ribosomes  A site.   2. The release factor induces the transfer of the peptidyl group from the  peptidyl-tRNA to water, rather than to another aa-tRNA, to yield an  uncharged tRNA in the P site and a free polypeptide that dissociates  from the ribosome.   3. A third factor, RF-3 (eRF in eukaryotes), a G protein, binds to the ri¬  bosome in complex with GDP. RF-3 binds to the same site on the ri¬  bosome as do EF-Tu and EF-G. In fact, the X-ray structure of RF-3 •  GDP resembles that of EF-Tu • GMPPNP (Fig. 27-28). Free RF-3 has  a greater affinity for GDP than GTP but on binding to the ribo-  some-RF-1/2 complex, it exchanges its bound GDP for GTP. The re¬  sulting change in the conformation of RF-3, as seen in cryo-EM  studies, causes it to bind more tightly to the ribosome and expel the  RF-1/2. RF-3 is not required for cell viability although it is necessary  for maximum growth rate; RF-3 only accelerates the dissociation of  RF-1/2 from the ribosome by ~ 5-fold.   4. The interaction of RF-3 * GTP with the ribosome stimulates it to hy¬  drolyze its bound GTP, much as occurs with EF-Tu • GTP and EF-G •  GTP. The resulting RF-3 • GDP then dissociates from the ribosome.  Subsequently, ribosomal recycling factor (RRF) binds in the riboso-  mal A site followed by EF-G • GTP. RRF, which was discovered by  Akira Kaji, is essential for cell viability.   5. EF-G hydrolyzes its bound GTP, which causes RRF to move to the P  site and the tRNAs occupying the P and E sites (the latter not shown in  Fig. 27-38) to be released. Finally, the small and large ribosomal subunits  separate, a process that is facilitated by the binding of IF-3 (Section 27-  4A), and RRF, EF-G • GDP, and mRNA are released. The ribosomal sub¬  units can then participate in a new round of initiation (Fig. 27-25).   RRF Binds in the Ribosomal A Site. The X-ray structure of 77 thermophilus  ribosomal recycling factor (RRF; Table 27-6), determined by Yoshikazu  Nakamura, reveals it to be a two-domain structure that resembles tRNA in its  overall shape (Fig. 27-39). The comparison of this structure with those of sev¬  eral other bacterial RRFs indicates that the two linkers connecting the RRF  domains are flexible such that domain II can rotate about the axis of the three-  helix bundle forming domain I.   The X-ray structure of the 77 thermophilus ribosome in complex with RRF  in its A site, the anticodon stem-loop (ASL) of tRNA Phe in its P site, tRNAf 4 ^    FIG. 27-39 X-Ray structure of T. thermophilus RRF. This monomeric protein is  drawn in ribbon form colored in rainbow order from its N-terminus (blue) to its C-terminus  (red). [Based on an X-ray structure by Yoshikazu Nakamura, The University of Tokyo, Japan.  PDBid 1EH1J   FIG. 27-40 X-Ray structure of the T.  thermophilus ribosome in complex with RRF.   The RRF and an mRNA with a UAG Stop codon  are in its A site, the anticodon stem-loop (ASL)  of tRNA Phe is in its P site, and tRNA f Met is in its  E site. The ribosomal RNAs are shown as  semitransparent surface diagrams with 23S RNA  cyan, 5S RNA blue-green, and 16S RNA  yellow. The ribosomal proteins are drawn in  ribbon form with 50S subunit proteins blue and  30S subunit proteins orange. The RRF, tRNAs,  and mRNA are represented by their surface  diagrams with domains I and II of RRF red and  blue, the mRNA magenta, the P-site ASL purple,  and the E-site tRNA f Met gray. [Courtesy of Venki  Ramakrishnan, MRC Laboratory of Molecular  Biology, Cambridge, UK. PDBids 2V46 and 2V47.]   • How was it determined that peptides  are synthesized from their N-terminus  to their C-terminus?   • Compare prokaryotic and eukaryotic  translation initiation with respect to  delivering an initiator tRNA to the  ribosome and locating the initiation codon.   • Summarize the roles of initiation, elonga¬  tion, and release factors in translation.   • Summarize the role of GTP hydrolysis in  promoting the efficiency of translation  initiation, decoding, translocation, and  chain termination.   • What mechanism ensures that the  ribosome translocates by exactly three  nucleotides?    in its E site, and an mRNA with a UAG Stop codon in the A site was deter¬  mined by Ramakrishnan (Fig. 27-40). Domain I of the RRF spans the A and  P sites of the 5OS ribosome, a position in which the tip of its domain I would  clash with a tRNA in the P site. This suggests that RRF binding forces a tRNA  bound in the P site into the P/E hybrid binding state (Section 27-4B).   Nonsense Suppressors Prevent Termination. A mutation that converts an  aminoacyl-coding (sense) codon to a Stop codon is known as a nonsense mu¬  tation and leads to the premature termination of translation. An organism  with such a mutation may be “rescued” by a second mutation in a tRNA gene  that causes the tRNA to recognize a Stop (nonsense) codon. This nonsense  suppressor tRNA carries the same amino acid as its wild-type progenitor and  appends it to the growing polypeptide at the Stop codon, thereby preventing  chain termination. For example, the E. coli suppressor known as su3 is a  tRNA Tyr whose anticodon has mutated from the wild-type GUA (which reads  the Tyr codons UAU and UAC) to CUA (which recognizes the Stop codon  UAG). An su3 + E. coli cell with an otherwise lethal mutation to UAG in a  gene coding for an essential protein would be viable if the replacement of the  wild-type amino acid residue by Tyr does not inactivate the protein.   How do cells tolerate a mutation that both eliminates a normal tRNA and  prevents the termination of polypeptide synthesis? They survive because the  mutated tRNA is usually a minor member of a set of isoaccepting tRNAs and  because nonsense suppressor tRNAs must compete with release factors for  binding to Stop codons. Consequently, many suppressor-rescued mutants grow  more slowly than wild-type cells.    • Chaperones bind to polypeptide chains as they emerge from the ribosome.   • The polypeptide chain may be covalently modified and translocated via the secre¬  tory pathway.    The tunnel that leads from the peptidyl transferase active site to the exterior of  the bacterial ribosome (Fig. 27-13^) is --TOO A long, enough to shelter a polypep¬  tide of ~30 residues. Once it emerges from the ribosome, the polypeptide   destination. Both processes occur with the aid of other proteins. In addition,  a nascent polypeptide may be covalently modified before it achieves its mature  form. In this section we review some of the events of posttranslational  protein processing.   The ribosome’s peptide exit tunnel is too narrow (~15 A wide) to permit the  formation of secondary structure other than perhaps helices, so a nascent  polypeptide cannot begin to fold until after it emerges from the ribosome.  However, folding commences well before translation is complete. Molecular  chaperones (Section 6-5B) bind to the N-terminus of nascent polypeptides to  prevent their aggregation, to facilitate their folding, and to promote their  proper association with other subunits.   A growing body of evidence suggests that ribosomal proteins play a role  in protein folding by recruiting chaperones. For example, in E. coli, a protein  known as trigger factor associates with the ribosomal protein L23, which is  located at the outlet of the peptide exit tunnel. Trigger factor recognizes rela¬  tively short hydrophobic protein segments when they first emerge from the ri¬  bosome. DnaK, a member of the Hsp70 family of chaperones, together with  DnaJ, an Hsp40 cochaperonin (Section 6-5B), can bind to and protect longer  polypeptides (DnaK and DnaJ were so named because they were discovered  through the isolation of mutants that do not support the growth of bacterio¬  phage X and hence were initially assumed to participate in DNA replication).  The activities of trigger factor and DnaK/J are somewhat redundant, as E. coli  can tolerate the deletion of either one. The loss of both, however, is lethal  above 30°C and is accompanied by the massive aggregation of newly synthe¬  sized proteins. Ultimately, the chaperonins GroEL and GroES facilitate pro¬  tein folding after translation termination (Section 6-5B).   The X-ray structure of E. coli trigger factor reveals an elongated multido¬  main protein that has a pronounced hydrophobic cleft along its central region.  The X-ray structure of trigger factors 144-residue ribosome-binding N-termi-  nal domain in complex with the 5OS subunit from H. marismortui reveals that  this domain binds near the exit of the ribosomes peptide tunnel. The super¬  position of this N-terminal domain with that in the X-ray structure of the in¬  tact 432-residue trigger factor generated a model of the interaction of trigger  factor with the 5OS subunit (Fig. 27-41).   In this model, trigger factor hunches over the exit of the ribosomes pep¬  tide tunnel with its hydrophobic concave face positioned to interact with a  newly synthesized polypeptide segment as it emerges from the tunnel.  Moreover, fluorescence energy transfer experiments show that on binding to  the ribosome, trigger factor undergoes a conformational expansion that fur¬  ther exposes its hydrophobic surface.   Unlike many other chaperones (e.g., DnaK/J and GroEL/ES), trigger factor  does not require ATP for its activation. Activated trigger factor dissociates from  the ribosome after ~10 s but may remain associated with the elongating  polypeptide for >30 s, during which time an additional molecule of trigger  factor can bind to the ribosome and shield hydrophobic sequences farther    FIG. 27-41 Model of the association of trigger factor with the 50S ribosomal  subunit. The 50S subunit is shown in a cutaway view that exposes its peptide exit tunnel  into which an a helix ( magenta ) extending from the peptidyl transferase center (PT) has  been modeled. The ribosomal RNA is pink and the ribosomal proteins are gray. Trigger  factor is drawn in worm form with its four domains colored, from N- to C-terminus, red,  yellow, green, and blue. [Courtesy of Nenad Ban, Eidgenossische Technische Hochschule,  Zurich, Switzerland.]    FIG. 27-42 Conversion of proinsulin to  insulin. The prohormone, with three disulfide  bonds, is proteolyzed at two sites ( arrows ) to  eliminate the C chain. The mature hormone  insulin consists of the disulfide-linked A and B  chains.    along the polypeptide chain. Apparently, trigger factor functions to shield hy¬  drophobic segments of newly synthesized and hence unfolded polypeptides so as to  prevent their misfolding and aggregation. Subsequently, many of the resulting par¬  tially folded proteins are handed off to other chaperones, such as GroEL/ES,  to complete the folding process. Eukaryotic cells lack a homolog of trigger  factor but contain other small chaperones that may have similar functions.   The list of known posttranslational modifications is long and includes the re¬  moval and/or derivatization of specific residues. For example, a polypeptides  N-terminal Met or fMet residue is usually excised. Other common posttrans¬  lational modifications are the hydroxylation of Pro and Lys residues in colla¬  gen (Section 6-1C), glycosylation (Sections 8-3C and 16-5), and the  prenylation and fatty acylation of membrane-anchored proteins (Section 9-3B).  Many covalent modifications, such as phosphorylation (Section 12-3B) and  palmitoylation (Section 9-3B), are reversible. Over 150 different types of side  chain modifications are known; these involve all side chains except those of  Ala, lie, Leu, and Val. The functions of such modifications are varied and in  many cases remain enigmatic.   A growing number of proteins are known to be ubiquitinated. Recall from  Section 21-IB that the attachment of the 76-residue protein ubiquitin marks  a protein for degradation by the proteasome. A homologous 97-residue pro¬  tein known as small ubiquitin-related modifier (SUMO) can be ligated to  a Lys residue, as is ubiquitin, to regulate protein function. But whereas ubiq-  uitination is usually associated with protein degradation, sumoylation appears  to play a role in determining protein localization inside cells.   Proteins that are synthesized as inactive precursors, called proproteins or  proenzymes, are activated by limited proteolysis. The zymogens of serine pro¬  teases are converted to active enzymes in that fashion (Section 11-5D). Proteins  that are translocated into the endoplasmic reticulum for export from the cell  typically contain a signal peptide that must be excised (Section 9-4D). Proteins  bearing a signal peptide are known as preproteins, or preproproteins if they  undergo additional proteolysis during their maturation. For example, the hor¬  mone insulin (Section 22-2) is synthesized as a preproprotein, which is con¬  verted to the proprotein proinsulin, a single 84-residue polypeptide. Proteolysis  at two sites generates the mature hormone, whose A and B chains remain linked  by disulfide bonds (Fig. 27-42). The 33-residue C chain is discarded.   The Signal Recognition Particle Is a Ribonucleoprotein. Many transmem¬  brane and secretory proteins are translocated into or completely through cel¬  lular membranes [the endoplasmic reticulum (ER) in eukaryotes and the  plasma membrane in prokaryotes] via a series of events known as the secre¬  tory pathway (Section 9-4D). As their N-terminal signal sequences emerge  from the ribosome, these proteins associate with the signal recognition parti¬  cle (SRP). Peptide chain elongation ceases on SRP binding to the ribosome   Its various double-helical segments (denoted HI through H8) and loops  (denoted LI.2 and L2), are drawn in red and yellow with Watson-Crick  base pairs represented by connecting lines and non-Watson-Crick base  pairs indicated by dots. The positions at which the various SRP proteins  bind to the 7S RNA are indicated in cyan, purple, and gray. [Courtesy of  Roland Beckmann, Ludwig-Maximilians-Universitat Munchen, Munich, Germany.]    but resumes after the SRP docks with its receptor on the ER membrane. The  SRP then dissociates from the ribosome, allowing the nascent polypeptide to  pass into or through the membrane via a protein pore called the translocon.   Like the ribosome, the spliceosome (Section 26-3B), and RNase P (Section  26-3D), the SRP is a ribonucleoprotein. Mammalian SRPs consist of six polypep¬  tides known as SRP9, SRP14, SRP19, SRP54, SRP68, and SRP72 (where the  numbers are their molecular masses in kD) and a 7S RNA (~300 nt; Fig. 27-43).  Many prokaryotic SRPs are much simpler; that in E. coli consists of a single  polypeptide named Ffh that is homologous to SRP54 (Ffh for Aifty-/bur  Aomolog) and a 4.5S RNA (114 nt) that, in part, is predicted  to have a secondary structure similar to that portion of the 7S  RNA to which SRP54 binds. Indeed, replacing SRP54 with  Ffh or vice versa yields functional SRPs, at least in vitro ,  thereby suggesting that the Ffh—4.5S RNA complex is a struc¬  turally minimized version of the eukaryotic SRP.   Roland Beckmann and Frank generated a model of the  ribosome—SRP complex by fitting the X-ray structures of  the yeast ribosome (Fig. 27-19) and several SRP fragments  to its 12-A resolution cryo-EM-based image (Fig. 27-44^).   The model indicates that the so-called S (signal  sequence-binding) domain of the ~270-A-long SRP RNA  binds at the base of the large ribosomal subunit next to  the exit of the tunnel through which the newly synthesized  polypeptide emerges, whereas its Alu domain (so called be¬  cause it contains A/z/-like sequences; Section 28-1C) wraps  around the large subunit to contact the ribosome at the in¬  terface between its large and small subunits. The model in¬  dicates that the 7S RNA consists mainly of a long  double-helical rod that is bent at two positions, hinge 1 and  hinge 2 (Fig. 27-44 £).    FIG. 27-44 Cryo-EM-based image of the wheat germ ribosome  in complex with canine SRP at 12 A resolution, (a) Surface diagram  showing the small ribosomal subunit in yellow, the large ribosomal subunit  in blue, the SRP in red, and a tRNA occupying the ribosomal P site in  green. The SRP’s S domain interacts with the polypeptide emerging from  the ribosome’s exit channel, and its Alu domain binds to the ribosome at  the elongation factor-binding site. ( b ) Molecular model of the SRP The  cryo-EM-based electron density is shown in transparent white and the  ribbon diagrams of the X-ray structures of SRP proteins and RNA  fragments that have been docked into it are colored as is indicated in Fig.   27-43. The signal sequence, modeled as an a helix, is green. Note that no  atomic resolution structure of the SRP68/72 heterodimer is available.    FIG. 27-45 Cryo-EM-based image of an  actively translating yeast ribosome-Sec61  complex at 9 A resolution. The structure is  represented by its surface diagram cut away  through the ribosome’s polypeptide exit tunnel.  The ribosome’s 60S subunit is cyan, the P-site  tRNA and its attached nascent chain (NC) are  green, and Sec61 is red. [Courtesy of Roland  Beckmann, Ludwig-Maximilians-Universitat  Munchen, Munich, Germany. PDBid 2WW9.]    SRP54 consists of three domains: its N-terminal N domain; its central G  domain, which contains the SRP s GTPase function; and its C-terminal M do¬  main [so called because it is rich in Met residues (25 of its 209 residues in hu¬  mans)] . The N domain, a bundle of four antiparallel a helices, closely associates  with the Ras-like G domain to form an assembly named SRP54NG that  mediates the SRP s interaction with the SRP receptor. The M domain (SRP54M),  which contacts the ribosome near the mouth of its peptide exit tunnel, con¬  tains a deep groove that binds the helical signal sequence. The groove is lined  almost entirely with hydrophobic residues including many of SRP54s Met  residues (the Met side chain has physical properties similar to that of an 72-  butyl group; Section 4-1C). Its flexible unbranched Met side chain “bristles”  presumably provide the groove with the plasticity to bind a variety of different  signal sequences so long as they are hydrophobic and form an a helix.   Structural and biochemical studies indicate that the eukaryotic SRP first  binds to the ribosome via its SRP54 protein. This interaction involves several  ribosomal proteins, including L23, to which trigger factor also binds (Section  27-5A). When an emerging signal peptide binds to SRP54M, the SRP alters  its conformation in a way that allows its Alu domain to fit into the site on the  ribosomes intersubunit interface where elongation factor eEF2 (the eukaryotic  counterpart of E. coli EF-G) also binds. This presumably interferes with the ri¬  bosomal elongation cycle (Fig. 27-27) and explains how the SRP arrests trans¬  lation. This hypothesis is corroborated by the observation that bacterial SRP s,  which do not arrest translation on binding to a ribosome, lack Alu domains.   Protein synthesis resumes after the SRP docks with its transmembrane re¬  ceptor and its place has been taken by the translocon, whose ribosomal bind¬  ing site largely overlaps that of the SRP. The central component of the  translocon, named Sec61 (Sec for ^cretion) in eukaryotes and the SecY com¬  plex in prokaryotes, is a heterotrimeric protein. Its a and y subunits, but not  its (3 subunit, are essential for channel function and are conserved across all  kingdoms of life (these subunits are respectively named Sec61a, Sec61(3, and  Sec61y in eukaryotes and SecY, SecE, and SecG in bacteria; the X-ray struc¬  ture of SecY is shown in Fig. 9-38). A cryo-EM-based structure of an actively  translating yeast ribosome-Sec61 complex (Fig. 27-45), determined by  Elisabet Mandon and Beckmann at 9 A resolution, reveals, as expected, that  a single Sec61 channel is positioned over the ribosomes polypeptide exit tun¬  nel, with the nascent polypeptide occupying both the ribosomes polypeptide  exit tunnel and Sec6Ts protein conducting channel.   partially processed glycoprotein bearing one terminal glucose residue  can bind to the chaperones calnexin or calreticulin. The bound  glycoprotein is then deglucosylated and processed further. An    improperly folded glycoprotein that becomes deglucosylated is  reglucosylated by an enzyme that is specific for unfolded  glycoproteins. This mechanism promotes the proper folding and  oligosaccharide processing of newly synthesized glycoproteins.    Glycosylation Acts as a Quality Control Mechanism. In eukaryotes,  polypeptides that are translocated into the ER by the secretory pathway un¬  dergo glycosylation. The cotranslational addition of a 14-residue oligosaccha¬  ride to an Asn residue is the first step in the production of TV-linked  glycoproteins (Section 8-3C). The attachment of the oligosaccharide verifies  that the protein is being successfully translocated from the cytosol to the lu¬  men of the endoplasmic reticulum. The subsequent removal of the two termi¬  nal glucose residues allows the protein to interact with a chaperone that recognizes  the partially processed oligosaccharide. The membrane-bound 572-residue  form of the chaperone, called calnexin, or its soluble 400-residue homolog  calreticulin, binds the immature (unfolded) glycoprotein to assist its folding  and to protect it from degradation or premature transfer to the Golgi apparatus  (where the late stages of oligosaccharide processing take place; Section 9-4E).   After the folded glycoprotein is released from calnexin/calreticulin, it pro¬  ceeds through the rest of the oligosaccharide-processing pathway. A glycopro¬  tein that is released before it has adopted its mature conformation can still  undergo the next step of processing, which is the removal of another glucose  residue. However, when this happens, a glycosyltransferase that recognizes only  unfolded glycoproteins reattaches a glucose residue to the immature oligosac¬  charide. As a result, the glycoprotein can bind again to calnexin/calreticulin  for another chance to fold properly (Fig. 27-46). Most glycoproteins undergo  reglucosylation at least once.   • What is the function of trigger factor?  Why must the number of trigger factor  molecules be at least as great as the  number of ribosomes?   • Describe the functions of the complexes  that interact with the ribosome near its  polypeptide exit channel.   • What is the function of the RNA  component of the SRP?   • The genetic code, by which nucleic acid sequences are translated  into amino acid sequences, is composed of three-nucleotide  codons that do not overlap and are read sequentially by the pro¬  tein-synthesizing machinery.   • The standard genetic code of 64 codons includes numerous syn¬  onyms, three Stop codons, and one initiation codon.   • All tRNAs have numerous chemically modified bases and a simi¬  lar cloverleaf secondary structure comprising an acceptor stem, D  arm, Tv|;C arm, anticodon arm, and variable arm. The three-  dimensional structures of tRNAs are likewise similar and are main¬  tained by stacking interactions and non-Watson-Crick hydrogen-  bonded cross-links.    • Aminoacyl-tRNA synthetases (aaRSs) catalyze the ATP- depend¬  ent attachment of an amino acid to the appropriate tRNA to yield  an aminoacyl-tRNA (aa-tRNA). The acceptor stem and  anticodon loop are common identity elements for tRNA-aaRS  interactions. The fidelity of aminoacylation is enhanced by  proofreading.   • Wobble pairing between mRNA codons and tRNA anticodons at  the third position accounts for much of the degeneracy of the ge¬  netic code. Within a species, the preferred codons correspond to  the most abundant species in each set of isoaccepting tRNAs.   • Ribosomes, which are large complexes of RNA and protein, have  a structure that is determined by their RNA components. A small  subunit and large subunit associate to form the intact ribosome,   which accommodates an aa-tRNA in the A site, a peptidyl-tRNA  in the P site, and a deacylated tRNA in the E site.   • During translation initiation, an initiator tRNA charged with  fMet (in prokaryotes) or Met (in eukaryotes), an mRNA with an  AUG initiation codon, and the ribosomal subunits assemble.  Initiation requires GTP-hydrolyzing initiation factors. The initi¬  ating AUG codon is identified in prokaryotes via the mRNA’s  Shine-Dalgarno sequence. In eukaryotes it is the mRNA’s initiat¬  ing AUG that is identified in a complex process by its proximity  to the mRNA’s 5' cap.   • A polypeptide chain is elongated from its N- to its C-terminus.  An aa-tRNA in complex with a GTP-hydrolyzing elongation fac¬  tor binds to the A site, where the ribosome senses correct    codon-anticodon pairing, a process whose accuracy is enhanced  by proofreading. The peptidyl transferase activity of the large sub¬  unit’s RNA catalyzes the attack of the A-site bound aa-tRNA’s  amino group on the peptidyl-tRNA in the P site. Following  transpeptidation, a second GTP-hydrolyzing elongation factor pro¬  motes the translocation of the new peptidyl-tRNA to the P site.   • As a polypeptide emerges from the ribosome, it begins to fold with  the assistance of chaperones. Proteins may undergo posttransla¬  tional processing that includes proteolysis, covalent modification,  translocation through a membrane, and glycosylation.    genetic code 962  codon 963  degeneracy 963  suppressor 963  reading frame 963  frameshift mutation 963  anticodon 964  synonym 966  Stop codon 966  acceptor stem 968    D arm 968  anticodon arm 968  Ti|;C arm 968  variable arm 969  aminoacyl—tRNA 971  aaRS 971    1. Could a single nucleotide deletion restore the function of a pro¬  tein-coding gene interrupted by the insertion of a 4-nt sequence?  Explain.   2. When a sequence of deoxynucleotides is entered into a transla¬  tion program (such as the ExPASy Translate tool at  http://www.expasy.org/tools/dna.html), six possible polypeptide  sequences are given as results. Explain.   3. List all possible codons present in a ribonucleotide polymer con¬  taining U and G in random sequence. Which amino acids are  encoded by this RNA?   4. Which amino acids are specified by codons that can be changed  to a UAG Stop codon by a single point mutation?   3. In the genetic code used by mitochondria in many species, UGA  codes for tryptophan rather than a Stop signal. How does mito¬  chondrial translation terminate?   6. Some mitochondria use a second codon, in addition to AUG, to  specify Met. Which codon (s) is (are) most likely to be used this  way?   7. A double-stranded fragment of viral DNA, one of whose strands  is shown below, encodes two peptides, called vir-1 and vir-2.  Adding this double-stranded DNA fragment to an in vitro tran¬  scription and translation system yields peptides of 10 residues  {vir-1) and 3 residues {vir-2).   (c) In a mutant viral strain, the T at position 23 has been re¬  placed with G. Determine the amino acid sequences of the  two peptides encoded by the mutant virus.   8. The sequence of the sense strand of a mammalian gene is   Determine the sequences of the mature RNA and the encoded  protein. Assume that transcription initiates at a G approximately  25 bp downstream of the TATAATA sequence, that each 5' splice  site has the sequence AG/GUAAGU, and that each 3' splice site  has the sequence CAG/G (where / marks the location of the splice).   9. The protein Xpot transports tRNAs out of the nucleus so that  they can be aminoacylated in the cytosol, (a) What tRNA struc¬  tural features is Xpot likely to recognize? (b) How does Xpot dis¬  tinguish mature tRNAs from pre-tRNAs?   10. The enzyme thiouridylase converts certain tRNA uridine residues  to 2-thiouridine. Draw the structure of this modified nucleotide.    11. IleRS uses a double-sieve mechanism to accurately produce  lie—tRNA Ile and prevent the synthesis of Val—tRNA Ile . Which  other pairs of amino acids differ in structure by a single carbon  and might have aaRSs that use a similar double-sieve proofread¬  ing mechanism?   12. Some mutations that do not alter the identity of the encoded  amino acid lead to diminished production of the protein. How  does the mutation affect translation?   13. Draw the structures of an I • U wobble pair and an I • C wob¬  ble pair.   14. In eukaryotes, the primary rRNA transcript is a 45S rRNA that  includes the sequences of the 18S, 5.8S, and 28S rRNAs sepa¬  rated by short spacers. What is the advantage of this operon-like  arrangement of rRNA genes?   13. Explain the significance of the observation that peptides such as  fMet-Leu-Phe “activate” the phagocytotic (particle-engulfing)  functions of mammalian leukocytes (white blood cells).   16. Explain why prokaryotic ribosomes can translate a circular  mRNA molecule, whereas eukaryotic ribosomes normally cannot,  even in the presence of the required cofactors.   17. Explain why the translation of a given mRNA can be inhibited  by a segment of its complementary sequence, a so-called anti-  sense RNA.   18. Eukaryotic initiation factor eIF2B is a guanine nucleotide exchange  factor. Explain why eIF2B would enhance the activity of eIF2.   19. EF-Tu binds all aminoacyl-tRNAs with approximately equal affin¬  ity so that it can deliver them to the ribosome with the same ef¬  ficiency. Based on the experimentally determined binding constants  for EF-Tu and correctly charged and mischarged aminoacyl-  tRNAs (see table), explain how the tRNA-EF-Tu recognition sys¬  tem could prevent the incorporation of the wrong amino acid  during translation.    Source: [LaRiviere, EJ., Wolfson, A.D., and Uhlenbeck, O.C., Science  294, 167 (2001).]    20. The antibiotic paromomycin binds to a ribosome and induces  the same conformational changes in 16S rRNA residues A1492  and A1493 as are induced by codon-anticodon pairing (Fig. 27-32).  Propose an explanation for the antibiotic effect of paromomycin.   21. The rate of the peptidyl transferase reaction increases as the pH  increases from 6 to 8. Explain these results in terms of its reac¬  tion mechanism.   22. It has been proposed that residue A2486 is protonated and there¬  fore stabilizes the tetrahedral reaction intermediate. Is this mech¬  anistic embellishment consistent with the observed pH effect  (Problem 21)? Explain.   23. Cells can make certain oligopeptides using conventional enzymes  rather than mRNA-directed translation by a ribosome. Draw the  product of the reaction catalyzed by L-glutamate:L-cysteine  gamma-ligase (y-glutamylcysteine synthetase).   24. The peptidyl transferase activity of the ribosome does not require  an additional source of free energy. Is the reaction catalyzed by  y-glutamylcysteine synthetase (Problem 23) also exergonic?   25. All cells contain an enzyme called peptidyl-tRNA hydrolase,  and cells that are deficient in the enzyme grow very slowly. What  is the probable function of the enzyme and why is it necessary?   26. Genetically engineered mRNAs that code for a stretch of basic  residues, such as poly(Lys), induce translation termination and  destruction of the nascent polypeptide. Explain how this response  would protect cells from the effect of faulty transcription that  produces mRNAs with mutated Stop codons.   27. Calculate the energy required, in ATP equivalents, to synthesize  a 100-residue protein from free amino acids in E. coli (assume  that the N-terminal Met remains attached to the polypeptide and  that no ribosomal proofreading occurs).   28. Design an mRNA with the necessary prokaryotic control sites  that codes for the octapeptide Lys-Pro-Ala-Gly-Thr-Glu-Asn-Ser.   Not all naturally occurring pep¬  tides are synthesized by ribosomes that decode messenger RNA.  How does nonribosomal peptide synthesis take place? How does  it differ from polyketide synthesis (Box 20-3)? What are some func¬  tions of these peptides?   by a mutation in an enzyme leading to the vitamin’s synthesis.   Judson, H.E, The Eighth Day of Creation, Expanded edition, Part II, Cold  Spring Harbor Laboratory Press (1996). [A fascinating historical nar¬  rative on the elucidation of the genetic code.]   Knight, R.D., Freeland, S.J., and Landweber, L.F., Selection, history and  chemistry: the three faces of the genetic code, Trends Biochem. Sci.  24, 241-247 (1999).   Yanofsky, C., Establishing the triplet nature of the genetic code, Cell 128,  815-818 (2007).   Ibba, M., Becker, H.D., Stathopoulos, C., Tumbula, D.L., and Soil, D.,  The adaptor hypothesis revisited, Trends Biochem. Sci. 25, 311-316  (1999). [Describes exceptions to the 20 aminoacyl synthetase rule.]  Nureki, O., Vassylyev, D.G., Tateno, M., Shimada, A., Nakama, T.,  Fukai, S., Konno, M., Hendrickson, T.L., Schimmel, P., and  Yokoyama, S., Enzyme structure with two catalytic sites for double¬  sieve selection of substrate, Science 280, 578-582 (1998). [The X-  ray structures of IleRS in complexes with isoleucine and valine.]   Ben-Shem, A., Jenner, L., Yusupova, G., and Yusupov, M., Crystal struc¬  ture of the eukaryotic ribosome, Science 330, 1203-1209 (2010).   Dintzis, H.M., The wandering pathway to determining N to C synthe¬  sis of proteins, Biochem. Mol. Biol. Educ. 34, 241-246 (2006). [A  personal history of how the direction of polypeptide synthesis was  determined.]   Green, R. and Lorsch, J.R., The path to perdition is paved with protons,  Cell 110, 663-668 (2002). [Discusses the difficulties in characteriz¬  ing the catalytic mechanism of the ribosomal peptidyl transferase.]   Jackson, R.J., Hellen, C.U.T., and Pestova, T.V., The mechanism of eu¬  karyotic translation and principles of its regulation, Nature Rev. Mol.  Cell Biol. 10, 113-127 (2010).   Kiel, M.C., Kaji, H., and Kaji, A., Ribosome recycling, Biochem. Mol.  Biol. Educ. 35, 40-44 (2007).   Korostelev, A. and Noller, H.F., The ribosome in focus: new structures  bring new insights, Trends Biochem. Sci. 32, 434-441 (2007).   Leung, E.K.Y., Suslov, N., Tuttle, N., Sengupta, R., and Piccirilli, J.A.,  The mechanism of peptidyl transfer catalysis by the ribosome, Annu.  Rev. Biochem. 80, 327-333 (2011).   Moore, P.B. and Steitz, T.A., RNA, the first macromolecular catalyst: the  ribosome is a ribozyme, Trends Biochem. Sci. 28, 411-418 (2003);  and The structural basis of large ribosomal subunit function, Annu.  Rev. Biochem. 72, 813-850 (2003).   Nissen, P., Kjeldgaard, M., and Nyborg, J., Macromolecular mimicry,  EMBO J. 19, 489-495 (2000). [Describes how translation factors  and other proteins function by adopting the same structures and in¬  teracting with the same binding sites as nucleic acids.]   Ogle, J.M., Brodersen, D.E., Clemons, W.M., Jr., Tarry, M.J., Carter,  A.P., and Ramakrishnan, V., Recognition of cognate transfer RNA  by the 30S ribosomal subunit, Science 292, 897-902 (2001). [Pre¬  sents X-ray structural evidence for an rRNA sensor that verifies cor¬  rect codon-anticodon pairing in the ribosome.]   Rodnina, M.V., Beringer, M., and Wintermeyer, W., How ribosomes  make peptide bonds, Trends Biochem. Sci. 32, 20-26 (2007); and  Rodnina, M.V. and Wintermeyer, W., The ribosome goes Nobel,  Trends Biochem. Sci. 35, 1-5 (2010).   Schmeing, T.M., and Ramakrishnan, V., What recent ribosome structures  have revealed about the mechanism of translation, Nature 461,  1234-1242 (2009). [Discusses the ribosomal structures responsible  for various phases of translation.]    Schmeing, T.M., Voorhees, R.M., Kelley, A.C., Gao, Y.-G., Murphy, EV.,  IV, Weir, J.R., and Ramakrishnan, V., The crystal structure of the  ribosome bound to EF-Tu and aminoacyl-tRNA; and Gao, Y.-G.,  Selmer, M., Dunham, M., Weixlbaumer, A., Kelley, A.C., and  Ramakrishnan, V., The structure of the ribosome with elongation  factor G trapped in the posttranslocational state, Science 326,  688-693 and 694-699 (2009).   Selmer, M., Dunham, C.M., Murphy, F.V., IV, Weixlbaumer, A., Petry,  S., Kelley, A.C., Weir, J.R., and Ramakrishnan, V., Structure of the  70S ribosome complexed with mRNA and tRNA, Science 313,  1935-1942 (2006); and Korostelev, A., Trakhanov, S., Laurberg, M.,  and Noller, H.F., Crystal structure of a 70S ribosome-tRNA com¬  plex reveals functional interactions and rearrangements, Cell 126,  1065-1077 (2006). [Presents high resolution (3.7 and 2.8 A) X-ray  structures of the T. thermophilus ribosome.]   Sievers, A., Beringer, M., Rodnina, M.V., and Wolfenden, R., The ribo¬  some as an entropy trap, Proc. Natl. Acad. Sci. 101, 7897-7901  (2004).   Valle, M., Zavialov, A., Li, W., Stagg, S.M., Sengupta, J., Nielsen, R.C.,  Nissen, P., Harvey, S.C., Ehrenberg, M., and Frank, J., Incorpora¬  tion of aminoacyl-tRNA into the ribosome as seen by cryo-electron  microscopy, Nature Struct. Biol. 10, 899-906 (2003).   Becker, T., et ah, Structure of monomeric yeast and mammalian Sec6l  complexes interacting with the translating ribosome, Science 326,  1369-1373 (2009).   Fedyukina, D.V. and Cavagnero, S., Protein folding at the exit tunnel,  Annu. Rev. Biophys. 40, 337-359 (2011).   Halic, M., Becker, T., Pool, M.R., Spahn, C.M.T., Grassucci, R.A., Frank,  J., and Beckmann, R., Structure of the signal recognition particle in¬  teracting with the elongation-arrested ribosome, Nature 427,  808-814 (2004).   Hard, F.U. and Hayer-Hartl, M., Molecular chaperones in the cytosol:  from nascent chain to folded protein, Science 295, 1852-1858  (2002). [Describes the structures, functions, and potential interac¬  tions of the various chaperone systems in eukaryotes, eubacteria, and  archaea.]   Williams, D.B., Beyond lectins: the calnexin/calreticulin chaperone sys¬  tem of the endoplasmic reticulum, J. Cell Sci. 119, 615-623 (2006).   The controlled expression of genetic information requires a multitude of factors that  interact specifically and nonspecifically with DNA. This painting schematically diagrams  transcription factors in the preinitiation complex whose formation in eukaryotes must  precede the transcription of DNA to mRNA. [Illustration, Irving Geis. Image from the  Irving Geis Collection/Howard Hughes Medical Institute. Rights owned by HHMI.  Reproduction by permission only.]    The faithful replication of DNA ensures that all the descendants of a single  cell contain virtually identical sets of genetic instructions. Yet individual cells  may differ—sometimes dramatically—from their progenitors, depending on  how those instructions are read. The expression of genetic information in a  given cell or organism, that is, the synthesis of RNA and proteins specified by  the DNA sequence, is neither random nor fully preprogrammed. Rather, the  information in an organisms genome must be tapped in an orderly fashion dur¬  ing development and yet must also be available to direct the organisms responses  to changes in internal or external conditions.   Much of the mystery surrounding the regulation of gene expression has  to do with how genetic information is organized and how it can be located  and accessed on an appropriate time scale. The complexity of the mechanisms  for transcribing and translating genes suggests the potential for even more  complicated systems for enhancing or inhibiting these processes. Indeed, we  have already seen numerous examples of how accessory protein factors influ¬  ence the rates or specificities of transcription and translation. In this chapter,  we consider some additional aspects of gene expression by examining a vari¬  ety of strategies used by prokaryotes and eukaryotes to control how genetic  information specifies cell structures and metabolic functions.   B Tumor Suppressors Prevent Cancer  C Apoptosis Is an Orderly Process  D Development Has a Molecular Basis   • An organism’s complexity increases with the number of its genes but not the size  of its genome.   Genomics, the study of organisms’ genomes, was established as a discipline with  the advent of techniques for rapidly sequencing enormous tracts of DNA, such  as make up the chromosomes of living things. Studies that once relied on the  hybridization of oligonucleotide probes to identify genes can now be conducted  by comparing nucleotide sequences deposited in databases (Section 5-3E). In fact,  a computerized database is the only practical format for storing the vast amount  of information provided by a whole-genome sequence. Even a simplified diagram  of the genome of the relatively simple bacterium Helicobacter pylori (a major cause  of peptic ulcers) reveals a bewildering array of genes (Fig. 28-1). Nevertheless,  the ability to sequence and map the entire genome of an organism makes it pos¬  sible to draw far-reaching conclusions about how many genes an organism con¬  tains, how they are organized, and how they serve the organism.   The rough correlation between the quantity of an organisms unique genetic  material (its C value) and the complexity of its morphology and metabolism  (Table 3-3) has numerous exceptions, known as the C-value paradox. For ex¬  ample, the genomes of lungfishes are 10 to 15 times larger than those of mam¬  mals (Fig. 28 -2). Some algae have genomes 10 times larger still. We know  that much of this “extra” DNA is unexpressed, but its function is largely a  matter of conjecture. The complete genomic sequences of numerous prokary¬  otes and eukaryotes, including data from some large, complicated genomes,  nevertheless indicate that the apparent number of genes, like the overall quan¬  tity of DNA, roughly parallels the organisms complexity (Table 28-1). Thus,  humans have —23,000 genes compared to E. colis —4300 genes.   Only a Small Portion of the Human Genome Encodes Proteins. All but   about 0.3% of the 3,038,000-kb human genome has been fully sequenced (the    FIG. 28-1 Map of the 1670-kb H. pylori  circular chromosome. The 1590 predicted  protein-coding sequences (91% of the genome)  are indicated in different colors on the two  outermost rings, which represent the two DNA  strands. The third and fourth rings indicate  intervening sequence elements and other  repeating sequences (2.3% of the genome). The  fifth and sixth rings show the genes for tRNAs,  rRNAs, and other small RNAs (0.7% of the  genome). Intergenic sequences account for 6%  of the genome. [Courtesy of J. Craig Venter,  Celera Genomics, Rockville, Maryland.]    FIG. 28-2 Range of haploid genome DNA contents in various  categories of organisms. The morphological complexity of the  organisms, as estimated from their number of different cell types,  increases from bottom to top. Haploid genome size roughly correlates    with complexity; exceptions are described by the C-value paradox.  [After Raff, R.A. and Kaufman, T.C., Embryos, Genes, and Evolution,  p. 314, Macmillan (1983).]    remaining DNA includes gene-poor regions near the centromeres and telo¬  meres). Only about 1.2% of the genome encodes proteins , and the 23,000 or so  protein-coding genes are distributed throughout the genome, sometimes in  clusters. Identifying the genes in human or other genomes involves several  approaches.   A gene can be identified by its homology to a previously described mRNA  or protein sequence. A protein-coding gene may also be identified as an open   Chapter 28 Regulation of Gene Expression reading frame (ORF), a sequence that is not interrupted by Stop codons and   that exhibits the same codon-usage preferences as other genes in the organ¬  ism. However, our incomplete knowledge of the features through which cells  recognize genes, combined with the fact that human genes consist of relatively  short exons (averaging —150 nt) separated by much longer introns (averaging  ~3500 nt and often much longer), has limited the success of computer-based  gene identification algorithms. Such algorithms therefore rely on sequence  alignments with expressed sequence tags (ESTs; cDNAs that have been re-  verse-transcribed from mRNAs) together with alignments with known genes  from other organisms. Genes may also be revealed by the presence of a CpG  island (Box 25-4). CpG dinucleotides are present in vertebrate genomes at  only about one-fifth their randomly expected frequency (because the sponta¬  neous deamination of m 5 C yields a normal T and therefore often results in a  CG —> TA mutation), but they appear in clusters, or islands, at near-normal  frequencies in the upstream regions of many genes (~56% of human genes).   The number of human genes is much lower than had once been predicted.  The discrepancy between current estimates—some predicting <20,000  genes—and earlier estimates of as many as 140,000 protein-coding, or struc¬  tural, genes is largely attributed to the high prevalence of alternative splicing  (Section 26-3B). In addition, variations in posttranslational processing  (Section 27-5B) enable a given DNA sequence to give rise to several function¬  ally distinct protein products.   The functions of many human genes have been identified through se¬  quence comparisons at the level both of protein families and of domains  (Fig. 28-3). Note that nearly 42% of these genes have unknown functions,  as is likewise the case with most other genomes of known sequence, includ¬  ing those of prokaryotes. Genes with no known function are called orphan  genes. Some of these sequences may represent novel genes whose protein  products have not yet been discovered. Others may simply be counterparts  of known genes but are too different in sequence to be recognized as such.   About three-quarters of all known human genes appear to have counterparts  in other species. About one-quarter are present only in other vertebrates, and one-  quarter are present in prokaryotes as well as eukaryotes. As expected, the human  genome contains approximately the same number of “housekeeping” genes (genes  required for the most fundamental cellular activities) as other eukaryotes. But it  includes relatively more genes for vertebrate-specific activities, such as those rel¬  evant to the immune system and neuronal and hormonal signaling pathways.    FIG. 28-3 Distribution of molecular  functions of the putative structural genes in  the human genome. Each wedge of the pie  chart lists, in parentheses, the percentage of the  genes assigned to the indicated category of  molecular function. The outer circle indicates the  general functional categories whereas the inner  circle provides a more detailed breakdown of the  categories. [Courtesy of J. Craig Venter, Celera  Genomics, Rockville, Maryland.]   structural protein (2.8%)  extracellular matrix (1.4%)  immunoglobulin (0.9%)  channel (1.3%)  motor (1.2%)    A Significant Portion of the Human Genome Is Transcribed to RNA.   Although the proportion of exonic DNA in the human genome is relatively  small (—1.4%), as much as 80% of the genome may be transcribed. This mass  of RNA includes the products of 4000 or so identified genes for tRNAs,  rRNAs, and other small RNAs, as well as tens of thousands of other noncod¬  ing RNAs (ncRNAs), many of which have no known function. Some ncRNAs  are polyadenylated, as are mRNAs (Section 26-3A), but do not correspond to  any known genes. One possibility is that this RNA represents transcriptional  noise arising from promoter-like elements scattered throughout the genome.  However, the observation that some ncRNAs exhibit conservation between  species and tissue-specific splice variants suggests that such RNA performs a  regulatory function.   Many Disease Genes Have Been Identified. The availability of human ge¬  nomic data has facilitated the identification of sequence variants that are as¬  sociated with particular diseases. Over 12,500 such variants have been catalogued  [in the Online Mendelian Inheritance in Man (OMIM) database  (http://www.ncbi.nlm.nih.gov/omim/)], including those for cystic fibrosis (Box  3-1), glycogen storage diseases (Box 16-2), and many hereditary forms of can¬  cer. However, monogenetic diseases are relatively rare. Most diseases result  from interactions among several genes and from environmental factors. One  goal of genomics is to identify genetic features that can be linked to suscep¬  tibility to disease or infection. To that end, a catalog of human DNA sequence  variations is being compiled as a database of single nucleotide polymor¬  phisms (SNPs; pronounced “snips”). A SNP, or single-base difference between  individuals, occurs about every —1250 bp on average. Over 115 million SNPs  have been described. Although less than 1% of them result in protein vari¬  ants, and probably fewer still have functional consequences, it is becoming in¬  creasingly apparent that SNPs are largely responsible for an individuals  susceptibility to many diseases as well as to adverse reactions to drugs (side ef¬  fects; Section 12-4). Moreover, SNPs can potentially serve as genetic markers  for nearby disease-related genes.   Genome-wide association studies (GWAS) attempt to link common ge¬  netic variants to specific diseases that have no obvious hereditary component.  Such studies typically involve examining the DNA of thousands of individu¬  als to obtain a dataset in which relatively rare sequence variations can be tab¬  ulated. For example, 39 loci (sites) have been shown to be associated with type  2 diabetes (Section 22-4B), and 71 loci with Crohn’s disease, an autoimmune  disorder. Although the increased risk of disease assigned to a particular allele  may be low, the entire set of variants can explain up to 50% of the heritabil-  ity of a disease. Moreover, the discovery of new genes associated with a con¬  dition can provide additional targets for drug development (Section 12-4).   Genes are not necessarily randomly distributed throughout an organisms  genome. For example, the average gene frequency in the human genome is  — 1 gene per 100 kb but ranges from 0 to 64 genes per 100 kb. Moreover,  some protein-coding genes and other chromosomal elements exhibit a certain  degree of organization. Prokaryotic genomes, for example, contain numerous  operons, in which, as we have seen (Section 26-IB), genes with related func¬  tions (e.g., encoding the proteins involved in a particular metabolic pathway)  occur close together, sometimes in the same order in which their encoded pro¬  teins act in a metabolic reaction sequence, and are transcribed onto a single  polycistronic mRNA.   Gene clusters also occur in both prokaryotes and eukaryotes. Although  most genes occur only once in an organisms haploid genome, genes such as  those for rRNA and tRNA, whose products are required in relatively large   FIG. 28-4 An electron micrograph of tandem arrays of actively transcribing 18S,  5.8S, and 28S rRNA genes from the newt Notophthalmus viridescens. The axial fibers  are DNA. The fibrillar “Christmas tree” matrices, which consist of newly synthesized RNA  strands in complex with proteins, outline each transcriptional unit. Note that the longest  ribonucleoprotein branches are only —10% of the length of their corresponding DNA.  Apparently, the RNA strands are compacted through secondary structure interactions  and/or protein associations. The matrix-free segments of DNA are the untranscribed  spacers. [Courtesy of Oscar L. Miller, Jr., and Barbara R. Beatty, University of Virginia.]    amounts, may occur in multiple copies. As we saw in Section 26-3C, large  rRNA transcripts are cleaved to yield the mature rRNA molecules.  Furthermore, the transcribed blocks of 18S, 5.8S, and 28S eukaryotic rRNA  genes are arranged in tandem repeats that are separated by nontranscribed  spacers (Fig. 28 -4). The rRNA genes, which may be distributed among sev¬  eral chromosomes, vary in haploid number from less than 50 to over 10,000,  depending on the species. Humans, for example, have 50 to 200 blocks of  rRNA genes spread over five chromosomes. tRNA genes are similarly reiter¬  ated and clustered.   Protein-coding genes almost never occur in multiple copies , presumably be¬  cause the repeated translation of a few mRNA transcripts provides adequate  amounts of most proteins. One exception is histone proteins, which are re¬  quired in large amounts during the short time when eukaryotic DNA synthe¬  sis occurs. Not only are histone genes reiterated (up to --TOO times in  Drosophila :), they often occur as sets of each of the five different histone genes  separated by nontranscribed sequences (Fig. 28-5). The gene order and the  direction of transcription in these quintets are preserved over large evolution¬  ary distances. The spacer sequences vary widely among species and, to a lim¬  ited extent, among the repeating quintets within a genome. In birds and  mammals, which contain 10 to 20 copies of each of the five histone genes,  the genes occur in clusters but in no particular order.   Other gene clusters contain genes of similar but not identical sequence.  For example, human globin genes are arranged in two clusters on separate  chromosomes (Fig. 28-6). The various genes are transcribed at different de¬  velopmental stages. Adult hemoglobin is an a 2 (3 2 tetramer, whereas the first  hemoglobin made by the human embryo is a £ 2 82 tetramer in which £ and 8  are a- and (3-like subunits, respectively (Fig. 28-7). At approximately 8 weeks  after conception, fetal hemoglobin containing a and y subunits appears. The  y subunit is gradually supplanted by (3 starting a few weeks before birth. Adult  human blood normally contains ~97% a 2 (3 2 hemoglobin, 2% a 2 8 2 (in which  8 is a (3 variant), and 1% a 2 y 2 .   The a-globin gene cluster (Fig. 28-6, top), which spans 28 kb, contains  three functional genes: the embryonic £ gene and two slightly different a  genes, al and a2, which encode identical polypeptides. The a cluster also  contains four pseudogenes (nontranscribed relics of ancient gene duplications):    FIG. 28-5 The organization and lengths  of the histone gene cluster repeating units in  a variety of organisms. Coding regions are  indicated in color, and spacers are gray. The  arrows denote the direction of transcription (the  top three organisms are distantly related sea  urchins).    FIG. 28-6 The organization of human globin genes. Red boxes represent active  genes; green boxes represent pseudogenes; yellow boxes represent repetitive sequences,  with the arrows indicating their relative orientations; and triangles represent Alu elements in  their relative orientations. [After Karlsson, S. and Nienhuis, A.W., Annu. Rev. Biochem. 54,  1074 (1985).]   i|/£, i|ia2, i|ial, and iJjO. The [5-globin gene cluster (Fig. 28-6, bottom ), which  spans >60 kb, contains five functional genes: the embryonic 8 gene, the  fetal genes G y and A y (duplicated genes encoding polypeptides that differ only  by having either Gly or Ala at position 136), and the adult genes 8 and (3.  The (3-globin cluster also contains one pseudogene, i(j| 3. Both a and (3 gene  clusters also include copies of repetitive DNA sequences (see below).   Approximately 11% of the E. coli genome consists of nontranscribed regions,  including the regulatory sequences that separate individual genes and sites that  govern the origin and termination of replication. In addition, bacterial  genomes typically contain insertion sequences (Section 25-6C) and the rem¬  nants of integrated bacteriophages.   The small sizes of prokaryotic genomes probably exert selective pressure against  the accumulation of useless DNA. Eukaryotic genomes, however, which are usu¬  ally much larger than prokaryotic genomes, apparently are not subject to the same  selective forces. About 30% of the yeast genome consists of nonexpressed se¬  quences, and the proportion is far greater in higher eukaryotes. Much of this  DNA consists of repetitive sequences that contribute significantly to the rela¬  tively large genomes of certain plants and amphibians. Some repetitive DNA  plays a structural role at the centromeres of eukaryotic chromosomes, the re¬  gions attached to the microtubular spindle during mitosis. These sequences  may help align chromosomes and facilitate recombination. The telomeres also   FIG. 28-7 The progression of human globin chain synthesis with fetal  development. Note that any red blood cell contains only one type each of a- and (3-like  subunits. [After Weatherall, D.J. and Clegg, J.B., The Thalassaemia Syndromes (3rd ed.),  p. 64, Blackwell Scientific Publications (1981).]    At least 14 human diseases are associated with repeated trinu¬  cleotides in certain genes. These trinudeotide repeats exhibit an  unusual genetic instability: Above a threshold of about 35 to 50  copies (100-150 bp), the repeats tend to expand with successive  generations. Because the overall length of the repeat typically  correlates with the age of onset of the disease, descendants of  an individual with a trinucleotide repeat disease tend to be more  severely affected and at an earlier age. The disease is therefore  said to exhibit genetic anticipation.   Some types of trinucleotide repeat diseases are caused by  massive expansion (usually to hundreds of copies) of a  trinucleotide in the noncoding region of a gene, for example, in a  region upstream of the transcription start site, in a 5' or 3' un¬  translated region (UTR), or in an intron (see table). These expan¬  sions generally affect gene expression. For example, myotonic  dystrophy results from aberrant expression of a protein kinase.  The severity of the symptoms, progressive muscle weakness and  wasting, correlate with the number of CTG repeats (>2000 in  some cases) in its gene’s 5' UTR.    Fragile X syndrome, the most common cause of mental  retardation after Down’s syndrome, is so named because the tip  of the X chromosome’s long arm is connected to the rest of the  chromosome by a slender thread that is easily broken. As in many  trinucleotide repeat diseases, the genetics of fragile X syndrome  are bizarre. The maternal grandfathers of individuals having fragile  X syndrome may be asymptomatic. Their daughters are likewise  asymptomatic, but these daughters’ children of either sex may  have the syndrome. Evidently, the fragile X defect is activated by  passage through a female.   The affected gene in fragile X syndrome, FMR1 (for fragile X  mental retardation 1 ), encodes a 632-residue RNA-binding protein  named FMRP (for FMR protein), which apparently functions in the  transport of certain mRNAs from the nucleus to the cytoplasm.  FMRP, which is highly conserved in vertebrates, is heavily ex¬  pressed in brain neurons, where a variety of evidence indicates  that it participates in the proper formation and/or function of  synapses. In the general population, the 5' untranslated region of  FMR1 contains a (CGG) n sequence with n ranging from 6 to 60  (such a gene is said to be polymorphic). However, in individuals  with fragile X syndrome, this triplet repeat has undergone an  astonishing expansion to values of n ranging from >200 to sev¬  eral thousand. Moreover, these triplet repeats differ in size among  siblings and often exhibit heterogeneity within an individual,  suggesting that they are somatically generated.   Some trinucleotide repeat diseases result from the moderate  expansion of a CAG triplet (which codes for Gin) in the protein¬  coding region of a gene. For example, in normal individuals, hunt-  ingtin, a polymorphic, —3150-residue protein of unknown  function, contains a stretch of 1 1 to 34 consecutive Gin residues  beginning 17 residues from its N-terminus. However, in Hunting¬  ton’s disease (HD), this poly(Gln) tract has expanded to between    are composed of repeating DNA sequences (Section 25-3C). A variety of neu¬  rological diseases result from excessively reiterated trinucleotide sequences (see  Box 28-1).   Nearly identical sequences up to 10 bp long that are tandemly repeated in  clusters may be present at >10 6 copies per haploid genome. Approximately  3% of the human genome consists of highly repetitive DNA sequences (also  known as short tandem repeats; STRs), with the greatest contribution (0.5%)  provided by dinucleotide repeats, most frequently CA and TA. STRs appear  to have arisen by template slippage during DNA replication. This occurs more  frequently with short repeats, which therefore have a high degree of length  polymorphism in the human population. Because the number of repeats at a  given STR locus varies between individuals, analysis of STRs is widely used  for DNA fingerprinting (Box 3-2).   Moderately Repetitive Sequences Arise from Transposons. Moderately  repetitive DNA (<10 6 copies per haploid genome) occurs in segments of 100  to several thousand base pairs that are interspersed with larger blocks of unique  DNA. Most of the repetitive sequences are the remnants of retrotransposons  (transposable elements that propagate through the intermediate synthesis of    37 and 86 repeats. Synthetic poly(Gln) aggregates as p sheets  that are linked by hydrogen bonds involving both their main chain  and side chain amide groups.   Individuals with HD, an autosomal dominant condition, suffer  progressive choreic (jerky and disordered) movements, cognitive  decline, and emotional disturbances over a 15- to 20-year course  that is invariably fatal. This devastating disease typically develops  late in life, around age 40, or earlier in individuals with high num¬  bers of trinucleotide repeats. One of the hallmarks of HD, as well  as of other neurodegenerative diseases such as Alzheimer’s dis¬  ease, is the deposition of insoluble protein aggregates in the  cytosol and nuclei of neurons (Section 6-5C). The appearance of  these intracellular aggregates (presumably huntingtin and/or its  proteolytic products in HD) often coincides with the onset of  neurological symptoms, although it is not clear whether protein  aggregation directly causes neuronal death and its attendant  symptoms. The long incubation period before the symptoms of HD  become evident is attributed to the lengthy nucleation time for  aggregate formation, much like what we have seen occurs in the  formation of amyloid fibrils (Section 6-5C).   The expansion of trinucleotide repeats occurs by an unknown  mechanism. One theory, that additional nucleotides are intro¬  duced by the slippage of DNA polymerases during replication,  is inconsistent with the gradual accumulation of trinucleotide  repeats over time in cells such as neurons that normally do not  divide. Another possibility is that the additional nucleotides are  introduced during DNA repair processes (which are ongoing  and independent of DNA replication) and may result from hairpin  formation within the trinucleotide repeat region. These  intrastrand base-paired structures (see figure) may lead to  misalignment of DNA strands and the polymerization of  additional nucleotides to fill in the gaps.    RNA; Section 25-6C). Around 42% of the human genome consists of three  types of retrotransposons (Table 28-2):   1. Long interspersed nuclear elements (LINEs) are 6- to 8-kb-long seg¬  ments. The great majority of these molecular parasites have accumu¬  lated mutations that render them transcriptionally inactive, but a few  still appear capable of further transposition. Several hereditary diseases  are caused by insertion of a LINE into a gene.   Chapter 28 Regulation of Gene Expression 2. Short interspersed nuclear elements (SINEs) consist of 100- to 400-bp   elements. The most common SINEs in the human genome are mem¬  bers of the Alu family, which are so named because most of their  —300-bp segments contain a cleavage site for the restriction endonu¬  clease Alul (AGCT; Table 3-2). The globin gene cluster contains sev¬  eral Alu elements (Fig. 28-6).   In addition, the human genome contains DNA transposons that resemble bac¬  terial transposons. Overall, —45% of the human genome consists of widely  dispersed and almost entirely inactive transposable elements.   No function has been unequivocally assigned to moderately repetitive  DNA, which therefore has been termed selfish or junk DNA. This DNA ap¬  parently is a molecular parasite that, over many generations, has disseminated  itself throughout the genome through transposition. The theory of natural se¬  lection suggests that the increased metabolic burden imposed by the replica¬  tion of an otherwise harmless selfish DNA would eventually lead to its  elimination. Yet for slowly growing eukaryotes, the relative disadvantage of  replicating an additional 100 bp of selfish DNA in an — 1-billion-bp genome  would be so slight that its rate of elimination would be balanced by its rate  of propagation. Because unexpressed sequences are subject to little selective pres¬  sure , they accumulate mutations at a greater rate than do expressed sequences.   DNA Polymorphisms Can Establish Genealogies. Unexpressed sequences,  which are subject to little selective pressure, evolve so much faster than ex¬  pressed sequences that they accumulate significant numbers of sequence    FIG. 28-8 A phylogenetic tree indicating  the lines of descent among 26 representative  human populations. The tree is based on the  sequences of 29 polymorphic loci. The major  groups of human populations are (A) Africans,   (D) Amerindians, and (E) Australopapuans. The  number beside each of the branches indicates  their relative lengths and is therefore indicative  of the genetic distances between the related  populations and thus the times between their  divergence. [Based on a drawing by Masatoshi  Nei, Pennsylvania State University.]    polymorphisms (variations) within a single species. Consequently, the evolu¬  tionary relationships among populations within a species can be established by  determining how a series of polymorphic DNA sequences are distributed  among them. For example, Masatoshi Nei inferred the genealogy of 26 repre¬  sentative human populations from around the world from the sequence vari¬  ations in 29 polymorphic loci. The resulting phylogenetic tree (Fig. 28-8;  opposite) indicates that non-African (Eurasian) populations are much more  closely related to each other than they are to African populations. Fossil evi¬  dence indicates that anatomically modern man arose in Africa —200,000 years  ago and spread throughout that continent. The phylogenetic tree therefore  suggests that all Eurasian populations are descended from a surprisingly small  “founder population” (perhaps only a few hundred individuals) that left Africa  — 100,000 years ago. A similar analysis indicates that the sickle-cell variant of  the (3-globin gene (Section 7-IE) arose on at least three separate occasions in  geographically distinct regions of Africa.   The lac repressor prevents or allows transcription of the lac operon.  cAMP stimulates transcription of catabolite-repressed operons.  Attenuation links amino acid availability to operon expression.    A complete genome sequence reveals the metabolic capabilities of an organ¬  ism, but gene sequences alone do not necessarily indicate when or where the en¬  coded molecules are produced. The route from gene sequence to fully functional  gene product offers many potential points for regulation, but in prokaryotes,  gene expression is primarily controlled at the level of transcription. This is perhaps  because prokaryotic mRNAs have lifetimes of only a few minutes, so transla¬  tional control is less necessary. In this section, we examine a few well-  documented examples of gene regulation in prokaryotes. In the following  section, we will consider how eukaryotic cells regulate gene expression.   Bacteria adapt to their environments by producing enzymes that metabolize  certain nutrients only when those substances are available. For example, E. coli  cells grown in the absence of lactose are initially unable to metabolize the  disaccharide. To do so they require two proteins: |3-galactosidase, which  catalyzes the hydrolysis of lactose to its component monosaccharides,   • What is the relationship between genome  size and gene number in different  organisms?   • What are some factors that allow humans  to get by with only slightly more genes  than invertebrates?   • What is the purpose of cataloging SNPs  and doing genome-wide association  studies?   • Summarize the relationship between  genome size, repetitive DNA, transcribed  DNA, and protein-coding exons in the  human genome.   • What is the source of the repetitive DNA  in the human genome?    and galactoside permease (also known as lactose permease; Section 10-3D),  which transports lactose into the cell. Cells grown in the absence of lactose  contain only a few molecules of the proteins. Yet a few minutes after lactose  is introduced into their medium, the cells increase the rate at which they syn¬  thesize the proteins by —1000-fold and maintain that pace until lactose is no  longer available. This ability to produce a series of proteins only when the sub¬  stances they metabolize are present permits the bacteria to adapt to their environ¬  ment without the debilitating need to continuously synthesize large quantities of  otherwise unnecessary enzymes.   Lactose or one of its metabolic products must somehow act as an inducer  to trigger the synthesis of the above proteins. The physiological inducer of the  lactose system, the lactose isomer 1,6-allolactose {at left), arises from lactoses  occasional transglycosylation by (3-galactosidase. Most in vitro studies of lactose  metabolism use isopropylthiogalactoside (IPTG; at left ), a synthetic inducer  that structurally resembles allolactose but is not degraded by (3-galactosidase.  Natural and synthetic inducers also stimulate the synthesis of thiogalactoside  transacetylase, an enzyme whose physiological role is unknown.   The genes specifying (3-galactosidase, galactoside permease, and thiogalac¬  toside transacetylase, designated Z , Y, and A, respectively, are contiguously  arranged in the lac operon (Fig. 26-5). All three structural genes are translated  from a single mRNA transcript. A nearby gene, /, encodes the lac repressor,  a protein that inhibits the synthesis of the three lac proteins.    lac Repressor Recognizes Operator Sequences. The target of the lac repres¬  sor is a region of the lac operon known as its operator, which lies near the be¬  ginning of the (3-galactosidase gene. In the absence of inducer, lac repressor  specifically binds to the operator to prevent the transcription of mRNA (Fig. 28-9#).  On binding inducer ; the repressor dissociates from the operator ; thereby permitting  the transcription and subsequent translation of the lac enzymes (Fig. 28-9 b).   The lac operon actually contains three operator sequences to which lac re¬  pressor binds with high affinity, known as 0 1? 0 2 , and 0 3 . O l5 the primary  repressor-binding site, was identified through its protection by lac repressor  from nuclease digestion. The 26-bp protected sequence lies within a nearly  twofold symmetrical sequence of 35 bp (Fig. 28-10). O x overlaps the tran¬  scription start site of the lacZ gene. The operator sequence 0 2 is centered 401  bp downstream, fully within the lacZ gene, and 0 3 is centered 93 bp upstream    FIG. 28-9 The expression of the lac operon. (a) In the   absence of inducer, the repressor (the product of the / gene) binds to  the operator (Ch), thereby preventing transcription of the lac operon  from the promoter (P). (b) On binding inducer, the repressor  dissociates from the operator, which permits the transcription and    subsequent translation of the lac structural genes (Z, Y, and A, which  respectively encode (3-galactosidase, lactose permease, and  thiogalactoside transacetylase).   Predict the effect of a mutation in I, P, O, Z, or Y.   FIG. 28-10 The base sequence of the lac operator 0 A . Its symmetry-related  regions, which comprise 28 of its 35 bp, are shaded in red.    of Oi, at the end of the lacl gene. Genetic engineering experiments show that  all three operator sequences must be present for maximum repression in vivo.   The observed rate constant for the binding of lac repressor to lac opera¬  tor is k ~ 10 10 M 1 • s 1 . This ' “on” rate is much greater than that calculated  for the diffusion-controlled process in solution: k ~ 10 7 M 1 • s 1 for mole¬  cules the size of lac repressor. Since it is impossible for a reaction to proceed  faster than its diffusion-controlled rate, lac repressor must not encounter op¬  erator from solution in a random three-dimensional search. Rather, it appears  that lac repressor finds its operator by nonspecific ally binding to DNA and slid¬  ing along it in a far more efficient one-dimensional search.   lac Repressor Binds Two DNA Segments Simultaneously. The isolation of  lac repressor, by Beno Miiller-Hill and Walter Gilbert in 1966, was exceed¬  ingly difficult because the repressor constitutes only —0.002% of the protein  in wild-type E. coli. Molecular cloning techniques (Section 3-5) later made it  possible to produce large quantities of lac repressor. Nevertheless, it was not  until 1996 that Ponzy Lu and Mitchell Lewis reported the complete three-  dimensional structure of the protein. Each 360-residue subunit of the repressor  homotetramer has four functional units (Fig. 28-11):   1. An N-terminal “headpiece,” which contains a helix—turn—helix (HTH)  motif that resembles those in other prokaryotic DNA-binding proteins  (Section 24-4B) and which specifically binds operator DNA sequences.   2. A linker, which contains a short a helix that acts as a hinge and also  binds DNA. In the absence of DNA, the hinge helices of the lac re¬  pressor tetramer are disordered, allowing the headpieces to move freely.   4. A C-terminal a helix, which is required for the quaternary structure  of lac repressor. In the tetramer, all four C-terminal helices associate.  Surprisingly, the repressor homotetramer does not exhibit the D 2  symmetry of nearly all homotetrameric proteins of known structure  (three mutually perpendicular twofold axes; Section 6-3) but is instead  V-shaped (with only twofold symmetry) and is therefore best consid¬  ered to be a dimer of dimers (Fig. 28-12).   The X-ray structure of a complex between lac repressor and  a 21-bp synthetic DNA containing a high-affinity binding se¬  quence reveals that each repressor tetramer binds two DNA seg¬  ments (Fig. 28-12). The repressors HTH motif fits snugly into  the major groove, bending the DNA so that it has a radius of cur¬  vature of 60 A. The two bound DNA segments are laterally sepa  rated by —25 A.    FIG. 28-1 1 Ribbon diagram of the lac  repressor monomer. The DNA-binding domain  containing its helix-turn-helix motif is red; the  hinge helix is yellow; the inducer-binding core is  light and dark blue; and the tetramerization helix  is purple. [Courtesy of Ponzy Lu and Mitchell  Lewis, University of Pennsylvania. PDBid 1 LBG.]    FIG. 28-1 2 X-Ray structure of the lac repressor tetramer bound to two  21-bp segments of DNA. The protein subunits are shown in ribbon form in yellow,  cyan, green, and orange and the DNA segments are drawn in space-filling form  C white, N blue, 0 red, and P orange. [Courtesy of Ponzy Lu and Mitchell Lewis  coordinates generated by Benjamin Wieder, University of Pennsylvania. PDBid 1LBG.]   FIG. 28-13 Model of the 93-bp loop  formed when the lac repressor tetramer  binds to 0-, and 0 3 . The proteins are  represented by their C a backbones, and the  DNA is shown in skeletal form with its  sugar-phosphate backbones traced by helical  ribbons. The model was constructed from the  X-ray structure of lac repressor (red) in complex  with two 21-bp operator DNA segments ( orange)  and the X-ray structure of CAP ( purple ) in  complex with its 30-bp target DNA (cyan). The  remainder of the DNA loop was generated by  applying a smooth curvature to B-DNA (white)  with the -10 and -35 regions of the lac  promoter highlighted in green. [Courtesy of  Ponzy Lu and Mitchell Lewis, University of  Pennsylvania.]    IPTG binds to the repressor core at the interface between the two do¬  mains colored light and dark blue in Fig. 28-11. This binding induces a con¬  formational change in the repressor dimer that is communicated through the  hinge helices to the headpieces. This causes the two DNA-binding domains  in each dimer to separate by ~3.5 A so that they can no longer simultane¬  ously bind DNA, thereby causing the repressor to dissociate from the DNA.   The lac repressor is an allosteric protein: IPTG binding to one subunit al¬  ters the DNA-binding activity of its dimeric partner (but not of the other  dimer in the repressor tetramer). Since the allosteric transition occurs within  the dimer, why does full repressor activity require a tetramer? Model-building  studies provide a plausible answer to this puzzle. A lac repressor tetramer simul¬  taneously binds to two operators so that it brings them together, forming a loop of  DNA either 93 or 401 bp long, depending on whether the repressor binds Oj and  0 3 or 0 2 and 0 2 . The formation of a stable looped structure may require ad¬  ditional DNA-binding proteins; one candidate is CAP (see below), a DNA-  binding protein that binds to the DNA between O x and 0 3 (Fig. 28-13). In  the model shown in Fig. 28-13, the /^promoter is part of the looped DNA.   It was widely assumed for years that lac repressor simply physically ob¬  structs the binding of RNA polymerase (RNAP) to the lac promoter. However,  experiments have demonstrated that RNAP can bind to the promoter in the  presence of the repressor but cannot properly initiate transcription.  Dissociation of the repressor in response to an inducer would allow unim¬  peded transcription. If RNAP were already bound to the lac promoter, tran¬  scription could begin immediately. Nevertheless, in the model shown in Fig.  28-13, the contact surface for RNAP is on the inside of the DNA loop, which  presumably would preclude the binding of RNAP. Further studies are needed  to resolve this apparent contradiction.   Glucose is E. colds metabolic fuel of choice; adequate amounts of glucose pre¬  vent the full expression of genes specifying proteins involved in the fermenta¬  tion of numerous other catabolites, including lactose, arabinose, and galactose,  even when they are present in high concentrations. This phenomenon, which  is known as catabolite repression, prevents the wasteful duplication of en-  ergy-producing enzyme systems. Catabolite repression is overcome in the ab¬  sence of glucose by a cAMP-dependent mechanism. cAMP levels are low in  the presence of glucose but rise when glucose becomes scarce.   CAP-cAMP Complex Stimulates the Transcription of Catabolite-Repressed  Operons. Certain E. coli mutants, in which the absence of glucose does not  relieve catabolite repression, are missing a cAMP-binding protein that is  synonymously named catabolite gene activator protein (CAP) or cAMP  receptor protein (CRP). CAP is a dimeric protein of identical 210-residue  subunits that undergoes a large conformational change on binding cAMP. The  CAP-cAMP complex, but not CAP alone, binds to the promoter region of  the lac operon (among others) and stimulates transcription in the absence  of repressor. CAP is therefore a positive regulator (it turns transcription  on), in contrast to lac repressor, which is a negative regulator (it turns tran¬  scription off).   The X-ray structure of CAP-cAMP in complex with a 30-bp segment of  DNA, whose sequence resembles the CAP-binding site, reveals that the CAP  dimer binds in successive turns of DNAs major groove via its two HTH mo¬  tifs so as to bend the DNA by ~90° around the protein dimer (Fig. 28-14^).  The bend arises from two ~45° kinks in the DNA between the fifth and sixth  bases out from the complex’s twofold axis in both directions and results in a  closing of the major groove and in an enormous widening of the minor groove  at each kink.   FIG. 28-14 X-Ray structures of CAP-cAMP-dsDNA complexes. The dsDNA and  cAMP in these twofold symmetric complexes are colored according to atom type with DNA  C white, cAMP C green, N blue, 0 red, and P orange, (a) CAP-cAMP in complex with a  palindromic 30-bp self-complementary DNA viewed with its twofold axis horizontal. The  protein is drawn in ribbon form with its identical subunits pink and blue and with their  C-terminal domains in darker shades. The DNA is shown in stick form with successive P  atoms in the same strand connected by orange rods and with the cAMP drawn in  space-filling form, (b) CAP-cAMP in complex with a 44-bp palindromic DNA and four  aCTD subunits. The DNA, CAP, and cAMP are viewed as in Part a with the DNA drawn in  space-filling form. The aCTD subunits are drawn in ribbon form with the aCTD CAP,DNA  green and the aCTD DNA dark green. [Part a based on an X-ray structure by Thomas Steitz,  Yale University. PDBid 1CGP. Part b based on an X-ray structure by Helen Berman and  Richard Ebright, Rutgers University. PDBid 1 LB2J © See Interactive Exercise 55.    Why is the CAP-cAMP complex necessary to stimulate the transcription  of its target operons? And how does it do so? The lac operon has a weak (low  efficiency) promoter; its —10 and — 35 sequences (TATGTT and TTTACA;  Fig. 26-6) differ significantly from the corresponding consensus sequences of  strong (high-efficiency) promoters (TATAAT and TTGACA; Fig. 26-6). Such  weak promoters evidently require some sort of help for efficient transcriptional  initiation.   Richard Ebright has shown that CAP interacts directly with RNAP via  the C-terminal domain of its 8 5-residue a subunit (aCTD) in a way that  stimulates RNAP to initiate transcription from a nearby promoter. The  aCTD, which is flexibly linked to the rest of the a subunit, binds dsDNA  nonspecifically but does so with higher affinity at A + T-rich sites. The  X-ray structure of CAP-cAMP in complex with the E. coli aCTD and a  44-bp palindromic DNA containing the 22-bp CAP-cAMP binding site  and 5'-AAAAAA-3' at each end, determined by Helen Berman and Ebright,  reveals how these components interact (Fig. 2S-\4b). The twofold symmet¬  ric CAP-cAMP-aCTD complex contains two differently located pairs of  aCTDs. Each member of the pair designated aCTD GAI ^ DNA binds both to CAP  and in the minor groove of a 6-bp segment of the DNA (5 , -AAAAAG-30  centered 19 bp from the center of the DNA (each member of the other pair   Chapter 28 Regulation of Gene Expression Q f aCTDs, designated aCTD DNA , interacts with the DNA but does not   contact other protein molecules). The common portions of the two CAP com¬  plexes pictured in Fig. 2S-l4a,b are closely superimposable, thereby indicat¬  ing that the conformation of CAP and its interaction with DNA are not  significantly altered by its association with the aCTD. Evidently, CAP-cAMP  transcriptionally activates RNAP via a simple “adhesive” mechanism that  facilitates and/or stabilizes its interaction with the promoter DNA.   The model for lac repressor binding (Fig. 28-13) paradoxically includes  CAP, which is an activator. This dual binding may be a mechanism for con¬  serving cellular energy in the absence of both glucose and lactose. If lactose  became available, lac repressor would dissociate, and CAP-cAMP would be  poised to promote transcription of the lac operon.   The E. coli trp operon encodes five polypeptides (which form three enzymes)  that mediate the synthesis of tryptophan from chorismate (Section 21-5B).  The five trp operon genes (A—E; Fig. 28-15) are coordinately expressed under  the control of the trp repressor, which binds L-tryptophan to form a complex  that specifically binds to the trp operator to reduce the rate of trp operon  transcription 70-fold (Section 24-4B). In this system, tryptophan acts as a co¬  repressor; its presence prevents what would be superfluous tryptophan  biosynthesis.   The trp repressor—operator system was at first thought to fully account for  the regulation of tryptophan biosynthesis in E. coli. However, the discovery of  trp deletion mutants located downstream from the operator ( trpO ) that in¬  crease trp operon expression sixfold indicated the existence of an additional  transcriptional control element. This element is located ~30 to 60 nucleotides    FIG. 28-1 5 Genetic map of the E. coli trp operon indicating synthesis of tryptophan (Section 21-5B). [After Yanofsky, C., J. Am.   the enzymes it specifies and the reactions they catalyze. The Med. Assoc. 218, 1027 (1971).]   upstream of the structural gene trpE in a 162-nucleotide leader sequence Section 2 Regulation of Prokaryotic   When tryptophan is scarce, the entire 6720-nucleotide polycistronic trp  mRNA, including the trpL sequence, is synthesized. When the encoded en¬  zymes begin synthesizing tryptophan, the rate of trp operon transcription de¬  creases as tryptophan binds to trp repressor. Of the trp mRNA that is  transcribed, however, an increasing proportion consists of only a 140-  nucleotide segment corresponding to the 5' end of trpL. The availability of  tryptophan therefore results in the premature termination of trp operon transcrip¬  tion. The control element responsible for this effect is consequently termed an  attenuator.   The trp Attenuator’s Transcription Terminator Is Masked when Tryptophan  Is Scarce. The attenuator transcript contains four complementary segments  that can form one of two sets of mutually exclusive base-paired hairpins (Fig.   28 - 16 ). Segments 3 and 4 together with the succeeding residues constitute a  transcription terminator (Section 26-ID): a G + C-rich hairpin followed by  several sequential U’s (compare with Fig. 26-10). Transcription rarely proceeds  beyond this termination site unless tryptophan is in short supply.   How does a shortage of tryptophan cause transcription to proceed past  the terminator? A section of the leader sequence, which includes segment 1 of  the attenuator, is translated to form a 14-residue polypeptide that contains two  consecutive Trp residues (Fig. 28-16, left). The position of this particularly rare  dipeptide (—1% of the residues in E. coli proteins are Trp) provided an im¬  portant clue to the mechanism of attenuation, as proposed by Charles   The formation of the base-paired 2 • 3 (antiterminator) hairpin (right)  precludes formation of the 1 • 2 and 3 • 4 (terminator) hairpins (left)  and vice versa. Attenuation results in the premature termination of   transcription immediately after nucleotide 140 when the 3 • 4 hairpin  is present The arrow indicates the mRNA site past which RNA  polymerase pauses until approached by an active ribosome. [After  Fisher, R.F. and Yanofsky, C., J. Biol. Chem. 258, 8147 (1983).]   (a) When Trp-tRNA Trp is abundant, the ribosome  translates trpL mRNA. The presence of the  ribosome on segment 2 prevents the formation  of the base-paired 2 • 3 hairpin. The 3 • 4  hairpin, an essential component of the  transcriptional terminator, can then form, thus  aborting transcription, (b) When Trp-tRNA Trp is  scarce, the ribosome stalls on the tandem Trp  codons of segment 1. This situation permits the  formation of the 2 • 3 hairpin, which precludes  the formation of the 3 • 4 hairpin. RNA  polymerase therefore transcribes through this  unformed terminator and continues transcribing  the trp operon.    Could attenuation occur if mRNA  segment 1 or segment 4 were missing?   Yanofsky (Fig. 28-17): An RNA polymerase that has escaped repression initi¬  ates trp operon transcription. Soon after the ribosomal initiation site of the  trpL sequence has been transcribed, a ribosome attaches to it and begins trans¬  lating the leader peptide. When tryptophan is abundant (i.e., there is a plen¬  tiful supply of Trp-tRNA Trp ), the ribosome follows closely behind the  transcribing RNA polymerase. Indeed, RNA polymerase pauses past position  92 of the transcript and continues transcribing only on the approach of a ri¬  bosome, thereby ensuring the close coupling of transcription and translation.  The progress of the ribosome prevents the formation of the 2 • 3 hairpin and  permits the formation of the 3 * 4 hairpin, which terminates transcription  (Fig. 28-1 7a). When tryptophan is scarce, however, the ribosome stalls at the  tandem UGG codons (which specify Trp) because of the lack of Trp—tRNA Trp .  As transcription continues, the newly synthesized segments 2 and 3 form a  hairpin because the stalled ribosome prevents the otherwise competitive  formation of the 1 • 2 hairpin (Fig. 28-17^). The 3 • 4 hairpin does not form,  allowing transcription to proceed past this region and into the remainder of  the trp operon. Thus, attenuation regulates trp operon transcription according  to the tryptophan supply.   The leader peptides of the five other amino acid—biosynthesizing operons  known to be regulated by attenuation are all rich in their corresponding amino  acid residues. For example, the E. coli his operon, which specifies enzymes  synthesizing histidine, has seven tandem His residues in its 16-residue leader  peptide, whereas the ilv operon, which specifies enzymes participating in  isoleucine, leucine, and valine biosynthesis, has five lie, three Leu, and six Val  residues in its 32-residue leader peptide. The leader transcripts of these oper¬  ons resemble that of the trp operon in their capacity to form two alternative  secondary structures, one of which contains a trailing transcription terminator.   We have just seen how the formation of secondary structure in a growing RNA  transcript can regulate gene expression though attenuation. The conforma¬  tional flexibility of mRNA also allows it to regulate genes by directly interact¬  ing with certain cellular metabolites, thereby eliminating the need for sensor  proteins such as the lac repressor, CAP, and the trp repressor.   In E. coli , the biosynthesis of thiamine pyrophosphate (TPP; Section 15-3B)  requires the action of several proteins whose levels vary according to the  cells need for TPP In at least two of the relevant genes the untranslated re¬  gions at the 5' end of the mRNA include a highly conserved sequence called  the thi box. The susceptibility of the thi box to chemical or enzymatic cleav¬  age, as Ronald Breaker showed, differs in the presence and absence of TPP,  suggesting that the RNA changes its secondary structure when TPP binds to  it (the binding of a metabolite by RNA is not unprecedented; synthetic  oligonucleotides known as aptamers bind certain molecules with high speci¬  ficity and affinity). The TPP-sensing mRNA element has been dubbed a  riboswitch.   The predicted secondary structure of the TPP-sensing riboswitch and its  proposed mechanism are shown in Fig. 28-18#. In the absence of TPP, the  mRNA assumes a conformation that allows a ribosome to begin translation.  In the presence of TPP, an alternative secondary structure masks its  Shine-Dalgarno sequence (Section 27-4A) so that the ribosome cannot initi¬  ate the mRNAs translation. Thus, the concentration of a metabolite can regu¬  late the expression of genes required for its synthesis. The X-ray structure of the  80-nt TPP-binding domain from the E. coli TPP-sensing riboswitch, deter¬  mined by Breaker and Dinshaw Patel, reveals an intricately folded RNA that  binds TPP in an extended conformation (Fig. 28-18^).   E. coli. (a) The predicted secondary structure of a 165-residue segment  at the 5' end of the thiM gene is shown in the absence (left) and  presence (right) of TPP. The TPP-binding conformation masks the  Shine-Dalgarno sequence (purple) required by the ribosome to initiate  translation at the AUG sequence (red) just downstream. [After Winkler,  W., Nahvi, A., and Breaker, R.R., Nature 419, 952 (2002).] (b) The  X-ray structure of the riboswitch’s TPP-sensing domain. The RNA is    drawn in cartoon form with its sugar-phosphate backbone  represented by an orange rod and its bases represented by paddles  with C green, N blue, and 0 red. The TPP is drawn in space-filling  form with C cyan, N blue, 0 red, and S yellow. Mg 2+ ions are  represented by lavender spheres. [Based on an X-ray structure by  Ronald Breaker, Yale University, and Dinshaw Patel, Memorial  Sloan-Kettering Cancer Center, New York, New York. PDBid 2GDI.]    • What is the regulatory advantage of  arranging genes in operons?   • Describe the regulation of the lac operon  by lac repressor and CAR   • Why does the lac operon include more  than one operator sequence?   • Compare regulation by the lac repressor  and the trp repressor.   • How does the conformation of mRNA  regulate gene expression by attenuation?   • Explain how riboswitches can turn gene  expression on and off.    Over 20 types of bacterial riboswitches have as yet been identified, includ¬  ing those that regulate the expression of enzymes involved in the synthesis of  coenzyme B 12 (Fig. 20-17), riboflavin (Fig. 14-13), and S-adenosylmethi-  onine (SAM; Fig. 21-18). These collectively regulate >2% of the genes in cer¬  tain bacteria. In some cases, metabolite binding to the mRNA controls the  formation of an internal transcription termination site so that transcription  beyond this site proceeds only when the metabolite is absent. In others, a ri-  bozyme that forms a portion of an mRNA is activated by the binding of a  metabolite to self-cleave, thereby inactivating the mRNA. Plants and fungi also  contain riboswitches. The fact that the interaction of riboswitches with their  effectors does not require the participation of proteins suggests that they are  relics of the RNA world (Box 24-3) and hence among the oldest regulatory  systems.   FIG. 28-19 Electron micrograph of a B  lymphocyte. The large body in the center of the  cell is the nucleus in which heterochromatin  (black) adheres to the inner nuclear membrane  and the transcriptionally active euchromatin  (gray) is more centrally located. The small black  body at the center of the nucleus is the  nucleolus. Also note the extensive rough  endoplasmic reticulum filling the cytoplasm of this  antibody-producing cell (Section 7-3). Compare  this figure with Fig. 1-8. [Don W. Fawcett/Photo  Researchers, Inc.]   • Chromatin structure, as influenced by chromatin-remodeling complexes, histone  modification, and DNA methylation, controls gene expression.   • Numerous proteins and DNA elements interact to regulate transcription initiation  in eukaryotes.   • Somatic recombination and hypermutation in immunoglobulin gene segments  contributes to antibody diversity.   The general principles that govern the expression of prokaryotic genes apply  also to eukaryotic genes: The expression of specific genes may be actively inhib¬  ited or stimulated through the ejfects of proteins that bind to DNA or RNA. As  in prokaryotes, the majority of known regulatory mechanisms act at the level  of gene transcription, but unlike prokaryotic control systems, the eukaryotic  mechanisms must contend with much larger amounts of DNA that is pack¬  aged in seemingly inaccessible structures. In this section, we describe a few of  the strategies whereby eukaryotic cells manage their genetic information.   The majority of DNA in multicellular organisms is not expressed. This in¬  cludes the portion of the genome that does not encode protein or RNA, as  well as the genes whose expression is inappropriate for a particular cell type.  Although nearly all the cells in an organism contain identical sets of DNA,  genes are expressed in a highly tissue-specific manner. For example, most pan¬  creatic cells synthesize and secrete digestive enzymes, but the pancreatic islet  cells synthesize insulin or glucagon instead (Section 22-2).   Nonexpressed DNA is typically highly condensed in a form known as het¬  erochromatin (Fig. 28-19). An extreme example of this is the complete in¬  activation of one of the two X chromosomes in female mammals (Box 28-2).  Transcriptionally active DNA, which is known as euchromatin, is less con¬  densed, presumably to provide access to the transcription machinery. In fact,  Harold Weintraub demonstrated that transcriptionally active chromatin is  more susceptible to digestion by pancreatic DNase I than is transcriptionally  inactive chromatin. Yet nuclease sensitivity apparently reflects a genes poten¬  tial for transcription rather than transcription itself.   In order for genetic information to be expressed, the transcription ma¬  chinery must gain access to the DNA, which is packaged in nucleosomes  (Section 24-5A). Not surprisingly, nucleosomes are not fixed structures but  can undergo remodeling. In addition, histones can be covalently modified to  alter how they interact with other cellular components.    Female mammalian cells contain two X chromosomes, whereas  male cells have one X and one Y chromosome. Female somatic  cells, however, maintain only one of their X chromosomes in a  transcriptionally active state. Consequently, both males and females  make approximately equal amounts of X chromosome-encoded  gene products, a phenomenon known as dosage compensation.   The inactive X chromosome is visible as a highly condensed  and darkly staining Barr body at the periphery of the cell nucleus.    In marsupials (pouched mammals such as kangaroos), the Barr  body is always the paternally inherited X chromosome, but in  placental mammals, one randomly selected X chromosome in  every somatic cell is inactivated when the embryo consists of only  a few cells. The progeny of each of these cells maintain the same  inactive X chromosome. Female placental animals are therefore  mosaics of cloned groups of cells in which the active X chromo¬  some is either paternally or maternally inherited. The calico cat, for  example, with its patches of black and yellow fur, is almost always  a female cat whose two X chromosomes specify the different  coat colors.   which has multiple regulatory sites together with the Xist gene,  which occurs only in placental mammals and is transcribed only  from the inactive X chromosome (Xi). The 25-kb Xist RNA binds  at or near XIC where it induces the heterochromatization of that  region of Xi via a process described below. The resulting het¬  erochromatin recruits additional Xist RNA to a neighboring site,  which in turn, causes its heterochromatization. This process propa¬  gates in both directions from XIC until Xi is “painted” over its  whole length by Xist RNA and the entire chromosome is het-    erochromatized and thereby transcriptionally inactivated at all sites  but Xist XIC is necessary and sufficient to initiate X inactivation  as is demonstrated by the observations that the insertion of XIC  into an autosome results in its inactivation and X chromosomes  that lack XIC are not inactivated.   Localized Xist RNA does not by itself inactivate Xi. However,  Xist RNA recruits enzyme systems that derivatize histones in a  way the inactivates their associated genes (histone modification is  discussed below). In addition, Xist RNA recruits the H2A variant  macroH2A1, which has a large C-terminal globular domain that  H2A lacks. Some or all of these changes are required for X chro¬  mosome inactivation and, moreover, are responsible for maintaining  Xi’s heterochromatic state in subsequent cell generations. In fact,  once an X chromosome has been inactivated, Xist RNA plays only  a minor role in the maintenance of this state.   What is the mechanism through which one X chromosome is  inactivated but not the other? XIC also contains a gene named  Tsix C Xist spelled backwards) that overlaps Xist but is transcribed  from the opposite DNA strand. Tsix RNA therefore forms a hybrid  RNA-DNA double helix with Xist that prevents its transcription  and vice versa. Thus Xi expresses Xist but not Tsix, whereas Xa  expresses Tsix but not Xist If Tsix is inactivated on one X chromo¬  some, that chromosome will be inactivated. Apparently, the balance  of transcription between Xist and Tsix selects which X chromo¬  some is inactivated.    Chromatin-Remodeling Complexes Shuffle Nucleosomes. Sequence-  specific DNA-binding proteins must gain access to their target DNAs before  they can bind to them. Yet nearly all DNA in eukaryotes is sequestered by  nucleosomes if not by higher order chromatin. How then do the proteins that  bind to DNA segments gain access to their target DNAs? The answer, which  has only become apparent since the mid-1990s, is that chromatin contains  ATP-driven complexes that remodel nucleosomes , that is, they somehow disrupt  the interactions between histones and DNA in nucleosomes to make the DNA  more accessible. This may cause the histone octamer to slide along the DNA  strand to a new location or even relocate to a different DNA strand. Thus,  these chromatin-remodeling complexes impose a “fluid” state on chromatin that  maintains the DNAs overall packaging but transiently exposes individual sequences  to interacting factors.    FIG. 28-20 Model of the RSC with a  bound nucleosome. The X-ray structure of the  nucleosome (with the proteins shown as orange  ribbons and the DNA backbone in light green)  was fitted into a cryo-EM-based image of the  RSC (gray). The two views show the back and  front of the complex. The DNA’s dyad (twofold)  axis is represented by a blue rod, and two  possible binding sites for the ATP-hydrolyzing  translocase subunit are indicated. [Courtesy of  Eva Nogales and Andrew Leschziner, University  of California at Berkeley.]    Chromatin-remodeling complexes consist of multiple subunits. The first  of them to be characterized was the yeast SWI/SNF complex, so called be¬  cause it is essential for mating type switching (SWI for swi tching defective)  and for growth on sucrose (SNF for mcrose ^on/ermenter). SWI/SNF, an  1150-kD complex of 11 different types of subunits, is essential for the expres¬  sion of only ~3% of yeast genes and is not required for cell viability. However,  a related complex named RSC (for remodels the structure of chromatin) is  — 100 times more abundant in yeast and is required for cell viability. RSC  shares two subunits with SWI/SNF, and many of their remaining subunits are  homologs, including their ATPase subunits. All eukaryotes contain multiple  chromatin-remodeling complexes.   Biochemical studies indicate that the RSC binds tightly to nucleosomes  in a 1:1 complex. Indeed, cryoelectron microscopy-based images of the yeast  RSC reveal that it has an irregular shape containing a large central cavity that  is remarkably complementary in size and shape to a single nucleosome core  particle (Fig. 28-20).   The simultaneous release of all of the many interactions holding DNA to  a histone octamer would require an enormous free energy input and hence is  unlikely to occur. How, then, do chromatin-remodeling complexes function?  Their various ATP-hydrolyzing translocase subunits share a region of homol¬  ogy with helicases (Section 25-2B), although they lack helicase activity.  Nevertheless, it seems plausible that, like helicases, chromatin-remodeling com¬  plexes “walk” up DNA strands as driven by ATP hydrolysis. If such a complex  were somehow tethered to a histone, this would put torsional strain on the  DNA in the nucleosome, thereby decreasing its local twist (DNA supercoiling  is discussed in Section 24-1C). The region of decreased twist could diffuse along  the DNA wrapped around the nucleosome, thereby transiently loosening the  histone octamers grip on a segment of DNA. The torsional strain might also  be partially accommodated as a writhe, which would lift a segment of DNA  off the nucleosome s surface. In either case, the resulting DNA distortion could  diffuse around the surface of the nucleosome in a wave that would locally and  transiently release the DNA from the histone octamer as it passed (Fig. 28-21).  Multiple cycles of ATP hydrolysis would send multiple DNA-loosening waves  around the nucleosome, thereby sliding the nucleosome along the DNA and  providing DNA-binding proteins access to the DNA.   HMG Proteins Are Architectural Proteins That Help Regulate Gene  Expression. The variation of a given genes transcriptional activity according  to cell type indicates that chromosomal proteins participate in the gene acti¬  vation process. Yet histones 5 chromosomal abundance and lack of variety make  it highly unlikely that they have the specificity required for this role. Among  the most common nonhistone protein components of chromatin are the mem¬  bers of the high mobility group (HMG), so named because of their high  electrophoretic mobilities in polyacrylamide gels. These highly conserved, low   The remodeling  complex binds to  DNA and  translocates the  DNA so that it is  undertwisted ahead   The undertwisted  DNA moves in a  one-dimensional  wave around the  nucleosome.   chromatin-remodeling complexes. The chromatin-remodeling movement of a fixed point on the DNA (yellow star). [After a drawing   complex (green) couples the free energy of ATP hydrolysis to the by Saha, A., Wittmeyer, J., and Cairns, B.R., Genes Dev. 16, 2120   translocation and concomitant twisting of the DNA in the nucleosome (2002).]    molecular mass (<30 kD) proteins, which have the unusual amino acid com¬  position of —25% basic side chains and 30% acidic side chains, are relatively  abundant with —1 HMG molecule per 10 to 15 nucleosomes.   The yeast HMG protein known as NHP6A contains an —80-residue  structural motif called an HMG box. Its NMR structure in complex with  DNA, determined by Juli Feigon, reveals that it causes the DNA to bend by  as much as 70° toward its major groove (Fig. 28-22). Such proteins presum¬  ably facilitate the binding of other regulatory proteins to the DNA and are   FIG. 28-22 NMR structure of yeast  NHP6A protein in complex with a 15-bp  DNA. The protein is drawn in ribbon form (cyan)  and the DNA is drawn in stick form colored  according to atom type (C green, N blue, O red,  and P yellow). The three helices of the HMG  box form an L shape, with the inside of the L  inserted into the DNA’s minor groove so as to  bend the DNA by —70° toward its major groove.  [Based on an NMR structure by Juli Feigon,  University of California at Los Angeles. PDBid  1J5NJ    FIG. 28-23 Histone modifications on the  nucleosome core particle. Posttranslational  modification sites on the four core histones are  indicated with colored symbols. The added  groups are ac = acetyl, me = methyl,  ph = phosphoryl, pr = propionyl, rib = ADP-ribose,  and Ub = ubiquitin. Cit represents citrulline  (deiminated arginine). Note that some residues  can be modified in multiple ways. This diagram is  a composite; not all modifications occur in all  organisms. [After a diagram by Ali Shilatifard,   ARTKQTARKSTGGKAPRKQLATKAARKS APATGGVK* • *K* • *K  2 3 4 8 91011 14 1718 23 26 2728 36 56 79    therefore described as architectural proteins. Other HMG proteins differ in  structure but can also alter DNA conformation by inducing it to bend,  straighten, unwind, or form loops. Some HMG proteins compete with his¬  tones for DNA, thereby altering nucleosomal structure.    Histones Are Covalently Modified. The posttranslational modifications to  which core histones are subject include the acetylation of specific Lys side  chains, the methylation of specific Lys and Arg side chains, the phosphoryla¬  tion of specific Ser and Thr side chains, and the ubiquitination of specific Lys  side chains (Fig. 28-23). Moreover, Lys side chains can be mono-, di-, and  trimethylated and Arg side chains can be mono- and both symmetrically and  asymmetrically dimethylated. All of these modifications can be undone by the  action of specific enzymes. Less frequent modifications include ADP-ribosylation  of Glu, addition of a propionyl group to Lys, and the conversion of Arg  residues to citrullyl residues.   The core histones’ N-terminal tails, as we have seen (Section 24-5A), are  implicated in stabilizing the structures of both core nucleosomes and higher-  order chromatin. Nearly all of the modifications listed above, except methyla-  tions, reduce (make more negative) the electronic charge of their attached side  chains and hence are likely to weaken histone-DNA interactions so as to pro¬  mote chromatin decondensation, although as we will see, this is not always  the case. Methyl groups, in contrast, increase the basicity and hydrophobicity  of the side chains to which they are linked and hence tend to stabilize chro¬  matin structure. Modified histone tails also interact with specific chromatin-  associated nonhistone proteins in a way that changes the transcriptional  accessibility of their associated genes. Internucleosome and histone-histone in¬  teractions are also likely to be affected by covalent modification of the glob¬  ular cores of the histone proteins, not just their tails.   The characterization of a variety of histone tail modifications led David Allis  to hypothesize that there is a histone code in which specific modifications evoke  certain chromatin-based functions and that these modifications act sequentially or in  combination to generate unique biological outcomes. For example, Fig. 28-24 indi¬  cates the distribution of the various histone marks (modifications) on nucleo¬  somes associated with a typical transcriptionally active human gene. It can be seen  from Figs. 28-23 and 28-24 that there are a vast number of possible combina¬  tions of histone marks, although the way they interact is largely unknown.    Histone Acetyltransferases Are Components of Multisubunit Transcriptional  Coactivators. Histone Lys side chains are acetylated in a sequence-specific    FIG. 28-24 Chromatin map of a typical transcriptionally active human gene. The   height of each colored strip at a given position indicates the prevalence of the  corresponding histone mark(s) on the nucleosome at that position. For example, H3K4me3  (trimethylated H3 Lys 4) and H2BK5ac (acetylated H2B Lys 5) are associated with the  nucleosomes flanking the transcriptional start site (squared off arrow) as well as portions of  enhancers and insulators (Section 28-3B), whereas H3R2me2a (asymmetrically  dimethylated H3 Arg 2) is associated with nucleosomes occupying the downstream region  of the gene’s protein encoding segment. The positions of nucleosomes containing the H2A  variant H2A.Z ( yellow ) are also indicated. [Modified from a drawing by Oliver Rando,  University of Massachusetts Medical School, and Howard Chang, Stanford University.]    manner by enzymes known as histone acetyltransferases (HATs), all of which  employ acetyl-CoA as their acetyl group donor:   Most if not all HATs function in vivo as members of often large (10-20 sub¬  units) multisubunit complexes, many of which were initially characterized as  transcriptional regulators. For example, TAF1, the largest subunit of the gen¬  eral transcription factor TFIID (TAF for TBP-^ssociated /actor, where TBP  is TATA-binding protein; Section 26-2C), is a HAT. Moreover, many HAT  complexes share subunits. Thus, the HAT complex named SAGA contains  TAF5, TAF6, TAF9, TAF10, and TAF12 as does the PCAF complex with  the exception that TAF5 and TAF6 are replaced in the PCAF complex by their  close homologs PAF65P and PAF65ot. Portions of TAF6, TAF9, and TAF12  are structural homologs of histones H3, H4, and H2B, respectively (Section  24-5). Consequently, these TAFs probably associate to form an architectural  element that is common to TFIID, SAGA, and the PCAF complex and hence  these complexes are likely to interact with TBP in a similar manner. The var¬  ious HAT complexes presumably target their component HATs to the pro¬  moters of active genes.   The X-ray structure of the HAT domain of Tetrahymena thermophila  GCN5 (residues 48-210 of the 418-residue protein) in complex with a  bisubstrate inhibitor was determined by Ronen Marmorstein. The bisubstrate    FIG. 28-25 X-Ray structure of Tetrahymena GCN5 in complex  with an inhibitor, (a) The bisubstrate inhibitor, a peptide-CoA conjugate,  consists of CoA covalently linked from its S atom via an isoproprionyl  group to the side chain of Lys 14 of the 20-residue N-terminal segment  of histone H3. (b) The protein is drawn in ribbon form in pink with its core  region magenta. The peptide-CoA conjugate is drawn in stick form colored  according to atom type (histone C cyan, isopropionyl group C orange, CoA  C green, N blue, 0 red, S yellow, and P gold). [Part b based on an X-ray  structure by Ronen Marmorstein, The Wistar Institute, Philadelphia,  Pennsylvania. PDBid 1MID.]    inhibitor (Fig. 28-25 d) consists of CoA covalently linked from its S atom via  an isopropionyl group (which mimics an acetyl group) to the side chain of Lys  14 (the acetyl group acceptor) of the 20-residue N-terminal segment of histone  H3. The enzyme (Fig. 28-25 £) is deeply clefted and contains a core region  common to all HATs of known structure (magenta in Fig. 28-25 B) that con¬  sists of a three-stranded antiparallel (3 sheet connected via an a helix to a fourth  (3 strand that forms a parallel interaction with the (3 sheet. Only six residues  of the histone tail, Gly 12 through Arg 17, are visible in the X-ray structure.  The CoA moiety binds in the enzymes cleft such that it is mainly contacted  by core residues. The comparison of this structure with other GCN5-containing  structures indicates that the cleft has closed down about the CoA moiety.    Bromodomains Recruit Coactivators to Acetylated Lys Residues in Histone  Tails. The different patterns of histone acetylation required for different func¬  tions (the histone code) suggest that the function of histone acetylation is more  complex than merely attenuating the charge-charge interactions between the  cationic histone N-terminal tails and anionic DNA. In fact, specific acetylation  patterns appear to be recognized by protein modules of transcriptional coacti¬  vators in much the same way that specific phosphorylated sequences are rec¬  ognized by protein modules such as the SH2 domain that mediates signal  transduction via protein kinase cascades (Section 13-2B). Thus, nearly all HAT-  associated transcriptional coactivators contain ~ 110-residue modules known as  bromodomains that specifically bind acetylated Lys residues on histones. For  example, GCN5 essentially consists of a HAT domain followed by a bromo-  domain, whereas TAF1 consists mainly of an N-terminal kinase domain followed  by a HAT domain and two tandem bromodomains.   The X-ray structure of human TAFls double bromodomain (residues  1359-1638 of the 1872-residue protein), determined by Robert Tjian, reveals  that it consists of two nearly identical antiparallel four-helix bundles (Fig. 28-26).  A variety of evidence, including NMR structures of single bromodomains in  complex with their target acetyl-Lys-containing peptides, indicates that the    acetyl-Lys-binding site of each bromodomain occurs in a deep hydrophobic  pocket that is located at the end of its four-helix bundle opposite its N- and  C-termini. The double bromodomains two binding pockets are separated by  ~25 A, which makes them ideally positioned to bind two acetyl-Lys residues  that are separated by 7 or 8 residues. In fact, the N-terminal tail of histone  H4 contains Lys residues at positions 5, 8, 12, and 16 (Fig. 28-23), whose  acetylation is correlated with increased transcriptional activity. Moreover, the  36-residue N-terminal peptide of histone H4, when fully acetylated, binds to  the TAF1 double bromodomain in 1:1 ratio with 70-fold higher affinity than  to single bromodomains but fails to bind when it is unacetylated.   The foregoing structure suggests that the TAF1 bromodomains serve to tar¬  get TFIID to promoters that are within or near nucleosomes [in contrast to  the previously held notion that TFIID targets PICs (preinitiation complexes)  to nucleosome-free regions]. Tjian has therefore postulated that the transcrip¬  tional initiation process begins with the recruitment of a HAT-containing coac¬  tivator complex by an upstream DNA-binding protein (Fig. 28-27). The HAT  could then acetylate the N-terminal histone tails of nearby nucleosomes, which  would recruit TFIID to an appropriately located promoter via the binding of  its TAF1 bromodomains to the acetyl-Lys residues. Moreover, the TAF1 HAT  activity could acetylate other nearby nucleosomes, thereby initiating a cascade  of acetylation events that would render the DNA template competent for tran¬  scriptional initiation.   Histone acetylation is a reversible process. The enzymes that remove the  acetyl groups from histones, the histone deacetylases (HDACs), promote  transcriptional repression and gene silencing. Eukaryotic cells from yeast to  humans typically contain numerous different HDACs; 10 HDACs have been  identified in yeast and 17 in humans.   Histone Methylation May Silence or Activate Genes. Histone methyla-  tion at both the Lys and Arg side chains of histone H3 and H4 N-termi¬  nal tails (Fig. 28-23) tends to silence the associated genes by inducing the   FIG. 28-26 X-Ray structure of the human  TAF1 double bromodomain. Each bromodomain  consists of an antiparallel four-helix bundle  whose helices are colored, from N- to C-termini,  red, yellow, green, and blue, with the remaining  portions of the protein orange. The acetyl-Lys  binding sites occupy deep hydrophobic pockets  at the end of each four-helix bundle opposite its  N- and C-termini. [Based on an X-ray structure  by Robert Tjian, University of California at  Berkeley. PDBid 1 EQFJ    FIG. 28-27 Simplified model for the  assembly of a transcriptional initiation complex  on chromatin-bound templates. Here the DNA  is represented by a blue worm, the histone  octamers around which the DNA wraps to form  nucleosomes are shown as gray spheres, and  their N-terminal histone tails are drawn as short  beaded chains with the black and red beads  representing unacetylated and acetylated Lys  residues. The transcription initiation site is  represented by the orange segment of the DNA  from which the green squared-off arrow points  downstream, (a) The process begins by the  recruitment of a HAT-containing transcriptional  coactivator complex (green) through its  interactions with a DNA-binding activator protein  (pink) that is bound to an upstream enhancer  (yellow). The HAT coactivator complex is thereby  positioned to acetylate the N-terminal tail on  nearby nucleosomes (curved arrows), (b) The  binding of its TAFI’s bromodomains to the  acetylated histone tails could then help recruit  TFIID (pink) to a nearby TATA box (gray). Further  acetylation of nearby histone tails by the TAFI’s  HAT domain could help recruit other basal  factors (yellow-green) and RNAP II (green) to  the promoter, thus inducing PIC formation.   Chapter 28 Regulation of Gene Expression formation of heterochromatin, although many forms of methylation are   instead associated with active genes (Fig. 28-24). The enzymes mediating  histone methylation, the histone methyltransferases (HMTs), all utilize  5-adenosylmethionine (SAM) as their methyl donor. Thus, the lysine HMTs,  the most extensively characterized HMTs, catalyze the following reaction:    These enzymes all have a so-called SET domain, which contains their cat¬  alytic sites.   The human lysine HMT named SET7/9 monomethylates Lys 4 of his¬  tone H3. The X-ray structure of the SET domain of SET7/9 (residues  108-366 of the 366-residue protein) in complex with SAM and the N-terminal  decapeptide of histone H3 in which Lys 4 is monomethylated was determined  by Steven Gamblin. Interestingly, SAM and the peptide substrate bind to op¬  posite sides of the protein (Fig. 28-28). However, there is a narrow tunnel  through the protein into which the Lys 4 side chain is inserted such that its  amine group is properly positioned for methylation by SAM. The arrangement  of the hydrogen bonding acceptors for the Lys amino group stabilize the  methyl-Lys side chain in its observed orientation about the C £ —bond, thus  sterically precluding the methyl-Lys group from assuming a conformation in  which it could be further methylated by SAM.   Methylated histones are bound by several kinds of protein domains that rec¬  ognize specific Lys residues with some preference for their mono-, di-, or trimethyl  forms. The so-called chromodomain and the motif known as a PHD finger  (for ydant /mmeo^/omain) both include binding sites for methylated Lys  residues that can be characterized as aromatic cages (Fig. 28-29). Interestingly, the  methyl-Lys-binding proteins often include additional histone code-reading mo¬  tifs such as a bromodomain, or a histone-modifying enzyme such as a deacetylase.   The chromodomain-containing heterochromatin protein 1 (HP1),  which binds dimethylated H3 Lys 9, contributes to gene silencing. The bound  HP1 recruits the HMT Suv39h, which methylates nearby nucleosomes at  their H3 Lys 9 residues, thereby recruiting additional HP1, etc. This mecha¬  nism could explain why heterochromatin has a tendency to spread so as to si¬  lence neighboring genes. In transcriptionally silent chromatin, the DNA itself  may also be methylated.   Methyltransferases Flip Their Target Bases Out of the DNA Double Helix.   The methylation of DNA, as we have seen (Box 25-4), switches off eukary¬  otic gene expression. Eukaryotic DNAs are methylated only on their cytosine  residues to form 5-methylcytosine (m 5 C) residues, a modification that occurs  largely on the palindromic CpG islands that commonly occur in the upstream    FIG. 28-28 X-Ray structure of the human histone methyltransferase SET7/9 in  complex with SAM and the histone H3 N-terminal decapeptide with its Lys 4  monomethylated. The protein is represented by its surface diagram colored according to  charge (blue most positive, red most negative, and gray neutral) and the H3 decapeptide is  represented by a green ribbon. Note the narrow tunnel through the protein in which the  methyl-Lys side chain is inserted. The inset above shows a close-up of this Lys access  channel containing the methyl-Lys side chain (green) as viewed from the opposite side of  the protein from the SAM-binding site. [Courtesy of Steven Gamblin, National Institute for  Medical Research, London, U.K. PDBid 109SJ    FIG. 28-29 Binding sites for histone trimethyl-Lys residues, (a) H3 trimethyl-Lys 9  bound to the chromodomain of Drosophila heterochromatin protein 1 (HP1) and (b) H3  trimethyl-Lys 4 bound to a human PHD finger. The H3 trimethyl-Lys residues are drawn  with C green and N blue and with the peptide backbone yellow. The aromatic side chains  of HP1 and the PHD finger that cage the trimethylammonium group are drawn with C  cyan, N blue, and 0 red. The dot surfaces delineate the van der Waals surfaces of the  aromatic side chains and the trimethylammonium groups. [Part a based on an X-ray  structure by Sepideh Khorasanizadeh, University of Virginia. PDBid 1KNE. Part b based on  an X-ray structure by David Allis, Rockefeller University, and Dinshaw Patel, Memorial  Sloan-Kettering Cancer Center, New York, New York. PDBid 2F6J.]    promoter regions of genes (Section 26-2B). The enzymes that mediate these  reactions, the DNA methyltransferases (DNA MTases), all use SAM as their  methyl donor. Cells also contain demethylases, which catalyze the reaction    In DNA MTase reactions, SAM reacts with the enzymes targeted cyto¬  sine base to form a tetrahedral intermediate in which the cytosine C5 is sub¬  stituted by both its original hydrogen atom and the donated methyl group.  This requires that the attacking methyl group approach C5 from above or be¬  low the cytosine ring. Yet the faces of nucleic acid bases are inaccessible within  the DNA double helix. Consequently, a DNA MTase must induce its target  cytosine base to flip out of the double helix to where it can be methylated by  SAM. Such a flipped-out intermediate is observed in the X-ray structure of  the DNA MTase from Haemophilus haemolyticus (M.Hhal), determined by  Richard Roberts and Xiaodong Cheng, in which the target cytosine base has  been replaced by 5-fluorocytosine (Fig. 28-30). Although the normally pres¬  ent C5—H atom is eliminated from the foregoing tetrahedral intermediate as    FIG. 28-30 X-Ray structure of the M.Hhal DNA methyltransferase. The protein,  represented by a semitransparent ribbon, is shown in complex with S-adenosylhomocysteine  and a dsDNA containing a methylated 5-fluorocytosine base at the enzyme’s target site.  The DNA is drawn in stick form colored according to atom type (C green, N blue, 0 red,  and P orange). Its methylated 5-fluorocytidine residue (C atoms cyan) has swung out of  the DNA helix into the enzyme’s active site pocket, where its C6 forms a covalent bond  with the S atom of an enzyme Cys residue (C atoms magenta and S yellow). The methyl  group and a fluorine atom at C5 (which prevents the methylation reaction from going to  completion) are represented by magenta and yellow-green spheres, respectively. The  position of the flipped-out cytosine base in the DNA double helix is occupied by the side  chain of a Gin residue (shown in space-filling form with C yellow), which hydrogen bonds  to the “orphaned” guanine base. The S-adenosylhomocysteine, which is SAM without its  methyl group, is drawn in stick form with its C atoms pink. [Based on an X-ray structure by  Richard Roberts, New England Biolabs, Beverly, Massachusetts, and Xiaodong Cheng, Cold  Spring Harbor Laboratory, Cold Spring Harbor, New York. PDBid 1MHTJ   pattern of methylation on a parental DNA strand  induces the corresponding methylation pattern in  the complementary strand. In this way, a stable  methylation pattern may be maintained in a cell  line.    H + to yield the methylation reactions m 5 C product, the high electronegativ¬  ity of fluorine prevents the C5—F atom from being eliminated as F + , thereby  trapping the tetrahedral intermediate on the enzyme. Certain DNA repair en¬  zymes that act on individual DNA bases also operate by flipping their target  base out the DNA helix (Section 25-5A).   DNA Methylation in Eukaryotes Is Self-Perpetuating. The palindromic na¬  ture of DNA methylation sites in eukaryotes permits the methylation pattern  on a parental DNA strand to direct the generation of the same pattern in its  daughter strand (Fig. 28-31). This maintenance methylation would result in  the stable “inheritance” of a methylation pattern in a cell line and hence cause  these cells to all have the same differentiated phenotype. Such changes to the  genome are described as being epigenetic (Greek: epi, on or beside) because  they provide an additional layer of information that specifies when and where  specific portions of the otherwise fixed genome are expressed [epigenetic  changes that we have already encountered are the lengthening of telomeres in  germ cells (Section 25-3C) and the inactivation of a specific X chromosome  in female mammals (Box 28-2)]. Epigenetic characteristics, as we will see, are  not bound by the laws of Mendelian inheritance.   There is considerable experimental evidence favoring the existence of  maintenance methylation, including the observation that artificially methyl¬  ated viral DNA, when taken up by eukaryotic cells, maintains its methyla¬  tion pattern for at least 30 cell generations. Maintenance methylation in  mammals appears to be mediated mainly by DNMT1 protein, which has a  strong preference for methylating hemimethylated substrate DNAs. In con¬  trast, prokaryotic DNA MTases such as M.Hhal do not differentiate between  hemimethylated and fully methylated substrate DNAs. The importance of  maintenance methylation is demonstrated by the observation that mice that  are homozygous for the deletion of the DNMTl gene die early in embryonic  development.   The pattern of DNA methylation in mammals varies in early embryonic  development. DNA methylation levels are high in mature gametes (sperm and  ova) but are nearly eliminated by the time a fertilized ovum has become a  blastocyst (a hollow ball of cells, the stage at which the embryo implants into  the uterine wall). After that stage, however, the embryos DNA methylation  levels globally rise until, by the time the embryo has reached the developmen¬  tal stage known as a gastrula, its DNA methylation levels have risen to adult  levels, where they remain for the lifetime of the animal. This de novo (new)  methylation appears to be mediated by two DNA MTases distinct from  DNMTl named DNMT3a and DNMT3b. An important exception to this  remethylation process is that the CpG islands of germline cells (cells that give  rise to sperm or ova) remain unmethylated. This ensures the faithful transmis¬  sion of the CpG islands to the succeeding generation in the face of the strong  mutagenic pressure of m 5 C deamination (which yields T, a mutation that mis¬  match repair occasionally fails to correct; Box 25-5). In female mammals, the  inactivation of the same X chromosome from cell generation to cell genera¬  tion is, at least in part, preserved by maintenance methylation (Box 28-2).   The change in DNA methylation levels (epigenetic reprogramming) dur¬  ing embryonic development suggests that the pattern of genetic expression dif¬  fers in embryonic and somatic cells. This explains the observed high failure  rate in cloning mammals (sheep, mice, cattle, etc.) by transferring the nucleus  of an adult cell into an enucleated oocyte (immature ovum). Few of these an¬  imals survive to birth, many of those that do so die shortly thereafter, and  most of the ~1% that do survive have a variety of abnormalities, most promi¬  nently an unusually large size. However, the survival of any embryos at all in¬  dicates that the oocyte has the remarkable capacity to epigenetically reprogram  somatic chromosomes (although it is rarely entirely successful in doing so) and  that mammalian embryos are relatively tolerant of epigenetic abnormalities.    Presumably, the reproductive cloning of humans from adult nuclei would re¬  sult in similar abnormalities and for this reason (in addition to social and eth¬  ical prohibitions) should not be attempted.   Genomic Imprinting Results from Differential DNA Methylation. It has been  known for thousands of years that maternal and paternal inheritance can dif¬  fer. For example, a mule (the offspring of a mare and a male donkey) and a  hinny (the offspring of a stallion and a female donkey) have obviously differ¬  ent physical characteristics, a hinny having shorter ears, a thicker mane and  tail, and stronger legs than a mule. This is because, in mammals only, certain  maternally and paternally supplied genes are differentially expressed, a phe¬  nomenon termed genomic imprinting. The genes that are subject to genomic  imprinting are, as Rudolph Jaenisch has shown, differentially methylated in  the two parents during gametogenesis, and the resulting different methylation  patterns are resistant to the wave of demethylation that occurs during the for¬  mation of the blastocyst and to the wave of de novo methylation that occurs  thereafter.   that an embryo derived from the transplantation of two male or two fe¬  male pronuclei into an ovum fails to develop (pronuclei are the nuclei of ma¬  ture sperm and ova before they fuse during fertilization). Inappropriate  imprinting is also associated with certain diseases. For example, Prader-Willi  syndrome (PWS), which is characterized by the failure to thrive in infancy,  small hands and feet, marked obesity, and variable mental retardation, is  caused by a >5000-kb deletion in a specific region of the paternally inherited  chromosome 15. In contrast, Angelman syndrome (AS), which is manifested  by severe mental retardation, a puppetlike ataxic (uncoordinated) gait, and  bouts of inappropriate laughter, is caused by a deletion of the same region  from the maternally inherited chromosome 15. These syndromes are also ex¬  hibited by those rare individuals who inherit both their chromosomes 15 from  their mothers for PWS and from their fathers for AS. Evidently, certain genes  on the deleted chromosomal region must be paternally inherited to avoid PWS  and others must be maternally inherited to avoid AS. Several other human  diseases are also associated with either maternal or paternal inheritance or lack  of it. Aberrant epigenetic programming may also play a role in tumor growth,  as cancer cells often exhibit abnormal patterns of DNA methylation.   In addition to the six general transcription factors described in Section 26-2C,  eukaryotic cells contain a host of other proteins that interact with DNA and/or  other transcription factors to stimulate or repress the transcription of class II  genes (genes that are transcribed by RNAP II). Some of these proteins bind to  any available DNA containing their target sequences, whereas others must be  activated and deactivated, often as part of a signal transduction pathway (Section  13-2B). These regulatory proteins are known as activators and repressors, and  their target DNA sites are known as enhancers and silencers, respectively.   An enhancer typically is not essential for transcription but greatly increases  its rate (conversely, silencers repress the transcription of their associated genes).  Unlike promoters, which are necessarily located a short distance from the tran¬  scription start site, enhancers (or silencers) need not have fixed positions and ori¬  entations (Section 26-2B). For example, William Rutter linked the upstream  (5'-flanking) sequences of either the insulin or the chymotrypsin gene to the  sequence encoding chloramphenicol acetyltransferase (CAT), an easily as¬  sayed enzyme not normally present in eukaryotic cells. A plasmid containing  the insulin gene sequences elicits expression of the CAT gene only when in¬  troduced into cultured cells that normally produce insulin. Likewise, the chy¬  motrypsin recombinants are active only in chymotrypsin-producing cells.   Chapter 28 Regulation of Gene Expression Dissection of the insulin gene control sequence indicates that the enhancer lies   between positions 2103 and 2333 and, in insulin-producing cells only, it stim¬  ulates the transcription of the CAT gene with little regard to its position and  orientation relative to the promoter. Because so many enhancers are located  upstream of transcribed sequences, the proteins that bind to them are often  called upstream transcription factors.   the PIC. How do upstream transcription factors stimulate (or inhibit) tran¬  scription? Evidently\ when these proteins bind to their target DNA sites in the  vicinity of a PIC (in some cases, many thousands of base pairs distant), they some¬  how activate (or repress) its component RNAP II to initiate transcription.  Transcription factors may bind cooperatively to each other and/or the PIC,  thereby synergistically stimulating (or repressing) transcriptional initiation.  Indeed, molecular cloning experiments indicate that many enhancers and si¬  lencers consist of segments (modules) whose individual deletion reduces but  does not eliminate enhancer/silencer activity. Such complex arrangements pre¬  sumably permit transcriptional control systems to respond to a variety of stimuli  in a graded manner. In some cases, however, several transcription factors to¬  gether with so-called architectural proteins cooperatively assemble on an  ~ 100-bp enhancer to form a multisubunit complex known as an enhanceo-  some, in which the absence of a single subunit all but eliminates its ability to  stimulate transcriptional initiation at the associated promoter. Thus, enhanceo-  somes function more like on/off switches rather than providing a graded re¬  sponse. Enhanceosomes may also contain coactivators and/or corepressors,  proteins that do not bind to DNA but, rather, interact with proteins that do  so to activate or repress transcription.   The functional properties of many upstream transcription factors are sur¬  prisingly simple. They typically consist of (at least) two domains:   1. A DNA-binding domain that specifically binds to the proteins target  DNA sequence (several such domains are described in Section 24-4C).   2. A domain containing the transcription factors activation function.  Sequence analysis indicates that many of these activation domains  have conspicuously acidic surface regions whose negative charges, if  mutationally increased or decreased, respectively raise or lower the tran¬  scription factors activity. This suggests that the associations between  these transcription factors and a PIC are mediated by relatively non¬  specific electrostatic interactions rather than by conformationally more  demanding hydrogen bonds. Other types of activation domains have  also been characterized, including those with Gin-rich regions and  those with Pro-rich regions.   The DNA-binding and activation functions of eukaryotic transcription  factors can be physically separated (which is why they are thought to occur  on different domains). In fact, a genetically engineered hybrid protein, con¬  taining the DNA-binding domain of one transcription factor and the activa¬  tion domain of a second, activates the same genes as the first transcription  factor. Moreover, it makes little functional difference as to whether the acti¬  vation domain is placed on the N-terminal side of the DNA-binding domain  or on its C-terminal side. This geometric permissiveness in the binding be¬  tween the activation domain and its target protein is also indicated by the ob¬  servation that transcription factors are largely insensitive to the orientations  and positions of their corresponding enhancers relative to the transcriptional  start site. Of course, the DNA between an enhancer and its distant transcrip¬  tional start site must be looped around for an enhancer-bound transcription fac¬  tor to interact with the promoter-bound PIC (Section 26-2C).   The synergy (cooperativity) of multiple transcription factors in initiating  transcription may be understood in terms of a simple recruitment model.    Suppose an enhancer-bound transcription factor increases the affinity with  which a PIC binds to the enhancers associated promoter so as to increase the  rate at which the PIC initiates transcription there by a factor of 10. Then, if  another transcription factor binding to a different enhancer subsite likewise  increases the initiation rate by a factor of 20, both transcription factors act¬  ing together will increase the initiation rate by a factor of 200. In this way, a  limited number of transcription factors can support a much larger number of tran¬  scription patterns. Transcriptional activation, according to this model, is essen¬  tially a mass action effect: The binding of a transcription factor to an enhancer  increases the transcription factors effective concentration at the associated pro¬  moter (the DNA holds the transcription factor in the vicinity of the promoter),  which consequently increases the rate at which the PIC binds to the promoter.  This explains why a transcription factor that is not bound to DNA (or even  lacks a DNA-binding domain) inhibits transcriptional initiation. Such unbound  transcription factors compete with DNA-bound transcription factors for their  target sites and thereby reduce the rate at which the PIC is recruited to the as¬  sociated promoter. This phenomenon, which is known as squelching, is ap¬  parently why transcription factors in the nucleus are almost always bound to  inhibitors unless they are actively engaged in transcriptional initiation.   Mediator Links Transcriptional Activators and RNAP II. Eukaryotic genomes  encode as many as several thousand transcriptional regulators for class II genes  (e.g., Fig. 28-3). However, activators fail to stimulate transcription by a recon¬  stituted PIC in vitro. Evidently, an additional factor is required to do so.  Indeed , genetic studies in yeast by Roger Kornberg led him to discover an ~ 21-  subunit , ~ 1200-kD complex named Mediator \ whose presence is required for  transcription from nearly all class II gene promoters in yeast. Mediator, which is  therefore considered to be a coactivator, binds to the C-terminal domain  (CTD) of RNAP IPs (T subunit (Section 26-2A). Further investigations re¬  vealed that multicellular organisms contain multisubunit complexes that func¬  tion similarly to yeast Mediator and which share many of their numerous  subunits. Moreover, many of their subunits are related, albeit distantly, to those  of yeast Mediator. Mediators apparently function as adaptors that bridge DNA-  bound transcriptional regulators and RNAP II so as to influence (induce or in¬  hibit) the formation of a stable PIC at the associated promoter. They thereby  function to integrate the various signals implied by the binding of these transcrip¬  tional regulators to their target DNAs.   The EM-based image of the yeast RNAP II-Mediator complex (Fig. 28-32),  determined by Francisco Asturias, reveals that Mediator consists of four  domains known as the head, middle, arm, and tail domains. The head do¬  main interacts closely with the RNAP II, although >75% of RNAP IPs    FIG. 28-32 EM-based structure of yeast  Mediator in complex with RNAP II. Mediator’s  structural modules are in different colors: head  (red), middle (green), tail (tan), and arm (purple).  The RNAP II is represented by its surface  diagram (orange). Note that Mediator binding  leaves RNAP M’s active site cleft fully accessible.  [Courtesy of Francisco Asturias, The Scripps  Research Institute, La Jolla, California.]   Chapter 28 Regulation of Gene Expression surface remains accessible for interaction with other components of the PIC.   The tail domain, however, appears not to contact RNAP II at all. Includ¬  ing Mediator, the transcriptional machinery for class II genes comprises  nearly —73 polypeptides with an aggregate molecular mass of —3.7 million  D. Nevertheless, as we have seen (Section 28-3A), this ribosome-sized assem¬  bly (the eukaryotic ribosome has a molecular mass of —4.2 million D; Table  27-5) requires considerable assistance from other large particles to gain access  to the DNA in chromatin.   Insulators Limit the Effects of Enhancers and Stop Heterochromatin  Spreading. Enhancers function independently of their position relative to the  promoter. So what prevents an enhancer from affecting the transcription of all  the genes in its chromosome? Conversely, heterochromatin, as we have seen  (Section 28-3A), appears to be self-nucleating. What prevents heterochromatin  from spreading into neighboring segments of euchromatin? Experiments in  which DNA sequences near Drosophila genes were rearranged revealed that  short (<2 kb) segments of DNA known as insulators define the boundaries  of functional units for transcription. Inserting an insulator between a gene and  its upstream enhancer blocks the effect of the enhancer on transcription.  Insulators may also prevent heterochromatin from spreading. For example, the  HS4 insulator in the chicken (3-globin gene cluster recruits HATs that acetyl-  ate H3 Lys 9 on nearby nucleosomes (which is associated with transcriptional  activity; Fig. 28-24), thereby blocking their methylation (which induces het¬  erochromatin formation). The mechanism of action of insulators is enigmatic.  Presumably, it is not the insulators themselves but the proteins that bind to  them that form the active insulator elements. Indeed, the HAT-recruiting ac¬  tivity of HS4 is distinct from its enhancer-blocking function, which is medi¬  ated by the binding of an 11-zinc finger protein named CTCF (for  CC CTC-binding /actor) to a different subsite of HS4 than that to which  HATs bind.   Many Signal Transduction Pathways Activate Transcription Factors. A variety  of signaling pathways, including some involving steroid hormones, heterotrimeric  G proteins, receptor tyrosine kinases (RTKs), and phosphoinositide cascades  (Chapter 13), result in the activation (or inactivation) of transcription factors.  In this way, extracellular factors such as hormones can influence gene expres¬  sion inside a cell. We have already seen that the Ras signaling cascade (Fig.  13-7) results in the phosphorylation of several transcription factors, including  Fos, Jun, and Myc, thereby modulating their activities. The sterol regulatory  element binding protein (SREBP), which regulates the expression of the genes  involved in cholesterol biosynthesis by binding to their sterol regulatory ele¬  ments (SREs; Section 20-7B), is also an inducible transcription factor. In the  following paragraphs, we discuss two additional examples of signaling path¬  ways that activate transcription factors.   The JAK-STAT Pathway Relays Cytokine-Based Signals. The protein  growth factors that regulate the differentiation, proliferation, and activities of  numerous types of cells, most conspicuously white blood cells, are known as  cytokines. The signal that certain cytokines have been extracellularly bound  by their cognate receptors is transmitted within the cell, as James Darnell elu¬  cidated, by the JAK-STAT pathway. Cytokine receptors form complexes with  proteins of the Janus kinase (JAK) family of nonreceptor tyrosine kinases  (NRTKs), so named because each of its four —1150-residue members has two  tyrosine kinase domains (Janus is the two-faced Roman god of gates and door¬  ways), although only the C-terminal domain is functional. STATs (for signal  transducers and activators of transcription) comprise a family of seven —800-  residue proteins that are the only known transcription factors whose activities  are regulated by Tyr phosphorylation and that have SH2 domains.   The JAK-STAT pathway functions as is diagrammed in Fig. 28-33: Section 3 Regulation of Eukaryotic   1. Ligand binding induces the cytokine receptor to dimerize (or, in some  cases, to trimerize or even tetramerize).   2. The cytokine receptors two associated JAKs are thereby brought into  apposition, whereon they reciprocally phosphorylate each other and  then their associated receptors, a process resembling the autophos¬  phorylation of dimerized RTKs (Section 13-2A).   3. STATs bind to the phospho-Tyr group on their cognate activated re¬  ceptor via their SH2 domain and are then phosphorylated on a con¬  served Tyr residue by the associated JAK.    FIG. 28-33 The JAK-STAT pathway for the intracellular relaying of cytokine signals.   [After Carpenter, L.R., Yancopoulos, G.D., and Stahl, N., Adv. Prot. Chem. 52, 109 (1999).]    FIG. 28-34 X-Ray structure of the  Stat3p homodimer bound to a 17-bp DNA  containing its 9-bp target sequence. The   complex is viewed along the helix axis of the  DNA, which is represented by a purple ribbon.  The protein’s various domains are drawn in  different colors. The phosphorylatable Tyr  residues are located in the SH2 domains  (yellow), which mediate the protein’s dimerization.  Note how the STAT dimer binds the DNA in a  sort of scissors grip. [Courtesy of Christoph  Muller, European Molecular Biology Laboratory,  Grenoble, France. PDBid 1BG1.]   4. Following their dissociation from the receptor, the phosphorylated  STATs homo- or heterodimerize via the association of their phospho-  Tyr residue with the SH2 domain on the opposing subunit.   5. The STAT dimers are then translocated to the nucleus, where the now  functional transcription factors specifically bind to 9-bp DNA seg¬  ments with the consensus sequence TTCCGGGAA, thereby inducing  the transcription of their associated genes.   Like many other components of signal transduction pathways, the STAT sig¬  nal is inactivated through the action of phosphatases so that a STAT typically  remains active for only a few minutes. The X-ray structure of the STAT named  Stat3P, determined by Christoph Muller, is shown in Fig. 28-34.   Nuclear Receptors Are Activated by Hormones. The nuclear receptor su¬  perfamily, which occurs in animals ranging from worms to humans, is com¬  posed of >150 proteins that bind a variety of hormones including steroids  (glucocorticoids, mineralocorticoids, estrogens, and androgens; Sections 9-IE  and 13-1C), thyroid hormones such as thyroxine (Fig. 4-15) that stimulate  metabolism, and vitamin D (Section 9-IE), all of which are nonpolar mole¬  cules. The nuclear receptors, many of which activate distinct but overlapping  sets of genes, share a conserved modular organization that includes, from N-  to C-terminus, a poorly conserved activation domain, a highly conserved  DNA-binding domain, a connecting hinge region, and a ligand-binding do¬  main. The DNA-binding domains each contain eight Cys residues that, in  groups of four, tetrahedrally coordinate two Zn 2+ ions. Many members of the  nuclear receptor superfamily recognize specific DNA segments known as hor¬  mone response elements (HREs) that have the half-site consensus sequences  5'-AGAACA-3' for steroid receptors and 5'-AGGTCA-3' for other nuclear re¬  ceptors. These sequences are arranged in direct repeats inverted re¬  peats and everted repeats (<— n—>), where n represents a 0- to 8-bp   spacer (usually 1-5 bp) to whose length a specific receptor is targeted. Steroid  receptors bind to their hormone response elements as homodimers, whereas  other nuclear receptors do so as homodimers, as heterodimers, and in a few  cases as monomers.   Steroid hormones, being nonpolar, readily pass through cell membranes  into the cytosol or nucleus, where they bind to their receptors (although in  some cases, they may also interact with cell-surface receptors). In the absence  of their cognate steroid, these receptors are part of large multiprotein    FIG. 28-35 X-Ray structure of the dimeric glucocorticoid receptor DNA-binding  domain in complex with DNA. The complex is viewed with its approximate twofold  molecular axis horizontal. Its 18-bp DNA is drawn in paddle form with the C atoms of its  two 6-bp GRE half-sites magenta, the C atoms of the remaining nucleotides green, N blue,  0 red, and P orange and with successive P atoms connected by orange rods. The protein  is shown in ribbon form with its lower subunit colored in rainbow order from its N-terminus  (blue) to its C-terminus (red) and upper subunit pink. Each subunit contains a zinc finger  (Section 24-4C), whose two Zn 2+ ions (cyan spheres) are tetrahedrally liganded by Cys  side chains shown in stick form with C green and S yellow. Note how the receptor’s two  N-terminal helices are inserted into adjacent major grooves of the DNA. However, only the  lower subunit binds to the DNA in a sequence-specific manner; the upper subunit binds to  the palindromic DNA one base pair closer to the center of the DNA molecule than does  the upper subunit and hence does not make sequence-specific contacts with the DNA.  [Based on an X-ray structure by Paul Sigler, Yale University. PDBid 1GLU.] Q See  Interactive Exercise 56.    complexes. Steroid binding releases these receptors, whereon they dimerize  and, if they are cytoplasmic, enter the nucleus, where they bind to their tar¬  get HREs so as to induce, or in some cases repress, the transcription of the  associated genes.   The X-ray structure of the 86-residue DNA-binding domain of rat glu¬  cocorticoid receptor (GR; which regulates carbohydrate, protein, and lipid  metabolism) in complex with DNA containing two ideal 6-bp glucocorticoid  response element (GRE) half-sites arranged in inverted repeats was deter¬  mined by Paul Sigler and Keith Yamamoto (Fig. 28-35). The protein forms  a symmetric dimer involving protein—protein contacts even though it exhibits  no tendency to dimerize in the absence of DNA (NMR measurements indi¬  cate that the contact region is flexible in solution). Each protein subunit con¬  sists of two structurally distinct modules, each nucleated by a Zn 2+  coordination center, that closely associate to form a compact globular fold.  The C-terminal module provides the entire dimerization interface and also  makes several contacts with the phosphate groups of the DNA backbone. The  N-terminal module, which is also anchored to the phosphate backbone, makes  all of the GR’s sequence-specific interactions with the GRE via three side  chains that extend from the N-terminal a helix, its recognition helix, which  is inserted into the GREs major groove.   Afterword. As we have seen, eukaryotic transcriptional initiation is an as-  toundingly complex process that involves large segments of DNA as well as  the synergistic participation of numerous multisubunit complexes comprising  several hundred often loosely or sequentially interacting polypeptides (i.e.,  histones, the PIC, Mediator, transcription factors, architectural factors, coacti¬  vators, corepressors, chromatin-remodeling complexes, and histone-modifying  enzymes). Despite the extensive characterization of many of these factors, we  are far from having a more than rudimentary understanding of how the var¬  ious components interact in vivo to transcribe only those genes required by  their cell under its particular circumstances in the appropriate amounts and  with the proper timing. Nevertheless, an increase in the complexity of tran¬  scriptional regulation—rather than an increase in gene number per se—is  likely to be at least partially responsible for the vast differences in morphol¬  ogy and behavior among nematodes, fruit flies, and humans, which have com¬  parable numbers of genes (Table 28-1).   Although most regulation of gene expression in eukaryotes occurs at the level  of transcription, additional control mechanisms act after an RNA transcript  has been synthesized. In this section, we consider several of these mechanisms,  including mRNA degradation, RNA interference, and control of translation   Chapter 28 Regulation of Gene Expression initiation. We have already discussed alternative mRNA splicing (Section 26-3B)   as a posttranscriptional mechanism for modulating gene expression through  the selection of different exons.   mRNAs Are Degraded at Different Rates. The range of mRNA stability in  eukaryotic cells, measured in half-lives, varies from a few minutes to many  hours or days. The mRNA molecules themselves appear to contain elements  that dictate their decay rates. These elements include the poly(A) tail, the 5' cap,  and sequences that are located within the coding region.   A major route for mRNA degradation begins with the progressive removal  of its poly(A) tail, a process catalyzed by deadenylases that appear to be located  throughout the cytosol. When the residual poly(A) tail is less than ~10 nt  long and hence no longer capable of interacting with poly (A)-binding protein  (Section 26-3A), the mRNA becomes a substrate for a decapping enzyme,  which hydrolytically excises the mRNAs m 7 GDP cap. This is possible because,  as we have seen (Section 27-4A), the translational initiation factor eIF4G in¬  teracts with both poly(A)-binding protein and cap-binding protein, thereby  circularizing the mRNA so that events at its 3' end can be coupled to events  at its 5' end. The decapped and deadenylated mRNA is then degraded by exo¬  nucleases, mainly the 1706-residue 5' —> 3' exonuclease Xrnl and the 3' —>  5' exonuclease complex named the exosome. A decapping enzyme, 5' —»  y exonucleases, and accessory proteins form a complex called a P body (P for  processing), that functions either to degrade mRNA or to store it in an inac¬  tive form.   Proteins that bind to AU-rich elements (AREs) in the 3' untranslated re¬  gion of the mRNA also appear to increase or decrease the rate of mRNA degra¬  dation, although their exact action is not understood. RNA secondary  structure and RNA-binding proteins, which may be susceptible to modifica¬  tion by cellular signaling pathways, play a role in regulating mRNA stability.  mRNAs that cannot be translated due to the presence of a premature Stop  codon are specifically targeted for degradation (Box 28-3).   The eukaryotic core exosome consists of single copies of nine different  subunits. Its X-ray structure (Fig. 28-36), determined by Christopher Lima,  reveals that six of these subunits, Rpr4l (Rpr for rRNA processing; the exo¬  some was discovered as an activity that processed the 3' ends of rRNAs),    Errors during DNA replication, transcription, or mRNA splicing can  give rise to a Stop codon either through substitution of one nu¬  cleotide for another or through frameshifting. “Premature” Stop  codons that interrupt a coding sequence may account for as many  as one-third of human genetic diseases. Interestingly, eukaryotic  mRNAs with premature Stop codons rarely produce the corre¬  sponding truncated polypeptide because the mRNA is destroyed  through nonsense-mediated decay (NMD) soon after its synthesis.   How do cells distinguish a premature Stop codon from a normal  Stop codon at the end of the coding sequence? Experiments sug¬  gest that the signal for NMD depends on RNA splicing. Following  intron removal, an exon-junction protein complex (EJC) contain¬  ing splicing factors and other components remains associated with  each exon-exon junction, even after the mRNA has been exported  to the cytosol for translation. An EJC following a Stop codon  apparently marks the transcript for destruction.   A revised model for NMD proposes that quality control occurs  before the mRNA leaves the nucleus. Although the bulk of    translation occurs in the cytosol, an estimated 10 to 15% of total  translation in mammalian cells takes place inside the nucleus.  During nuclear translation, a ribosome paused at a Stop codon  may be converted to a “surveillance” complex that then scans the  rest of the mRNA for the presence of an EJC. If an EJC is de¬  tected, the mRNA is degraded by removal of its 5' cap and 3'  poly(A) tail and by the action of exonucleases.   Two other quality-control mechanisms also depend on transla¬  tion. mRNAs that lack Stop codons entirely are subject to non¬  stop decay. When the ribosome reaches the 3' end of the  defective mRNA, a protein known as Ski7p, which structurally  resembles the release factor eRF3, binds to the A site and re¬  cruits an exosome to degrade the mRNA. In yeast, a mechanism  known as no-go decay deals with mRNAs that include an obsta¬  cle such as a stable stem-loop that halts translation. In this case,  the mRNA is cleaved by an endonuclease associated with the  stalled ribosome.    FIG. 28-36 X-Ray structure of the human  core exosome. The protein complex is drawn in  ribbon form embedded in its semitransparent  molecular surface, with each of its nine different  subunits separately colored. The view is toward  the face of the six-membered ring of subunits  opposite that to which the three other subunits  bind. [Based on an X-ray structure by  Christopher Lima, Sloan-Kettering Institute, New  York, New York. PDBid 2NN6.]    Rrp42, Mtr3, Rrp43, Rrp46, and Rrp45, form a six-membered ring with  the remaining three subunits, Rrp4, Csl4, and Rrp40, bound to the same  face of the ring. The subunits are arranged such that the core exosome con¬  tains an ~9-A-wide central channel that allows the entrance of only single-  stranded RNAs.   The archaeal exosome appears to be a simpler version of the eukaryotic  core exosome. Its six-membered ring consists of only two types of subunit,  Rrp4l and Rrp42, that alternate around the ring, with three copies of Rrp4  bound to the same face of the ring. Only Rrp4l contains an active site al¬  though Rrp42 is required for activity. Not surprisingly, eukaryotic Rrp4, Mtr3,  and Rrp46 are homologs of archaeal Rrp4l, eukaryotic Rrp42, Rrp43, and  Rrp45 are homologs of archaeal Rrp42, and eukaryotic Rrp4, Csl4, and Rrp40  are homologs of archaeal Rrp4. Nevertheless, despite the fact that each of its  core subunits is essential for viability, eukaryotic core exosomes, from yeast to  humans, are catalytically inactive. However, the core exosome associates with  two 3'-exonucleases, Rrp6 and Rrp44, whose catalytically inactive mutants are  individually viable in yeast but lethal in combination. Moreover, the core ex¬  osome interacts with numerous mostly multisubunit cofactors that carry out  a variety of RNA processing activities in both the nucleus and the cytosol.  Thus, the eukaryotic core exosome appears to be a structural platform on  which many RNA processing enzymes can be mounted.   RNA Interference Is a Type of Posttranscriptional Gene Silencing. Since  the 1990s, it has become increasingly clear that noncoding RNAs can have  important roles in controlling gene expression. One of the first indications of  this phenomenon occurred in Richard Jorgensens attempt to genetically engi¬  neer more vividly purple petunias by introducing extra copies of the gene that  directs the synthesis of the purple pigment. Unexpectedly, the resulting trans¬  genic plants had variegated or entirely white flowers. Apparently, the purple¬  making genes switched each other off. This result was at first attributed to the  well-known phenomenon in which antisense RNA (RNA that is complemen¬  tary to a portion of an mRNA) prevents the translation of the corresponding  mRNA because the ribosome cannot translate double-stranded RNA.  However, injecting sense RNA (RNA with the same sequence as the mRNA)  into experimental organisms such as the nematode worm C. elegans also  blocked protein production. Since the added RNA somehow interferes with  gene expression, this phenomenon is known as RNA interference (RNAi).  RNAi is now known to occur in all eukaryotes except bakers yeast.   Chapter 28 Regulation of Gene Expression Further experiments by Andrew Fire and Craig Mello revealed that double-   stranded RNA is even more effective than either of its component strands  alone at interfering with gene expression. In fact, RNAi can be induced by  just a few molecules of double-stranded RNA, indicating that RNAi is a cat¬  alytic rather than a stoichiometric phenomenon.   RNA interference is not merely an artifact of genetic engineering. In many  cells, naturally occurring small RNA molecules, called short interfering RNAs  (siRNAs) or micro RNAs (miRNAs), depending on their origin, down-  regulate gene expression by binding to complementary mRNA molecules.  Thousands of miRNAs, ranging in length from 18 to 25 nucleotides, have  been identified in mammals, although their mRNA targets are only just be¬  ginning to be characterized.   Several Nucleases Participate in RNAi. Work with exogenous double-  stranded RNA in C. elegans and Drosophila has led to the elucidation of the  following pathway for RNAi (Fig. 28-37):   1. The trigger double-stranded RNA, as Phillip Zamore discovered, is  chopped up into —21- to 23-nt fragments (that is, siRNAs), each of    L RNA-induced silencing complex  2 i(RISC) binds to the siRNA and  f separates its strands.    I I I I I I I I I I I I I + I I I I I I I I I I I I I I I I I    FIG. 28-37 A mechanism of RNA interference. ATP is required unwinding of double-stranded RNA. Depending on the species, the   for Dicer-catalyzed cleavage of RNA and for RISC-associated helicase mRNA may not be completely degraded.    whose strands has a 5 r phosphate and a 2-nt overhang at its 3' end.  The cleavage reaction is catalyzed by an ATP-dependent RNase named  Dicer, a homodimer of ~ 1900-residue subunits in animals that is a  member of the RNase III family of double-strand-specific RNA en¬  donucleases.   2. An siRNA is transferred to a 250- to 500-kD enzyme complex known  as the RNA-induced silencing complex (RISC). RISC has at least  four protein components, one of which is an ATP-dependent RNA he-  licase that separates the two strands of the siRNA. The strand whose  5' end has the lower free energy of binding, the guide RNA, is bound  by the RISC whereas its complementary strand, the passenger RNA,  is cleaved and discarded. In some species, but apparently not in hu¬  mans, the original siRNA signal is amplified by the action of an RNA-  dependent RNA polymerase.   3. The guide RNA recruits the RISC complex to an mRNA with the  complementary sequence.   4. An RNase III component of RISC known as Argonaute (AGO; also  called Slicer) cleaves the mRNA opposite the bound guide RNA. The  cleaved mRNA is then further degraded by cellular nucleases, thereby  preventing its translation.   Although the molecular machinery involved in RNAi is highly conserved  from yeast to humans, variations abound, even within a single organism. For  example, different Dicer enzymes can potentially generate different lengths of  siRNAs, which may differ in their gene-silencing activities. The exact length  of the siRNA produced by Dicer, or the degree to which its sequence exactly  complements the mRNA sequence, may determine whether a corresponding  mRNA is completely degraded or just rendered nontranslatable.   The X-ray structure of Dicer from the parasitic protozoan Giardia intesti-  nalis , determined by Jennifer Doudna, reveals a hatchet-shaped enzyme with  two RNase active sites, each of which includes four conserved acidic residues  that bind two Mg 2+ ions. The active sites are 17.5 A apart, the width of  dsRNAs major grove, and thus appear positioned to cleave the two strands of  a bound dsRNA (Fig. 28-38). The so-called PAZ domain (bottom of Fig.  28-38) specifically binds dsRNA ends that have a 3' two-nucleotide overhang.  The distance between this binding site and the closest RNase active site is 65 A,  the length of a 25-bp dsRNA.   The ease with which RNAi can be induced by exogenous RNAs suggests  that the pathway may have arisen as a defense against viruses. In many eu¬  karyotes, the major source of double-stranded RNA is RNA viruses, many of  which encode their own RNA-dependent RNA polymerase to convert their  single-stranded genome into double-stranded RNA. In fact, many plant viruses  contain genes that suppress various steps of RNAi and that are therefore es¬  sential for pathogenesis.   Micro RNAs Regulate Gene Expression. A wide variety of eukaryotes, includ¬  ing plants, nematodes, flies, fish, and mammals, use RNAi to control gene expres¬  sion. Certain mRNAs expressed by these organisms contain ~70-nt, imperfectly    FIG. 28-38 X-Ray structure of Dicer from G. intestinalis. The protein is represented  by its molecular surface colored according to its surface charge with red negative, blue  positive, and white neutral. Bound Mg 2+ ions, which are represented by green spheres,  mark the active site of each of the protein’s two RNase domains. A dsRNA has been  modeled into the structure with its 3' overhang entering the PAZ domain’s binding pocket  (asterisk). The white arrows point to the dsRNA’s scissile phosphate groups. [Courtesy of  Jennifer Doudna, University of California at Berkeley. PDBid 2FFL]   FIG. 28-39 The generation of miRNAs  from pri-miRNAs and their RISC-mediated  binding to target mRNAs. See the text for  details.   base-paired, stem-loop structures that are excised by a 1374-residue RNase III  named Drosha (Fig. 28-39). The stem-loops are exported from the nucleus  to the cytosol where they are cleaved by Dicer to liberate — 22-bp dsRNAs  known as micro RNAs (miRNAs; so called to differentiate these endogenous  RNAs from the exogenous siRNAs). The transcripts from which miRNAs are  derived are known as pri-miRNAs (pri for granary), whereas the stem-loops  are called pre-miRNAs (pre for precursor). Pre-miRNAs can be located within  both the introns and, less commonly, the exons of a pri-miRNA. The miRNAs  bind to RISC in which they function to identify the tens to hundreds of mRNAs  containing segments that are partially complementary to the miRNA.   The RISC-bound miRNA binds to its target site, which is usually in the  y untranslated region (3'UTR) of an mRNA. A lack of perfect complemen¬  tarity prevents Argonaute from cleaving the mRNA (Argonaute catalyzes slic¬  ing only if there is perfect complementarity to the miRNAs so-called seed  sequence, which consists of nucleotides 2-8 from its 5' end), and in fact, some  species of Argonaute lack the catalytic residues to do so. Instead, miRNA-  mediated silencing is thought to occur through the removal of its target  mRNAs poly(A) tail or its m 7 G cap, which leads to the mRNAs degradation,  and/or the RISC-mediated repression of the target mRNAs translation by  interfering with ribosomal initiation (Section 27-4A) and sequestering or de¬  grading the mRNA in P bodies.   In 1993, Victor Ambros discovered the first known miRNA, which is en¬  coded by the lin-4 gene of C. elegans (Fig. 28-40 #). The lin-4 gene was known  to control the timing of larval development, although at the time it was  thought that it encoded a protein that repressed the expression of the lin-14  gene. In fact, the lin-4 miRNA is complementary to seven sites on the 3'UTR  of the lin-14 gene, which had previously been shown to mediate the repres¬  sion of lin-14 by the lin-4 gene product. A puzzling observation at the time  was that this regulation greatly reduces the amount of LIN-14 protein pro¬  duced without altering the level of lin-14 mRNA. These findings were even¬  tually followed by the discovery that the C. elegans let-7 gene encodes what is  now known to be an miRNA (Fig. 28-40 £) that controls the transition from  larval to adult stages of development. Subsequently, let-7 homologs were iden¬  tified in the Drosophila and human genomes and let-7 RNA was detected in  these organisms as well as in numerous other animals.   Both the lin-4 and let-7 miRNAs were discovered by genetic analyses.  However most of the nearly 10,000 miRNAs in plants and animals that are  now known, including those in Fig. 28-40c, were identified through bioinfor-  matic approaches (Section 6-2E). Nearly all miRNAs are conserved among  closely related animals (e.g., mice and humans) and many are more broadly  conserved throughout animal lineages (e.g., over one-third of the 174 C. ele¬  gans miRNAs have homologs in humans). The significance of miRNAs is indi¬  cated by the fact that humans express over 720 miRNAs that participate in  regulating — 30% of their protein-coding genes.    RNAi Has Numerous Applications. The exquisite specificity of RNAi has  made it the method of choice for “knocking out” specific genes in plants and  invertebrates. For example, in C. elegans , RNAi has been used to systemati¬  cally inactivate over 16,000 of its —19,000 protein-coding genes in an attempt  to assign a function to each gene. C. elegans is particularly amenable to the  RNAi approach, since these worms eat E. coli cells, and it is relatively easy to  genetically engineer the bacterial cells to express double-stranded RNA that  becomes part of the worms’ diet. One limitation of the RNAi method is that  only gene inactivation—rather than gene activation—can be examined.   Manipulating the RNAi pathway in mammals is more problematic, mainly  due to the difficulty of delivering double-stranded RNA to cells and eliciting  more than transient gene silencing. To complicate matters, mammalian cells have  additional pathways for dealing with foreign RNA, which result in nonspecific    degradation of RNA and cessation of all translation (these responses probably  also help prevent infection by RNA viruses). Nevertheless, experiments have  demonstrated that it is possible to use RNAi to block the livers inflammatory  response to a hepatitis virus, at least in mice, and to prevent HIV replication in  cultured human cells. One challenge for the future is to devise protocols for  more specific and longer-lasting gene silencing that would make it possible to  prevent viral infections or to block the effects of disease-causing mutant genes.    mRNA Translation May Be Controlled. In some cells, altering the rates of  mRNA production or degradation does not provide the necessary level of con¬  trol. For example, the early embryonic development of sea urchins, insects,  and frogs depends on the rapid translation of mRNA that has been stockpiled  in the oocyte. The mRNA is stored in inactive form in association with pro¬  teins but on fertilization becomes available for translation. This permits em-  bryogenesis to commence immediately, without waiting for mRNAs from  paternally supplied genes to be synthesized.   Globin synthesis in reticulocytes (immature red blood cells) also proceeds  rapidly, but only if heme is available. The inhibition of globin synthesis oc¬  curs at the level of translation initiation. In the absence of heme, reticulocytes  accumulate a protein, heme-regulated inhibitor (HRI). HRI is a kinase  that phosphorylates a specific Ser residue, Ser 51, on the a subunit of eIF2  (the initiation factor that delivers GTP and Met—tRNAi Met to the ribosome;  Section 27-4A).   Phosphorylated eIF2 participates in translation initiation in much the  same way as unphosphorylated eIF2, but it is not regenerated normally. At  the completion of the initiation process, unmodified eIF2 exchanges its bound  GDP for GTP in a reaction mediated by another initiation factor, eIF2B:    Phosphorylated eIF2 forms a much tighter complex with eIF2B than does un¬  phosphorylated eIF2. This sequesters eIF2B (Fig. 28-41), which is present in  lesser amounts than is eIF2, thereby preventing regeneration of the eIF2 • GTP  required for translation.   U G-U  A-U  G-C  U-A  G-C  U-A  G-C  A C  A-U  < C-G  | U-G  * C-G  C    C A U  U A  A A  U-A  A-U  U-G  C-G  A-ir  -A   B —A A  U-A  C A  C-G  U-A  U-A  C-G  -U  U-A  A-U  C-G  G-U  U-A  C-G  G A  A A  G-C  C-G    FIG. 28-40 The predicted stem-loops  of some pre-miRNAs. The miRNAs contained  in these pre-mRNAs, all of which are from C.  elegans, are red. (a) lin-4, (b) let-7, and (c) miR-1  and miR-1* (in blue), which are largely  complementary to each other.   Chapter 28 Regulation of Gene Expression J n the presence of heme, the heme-binding sites in HRI are occupied,   which inactivates the kinase. The eIF2 molecules that are already phosphory-  lated are reactivated through the action of eIF2 phosphatase, which is unaf¬  fected by heme. The reticulocyte thereby coordinates its synthesis of globin  and heme.   As described in Section 7-3B, an individual can generate an extraordinary  number of different antibody molecules from a limited number of im¬  munoglobulin gene segments. This feat is accomplished through an unusual  mutation process and through somatic recombination in differentiating cells  of the immune system. Homologous recombination (Section 25-6A), a fun¬  damental feature of reproduction in multicellular organisms, occurs only in  germline cells. The mechanism whereby a developing B lymphocyte “selects”  light and heavy chain gene segments for expression is described below. Similar  events occur in T lymphocytes to generate a large number of unique T cell  receptors.   Immunoglobulin Chain Genes Are Assembled from Multiple Gene  Segments. One of the two types of immunoglobulin light chains, the K  chain, is encoded by four exons (Fig. 28-42):   1. A leader or Z K segment, which encodes a 17- to 20-residue hydropho¬  bic signal peptide. This polypeptide directs newly synthesized k chains  to the endoplasmic reticulum and is then excised (Section 9-4D).   2. A V K segment, which encodes the first 95 residues of the k chains  108-residue variable region.   4. The C K segment, which encodes the k chains constant region.   In embryonic tissues (which do not make antibodies), the exons occur in clus¬  ters. The human k chain gene family contains ~40 functional L K and V K seg¬  ments, separated by introns, with the L K - V K units separated from each other by  ~7-kb spacers. This sequence of exon pairs is followed, well downstream, by 5  J K segments at intervals of ~300 bp, a 2.4-kb spacer, and a single C K segment.   The assembly of a k chain mRNA is a complex process involving both so¬  matic recombination and selective mRNA splicing (Section 26-3B) over several  cell generations. The first step of this process, which occurs in B cell progenitor    differentiation, a single Z_ K - V K unit is joined to a J K unit through  somatic recombination. In the progeny B cells, the rearranged gene is  transcribed (2) and spliced (3) to join the previously selected Z_ K , V K ,  and J K exons with the C K exon. This diagram is not drawn to scale.    cells, is an intrachromosomal recombination that joins an L K -V K unit to a J K  segment and deletes the intervening DNA sequences (Fig. 28-42). Then, in  later cell generations, the entire modified gene is transcribed and the resulting  transcript is selectively spliced so as to join the L K -V K -J K unit to the C K seg¬  ment. The L k and V K segments are also spliced together in this step, yielding  an mRNA that encodes one of each of the four elements of a k chain gene.   The joining of 1 of 40 functional V K segments to 1 of 5 J K segments can  generate only 40 X 5 — 200 different k chains, far less than the number ob¬  served. However, studies of many joining events involving the same V K and  J K segments revealed that the VI] recombination site is not precisely defined; the  two gene segments can join at dijferent points (Fig. 28-43). Consequently, the  amino acids specified by the codons in the vicinity of the VI] joint, which cor¬  responds to the third hypervariable loop (Fig. 7-40), depend on which nu¬  cleotides are supplied by the germline V K segment and which are supplied by  the germline ] K segment. Assuming that this junctional flexibility increases the  possible k chain diversity 10-fold, the expected number of possible different  k chains is increased to around 40 X 5 X 10 = 2000.   The other type of immunoglobulin light chain, the \ chain, is similarly  encoded by a gene family containing LV and C segments whose re¬  combination likewise yields a large number of possible polypeptides.   Heavy chain genes are assembled in much the same way as are light chain  genes but with the additional inclusion of an ~ 13-bp diversity or D segment  between their V H and / H segments. The human heavy chain gene family  consists of clusters of ~65 different functional L H -V H units, ~27 D segments,  6 / H segments, and 9 C H segments (Fig. 28-44). Germline Vh, A and / H  segments are joined in a particular order (D is joined to / H before Vh is joined  to A/h)> and the joining sites are subject to the same junctional flexibility as  are light chain VI] sites. Thus the number of possible different heavy chains  is around 65 X 27 X 6 X 9 X 10 2 = 9.5 X 10 6 .   The point at which the V K and J K sequences  somatically recombine varies by several  nucleotides, thereby giving rise to different  nucleotide sequences in the active k gene. In  this example of junctional flexibility, the second  amino acid can be Ser, Gly, or Asp.    Somatic Recombination Occurs at Specific Sequences. Highly conserved  sequences flanking the V, D, and J gene segments act as recombination signal   FIG. 28-44 The organization and rearrangement of the  heavy chain gene family in humans. This gene family consists of  —65 sequential pairs of Z_ H and l/ H gene segments followed by —27  D segments, 6 T H segments, and 9 C H segments (one for each class  or subclass of heavy chains; Table 7-2). During lymphocyte    differentiation, an Z_ H -l/ H unit is joined to a D segment and a 7 H  segment. In the process, the D segment becomes flanked by short  stretches of random sequence called N regions. In the B cell and its  progeny, transcription and splicing join the Z_ H -I/ H -A/-D-AK/ H unit to  one of the 9 C H gene segments. This diagram is not drawn to scale.   elements consist of 7-bp and 9-bp conserved sequences separated by  23-bp or 12-bp spacers (white and black arrowheads), (b) Locations of  12-RSS (black) and 23-RSS (white) elements relative to light chain and  heavy chain gene segments. Recombination, represented by the  double-headed arrows, always links a gene segment with a 12-RSS to a  gene segment with a 23-RSS, thereby ensuring that the V K /J K and  I / h /D/J h gene sequences are properly assembled.    sequences (RSS; Fig. 28-45 tf). Each RSS consists of a palindromic heptamer  and an AT-rich nonamer separated by either 12 bp (corresponding to ~ 1 turn  of the DNA helix) or 23 bp (corresponding to ~2 helical turns). The recom¬  bination machinery joins a segment with a so-called 12-RSS to a segment with  a 23-RSS. Because each V K gene segment has a 12-RSS and each J K segment  has a 23-RSS, somatic recombination cannot join two V K segments or two J K  segments. Similarly, the distribution of 12-RSS and 23-RSS elements in heavy  chain gene segments is such that V H must join to D , and D to / H , which pre¬  cludes VulJu joining (Fig. 28-45 b).   Recombination requires enzymes that are synthesized only in developing  lymphocytes, as well as ubiquitous DNA repair proteins. All V(D)J joining  reactions are catalyzed by an evolutionarily conserved V(D)J recombinase sys¬  tem. Indeed, David Baltimore discovered two proteins, RAG1 and RAG2  (RAG for recombination activating genes), that work in concert to recognize  the RSS elements of two different gene segments, align them, and catalyze  DNA cleavage. This process is assisted by HMG proteins HMG1 and HMG2,  which appear to bend the DNA. Under laboratory conditions, RAG1 and  RAG2 can catalyze transposition, that is, the relocation of a DNA segment to  another DNA molecule (Section 25-6C), consistent with the proposal that the  RAG proteins originated as a transposon that appeared in vertebrates around  450 million years ago.   During recombination, the two RSS elements are neatly excised, but their  adjacent coding sequences may be trimmed by nucleases and unevenly joined,  thereby contributing to junctional flexibility (Fig. 28-43). The trimming and  ligating reactions are not unique to lymphocytes but apparently make use of  a variety of DNA repair enzymes that function in all cells (Section 25-5).   In addition, a few nucleotides may be added to the recombination joints,  either as part of the DNA-repair process or, in the case of heavy chain gene  assembly, by the action of the lymphocyte-specific enzyme terminal deoxynu-  cleotidyl transferase. This DNA polymerase is unusual in that it does not re¬  quire a template. It may add up to 15 nucleotides at the V^/ D H and Z) H // H  joints, which, of course, greatly increases the potential for sequence diversity  in the encoded variable region of the immunoglobulin chain.   Hypermutation Is a Further Source of Antibody Diversity. Despite the enor¬  mous antibody diversity generated by somatic recombination, immunoglobu¬  lins are subject to even more variation that arises as the B cells undergo cell  division. The amino acid sequences of the variable regions of both heavy and    light chains are more diverse than is expected from their original germline nu¬  cleotide sequences. Indeed, these regions mutate at rates of up to 10 3 base  changes per nucleotide per cell generation, rates that are at least a millionfold  higher than the rates of spontaneous mutation in other genes. This somatic  hypermutation of immunoglobulin gene segments is initiated by the expres¬  sion, in proliferating B cells, of a cytidine deaminase named activation-  induced deaminase (AID). The resulting uracil may then participate in  normal DNA replication, yielding a C • G to T • A transition (Section 25-4A).  More frequently, however, the uracil is excised by uracil-DNA glycosylase  (Section 25-5B) and replaced by a base excision repair (BER; Section 25-5B), a  process that employs error-prone translesion DNA polymerases (Section 25-5E).  The mutations occur throughout the VIJ and VIDIJ regions but are concen¬  trated at the sequences corresponding to the three hypervariable loops of each  chain. DNA sequences in other genes do not seem to be affected.   Somatic hypermutation acts over many cell generations after antigen stim¬  ulation. Because B cells producing antibodies with high affinity for their anti¬  gen tend to proliferate faster than B cells producing poorly binding antibodies,  the pool of antibody molecules becomes exquisitely tailored to a particular  antigen over time. The diversity arising from somatic recombination, with its  junctional flexibility, and from hypermutation thereby permits an individuals  immune system to cope, in a kind of Darwinian struggle, with the rapid mu¬  tation rates of pathogenic microorganisms.   • Cyclins and cyclin-dependent kinases regulate progress through the cell cycle.   • Tumor suppressors such as p53 alter gene expression to arrest the cell cycle.   • Apoptosis leads to cell death in response to internal or external signals.    In addition to selecting which genes to express from among the thousands that  are present in its genome, a cell must decide when to express them. Thus a  cell can undertake a particular developmental program or respond effectively  to changing conditions. The events of the eukaryotic cell cycle, encompassing  normal and abnormal cell reproduction, as well as cell death, illustrate how  particular gene products regulate a cells activities at different times.   • Describe how histone modifications can  affect the structure of nucleosomes and  the function of transcription factors.   • Summarize the role of DNA methylation in  epigenetic inheritance and imprinting.   • Explain why eukaryotic gene expression  requires proteins in addition to the six  general transcription factors.   • Describe how antibody diversity is  generated. List the responsible enzymatic  activities that are unique to lymphocytes.   The cell cycle, the general sequence of events that occur during the lifetime  of a eukaryotic cell, is divided into four distinct phases (Fig. 28-46):   1. Mitosis and cell division occur during the relatively brief M  phase (for mitosis).   2. This is followed by the Gj phase (for gap), which covers  the longest part of the cell cycle.   3. Gi gives way to the S phase (for synthesis), which, in  contrast to events in prokaryotes, is the only period in the  cell cycle when DNA is synthesized.   4. During the relatively short G 2 phase, the now tetraploid  cell prepares for mitosis. It then enters M phase once again  and thereby commences a new round of the cell cycle.    FIG. 28-46 The eukaryotic cell cycle. Cells may enter a quiescent phase  (G 0 ) rather than continuing about the cycle.   Chapter 28 Regulation of Gene Expression The cell cycle for cells in culture typically occupies a 16- to 24-h period. In   contrast, cell cycle times for the different types of cells in a multicellular or¬  ganism may vary from as little as 8 h to >100 days. Most of this variation  occurs in the G x phase. Moreover, many terminally differentiated cells, such  as neurons or muscle cells, never divide; they assume a quiescent state known  as the G 0 phase.   Progression through the cell cycle is triggered by external as well as inter¬  nal signals. In addition, the cell cycle has a series of checkpoints that moni¬  tor its progress and the health of the cell and arrest the cell cycle if certain  conditions have not been satisfied. For example, G 2 has a checkpoint that pre¬  vents the initiation of M until all of the cells DNA has been replicated,  thereby ensuring that both daughter cells will receive a full complement of  DNA. Similarly, a checkpoint in M prevents mitosis until all chromosomes  have properly attached to the mitotic spindle (if this were not the case, even  for one chromosome, one daughter cell would lack this chromosome and the  other would have two, both deleterious if not lethal conditions). Checkpoints  in Gi and S also arrest the cell cycle in response to damaged DNA so as the  give the cell time to repair the damage (Section 25-5). In the cells of multi¬  cellular organisms, if after a time the checkpoint conditions have not been  met, the cell may be directed to commit suicide, a process named apoptosis  (see below), thereby preventing the proliferation of an irreparably damaged  and hence dangerous (e.g., cancerous) cell.   The Activities of Cyclin-Dependent Protein Kinases Change throughout  the Cell Cycle. The progression of a cell through the cell cycle is regulated  by proteins known as cyclins and cyclin-dependent protein kinases (Cdks).   Cyclins are so named because they are synthesized during one phase of the  cell cycle and are completely degraded during a succeeding phase (protein  degradation is discussed in Section 21-1). A particular cyclin specifically binds  to and thereby activates its corresponding Cdk(s), which are Ser/Thr protein  kinases, to phosphorylate their target nuclear proteins. These proteins, which  include histone HI, several oncogene proteins (see below), and proteins in¬  volved in nuclear disassembly and cytoskeletal rearrangement, are thereby ac¬  tivated to carry out the processes making up that phase of the cell cycle.   The human Cdk named Cdk2 is activated by the binding of cyclin A  and by phosphorylation of its Thr 160 as catalyzed by Cdk-activating kinase  (CAK; this kinase is itself a complex of cyclin H and Cdk7). Phosphorylation  of Cdk2 residues Thr 14 and Tyr 15 negatively regulates Cdk2 activity. The  X-ray structure of unphosphorylated Cdk2 in complex with ATP but in the  absence of cyclin A (Fig. 28-47#) closely resembles that of the catalytic sub¬  unit of protein kinase A (PKA; Fig. 13-21). However, Cdk2 is inactive as a  kinase in part because access of protein substrates to the y-phosphate of the  bound ATP is blocked by a 19-residue protein loop dubbed the T loop (which  contains Thr 160). Phosphorylation of Thr 160 and binding of cyclin A alter  the conformation of Cdk2 (Fig. 28-47£). In particular, the N-terminal a he¬  lix of Cdk2, which contains the PSTAIRE sequence motif characteristic of the  Cdk family, rotates about its axis by 90° and moves several angstroms in or¬  der to position several residues in a catalytically active arrangement. In addi¬  tion, Cdk2’s T loop undergoes a dramatic reorganization, involving position  shifts of up to 21 A, that allows protein substrates access to the active site.  The phosphate group on Thr 160 fits snugly into a positively charged pocket  composed of three Arg residues that forms, in part, on cyclin A binding.   In addition to their control by phosphorylation/dephosphorylation and by  the binding of the appropriate cyclin, Cdk activities are regulated by cyclin-  dependent kinase inhibitors (CKIs), which arrest the cell cycle in response to  such antiproliferative signals as contact with other cells, DNA damage, terminal  differentiation, and senescence (in which cell cycle arrest is permanent). The im¬  portance of CKIs is indicated by their frequent alterations in cancer, which is   FIG. 28-47 X-Ray structure of human cyclin-dependent  kinase 2 (Cdk2). (a) Cdk2 in complex with ATP. The protein is shown  in the “standard” protein kinase orientation with its N-terminal lobe  pink, its C-terminal lobe cyan, its PSTAIRE helix (residues 45-56)  magenta, and its T loop (residues 152-170) orange. The ATP is  shown in space-filling form and the phosphorylatable side chains of  Thr 14, Tyr 15, and Thr 160 are shown in stick form, all colored  according to atom type (C green, N blue, 0 red, and P yellow). [Based  on an X-ray structure by Sung-Hou Kim, University of California at  Berkeley. PDBid 1HCK.] (b) The complex of Thr 160-phosphorylated  Cdk2 with cyclin A and ATP. The Cdk2 and ATP are represented as in  Part a and viewed similarly. The cyclin A is colored light and dark   green. The phosphoryl group at Thr 160 of Cdk2 is drawn in  space-filling form. Note how the binding of cyclin A together with the  phosphorylation of Thr 160 has caused a major structural reorganization  of the T loop and the Cdk2 N-terminal lobe, including its PSTAIRE  helix. Also note the different conformations of the ATP triphosphate  group in the two structures. [Based on an X-ray structure by Nikola  Pavletich, Memorial Sloan-Kettering Cancer Center, New York, New  York. PDBid 1JST.] © See Interactive Exercise 57.    Compare the structure of Cdk2 with that of protein kinase A  (PKA; Fig. 13-21).    manifest as uncontrolled cell division. The pathways by which sensor proteins  monitor cellular conditions and activate or inactivate Cdks are not yet fully  understood. However, considerable information has been provided by studies  of proteins, including p53 and pRb (see below), whose mutations are associ¬  ated with loss of cell cycle control.   Individuals with the rare inherited condition known as Li-Fraumeni syn¬  drome are highly susceptible to a variety of malignant tumors, particularly  breast cancer, which they often develop before their thirtieth birthdays. These  individuals have germline mutations in their p53 gene, which suggests that its  normal protein product, p53 (a polypeptide with a nominal mass of 53 kD),  is a tumor suppressor. In other words, p53 functions to restrain the uninhib¬  ited cell proliferation that is characteristic of cancer. Indeed, the p53 gene is the  most commonly altered gene in human cancers; ~50% of human cancers con¬  tain a mutation in p53, and many other oncogenic (cancer-causing) mutations  occur in genes that encode proteins that directly or indirectly interact with  p53. Evidently, p53 functions as a “molecular policeman” in monitoring  genome integrity.   Chapter 28 Regulation of Gene Expression p 53 Arrests the Cell Cycle in G 2 . Despite the central role of p53 in prevent¬   ing tumor formation, the way it does so has only gradually come to light. p53  is specifically bound by Mdm2 protein, a ubiquitin-protein ligase (E3) that  specifically ubiquitinates p53, thereby marking it for proteolytic degradation  by the proteasome (Section 21-1). Consequently, the amplification (generation  of several copies) of the mdm2 gene, which occurs in >35% of human sarco¬  mas (none of which have a mutated p53 gene; sarcomas are malignancies of  connective tissues such as muscle, tendon, and bone), results in an increased  rate of degradation of p53, thereby predisposing cells to malignant transforma¬  tion. Thus, an additional way that oncogenes (cancer-causing genes; Box 13-3) can  cause cancer is by inactivating normal tumor suppressors.   p53 is an efficient transcriptional activator. Indeed, all point-mutated forms  of p53 that are implicated in cancer have lost their sequence-specific DNA-  binding properties. How, then, does p53 function as a tumor suppressor? A  clue to this riddle came from the observation that the treatment of cells with  DNA-damaging ionizing radiation results in the accumulation of normal p53.  This led to the discovery that the activated protein kinases ATM and Chk2  both phosphorylate p53 [ATM for ataxia telangiectasia mutated (ataxia  telangiectasia is a rare genetic disease characterized by a progressive loss of  motor control, growth retardation, premature aging, and a greatly increased  risk of cancer); Chk for checkpoint hinase]. ATM and Chk2 had previously  been shown to be activated by and participate in a phosphorylation cascade  that induces cell cycle arrest at the G 2 checkpoint on the detection of dam¬  aged or unreplicated DNA. The phosphorylation of p53 prevents its binding  by Mdm2 and hence increases the otherwise low level of p53 in the nucleus.  Although p53 does not initiate cell cycle arrest in G 2 , its presence is required to  prolong this process. It does so by activating the transcription of the gene encoding  the cyclin-dependent kinase inhibitor (CKI) named p21 Cipl , which binds to sev¬  eral Cdk-cyclin complexes so as to inhibit both the GfS and G 2 /M transitions.   Cells that are irreparably damaged synthesize excessive levels of p53, which  in turn induces these cells to commit suicide by activating the expression of  several of the proteins that participate in apoptosis (see below). In the absence  of p53 activation, cells control the level of p53 through a feedback loop in  which p53 stimulates the transcription of the mdm2 gene.   In addition to the foregoing, p53 represses the expression of numerous  genes. One mechanism through which it does so is by directly or indirectly  stimulating the transcription of certain microRNAs (miRNAs), which in turn  repress the expression of a variety of pro-proliferative proteins (Section 28-3C).  The most conspicuous of these miRNAs is miR-34a, which is implicated in  inducing senescence and facilitating apoptosis (cell death; see below).   A second repressive mechanism, which has only recently come to light,  involves a large intergenic noncoding RNA (lincRNA) known as lincRNA-  p21 (so named because its gene neighbors that of p21, although the signifi¬  cance of this, if any, is unknown). LincRNAs are large (>5 kb), evolutionarily  conserved RNAs that, like mRNAs, are transcribed by RNAP II, 5'-capped,  and polyadenylated, but do not encode proteins. Xist and Tsix RNAs (Box 28-2)  are examples of lincRNAs that we have encountered. Over 3000 lincRNAs  occur in the mammalian genome, many of which appear to participate in tran¬  scriptional control. The transcription of lincRNA-p21, as John Rinn discov¬  ered, is induced by p53. LincRNA-p21 forms a complex with heterogeneous  nuclear ribonucleoprotein K (hnRNP-K; hnRNPs are proteins that bind  hnRNAs) that represses the transcription of hundreds of genes, many of which  are pro-proliferative, although how this complex does so is poorly understood.   The X-Ray Structure of p53 Explains Its Oncogenic Mutations. p53 is a   tetramer of identical 393-residue subunits, which each contain a sequence-  specific DNA-binding core. The X-ray structure of this domain (residues  102-313) in complex with a 21-bp target DNA sequence, determined by    FIG. 28-48 X-Ray structure of the  DNA-binding domain of human p53 in  complex with its target DNA. The DNA is   drawn in stick form colored according to atom  type (C green, N blue, 0 red, and P orange) with  its successive P atoms joined by orange rods.  The protein is shown in ribbon form colored in  rainbow order from its N-terminus (blue) to its  C-terminus (red). A tetrahedrally liganded Zn 2+  ion is represented by a magenta sphere, and the  side chains of the six most frequently mutated  residues in human tumors are shown in stick  form with C yellow and identified with their  one-letter codes. [Based on an X-ray structure  by Nikola Pavletich, Memorial Sloan-Kettering  Cancer Center, New York, New York. PDBid  1TSRJ O See Interactive Exercise 58.    Nikola Pavletich, is shown in Fig. 28-48. The p53 DNA-binding motif does  not resemble any other that has previously been characterized (Sections 24-4B  and 24-4C). It makes sequence-specific contacts with the bases of the DNA  in its major groove (lower right of Fig. 28-48). In addition, the side chain of  Arg 248 extends into the DNAs minor groove (upper right of Fig. 28-48).  The protein also contacts the DNA backbone between the major and minor  grooves in this region (notably with Arg 273).   The structures most striking feature is that its DNA-binding motif consists  of conserved regions comprising the most frequently mutated residues in the >1000  p53 variants found in human tumors. Among them are one Gly and five Arg  residues (highlighted in yellow in Fig. 28-48) whose mutations collectively ac¬  count for over 40% of the p53 variants in tumors. The two most frequently  mutated residues, Arg 248 and Arg 273, as we saw, directly contact the DNA.  The other four “mutational hotspot” residues appear to play a critical role in  structurally stabilizing p53’s DNA-binding surface. The relatively sparse sec¬  ondary structure in the polypeptide segments forming this surface (one helix  and three loops) accounts for this high mutational sensitivity: Its structural in¬  tegrity mostly relies on specific side chain-side chain and side chain-backbone  interactions.   p53 Is a Sensor That Integrates Information from Several Pathways. p53   may be activated by several other pathways. For example, aberrant growth sig¬  nals, including those generated by oncogenic variants of Ras signaling cascade  components (Fig. 13-7) such as Ras, cause the inappropriate activation of a  variety of transcription factors. One of them, Myc, activates the transcription  of the gene encoding pl4 ARF , which binds to Mdm2 and thereby inhibits its  activity. This prevents the degradation of p53 and hence triggers the p53-  dependent transcriptional programs leading to cell cycle arrest as well as apop¬  tosis. Evidently, pl4 ARF acts as part of a p5 3-dependent fail-safe system to  counteract hyperproliferative signals.   A third activation pathway for p53 is induced by a wide variety of DNA-  damaging chemotherapeutic agents, protein kinase inhibitors, and UV radi¬  ation. These activate a protein kinase named ATR to phosphorylate p53 so    as to reduce its affinity for Mdm2 in much the same way as do ATM and  Chk2. p53 is also subject to a rich variety of reversible posttranslational  modifications that markedly influence the expression of its target genes, in¬  cluding acetylation at several Lys residues, glycosylation, and sumoylation  (Section 27-5B), in addition to its phosphorylation at multiple Ser/Thr  residues and ubiquitination.   p53, as we have only glimpsed, is the recipient of a vast number of intracel¬  lular signals and, in turn, controls the activities of a large number of downstream  regulators. One way to understand the operation of this highly complex and  interconnected network is in analogy with the Internet. In the Internet (cell),  a small number of highly connected servers or hubs (“master” proteins) trans¬  mit information to/from a large number of computers or nodes (other pro¬  teins) that directly interact with only a few other nodes (proteins). In such a  network, overall performance is largely unperturbed by the inactivation of one  of the nodes (other proteins). However, the inactivation of a hub (“master”  protein) will greatly impact system performance. p53 is a “master” protein,  that is, it is analogous to a hub. Inactivation of one of the many proteins that  influences its performance or one of the many proteins whose activity it in¬  fluences usually has little effect on cellular events due to the cells redundant  and highly interconnected components. However, the inactivation of p53 or  several of its most closely associated proteins (e.g., Mdm2) disrupts the cells  responses to DNA damage and tumor-predisposing stresses, thereby leading to  tumor formation.   T Loss of pRb Protein Leads to Cancer. Retinoblastoma, a cancer of the  developing retina that affects infants and young children, is associated with  the loss of the Rb gene, which encodes the tumor suppressor pRb. This 928-  residue DNA-binding protein interacts with the E2F family of transcription  factors, which has six members in mammals. E2F proteins induce the tran¬  scription of genes that encode proteins required for entry into S phase. pRb  can be phosphorylated at as many as 16 of its Ser/Thr residues by various  Cdk-cyclin complexes (the various complexes phosphorylate different sets of  sites on pRb). In nonproliferating cells (those in early G^, pRb is hypophos-  phorylated. In that state, it binds to E2F so as to prevent it from activating  transcription at the promoters to which it is bound. In response to a mito¬  genic signal (a signal that induces mitosis), the levels of D-type cyclins in¬  crease, which triggers phosphorylation of pRb by Cdk4/6-cyclin D  complexes. Hyperphosphorylated pRb releases E2F, which then induces the  expression of genes that promote cell cycle progression, including genes for  additional cyclins and Cdks. The E2F-binding site of pRb is the major site of  Rb gene alterations in tumors.   Programmed cell death or apoptosis (Greek: falling off, as leaves from a tree),  which was first described by John Kerr in the late 1960s, is a normal part of  development as well as maintenance and defense of the adult animal body. For  example, in many vertebrates, the digits of the developing hands and feet are  initially connected by webbing that is eliminated by programmed cell death  (Fig. 28-49), as are the tails of tadpoles and the larval tissues of insects  (Section 28-4D) during their metamorphoses into adults. In the adult human  body, which consists of nearly 10 14 cells, an estimated 10 11 cells are elimi-   FIG. 28-49 Programmed cell death in the embryonic mouse paw. At day 12.5 of   development, its digits are fully connected by webbing. At day 13.5, the webbing has begun  to die. By day 14.5, the apoptotic process is complete. [Reproduced by permission from  Wood, W., Turmaine, M., Weber, R., Camp, V., Maki, R.A., McKercher, S.R., and Martin,   nated each day through programmed cell death (which closely matches the Section 4 The Cell Cycle, Cancer, Apoptosis,   we annually lose in this manner approaches that of our entire body. The im¬  mune system eliminates virus-infected cells, in part by inducing them to un¬  dergo apoptosis, in order to prevent viral replication. Cells with irreparably  damaged DNA and hence at risk for malignant transformation undergo apop¬  tosis, thereby protecting the entire organism from cancer. In fact, as Martin  Raff pointed out, apoptosis appears to be the default option for animal cells:   Unless they continually receive external hormonal and/or neuronal signals not to  commit suicide , they will do so. Thus, adult organs maintain their constant size  by balancing cell proliferation with apoptosis. Not surprisingly, therefore, in¬  appropriate apoptosis has been implicated in several neurodegenerative diseases  including Alzheimer s disease (Section 6-5C), Parkinsons disease (Section 21-6B),  and Huntingtons disease (Box 28-1), as well as much of the damage caused  by stroke and heart attack.   Apoptosis is qualitatively different from necrosis, the type of cell death  caused by trauma (e.g., lack of oxygen, extremes of temperature, and mechani¬  cal injury). Cells undergoing necrosis essentially explode: They and their mem¬  brane-enclosed organelles swell as water rushes in through their compromised  membranes, releasing lytic enzymes that digest the cell contents until the cell ly¬  ses, spilling its contents into the surrounding region (Box 18-5). The cytokines  that the cell releases often induce an inflammatory response (which can damage  surrounding cells). In contrast, apoptosis begins with the loss of intercellular  contacts by an apparently healthy cell followed by its shrinkage, the condensa¬  tion of its chromatin at the nuclear periphery, the collapse of its cytoskeleton,  the dissolution of its nuclear envelope, the fragmentation of its DNA, and vio¬  lent blebbing (blistering) of its plasma membrane. Eventually, the cell disinte¬  grates into numerous membrane-enclosed apoptotic bodies that are  phagocytosed (engulfed) by neighboring cells as well as by roving macrophages  without spilling the cell contents and hence not inducing an inflammatory re¬  sponse. The phagocytotic cells recognize the apoptotic bodies through “eat me”  signals. In mammals, this signal is the phospholipid phosphatidylserine, which  in the plasma membrane of healthy cells occurs only in its inner leaflet (Fig.   9-32) but in apoptotic cells is translocated to the outer leaflet.   Caspases Participate in Apoptosis. Apoptosis involves a family of proteases  known as caspases (for cysteinyl ^artate-specific prot eases), which are cys¬  teine proteases whose mechanism resembles that of serine proteases  (Section 11-5) but with Cys replacing the active site Ser. Caspases cleave  target polypeptides after an Asp residue.   Caspases are a 2 (3 2 heterotetramers that consist of two large a  subunits (—300 residues) and two small (3 subunits (—100 residues).   They are expressed as single-chained zymogens (procaspases) that  are activated by proteolytic excision of their N-terminal  prodomains and proteolytic separation of their a and (3 subunits.   The activating cleavage sites all follow Asp residues and are, in  fact, targets for caspases, suggesting that caspase activation may  either be autocatalytic or be catalyzed by another caspase.   The X-ray structure of human caspase-7 (Fig. 28-50), deter¬  mined by Keith Wilson and Paul Charifson, reveals that each a(3    FIG. 28-50 X-Ray structure of caspase-7 in complex with a tetrapeptide  aldehyde inhibitor. The a 2 p 2 heterotetrameric enzyme is viewed along its twofold  axis with its large (a) subunits orange and gold and its small (|3) subunits cyan and light  blue. The acetyl-Asp-Glu-Val-Asp-CHO inhibitor is drawn in stick form with C green, N blue,  and 0 red. [Based on an X-ray structure by Keith Wilson and Paul Charifson, Vertex  Pharmaceuticals, Cambridge, Massachusetts. PDBid 1F1J.]   Chapter 28 Regulation of Gene Expression heterodimer contains a six-stranded (3 sheet flanked by five a helices that are   approximately parallel to the (3 strands. The (3 sheet is continued across the  enzymes twofold axis to form a twisted 12-stranded (3 sheet. The active site  of each a(3 heterodimer is located at the C-terminal ends of its parallel (3  strands. The structures of other caspases differ mainly in the conformations of  the four loops forming their active sites. In the inactive procaspase-7, these  loops are folded so as to obliterate the active site.   Over 60 cellular proteins have been identified as caspase substrates.  These include cytoskeletal proteins, proteins involved in cell cycle regulation  (including cyclin A, p21 Clpl , ATM, and pRb), proteins that participate in  DNA replication, transcription factors, and proteins that participate in sig¬  nal transduction. Nevertheless, how the cleavage of these numerous proteins  causes the morphological changes that cells undergo during apoptosis is un¬  clear. The induction of apoptosis also causes the rapid degradation of DNA  by the action of caspase-activated DNase. Presumably, DNA degradation  prevents the genetic transformation of other cells that subsequently phago-  cytose apoptotic bodies containing viral DNA or damaged chromosomal  DNA.   Apoptosis Is Triggered Extracellularly or Intracellularly. Apoptosis in a  given cell may be induced either by externally supplied signals in the so-called  extrinsic pathway (death by commission) or by the absence of external sig¬  nals that inhibit apoptosis in the so-called intrinsic pathway (death by omis¬  sion). The extrinsic pathway is initiated by the association of a cell destined  to undergo apoptosis with a cell that has selected it to do so. In what is  perhaps the best characterized such pathway (Fig. 28-51), a transmem¬  brane protein named Fas ligand (FasL) that projects from the plasma  membrane of the inducing cell, a so-called death ligand, binds to a trans¬  membrane protein known as Fas that projects from the plasma membrane  of the apoptotic cell, a so-called death receptor. Fas ligand is a ho-  motrimeric protein whose binding to three Fas molecules causes the Fas  cytoplasmic domains to trimerize. The trimerized Fas recruits three mole¬  cules of a 208-residue adaptor protein named FADD (for Aas-associating  ^/eath domain-containing protein), which in turn recruits procaspase-8  and procaspase-10. The consequent clustering of procaspase-8 and pro-  caspase-10 results in the proteolytic autoactivation of these zymogens,  thereby generating caspases-8 and -10, which are termed initiator cas¬  pases. This is because these enzymes then activate caspase-3, which is  known as an effector (executioner) caspase because its actions cause the  cell to undergo apoptosis.   The intrinsic pathway for initiating apoptosis follows a slightly different  route to caspase-3 activation. Most animal cells are continuously bathed in an  extracellular soup, generated in part by neighboring cells, that contains a wide  variety of substances that regulate the cells growth, differentiation, activity,  and survival. The withdrawal of this chemical support for its survival or the  loss of direct cell—cell interactions induces a cell to undergo apoptosis via the  intrinsic pathway. The initial step of this pathway appears to be the activation  of one or more members of the Bcl-2 family (so named because its founding  member is involved in B cell /ymphoma). Association of certain Bcl-2 proteins  with the mitochondrion causes it to release cytochrome c (Section 18-2E) from  the intermembrane space into the cytosol. It is not clear how cytochrome c  exits the mitochondrion. It may traverse the outer membrane via a newly  formed pore or via an existing pore whose conformation is altered to accom¬  modate the ~12-kD cytochrome, or the outer membrane may rupture as a  result of Bcl-2 activity.   The cytochrome c combines with a 1248-residue protein called Apaf-1  (for apoptosis protease-activating /actor- 1) and ATP or dATP to form an   The trimeric death ligand binds to  the death receptor, causing its (l  death domains (DD) to trimerize    This recruits adaptors,  which bind via their DDs to  the death receptor DDs.    The adaptors recruit an  initiator caspase zymogen  via interactions between  their death effector  domains (DEDs).    This autoactivates the initiator  caspase zymogen to yield a (4  heterotetrameric initiator caspase.    What features of this pathway would help prevent the accidental triggering of apoptosis?    FIG. 28-52 Cryoelectron microscopy-based image of the  human apoptosome at 9.5 A resolution, (a) In its top view, the  particle is viewed along its sevenfold axis of symmetry, (b) The side  view reveals the flattened nature of this ~250-A-diameter  wheel-shaped particle. Cytochrome c forms the bulge protruding from  the top of the large knob at the end of every arm. The magenta disk  is of lower resolution than the rest of the structure and hence  appears to be flexibly linked to it. [Courtesy of Christopher Akey,  Boston University School of Medicine.]    — 1100-kD wheel-shaped complex named the apoptosome (Fig. 28-52). The  apoptosome binds several molecules of procaspase-9 in a manner that induces  their autoactivation. The resulting caspase-9, still bound to the apoptosome,  then activates procaspase-3 to instigate cell death.   Perhaps the most awe-inspiring event in biology is the growth and develop¬  ment of a fertilized ovum to form an extensively differentiated multicellular  organism. No outside instruction is required to do so; fertilized ova contain  all the information necessary to form complex multicellular organisms such as hu¬  man beings. Much of what we know about the molecular basis of cell differ¬  entiation is based on studies of the fruit fly Drosophila melanogaster. We  therefore begin this section with a synopsis of Drosophila embryogenesis.   The Drosophila Embryo Is Divided into Segments. Almost immediately after  the Drosophila egg (Fig. 28-53 d) is laid (which, rather than the earlier fertiliza¬  tion, triggers development), it commences a series of rapid, synchronized nuclear  divisions, one every 6 to 10 min. Here, the nuclear division process is not ac¬  companied by the formation of new cell membranes; the nuclei continue shar¬  ing their common cytoplasm to form a so-called syncytium (Fig. 28-53 b). After  the eighth round of nuclear division, the —256 nuclei begin to migrate toward  the cortex (outer layer) of the egg where, by around the eleventh nuclear divi¬  sion, they have formed a single layer surrounding a yolk-rich core (Fig. 28-53 c;  the germ-cell progenitors, the pole cells, are set aside after the ninth division). At  this stage, the mitotic cycle time begins to lengthen while the nuclear genes,  which have heretofore been fully engaged in DNA replication, become transcrip¬  tionally active (a freshly laid egg contains an enormous store of mRNA that has  been contributed by the developing oocytes surrounding “nurse” cells). In the  fourteenth nuclear division cycle, which lasts —60 min, the eggs plasma mem¬  brane invaginates around each of the —6000 nuclei to yield a cellular monolayer  called a blastoderm (Fig. 28-53 d). At this point, after —2.5 h of development,  transcriptional activity reaches its maximum and mitotic synchrony is lost.   During the next few hours, the embryo undergoes gastrulation (migra¬  tion of cells to form a triple-layered structure) and organogenesis. A striking  aspect of this remarkable process, in Drosophila as well as in higher animals,  is the division of the embryo into a series of segments corresponding to the  adult organisms organization (Fig. 28-53<?). The Drosophila embryo has at least    FIG. 28-53 Development in Drosophila. The various stages are  explained in the text. Note that the embryos and newly hatched larva are  all the same size, —0.5 mm long. The adult is, of course, much larger.   The approximate numbers of cells in the early stages of development are  given in parentheses.    three segments that eventually merge to form its head (Md, Mx,  and Lb for mandibulary, maxillary, and labial), three thoracic  segments (T1-T3), and eight abdominal segments (A1-A8).  As development continues, the embryo elongates and several  of its abdominal segments fold over its thoracic segments (Fig.  28-53 f). At this stage, the segments become subdivided into  anterior (forward) and posterior (rear) compartments. The em¬  bryo then shortens and unfolds to form a larva that hatches 1  day after beginning development (Fig. 28-53 g). Over the next  5 days, the larva feeds, grows, molts twice, pupates, and com¬  mences metamorphosis to form an adult (Fig. 28-53 h). In the  latter process, the larval epidermis is almost entirely replaced  (through apoptosis) by the outgrowth of apparently undifferen¬  tiated patches of larval epithelium known as imaginal disks  that are committed to their developmental fates as early as the  blastoderm stage. These structures, which maintain the larva’s  segmental boundaries, form the adults legs, wings, antennae,  eyes, etc. About 10 days after commencing development, the  adult emerges and, within a few hours, initiates a new repro¬  ductive cycle.   Developmental Patterns Are Genetically Mediated. What is  the mechanism of embryonic pattern formation? Much of what  we know about this process stems from genetic analyses of a se¬  ries of bizarre mutations in three classes of Drosophila genes that  normally specify progressively finer regions of cellular special¬  ization in the developing embryo:   1. Maternal-effect genes , which define the embryo's polarity,  that is, its anteroposterior (head to tail) and dorsoventral  (back to belly) axes. Mutations of these genes globally al¬  ter the embryonic body pattern, producing, for example,  nonviable embryos with two anterior or two posterior  ends pointing in opposite directions.   2. Segmentation genes , which specify the correct number  and polarity of embryonic body segments. Investigations by  Christiane Niisslein-Volhard and Eric Wieschaus led to  their subclassification as follows:   (a) Gap genes, the first of a developing embryo’s to be  transcribed, are so named because their mutations  result in gaps in the embryos segmentation pattern.  Embryos with defective hunchback (hb) genes, for  example, lack mouthparts and thorax structures.   (b) Pair-rule genes specify the division of the embryo’s  broad gap domains into segments. These genes are  so named because their mutations usually delete  portions of every second segment.   (c) Segment polarity genes specify the polarities of the  developing segments. Thus, homozygous engrailed  (en) mutants lack the posterior compartment of  each segment.   Drosophila, (a) Head and thorax of a wild-type  adult fly (left) and one that is homozygous for a  mutant form of the homeotic Antennapedia (antp)  gene (right). The mutant gene is inappropriately  expressed in the imaginal disks that normally  form antennae (where the wild-type antp gene is  not expressed) so that they develop as the legs  that normally occur only on segment T2.  [Courtesy of Gines Morata, Universidad  Autonoma de Madrid, Spain.] (b) A four-winged  Drosophila (it normally has two wings) that  results from the presence of three mutations in  the bithorax complex. These mutations cause the  normally haltere-bearing segment T3 to develop  as if it were the wing-bearing segment T2.    3. Homeotic selector genes, which specify segmental identity.  Mutations of homeotic selector genes transform one body part  into another. For instance, Antennapedia (antp, antenna-foot)  mutants have legs in place of antennae (Fig. 28-54#), whereas  the mutations bithorax (bx), anterior bithorax (abx), and  postbithorax (pbx) each transform sections of halteres (vestig¬  ial wings that function as balancers), which normally occur  only on segment T3, to the corresponding sections of wings,  which normally occur only on segment T2 (Fig. 28-54 £).    The properties of maternal-effect gene mutants suggest that mater¬  nal-effect genes specify substances known as morphogens whose distri¬  butions in the egg cytoplasm define the future embryos spatial  coordinate system. Indeed, immunofluorescence studies by Niisslein-  Volhard have demonstrated that the product of the bicoid (bed)  gene is distributed in a gradient that decreases toward the posterior  end of the normal embryo (Fig. 28-55 d), whereas embryos with  ^/-deficient mothers lack this gradient. The gradient arises through  the secretion, by ovarian nurse cells, of bed mRNA into the ante¬  rior end of the oocyte during oogenesis. The nanos gene mRNA is  similarly deposited near the eggs posterior pole. The bed and nanos  gene products regulate the expression of specific gap genes. Some  other maternal-effect genes specify proteins that trap the localized  mRNAs in their area of deposition. This explains why early em¬  bryos produced by females homozygous for maternal-effect muta¬  tions can often be “rescued” by the injection of cytoplasm, or  sometimes just the mRNA, from early wild-type embryos.   The mRNA of the gap gene hunchback (hb) is deposited uni¬  formly in the unfertilized egg (Fig. 28-55 a). However, Bicoid pro¬  tein activates the transcription of the embryonic hb gene, whereas  Nanos protein inhibits the translation of hb mRNA.  Consequently, Hunchback protein becomes distributed in a gra¬  dient that decreases from anterior to posterior (Fig. 28-55 £). Footprinting stud¬  ies have demonstrated that Bicoid protein binds to five homologous sites  (consensus sequence TCTAATCCC) in the hb genes upstream promoter region.   Hunchback protein controls the expression of several other gap genes (Fig.  28-55 c,d): High levels of Hunchback protein induce giant expression;  Kriippel (German: cripple) is expressed where the level of Hunchback protein  begins to decline; knirps (German: pigmy) is expressed at even lower levels of  Hunchback protein; and giant is again activated in regions where Hunchback  protein is undetectable. These patterns of gene expression are stabilized and  maintained by additional interactions. For example, Kriippel protein binds  to the promoters of the hb gene, which it activates, and the knirps gene, which  it represses. Conversely, Knirps protein represses the Kriippel gene. This mu¬  tual repression is thought to be responsible for the sharp boundaries between  the various gap domains.   Pair-rule genes are expressed in sets of seven stripes, each just a few nu¬  clei wide, along the early embryos anterior-posterior axis (Fig. 28-56). The  gap gene products directly control three primary pair-rule genes: hairy,  even-skipped (eve), and runt. The promoters of most primary pair-rule genes  consist of a series of modules, each of which contains a particular arrange¬  ment of activating and inhibitory binding sites for the various gap gene pro¬  teins. As a result, the expression of a pair-rule gene reflects the combination  of gap gene proteins present, giving rise to a “zebra stripe” pattern. As with  the gap genes, the patterns of expression of the primary pair-rule genes be¬  come stabilized through interactions among themselves. The primary pair-  rule gene products also induce or inhibit the expression of five secondary  pair-rule genes including fushi tarazu (ftz; Japanese for not enough   FIG. 28-55 The formation and effects of the Hunchback protein  gradient in Drosophila embryos, (a) The unfertilized egg contains  maternally supplied bicoid and nanos mRNAs placed at its anterior and  posterior poles, together with a uniform distribution of hunchback mRNA.   (b) On fertilization, the three mRNAs are translated. Bicoid and Nanos  proteins are not bound in place as are their mRNAs and hence their  gradients are broader than those of the mRNAs. Bicoid protein stimulates  the translation of hunchback mRNA whereas Nanos protein inhibits its  translation, resulting in a gradient of Hunchback protein that decreases  nonlinearly from anterior to posterior, (c) Specific concentrations of  Hunchback protein induce the transcription of the giant, Kruppel, and knirps  genes. The gradient of Hunchback protein thereby specifies the positions  at which the latter mRNAs are synthesized, (d) A photomicrograph of a  Drosophila embryo ( anterior end left) that has been immunofluorescently  stained for both Hunchback (green) and Kruppel proteins (red). The region  where the proteins overlap is yellow. [Parts a, b, and c after Gilbert, S. F.,  Developmental Biology (5th ed.), pp. 550 and 565, Sinauer Associates  (1997); Part d courtesy of Jim Langeland, Stephen Paddock, and Sean  Carroll, Howard Hughes Medical Institute, University of Wisconsin—   segments). Thus, as Walter Gehring demonstrated , ftz transcripts first appear  in the nuclei lining the cortical cytoplasm during the embryos tenth nuclear  division cycle. By the fourteenth division cycle, when the cellular blastoderm  forms, ftz is expressed in a pattern of seven belts around the blastoderm, each  3 or 4 cells wide (Fig. 28-56).   The expression of eight known segment polarity genes is initiated by pair-  rule gene products. For example, by the thirteenth nuclear division cycle, as  Thomas Kornberg demonstrated, engrailed (en) transcripts become detectable  but are more or less evenly distributed throughout the embryonic cortex.  However, since en is preferentially expressed in nuclei containing high concen¬  trations of either Eve or Ftz proteins, by the fourteenth cycle they form a strik¬  ing pattern of 14 stripes around the blastoderm (half the spacing of ftz  expression). The en gene product thereby induces the posterior half of each  segment to develop in a different fashion from its anterior half.   FIG. 28-56 Drosophila embryo stained  for pair-rule genes. The Fushi tarazu  protein (Ftz) is stained brown, and the Eve  protein is stained gray. These proteins are each  expressed in seven stripes. [Courtesy of Peter  Lawrence, MRC Laboratory of Molecular Biology,  Cambridge, U.K.]   Homeotic Genes Direct Development of Individual Body Parts. The struc¬  tural components of developmental^ analogous body parts, say Drosophila an¬  tennae and legs, are nearly identical; only their organizations differ (Fig. 28-57).  Consequently, developmental genes must control the pattern of structural gene expres¬  sion rather than simply turning the genes on or off.   The Drosophila homeotic selector genes map into two large gene families:  the bithorax complex (BX-C), which controls differentiation in the thoracic  and abdominal segments, and the antennapedia complex (ANT-C), which  primarily affects head and thoracic segments. Homozygous mutations in BX-C  cause one or more segments to develop as if they were more anterior segments (e.g.,  segment T3 develops as if it were segment T2; Fig. 28-54 £). The entire dele¬  tion of BX-C causes all segments posterior to T2 to resemble T2; apparently  T2 is the developmental “ground state” of the more distal segments. The evo¬  lution of homeotic gene families, it is thought, permitted arthropods (the phy¬  lum containing insects) to arise from the more primitive annelids (segmented  worms) in which all segments are nearly alike.   Detailed genetic analysis of BX-C led Edward B. Lewis to formulate a  model for segmental differentiation (Fig. 28-58): BX-C ’ Lewis proposed, con¬  tains at least one gene for each segment from T3 to A8 (numbered 0 to 8 in  Fig. 28-58). Starting with segment T3, progressively more posterior segments  express successively more BX-C genes until, in segment A8, all of the genes are  expressed. Such a pattern of gene expression may result from a gradient in the  concentration of a BX-C repressor that decreases from the anterior to the pos¬  terior end. The developmental fate of a segment is thereby determined by its  position in the embryo. Subsequent sequence analysis revealed that the nine  “genes” in the Lewis model are actually enhancer elements on three BX-C genes.   In characterizing the Antennapedia (antp) gene, Gehring and Matthew  Scott independently discovered that antp cDNA hybridizes to both the antp  and the ftz genes, indicating that these genes share a common base sequence.  Subsequent experiments revealed that a similar sequence, called a homeo-  domain or homeobox, occurs in many Drosophila homeotic genes. These se¬  quences, which are 70 to 90% identical to one another, encode even more  identical 60-residue polypeptide segments.   Further hybridization studies using homeodomain probes led to the truly  astonishing finding that homeodomains are also present in the genomes of many  animals. Homeodomain-containing genes have collectively become known as  Hox genes. In vertebrates, they are organized in four clusters of 9 to 11 genes,  each located on a separate chromosome and spanning more than 100 kb. In    H T1 T2 T3 A1 A2 A3 A4 A5 A6 A7 A8  Segments    FIG. 28-58 Model for the differentiation  of embryonic segments in Drosophila.   Segments T2, T3, and A1-8, as the lower  drawing indicates, are each characterized by a  unique combination of active ( purple circles) and  inactive ( yellow circles) BX-C “genes.” These  “genes” (which are really enhancer elements),  here numbered 0 to 8, are thought to be  sequentially activated from anterior to posterior  in the embryo so that segment T2, the  developmental^ most primitive segment, has no  active BX-C genes, while in segment A8, all of  them are active. [After Ingham, P., Trends Genet  1, 113 (1985).]    contrast, Drosophila, as we saw, have two Hox clusters, whereas nematodes,  which are evolutionarily more primitive than insects, have only one Hox clus¬  ter. The various Hox clusters, as well as their component genes, almost cer¬  tainly arose through a series of gene duplications.   Hox Genes Encode Transcription Factors. Some Hox genes are remarkably sim¬  ilar; for example, the homeodomains of the Drosophila antp gene and the frog  MM3 gene encode polypeptides that have 59 of their 60 amino acids in com¬  mon. Since vertebrates and invertebrates diverged over 600 million years ago, this  strongly suggests that the product of the homeodomain has an essential function.   The polypeptide encoded by the homeodomain of the Drosophila engrailed  gene specifically binds to the DNA sequences just upstream from the tran¬  scription start sites of both the en and the ftz genes. Moreover, fusing the ftz  genes upstream sequence to other genes imposes fizs pattern of stripes (Fig.  28-56) on the expression of these genes in Drosophila embryos. These obser¬  vations suggest that homeodomain-containing genes encode transcription factors  that regulate the expression of other genes.   Thomas Kornberg and Carl Pabo determined the X-ray structure  of the 61-residue homeodomain from the Drosophila Engrailed pro¬  tein in complex with a 21-bp DNA (Fig. 28-59). The homeodomain  consists largely of three a helices, the last two of which form an HTH    FIG. 28-59 X-Ray structure of the Engrailed protein homeodomain  in complex with its target DNA. The protein is shown in ribbon form colored  in rainbow order from its N-terminus (blue) to its C-terminus (red). The DNA is  shown in stick form colored according to atom type (C green, N blue, 0 red, and  P orange) with the nucleotides forming its TAAT subsite highlighted with C atoms  cyan. Successive P atoms in the same chain are linked by orange rods. Note that  the protein’s recognition helix, its C-terminal helix (red and orange), is bound in the  DNA’s major groove, and that the protein’s N-terminal segment (blue) is inserted into  the DNA’s minor groove. [Based on an X-ray structure by Carl Pabo, MIT. PDBid  1HDDJ O See Interactive Exercise 59.    FIG. 28-60 Pattern of expression of the  Hox-3.1 gene in a 12.5-day-old mouse  embryo. The protein-coding portion of the  Hox-3.1 gene was replaced by the lacZ gene.  The regions of this transgenic embryo in which  Hox-3.1 is expressed are revealed by  soaking the embryo in a buffer containing a  substance (X-gal; Section 3-5A) that turns blue  when hydrolyzed by the lacZ gene product,  p-galactosidase. [Courtesy Yvan Lallemand and  Phillipe Brulet, College de France and the  Institut Pasteur, France.]   • Summarize the stages of the cell cycle.  What is the role of cyclins?   • Summarize the sequence of events in  Drosophila embryogenesis and explain the  functions of the substances that control  these events.    motif that is closely superimposable on the HTH motifs of prokaryotic re¬  pressors (Section 24-4B).   Vertebrate Hox genes, like those of Drosophila , are expressed in specific pat¬  terns and at particular stages during embryogenesis. That the Hox genes directly  specify the identities and fates of embryonic cells was shown, for example, by  the following experiment. Mouse embryos were made transgenic for the Hox-  1.1 gene that had been placed under the control of a promoter that is active  throughout the body even though Hox-1.1 is normally expressed only below the  neck. The resulting mice had severe craniofacial abnormalities such as a cleft  palate and an extra vertebra and an intervertebral disk at the base of the skull.  Some also had an extra pair of ribs in the neck region. Thus, the altered expres¬  sion of the Hox-1.1 gene induced a homeotic mutation, that is, a change in the  development pattern, analogous to those observed in Drosophila (Fig. 28-54).   Homozygotic mice whose Hox-3.1 coding sequence has been replaced with  that of lacZ are born alive but usually die within a few days. They exhibit  skeletal deformities in their trunk regions in which several skeletal segments  are transformed into the likenesses of more anterior segments. The pattern of  P-galactosidase activity, as colorimetrically detected through the use of a sub¬  strate analog whose hydrolysis products are blue (Fig. 28-60), indicates that  Hox-3.1 deletion modifies the properties but not the positions of the embry¬  onic cells that normally express Hox-3.1.   • Genomic DNA sequence data reveal the total number of genes,  their probable functions, and possible links to disease.   • Certain genes are found in clusters, for example, those in bac¬  terial operons, rRNA and tRNA genes, and eukaryotic histone  genes. The human globin gene clusters contain genes expressed at  different developmental stages.    DNA, which include control regions for replication and transcrip¬  tion. The genomes of higher eukaryotes contain much larger pro¬  portions of nontranscribed DNA in the form of repetitive  sequences, many of them the remnants of transposons.   • Prokaryotic gene expression is controlled primarily at the level of  transcription. Regulation of the lac operon is mediated by the   binding of the lac repressor to its operator sequences. This bind¬  ing, which prevents transcription of the operon, is reversed by the  binding to the lac repressor of an inducer whose presence signals  the availability of lactose, the substrate for the lac operon-encoded  enzymes.   • In catabolite repression, a complex of CAP and cAMP, which sig¬  nals the scarcity of glucose, binds to its target DNA to stimulate  the transcription of genes encoding proteins that participate in the  metabolism of other sugars.   • Attenuation is a mechanism whereby the translation-dependent  formation of alternate mRNA secondary structures in an operon’s  leader sequence determines whether transcription proceeds or ter¬  minates. In a riboswitch, metabolite binding to an mRNA regu¬  lates gene expression.   • The expression of eukaryotic genetic information involves the  repositioning of nucleosomes and the covalent modification of  histones as part of the histone code that is read by regulatory  proteins. DNA methylation allows epigenetic inheritance.   • In eukaryotes, activators and repressors, which bind to DNA en¬  hancers and silencers, act cooperatively to regulate the rate of tran¬  scription initiation. Some of these transcription factors are  activated by hormone signaling.   • Eukaryotic gene expression may also be controlled by variable rates  of mRNA degradation and regulation of translation initiation.   RNA interference (RNAi) is a posttranscriptional gene silencing  pathway in which an RNA segment directs the selective degrada¬  tion of a complementary mRNA. MicroRNAs (miRNAs) are tran¬  scriptional regulators.   • Antibody diversity results from somatic recombination involving  the selection of individual members of clustered gene sequences  encoding the different segments of the immunoglobulin light and  heavy chains. Diversity is augmented by imprecise V/D/J joining  and by somatic hypermutation.   • The eukaryotic cell cycle is governed by cyclin-dependent kinases,  whose targets include the tumor suppressors p53 and pRb. p53  functions as a transcriptional activator that detects a variety of  pathological states such as DNA damage and thereon helps main¬  tain cell cycle arrest and, if the damage cannot be repaired, apop¬  tosis (programmed cell death).   • Extracellular or intracellular signals may trigger apoptosis, a cell-  suicide pathway that requires the activities of various caspases.   • The development of the Drosophila embryo is controlled by ma¬  ternal-effect genes, which define the embryo’s polarity; gap, pair-  rule, and segment polarity genes, which specify the number and  polarity of embryonic body segments; and homeotic selector genes  (Hox genes), which encode transcription factors that regulate the  expression of genes and therefore govern cell differentiation. Hox  genes similarly regulate vertebrate development.    gene expression 1013  genomics 1014  C value 1014  C-value paradox 1014  ORF 1016  EST 1016  CpG island 1016  orphan gene 1016  ncRNA 1017  SNP 1017  GWAS 1017  gene cluster 1017  centromere 1019  genetic anticipation 1020  highly repetitive DNA 1020  STR 1020  moderately repetitive  DNA 1022  selfish DNA 1022  polymorphism 1023    inducer 1024  repressor 1024  operator 1024  catabolite repression 1026  co repressor 1028  leader sequence 1029  attenuation 1029  ap tamer 1031  riboswitch 1031  heterochromatin 1032  euchromatin 1032  chromatin-remodeling  complex 1033  Barr body 1 033  HMG protein 1034  histone code 1036  HAT 1037  bromodomain 1038  chromodomain 1040  epigenetics 1042    genomic imprinting 1 043  enhancer 1043  silencer 1043  enhanceosome 1044  squelching 1045  insulator 1046  JAK-STAT 1046  hormone response  element 1048  exosome 1050  P body 1050   nonsense-mediated decay 1050  antisense RNA 1051  RNAi 1051  siRNA 1052  miRNA 1052  somatic recombination 1056  somatic hypermutation 1059  cell cycle 1059    cyclin 1060  tumor suppressor 1061  lincRNA 1062  apoptosis 1064  necrosis 1065  caspase 1065  syncytium 1068  blastoderm 1068  gastrulation 1068  imaginal disk 1069  maternal-effect gene 1069  segmentation gene 1069  gap gene 1069  pair-rule gene 1069  segment polarity gene 1069  homeotic selector gene 1070  morphogen 1070  homeodomain 1072  Hox gene 1072    1. DNA isolated from an organism can be sheared into fragments  of uniform size (—1000 bp), heated to separate the strands, then  cooled to allow complementary strands to reanneal. The renatu-  ration process can be followed over time. Explain why the rena-  turation of E. coli DNA is a monophasic process whereas the    renaturation of human DNA is biphasic (an initial rapid phase  followed by a slower phase).   2. Explain why the organization of genes in operons facilitates the  assignment of functions to previously unidentified ORFs in a  bacterial genome.   3. The genome of Daphnia pulex , a small freshwater crustacean, in¬  cludes approximately 30,000 genes in —200,000 kb of DNA.  How does the genome compare to that of Drosophila  melanogaster, another arthropod, and to that of humans?   4. The marine alga Ostreococcus tauri has an unusually high gene  density for a eukaryote: —8000 genes in an — 13,000-kb genome.  Assuming that intergene distances are negligible, what is the av¬  erage size of an O. tauri gene? How does this compare to the av¬  erage gene size in the prokaryote E. col?.   3. A possible mechanism for the pathology of trinucleotide repeat  diseases is aberrant translation in which all three reading frames  are used. Which repeating amino acid residues will result from  the translation of transcripts made from repeating DNA se¬  quences of (a) CAG and (b) CTG?   6. In addition to SNPs, humans exhibit copy-number polymor¬  phisms (CNPs), which result from relatively large deletions or in¬  sertions of certain segments of the genome. One study showed  that individuals differed by 11 CNPs with an average length of  463 kb. Calculate the fraction of the human genome represented  by copy-number variants.   7. Explain why inactivation of the O x sequence of the lac operator  almost completely abolishes repression of the lac operon.   8. Explain why (a) inactivation of the 0 2 or 0 3 sequence of the lac  operon causes only a twofold loss in repression, and (b) inactiva¬  tion of both 0 2 and 0 3 reduces repression —70-fold.   9. Why do E. coli cells with a defective lacZ gene fail to show galac-  toside permease activity after the addition of lactose in the ab¬  sence of glucose?   10. Describe the probable genetic defect that abolishes the sensitiv¬  ity of the lac operon to the absence of glucose when other meta¬  bolic operons continue to be sensitive to the absence of glucose.   12. Predict the effect of deleting the leader peptide sequence on reg¬  ulation of the trp operon.   13. Red-green color blindness is caused by an X-linked recessive ge¬  netic defect. Hence females rarely exhibit the red-green color¬  blind phenotype but may be carriers of the defective gene. When  a narrow beam of red or green light is projected onto some ar¬  eas of the retina of such a female carrier, she can readily differ¬  entiate the two colors but on other areas she has difficulty in  doing so. Explain.   14. Why is transcriptionally active chromatin —10 times more sus¬  ceptible to cleavage by DNase I than transcriptionally silent chro¬  matin?   15. Draw the molecular formula of the covalently modified histone  side chain of acetyllysine. How does this modification alter the  chemical properties of the side chain?   16. Draw the molecular formulas of the covalently modified histone  side chains of methyllysine and methylarginine. How do these  modifications alter the chemical properties of the side chains?   17. Explain why a deficiency of the vitamin folic acid could lead to  undermethylation of histones and DNA.   18. Monomethylation of histone Arg residues can be reversed by the  action of a peptidylarginine deiminase, which requires water to  remove the methyl group along with the imino group of Arg.    Draw the structure of the resulting amino acid side chain and  identify this nonstandard amino acid.   19. Is it possible for a transcription enhancer to be located within  the protein-coding sequence of a gene? Explain.   20. Proliferation and differentiation of T and B lymphocytes during  an immune response are mediated by signaling that leads to phos¬  phorylation of a protein called IkB. Unphosphorylated IkB binds  to and inhibits the transcription factor NF-kB. Phosphorylated  IkB releases NF-kB and is then ubiquitinated and degraded by  a proteasome. (a) Explain why this mechanism for activating  NF-kB allows rapid changes in gene expression, (b) Explain why  directly phosphorylating NF-kB might not be an effective mech¬  anism for altering gene expression, (c) A protein produced by  Yersinia pestis (the bacterium that causes plague) removes the  ubiquitin molecules from IkB. How would this affect the im¬  mune response to the Yersinia infection?   21. Explain why natural selection has favored the instability of RNA.   22. Explain why RNAi would be a less efficient mechanism for reg¬  ulating the expression of specific genes if Dicer hydrolyzed  double-stranded RNA every 11 bp rather than every 22 bp.   23. How many different heavy chain variable regions can theoreti¬  cally be generated by somatic recombination in humans (ignore  junctional flexibility)? If each of these heavy chains could com¬  bine with any of the —2000 k light chains, how many different  immunoglobulins could be produced?   24. VID!J recombination frequently yields a gene whose mRNA can¬  not be successfully translated into an immunoglobulin chain.  What aspect of somatic recombination is likely to produce non¬  productive gene rearrangement?   25. B cells suppress the expression of all but one heavy chain allele  and one light chain allele, a process known as allelic exclusion,  by inhibiting the further somatic recombination of heavy and  light chain genes after a productive recombination has occurred.  Describe the aberrant properties of immunoglobulins produced  by a B cell in which allelic exclusion is defective.   26. Explain what might happen if the activation-induced cytidine  deaminase (AID) were activated in a cell other than a B cell.   27. Why is it disadvantageous for single-celled eukaryotes such as  yeast to undergo apoptosis?   28. During apoptosis, phosphatidylserine (Section 9-1C) undergoes  transverse movement (flip-flop). 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Schmid, M. and Jensen, T.H., The exosome: a multipurpose RNA-decay  machine, Trends Biochem. Sci. 33, 501-510 (2008).   Shabbazian, M. and Grunstein, M., Functions of site-specific histone  acetylation and deacetylation. Annu. Rev. Biochem. 76 , 75-100  (2007).   Shilatifard, A., Chromatin modifications by methylation and ubiquitina-  tion: Implication in the regulation of gene expression. Annu. Rev.  Biochem. 75, 243-269 (2006).   Suganuma, T. and Workman, J.L., Signals and combinatorial functions of  histone modifications, Annu. Rev. Biochem. 80 , 473-499 (2011).   Turner, B.M., Cellular memory and the histone code, Cell 111 , 285-291  ( 2002 ).    1. A Thiol (sulfhydryl) group  B Carbonyl group  C Amide linkage   Volume = (4/3 ) , tt^ = (4/3)tt(3 X 10 7 m ) 3   = 3.24 X 10“ 19 m 3 = 3.24 X 10“ 16 L  Moles of protein = (2 molecules)/(6.022 X 10 23 molecules • mol x )    Concentration = (3.32 X 10 24 mol)/(5.24 X 10 16 L)   5. (a) Liquid water; (b) ice has less entropy at the lower temperature.  7. (a) T= 273 + 10 = 283 K  AC = AH - TAS  AG = 15 kj - (283 K)(0.05 kj • K _1 )   A G is greater than zero, so the reaction is not spontaneous.   9. In order for AC to have a negative value (a spontaneous reaction),  TAS must be greater than AH.    50 + x = 25(50 - x)  50 + x = 1250 - 25x  26x = 1200   [R] = 50 |xM + 46.15 fxM = 96.15 fxM  [Q] = 50 |xM - 46.15 fxM = 3.85 fxM  17. This strategy will NOT work because Reaction 1 has a negative  enthalpy change, releasing heat, and will therefore become more  favorable with decreasing temperature, whereas Reaction 2, which has  a positive enthalpy change, will become less favorable. Thus decreas¬  ing the temperature will favor Reaction 1, not Reaction 2. In order  to make Reaction 2 more favorable, the temperature must be raised.   To calculate the amount that the temperature must be raised,  Equation 1-18 may be used as follows:   Hence to increase K 2 IK x from 1 to 10, the temperature must be  raised from 298 K to 332 K.    R is 25 times greater than the concentration of Q. When equal con¬  centrations of Q and R are mixed, molecules of Q will be converted  to molecules of R.   (b) Let x = amount of Q converted to R, so that [R] will be 50 |JiM    1 . (a) Donors: NH 1 , NH 2 at C 2 , NH9; acceptors: N3, O at C 6 , N7.  (b) Donors: NH + , NH 2 at C4; acceptors: O at C 2 , N3. (c) Donors:  NH 3 group, OH group; acceptors: COO group, OH group.   5. From most soluble (most polar) to least soluble (least polar): c, b, e, a, d.   7. Water molecules move from inside the dialysis bag to the surround¬  ing seawater by osmosis. Ions from the seawater diffuse into the dial¬  ysis bag. At equilibrium, the compositions of the solutions inside and  outside the dialysis bag are identical. If the membrane were solute-  impermeable, essentially all the water would leave the dialysis bag.   9. The high solute concentration of honey tends to draw water out of  microorganisms by osmosis, thereby preventing their growth.   11 . The high concetration of bicarbonate in the dialysate means that  some bicarbonate will diffuse from the dialysate across the dialysis  membrane into the patients blood, where it will combine with and  neutralize excess protons.    13. pH 4, NHj; pH 8 , NH 4 ; pH 11, NH 3 .   17. The increase in [H + ] due to the addition of HC1 is (30 mL)  (1 mM)/(250 mL) = 0.2 mM = 2 X 10 -4 M. Because the [H + ] of  pure water, 10 7 M, is relatively insignificant, the pH of the solu¬  tion is equal to — log(2 X 10 4 ) or 3.7.   19. The pi 6 corresponding to the equilibrium between H 2 PO 4 (HA) and  HPO 4 (A ) is 6.82 (Table 2-4). The concentration of A is  (50 mL)(2.0 M)/(200 mL) = 0.5 M, and the concentration of HA  is (25 mL)(2.0 M)/(200 mL) = 0.25 M. Substitute these values into  the Henderson-Hasselbalch equation (Eq. 2-9):   21. Let HA = sodium succinate and A = disodium succinate.   [A - ] + [HA] = 0.05 M, so [A - ] = 0.05 M - [HA]   log([A“]/[HA]) = pH - p K= 6.0 - 5.64 = 0.36   [A - ] = 0.05 M - 0.015 M = 0.035 M   grams of sodium succinate = (0.015 mol • L -1 )(l40 g • mol -1 ) X  (1 L) = 2.1 g   grams of disodium succinate = (0.035 mol • L -1 )(162 g • mol -1 )  X (1 L) = 5.7 g   23. The standard free energy change can be calculated using Eq. 1-16  and the value of K from Table 2-4.   = -(8.314 J • K -1 • mol -1 )(298 K) ln(3.39 X 10 -8 )   = 42,600 J • mol 1 = 42.6 kj • mol 1   25. The dissociation of TrisH + to its basic form and H + is associated  with a large, positive enthalpy change. Consequently, heat is taken  up by the reactant on dissociation. When the temperature is lowered,  there is less heat available for this process, shifting the equilibrium  constant toward the associated form (the effect of temperature on the  equilibrium constant of a reaction is given by Eq. 1-18). To avoid  this problem, the buffer should be prepared at the same temperature  as its planned use.   7. Since the haploid genome contains 21 % G, it must contain 21 % C  (because G = C) and 58% A + T (or 29% A and 29% T, because  A = T). Each cell is diploid, containing 90,000 kb or 9 X 10 7 bases.  Therefore,    A = T = (0.29)(9 X 10 7 ) = 2.61 X 10 7 bases  C = G = (0.21)(9 X 10 7 ) = 1.89 X 10 7 bases    13. The high pH eliminates hydrogen bonds between bases, making it  easier to separate the strands of DNA.   15. According to the central dogma, DNA serves as a template for RNA  synthesis. The HIV enzyme, called reverse transcriptase, works in re¬  verse by synthesizing DNA from an RNA template.   19. (a) Newly synthesized chains would be terminated less frequently, so  the bands representing truncated fragments on the sequencing gel  would appear faint.   (b) Chain termination would occur more frequently, so longer frag¬  ments would be less abundant.    21. In O. tauri , the gene density is 8000 genes/13,000 kb = 0.62, a value  that is somewhat lower than that of the prokaryote E. coli (4300  genes/4639 kb = 0.93) but less than that of the plant A. thaliana  (25,500 genes/119,200 kb = 0 . 21 ).   23. The C. elegans genome contains 97,000 kb, so  f= 5/97,000 = 5.2 X 10 -5   N = log(l - 0.99)/log(l - 5.2 X 10 -5 )   25. The genomic library contains DNA sequences corresponding to all  the organism’s DNA, which includes genes and nontranscribed se¬  quences. A cDNA library represents only the DNA sequences that  are transcribed into mRNA.   27. (a) Only single DNA strands of variable length extending from the re¬  maining primer would be obtained. The number of these strands would  increase linearly with the number of cycles rather than geometrically,  (b) PCR would yield a mixture of DNA segments whose lengths corre¬  spond to the distance between the position of the primer with a single  binding site and the various sites where the multispecific primer binds.  29. ATAGGCATAGGC and CTGACCAGCGCC.   1. Gly and Ala, Ser and Thr; Val, Leu and lie; Asn and Gin; Asp and  Glu.   7. Hydrogen bond donors: a-amino group, amide nitrogen. Hydrogen  bond acceptors: a-carboxylate group, amide carbonyl.   9. The polypeptide would be even less soluble than free Tyr, because  most of the amino and carboxylate groups that interact with water  and make Tyr at least slightly soluble are lost in forming the peptide  bonds in poly(Tyr).   27. Isopeptide bonds can form from the side chain amino group of Lys  and from the side chain carboxylate groups of Asp and Glu.   3. Peptide B, because it contains more Trp and other aromatic residues.   5. Lowering the pH from 7.0 to 5.0 would promote the precipitation  of protein Q because the protein will be least soluble when its net  charge is zero (when pH = p/).   9. The protein behaves like a larger protein during gel filtration, sug¬  gesting that it has an elongated shape. The mass determined by SDS-  PAGE is more accurate since the mobility of a denatured SDS-coated  protein depends only on its size.   1 1. Because protein 1 has a greater proportion of hydrophobic residues  (Ala, lie, Pro, Val) than do proteins 2 and 3, hydrophobic interac¬  tion chromatography could be used to isolate it.    (a) The pA°s of the ionizable side chains (Table 4-1) are 3.90 (Asp)  and 10.54 (Lys); assume that the terminal Lys carboxyl group has a  p K of 3.5 and the terminal Ala amino group has a p K of 8.0  (Section 4-ID). The p/is approximately midway between the pAs  of the two ionizations involving the neutral species (the p K of Asp  and the N-terminal p K)\   (b) The net charge at pH 7.0 is 0 (as drawn above).   17. (a) chiral; (b) nonchiral; (c) chiral; (d) chiral; (e) nonchiral; (f) chiral;  (g) chiral.   25. (a) Serine (A^-acetylserine); (b) lysine (5-hydroxylysine); (c) methion¬  ine (A^-formylmethionine)    (b) The DEAE chromatography step results in only a 47% yield,  while the affinity chromatography step results in an 80% yield from  the step before it. The DEAE chromatography step therefore results  in the greatest loss of Mb.   (c) The DEAE chromatography step results in a 1.3-fold purification,  while the affinity chromatography step results in a 87-fold purifica¬  tion from the previous step. The affinity chromatography step there¬  fore results in the greatest purification of Mb.   (d) The affinity chromatography step is the best choice for a one-step  purification of Mb in this example.   17. Dansyl chloride reacts with primary amino groups, including the  8 -amino group of Lys residues.   ' 19. Thermolysin would yield the most fragments (9) and endopeptidase  V8 would yield the fewest (2).   21 . (a) The positive charges are caused by the protonation of basic side  chains (H, K, and R) and the N-terminal amino groups of the  protein.   (b) There are 1 H, 6 K, 11 R, and 1 NH 2 at the N-terminus.  Therefore, the maximum number of positive charges that can be  obtained is 19.   27. Because the side chain of Gly is only an H atom, it often occurs in  a protein at a position where no other residue can fit. Consequently,  Gly can take the place of a larger residue more easily than a larger  residue, such as Val, can take the place of Gly.   3 . (100 residues)(1 a-helical turn/3.6 residues)(3.1 A/keratin turn) =  142 A   7. The reducing conditions promote cleavage of the disulfide bonds that  cross-link a keratin molecules. This helps the larvae digest the wool  clothing that they eat.   9. Collagens primary structure is its amino acid sequence, which is a  repeating triplet of mostly Gly-Pro-Hyp. Its secondary structure is  the left-handed helical conformation characteristic of its repeating se¬  quence. Its tertiary structure is essentially the same as its secondary  structure, since most of the protein consists of one type of second¬  ary structure. Collagens quaternary structure is the arrangement of  its three chains in a right-handed triple helix.   11. A fibrous protein such as a keratin does not have a discrete globu¬  lar core. Most of the residues in its coiled coil structure are exposed  to the solvent. The exception is the strip of nonpolar side chains at  the interface of the two coils.   13. Peptide c is most likely to form an a helix with its three charged  residues (Lys, Glu, and Arg) aligned on one face of the helix. Peptide  a has adjacent basic residues (Arg and Lys), which would destabilize  a helix. Peptide b contains Gly and Pro, both of which are helix¬  breaking (Table 6 - 1 ).   15. In a protein crystal, the residues at the end of a polypeptide chain  may experience fewer intramolecular contacts and therefore tend to  be less ordered (more mobile in the crystal). If their disorder prevents  them from generating a coherent diffraction pattern, it may be im¬  possible to map their electron density.   17. (a) C 4 and Z) 2 ; (b) C 6 and Z) 3 .   19. (a) Phe. Ala and Phe are both hydrophobic, but Phe is much larger  and might not fit as well in Vais place.   (b) Asp. Replacing a positively charged Lys residue with an oppositely  charged Asp residue is likely to be more disruptive.   (c) Glu. The amide-containing Asn would be a better substitute for  Gin than the acidic Glu.   (d) His. Pro’s constrained geometry is best approximated by Gly,  which lacks a side chain, rather than a residue with a bulkier side  chain such as His.   23. At physiological pH, the positively charged Lys side chains repel each  other. Increasing the pH above their pi6(>10.5) would neutralize the  side chains and allow an a helix to form.   25. The molecular mass of 0 2 is 32 D. Hence the ratio of the masses of  hemoglobin and 4 0 2 , which is equal to the ratio of their volumes,  is 65,000/(4 X 32) = 508. The 70-kg office worker has a volume  of 70 kg X 1 cm 3 /g X (1000 g/kg) X (1 m/100 cm ) 3 = 0.070 m 3 .  Hence the ratio of the volumes of the office and the office worker is  (4 X 4 X 3)/0.070 = 686 . These ratios are similar in magnitude,  which you may not have expected.   27. At elevated temperatures, more protein denaturation is likely, so it is  advantageous for a cell to increase the rate at which it can eliminate  these nonfunctional proteins.   29. The brains of the Alzheimer’s-prone mice were already burdened  with accumulated A|3, so the PrP 3c aggregation augmented the  brain damage, leading to earlier onset of symptoms than in mice  whose brains were not already damaged by A(3 accumulation.  (There is also some evidence that A(3 oligomers may interact di¬  rectly with PrP.)    The difference in Yq 2 values is 0.78 — 0.26 = 0.52. Therefore, in  active muscle cells, myoglobin can transport a significant amount of  0 2 by diffusion from the cell surface to the mitochondria.   5. (a) For hemoglobin, p 50 = 26 torr. Let the Hill coefficient, n,  equal 3.    7. (a) Vitamin O is useless because the body’s capacity to absorb oxy¬  gen is not limited by the amount of oxygen available but by the abil¬  ity of hemoglobin to bind and transport 0 2 . Furthermore, oxygen is  normally introduced into the body via the lungs, so it is unlikely that  the gastrointestinal tract would have an efficient mechanism for ex¬  tracting oxygen.   (b) The fact that oxygen delivery in vertebrates requires a dedicated  0 2 -binding protein (hemoglobin) indicates that dissolved oxygen by  itself cannot attain the high concentrations required. Moreover, a  few drops of vitamin O would make an insignificant contribution  to the amount of oxygen already present in a much larger volume  of blood.   11. (a) Hyperventilation eliminates C0 2 , but it does not significantly af¬  fect the 0 2 concentration, since the hemoglobin in arterial blood is  already essentially saturated with oxygen.   (b) The removal of C0 2 also removes protons, according to the  reaction   H + + HCO^ H 2 0 + CO 2  The resulting increase in blood pH would increase the 0 2 affinity of  hemoglobin through the Bohr effect. The net result would be that  less oxygen could be delivered to the tissues until the C0 2 balance  was restored. Thus, hyperventilation has the opposite of the intended  effect (note that since hyperventilation suppresses the urge to breathe,  doing so may cause the diver to lose consciousness due to lack of 0 2  and hence drown).   13. (a) Lower; (b) higher. The Asp 99(3His mutation of hemoglobin  Yakima disrupts a hydrogen bond at the ot 1 -|3 2 interface of the T  state (Fig. 7-9a), causing the T R equilibrium to shift toward R  state (lower p 50 ). The Asn 102(3 —» Thr of hemoglobin Kansas causes  the opposite shift in the T R equilibrium by abolishing an R-state  hydrogen bond (Fig. 7-9(7).   13. The increased BPG helps the remaining erythrocytes deliver 0 2 to  tissues. However, BPG stabilizes the T conformation of hemoglobin,  so it promotes sickling and therefore aggravates the disease.   17. Myosin is both fibrous and globular. Its two heads are globular, with  several layers of secondary structure. Its tail, however, consists of a  lengthy, fibrous coiled coil.   19. Because many myosin heads bind along a thin filament where it over¬  laps a thick filament, and because the myosin molecules do not exe¬  cute their power strokes simultaneously, the thick and thin filaments  can move past each other by more than 100 A in the interval be¬  tween power strokes of an individual myosin molecule.   21 . Micro filaments consist entirely of actin subunits that are assembled  in a head-to-tail fashion so that the polarity of the subunits is pre¬  served in the fully assembled fiber. In keratin filaments, however,  successive heterodimers align in an antiparallel fashion, so that in  a fully assembled intermediate filament, half the molecules are ori¬  ented in one direction and half are oriented in the opposite direc¬  tion (Fig. 6-16).    Solutions to Odd Problems SP-5  23. (a) 130-200 kD; (b) 150-200 kD; (c) -23 kD and 53-75 kD  25. The loops are on the surface of the domain, so they can tolerate more  amino acid substitutions. Amino acid changes in the (3 sheets would  be more likely to destabilize the domain.   27. The antigenic site in the native protein usually consists of several pep¬  tide segments that are no longer contiguous when the tertiary struc¬  ture of the protein is disrupted.   29. Cleaving the IgA molecules into separate Fab fragments destroys their  ability to cross-link antigens. Although the Fab fragments can still  bind to the bacteria, the bacteria will not become trapped in a cross-  linked network and can still initiate an infection.   11. (a) a-D-glucose-(l —» l)-a-D-glucose or a-D-glucose-(l 1)-(3-d-  glucose. (b) The numerous hydrogen-bonding —OH groups of the  disaccharide act as substitutes for water molecules. Because trehalose  is a nonreducing sugar, it is unlikely to participate in oxidation-  reduction reactions with other biomolecules when it is present at high  concentrations.   19. Glucosamine is a building block of certain glycosaminoglycan com¬  ponents of proteoglycans (Fig. 8-12). Boosting the body’s supply of  glucosamine might slow the progression of the disease osteoarthritis,  which is characterized by the degradation of proteoglycan-rich artic¬  ular (relating to a joint) cartilage.   23. The cationic Ca 2+ ions shield the negative charges of the gly-  cosaminoglycans in the proteoglycans, so that they can be stored in  a relatively small volume inside the cell. When the Ca 2+ ions are  pumped out, the repulsion between the glycosaminoglycan chains  causes them to expand in the extracellular space.   1 . trans-O leic acid has a higher melting point because, in the solid state,  its hydrocarbon chains pack together more tightly than those of cis-  oleic acid.    H 3 C(CH 2 ) 7 CH= CH(CH 2 ) 7 —c—o—ch    5. Of the 4X4= 16 pairs of fatty acid residues at Cl and C3, only  10 are unique because a molecule with different substituents at Cl  and C 3 is identical to the molecule with the reverse substitution or¬  der. However, C 2 may have any of the four substituents for a total  of 4 X 10 = 40 different triacylglycerols.    (b) To convert sphingomyelin to sphingosine- 1 -phosphate, a portion  of the head group (typically choline or ethanolamine) must be hy¬  drolytically cleaved, leaving a phosphate group, and the amide link¬  age to the fatty acyl group must be hydrolyzed.   11. Both DNA and phospholipids have exposed phosphate groups that  are recognized by the antibodies.   13. Eicosanoids synthesized from arachidonic acid are necessary for in¬  tercellular communication. Cultured cells do not need such commu¬  nication and therefore do not require linoleic acid.   15. Triacylglycerols lack polar head groups, so they do not orient them¬  selves in a bilayer with their acyl chains inward and their glycerol  moiety toward the surface.   17. (a) Saturated; (b) long-chain. By increasing the proportion of satu¬  rated and long-chain fatty acids, which have higher melting points,  the bacteria can maintain constant membrane fluidity at the higher  temperature.   (c) The additional residues form a helix, which partially satisfies back¬  bone hydrogen bonding requirements, where the lipid head groups  do not offer hydrogen bonding partners.   23. The mutant signal peptidase would cleave many preproteins within  their signal peptides, which often contain Leu-Leu sequences. This  would not affect translocation into the ER, since signal peptidase  acts after the signal peptide enters the ER lumen. Proteins lacking  the Leu-Leu sequence would retain their signal peptides. These  proteins, and those with abnormally cleaved signal sequences,  would be more likely to fold abnormally and therefore function  abnormally.    = -1270 J-mol 1 = -1.27 kj- mol 1  3. Use Equation 10-3 and let Z = 2 and T = 310 K:   = (8.314 J • K -1 • mol -1 )(310 K) ln(10 - 7 )/(10 -3 )  + (2)(96,485 J • V -1 • mol _1 )( —0.050 V)    The negative value of AC indicates a thermodynamically favor¬  able process.   = (8.314 J • K -1 • mol -1 )(310 K) ln(10“ 7 )/(10“ 3 )   + (2)(96,485 J • V -1 • mo r 1 )( + 0.150 V)    = —23,700 J* mol 1 + 28,900 J* mol 1  = +5,200 J • mol 1 = +5.2kJ*mol 1  The positive value of A G indicates a thermodynamically unfavorable  process.    7. K + transport ceases because the ionophore-K + complex cannot dif¬  fuse through the membrane when the lipids are immobilized in a gel¬  like state.   9. An eight-stranded (3 barrel has a solid core. A P barrel with 16 or  18 strands has a diameter large enough to accommodate a pore for  solute transport.   11 . The presence of a series of K + ions within the channel prevents  water molecules from forming a hydrogen-bonded chain.   (b) Miltefosine is amphipathic and therefore cannot cross the parasite  cell membrane by diffusion. Since it is not a normal cell component,  it probably does not have a dedicated active transporter. It most likely  enters the cell via a passive transport protein.   (c) This amphipathic molecule most likely accumulates in mem¬  branes, with its hydrophobic tail buried in the bilayer and its polar  head group exposed to the solvent.   (d) The protein recognizes the phosphocholine head group, which  also occurs in some sphingolipids and some glycerophospholipids.  Since the protein does not bind all phospholipids or triacylglycerols,  it does not recognize the hydrocarbon tail.   13. (a) The data do not indicate the involvement of a transport protein,  since the rate of transport does not approach a maximum as [X]  increases.   (b) To verify that a transport protein is involved, increase [X] to  demonstrate saturation of the transporter at high [X], or add a struc¬  tural analog of X to compete with X for binding to the transporter,  resulting in a lower flux of X.   17. A channel provides an open pore across the membrane, whereas a  pump operates by changing its conformation in an ATP-dependent  manner. The additional time required for ATP hydrolysis and pro¬  tein conformation changes causes ion movement through a pump to  be slower than through a channel.   19. (a) pH = —log[H + ] = — log(0.15) = 0.82   The pH of the secreted HC1 is over 6 pH units lower than the  cytosolic pH, which corresponds to a [H + ] of ~4 X 10 8 M.   (b) C0 2 + H 2 0 HCO 3 + H +    7. At 23°C, every 10-fold increase in rate corresponds to a decrease of  about 5.7 kj • mol 1 in AG 7 *. For the nuclease, with a rate enhance¬  ment on the order of 10 14 , AG* is lowered about 14 X 5.7 kj • mol l ,  or about 80 kj • mol \ Alternatively, since the rate enhancement,  k, is given by k = ^ AAGcat/i?r ,    21 . In the absence of ATP, Na + extrusion by the (Na + -K + )-ATPase  would cease, so no glucose could enter the cell by the Na + -glucose  symport. The glucose in the cell would then exit via the passive-  mediated glucose transporter, and the cellular [glucose] would decrease  until it matched the extracellular [glucose] (of course, the cell would  probably osmotically burst before this could occur).   5. There are three transition states (X*) and two intermediates (I). The  reaction is not thermodynamically favorable because the free energy  of the products is greater than that of the reactants.   11 . As the temperature increases, thermal energy boosts the proportion  of reactants that can achieve the transition state per unit time, so the  rate increases. Above an optimal temperature, the enzyme becomes  denatured and rapidly loses catalytic activity (recall that proteins are  typically only marginally stable; Section 6-4).   13. Glu has a piTof ~4 and, in its ionized form, acts as a base catalyst. Lys  has a piTof ~10 and, in its protonated form, acts as an acid catalyst.   15. Inhibition by various metal ions suggests that urease requires a metal  ion for catalysis, whose replacement by Hg, Co, or Cd inactivates the  enzyme. However, the inhibitory metal ions could also disrupt the  enzyme’s structure by binding somewhere other than the active site,  so this inhibitory effect does not prove that urease acts via metal ion  catalysis. (In fact, urease activity requires two catalytic Ni ions.)   17. The preferential binding of the transition state to an enzyme is an  important (often the most important) part of an enzyme’s catalytic  mechanism. Hence, the substrate binding site is the catalytic site.   19. The lysozyme active site is arranged to cleave oligosaccharides be¬  tween the fourth and fifth residues. Moreover, since the lysozyme ac¬  tive site can bind at least six monosaccharide units, (NAG ) 6 would  be more tightly bound to the enzyme than (NAG) 4 , and this addi¬  tional binding free energy would be applied to distorting the D ring  to its half-chair conformation, thereby facilitating the reaction.   21 . Asp 101 and Arg 114 form hydrogen bonds with the substrate mol¬  ecule (Fig. 11-19). Ala cannot form these hydrogen bonds, so the  substituted enzyme is less active.    27. Yes. An enzyme decreases the activation energy barrier for both the  forward and the reverse directions of a reaction.   29. If the soybean trypsin inhibitor were not removed from tofu, it would  inhibit the trypsin in the intestine. At best, this would reduce the  nutritional value of the meal by rendering its protein indigestible. It  might very well also lead to intestinal upset.   31. Activated factor IXa leads, via several steps, to the activation of the fi¬  nal coagulation protease, thrombin. The absence of factor IX therefore  slows the production of thrombin, delaying clot formation, and caus¬  ing the bleeding of hemophilia. A1 though activated factor XIa also leads  to thrombin production, factor XI plays no role until it is activated by  thrombin itself. By this point, coagulation is already well underway, so  a deficiency of factor XI does not significantly delay coagulation.   = 2.5 X 10 4 min 1  (b) The reaction has a molecularity of 1.   3. From Eq. 12-7, [A] = [A] 0 e kt . Since q /2 = 0.693/ k, k = 0.693/14 d  = 0.03 d -1 . (a) 7 mmol; (b) 5 pmol; (c) 3.5 pmol; (d) 0.3 pmol.  5. Only a plot of 1/[reactant] versus t gives a straight line, so the reac¬  tion is second order. The slope, k , is 0.15 mM -1 • s -1 .    9. v Q = V max [S\/(K M + [S\)  v Q /V max = [S]/(K m + [S])   0.05[S] = 0.95 K m  [S] = (0.95/0.05 )K m = 19 K m  11. Construct a Lineweaver-Burk plot.   K m = — Hx -intercept = — 1 /(—4 pM -1 ) = 0.25 pM  Knax = 1/y-intercept = 1/(0 .8 mM 1 • s) = 1.25 mM-s 1  13. Velocity measurements can be made using any convenient unit of  change per unit of time. K M is, by definition, a substrate concentra¬  tion (the concentration when v 0 = V max /2), so its value does not re¬  flect how the velocity is measured.    15. Comparing the two data points, since a 100-fold increase in substrate  concentration only produces a 10 -fold increase in reaction velocity,  it appears that when [S] = 100 mM, the velocity is close to V^ax*   Therefore, assume that V max ~ 50 |j,M • s 1 and use the other data  point to estimate K M using the Michaelis-Menten equation:   The true must be greater than the estimated value, so the value  of K m is an underestimate of the true K M .    17. The enzyme concentration is comparable to the lowest substrate  concentration and therefore does not meet the requirement that  [E] « [S]. You could fix this problem by decreasing the amount of  enzyme used for each measurement.   19. (a) TV-Acetyltyrosine ethyl ester, with the lower value of K M , has  greater apparent affinity for chymotrypsin. (b) The value of is  not related to the value of K M , so no conclusion can be drawn.   21. A* will appear if the reaction follows a Ping Pong mechanism, since  a double-displacement reaction can exchange an isotope from P back  to A in the absence of B.    23. By irreversibly reacting with chymotrypsin’s active site, DIPF would  decrease [E] T . The apparent V max would decrease since V msLX =  k cax [E]t* K m would not be affected since the uninhibited enzyme  would bind substrate normally.   25. If an irreversible inhibitor is present, the enzyme solutions activity would  be exactly 100 times lower when the sample is diluted 100-fold. Dilution  would not significantly change the enzymes degree of inhibition.   27. The lines of the double-reciprocal plots intersect to the left of the llv 0  axis (on the 1/[S] axis). Hence, inhibition is mixed (with a = a').    29. (a) Inhibition is most likely mixed (noncompetitive) with a = a'  since it is reversible and only V^ax is affected.   (b) Since = 0.8 V max , 80% of the enzyme remains uninhibited.  Therefore, 20% of the enzyme molecules have bound inhibitor.   (c) As indicated in Table 12-2 for mixed inhibition, = V max /a '.  Thus,   1. Somatostatin is a 14-residue peptide hormone and is not lipid-soluble,  so it would require a cell-surface receptor.   3. (a) Norepinephrine is synthesized by the decarboxylation of Tyr and  by hydroxylation of its |3 carbon and its phenyl group, (b) Epinephrine  is derived from norepinephrine by A^methylation.   5. A plot of fractional saturation (Y) versus [L] yields a hyperbola where  the estimated value of 0.5 Y corresponds to a ligand concentration, or  Af, of about 3.5 |xM.   7. Use Equation 13-8, letting [L] = 1 |xM and [B] = [I 50 ] = 2.5 |xM,   11. The PH domain allows the IRS to be localized to the intracellular  leaflet of the cell membrane, close to the insulin receptor and ready  to participate in signal transduction. The PTB domain, like an SH2  domain, recognizes phospho-Tyr residues, in this case on the au-  tophosphorylated insulin receptor. As a result, the IRS can be acti¬  vated following insulin binding to its receptor. The phospho-Tyr  residues on the IRS itself are recognition points for SH2-containing  proteins, which can thereby become activated.   13. (a) Yes, because the binding of Src’s SH3 domain to the linker that  connects its SH 2 domain to the N-terminal lobe of its PTK domain  is required for Src to maintain its autoinhibited conformation.  Hence, the deletion of this SH3 domain would constitutively acti¬  vate Src, thereby driving the cell with this mutation to a state of un¬  restrained proliferation.   (b) No, because the phosphorylation of Tyr 416 is required for the  activation of Src and hence the Y416F mutation would inhibit the  mutant cell’s proliferation.   13. The SH 2 domain allows the enzyme to bind to phospho-Tyr residues  on its target proteins. Because targets typically contain more than one  phospho-Tyr group, the phosphatase can recognize and bind to one  site on the target protein while dephosphorylating another site on the  same protein.   17. Transformation to the cancerous state results from several genetic  changes in a cell. Thus, a single oncogene supplied to an otherwise nor¬  mal cell will be insufficient to transform it. However, an immortalized  cell already has some of the genetic changes necessary for transforma¬  tion (malignant cells are also immortal). In such cells, the additional  oncogene may be all they require to complete their transformation.   19. Because the GTP analog cannot be hydrolyzed, G a remains active.  Analog binding to G s therefore increases cAMP production. Analog  binding to Gj decreases cAMP production.   21 . Like cholera toxin, B. anthracis EF leads to the overproduction of cAMP,  which triggers the release of fluid from cells. The result is edema.   23. When G sa catalyzes the hydrolysis of its bound GTP to GDP + P z ,  the Arg side chain that cholera toxin ADP-ribosylates functions to  stabilize the transition states developing negative charge. The ADP-  ribosylated G sa therefore hydrolyzes its bound GTP at a greatly re¬  duced rate and hence remains activated far longer than normal G sa .  Mutating this Arg will have a similar effect. Consequently, in cells in  which a G sa • GTP functions to induce cell proliferation, mutating  this Arg will drive the cell into a state of unrestrained proliferation,  a requirement for the malignant transformation of the cell. The gene  encoding such a G sa subunit is a proto-oncogene and the mutation  of its Arg residue converts it to an oncogene.   23. No. Although the diacylglycerol second messengers are identical,  phosphatidylethanolamine does not generate an IP 3 second messen¬  ger that triggers the release of Ca 2+ , which in turn alters protein ki¬  nase C activity.   29. Li + blocks the conversion of phosphorylated inositol species to inositol,  thereby preventing the recycling of IP 3 and its degradation products back  to inositol. This in turn prevents the synthesis of phosphatidylinositol  and PIP 2 , the precursor of the IP 3 second messenger.   1. A heterotroph relies on other organisms for food, which may include  substances the heterotroph cannot synthesize (including vitamins).  An autotroph can produce all the molecules it needs.   3. Arsenic, which resembles phosphorus, is incorporated into nucleic  acids and other compounds that ordinarily contain phosphate groups.  5. C, D, A, E, B  7. (a) K= e ~^ G °'' RT   (b) The large change in free energy makes the citrate synthase reac¬  tion irreversible, so it could (and does; Section 17-4B) serve as a con¬  trol point for the citric acid cycle.    The theoretical maximum yield of ATP is equivalent to (AG 0 ' for  fuel oxidation)/(AG°' for ATP synthesis) = ( — 2850 kj • mol -1 )/  (—30.5 kj • mol x ) ~ 93 ATP    At pH 6 , the phosphate groups are more ionized than they are at pH  5 , which increases their electrostatic repulsion and therefore increases  the magnitude of AG for hydrolysis (makes it more negative).   Calculating AG for the reaction ATP + creatine v phosphocre-  atine + ADP, using Eq. 14-1:   = 12.6 kj- mol -1 + (8.3145 J • K -1 • mol -1 )(298 K)  ' (2.5 mM)(0.15 m M) N   = 12.6 kj • mol 1 — 5.9 kj • mol 1 = 6.7 kj • mol    Since AG > 0 , the reaction will proceed in the opposite direction as  written above, that is, in the direction of ATP synthesis.   The more positive the reduction potential, the greater the oxidizing  power. From Table 14-4,    The ubiquinone half-reaction has a higher reduction potential (0.045  V) than the NAD + half-reaction (—0.315 V). Therefore, electrons  will flow from NADH (which becomes oxidized) to ubiquinone  (which becomes reduced).   Because A^°' > 0, AG°' < 0 and the reaction will spontaneously  proceed as written.   2 cyto c (Fe 3+ ) + ubiquinol —> 2 cyto c (Fe 2+ ) + ubiquinone + 2 H +  Using the data in Table 14-4,   Probably not. Although all cells carry out a similar set of basic meta¬  bolic reactions, the enzymes that catalyze the reactions have different  amino acid sequences and hence different gene sequences. cDNAs  produced from mammalian mRNAs would be unlikely to hybridize  with bacterial DNA segments.   1. (a) Reactions 1, 3, 7, and 10; (b) Reactions 2 , 5, and 8 ; (c) Reaction  6 ; (d) Reaction 9; (e) Reaction 4.    Cl of DHAP and Cl of GAP are achiral but become chiral in FBP  (as C3 and C4). There are four stereoisomeric products that differ in  configuration at C3 and C4: fructose- 1 , 6 -bisphosphate, psicose- 1 , 6 -  bisphosphate, tagatose-l, 6 -bisphosphate, and sorbose- 1 , 6 -bisphosphate  (see Fig. 8 - 2 ).   (a) Glucose + 2 NAD + + 2 ADP + 2 P,   2 pyruvate + 2 NADH + 2 ATP + 2 H 2 0   2 pyruvate + 2 NADH + 2 ADP—AsO^ - + 2 H 2 0  2 ADP — AsOg - + 2 H 2 Q —> 2 ADP + AsOj~_   (c) Arsenate is a poison because it uncouples ATP generation from  glycolysis. Consequently, glycolytic energy generation cannot occur.  The reaction intermediate is glucose- 1 , 6 -bisphosphate (G 1 , 6 P).  Alcoholic fermentation, unlike homolactic fermentation, includes a  step (the pyruvate decarboxylase reaction) in which a carbon is lost  as C0 2 . Because the C0 2 diffuses (bubbles) away, the reaction can¬  not proceed in reverse.   AG values differ from AG°' values because AG = AG°' + RT  In [products]/[reactants] and cellular reactants and products are not in  their standard states.   When [GAP] = 10 “ 4 M, [DHAP] = 5.5 X 10 “ 4 M. According to  Eq. 1-17,   [GAP] [DHAP] _ —(22,800 J* mol -l )/(8.3145 J* K -1 • mol -1 X310 K)  [FBP]   [FBP]/[GAP] = (3.8 X 10“ 4 M)/(1(T 4 M) = 3.8   Pyruvate kinase regulation is important for controlling the flux of  metabolites, such as fructose (in liver), which enter glycolysis after  the PFK step.   Galactokinase, which catalyzes a highly exergonic reaction, is a possible  control point. Because galactose enters the glycolytic pathway as glucose-  6 -phosphate, the PFK reaction is also likely to be a major control point,  (a) Glycerol can be converted to the glycolytic intermediate DHAP  by the activity of glycerol kinase and glycerol phosphate dehydroge¬  nase (Fig. 15-27). (b) One ATP is consumed by the glycerol kinase  reaction, but two ATP are produced (by the PGK and PK reactions),  for a net yield of one ATP per glycerol (this does not count the ATP  that might be generated through oxidative phosphorylation from the  NADH produced in the glycerol phosphate dehydrogenase reaction).  The three glucose molecules that proceed through glycolysis yield 6  ATP. The bypass through the pentose phosphate pathway results in  a yield of 5 ATP.   25. Transketolase transfers 2 -carbon units from a ketose to an aldose, so  the products are a 3 -carbon sugar and a 7 -carbon sugar.   27. Even when the flux of glucose through glycolysis and hence the cit¬  ric acid cycle is blocked, glucose can be oxidized by the pentose phos¬  phate pathway, with the generation of C0 2 .   29. (a) Reaction 8 , (b) reaction 5, (c) reaction 1, (d) reaction 2,  (e) reaction 3 .   1. Glycogen is broken down when the cell needs to catabolize glucose  to produce ATP. The G1P generated by the glycogen phosphorylase  reaction is quickly isomerized to G 6 P and enters glycolysis. The con¬  tinual consumption of G 1 P “pulls” the phosphorylase reaction for¬  ward, making it thermodynamically favorable.   3. Phosphoglucokinase activity generates G1,6P, which is necessary to  “prime” phosphoglucomutase that has become dephosphorylated  and thereby inactivated through the loss of its G 1 , 6 P reaction  intermediate.   5. The conversion of circulating glucose to lactate in the muscle gen¬  erates 2 ATP. If muscle glycogen could be mobilized, the energy  yield would be 3 ATP, since phosphorolysis of glycogen bypasses  the hexokinase-catalyzed step that consumes ATP in the first stage  of glycolysis.   7. In the course of glucose catabolism, a detour through glycogen syn¬  thesis and glycogen breakdown begins and ends with G 6 P. The en¬  ergy cost of this detour is 1 ATP equivalent, consumed in the  UDP-glucose pyrophosphorylase step. The overall energy lost is  therefore 1/32 or ~ 3 %.   9. A glycogen molecule with 28 tiers would represent the most efficient  arrangement for storing glucose, and its outermost tier would con¬  tain considerably more glucose residues than a glycogen molecule  with only 12 tiers. However, densely packed glucose residues would  be inaccessible to phosphorylase. In fact, such a dense glycogen mol¬  ecule could not be synthesized because glycogen synthase and branch¬  ing enzyme would have no room to operate (see Box 16-3 for a  discussion of glycogen structure).   11. (a) Circulating [glucose] is high because cells do not respond to the  insulin signal to take up glucose.   (b) Insulin is unable to activate phosphoprotein phosphatase-1 in  muscle, so glycogen synthesis is not stimulated. Moreover, glycogen  synthesis is much reduced by the lack of available glucose in the cell.   2 pyruvate + 2 NADH + 4 H + +2 ATP + 2 H 2 0  The equation for gluconeogenesis is   2 Pyruvate + 2 NADH + 4 H + +4 ATP + 2 GTP + 6 H 2 0   2 ATP + 2 GTP + 4 H 2 0 —> 2 ADP + 2 GDP + 4 P,   17. (a) Aspartate can be transaminated to produce oxaloacetate, a gluco¬  neogenic precursor, (b) To convert 2 aspartate to glucose, 2 GTP are  consumed in the PEPCK reaction and 2 ATP are consumed in the  phosphoglycerate kinase reaction, for a total of 4 ATP equivalents.   19. (a) At the beginning of a fast, blood glucose levels are normal, be¬  cause dietary sources or glycogenolysis can supply glucose, (b) After  a fast, the blood glucose levels are very low, because dietary glucose  and glycogen have been depleted and gluconeogenesis is impaired due  to the fructose- 1 , 6 -bisphosphatase deficiency.   21. A high level of AMP results from a high rate of ATP consumption  in the cell, so it would act to promote flux through ATP-generating  pathways such as glycolysis. Therefore, AMP would be expected  to inhibit the activity of the gluconeogenic enzyme fructose-1,  6-bisphosphatase.   1. The labeled carbon becomes C4 of the succinyl moiety of succinyl-  CoA. Because succinate is symmetrical, the label appears at Cl and  C4 of succinate. When the resulting oxaloacetate begins the sec¬  ond round, the labeled carbons appear as l4 C0 2 in the isocitrate  dehydrogenase and the a-ketoglutarate dehydrogenase reactions  (see Fig. 17-2).   3. In mammals, pyruvate can be converted to lactate (reduction), to ala¬  nine (transamination), to acetyl-CoA (oxidative decarboxylation), and  to oxaloacetate (carboxylation). In yeast, pyruvate is also converted  to acetaldehyde (decarboxylation).   3. The decarboxylation step is most likely to be metabolically irre¬  versible since the C0 2 product is rapidly hydrated to bicarbonate.  The reverse reaction, a carboxylation, requires the input of free en¬  ergy to become favorable (Section 16-4A). The other four reactions  are transfer reactions or oxidation-reduction reactions (transfer of  electrons) that are more easily reversed.   11. NAD + (%°' = —0.313 V) does not have a high enough reduction  potential to support oxidation of succinate to fumarate i%°' =  + 0.031 V); that is, the succinate dehydrogenase reaction has insuf¬  ficient free energy to reduce NAD + . Enzyme-bound FAD is   13. Competitive inhibition can be overcome by adding more substrate,  in this case succinate. Oxaloacetate overcomes malonate inhibition  because it is converted to succinate by the reactions of the citric  acid cycle.   15. The A G°' value is the sum of the A G°' values for the malate dehy¬  drogenase reaction (29. 7 kj • mol -1 ) and the citrate synthase reac¬  tion ( — 31. 5 kj • mol x ): —1.8 kj • mol \    17. From Table 17-2, for the succinate dehydrogenase reaction, A G°' =  6 kj • mol -1 and A G = ~0.   _ -(6000 J • mol -1 )/(8.3145 J • K 1 • mol -1 )(310 K)   19. The phosphofructokinase reaction is the major flux-control point for  glycolysis. Inhibiting phosphofructokinase slows the entire pathway,  so the production of acetyl-CoA by glycolysis followed by the pyru¬  vate dehydrogenase complex can be decreased when the citric acid  cycle is operating at maximum capacity and the citrate concentration  is high. As citric acid cycle intermediates are consumed in synthetic  pathways, the citrate concentration drops, relieving phosphofructo¬  kinase inhibition and allowing glycolysis to proceed in order to replen¬  ish the citric acid cycle intermediates.   21. (a) Because citric acid cycle intermediates such as citrate and succinyl-  CoA are precursors for the biosynthesis of other compounds, anaer¬  obes must be able to synthesize them.   (b) These organisms do not need a complete citric acid cycle, which  would yield reduced coenzymes that must be reoxidized.   25. To synthesize citrate, pyruvate must be converted to oxaloacetate by  pyruvate carboxylase:   Pyruvate + C0 2 + ATP + H 2 0 —» oxaloacetate + ADP + P z  A second pyruvate is converted to acetyl-CoA by pyruvate dehydro¬  genase:   Pyruvate + CoASH + NAD + —> acetyl-CoA + C0 2 + NADH  The acetyl-CoA then combines with oxaloacetate to produce  citrate:   Oxaloacetate + acetyl-CoA + H 2 0 —> citrate + CoASH + H +  The net reaction is   2 Pyruvate + ATP + NAD + + 2 H 2 0 ->   citrate + ADP + P z + NADH + H +  27. Oxaloacetate + FADH 2 + NADH + H + —»   1. Mitochondria with more cristae have more surface area and therefore  more proteins for electron transport and oxidative phosphorylation.  Tissues with a high demand for ATP synthesis (such as heart) con¬  tain mitochondria with more cristae than tissues with lower demand  for oxidative phosphorylation (such as liver).   3. When NADH participates in the glycerophosphate shuttle, the elec¬  trons of NADH flow to FAD and then to CoQ, bypassing Complex  I. Thus, about 1.5 ATP are synthesized per NADH.   \ 0 2 + 2 H + + 2 H 2 0 %°' = 0.815 V   Since the 0 2 /H 2 0 half-reaction has the more positive A^°', the FAD  half-reaction is reversed and the overall reaction is  \ 0 2 + FADH 2 H 2 0 + FAD  \%°'= 0.815 V - (-0.219 V) = 1.034 V  Since l±G°'= -n& A^°',   A G°'= -(2)(96,485 J • V -1 • mol -1 )(1.034 V) = -200 kj • mol -1   The maximum number of ATPs that could be synthesized under stan¬  dard conditions is therefore 200 kj • mol V30.5 kj • mol 1 = 6.6  mol ATP/mol FADH 2 oxidized by 0 2 .   7. NO 3 is the electron acceptor and NADH is the electron donor. Their  standard reduction potentials are listed in Table 14-4.   = (0.42 V) - (-0.315 V) = 0.735 V  A G = -n®\%   = —142,000 J • mol 1 = —142 kj • mol 1  Since the standard free energy change for ATP synthesis is 30.5  kj • mol l , approximately 142/30.5 = 4.6 ATP could be synthesized.   (a) 0 2 consumption ceases because amytal blocks electron transport  in Complex I.   (b) Electrons from succinate bypass the amytal block by entering the  electron-transport chain at Complex II and thereby restore electron  transport through Complexes III and IV.   11. % may differ from %°\ depending on the redox center’s microenvi¬  ronment and the concentrations of reactants and products. In addi¬  tion, the tight coupling between successive electron transfers within  a complex may “pull” electrons so that the overall process is sponta¬  neous.   15. For the transport of a proton from outside to inside (Eq. 18-1),   AC = 2.3 RT[pU {side 1) - pH {side 2)] + Z®. A'P  The difference in pH is —1.4. Since an ion is transported from the  positive to the negative side of the membrane, A'T is negative.   A G= (2.3)(8.314 J • KC 1 • mor 1 )(298 K)( -1.4)   A G= -7980 J-mol -1 - 5790 J-mol -1 = -13.8 kj • mol -1  17. The import of ADP (net charge —3) and the export of ATP (net  charge —4) represents a loss of negative charge from inside the mi¬  tochondrion. This decreases the difference in electrical charge across  the membrane, since the outside is positive due to the translocation  of protons during electron transport. Consequently, the electrochem¬  ical gradient is diminished by the activity of the ADP-ATP translo¬  cator. The activity of the P z -H symport protein diminishes the  proton gradient by allowing protons from the intermembrane space  to re-enter the matrix.   19. In an ATP synthase with more c subunits, more proton translocation  events are required to drive one complete rotation of the r-ring.  Consequently, more substrate oxidation (0 2 consumption) is re¬  quired to synthesize three ATP (the yield of one cycle of the rotary  engine), and the P/O ratio is lower.   21. The protonation and subsequent deprotonation of Asp 61 of the  F 1 F 0 -ATPase’s c subunits induces the rotation of the c-ring, which in  turn, mechanically drives the synthesis of ATP. DCCD reacts with  Asp 61 in a manner that prevents it from binding a proton and  thereby prevents the synthesis of ATP.   23. DNP and related compounds dissipate the proton gradient required  for ATP synthesis. The dissipation of the gradient decreases the rate  of synthesis of ATP, decreasing the ATP mass action ratio. Decreasing  this ratio relieves the inhibition of the electron transport chain, caus¬  ing an increase in metabolic rate.   25. The switch to aerobic metabolism allows ATP to be produced by ox¬  idative phosphorylation. The phosphorylation of ADP increases the  [ATP]/[ADP] ratio, which then increases the [NADH]/[NAD + ] ra¬  tio because a high ATP mass action ratio slows electron transport.  The increases in [ATP] and [NADH] inhibit their target enzymes in  glycolysis and the citric acid cycle (Fig. 18-30) and thereby slow these  processes.   27. The dead algae are a source of food for aerobic microorganisms lower  in the water column. As the growth of these organisms increases, the  rate of respiration and 0 2 consumption increase to the point where  the concentration of 0 2 in the water becomes too low to sustain  larger aerobic organisms.   29. Glucose is shunted through the pentose phosphate pathway to pro¬  vide NADPH, whose electrons are required to reduce 0 2 to 0 2 *.   1. 2 H 2 0 + 2 NADP + -> 2 NADPH + 2 H + + 0 2   3. The color of the seawater indicates that the photosynthetic pigments  of the algae absorb colors of visible light other than red.   5. The energy per photon is E = hc /\, so the energy per mole of  photons (X = 700 nm) is  E= NhdX   = (6.022 X 10 23 mol _1 )(6.626 X 10“ 34 J-s)   (2.998 X 10 8 m • s 1 )/(7 X 10~ 7 m)   7. The order of action is water-plastoquinone oxidoreductase  (Photosystem II), plastoquinone-plastocyanin oxidoreductase (cy¬  tochrome and plastocyanin-ferredoxin oxidoreductase   9. Both systems mediate cyclic electron flows. The photooxidized bac¬  terial reaction center passes electrons through a series of electron car¬  riers so that electrons return to the reaction center (e.g., P960 + ) and  restore it to its original state. During cyclic electron flow in PSI, elec¬  trons from photooxidized P700 are transferred to cytochrome b 6 f  and, via plastoquinone and plastocyanin, back to P700 + . In both  cases, there is no net change in the redox state of the reaction cen¬  ter, but the light-driven electron movements are accompanied by the  transmembrane movement of protons.   13. One mole of photons of red light (X = 700 nm) has an energy of  171 kj. Therefore, 438/171 = 2.6 moles of photons are theoretically  required to drive the oxidation of H 2 0 by NADP to form one mole  of 0 2 .   13. The buildup of the proton gradient is indicative of a high level of  activity of the photosystems. A steep gradient could therefore trig¬  ger photoprotective activity to prevent further photooxidation  when the proton-translocating machinery is operating at maximal  capacity.   17. Because the light-dependent reactions (measured as 0 2 produced by  PSII) and the light-independent reactions (measured as C0 2 fixed by  the Calvin cycle) are only indirectly linked via ATP and NADPH,  they may vary. Cyclic electron flow, which increases ATP production  without increasing NADPH production, may increase the amount of  0 2 produced without increasing C0 2 fixation.   19. Because chloroplast cytochrome b G f is functionally and structurally  similar to mitochondrial Complex III, myxothiazol would be ex¬  pected to block electron transport in the chloroplast. As a result, the  Q cycle would not function and no proton gradient would be gen¬  erated. No ATP would be produced and no electrons would reach  NADP + .   23. An uncoupler dissipates the transmembrane proton gradient by pro¬  viding a route for proton translocation other than ATP synthase.  Therefore, chloroplast ATP production would decrease.   25. After the light is turned off, ATP and NADPH levels fall as these  substances are used up in the Calvin cycle without being replaced by  the light reactions. The RuBP level drops because it is consumed by  the RuBP carboxylase reaction (which requires neither ATP nor  NADPH) and its replenishment is blocked by the lack of ATP for  the phosphoribulokinase reaction.   27. The net synthesis of 2 GAP from 6 C0 2 in the initial stage of the  Calvin cycle (Fig. 19-26) consumes 18 ATP and 12 NADPH (equiv¬  alent to 30 ATP). The conversion of 2 GAP to glucose-6-phosphate  (G6P) by gluconeogenesis does not require energy input (Section 16-  4B), nor does the isomerization of G6P to glucose-1-phosphate  (G1P). The activation of G1P to its nucleotide derivative consumes  2 ATP equivalents (Section 16-5), but ADP is released when the glu¬  cose residue is incorporated into starch. These steps represent an over¬  all energy investment of 18 + 30 + 1 = 49 ATP.   Starch breakdown by phosphorolysis yields G1P, whose subse¬  quent degradation by glycolysis yields 3 ATP, 2 NADH (equivalent  to 5 ATP), and 2 pyruvate. Complete oxidation of 2 pyruvate to 6  C0 2 by the pyruvate dehydrogenase reaction and the citric acid cy¬  cle (Section 17-1) yields 8 NADH (equivalent to 20 ATP), 2 FADH 2  (equivalent to 3 ATP), and 2 GTP (equivalent to 2 ATP). The over¬  all ATP yield is therefore 3 + 5 + 20 + 3 + 2 = 33 ATP.   The ratio of energy spent to energy recovered is 49/33 = 1.5.   29. The carbonic anhydrase catalyzes the conversion of bicarbonate to  C0 2 , which is the substrate for RuBP carboxylase.   31. An increase in [0 2 ] increases the oxygenase activity of RuBP carboxylase-  oxygenase and therefore lowers the efficiency of C0 2 fixation.   33. The increased availability of the substrate C0 2 would increase the  rate of photosynthesis. Because C4 plants spend relatively more energy  to acquire C0 2 for the Calvin cycle, C 3 plants might have the  advantage when C0 2 is more accessible.   3. Lipoprotein B, with a greater proportion of protein, has a higher  density.   5. The products are heptadecane (C 17 H 36 ) and pentadecane (C 15 H 32 ).   7. A defect in carnitine palmitoyl transferase II prevents normal trans¬  port of activated fatty acids into the mitochondria for (3 oxidation.  Tissues such as muscle that use fatty acids as metabolic fuels there¬  fore cannot generate ATP as needed.   9. The first three steps of (3 oxidation resemble the reactions that con¬  vert succinate to oxaloacetate (Sections 17-3F—17-3H).    11. There are not as many usable nutritional calories per gram in un¬  saturated fatty acids as there are in saturated fatty acids. This is be¬  cause oxidation of fatty acids containing double bonds yields fewer  reduced coenzymes whose oxidation drives the synthesis of ATP. In  the oxidation of fatty acids with a double bond at an odd-numbered  carbon, the enoyl-CoA isomerase reaction bypasses the acyl-CoA  dehydrogenase reaction and therefore does not generate FADH 2  (equivalent to 1.5 ATP). A double bond at an even-numbered  carbon must be reduced by NADPH (equivalent to the loss of  2.5 ATP).   13. Oxidation of odd-chain fatty acids generates succinyl-CoA, an inter¬  mediate of the citric acid cycle. Because the citric acid cycle operates  as a multistep catalyst to convert acetyl groups to C0 2 , increasing  the concentration of a cycle intermediate can increase the catalytic  activity of the cycle.   15. (a) Phytanate can be esterified to CoA, but the methyl group at the  (3 position prevents the dehydrogenation catalyzed by hydroxyacyl-  CoA dehydrogenase (reaction 3 of the (3 oxidation pathway).   17. 3-Ketoacyl-CoA transferase is required to convert ketone bodies to  acetyl-CoA. If the liver contained this enzyme, it would be unable to  supply ketone bodies as fuels for other tissues.   19. Palmitate (C 16 ) synthesis requires 14 NADPH. The transport of 8  acetyl-CoA to the cytosol by the tricarboxylate transport system sup¬  plies 8 NADPH (Fig. 20-24), which represents 8/14 X 100 = 37%  of the required NADPH.   21 . The label does not appear in palmitate because l4 C0 2 is released in  Reaction 2b of fatty acid synthesis (Fig. 20-26).   23. The synthesis of stearate (18:0) from mitochondrial acetyl-CoA re¬  quires 9 ATP to transport 9 acetyl-CoA from the mitochondria to  the cytosol. Seven rounds of fatty acid synthesis consume 7 ATP (in  the acetyl-CoA carboxylase reaction) and 14 NADPH (equivalent to  33 ATP). Elongation of palmitate to stearate requires 1 NADH and  1 NADPH (equivalent to 5 ATP). The energy cost is therefore 9 +  7 + 35 + 5 = 56 ATP.   The degradation of stearate to 9 acetyl-CoA consumes 2 ATP (in  the acyl-CoA synthetase reaction) but generates, in eight rounds of  (3 oxidation, 8 FADH 2 (equivalent to 12 ATP) and 8 NADH (equiv¬  alent to 20 ATP). Thus, the energy yield is 12 + 20 — 2 = 30 ATP.  This represents only about half of the energy consumed in synthesiz¬  ing stearate (30 ATP versus 56 ATP).   25. Palmitate biosynthesis consumes 7 ATP and 14 NADPH (equivalent  to 35 ATP). The addition of four more 2-carbon units as acetyl-CoA  in the mitochondrion (Fig. 20-28) consumes 4 NADH (equivalent  to 10 ATP) and 4 NADPH (equivalent to 10 ATP), so that a total  of 62 ATP are consumed.   27. The breakdown of glucose by glycolysis generates the dihydroxyace-  tone phosphate that becomes the glycerol backbone of triacylglycerols  (Fig. 20-29).   29. ACC catalyzes the first committed step of fatty acid synthesis, so  blocking this step might decrease the fatty acids available for storage  as triacylglycerols (fat). The malonyl-CoA produced in the ACC re¬  action inhibits import of fatty acyl-CoA into the mitochondria, so  lowering the level of malonyl-CoA might help promote fatty acid ox¬  idation and reduce fat accumulation.   1. Proteasome-dependent proteolysis requires ATP to activate ubiquitin  in the first step of linking ubiquitin to the target protein (Fig. 21-2)  and for denaturing the protein as it enters the proteasome.   3. The proteasome would facilitate the degradation of intracellular proteins  that have been damaged or denatured by heat or oxidation and would  therefore help the cell eliminate these nonfuctional and possibly toxic  proteins so that they could be replaced by newly synthesized proteins.   5. A glutamate receptor would have helped human ancestors recognize  protein-rich foods, because foods containing significant amounts of  protein also contain relatively large amounts of glutamate, one of the  most abundant amino acids.   7. The urea cycle transforms excess nitrogen from protein breakdown to  an excretable form, urea. In a deficiency of a urea cycle enzyme, the  preceding urea cycle intermediates may build up to a toxic level. A low-  protein diet minimizes the amount of nitrogen that enters the urea cy¬  cle and therefore reduces the concentrations of the toxic intermediates.   9. (a) Three ATP are converted to 2 ADP and AMP + PP Z , for a total  of 4 ATP equivalents.   (b) The fumarate produced in the urea cycle can be converted to malate  and then to pyruvate by malic enzyme, generating NADPH (equiva¬  lent to 2.5 ATP). Conversion of pyruvate to acetyl-CoA generates  NADH (2.5 ATP equivalents), and the oxidation of the acetyl-CoA by  the citric acid cycle yields another 10 ATP, for a total of 15 ATP.   11. (a) Ala, Arg, Asn, Asp, Cys, Gin, Glu, Gly, His, Met, Pro, Ser, and Val   13. The 8-amino group is removed by the addition of a-ketoglutarate  followed by the departure of glutamate (Fig. 21-22, Reactions 1 and 2).  The a-amino group is eliminated when a-aminoadipate undergoes  transamination with a-ketoglutarate (Fig. 21-22, Reaction 4).   15. Since the three reactions converting tiglyl-CoA to acetyl-CoA and  propionyl-CoA are analogous to those of fatty acid oxidation (|3 ox¬  idation; Fig. 20-12), the reactions are  O   17. Tyrosine is derived from the essential amino acid phenylalanine, and  cysteine is derived from the essential amino acid methionine. A diet  lacking sufficient phenylalanine and methionine will lead to short¬  ages of tyrosine and cysteine also.   21 . In the absence of uridylyl-removing enzyme, adenylyltransferase • P n  will be fully uridylylated, since there is no mechanism for removing  the uridylyl groups once they are attached. Uridylylated adenylyl¬  transferase • P n adenylylates glutamine synthetase, which activates it.  Hence, the defective E. coli cells will have a hyperactive glutamine syn¬  thetase and thus a higher than normal glutamine concentration.  Reactions requiring glutamine will therefore be accelerated, thereby de¬  pleting glutamate and the citric acid cycle intermediate a-ketoglutarate.  Consequently, biosynthetic reactions requiring transamination, as well  as energy metabolism, will be suppressed.   23. The pigment coloring skin and hair is melanin, which is synthesized  from tyrosine. When tyrosine is in short supply, as when dietary pro¬  tein is not available, melanin cannot be synthesized in normal  amounts, and the skin and hair become depigmented.   27. The standard nitrogenase reaction, N 2 —> NH 3 , also produces H 2 .  This H 2 is used to reduce CO to C 2 H 6 and C 3 H 8 .   1 . ATP generating pathways such as glycolysis and fatty acid oxidation  require an initial investment of ATP (the hexokinase and phospho-  fructokinase steps of glycolysis and the acyl-CoA synthetase activa¬  tion step that precedes (3 oxidation). This “priming” cannot occur  when ATP has been exhausted.   3. Rapidly growing cancer cells need ATP as well as the raw materi¬  als for synthesizing nucleic acids, proteins, carbohydrates, and  lipids. While providing some ATP, catabolism of glucose can also  supply ribose (via the pentose phosphate pathway) for nucleotide  biosynthesis, pyruvate that can be converted to acetyl-CoA (via the  pyruvate dehydrogenase reaction) for fatty acid synthesis, and ox-  aloacetate (via the pyruvate carboxylase reaction) for amino acid  synthesis.   3. At high altitude, less oxygen is available for aerobic metabolism, so  glycolysis, an anaerobic pathway, would become relatively more im¬  portant in active muscles. An increase in GLUTl would increase the  intracellular glucose concentration, and an increase in PFK would  increase the flux of glucose through the pathway.   7. GLUT 2 has a higher K M than GLUTl so that the rate of glucose  entry into liver cells can vary directly with the concentration of glu¬  cose in the blood. A transporter with a high K M is less likely to be  saturated with its ligand and therefore would not limit the rate of  transport.   9. The portal vein delivers NHj-rich blood from the intestine directly  to the liver, which can convert it to urea (only the liver carries out  the urea cycle; Section 21-3). The remaining NH 4 is carried through  the circulation to other tissues, where glutamine synthetase converts  glutamate to glutamine (Section 21-5A).   11. Insulin promotes the uptake of glucose via the increase in GLUT4  receptors on the adipocyte surface. A source of glucose is necessary  to supply the glycerol- 3 -phosphate backbone of triacylglycerols.   13. Because fatty acids, like glucose, are metabolic fuels, it makes meta¬  bolic sense for them to stimulate insulin release, which is a signal of  abundant fuel.    15. Insulin activates ATP-citrate lyase, which is the enzyme that converts  citrate to oxaloacetate and acetyl-CoA (Section 20-4A). The activity  of this enzyme is essential for making acetyl units available for fatty  acid biosynthesis in the cytosol. The acetyl units, generated from  pyruvate in the mitochondria, combine with oxaloacetate to form cit¬  rate, which can then be transported from the mitochondria to the  cytosol for reconversion to acetyl-CoA.   17. Adipose tissue synthesizes and releases the polypeptide hormones  adiponectin, leptin, and resistin.   19. Since PYY 3 _ 36 is a peptide hormone, it would be digested if taken  orally. Introducing it directly into the bloodstream avoids degra¬  dation.   21. Ingesting glucose while in the resting state causes the pancreas to re¬  lease insulin. This stimulates the liver, muscle, and adipose tissue to  synthesize glycogen, fat, and protein from the excess nutrients while  inhibiting the breakdown of these metabolic fuels. Hence, ingesting  glucose before a race will gear the runners metabolism for resting  rather than for running.   23. Type I diabetics lack (3 cells that produce insulin, so providing the  hormone is an effective treatment for the disorder. In type II dia¬  betes, cells do not respond efficiently to insulin. Increasing the avail¬  ability of the hormone may boost its signaling activity in some  patients, but in the majority of type II diabetics, insulin levels are al¬  ready elevated and further increases are ineffective.   25. PFK-2 catalyzes the production of fructose-2, 6 -bisphosphate, so in¬  creasing PFK-2 activity would increase the concentration of this ac¬  tivator of phosphofructokinase. The effect would be increased flux of  glucose through glycolysis, which would help lower the concentra¬  tion of glucose in the blood.   1. Following aspartate addition to IMP, adenylosuccinate lyase removes  fumarate, leaving an amino group. In the urea cycle, following the  addition of aspartate to citrulline, argininosuccinase removes fu¬  marate, leaving an amino group.   3. Amidophosphoribosyl transferase (step 2 of IMP synthesis), FGAM  synthetase (step 5 of IMP synthesis), GMP synthetase (GMP synthe¬  sis), carbamoyl phosphate synthetase II (step 1 of UMP synthesis),  and CTP synthetase (CTP synthesis).   7. UTP functions as a feedback inhibitor of its own synthesis, to pre¬  vent the cell from synthesizing too many pyrimidine nucleotides.  ATP activates pyrimidine nucleotide synthesis so that when ATP con¬  centrations are high, the production of other nucleotides will increase  to match it.   9. Hydroxyurea destroys the tyrosyl radical that is essential for the ac¬  tivity of ribonucleotide reductase. Tumor cells are generally fast¬  growing and cannot survive without this enzyme, which supplies  dNTPs for nucleic acid synthesis. In contrast, most normal cells  grow slowly, if at all, and hence have less need for nucleic acid syn¬  thesis.   11 . Threonine is broken down to acetyl-CoA and glycine, either directly  via the serine hydroxymethyltransferase reaction (reaction 5 in  Fig. 21-14) or through the intermediacy of a-amino-(3-ketobutyrate  via the threonine dehydrogenase reaction (reaction 6 of Fig. 21-14)  followed by the a-amino-(3-ketobutyrate lyase reaction (reaction 7 of  Fig. 21-14). The acetyl-CoA can enter the citric acid cycle to pro¬  duce considerable ATP via oxidative phosphorylation. The glycine is  a substrate of the glycine cleavage system, which generates TV 5 ,^ 10 -    methylene-THF (reaction 3 of Fig. 21-14), the methyl-group donor  required for thymidylate synthesis.   13. FdUMP and methotrexate kill rapidly proliferating cells, such as can¬  cer cells and those of hair follicles. Consequently, hair falls out.   13. The synthesis of histidine and methionine requires THF. The cell’s  THF is converted to DHF by the thymidylate synthase reaction, but  in the presence of methotrexate, THF cannot be regenerated.   17. Trimethoprim binds to bacterial dihydro folate reductase but does not  permanently inactivate the enzyme. Therefore, it is not a mechanism-  based inhibitor.   19. In muscles, the purine nucleotide cycle functions to convert aspar¬  tate to fumarate to boost the capacity of the citric acid cycle. If glu¬  tamate dehydrogenase activity were high, then it would combine the  NHJ produced by the purine nucleotide cycle with a-ketoglutarate  to yield glutamate, a reaction that depletes a citric acid cycle inter¬  mediate.   21 . Fumarate is converted to malate by fumarase; malic enzyme decar-  boxylates malate to produce pyruvate; pyruvate is converted to acetyl-  CoA and C0 2 by pyruvate dehydrogenase; and the citric acid cycle  oxidizes the acetyl group to 2 C0 2 .   23. Uracil and thymine accumulate in the urine because they cannot be  further degraded in the absence of the dihydropyrimidine dehydro¬  genase (Fig. 23-24).    Solutions to Odd Problems SP-17  3. Since amino acids have an average molecular mass of ~ 110 D, the  50-kD protein contains 30,000 D 110 D/residue = —455  residues. These residues are encoded by 455 X 3 = 1365 nucleotides.  In B-DNA, the rise per base pair is 3.4 A, so the contour length of  1365 bp is 3.4 A/bp X 1365 bp = 4641 A, or 0.46 pan.   The helix diameter is smaller in this alternate arrangement  (Hoogsteen base pairing).   1. Hypoxanthine pairs with cytosine in much the same way as does  guanine.   Assuming all the DNAs in Figure 24-8 contain the same number of  base pairs, the most supercoiled structure would move farthest dur¬  ing electrophoresis, because its compact structure allows it to move  fastest through the agarose matrix.   L = T + W. For the constrained DNA circle, W = 0 so that  L = T = 207. For the unconstrained DNA circle, L = 207 since  this quantity is invariant, T = 2310 bp/(10.5 bp/turn) = 220,  and W = L — T = 207 — 220 = —13. For the constrained DNA  circle, a = W/ T = 0/207 = 0. For the unconstrained DNA circle,  cr = —13/220 = —0.059 (a value that is typical of naturally occur¬  ring DNA circles in vivo).   15. Its T m decreases because the charges on the phosphate groups are less  shielded from each other at lower ionic strength and hence repel each  other more strongly, thereby destabilizing the double helix.   17. The segment with 20% A residues (i.e., 40% A • T base pairs) con¬  tains 60% G • C base pairs and therefore melts at a higher tempera¬  ture than a segment with 30% A residues (i.e., 40% G • C base pairs).    23. The target sequence consists of 6 symmetry-related base pairs. Since  there are 4 possible base pairs (A • T, T • A, G • C, and C • G), the  probability that any two base pairs are randomly related by symme¬  try is 1/4. Hence, the probability of finding all 6 pairs of base pairs  by random chance is (1/4) 5 6 = 2.4 X 10 -4 .   23. Because they interact closely with DNA, protamines must be rich in  basic amino acids. In fact, they are particularly rich in arginine.   27. (a) The contour length is 3 X 10 7 bp X 3.4 A/bp = 1.7 X 10 8  A = 17 mm.   (b) A nucleosome, which binds —200 bp, compresses the DNA to an  80-A-high supercoil. The length of the DNA is therefore (80 A/200 bp)  X (5 X 10 7 bp) = 2 X 10 7 A = 2 mm.   29. Histones are required in large amounts during a relatively short pe¬  riod when DNA is replicated prior to cell division. The large num¬  ber of histone genes allows the efficient production of histones.    1. Okazaki fragments are 1000 to 2000 nt long, and the E. coli chro¬  mosome contains 4.6 X 10 6 bp. Therefore, E. coli chromosomal  replication requires 2300 to 4600 Okazaki fragments.   3. As indicated in Fig. a {below), nucleotides would be added to a  polynucleotide strand by attack of the 3'-OH of the incoming nu¬  cleotide on the 5' triphosphate group of the growing strand with the  elimination of PP Z . The hydrolytic removal of a mispaired nucleotide  by the 5' —> 3' exonuclease activity (Fig. b, below) would leave only  an OH group or monophosphate group at the 5' end of the DNA  chain. This would require an additional activation step before further  chain elongation could commence.   cause it removes RNA primers and replaces them with DNA. Absence  of this activity would be lethal.   7. When DNA polymerase begins synthesizing a new strand, it binds   the template DNA to which an RNA primer is already base paired.    In order to extend the primer, the polymerase active site must  accommodate the DNA-RNA hybrid helix, which has an A-DNA-  like structure (Fig. 24-4).   AT-rich DNA is less stable than GC-rich DNA and therefore would  more readily melt apart, a requirement for initiating replication.  DNA polymerase could extend a primer that entered its active site,  but polymerization would not be highly processive unless the sliding  clamp was in place. If the primer first associates with the clamp,  which then interacts with DNA polymerase, the primer can be ex¬  tended in a processive and therefore more efficient manner.   The E. coli replication system can fully replicate only circular DNAs.  Bacteria do not have a mechanism (e.g., telomerase-catalyzed exten¬  sion of telomeres) for replicating the extreme 3' ends of linear tem¬  plate strands.   After adding the 6-nt telomere sequence, telomerase translocates to  the new 3' end to add another repeat. The RNA template includes  a region of overlap (~3 nt) that helps position the enzyme and tem¬  plate for the next addition of nucleotides (see Fig. 25-26).    (b) When 5BU incorporated into DNA pairs with G, the result is an  A • T —> G • C transition after two more rounds of DNA replication:  A • T —» A • 5BU —> G • 5BU —» G • C    (a) The original A • T base pair becomes an I • T base pair. When  the DNA replicates, the template T will pair with A, and the tem¬  plate I will pair with C. Thus, one daughter cell will be normal and  one will have an abnormal I • C base pair, (b) After the second round  of cell division, two cells will be normal (A • T base pairs). One cell  will have an abnormal I • C base pair, and one will have a mutated  C • G base pair.   The half-life, t m = 0.693/rate = 0.693/(1.7 X 1(T 13 s _1 ) = 4.1 X  10 12 s or 130,000 years.   The triphosphatase destroys nucleotides containing the modified base  before they can be incorporated into DNA during replication.   Since thymine is a normal DNA base, the repair systems cannot de¬  termine whether such a T or its opposing G is the mutated base.  Consequently, only about half of the deaminated 5-methylcytosines  are correctly repaired.   Base excision repair. The deaminated base can be recognized because  hypoxanthine does not normally occur in DNA.   E. coli contains a low concentration of dUTP, which DNA poly¬  merase incorporates into DNA in place of dTTP. The resulting uracil    bases are rapidly excised by uracil-DNA glycosylase followed by nu¬  cleotide excision repair (NER), which temporarily causes a break in  the DNA chain. DNA that is isolated before DNA polymerase I and  DNA ligase can complete the repair process would be fragmented.  However, in the absence of a functional uracil-DNA glycosylase, the  inappropriate uracil residues would remain in place, and hence lead¬  ing strand DNA would be free of breaks. The lagging strand, being  synthesized discontinuously, would still contain breaks, although  fewer than otherwise.   31. DNA polymerase iq can synthesize a complementary DNA strand,  but the thymine dimer is still present. It can be repaired later by the  NER pathway.   33. (a) Loss of the helicase DnaB, which unwinds DNA for replication,  would be lethal.   (b) Loss of Pol I would prevent the excision of RNA primers and  would therefore be lethal.   (c) SSB prevents reannealing of separated single strands. Loss of SSB  would be lethal.   (d) RecA protein mediates the SOS response and homologous  recombination. Loss of RecA would be harmful but not necessarily  lethal.   (b) Because it lacks a 3'-OH group, the cordycepin incorporated into  a growing RNA chain cannot support further chain elongation in the  3 ' —> 3' direction.   3. The top strand is the sense strand. Its TATGAT segment differs by  only one base from the TATAAT consensus sequence of the pro¬  moters — 10 sequence; its TTTACA sequence differs by only one  base from the TTGACA consensus sequence of the promoters —33  sequence and is appropriately located ~25 nt to the 5' side of the  — 10 sequence; and the initiating G nucleotide is the only purine that  is located ~10 nt downstream of the —10 sequence.   5. G • C base pairs are more stable than A • T base pairs. Hence, the  more G • C base pairs that the promoter contains, the more difficult  it is to form the open complex during transcription initiation.   7. (a) Operons allow cells to turn sets of related genes on and off to¬  gether, thereby maximizing efficiency, since all the necessary genes are  expressed at the same time and in the same amount.   (b) In eukaryotes, genes in different locations can be turned on or  off at the same time if they share the same transcriptional regulatory  sequences, such as core promoter elements and enhancers that inter¬  act with the same transcription factors.   9. Increasing the error rate of transcription increases the chances of in¬  troducing a mutation that prevents the virus from completing its life  cycle in the host cell.   11 . In the presence of bicyclomycin, transcription of Rho-dependent  genes does not terminate, causing read-through into adjacent cod¬  ing regions. This results in the transcription of the adjacent gene(s),  often causing the inappropriate expression of the corresponding  protein(s).   13. By introducing a T7 promoter into recombinant DNA and using  T7 RNAP, genetic engineers can control the expression of a spe¬  cific gene without interference from other RNAP enzymes or other    Solutions to Odd Problems SP-19  promoter sequences that might be present in the experimental  system.   15. The cell lysates can be applied to a column containing a matrix with  immobilized poly(dT). The poly(A) tails of processed mRNAs will  bind to the poly(dT) while other cellular components are washed  away. The mRNAs can be eluted by decreasing the salt concentration  to destabilize the A • T base pairs.   17. (a) The phosphate groups of the phosphodiester backbone of the  mRNA will be labeled at all sites where a-[ 32 P]ATP is used as a sub¬  strate by RNA polymerase.   (b) 32 P will appear only at the 5' end of mRNA molecules that  have A as the first residue (this residue retains its a and (3 phos¬  phates). In all other cases where (3-[ 32 P]ATP is used as a substrate  for RNA synthesis, the (3 and y phosphates are released as PP Z (see  Fig. 26-7).   (c) No 32 P will appear in the RNA chain. During polymerization,  the (3 and y phosphates are released as PP Z . The terminal (y) phos¬  phate of an A residue at the 5' end of an RNA molecule is removed  during the capping process.   19. The active site of poly (A) polymerase is narrower because it does not  need to accommodate a template strand.   21 . The mRNA splicing reaction, which requires no free energy input  and results in no loss of phosphodiester bonds, is theoretically re¬  versible in vitro. However, the degradation of the excised intron  makes the reaction irreversible in the cell.   23. Inhibition of snRNA processing interferes with mRNA splicing. As  a result, host mRNA cannot be translated, so the host ribosomes will  synthesize only viral proteins.   1. A 4-nt insertion would add one codon and shift the gene’s reading  frame by one nucleotide. The proper reading frame could be restored  by deleting a nucleotide. Gene function, however, would not be  restored if (a) the 4-nt insertion interrupted the codon for a func¬  tionally critical amino acid; (b) the 4-nt insertion created a codon for  a structure-breaking amino acid; (c) the 4-nt insertion introduced a  Stop codon early in the gene; or (d) the 1-nt deletion occurred far  from the 4-nt insertion so that even though the reading frame was  restored, a long stretch of frame-shifted codons separated the inser¬  tion and deletion points.   3. The possible codons are UUU, UUG, UGU, GUU, UGG, GUG,  GGU, and GGG. The encoded amino acids are Phe, Leu, Cys, Val,  Trp, and Gly (Table 27-1).   5. There are still two other Stop codons, UAA and UAG, to terminate  translation.   7. (a) Each ORF begins with an initiation codon (ATG) and ends with  a Stop codon (TGA):   ATGCTCAACTATATGTGA encodes vir-2 and ATGCCGCAT-  GCTCTGTTAATCACATATAGTTGA on the complementary strand  encodes vir-1.   9. (a) Like an aaRS, Xpot must recognize features of tRNA structure  that are present in all tRNAs, such as the acceptor stem and the Tv|iC  loop, (b) Xpot can distinguish mature and pre-tRNAs because ma¬  ture tRNAs have a processed 5' end with a single phosphate group,  and the 3' end must be a —CCA sequence (see Fig. 27-3).   11. Gly and Ala; Val and Leu; Ser and Thr, Asn and Gin; and Asp  and Glu.   13. Only newly synthesized bacterial polypeptides have fMet at their  N-terminus. Consequently, the appearance of fMet in a mammalian  system signifies the presence of invading bacteria. Leukocytes that  recognize the fMet residue can therefore combat these bacteria  through phagocytosis.   17. Ribosomes cannot translate double-stranded RNA, so the base pair¬  ing of a complementary antisense RNA to an mRNA prevents its  translation.   19. As expected, the correctly charged tRNAs (Ala-tRNA Ala and  Gln-tRNA Gln ) bind to EF-Tu with approximately the same affin¬  ity, so they are delivered to the ribosomal A site with the same  efficiency. The mischarged Ala-tRNA Gln binds to EF-Tu much more  loosely, indicating that it may dissociate from EF-Tu before it  reaches the ribosome. The mischarged Gln-tRNA Ala binds to  EF-Tu much more tightly, indicating that EF-Tu may not be able  to dissociate from it at the ribosome. These results suggest that ei¬  ther a higher or a lower binding affinity could affect the ability of  EF-Tu to carry out its function, which would decrease the rate at  which mischarged aminoacyl- tRNAs bind to the ribosomal A site  during translation.   21 . Transpeptidation involves the nucleophilic attack of the amino group  of the aminoacyl-tRNA on the carbonyl carbon of the  peptidyl-tRNA. As the pH increases, the amino group becomes more  nucleophilic (less likely to be protonated).   23. The enzyme hydrolyzes peptidyl-tRNA molecules that dissociate  from a ribosome before normal translation termination takes place.  Because peptide synthesis is prematurely halted, the resulting  polypeptide, which is still linked to tRNA, is likely to be nonfunc¬  tional. Peptidyl-tRNA hydrolase is necessary for recycling the amino  acids and the tRNA.   27. Aminoacylation occurs via pyrophosphate cleavage of ATP, and hence  the aminoacylation of 100 tRNAs requires 200 ATP equivalents;  translation initiation requires 1 GTP (1 ATP equivalent); 99 cycles of  elongation require 99 GTP (99 ATP equivalents) for EF-Tu action;  99 cycles of ribosomal translocation require 99 GTP (99 ATP equiv¬  alents) for EF-G action; and translation termination requires 1 GTP  (1 ATP equivalent), bringing the total energy cost to 200 + 1 + 99  + 99 + 1 = 400 ATP equivalents.   1. Virtually all the DNA sequences in E. coli are present as single copies,  so the renaturation of E. coli DNA is a straightforward process of  each fragment reassociating with its complementary strand. In con¬  trast, the human genome contains many repetitive DNA sequences.  The many DNA fragments containing these sequences find each  other to form double-stranded regions (renature) much faster than  the single-copy DNA sequences that are also present, giving rise to a  biphasic renaturation curve.   3. The Daphnia and Drosophila genomes are similar in size (200,000 kb  versus 180,000 kb), but Daphnia contains far more genes (~30,000  versus ~ 13,000). The Daphnia genome is much smaller than the hu¬  man genome (200,000 kb versus 3,038,000 kb) but appears to con¬  tain more genes (~30,000 versus ~23,000).   5. (a) Translation of CAG repeats will yield polypeptides containing  polyglutamine (from the CAG codon), polyserine (from the AGC  codon), and polyalanine (from the GCA codon), (b) Translation of  CTG repeats will yield polypeptides containing polyleucine (from the  CUG codon), polycysteine (from the UGC codon), and polyalanine  (from the GCU codon).   7 . Oi is the primary repressor-binding site, so lac repressor cannot sta¬  bly bind to the operator in its absence and repression cannot occur.   9. In the absence of p-galactosidase (the product of the lacZ gene), lac¬  tose is not converted to the inducer allolactose. Consequently, lac en¬  zymes, including galactoside permease, are not synthesized.   11. In eukaryotes, transcription takes place in the nucleus and transla¬  tion occurs in the cytoplasm. Hence, in eukaryotes, ribosomes are  never in contact with nascent mRNAs, an essential aspect of the at¬  tenuation mechanism in prokaryotes.   13. Red-green color blindness is conferred by a mutation in an  X-linked gene so that female carriers of the condition, who do not  appear to be red-green colorblind, have one wild-type gene and one  mutated gene. In placental mammals such as humans, females are  mosaics of clones of cells in which only one of their two X chro¬  mosomes is transcriptionally active. Hence in a female carrier of  red-green color blindness, the transcriptionally active X chromo¬  some in some clones will contain the wild-type gene and the oth¬  ers will contain the mutated gene. The former type of retinal clone  is able to differentiate red and green light, whereas the latter type  of retinal clone is unable to do so. Apparently, these retinal clones  are small enough so that a narrow beam of light is necessary to sep¬  arately interrogate them.   In acetyllysine, the cationic side chain of Lys has been converted to  a polar but uncharged side chain.   17. Histone and DNA methylation requires S-adenosylmethionine  (SAM), which becomes S-adenosylhomocysteine after it gives up its  methyl group (Fig. 21-18). S-Adenosylhomocysteine is converted  back to methionine, the precursor of SAM, in a reaction in which  the methyl group is donated by the folic acid derivative tetrahydro-  folate (THF; Fig. 21-18). A shortage of this cofactor could limit cel¬  lular production of SAM, which would result in the  undermethylation of histones and DNA.    19. A sequence located downstream of the genes promoter (i.e., within  the coding region) could regulate gene expression if it were recog¬  nized by the appropriate transcription factor such that the resulting  DNA-protein complex successfully recruited RNA polymerase to the  promoter.   21. The susceptibility of RNA to degradation in vivo makes it possible  to regulate gene expression by adjusting the rate of mRNA degrada¬  tion. If mRNA were very stable, it might continue to direct transla¬  tion even when the cell no longer needed the encoded protein.   23. Since there are 63 Vh, 27 D, and 6 / H segments that can be used to  assemble the coding sequence of the variable region of the heavy  chain, somatic recombination could theoretically generate 63 X 27  X 6 = 10,530 heavy chain genes (junctional flexibility would in¬  crease this number). Since each immunoglobulin molecule contains  two identical heavy chains and two identical light chains, the possi¬  ble number of immunoglobulins would be 10530 X 2000 = ~21  million.   25. B cells are diploid [have two sets of genes specifying heavy chains  and fours sets of genes specifying light chains (two k’s and two  X’s)]. Hence, if allelic exclusion were defective, they would con¬  tinue to rearrange gene segments even after functional heavy and  light chain genes had been assembled. Such a B cell could produce  more than one type of heavy and light chain. The resulting mixed  chain immunoglobulins would be unable to cross-link antigens  since the two antigen-binding sites would have different binding  specificities.   27. In multicellular organisms, apoptosis of damaged cells minimizes  damage to the entire organism. For a single-celled organism, survival  of a genetically damaged cell is preferable in a Darwinian sense to its  death.   29. The esc gene is apparently a maternal-effect gene. Thus, the proper  distribution of the esc gene product in the fertilized egg, which is ma¬  ternally specified, is sufficient to permit normal embryonic develop¬  ment regardless of the embryos genotype.    Numbers and Greek letters are alphabetized as if they were spelled out.   ABC transporter. A member of a large family of transmembrane proteins  that use the free energy of ATP to mediate the transport of polar or  nonpolar substances across a membrane.   ABO blood group antigens. The oligosaccharide components of  glycolipids on the surfaces of erythrocytes and other cells.   Absolute configuration. The spatial arrangement of chemical groups  around a chiral center.   Absorbance ( A ). A function of the amount of light transmitted through a  solution  relative to the incident light (7 0 ) at a given wavelength:   Absorptivity (s). A constant that relates the absorbance of a solution to  the concentration of the solute at a given wavelength. Also called the  extinction coefficient.   Acceptor stem. The base-paired region of a tRNA molecule that contains  the 5 ' end and the 3' end, to which an amino acid is attached.   Accessory pigment. A molecule in a photosynthetic system that absorbs  light at wavelengths other than those absorbed by chlorophyll.   Acid-base catalysis. A catalytic mechanism in which partial proton transfer  from an acid or partial proton abstraction by a base lowers the free  energy of a reaction’s transition state. See also general acid catalysis and  general base catalysis.   Acidic solution. A solution whose pH is less than 7.0 ([H + ] > 10 -7 M).   Acidosis. A pathological condition in which the pH of the blood drops  below its normal value of 7.4.   Action potential. The wave of transient depolarization and repolarization  that constitutes the electrical signal generated by a nerve cell.   Active site. The region of an enzyme in which catalysis takes place.   Active transport. The transmembrane movement of a substance from low  to high concentrations by a protein that couples this endergonic  transport to an exergonic process such as ATP hydrolysis.   Activity. A solute’s concentration, corrected for its nonideal behavior at  concentrations greater than infinite dilution.   Acyl-carrier protein. A phosphopantetheine-containing protein that binds  the intermediates of fatty acid synthesis as thioesters.   Acyl-enzyme intermediate. An intermediate of peptide bond hydrolysis in  which the carbonyl carbon of the scissile bond is covalently bound to the  enzyme nucleophile that attacked it.   Acyl group. A portion of a molecule with the formula —COR, where R is  an alkyl group.   hormone binding to a cell-surface receptor activates a G protein that in  turn stimulates adenylate cyclase to synthesize the second messenger  3 ',5 '-cyclic AMP (cAMP) from ATP.   Adipocyte. Fat cell, which is specialized for the synthesis and storage of  triacylglycerols from free fatty acids.   ADP-ATP translocator. A membrane transport protein with an adenine  nucleotide—binding site that alternately allows ADP to enter and ATP to  exit the mitochondrial matrix.   Adrenergic receptor. A cell-surface receptor that binds and responds to  adrenal hormones such as epinephrine and norepinephrine. Also called  an adrenoreceptor.    Aerobe. An organism that uses 0 2 as an oxidizing agent for nutrient  breakdown.   Affinity chromatography. A procedure in which a molecule is separated  from a mixture of other molecules by its ability to bind specifically to an  immobilized ligand. See also metal chelate affinity chromatography.   Affinity labeling. A technique in which a labeled substrate analog reacts  irreversibly with, and can thereby be used to identify, a group in an  enzyme’s active site.   Agarose. Linear carbohydrate polymers, made by red algae, that form a  loose mesh.   Agonist. A substance that binds to a receptor so as to evoke a cellular  response.   Alcoholic fermentation. A metabolic pathway that synthesizes ethanol  from pyruvate through decarboxylation and reduction.   Alditol. A sugar produced by reduction of an aldose or ketose to a  polyhydroxy alcohol.   Aldol cleavage. A carbon-carbon cleavage reaction of an aldol (an aldehyde  or ketone with a (3 hydroxyl group) that yields smaller carbonyl  compounds.   Aldonic acid. A sugar produced by oxidation of an aldose aldehyde group  to a carboxylic acid group.   Alkalosis. A pathological condition in which the pH of the blood rises  above its normal value of 7.4.   Allele. An alternate form of a gene; diploid organisms contain two alleles  for each gene, which may or may not be identical.   Allosteric effector. A small molecule whose binding to a protein affects the  function of another site on the protein.   Allosteric interaction. The binding of ligand at one site in a   macromolecule that affects the binding of other ligands at other sites in  the molecule. See also cooperative binding.   olol motif. A protein motif consisting of two a helices packed against each  other with their axes inclined.   ot/p barrel. A (3 barrel in which successive parallel (3 strands are connected  by a helices such that a barrel of a helices surrounds the (3 barrel.   ot carbon. The carbon atom of an amino acid to which the amino and  carboxylic acid groups are attached.   a cell. A pancreatic islet cell that secretes the hormone glucagon in  response to low blood glucose levels.   ot helix. A regular secondary structure of polypeptides, with 3.6 residues  per right-handed turn, a pitch of 3.4 A, and hydrogen bonds between  each backbone N—H group and the backbone C=0 group that is four  residues earlier.   Alternative splicing. The tissue-specific patterns of splicing of a given pre-  mRNA that result in variations in the excision and retention of introns  and exons.   Alzheimer’s disease. A neurodegenerative disease characterized by the  precipitation of (3 amyloid protein in the brain.   Ames test. A method for assessing the mutagenicity of a compound from  its ability to cause genetically defective strains of bacteria to revert to  normal growth.   Amino acid. A compound consisting of a carbon atom to which are  attached a primary amino group, a carboxylic acid group, a side chain  (R group), and an H atom. Also called an a-amino acid.   Amino group. A portion of a molecule with the formula —NH 2 , —NHR,  or —NR 2 , where R is an alkyl group. Amino groups are usually  protonated at physiological pH.   Amino sugar. A sugar in which one or more OH groups are replaced by  an amino group, which is often acetylated.   Amino terminus. The end of a polypeptide that has a free amino group.  Also called the N-terminus.   Aminoacyl site (A site). The ribosomal site where a tRNA with an  attached aminoacyl group binds during protein synthesis.   Aminoacyl-tRNA (aa-tRNA). The covalent ester complex between an  “activated” amino acid and a tRNA molecule.   Aminoacyl-tRNA synthetase (aaRS). An enzyme that catalyzes the ATP-  dependent esterification of an amino acid to a tRNA with high  specificity for the amino acid and the tRNA molecule.   Amphibolic. A term to describe a metabolic process that can be either  catabolic or anabolic.   Amphiphilic substance. A substance that contains both polar and nonpolar  regions and is therefore both hydrophilic and hydrophobic. Also called  an amphipathic substance.   Amyloid. Insoluble extracellular aggregates of fibrous protein that  characterize such diseases as Alzheimer’s disease and the transmissible  spongiform encephalopathies.   Anabolism. The reactions by which biomolecules are synthesized from  simpler components.   Anaerobe. An organism that does not use 0 2 as an oxidizing agent for  nutrient breakdown. An obligate anaerobe cannot grow in the presence  of 0 2 , whereas a facultative anaerobe can grow in the presence or  absence of 0 2 .   Anaplerotic reaction. A reaction that replenishes the intermediates of a  metabolic pathway.   Androgen. A steroid that functions primarily as a male sex hormone.   Angina pectoris. Chest pain due to insufficient blood supply to the heart.  Often just called angina.   Anion exchanger. A cationic matrix used to bind anionic molecules in ion  exchange chromatography.   Anneal. To maintain conditions that allow loose base pairing between  complementary single polynucleotide strands so that properly paired  double-stranded segments form.   Anomeric carbon. The carbonyl carbon of a monosaccharide unit, which  becomes a chiral center when the sugar cyclizes to a hemiacetal or  hemiketal.   Anomers. Sugars that differ only in the configuration around the anomeric  carbon. In the a anomer, the OH substituent of the anomeric carbon is  on the opposite side of the ring from the CH 2 OH group at the chiral  center that designates the D or L configuration. In the (3 anomer, the  OH substituent is on the same side.   Antagonist. A substance that binds to a receptor but does not elicit a  cellular response.   Antenna chlorophyll. A chlorophyll group that absorbs light energy and  passes it on to a photosynthetic reaction center by exciton transfer.   Anti conformation. A purine or pyrimidine nucleotide conformation in  which the ribose and the base point away from each other. See also syn  conformation.   Antibody. A protein produced by an animal’s immune system in response  to the introduction of a foreign substance (an antigen); it contains at  least one pair each of identical heavy and light chains. Also called an  immunoglobulin (Ig).   Anticodon. The sequence of three nucleotides in tRNA that recognizes an  mRNA codon through complementary base pairing.   Anticodon arm. The conserved stem-loop structure in a tRNA molecule  that includes the anticodon.   Antigen. A substance that elicits an immune response (production of  antibodies) when introduced into an animal; it is specifically recognized  by an antibody.    Antioxidant. A substance that destroys reactive oxygen species such as 0 2 *  or -OH.   Antiport. A transmembrane channel that simultaneously transports two  molecules or ions in opposite directions. See also symport and uniport.   Antisense RNA. A single-stranded RNA molecule that forms a double-  stranded structure with a complementary mRNA so as to block its  translation into protein.   Antisense strand. The DNA strand that serves as a template for  transcription; it is complementary to the RNA. Also called the  noncoding strand.   AP site. An apurinic or apyrimidinic site; the deoxyribose residue  remaining after the removal of a base from a DNA strand.   Apoenzyme. An enzyme that is inactive due to the absence of a cofactor.   Apolipoprotein. A protein component of a lipoprotein. Also called an  apoprotein.   Apoprotein. A protein without the prosthetic group or metal ion that  renders it fully functional. See also apoenzyme and apolipoprotein.   Apoptosis. Cell death by a regulated process in which the cell shrinks and  fragments into membrane-bounded portions for phagocytosis by other  cells. See also necrosis.   Aptamer. A nucleic acid whose conformation allows it to bind a particular  ligand with high specificity and high affinity.   Aquaporin. A membrane protein that mediates the rapid diffusion of water  molecules but not protons or other ions across a membrane.   Archaea. One of the two major groups of prokaryotes (the other is  eubacteria). Also known as archaebacteria.   Assay. A laboratory technique for detecting, and in many cases quantifying,  a macromolecule or its activity.   Atherosclerosis. A disease characterized by the formation of cholesterol-  containing fibrous plaques in the walls of blood vessels, leading to loss of  elasticity and blockage of blood flow.   Atomic force microscopy (AFM). A technique for determining molecular  topography at subnanomolar resolution by passing a mechanical probe  over a surface and monitoring its deflection by optical or electronic  means.   ATP mass action ratio. The ratio [ATP]/[ADP][PJ, which influences the  rate of electron transport and oxidative phosphorylation.   ATPase. An enzyme that catalyzes the hydrolysis of ATP to ADP + P z or  the reverse reaction.   Attenuation. An RNA-based mechanism in prokaryotes for regulating gene  expression in which the availability of an amino acid determines whether  an operon (consisting of genes for the enzymes that synthesize the amino  acid) is transcribed.   Attenuator. A prokaryotic control element that governs transcription of an  operon according to the availability of an amino acid synthesized by the  proteins encoded by the operon.   Autocatalytic reaction. A reaction in which a product molecule can act as  a catalyst for the same reaction; the reactant molecule therefore appears  to catalyze its own reaction.   Autoimmune disease. A disease in which the immune system has lost  some of its self-tolerance and attacks certain self-antigens.   Autolysis. An autocatalytic process in which a molecule catalyzes its own  degradation.   Autophagy. The process in which damaged or unneeded organelles are  enclosed by a membrane and the resulting vesicle fuses with a lysosome  that digests its contents.   Autoradiography. A process in which X-ray film records the positions of  radioactive entities, such as proteins or nucleic acids, that have been  immobilized in a matrix such as a nitrocellulose membrane or an  electrophoretic gel.    Autotroph. An organism that can synthesize all its cellular components  from simple molecules using the energy obtained from sunlight  (photoautotroph) or from the oxidation of inorganic compounds  (chemolithotroph).   Axial substituent. A group that extends approximately perpendicularly from  the plane of the ring to which it is bonded. See also equatorial substituent.   Backbone. The atoms that form the repeating linkages between successive  residues of a polymeric molecule, exclusive of the side chains. Also called  the main chain.   Bacteria. The organisms comprising the two major groups of prokaryotes,  the archaea and the eubacteria.   Bacterial artificial chromosome (BAC). A plasmid-derived DNA molecule  that can replicate in a bacterial cell. BACs are commonly used as cloning  vectors.   Bacteriophage. A virus specific for bacteria. Also known as a phage.   Barr body. The condensed and darkly staining inactive X chromosome in  the nucleus of a female mammalian cell.   Base. ( 1 ) A substance that can accept a proton. ( 2 ) A purine or pyrimidine  component of a nucleoside, nucleotide, or nucleic acid.   Base excision repair (BER). The removal and replacement of a damaged  nucleotide in DNA that is initiated by the removal of its base.   Base pair. The specific hydrogen-bonded association between nucleic acid  bases. The Watson-Crick base pairs are A • T and G • C.   Basic helix-loop-helix (bHLH) motif. A eukaryotic protein motif that  includes a basic DNA-binding region followed by two amphipathic  helices connected by a loop. The second helix, which mediates protein  dimerization, is often continuous with a leucine zipper motif.   Basic solution. A solution whose pH is greater than 7.0 ([H + ] < 10 -7 M).   Beer-Lambert law. The equation that describes the relationship between a  solutes absorbance (A) and its concentration (c): A = ec/where e is the  solutes molar absorptivity and / is the length of the light path.   Beriberi. A disease caused by a deficiency of thiamine (vitamin B^, which  is a precursor of the cofactor thiamine pyrophosphate.   pctp motif. A protein motif consisting of an a helix connecting two  parallel strands of a p sheet.   P barrel. A protein motif consisting of a p sheet rolled into a cylinder.   P bulge. An irregularity in a p sheet resulting from an extra residue that is  not hydrogen bonded to a neighboring chain.   P cell. A pancreatic islet cell that secretes the hormone insulin in response  to high blood glucose levels.   P hairpin. A protein motif in which two antiparallel p strands are  connected by a reverse turn.   P oxidation. A series of enzyme-catalyzed reactions in which fatty acids are  progressively degraded by the removal of two-carbon units as acetyl-CoA.   P sheet. A regular secondary structure in which extended polypeptide  chains form interstrand hydrogen bonds. In parallel p sheets, the  polypeptide chains all run in the same direction; in antiparallel p sheets,  neighboring chains run in opposite directions.   Bilayer. An ordered, double layer of amphiphilic molecules in which polar  segments point toward the two solvent-exposed surfaces and the  nonpolar segments associate in the center.   Bile acid (bile salt). An amphiphilic cholesterol derivative that acts as a  detergent to solubilize lipids for digestion and absorption.   Binding change mechanism. The mechanism whereby the subunits of the  FjFq-ATP synthase adopt three successive conformations to convert ADP  + P* to ATP as driven by the dissipation of the transmembrane proton  gradient.   Bioavailability. A measure of the fraction of a drug that reaches its target  tissue, which depends on the drug’s dosage and pharmacokinetics.   Biochemical standard state. A set of conditions including unit activities of  the species of interest, a temperature of 25°C, a pressure of 1 atm, and a  pH of 7.0.   Biofilm. A surface-associated aggregate of bacterial cells and extracellular  polysaccharides that protect the cells from environmental assault.   Bioinformatics. The study of biological information in the form of  molecular sequences and structures; e.g., structural bioinformatics.   Biopterin. A pterin derivative that functions as a cofactor in the  hydroxylation of phenylalanine to produce tyrosine.   Blastoderm. The single layer of cells surrounding a yolklike core that forms  during the early development of an insect larva.   Blunt ends. The fully base-paired ends of a DNA fragment that has been  cleaved by a restriction endonuclease that cuts the DNA strands at  opposing sites.   Bohr effect. The decrease in 0 2 binding affinity of hemoglobin in response  to a decrease in pH.   bp. Base pair, the two hydrogen-bonded bases that form the repeating units  of double helical nucleic acids. Thousands of base pairs (kilobase pairs)  are abbreviated kb.   Bradford assay. A spectroscopic technique for determining protein  concentration in solution from the absorbance of a dye bound to the  protein.   Branch migration. The movement of a crossover point in a Holliday  junction during DNA recombination.   Bromodomain. A protein module that binds acetylated Lys residues in  histones.   (Brownian) back-and-forth fluctuations are converted to coherent forward  motion by inhibiting the backward motion.   Buffer. A solution of a weak acid and its conjugate base in approximately  equal quantities. Such a solution resists changes in pH on the addition  of acid or base.   Buffering capacity. The ability of a buffer solution to resist pH changes on  addition of acid or base. Buffers are most useful when the pH is within  one unit of its component acid’s p K   C value. A measure of the quantity of an organism’s unique genetic  material.   C-value paradox. The occurrence of exceptions to the rule that an   organism’s DNA content (C value) is correlated to the complexity of its  morphology and metabolism.   Cahn-Ingold-Prelog system (RS system). A system for unambiguously  describing the configurations of molecules with one or more asymmetric  centers by assigning a priority ranking to the substituent groups of each  asymmetric center.   Calmodulin (CaM). A small Ca 2+ -binding protein that binds to  other proteins in the presence of Ca 2+ and thereby regulates their  activities.   Calvin cycle. The sequence of photosynthetic dark reactions in which  ribulose-5-phosphate is carboxylated, converted to three-carbon  carbohydrate precursors, and regenerated. Also called the reductive  pentose phosphate cycle.   Cap. A 7-methylguanosine residue that is posttranscriptionally appended to  the 3' end of a eukaryotic mRNA.   Capillary electrophoresis (CE). Electrophoretic procedures carried out in  small diameter capillary tubes.   Carbamate. The product of a reaction between C0 2 and an amino group:  —NH—COO - .    Carbohydrate. A compound with the formula (C • H 2 0)^ where n ^ 3.   Carbonyl group. A portion of a molecule with the formula .C — O.    Carboxyl group. A portion of a molecule with the formula —COOH.  Carboxyl groups are usually ionized to carboxylate groups (COO ) at  physiological pH.   Carboxyl terminus. The end of a polypeptide that has a free carboxylate  group. Also called the C-terminus.   Carcinogen. An agent that damages DNA so as to induce a mutation that  leads to uncontrolled cell proliferation (cancer).   Caspase. A heterotetrameric cysteine protease that hydrolyzes cellular  proteins, including caspase zymogens, as part of the process of  apoptosis.   Catabolism. The degradative metabolic reactions in which nutrients and  cell constituents are broken down for energy and raw materials.   Catabolite repression. A phenomenon in bacteria in which the presence of  glucose prevents the expression of genes involved in the metabolism of  other fuels.   Catalyst. A substance that promotes a chemical reaction without itself  undergoing permanent change. A catalyst increases the rate at which a  reaction approaches equilibrium but does not affect the free energy  change of the reaction.   Catalytic perfection. The ability of an enzyme to catalyze a reaction as fast  as diffusion allows it to bind its substrates.   Catalytic triad. The hydrogen-bonded Ser, His, and Asp residues that  participate in catalysis in serine proteases and related enzymes.   Cataplerotic reaction. A reaction that drains the intermediates of a  metabolic pathway.   Catecholamine. A hydroxylated tyrosine derivative, such as dopamine,  norepinephrine, or epinephrine.   Catenate. To interlink circular DNA molecules like the links of a chain.   Cation exchanger. An anionic matrix used to bind cationic molecules in  ion exchange chromatography.   CCAAT box. A eukaryotic promoter element with the consensus sequence  CCAAT that, in many structural genes, is located 70 to 90 nucleotides  upstream from the transcription start site.   Cell cycle. The sequence of events between eukaryotic cell divisions; it  includes mitosis and cell division (M phase), a gap stage (G t phase), a  period of DNA synthesis (S phase), and a second gap stage (G 2 phase)  before the next M phase.   Central dogma of molecular biology. The paradigm that DNA directs its  own replication as well as its transcription to RNA, which is then  translated into a polypeptide. The flow of information is from DNA to  RNA to protein.   Centromere. The eukaryotic chromosomal region that attaches to the  mitotic spindle during cell division; it contains high concentrations of  repetitive DNA.   Ceramide. A sphingosine derivative with an acyl group attached to its  amino group.   Cerebroside. A ceramide with a sugar residue as a head group.   C 4 plant. A plant in which photosynthesis relies on C0 2 that has been  concentrated by incorporating it into oxaloacetate (a C 4 compound).   Chain-terminator procedure. A technique for determining the nucleotide  sequence of a DNA using dideoxy nucleotides so as to yield a collection  of daughter strands of all different lengths. Also called the dideoxy  method.   Channeling. The transfer of an intermediate product from one enzyme  active site to another in such a way that the intermediate remains  protected by the protein.   Chaotropic agent. A substance that increases the solubility of nonpolar  substances in water and thereby tends to denature proteins.   Chargaff’s rules. The observation, first made by Erwin Chargaff, that  duplex DNA has equal numbers of adenine and thymine residues and  equal numbers of guanine and cytosine residues.    Chemiosmotic theory. The postulate, now known to be correct, that the  free energy of electron transport is conserved by the formation of a  transmembrane proton gradient. The electrochemical potential of the  gradient is used to drive ATP synthesis.   Chemolithotroph. An autotrophic organism that obtains energy from the  oxidation of inorganic compounds.   Chiral center. An atom whose substituents are arranged such that it is not  superimposable on its mirror image. Also called an asymmetric center.   Chirality. The property of having one or more chiral centers. A chiral  molecule cannot be superimposed on its mirror image.   Chromatin. The complex of DNA and protein that comprises the  eukaryotic chromosomes.   Chromatin-remodeling complex. An ATP-dependent multisubunit protein  in eukaryotes that transiently disrupts DNA-histone interactions so as to  alter the accessibility of DNA in nucleosomes.   Chromatography. A technique for separating the components of a mixture  of molecules based on their partition between a mobile solvent phase  and a porous matrix (stationary phase).   Chromodomain. A protein module that binds methylated Lys residues in  histones.   Chromosome. The complex of protein and a single DNA molecule that  comprises some or all of an organism’s genome.   Chylomicrons. Lipoprotein particles that transport dietary triacylglycerols  and cholesterol from the intestines to the tissues.   Cis conformation. An arrangement of the peptide group in which  successive C a atoms are on the same side of the peptide bond.   Cis peptide. A conformation in which successive C a atoms are on the  same side of the peptide bond.   Citric acid cycle. A set of eight enzymatic reactions, arranged in a cycle, in  which free energy in the form of ATP, NADH, and FADH 2 is recovered  from the oxidation of the acetyl group of acetyl-CoA to C0 2 . Also  called the Krebs cycle and the tricarboxylic acid (TCA) cycle.   Clathrin. A three-legged protein that polymerizes to form a polyhedral  structure defining the shape of membranous vesicles that travel between  the plasma membrane and intracellular organelles such as the Golgi  apparatus.   Clinical trials. A three-phase series of tests of a drug’s safety, effectiveness,  and side effects in human subjects.   Clone. A collection of identical cells derived from a single ancestor.   Cloning. The production of exact copies of a DNA segment or the  organism that harbors it.   Cloning vector. A DNA molecule such as a plasmid, virus, or artificial  chromosome that can accommodate a segment of foreign DNA for  cloning.   Closed system. A thermodynamic system that can exchange energy but not  matter with its surroundings.   Coated vesicle. A membranous intracellular transport vesicle that is encased  by clathrin or another coat protein.   Codon. The sequence of three nucleotides in DNA or RNA that specifies a  single amino acid.   Coenzyme. A small organic molecule that is required for the catalytic  activity of an enzyme. A coenzyme may be either a cosubstrate or a  prosthetic group.   Coenzyme Q (CoQ). An isoprenoid that functions in electron-transport  pathways as a lipid-soluble electron carrier. Also called ubiquinone.   Cofactor. A small organic molecule (coenzyme) or metal ion that is  required for the catalytic activity of an enzyme.   Coiled coil. An arrangement of polypeptide chains in which two a helices  wind around each other, as in a keratin.   Cointegrate. The product of the fusion of two plasmids, which occurs as  an intermediate in transposition.    Colligative property. A physical property, such as freezing point depression  or osmotic pressure, that depends on the concentration of a dissolved  substance rather than on its chemical nature.   Colony hybridization. A procedure in which DNA from multiple cell  colonies is transferred to a membrane or filter and incubated with a  DNA or RNA probe to test for the presence of a desired DNA fragment  in the cell colonies. Also called in situ hybridization.   Combinatorial chemistry. A method for rapidly and inexpensively  synthesizing large numbers of related compounds by systematically  varying a portion of their structure.   Compartmentation. The division of a cell into smaller functionally discrete  systems.   Competitive inhibition. A form of enzyme inhibition in which a substance  competes with the substrate for binding to the enzyme active site and  thereby appears to increase K M .   Complementary DNA (cDNA). A DNA molecule, usually synthesized by  the action of reverse transcriptase, that is complementary to an mRNA  molecule.   Composite transposon. A genetic sequence that may include a variety of  genes and is flanked by IS-like elements; such transposons apparently  arose by the association of two independent IS elements.   Condensation reaction. The formation of a covalent bond between two  molecules, during which the elements of water are lost; the reverse of  hydrolysis.   Conjugate acid. The compound that forms when a base accepts a proton.   Conjugate base. The compound that forms when an acid donates  a proton.   Conjugate redox pair. An electron donor and acceptor that form a half¬  reaction. Also called a redox couple.   Conservative replication. A hypothetical mode of DNA duplication in  which the parental molecule remains intact and both strands of the  daughter duplex are newly synthesized.   Conservative substitution. A change of an amino acid residue in a protein  to one with similar properties, e.g., Leu to lie or Asp to Glu.   Constant region. The C-terminal portion of an antibody (immunoglobulin)  subunit, which does not exhibit the high sequence variability of the  antigen-recognizing (variable) region of the antibody.   Constitutive enzyme. An enzyme that is synthesized at a more or less  steady rate and that is required for basic cell function. Also called a  housekeeping enzyme. See also inducible enzyme.   Contact inhibition. The inhibition of proliferation in cultured animal cells  when the cells touch each other.   Contour map. A map containing lines (contours) that trace positions of  equal value of some property of the map (e.g., height above sea level,  electron density).   Convergent evolution. The independent development of similar  characteristics in unrelated species or proteins.   Cooperative binding. A situation in which the binding of a ligand at one  site on a macromolecule affects the affinity of other sites for the same  ligand. Both negative and positive cooperativity occur. See also allosteric  interaction.   Corepressor. A substance that acts together with a protein repressor to  block gene transcription.   Cori cycle. An interorgan metabolic pathway in which lactate produced by  glycolysis in the muscles is transported via the bloodstream to the liver,  where it is used for gluconeogenesis. The resulting glucose returns to the  muscles.   Cosubstrate. A coenzyme that is only transiently associated with an enzyme  so that it functions as a substrate.   Coupled enzymatic reaction. A technique in which the activity of an  enzyme is measured by the ability of a second enzyme to use the product  of the first enzymatic reaction to produce a detectable product.   Covalent catalysis. A catalytic mechanism in which the transient formation  of a covalent bond between the catalyst and a reactant lowers the free  energy of a reaction’s transition state.   CpG island. A cluster of CG dinucleotides located just upstream of many  vertebrate genes; such sequences occur elsewhere in the genome at only  one-fifth their randomly expected frequency.   Crassulacean acid metabolism (CAM). A variation of the C 4  photosynthetic cycle in which C0 2 is temporarily stored as  malate.   Cross talk. The interactions of different signal transduction pathways  through activation of the same signaling components, generation of a  common second messenger, or similar patterns of target protein  phosphorylation.   Cryoelectron microscopy (cryo-EM). A technique in electron microscopy  in which a sample is rapidly frozen to very low temperatures so that it  retains its native shape to a greater extent than in conventional electron  microscopy.   C 3 plant. A plant in which photosynthesis proceeds by the incorporation  of C0 2 into three-carbon compounds.   C3 '-endo. A ribose conformation in which C3' is displaced toward the  same side of the ring as C3k   C2 ' -endo. A ribose conformation in which C2' is displaced toward the  same side of the ring as C5'.   Curved arrow convention. A notation for indicating the movement of an  electron pair in a chemical reaction by drawing a curved arrow  emanating from the electrons and pointing to the electron-deficient  center that attracts the electron pair.   Cyanosis. A bluish skin color indicating the presence of deoxyhemoglobin  in the arterial blood.   Cyclic symmetry. A type of symmetry in which the asymmetric units of a  symmetric object are related by a single axis of rotation.   Cyclin. A member of a family of proteins that participate in regulating the  stages of the cell cycle and whose concentrations change dramatically  over the course of the cell cycle.   Cytochrome. A redox-active protein that carries electrons via a prosthetic  Fe-containing heme group.   Cytochromes P450. Heme-containing monooxygenases that catalyze the  addition of OH groups to drugs and toxins in order to detoxify them  and facilitate their excretion.   Cytoskeleton. The network of intracellular fibers that gives a cell its shape  and structural rigidity.   Cytosol. The contents of a cell excluding its nucleus and other membrane-  bounded organelles.   D. Dalton, a unit of molecular mass; 1/12th the mass of a 12 C atom.   D arm. A conserved stem-loop structure in a tRNA molecule that usually  contains the modified base dihydrouracil.   Dark reactions. The portion of photosynthesis in which NADPH and ATP  produced by the light reactions are used to incorporate C0 2 into  carbohydrates.   Debranching. The enzymatic removal of side chains from a branched  polymer such as glycogen.   Density gradient. A solution used in ultracentrifugation, in which the  concentration of a substance such as sucrose or CsCl increases from the  top to the bottom of the centrifuge tube.   Deoxy sugar. A monosaccharide produced by replacement of an OH group  by H.    Deoxyribonucleotide. A nucleotide in which the pentose is 2'-deoxyribose.  Also known as a deoxynucleotide.   Desaturase. An enzyme that introduces double bonds into a fatty acid.   Desensitization. A cell’s or organism’s adaptation to a long-term stimulus  through a reduced response to the stimulus.   Dextrorotatory. Rotating the plane of plane-polarized light clockwise from  the point of view of the observer; the opposite of levorotatory.   Diabetes mellitus. A disease in which the pancreas does not secrete  sufficient insulin (also called type 1, insulin-dependent, or  juvenile-onset diabetes) or in which the body has insufficient response  to circulating insulin (type 2, non-insulin-dependent, or maturity-  onset diabetes). Diabetes is characterized by elevated levels of glucose  in the blood.   Dialysis. A procedure in which solvent molecules and solutes smaller than  the pores in a semipermeable membrane freely exchange with the bulk  medium, while larger solutes are retained, thereby changing the solution  in which the larger molecules are dissolved.   Diffraction pattern. The record of the destructive and constructive  interferences of radiation scattered from an object. In X-ray  crystallography, this takes the form of a series of discrete spots resulting  from a collimated beam of X-rays scattering from a single crystal.   Diffusion-controlled limit. The theoretical maximum rate of an enzymatic  reaction in solution, about 10 8 to 10 9 M -1 • s" 1 .   Dihedral symmetry. A type of symmetry in which the asymmetric units  are related by a twofold rotation axis that intersects another rotation axis  at a right angle.   Dinucleotide binding fold. A protein structural motif consisting of two  (3a(3ap units, which binds a dinucleotide such as NAD + . Also called a  Rossmann fold.   Dipolar ion. A compound bearing oppositely charged groups. Also called a  zwitterion.   Disaccharide. A carbohydrate consisting of two monosaccharides linked by  a glycosidic bond.   Dissociation constant ( K ). The ratio of the products of the concentrations  of the dissociated species to those of their parent compounds at  equilibrium.   DNA. Deoxyribonucleic acid. A polymer of deoxynucleotides whose  sequence of bases encodes genetic information in all living cells.   DNA fingerprinting. A technique for distinguishing individuals on the  basis of DNA polymorphisms, such as the number of short tandem  repeats (STR).   DNA glycosylase. An enzyme that initiates base excision repair of DNA by  cleaving the glycosidic bond that links a nucleotide base to ribose.   DNA library. A set of cloned DNA fragments representing some or all of  an organism’s genome.   DNA microarray. A set of DNA segments of known sequence that are  immobilized on a solid support for the purpose of hybridizing them with  nucleic acids in test samples. Also called a DNA chip.   Dolichol. An long-chain polyisoprenoid that serves as a lipid-soluble carrier  of an AHinked oligosaccharide during its synthesis in the endoplasmic  reticulum.   Domain. A group of one or a few polypeptide segments of about 40-200  residues that folds into a globular unit.    Double-displacement reaction. A reaction in which a substrate binds and  a product is released in the first stage, and another substrate binds and  another product is released in the second stage.   Drug-drug interactions. Increases or decreases in the bioavailability of a  drug caused by the metabolic effects of another drug.   EC classification. The Enzyme Commission’s system for classifying and  numbering enzymes according to the type of reaction catalyzed.   Edman degradation. A chemical reaction that is used for the stepwise  removal and identification of the N-terminal residues of a polypeptide.   EF hand. A widespread helix-loop-helix structural motif that forms a  Ca 2+ -binding site.   Eicosanoids. C 2 o compounds derived from the C 2 o fatty acid arachidonic  acid and which act as local mediators. Prostaglandins, prostacyclins,  thromboxanes, leukotrienes, and lipoxins are eicosanoids.   separate compartments linked by a wire for transporting electrons and a  salt bridge for maintaining electrical neutrality; the simultaneous activity  of the half-reactions forms a complete oxidation-reduction reaction.   Electrochemical potential. The partial molar free energy of a substance  (chemical potential) in the presence of an electrical potential.   Electrogenic transport. The transmembrane movement of a charged  substance in a way that generates a charge difference across the  membrane.   Electron crystallography. A technique for determining molecular structure,  in which the electron beam of an electron microscope is used to elicit  diffraction from a two-dimensional crystal of the molecules of interest.   Electron density. The distribution of electrons that gives rise to a  diffraction pattern in X-ray crystallography.   Electron-transport chain. A series of membrane-associated electron carriers  that pass electrons from reduced coenzymes (NADH and FADH 2 ) to  molecular oxygen so as to recover free energy for the synthesis of ATP.   Electrophile. A group that contains an unfilled valence electron shell, or  contains an electron-deficient atom. An electrophile (electron-lover) reacts  readily with a nucleophile (nucleus-lover).   Electrospray ionization (ESI). A method for vaporizing macromolecules  for their mass spectrometry in which a macromolecular solution is  sprayed from a narrow capillary at high voltage to produce fine, highly  charged droplets that rapidly evaporate leaving the now charged  macromolecule in the gas phase.   Electrostatic catalysis. A catalytic mechanism in which the distribution of  charges about the catalytic site lowers the free energy of a reaction’s  transition state.   Elementary reaction. A simple one-step chemical process, several of which  may occur in sequence in a chemical reaction.   Elongase. An enzyme that adds acetyl units to a fatty acid previously  synthesized by fatty acid synthase.   Elongation factor. A protein that interacts with tRNA and/or the ribosome  during polypeptide synthesis.   Eluant. The solution used to wash material through a chromatographic  column.   Elution. The process of dislodging a molecule that has bound to a  chromatographic matrix.   Emergent property. A property of a complex system that is not   attributable to any of its individual components but becomes apparent  when all components are present.   Enantiomers. Molecules that are nonsuperimposable mirror images of one  another. Enantiomers are a type of stereoisomer.    Endergonic process. A process that has an overall positive free energy  change (a nonspontaneous process).   Endocrine gland. A tissue in higher animals that synthesizes and releases  hormones into the bloodstream.   Endocytosis. The internalization of extracellular material through the  formation of a vesicle that buds off from the plasma membrane; the  opposite of exocytosis. See also receptor-mediated endocytosis.   Endoglycosidase. An enzyme that catalyzes the hydrolysis of the glycosidic  bonds between two monosaccharide units within a polysaccharide.   Endonuclease. An enzyme that catalyzes the hydrolysis of the  phosphodiester bonds between two nucleotide residues within a  polynucleotide strand.   Endopeptidase. An enzyme that catalyzes the hydrolysis of a peptide bond  within a polypeptide chain.   Endoplasmic reticulum (ER). A labyrinthine membranous organelle in  eukaryotic cells in which membrane lipids are synthesized and proteins  destined for secretion, membrane insertion, and residence in certain  organelles undergo posttranslational modification.   Endosome. A membrane-bounded vesicle that receives materials that the  cell ingests via receptor-mediated endocytosis and passes them to  lysosomes for degradation.   Enediol intermediate. A reaction intermediate containing a carbon-carbon  double bond and a hydroxyl group attached to each carbon.   Energy coupling. The conservation of the free energy of electron transport  in a form that can be used to synthesize ATP from ADP + P z .   Enhanceosome. A complex containing several transcription factors that  regulates gene expression in eukaryotes.   Enhancer. A eukaryotic DNA sequence located some distance from the  transcription start site, where an activator of transcription may bind.   Enthalpy (H ). A thermodynamic quantity, H = U + PV, that is  equivalent to the heat absorbed at constant pressure ( q P ).   Entropy (5). A measure of the degree of randomness or disorder of a  system. It is defined as S = k B \n W, where k B is the Boltzmann  constant and W is the number of equivalent ways the system can be  arranged in its particular state.   Enzyme. A biological catalyst. Most enzymes are proteins; a few are RNA.   Enzyme-linked immunosorbent assay (ELISA). A technique in which a  molecule is detected, and in many cases quantified, by its ability to bind  an antibody to which an enzyme with an easily detected reaction product  is attached.   Enzyme saturation. A state in which the substrate concentration is so high  that essentially all the enzyme molecules are in their fully active ES form.   Epigenetics. The inheritance of patterns of gene expression that are  maintained from generation to generation independent of DNA’s base  sequence. This occurs, for example, through the methylation of DNA.   Epimers. Sugars that differ only by the configuration at one C atom  (excluding the anomeric carbon).   Equatorial substituent. A group that extends largely in the plane of the  ring to which it is bonded. See also axial substituent.   Equilibrium. The point in a process at which the forward and reverse  reaction rates are exactly balanced so that it undergoes no net change.   Equilibrium constant (K eq ). The ratio, at equilibrium, of the product of the  concentrations of reaction products to those of its reactants. K eq is related  to A G° for the reaction: AG 0 = —RT In K eq . Usually abbreviated K.   Erythrocyte. A red blood cell, which functions to transport 0 2 to the  tissues. It is essentially a membranous sack of hemoglobin.   Erythrocyte ghost. Membranous particles derived from erythrocytes, which  retain their original shape but are devoid of cytoplasm.   ES complex. The enzyme—substrate complex, whose formation is a key  component of the Michaelis—Menten model of enzyme action. Also  called the Michaelis complex.   Essential amino acid. An amino acid that an animal cannot synthesize and  must therefore obtain in its diet.    Essential fatty acid. A fatty acid that an animal cannot synthesize and  must therefore obtain in its diet.   Ester group. A portion of a molecule with the formula —COOR, where R  is an alkyl group.   Estrogen. A steroid that functions primarily as a female sex hormone.   Ether. An organic compound in which two C atoms are linked by an  O atom; —C—O—C—.   Eubacteria. One of the two major groups of prokaryotes (the other is  archaea).   Euchromatin. The transcriptionally active, relatively uncondensed  chromatin in a eukaryotic cell.   Eukaryote. An organism consisting of a cell (or cells) whose genetic  material is contained in a membrane-bounded nucleus.   Evolution. The gradual alteration of an organism or one of its components  through natural selection as a result of genetic changes that are passed  from parent to offspring.   Exciton transfer. A mode of decay of an energetically excited molecule, in  which electronic energy is transferred to a nearby unexcited molecule.  Also known as resonance energy transfer.   Exergonic process. A process that has an overall negative free energy  change (a spontaneous process).   Exit site (E site). The ribosomal binding site that accommodates a tRNA  molecule that has previously transferred its peptidyl group to an  incoming aminoacyl-tRNA and is ready to dissociate from the ribosome.   Exocytosis. The release outside the cell of a vesicle’s contents through the  fusion of the vesicle membrane with the plasma membrane; the opposite  of endocytosis.   Exoglycosidase. An enzyme that catalyzes the hydrolytic excision of a  monosaccharide unit from the end of a polysaccharide.   Exon. A portion of a gene that appears in both the primary and mature  mRNA transcripts. Also called an expressed sequence.   Exonuclease. An enzyme that catalyzes the hydrolytic excision of a  nucleotide residue from one end of a polynucleotide strand.   Exopeptidase. An enzyme that catalyzes the hydrolytic excision of an  amino acid residue from one end of a polypeptide chain.   Exosome. A complex containing multiple ribonucleases and other proteins  that participate in eukaryotic and archaeal RNA processing.   Expressed sequence tag (EST). A cDNA segment corresponding to a  cellular mRNA, that can be used to identify genes that are transcribed.   Expression vector. A plasmid containing the transcription and translation  control sequences required for the production of a foreign DNA gene  product (RNA or protein) in a host cell.   SF. Faraday, the electrical charge of one mole of electrons (96,485 C * mol ! ).   Fab fragment. A proteolytic fragment of an antibody molecule that  contains the antigen-binding site. See also Fc fragment.   Fat. A mixture of triacylglycerols that is solid at room temperature.   Fatty acid. A carboxylic acid with a long-chain hydrocarbon side group.   Fc fragment. A proteolytic fragment of an antibody molecule that contains  the two C-terminal domains of its two heavy chains. See also Fab  fragment.   Feedback inhibition. The inhibition of an early step in a reaction sequence  by the product of a later step.   Feedforward activation. The activation of a later step in a reaction  sequence by the product of an earlier step.   Fibrous protein. A protein characterized by a stiff, elongated conformation,  that tends to form fibers.    First-order reaction. A reaction whose rate is proportional to the  concentration of a single reactant.   Fischer convention. A system for describing the absolute configurations of  chiral molecules by relating their structures to that of D- or L-  glyceraldehyde.   Fischer projection. A graphical convention for specifying molecular  configuration in which horizontal lines represent bonds that extend  above the plane of the drawing and vertical lines represent bonds that  extend below the plane of the drawing.   5' end. The terminus of a polynucleotide whose C5' is not esterified to  another nucleotide residue.   Flippase. An enzyme that catalyzes the translocation of a membrane lipid  across a lipid bilayer (a flip-flop).   Fluid mosaic model. A model of biological membranes in which  integral membrane proteins float and diffuse laterally in a fluid  lipid bilayer.   Fluorescence. A mode of decay of an excited molecule, in which electronic  energy is emitted in the form of a photon.   Fluorescence recovery after photobleaching (FRAP). A technique for  assessing the diffusion of membrane components from the rate at which  the fluorescently labeled component moves into an area previously  bleached by an intense pulse of laser light.   Flux. (1) The rate of flow of metabolites through a metabolic pathway, /.  (2) The rate of transport per unit area.   fMet. The formylated methionine that initiates ribosomal polypeptide  synthesis in prokaryotes.   FiFo-ATPase. A multisubunit protein consisting of a proton-translocating  membrane-embedded component (F 0 ) linked to a soluble catalytic  component (F x ) that catalyzes ATP synthesis in the presence of a  protonmotive force. Also called proton-pumping ATP synthase.   Footprinting. A procedure in which the DNA sequence to which a protein  binds is identified by determining which bases are protected by the  protein from chemical or enzymatic modification.   Fractional saturation (F). The fraction of a protein’s ligand-binding sites  that are occupied by ligand. For example, Yq 2 is the fractional saturation  of a protein’s oxygen-binding sites.   Fractionation procedure. A laboratory technique for separating the  components of a mixture of molecules through differences in their  chemical and physical properties.   Frameshifit mutation. An insertion or deletion of nucleotides in DNA that  alters the sequential reading (the frame) of sets of three nucleotides  (codons) during translation.   Free energy (G). A thermodynamic quantity, G — H — TS, whose change  at constant pressure is indicative of the spontaneity of a process. For  spontaneous processes, AG < 0, whereas for a process at equilibrium,  AG = 0. Also called the Gibbs free energy.   Free energy of activation (AG*). The free energy of the transition state  minus the free energies of the reactants in a chemical reaction.   Functional group. A portion of a molecule that participates in interactions  with other substances. Common functional groups in biochemistry are  acyl, amido, amino, carbonyl, carboxyl, diphosphoryl (pyrophosphoryl),  ester, ether, hydroxyl, imino, phosphoryl, and sulfhydryl groups.   G*. The free energy of the transition state. See also free energy of  activation.   G protein. A guanine nucleotide-binding protein, most of which are  involved in signal transduction, that is inactive when it binds GDP and  active when it binds GTP. The GTPase activity of the G protein limits  its own activity. Heterotrimeric G proteins consist of three subunits,    which dissociate to form G a (to which GTP binds) and Gp 7  components on activation.   G-quartet. A cyclic tetramer of hydrogen-bonded guanine groups. Stacks of  G-quartets result from the antiparallel association of G-rich telomeric  DNA hairpin structures.   Ganglioside. A ceramide whose head group is an oligosaccharide  containing at least one sialic acid residue.   Gap junction. An intercellular channel for ions and small molecules that is  formed by protein complexes in the membranes of apposed cells.   Gastrulation. The stage of embryonic development in which cells migrate  to form a triple-layered structure.   Gates and fences model. A model for membrane structure that includes  cytoskeletal proteins that prevent or limit the free diffusion of other  membrane proteins.   Gating. The opening and closing of a transmembrane channel in response  to a signal such as mechanical stimulation, ligand binding, presence of a  signaling molecule, or change in membrane voltage.   Gel electrophoresis. A procedure in which macromolecules are separated  on the basis of charge or size by their differential migration through a  gel-like matrix under the influence of an applied electric field. In  polyacrylamide gel electrophoresis (PAGE), the matrix is cross-linked  polyacrylamide. Agarose gels are used to separate molecules of very large  masses, such as DNAs. See also SDS-PAGE and pulsed-field gel  electrophoresis.   Gel filtration chromatography. A procedure in which macromolecules are  separated on the basis of their size and shape. Also called size exclusion  or molecular sieve chromatography.   Gene. A unique sequence of nucleotides that encodes a polypeptide or  RNA; it may include nontranscribed and nontranslated sequences, some  of which have regulatory functions.   Gene cluster. A region of DNA containing multiple copies of genes,  usually tRNA or rRNA genes, whose products are required in large  amounts.   Gene duplication. An event, such as aberrant crossover, that gives rise to  two copies of a gene on the same chromosome, each of which can then  evolve independently.   Gene expression. The decoding, via transcription and translation, of the  information contained in a gene to yield a functional RNA or protein  product.   Gene knockout. A genetic engineering process that deletes or inactivates a  specific gene in an animal.   Gene product. The RNA or protein that is encoded by a gene and that is  the end point of the gene’s expression through transcription and  translation.   Gene therapy. The transfer of genetic material to the cells of an individual  in order to produce a therapeutic effect.   General acid catalysis. A catalytic mechanism in which partial proton  transfer from an acid lowers the free energy of a reaction’s transition  state.   General base catalysis. A catalytic mechanism in which partial proton  abstraction by a base lowers the free energy of a reaction’s transition  state.   General transcription factor (GTF). One of a set of eukaryotic proteins  that are required for the synthesis of all mRNAs.   Genetic anticipation. A pattern of inheritance of a genetic disease, in  which the age of onset of symptoms decreases with each generation.   Genetic code. The correspondence between the sequence of nucleotides in  a nucleic acid and the sequence of amino acids in a polypeptide; a series  of three nucleotides (a codon) specifies an amino acid.   Genome-wide association study (GWAS). A systematic examination of  genetic differences between individuals with and without a disease in  order to identify DNA sequence variations that might contribute to the  disease.    Genomic library. A set of cloned DNA fragments representing an  organism’s entire genome.   Genomics. The study of the size, organization, and gene content of  organisms’ genomes.   Globular protein. A water-soluble protein characterized by a compact,  highly folded structure.   Glucocorticoid. A steroid hormone that affects a range of metabolic  pathways and the inflammatory response.   Glucogenic amino acid. An amino acid whose degradation yields a  gluconeogenic precursor. See also ketogenic amino acid.   Glucose-alanine cycle. An interorgan metabolic pathway that transports  nitrogen to the liver in which pyruvate produced by glycolysis in the  muscles is converted to alanine and transported to the liver. There the  alanine is converted back to pyruvate and its amino group is used to  synthesize urea for excretion. The pyruvate is converted, via  gluconeogenesis, to glucose, which is returned to the muscles.   Glucose-fatty acid cycle. The downregulation of glycolysis by fatty acid  oxidation, caused by the acetyl-CoA—induced inhibition of  phosphofructokinase by citrate. Also called the Randle cycle (although it  is not actually a cycle).   interconversion of dihydroxyacetone phosphate and 3-phosphoglycerol to  transport cytosolic reducing equivalents into the mitochondria.   Glycerophospholipid. An amphiphilic lipid in which two fatty acyl groups  are attached to a glycerol-3-phosphate whose phosphate group is linked  to a polar group. Also called a phosphoglyceride.   Glycoconjugate. A molecule, such as a glycolipid or glycoprotein, that  contains covalently linked carbohydrate.   Glycoforms. Glycoproteins that differ in the sequence, location, and  number of covalently attached carbohydrates.   Glycogen. An a(l^-6) branched polymer of a(1^4)-linked glucose  residues that serves as a glucose storage molecule in animals.   Glycogen storage disease. An inherited disorder of glycogen metabolism  affecting the size and structure of glycogen molecules or their  mobilization in the muscle and/or liver.   Glycolysis. The 10-reaction pathway by which glucose is broken down to 2  pyruvate with the concomitant production of 2 ATP and the reduction  of 2 NAD + to 2 NADH.   Glycomics. The study of the structures and functions of all of a cell’s  carbohydrates, including large glycans and the small oligosaccharides of  glycoproteins.   Glycosaminoglycan. An unbranched polysaccharide consisting of  alternating residues of uronic acid and hexosamine.   Glycoside. A molecule containing a saccharide and another molecule linked  by a glycosidic bond to the anomeric carbon in the a configuration (a-  glycoside) or (3 configuration ((3-glycoside).   Glycosidic bond. The covalent linkage (acetal or ketal) between the  anomeric carbon of a saccharide and an alcohol (O-glycosidic bond) or  an amine (A^-glycosidic bond). Glycosidic bonds link the monosaccharide  residues of a polysaccharide.   Glycosylation. The attachment of carbohydrate chains to a protein or lipid  through N- or O-glycosidic linkages.   Glyoxylate pathway. A variation of the citric acid cycle in plants that  allows acetyl-CoA to be converted quantitatively to gluconeogenic  precursors.   Glyoxysome. A membrane-bounded plant organelle in which the reactions  of the glyoxylate cycle take place. It is a specialized type of peroxisome.   Golgi apparatus. A eukaryotic organelle consisting of a set of flattened  membranous sacs in which newly synthesized proteins and lipids are  modified.   Gout. A disease characterized by elevated levels of uric acid, usually the  result of impaired uric acid excretion. Its most common manifestation is  painful arthritic joint inflammation caused by the deposition of sodium  urate.   GPCR. G protein—coupled receptor, a cell-surface protein with seven  transmembrane helices that interacts with an associated G protein on  ligand binding.   GPI-linked protein. A protein that is anchored in a membrane via a  covalently linked glycosylphosphatidylinositol (GPI) group.   Gram-negative bacterium. A bacterium that does not take up Gram stain,  indicating that its cell wall is surrounded by a complex outer membrane  that excludes Gram stain.   Gram-positive bacterium. A bacterium that takes up Gram stain,  indicating that its outermost layer is a cell wall.   Grana (sing, granum). The stacked disks of the thylakoid in a chloroplast.   Group I intron. An intron in an rRNA molecule whose self-splicing reaction  requires a guanine nucleotide and generates a cyclized intron product.   Group II intron. An intron in a eukaryotic rRNA molecule whose self¬  splicing reaction does not require a free nucleotide and generates a lariat  intron product.   Growth factor. A protein hormone that stimulates the proliferation and  differentiation of its target cells.   GTPase. An enzyme that catalyzes the hydrolysis of GTP to GDP + P z .   Guide RNA (gRNA). Small RNA molecules that pair with an immature  mRNA to direct its posttranscriptional editing.   Half-reaction. The single oxidation or reduction process, involving an  electron donor and its conjugate electron acceptor, that occurs in  electrical cells but requires direct contact with another such reaction to  form a complete oxidation-reduction reaction.   Half-time (£ 1/2 ). The time required for half the reactant initially present to  undergo reaction. Also called half-life.   Haworth projection. A representation of a sugar ring in which ring bonds  that project in front of the plane of the paper are drawn as thick lines  and ring bonds that project behind the plane of the paper are drawn as  thin lines.   Helix cap. A protein structural element in which the side chain of a  residue preceding or succeeding a helix folds back to form a hydrogen  bond with the backbone of one of the helix’s four terminal residues.   Helix-turn-helix (HTH) motif. An ~ 20-residue protein motif that forms  two a helices that cross at an angle of —120°. The motif, which occurs  in numerous prokaryotic DNA-binding proteins, binds in DNA’s major  groove to specific base sequences.   Heme. A porphyrin derivative whose central Fe(II) atom is the site of  reversible oxygen binding (in myoglobin and hemoglobin) or  oxidation-reduction (in cytochromes).   Hemiacetal. The product of the reaction between an alcohol and the  carbonyl group of an aldehyde.   Hemiketal. The product of the reaction between an alcohol and the  carbonyl group of a ketone.   Hemolytic anemia. Loss of red blood cells through their lysis (destruction)  in the bloodstream.   Henderson-Hasselbalch equation. The mathematical expression of the  relationship between the pH of a solution of a weak acid and its p K\ pH  = P K+ log([A"]/[HA]).   polypeptide in which residues a and d are hydrophobic and thus line one  face of the helix, thereby inducing the formation of a coiled coil with a  second similar helix.   Heterogeneous nuclear RNA (hnRNA). Eukaryotic mRNA primary  transcripts whose introns have not yet been excised.   Heterologous DNA. A segment of DNA consisting of imperfectly  complementary strands.   Heterolytic cleavage. Cleavage of a bond in which one of two chemically  bonded atoms acquires both of the electrons that formed the bond.   Heteropolysaccharide. A polysaccharide consisting of more than one type  of monosaccharide.   Heterotroph. An organism that obtains free energy from the oxidation of  organic compounds produced by other organisms.   “High-energy” intermediate. A substance whose degradation is highly  exergonic (yields at least as much free energy as is required to synthesize  ATP from ADP + P z ; >30.5 kj • mol -1 under standard biochemical  conditions). Also called an “energy-rich” compound.   High-performance liquid chromatography (HPLC). An automated  chromatographic procedure for fractionating molecules using precisely  fabricated matrix materials and pressurized flows of precisely mixed solvents.   Highly repetitive DNA. Clusters of nearly identical sequences of up to 10 bp  that are repeated thousands of times; the sequences are present at >10 6  copies per haploid genome. Also known as short tandem repeats (STRs).   Hill coefficient. The exponent in the Hill equation. It provides a measure  of the degree of cooperative binding of a ligand to a molecule.   Hill equation. A mathematical expression for the degree of saturation of  ligand binding to a molecule with multiple binding sites as a function of  the ligand concentration.   Histone acetyltransferase (HAT). An enzyme that catalyzes the sequence-  specific acetylation of histones so as to regulate gene transcription.   Histone code. The correlation between the pattern of histone modification  and the transcriptional activity of the associated DNA.   Histones. Highly conserved basic proteins that constitute the protein core  to which DNA is bound to form a nucleosome.   HIV. Human immunodeficiency virus, the causative agent of acquired  immunodeficiency syndrome (AIDS).   HMG protein. A member of the high mobility group (HMG) of   nonhistone chromosomal proteins whose abundant charged groups give  them high electrophoretic mobility.   Holliday junction. The four-stranded structure that forms as an  intermediate in DNA recombination.   Homeodomain. An ~ 60-amino acid DNA-binding motif common to many  genes that specify the identities and fates of embryonic cells; such genes  encode transcription factors. Also called a homeobox.   Homeostasis. The maintenance of a steady state in an organism. See also  metabolic homeostasis.   Homeotic selector genes. Insect genes that specify the identities of body  segments.   Homolactic fermentation. The reduction of pyruvate to lactate with the  concomitant oxidation of NADH to NAD + .   Homologous end-joining. A pathway in which DNA with double-strand  breaks is repaired nonmutagenically through recombination with an  intact homologous chromosome.    Homologous proteins. Proteins that resemble each other due to their  evolution from a common ancestor.   Homolytic cleavage. Cleavage of a bond in which each participating atom  acquires one of the electrons that formed the bond.   Hoogsteen base pair. A form of base pairing in which thymine or uracil  atom N3 hydrogen bonds to adenine atom N7 and adenine N6 hydrogen  bonds to thymine or uracil 04. See also Watson-Crick base pair.   Hormone. A substance (e.g., a peptide or a steroid) that is secreted by one  tissue and which induces a physiological response (e.g., growth and  metabolism) in other tissues.   Hormone response element (HRE). A DNA sequence to which a  hormone—receptor complex binds so as to enhance or repress the  transcription of an associated gene.   Hormone-sensitive lipase. An adipose tissue enzyme that releases fatty  acids from triacylglycerols in response to a hormonally generated increase  in cAMP. Also known as hormone-sensitive triacylglycerol lipase.   Hox gene. A gene encoding a transcription factor that includes a  homeodomain and which specifies segmental identity.   Humoral immunity. Immunity mediated by antibodies (immunoglobulins)  produced by B lymphocytes (B cells).   Hybridization. The formation of double-stranded segments of  complementary DNA and/or RNA sequences.   Hybridoma. The cell clones produced by the fusion of an antibody-   producing lymphocyte and an immortal myeloma cell. These are the cells  that produce monoclonal antibodies.   Hydration. The molecular state of being surrounded by and interacting  with several layers of solvent water molecules, that is, solvated by water.   Hydrogen bond. A largely electrostatic interaction between a weakly acidic  donor group such as O—H or N—H and a weakly basic acceptor atom  such as O or N.   Hydrolysis. The cleavage of a covalent bond accomplished by adding the  elements of water; the reverse of a condensation.   Hydronium ion. A proton associated with a water molecule, H 3 0 + .   hydrophilicity of an amino acid residue; it is indicative of the likelihood  of finding that residue in a protein interior.   Hydrophilic substance. A substance whose high polarity allows it to  readily interact with water molecules and thereby dissolve in water.   Hydrophobic collapse. A driving force in protein folding, resulting from  the tendency of hydrophobic residues to avoid contact with water and  hence form the protein core.   Hydrophobic effect. The tendency of water to minimize its contacts with  nonpolar substances, thereby inducing the substances to aggregate.   molecules are selectively retained on a nonpolar matrix by virtue of their  hydrophobicity.   Hydrophobic substance. A substance whose nonpolar nature reduces its  ability to be solvated by water molecules. Hydrophobic substances tend  to be soluble in nonpolar solvents but not in water.   Hydroxide ion. OH , a product of the ionization of a water molecule.   Hyperammonemia. Elevated levels of ammonia in the blood, a toxic  situation.   Hyperbolic curve. The graphical representation of the mathematical equation  that describes the noncooperative binding of a ligand to a molecule or the  rate of a reaction catalyzed by a Michaelis-Menten enzyme.    Hypercholesterolemia. High levels of cholesterol in the blood, a risk factor  for heart disease. Familial hypercholesterolemia usually results from an  inherited defect in the LDL receptor.   Hyperchromic effect. The increase in duplex DNA’s ultraviolet absorbance  resulting from the loss of stacking interactions as the DNA denatures.   Hyperpolarization. The decrease of membrane potential below the  resting state that occurs during the latter portion of an action  potential in nerve cells.   Hypervariable residue. An amino acid residue occupying a position in a  protein that is occupied by many different residues among evolutionarily  related proteins. The opposite of a hypervariable residue is an invariant  residue.   Hypoxia. A condition in which the oxygen level in the blood is lower than  normal.   I-cell disease. A hereditary deficiency in a lysosomal hydrolase that causes  the accumulation of glycosaminoglycan and glycolipid inclusions in the  lysosomes.   Imaginal disk. A patch of apparently undifferentiated but developmentally  committed cells in an insect larva that ultimately gives rise to a specific  external structure in the adult. ^   Imino group. A portion of a molecule with the formula C — NH.   infection by producing antibodies and by killing pathogens and  infected host cells.   Immunoaffinity chromatography. A procedure in which a molecule is  separated from a mixture of other molecules by its ability to bind  specifically to an immobilized antibody.   Immunoassay. A procedure for detecting, and in some cases quantifying,  the activity of a macromolecule by using an antibody or mixture of  antibodies that reacts specifically with that substance.   Immunoblot. A technique in which a molecule immobilized on a  membrane filter can be detected through its ability to bind to an  antibody directed against it. A Western blot is an immunoblot to detect  an immobilized protein after electrophoresis.   Immunofluorescence microscopy. A technique in microscopy in which a  fluorescent-tagged antibody is used to reveal the presence of the antigen  to which it binds.   Immunoglobulin fold. A disulfide-linked domain consisting of a sandwich  of a three-stranded and a four-stranded antiparallel (3 sheet that occurs in  antibody molecules.   Imprinting. The differential expression of maternal and paternal genes  according to their patterns of DNA methylation.   in silico. In a computer simulation (electronic circuits are mainly  silicon-based).   Inactivator. An inhibitor that reacts irreversibly with an enzyme so as to  inactivate it.   Indirect readout. The ability of a DNA-binding protein to detect its target  base sequence though the sequence-dependent conformation and/or  flexibility of its DNA backbone rather than through direct interaction  with its bases.   Induced fit. An interaction between a protein and its ligand, which  induces a conformational change in the protein that increases the  protein’s affinity for the ligand.   Inducible enzyme. An enzyme that is synthesized only when required by  the cell. See also constitutive enzyme.   Inhibitor. A substance that reduces an enzyme’s activity by affecting its  substrate binding or turnover number.   Initiation factor. A protein that interacts with mRNA and/or the ribosome  and which is required to initiate translation.   Insertion sequence. A simple transposon that is flanked by short inverted  repeats. Also called an IS element.   Insertion/deletion mutation. A genetic change resulting from the addition  or loss of nucleotides; also called an indel.   Insulator. A segment of DNA that delimits the effective range of a  transcription-regulating element.   Insulin resistance. The decreased ability of cells to respond to insulin,  which results in a decrease in their ability to take up glucose.   Integral protein. A membrane protein that is embedded in the lipid  bilayer and can be separated from it only by treatment with agents that  disrupt membranes. Also called an intrinsic protein.   Intercalation agent. A substance, usually a planar aromatic cation, that  slips in between the stacked bases of a double-stranded polynucleotide.   Interfacial activation. The increase in activity when a lipid-specific enzyme  contacts the lipid-water interface.   Intermembrane space. The compartment between the inner and outer  mitochondrial membranes. Because of the porosity of the outer  membrane, the intermembrane space is equivalent to the cytosol in its  small-molecule composition.   Internal conversion. A mode of decay of an excited molecule, in which  electronic energy is converted to heat (the kinetic energy of molecular  motion).   Intrinsic terminator. A conserved G + C-rich hairpin followed by a  U-rich segment that causes Rho-independent transcription termination  in E. coli.   Intrinsically disordered protein. A protein whose tertiary structure  includes highly flexible extended segments that can adopt different  conformations.   Intron. A portion of a gene that is transcribed but excised prior to  translation. Also called an intervening sequence.   Invariant residue. A residue in a protein that is the same in all   evolutionarily related proteins. The opposite of an invariant residue is a  hypervariable residue.   Ion exchange chromatography. A fractionation procedure in which ions  are selectively retained by a matrix bearing oppositely charged groups.   Ion pair. An electrostatic interaction between two ionic groups of opposite  charge. In proteins, it is also called a salt bridge.   Ionophore. An organic molecule, often an antibiotic, that increases the  permeability of a membrane to a particular ion. A carrier ionophore  diffuses with its ion through the membrane, whereas a channel-forming  ionophore forms a transmembrane pore.   Iron-sulfur protein. A protein that contains a prosthetic group consisting most  commonly of equal numbers of iron and sulfur ions (i.e., [2Fe-2S] and  [4Fe-4S]) and that usually participates in oxidation-reduction reactions.   Ischemia. The blockage of blood flow to a tissue or organ.   Isoaccepting tRNA. A tRNA that carries the same amino acid as another  tRNA.   Isoelectric focusing (IEF). Electrophoresis through a stable pH gradient  such that a charged molecule migrates to a position corresponding to its  isoelectric point.   Isoelectric point (p/). The pH at which a molecule has no net charge and  hence does not migrate in an electric field.   Isolated system. A thermodynamic system that cannot exchange matter or  energy with its surroundings.   Isopeptide bond. An amide linkage between an a-carboxylate group of an  amino acid and the e-amino group of Lys, or between the a-amino group  of an amino acid and the (3- or y-carboxylate group of Asp or Glu.   Isoprenoid. A lipid consisting of five-carbon units with the same carbon  skeleton as isoprene.   Isozymes. Enzymes that catalyze the same reaction but are encoded by  different genes. Also called isoforms.   JAK-STAT pathway. A signal transduction pathway involving a JAK (Janus  kinase) protein and a STAT (signal tranducer and activator of  transcription) protein that relays cytokine-based signals.   Jaundice. A yellowing of the skin and whites of the eyes as a result of the  deposition of the heme degradation product bilirubin in those tissues. It  is a symptom of liver dysfunction, bile-duct obstruction, or a high rate  of red cell destruction.   & B . Boltzmann constant (1.38 07 X 10 -23 J • K -1 ); it is equivalent to R/N,  where R is the gas constant and N is Avogadro’s number.   k cat . The catalytic constant for an enzymatic reaction, equivalent to the  ratio of the maximal velocity (V ma3 ^) and the enzyme concentration  ([E] t ). Also called the turnover number.   k CSLt IK M ' The apparent second-order rate constant for an enzyme-catalyzed  reaction; it is a measure of an enzyme’s catalytic efficiency.   Ketogenic amino acid. An amino acid whose degradation in animals yields  compounds that can be converted to fatty acids or ketone bodies. See  also glucogenic amino acid.   Ketone bodies. Acetoacetate, D-p-hydroxybutyrate, and acetone; these  compounds are produced from acetyl-CoA by the liver for use as  metabolic fuels in peripheral tissues.   Ketosis. A potentially pathological condition in which ketone bodies are  produced in excess of their utilization.   Kinase. An enzyme that transfers a phosphoryl group between ATP and  another molecule.   phosphorylation activates the next kinase in a series, thereby amplifying  the effect of the initial kinase.   Km p The apparent (observed) Michaelis constant for an enzyme-catalyzed  reaction, which may differ from the true value due to the presence of an  enzyme inhibitor.   k-x. The rate constant for a reverse reaction, e.g., the breakdown of the ES  complex to E + S.   k 2 . The rate constant for the second step of a simple enzyme-catalyzed  reaction that follows Michaelis—Menten kinetics, that is, the conversion  of the ES complex to E + P.   Lactose intolerance. The inability to digest the disaccharide lactose due to  a deficiency of the enzyme p-galactosidase (lactase).    Lagging strand. A newly synthesized DNA strand that extends 3' —> 5' in  the direction of travel of the replication fork. This strand is synthesized  as a series of discontinuous fragments that are later joined.   Large intergenic noncoding RNA (lincRNA). Large nontranslated  eukaryotic RNAs that are transcribed by RNA polymerase II.   Lateral diffusion. The movement of a lipid within one leaflet of a bilayer.   Lead compound. A drug molecule that serves as the starting point for the  development of more effective drug molecules.   Leader sequence. (1) A nucleotide sequence that precedes the coding  region of an mRNA. (2) A signal sequence.   Leading strand. A newly synthesized DNA strand that extends 3 ' —> 3' in  the direction of travel of the replication fork. This strand is synthesized  continuously.   Le Chatelier’s principle. The phenomenon that a change in concentration,  temperature, volume, or pressure in a system at equilibrium causes the  equilibrium position to shift so as to counteract the change.   Lesch-Nyhan syndrome. A genetic disease caused by the deficiency of  hypoxanthine-guanine phosphoribosyltransferase (HGPRT), an enzyme  required for purine salvage reactions. Affected individuals produce  excessive uric acid and exhibit neurological abnormalities.   Leucine zipper. A protein structural motif in which two a helices, each  with a hydrophobic strip along one side, associate as a coiled coil. This  motif, which has a Leu at nearly every seventh residue, mediates the  association of many types of DNA-binding proteins.   Levorotatory. Rotating the plane of polarized light counterclockwise from  the point of view of the observer; the opposite of dextrorotatory.   Ligand. (1) A small molecule that binds to a larger molecule. (2) A  molecule or ion bound to a metal ion.   Ligand-gated channel. A channel whose opening and closing (gating) is  controlled by the binding of a specific molecule (ligand) Also called a  signal-gated channel.   Ligase. An enzyme that catalyzes bond formation coupled with the  hydrolysis of ATP.   Ligation. The joining together of two molecules such as two DNA  segments.   Light-harvesting complex (LHC). A pigment-containing membrane  protein that collects light energy and transfers it to a photosynthetic  reaction center.   Light reactions. The portion of photosynthesis in which specialized  pigment molecules capture light energy and are thereby oxidized.  Electrons are transferred to generate NADPH and a transmembrane  proton gradient that drives ATP synthesis.   Limited proteolysis. A technique in which a polypeptide is incompletely  digested by proteases.   Lineweaver-Burk plot. A graph of a rearrangement of the  Michaelis-Menten equation to a linear form that permits the  determination of K M and V max . Also called a double-reciprocal plot.   Linker DNA. The ~ 35-bp segment of DNA that links nucleosome core  particles in chromatin.   Linking number (Z). The number of times that one strand of a covalently  closed circular double-stranded DNA winds around the other; it cannot  be changed without breaking covalent bonds.   Lipid. Any member of a broad class of biological molecules that are largely  or wholly hydrophobic and therefore tend to be insoluble in water but  soluble in organic solvents such as hexane.   Lipid-linked protein. A protein that is anchored to a biological membrane  via a covalently attached lipid such as a farnesyl, geranylgeranyl,  myristoyl, palmitoyl, or glycosylphosphatidylinositol group.    Lipid raft. A semicrystalline region of a cell membrane containing tightly  packed glycosphingolipids and cholesterol.   Lipid storage disease. A defect in a lipid-degrading enzyme that causes the  substrate for the enzyme to accumulate in lysosomes.   Lipoprotein. A globular particle consisting of a nonpolar lipid core  surrounded by an amphiphilic coat of protein, phospholipid, and  cholesterol. Lipoproteins, which transport lipids between tissues via the  bloodstream, are classified by their density as high, low, intermediate,  and very low density lipoproteins (HDL, LDL, IDL, and VLDL).   Lipoyllysyl arm. An extended structure, consisting of lipoic acid linked to  a lysine side chain, that delivers intermediates between active sites in  multienzyme complexes such as the pyruvate dehydrogenase complex.   London dispersion forces. The weak attractive forces between electrically  neutral molecules in close proximity, which arise from electrostatic  interactions among their fluctuating dipoles.   Low-barrier hydrogen bond (LBHB). An unusually short and strong  hydrogen bond that forms when the donor and acceptor groups have  nearly equal piL values so that the hydrogen atom is equally shared  between them.   Lung surfactant. The amphipathic protein and lipid mixture that prevents  collapse of the lung alveoli (microscopic air spaces) on the expiration of air.   Lyase. An enzyme that catalyzes the elimination of a group to form a  double bond.   Lymphocyte. Types of white blood cells that mediate the immune response.  Mammalian B lymphocytes develop in the bone marrow, and T  lymphocytes in the thymus.   Lysophospholipid. A glycerophospholipid derivative lacking a fatty acyl  group at position C2 whose detergentlike properties can disrupt cell  membranes.   Lysosome. A membrane-bounded organelle in a eukaryotic cell that  contains a battery of hydrolytic enzymes and which functions to digest  ingested material and to recycle cell components.   Macronutrient. A nutrient that is required in relatively large amounts, such  as proteins, carbohydrates, and fats. See also micronutrient.   Major groove. The groove on a DNA double helix whose bounding  glycosidic bonds from each base pair form an angle of >180°. In  B-DNA, this groove is wider than the minor groove.   Malaria. A mosquito-borne disease caused by protozoa of the genus  Plasmodium , most notably Plasmodium falciparum, which reside in red  blood cells during much of their life cycle.   Malate-aspartate shuttle. A metabolic circuit that uses the malate and  aspartate transporters and the interconversion of malate, oxaloacetate,  and aspartate to ferry reducing equivalents into the mitochondrion.   Malignant tumor. A mass of cells that proliferate uncontrollably; a cancer.   Mass spectrometry. A technique for identifying molecules by measuring  the mass-to-charge ratios of molecular ions in the gas phase.   Maternal-effect genes. Insect genes whose mRNA or protein products are  deposited by the mother in the ovum and which define the polarity of  the embryonic body.   Matrix. The gel-like solution of enzymes, substrates, cofactors, and ions in  the interior of the mitochondrion.   Mechanism-based inhibitor. A molecule that chemically inactivates an  enzyme only after undergoing part or all of its normal catalytic reaction.  Also called a suicide substrate.   Mechanosensitive channel. A channel whose opening and closing (gating)  is controlled by mechanical stimuli such as touch, sound, and changes in  osmotic pressure.   Mediated transport. The transmembrane movement of a substance through  the action of a specific carrier protein; the opposite of nonmediated  transport.   Melting temperature (7^). The midpoint temperature of the melting  curve for the thermal denaturation of a macromolecule.    Membrane-enveloped virus. A virus produced by budding from the  surface of a host cell such that the viral particle is surrounded by a  membrane derived from the host cell.   Memory B cell. A B cell that can recognize its corresponding antigen and  rapidly proliferate to produce specific antibodies weeks to years after the  antigen was first encountered.   Messenger RNA (mRNA). A ribonucleic acid whose sequence is  complementary to that of a protein-coding gene in DNA. In the  ribosome, mRNA directs the polymerization of amino acids to form a  polypeptide with the corresponding sequence.   Metabolic fuel. A molecule that can be oxidized to provide free energy for  an organism.   Metabolic homeostasis. The ability of an organism to maintain a balance  between energy inflow and output.   Metabolic syndrome. An obesity-related disorder that includes insulin  resistance, hypertension, and atherosclerosis.   Metabolomics. The study of all the metabolites produced by a cell under a  given set of conditions, including their concentrations and functions.   Metagenomic sequencing. The analysis of all the DNA sequences in a  sample containing multiple organisms.   Metal chelate affinity chromatography. A procedure in which a molecule  bearing metal-chelating groups is separated from a mixture of other  molecules by its ability to bind to metal ions attached to a  chromatographic matrix.   Metal ion catalysis. A catalytic mechanism that requires the presence of a  metal ion to lower the free energy of a reaction’s transition state.   Metalloenzyme. An enzyme that contains a tightly bound metal ion  cofactor, typically a transition metal ion such as Fe 2+ , Zn 2+ , or Mn 2+ .   Micelle. A globular aggregate of amphiphilic molecules in aqueous solution  that are oriented such that polar segments form the surface of the  aggregate and the nonpolar segments form a core that is out of contact  with the solvent.   Michaelis constant (K M ). For an enzyme that follows the   Michaelis-Menten model, K M = ( k_\ + k-flk i; K M is equal to the  substrate concentration at which the reaction velocity is half-maximal.   Michaelis-Menten equation. A mathematical expression that describes the  initial rate of an enzymatic reaction  in terms of the substrate  concentration ([S]), the enzymes maximal velocity (V^), and its  Michaelis constant ( K M ): v 0 = V mzx [S]I(K M + [S]).   Micro RNA (miRNA). An endogenous RNA of ~22 nucleotides that  inhibits gene expression through RNA interference. See also short  interfering RNA (siRNA).   Microbiome. The collection of microorganisms that live in or on the  human body.   Micronutrient. A nutrient that is required in relatively small amounts, such  as vitamins and minerals. See also macronutrient.   Mineral. An inorganic substance required for metabolic activity, including  sodium, potassium, chloride, and calcium. Minerals such as iron, copper,  and zinc, which are required in small amounts, are known as trace  elements.   Mineralocorticoid. A steroid hormone that regulates the excretion of salt  and water by the kidneys.   Minor groove. The groove on a DNA double helix whose bounding  glycosidic bonds from each base pair form an angle of <180°. In  B-DNA, this groove is narrower than the major groove.    end. The end of a polymeric fdament where growth is slower. See also   end.   Mismatch repair (MMR). A postreplication process, in which mispaired  nucleotides are excised and replaced, that distinguishes between the  parental (correct) and daughter (incorrect) strands of DNA.   Mitochondria {sing, mitochondrion). The double-membrane-enveloped  eukaryotic organelles in which aerobic metabolic reactions occur,  including those of the citric acid cycle, fatty acid oxidation, and  oxidative phosphorylation.   Mixed inhibition. A form of enzyme inhibition in which an inhibitor binds  to both the enzyme and the enzyme-substrate complex and thereby  differently affects K M and V^. Also called noncompetitive inhibition.   Moderately repetitive DNA. Segments of hundreds to thousands of base  pairs that are present at <10 6 copies per haploid genome.   Modification methylase. A bacterial enzyme that methylates a specific  sequence of DNA as part of a restriction-modification system.   Molecular chaperone. A protein that binds to unfolded or misfolded  proteins so as to promote normal folding and the formation of native  quaternary structure. Also known as a heat shock protein (Hsp).   Molecularity. The number of molecules that participate in an elementary  chemical reaction.   Molten globule. A collapsed but conformationally mobile intermediate in  protein folding that has much of the native protein’s secondary structure  but little of its tertiary structure.   Monoclonal antibody. A single species of antibody molecule produced by  a clone of hybridoma cells, which are derived by the fusion of a  myeloma cell with a lymphocyte producing that antibody.   Monomer. (1) A structural unit from which a polymer is built up. (2) A  single subunit or protomer of a multisubunit protein.   Morphogen. A substance whose distribution in an embryo directs, in part,  the embryo’s developmental pattern.   Motor protein. An intracellular protein that couples the free energy of  ATP hydrolysis to molecular movement relative to another protein that  often acts as a track for the linear movement of the motor protein.   M r Relative molecular mass. A dimensionless quantity that is defined as the  ratio of the mass of a particle to 1/12th the mass of a 12 C atom. Also  known as molecular weight. It is numerically equal to the grams/mole of  a compound.   Multienzyme complex. A group of noncovalently associated enzymes that  catalyze two or more sequential steps in a metabolic pathway.   Multiple myeloma. A disease in which a cancerous B cell proliferates and  produces massive quantities of a single antibody known as a myeloma  protein.   Multisubunit protein. A protein consisting of more than one polypeptide  chain (subunit).   Mutase. An enzyme that catalyzes the transfer of a functional group from  one position to another on a molecule.   Myocardial infarction. The death of heart tissue caused by the loss of  blood supply (a heart attack).   Myofibril. The bundle of fibers that are arranged in register in striated  muscle cells.   Myristoylation. The attachment of a myristoyl group to a protein to form  a lipid-linked protein.    N-end rule. The correlation between the identity of a polypeptide’s  N-terminal residue and its half-life in the cell.   AHinked oligosaccharide. An oligosaccharide linked via a glycosidic bond  to the amide group of a protein Asn residue in the sequence Asn-X-  Ser/Thr.   Natural selection. The evolutionary process by which the continued  existence of a replicating entity depends on its ability to survive and  reproduce under the existing conditions.   Near-equilibrium reaction. A reaction whose A G value is close to zero, so  that it can operate in either direction depending on the substrate and  product concentrations.   disintegration of the cell and the release of proinflammatory substances.  See also apoptosis.   Nernst equation. An expression of the relationship between reduction  potential difference (A^) and the concentrations of the electron  donors and acceptors (A, B):   \%=\%°-RTIn® In ([A red ][B ox ]/[A ox ][B red ]).   Neurotransmitter. A substance released by a nerve cell that alters the  activity of a neighboring nerve cell.   Neutral drift. Evolutionary changes that become fixed at random rather  than through natural selection.   Neutral solution. A solution whose pH is equal to 7.0 ([H + ] = 10 -7 M).   Nick translation. The progressive movement of a single-strand break (nick)  in duplex DNA through the coordinated actions of a 5' —» 3'  exonuclease function that removes residues from the 5' side of the break  and a polymerase function that adds residues to the 3 ’ side.   Nitrogen assimilation. The incorporation of fixed nitrogen (e.g., ammonia)  into a biological molecule such as an amino acid.   Nitrogen cycle. The series of reactions in which N 2 and ammonia are  interconverted, often via nitrate and nitrite, by various organisms.   Nitrogen fixation. The process by which atmospheric N 2 is converted to a  biologically useful form such as NH 3 .   Noncoding RNA (ncRNA). An RNA molecule, such as rRNA, tRNA, or  another small RNA, that is not translated.   Noncompetitive inhibition. (1) A synonym for mixed inhibition. (2) A  special case of mixed inhibition in which the inhibitor binds the enzyme  and enzyme-substrate complex with equal affinities (A[ = K{), thereby  reducing the apparent value of but leaving K M unchanged.   Noncooperative binding. A situation in which binding of a ligand to a  macromolecule does not affect the affinities of other binding sites on the  same molecule.   Nonessential amino acid. An amino acid that animals can synthesize from  common intermediates.   Nonhomologous end-joining (NHEJ). An error-prone pathway for  repairing DNA with double-strand breaks.   Nonmediated transport. The transmembrane movement of a substance  through simple diffusion; the opposite of mediated transport.   Nonpolar molecule. A molecule that lacks a group with a permanent  dipole.   Nonreceptor tyrosine kinase. An intracellular tyrosine kinase that is  indirectly activated by ligand binding to a receptor.   Nonrepetitive structure. A segment of a polymer in which the backbone  has an ordered arrangement that is not characterized by a repeating  conformation.   Nonsense-mediated decay (NMD). The degradation of an mRNA that  contains a premature Stop codon.   Nonsense mutation. A mutation that converts a codon that specifies an  amino acid to a Stop codon, thereby causing the premature termination  of translation.   Nonsense suppressor tRNA. A mutated tRNA that recognizes a Stop  codon so that its attached aminoacyl group is appended to the  polypeptide chain; it mitigates the effect of a nonsense mutation in a  structural gene.   Northern blotting. A procedure for identifying an RNA containing a  particular base sequence through its ability to hybridize with a  complementary single-stranded segment of DNA or RNA. See also  Southern blotting.   Nuclear magnetic resonance (NMR). A spectroscopic method for  characterizing atomic and molecular properties based on the signals  emitted by radiofrequency-excited atomic nuclei in a magnetic field. It  can be used to determine the three-dimensional molecular structure of a  protein or nucleic acid.   Nucleic acid. A polymer of nucleotide residues. The major nucleic acids are  deoxyribonucleic acid (DNA) and ribonucleic acid (RNA). Also known  as a polynucleotide.   nucleus, where ribosomal RNA (rRNA) is transcribed and processed and  where ribosomes are assembled.   Nucleophile. A group that contains unshared electron pairs that readily  reacts with an electron-deficient group (electrophile). A nucleophile  (nucleus-lover) reacts with an electrophile (electron-lover).   Nucleoside. A compound consisting of a nitrogenous base and a five-  carbon sugar (ribose or deoxyribose) in A^-glycosidic linkage.   Nucleosome. The complex of a histone octamer and ~200 bp of DNA  that forms the lowest level of DNA organization in the eukaryotic  chromosome.   Nucleosome core particle. The complex of histones and —146 bp of DNA  that forms a compact disk-shaped particle in which the DNA is wound  in ~2 helical turns around the outside of the histone octamer.   Nucleotide. A compound consisting of a nucleoside esterified to one or  more phosphate groups. Nucleotides are the monomeric units of nucleic  acids.   Nucleotide excision repair (NER). A multistep process in which a portion  of DNA containing a lesion is excised and replaced by normal DNA.   Nucleotide sugar. A saccharide linked to a nucleotide by a phosphoester  bond, the cleavage of which drives the formation of a glycosidic  bond.   Nucleus. The membrane-enveloped organelle in which the eukaryotic cell’s  genetic material is located.   Nutrition. The intake and utilization of food as a source of raw materials  and free energy.   O-linked oligosaccharide. An oligosaccharide linked via a glycosidic bond  to the hydroxyl group of a protein Ser or Thr side chain.   Oil. A mixture of triacylglycerols that is liquid at room temperature.   Okazaki fragments. The short segments of DNA formed in the  discontinuous lagging-strand synthesis of DNA.   Oligomer. (1) A short polymer consisting of a few linked monomer units.  (2) A protein consisting of a few protomers (subunits).   Oligosaccharide processing. The cellular pathway in which a newly  glycosylated protein undergoes the enzymatic removal and addition of  monosaccharide residues.   Oncogene. A mutant version of a normal gene (a proto-oncogene), which  may be acquired through viral infection; it interferes with the  mechanisms that normally control cell growth and differentiation and  thereby contributes to uncontrolled proliferation (cancer).   Open complex. The separated DNA strands at the transcription start site.   Open reading frame (ORF). A portion of the genome that potentially  codes for a protein. This sequence of nucleotides begins with a Start  codon, ends with a Stop codon, contains no internal Stop codons, is  flanked by the proper control sequences, and exhibits the same codon-  usage preference as other genes in the organism.   Open system. A thermodynamic system that can exchange matter and  energy with its surroundings.   Operator. A DNA sequence at or near the transcription start site of a gene,  to which a repressor binds so as to control transcription of the gene.   Operon. A prokaryotic genetic unit that consists of several genes with  related functions that are transcribed as a single mRNA molecule.   Optical activity. The ability of a molecule to rotate the plane of polarized  light. Molecules with optical activity are chiral.   Ordered mechanism. A sequential reaction with a compulsory order of  substrate addition to the enzyme.   Organelle. A differentiated structure within a eukaryotic cell, such as a  mitochondrion, ribosome, or lysosome, that performs specific functions.   Orphan gene. A gene, usually identified through genome sequencing, with  no known function.   Orthologous genes. Evolutionarily related genes in different species that  have the same function.   Orthophosphate cleavage. The hydrolysis of ATP that yields ADP + P z .   Osmosis. The movement of solvent across a semipermeable membrane from a  region of low solute concentration to a region of high solute concentration.   Osmotic pressure. The pressure that must be applied to a solution  containing a high concentration of solute to prevent the net flow of  solvent across a semipermeable membrane separating it from a solution  with a lower concentration of solute. The osmotic pressure of a 1 M  solution of any solute separated from solvent by a semipermeable  membrane is ideally 22.4 atm.   Overproducer. A genetically engineered organism that produces massive  quantities of a foreign DNA gene product.   Oxidation. The loss of electrons. Oxidation of a substance is accompanied  by the reduction of another substance.   Oxidative phosphorylation. The process by which the free energy  obtained from the oxidation of metabolic fuels is used to generate  ATP from ADP + P z ,   Oxidizing agent. A substance that can accept electrons from other  substances, thereby oxidizing them and becoming reduced.   Oxonium ion. A resonance-stabilized and oxygen-containing carbocation  such as occurs during the lysozyme-catalyzed hydrolysis of a glycoside.   Oxyanion hole. A structure in an enzyme active site that preferentially  binds and thereby stabilizes the oxyanionic tetrahedral transition state of  the reaction.   Oxygen debt. The postexertion continued elevation in 0 2 consumption  that is required to replenish the ATP consumed by the liver during  operation of the Cori cycle.   Oxygenation. The binding of molecular oxygen, e.g., to a heme group.   P body. A complex of proteins that stores or degrades eukaryotic mRNAs.   P/O ratio. The ratio of the number of molecules of ATP synthesized from  ADP + P z to the number of atoms of oxygen reduced.   Palindrome. A word or phrase or a nucleotide sequence that reads the  same forward or backward.   Palmitoylation. The attachment of a palmitoyl group to a protein to form  a lipid-linked protein.   Paralogous genes. Related genes in the same organism derived from a gene  duplication event.   Partial oxygen pressure (p0 2 ). The concentration of gaseous 0 2 in units  of pressure (e.g., torr).   Passive-mediated transport. The thermodynamically spontaneous carrier-  mediated transmembrane movement of a substance from high to low  concentration. Also called facilitated diffusion.   Pasteur effect. The greatly increased sugar consumption of yeast grown  under anaerobic conditions compared to that of yeast grown under  aerobic conditions.   Pellagra. The vitamin deficiency disease resulting from a lack of niacin  (nicotinic acid) or nicotinamide, precursors of the nicotinamide-  containing cofactors NAD + and NADP + .   Pentose phosphate pathway. A pathway for glucose degradation that yields  ribose-5-phosphate and NADPH. Also called the hexose monophosphate  shunt.   Peptidase. An enzyme that hydrolyzes peptide bonds. Also called a protease.   Peptide bond. An amide linkage between the a-amino group of one amino  acid and the a-carboxylate group of another. Peptide bonds link the  amino acid residues in a polypeptide.   Peptidoglycans. The cross-linked bag-shaped macromolecules consisting of  polysaccharide and polypeptide chains that form bacterial cell walls.   Peptidyl site (P site). The ribosomal site that accommodates a tRNA with  an attached peptidyl group during protein synthesis.   Peptidyl transferase. The catalytic activity of the ribosome, which carries  out peptide bond synthesis by promoting the nucleophilic attack of an  incoming aminoacyl group on the growing peptidyl group.   Peptidyl-tRNA. The covalent complex between a tRNA molecule and a  growing polypeptide chain during protein synthesis.   Peripheral protein. A protein that is weakly associated with the surface of  a biological membrane. Also called an extrinsic protein.   Periplasmic compartment. The space between the inner membrane and  the outer membrane of gram-negative bacteria.   Perutz mechanism. A model for the cooperative binding of oxygen to  hemoglobin, in which 0 2 binding causes the protein to shift  conformation from the deoxy (T state) to the oxy (R state).   /> 50 . For a gaseous ligand, the ligand concentration, in units of pressure  (e.g., torr), at which a binding protein such as hemoglobin is half-  saturated with ligand.   pH. A quantity used to express the acidity of a solution, equal to  — log[H + ].   Pharmacogenomics. The study of how an individual’s genetic makeup  influences a drug’s effectiveness.   Pharmacokinetics. The behavior of a drug in the body over time,   (f> (phi). The torsion angle that describes the rotation around the C a —N  bond in a peptide group; the dihedral angle made by the bonds  connecting the C—N—C a —C atoms in a peptide chain.   Phosphagen. A phosphoguanidine whose phosphoryl group-transfer  potential is greater than that of ATP; these compounds can therefore  phosphorylate ADP to generate ATP.   Phosphatase. An enzyme that hydrolyzes phosphoryl ester groups. See also  protein phosphatase.   Phosphatidic acid. The simplest glycerophospholipid, consisting of two  fatty acyl groups attached to glycerol-3-phosphate.    Phosphodiester bond. The linkage in which a phosphate group is esterified  to two alcohol groups, e.g., the phosphate groups that join the adjacent  nucleoside residues in a polynucleotide.   Phosphoinositide pathway. A signal transduction pathway in which  hormone binding to a cell-surface receptor induces phospholipase C to  catalyze the hydrolysis of phosphatidylinositol-4,5-bisphosphate (PIP 2 ),  which yields inositol-1,4,3-trisphosphate (IP 3 ) and 1,2-diacylglycerol  (DAG), both of which are second messengers.   Phospholipase. An enzyme that hydrolyzes one or more bonds of a  glycerophospholipid.   Phosphorolysis. The cleavage of a chemical bond by the substitution of a  phosphate group rather than water.   Phosphoryl group. A portion of a molecule with the formula —P0 3 H 2 .   Phosphoryl group-transfer potential. A measure of the tendency of a  phosphorylated compound to transfer its phosphoryl group to water; the  opposite of its free energy of hydrolysis.   Photon. A packet (quantum) of light energy. See also Planck’s law.   Photooxidation. A mode of decay of an excited molecule, in which  oxidation occurs through the transfer of an electron to an acceptor  molecule.   Photophosphorylation. The synthesis of ATP from ADP + P z coupled to  the dissipation of a proton gradient that has been generated through  light-driven electron transport.   Photoreactivation. The conversion of pyrimidine dimers, a form of DNA  damage, to monomers using light energy.   Photorespiration. The consumption of 0 2 and evolution of C0 2 by plants  (a dissipation of the products of photosynthesis), resulting from the  competition between 0 2 and C0 2 for binding to ribulose bisphosphate  carboxylase.   Photosynthesis. The reduction of C0 2 to (CH 2 0)^ in plants and bacteria  as driven by light energy.   Photosynthetic reaction center. The pigment-containing protein complex  that undergoes photooxidation during the light reactions of  photosynthesis.   Phylogenetic tree. A reconstruction of the probable paths of evolution of a  set of related organisms, usually based on sequence variations among  their homologous proteins and nucleic acids; a sort of family tree.   Ping Pong reaction. A group-transfer reaction in which one or more  products are released before all substrates have bound to the enzyme.   Pitch. The distance a helix rises along its axis per turn; 3.4 A for an a  helix, 34 A for B-DNA.   pKA quantity used to express the tendency for an acid to donate a proton  (dissociate); equal to — log K, where Kis the acid’s dissociation constant.  Also known as p K a .   Planck’s law. An expression for the energy ( E) of a photon: E = hc/\ =  hv, where c is the speed of light, X is its wavelength, v is its frequency,  and h is Planck’s constant (6.626 X 10 34 J • s).   Plaque. (1) A region of lysed cells on a “lawn” of cultured bacteria, which  indicates the presence of infectious bacteriophage. (2) A deposit of  insoluble material in an animal’s tissues.   Plasmalogen. A glycerophospholipid in which the Cl substituent is  attached via an ether rather than an ester linkage.   Plasmid. A small circular DNA molecule that autonomously replicates in a  bacterial or yeast cell. Plasmids are often modified for use as cloning  vectors.   Point mutation. The substitution of one base for another in DNA. Point  mutations may arise from mispairing during DNA replication or from  chemical alterations of existing bases.   Polar molecule. A molecule with one or more groups that have permanent  dipoles.   Polarimeter. A device that measures the optical rotation of a solution. It  can be used to determine the optical activity of a substance.   Poly(A) tail. The sequence of adenylate residues that is posttranscriptionally  appended to the 3' end of eukaryotic mRNAs.   Polycistronic mRNA. The RNA transcript of a bacterial operon. It encodes  several polypeptides.   modification of 2- to 4-carbon units, catalyzed by an enzyme containing  multiple active sites arranged in modular fashion.   Polymer. A molecule consisting of numerous smaller units that are linked  together in an organized manner. Polymers may be linear or branched  and may consist of one or more kinds of structural units (monomers).   Polymerase. An enzyme that catalyzes the addition of nucleotide residues  to a polynucleotide through nucleophilic attack of the chains 3 r -OH  group on the a-phosphoryl group of the incoming nucleoside  triphosphate. DNA- and RNA-directed polymerases require a template  molecule with which the incoming nucleotide must base pair.   Polymerase chain reaction (PCR). A procedure for amplifying a segment  of DNA by repeated rounds of replication centered between primers that  hybridize with the two ends of the DNA segment of interest.   Polymorphism. A variation in DNA or amino acid sequences between  individuals.   Polypeptide. A polymer consisting of amino acid residues linked in linear  fashion by peptide bonds.   Polyprotic acid. A substance with more than one proton that can ionize.  Polyprotic acids have multiple ionization states.   Polyribosome. An mRNA transcript bearing multiple ribosomes in the  process of carrying out translation. Also called a polysome.   Polysaccharide. A polymeric carbohydrate consisting of multiple  monosaccharide residues. Also called a glycan.   Polyunsaturated fatty acid. A fatty acid that contains more than one  double bond in its hydrocarbon chain.   Porphyrias. Genetic defects in heme biosynthesis that result in the  accumulation of porphyrins.   Posttranscriptional modification. The removal or addition of nucleotide  residues or their modification following the synthesis of RNA.   Posttranslational processing. The removal or derivatization of amino acid  residues following their incorporation into a polypeptide, or the cleavage  of a polypeptide.   Prebiotic era. The period of time between the formation of the earth  ~4.6 billion years ago and the appearance of living organisms at least  3.3 billion years ago.   Precursor. An entity that gives rise, through a process such as evolution or  chemical reaction, to another entity.   Preinitiation complex (PIC). An assembly of eukaryotic transcription  factors bound to DNA that renders the DNA available for transcription  by RNA polymerase.   Prenylation. The attachment of an isoprenoid group to a protein to form a  lipid-linked protein.   Preproprotein. A protein bearing both a signal peptide (preprotein) and a  propeptide (proprotein).   Preprotein. A protein bearing a signal peptide that is cleaved off following  the translocation of the protein through the endoplasmic reticulum  membrane.   Pribnow box. The prokaryotic promoter element with the consensus  sequence TATAAT that is centered at around the — 10 position relative  to the transcription start site.   Primary active transport. Transmembrane transport that is driven by the  exergonic hydrolysis of ATP.   Primary transcript. The immediate product of transcription, which may be  modified before becoming fully functional.   Primase. The RNA polymerase responsible for synthesizing the RNA  segment that primes DNA synthesis.   Primer. An oligonucleotide that serves as a starting point for additional  polymerization reactions catalyzed by DNA polymerase to form a  polynucleotide. A primer base pairs with a segment of a template  polynucleotide strand so as to form a short double-stranded segment that  can then be extended through template-directed polymerization.   Primosome. The protein complex that synthesizes the RNA primers in  DNA synthesis.   Prion. A protein whose misfolding causes it to aggregate and produce the  neurodegenerative symptoms of transmissible spongiform  encephalopathies and related diseases. Prions act as infectious agents by  inducing their corresponding properly folded proteins to misfold and  thereby form more prions.   Probe. A labeled single-stranded DNA or RNA segment that can hybridize  with a DNA or RNA of interest in a screening procedure.   Prochirality. A property of certain nonchiral molecules such that they  contain a group whose substitution by another group yields a chiral  molecule.   Product inhibition. A case of enzyme inhibition in which product that  accumulates during the course of the reaction competes with substrate  for binding to the active site.   Prokaryote. A unicellular organism that lacks a membrane-bounded  nucleus. All bacteria are prokaryotes.   Promoter. The DNA sequence at which RNA polymerase binds to initiate  transcription.   Proofreading. An additional catalytic activity of an enzyme, which acts to  correct errors made by the primary enzymatic activity.   Propeptide. A polypeptide segment of an immature protein that must be  proteolytically excised to activate the protein.   Proprotein. The inactive precursor of a protein that, to become fully  active, must undergo limited proteolysis to excise its propeptide.   Prosthetic group. A cofactor that is permanently (often covalently)  associated with an enzyme.   Proteasome. A multiprotein complex with a hollow cylindrical core in  which polyubiquitin-tagged cellular proteins are degraded to peptides  (recycled) in an ATP-dependent process.   Protein. A macromolecule that consists of one or more polypeptide chains.   Protein kinase. An enzyme that catalyzes the transfer of a phosphoryl  group from ATP to the OH group of a protein Ser, Thr, or Tyr residue.   Protein phosphatase. An enzyme that catalyzes the hydrolytic excision of  phosphoryl groups from proteins.   Proteomics. The study of all of a cell’s proteins, including their   Proto-oncogene. The normal cellular analog of an oncogene; the mutation  of a proto-oncogene may yield an oncogene that contributes to  uncontrolled cell proliferation (cancer).   Protomer. One of two or more identical units of an oligomeric protein. A  protomer may consist of one or more polypeptide chains.   Proton jumping. The sequential transfer of protons between hydrogen-  bonded water molecules. Proton jumping is largely responsible for the  rapid rate at which hydronium and hydroxyl ions appear to move  through an aqueous solution.   Proton wire. A group of hydrogen-bonded protein groups and water  molecules that serves as a conduit for protons to traverse a  transmembrane protein via proton jumping.   Protonmotive force (pmf). The free energy of the electrochemical proton  gradient that forms during electron transport.   Proximity effect. A catalytic mechanism in which a reaction’s free energy of  activation is reduced by the prior bringing together of its reacting  groups.   PRPP. 5-Phosphoribosyl-a-pyrophosphate, an “activated” form of ribose  that serves as a precursor in the synthesis of histidine, tyrosine, and  purine and pyrimidine nucleotides.   Pseudo-first-order reaction. A bimolecular reaction whose rate appears to  be proportional to the concentration of only a single reactant because the  second reactant is present in large excess.   Pseudogene. An unexpressed sequence of DNA that is apparently the  defective remnant of a duplicated gene.   i|i (psi). The torsion angle that describes the rotational position around the  C a —C bond in a peptide group; the dihedral angle made by the bonds  connecting the N—C a —C—N atoms in a peptide chain.   PSI. Photosystem I, the protein complex that reduces NADP + during the  light reactions of photosynthesis.   PSII. Photosystem II, the protein complex that oxidizes H 2 0 to 0 2 during  the light reactions of photosynthesis.   Pulse-labeling. A technique for tracing metabolic fates, in which cells or a  reacting system are exposed briefly to high levels of a labeled compound.   procedure in which electrodes arrayed around the periphery of an  agarose slab gel are sequentially pulsed so that DNA molecules must  continually reorient, thereby allowing very large molecules to be  separated by size.   Purine nucleotide cycle. The conversion of aspartate to fumarate, which  replenishes citric acid cycle intermediates, through the deamination of  AMP to IMP.   heterocyclic compound. Adenine and guanine, two of the nitrogenous  bases of nucleotides, are purines.   Pyrimidine dimer. The cyclobutane-containing structure resulting from UV  irradiation of adjacent thymine or cytosine residues in the same strand of  DNA.   Pyrimidines. Derivatives of the compound pyrimidine, a planar aromatic,  heterocyclic compound. Cytosine, uracil, and thymine, three of the  nitrogenous bases of nucleotides, are pyrimidines.   Pyrophosphate cleavage. The hydrolysis of ATP that yields AMP + PP Z .   Pyrosequencing. A procedure for determining the sequence of nucleotides  in DNA by copying a DNA strand in such a way that a flash of light is  generated when the correct nucleotide is incorporated.   Q cycle. The cyclic flow of electrons accompanied by the transport of  protons, involving a stable semiquinone intermediate of CoQ in  Complex III of mitochondrial electron transport and in photosynthetic  electron transport.   q P , The thermodynamic term for heat absorbed at constant pressure.   Quantum yield. The ratio of molecules reacted to photons absorbed in a  light-induced reaction.    R group. A symbol for a variable portion of an organic molecule, such as  the side chain of an amino acid.   R state. One of two conformations of an allosteric protein; the other is the  T state. The R state is customarily the catalytically more active state.   Racemic mixture. A sample of a compound in which both enantiomers are  present in equal amounts.   Radioimmunoassay (RIA). A technique for measuring the concentration of  a molecule based on its ability to block the binding of a small amount  of the radioactively labeled molecule to its corresponding antibody.   Ramachandran diagram. A plot of cf> versus i|i that indicates the sterically  allowed conformations of a polypeptide.   Random mechanism. A sequential reaction without a compulsory order of  substrate addition to the enzyme.   Rate constant ( k ). The proportionality constant between the velocity of a  chemical reaction and the concentration(s) of the reactant(s).   Rate-determining step. The step with the highest transition state free  energy in a multistep reaction; the slowest step.   Rate enhancement. The ratio of the rates of a catalyzed to an uncatalyzed  chemical reaction.   Rate equation. A mathematical expression for the time-dependent progress  of a reaction as a function of reactant concentration.   Reaction coordinate. The path of minimum free energy for the progress of  a reaction.   Reaction order. The sum of the exponents of the concentration terms that  appear in a reaction’s rate equation.   Reactive oxygen species (ROS). Partially reduced forms of 0 2 , such as  superoxide (0 2 -) and the hydroxyl radical (-OH), that react with and  therefore damage cellular components.   Reading frame. The grouping of nucleotides in sets of three whose  sequence corresponds to a polypeptide sequence.   Receptor. A binding protein that is specific for its ligand and elicits a  discrete biochemical effect when its ligand is bound.   Receptor-mediated endocytosis. A process in which an extracellular ligand  binds to a specific cell-surface receptor and the resulting receptor-ligand  complex is engulfed by the cell.   Receptor tyrosine kinase. A hormone receptor whose intracellular domain  is activated, as a result of hormone binding, to phosphorylate tyrosine  residues on other proteins and/or on other subunits of the same receptor.   Recombinant. A DNA molecule constructed by combining DNA from  different sources. Also called a chimera.   modification of specific DNA sequences. Also called molecular cloning  or genetic engineering.   Recombination. The exchange of polynucleotide strands between separate  DNA segments. Homologous recombination occurs between DNA  segments with extensive homology, whereas site-specific recombination  occurs between two short, specific DNA sequences.   Recombination repair. A mechanism for repairing damaged DNA, in  which recombination exchanges a portion of a damaged strand for a  homologous segment that can then serve as a template for the  replacement of the damaged bases.   Reducing agent. A substance that can donate electrons, thereby reducing  another substance and becoming oxidized.   Reducing equivalent. A term used to describe the number of electrons that  are transferred from one molecule to another during a redox reaction.   Reducing sugar. A saccharide bearing an anomeric carbon that has not  formed a glycosidic bond and can therefore reduce mild oxidizing agents.   Reduction. The gain of electrons. Reduction of a substance is accompanied  by the oxidation of another substance.    Reduction potential . A measure of the tendency of a substance to gain  electrons.   Regular secondary structure. A segment of a polymer in which the  backbone adopts a regularly repeating conformation.   Release factor. A protein that recognizes a Stop codon and thereby helps  induce ribosomes to terminate polypeptide synthesis.   Renaturation. The refolding of a denatured macromolecule so as to regain  its native conformation.   Repetitive DNA. Stretches of DNA of up to several thousand bases that  occur in multiple copies in an organism’s genome; they are often  arranged in tandem.   Replica plating. The transfer of yeast colonies, bacterial colonies, or phage  plaques from a culture plate to another culture plate, a membrane, or a  filter in a manner that preserves the distribution of the cells on the  original plate.   Replication. The process of making an identical copy of a DNA molecule.  During DNA replication, the parental polynucleotide strands separate so  that each can direct the synthesis of a complementary daughter strand,  resulting in two complete DNA double helices.   Replication fork. The branch point in a replicating DNA molecule at  which the two strands of the parental molecule are separated and serve as  templates for the synthesis of the daughter strands.   Replicon. A unit of eukaryotic DNA that is replicated from one replication  origin.   Replisome. The DNA polymerase—containing protein assembly that  catalyzes the synthesis of both the leading and lagging strands of DNA  at the replication fork.   Repolarization. The recovery of membrane potential that occurs during  electrical signaling in cells such as neurons.   Repressor. A protein that binds at or near a gene so as to prevent its  transcription.   Respiratory distress syndrome. Difficulty in breathing in prematurely born  infants, caused by alveolar collapse resulting from insufficient synthesis of  lung surfactant.   Restriction endonuclease. A bacterial enzyme that recognizes a specific  DNA sequence and cleaves the DNA as part of a restriction-  modification system.   Restriction-modification system. A matched pair of bacterial enzymes that  recognize a specific DNA sequence: a modification methylase that  methylates bases in that sequence, and a restriction endonuclease that  cleaves the DNA if it has not been methylated in that sequence. It is a  defensive system that eliminates foreign (e.g., viral) DNA.   Retrotransposon. A transposon whose sequence and mechanism of  transposition suggest that it arose from a retrovirus.   Retrovirus. A virus whose genetic material is RNA that must be reverse-  transcribed to double-stranded DNA during host cell infection.   Reverse transcriptase. A DNA polymerase that uses RNA as its template.   Reverse turn. A polypeptide conformation in which the chain makes an  abrupt reversal in direction; usually consisting of four successive residues.  Also called a |3 bend.   Rho factor. A prokaryotic helicase that separates DNA and RNA to  promote transcription termination.   Ribosomal RNA (rRNA). The RNA molecules that constitute the bulk  of the ribosome, the site of polypeptide synthesis. rRNA provides  structural scaffolding for the ribosome and catalyzes peptide bond  formation.   Ribosome. The organelle that synthesizes polypeptides under the direction  of mRNA. It consists of around two-thirds RNA and one-third protein.   Riboswitch. An mRNA structure that regulates gene expression through  alterations in its structure triggered by the presence of the metabolite  that is synthesized by the encoded protein.   Rickets. The vitamin D-deficiency disease in children that is characterized  by stunted growth and deformed bones.   Rigor mortis. The stiffening of muscles after death due to lack of ATP.   RNA. Ribonucleic acid. A polymer of ribonucleotides. The major forms of  RNA include messenger RNA (mRNA), transfer RNA (tRNA), and  ribosomal RNA (rRNA).   RNA editing. The posttranscriptional insertion, deletion, or alteration of  bases in mRNA.   RNA interference (RNAi). A form of posttranscriptional gene regulation in  which a short double-stranded RNA segment triggers the degradation of  the homologous mRNA molecule.   RNAP. RNA polymerase, the enzyme that synthesizes RNA using a DNA  template.   Rotational symmetry. A type of symmetry in which the asymmetric units  of a symmetric object can be brought into coincidence through rotation.   Rough endoplasmic reticulum (RER). That portion of the endoplasmic  reticulum associated with ribosomes; it is the site of synthesis of  membrane proteins and proteins destined for secretion or residence in  certain organelles.   S. Svedberg, a unit for the sedimentation coefficient, equivalent to 10 13 s.   Salting in. The increase in solubility of a protein (or other molecule) with  increasing (low) salt concentration.   Salting out. The decrease in solubility of a protein (or other molecule)  with increasing (high) salt concentration.   Salvage pathway. A metabolic pathway for converting free purines and  pyrimidines to their nucleotide forms.   SAM. 5-Adenosylmethionine, a nucleotide cofactor that functions mostly as  a methyl group donor. Also called AdoMet.   Sarcomere. The repeating unit of a myofibril, which contains thin and  thick filaments that slide past each other during muscle contraction.   Saturated fatty acid. A fatty acid that does not contain any double bonds  in its hydrocarbon chain.   Saturation. The state in which all of a macromolecule’s ligand-binding sites  are occupied by ligand. See also enzyme saturation and saturated fatty acid.   Schiff base. An imine that forms between an amine and an aldehyde or  ketone.   Screening. A technique for identifying clones that contain a desired gene.   Scurvy. A disease caused by vitamin C (ascorbic acid) deficiency, which  results in inadequate formation of 4-hydroxyprolyl residues in collagen,  thereby reducing collagen’s stability.   SDS-PAGE. Polyacrylamide gel electrophoresis (PAGE) in the presence of  the detergent sodium dodecyl sulfate (SDS), which denatures and  imparts a uniform charge density to polypeptides and thereby permits  them to be fractionated on the basis of size rather than inherent charge.   Second messenger. An intracellular ion or molecule that acts as a signal for  an extracellular event such as ligand binding to a cell-surface receptor.   Second-order reaction. A reaction whose rate is proportional to the square  of the concentration of one reactant or to the product of the  concentrations of two reactants.   Secondary active transport. Transmembrane transport that is driven by the  energy stored in an electrochemical gradient, which itself is generated  utilizing the free energy of ATP hydrolysis or electron transport.   backbone atoms without regard to the conformations of its substituent  side chains, a helices and (3 sheets are common secondary structural  elements of proteins.   Secretory pathway. The series of steps in which a protein destined for  secretion, membrane insertion, or residence in certain organelles is  recognized by the signal recognition particle as it emerges from the  ribosome, has its signal peptide excised by a signal peptidase, and is  translocated across the endoplasmic reticulum membrane or inserted into  it via a translocon.   Sedimentation coefficient. A measure of a particle’s rate of sedimentation  in an ultracentrifuge, usually expressed in Svedberg units (S).   Segmentation genes. Insect genes that specify the correct number and  polarity of body segments. Gap genes, pair-rule genes, and segment  polarity genes are all segmentation genes.   Selectable marker. A gene whose product has an activity, such as antibiotic  resistance, such that, under the appropriate conditions, cells harboring  the gene can be distinguished from those that lack the gene.   Selfish DNA. Genomic DNA that has no apparent function. Also called  junk DNA.   Semiconservative replication. The natural mode of DNA duplication in  which each new duplex molecule contains one strand from the parent  molecule and one newly synthesized strand.   Semidiscontinuous replication. The mode of DNA replication in which  one strand is replicated as a continuous polynucleotide strand (the  leading strand) while the other is replicated as a series of discontinuous  fragments (Okazaki fragments) that are later joined (the lagging strand).   Sense strand. The DNA strand complementary to the strand that is   transcribed; it has the same base sequence (except for the replacement of  U with T) as the synthesized RNA. Also called the coding strand.   Sequential model of allosterism. A model for allosteric behavior in which  the subunits of an oligomeric protein change conformation in a stepwise  manner as the number of bound ligands increases.   Sequential reaction. A reaction in which all substrates must combine with  the enzyme before a reaction can occur; it can proceed by an Ordered or  Random mechanism.   Serine protease. A peptide-hydrolyzing enzyme characterized by a reactive  Ser residue in its active site.   Severe combined immunodeficiency disease (SCID). An inherited disease  that greatly impairs the immune system. One such defect is a deficiency  of the enzyme adenosine deaminase.   Shear degradation. The fragmentation of DNA by the mechanical force of  shaking or stirring.   Shine-Dalgarno sequence. A purine-rich sequence ~10 nucleotides  upstream from the start codon of many prokaryotic mRNAs that is  partially complementary to the 3' end of the 16S rRNA. This sequence  helps position the ribosome to initiate translation.   Short interfering RNA (siRNA). An exogenous RNA of 18 to 25   nucleotides that inhibits gene expression through RNA interference. See  also microRNA.   Shotgun cloning. The cloning of an organism’s genome in the form of a  set of random fragments.   deformed and damaged by the presence of a mutant hemoglobin (Glu  6(3 —> Val) that in its deoxy form polymerizes into fibers.   cr factor. A subunit of the bacterial RNA polymerase holoenzyme that  recognizes a gene’s promoter and is released once chain initiation has  occurred.   Sigmoidal curve. The S-shaped graphical representation of the cooperative  binding of a ligand to a molecule.    Signal peptide. A short (13—36 residues) N-terminal peptide sequence that  targets a nascent secretory or transmembrane protein to the endoplasmic  reticulum (in eukaryotes) or plasma membrane (in prokaryotes). This  leader peptide is subsequently cleaved away by a signal peptidase.   Signal recognition particle (SRP). A protein-RNA complex that binds to  the signal peptide of a nascent transmembrane or secretory protein and  escorts it to the endoplasmic reticulum (in eukaryotes) or plasma  membrane (in prokaryotes) for translocation through the membrane.   Signal transduction. The transmittal of an extracellular signal to the cell  interior by the binding of a ligand to a receptor so as to elicit a cellular  response through the activation of a sequence of intracellular events that  often include the generation of second messengers.   Silencer. A DNA sequence some distance from the transcription start site,  where a repressor of transcription may bind.   Single-displacement reaction. A reaction in which a group is transferred  from one molecule to another in a concerted fashion (with no  intermediates).   Single nucleotide polymorphism (SNP). A single base difference in the  genomes of two individuals; such differences occur every 1250 bp on  average in the human genome.   molecules of which coat single-stranded DNA during replication so as to  prevent the re-formation of double-stranded DNA.   Site-directed mutagenesis. A technique in which a cloned gene is mutated  in a specific manner.   Sliding filament model. A mechanism for muscle contraction in which  interdigitated thin and thick filaments move past each other so as to  shorten the overall length of a sarcomere.   Small nuclear ribonucleoprotein (snRNP). A complex of protein and  small nuclear RNA that participates in mRNA splicing.   Small nuclear RNA (snRNA). Highly conserved 60- to 300-nt RNAs that  participate in mRNA splicing.   Small nucleolar RNA (snoRNA). Eukaryotic RNA molecules of 70 to 100  nt that pair with immature rRNAs to direct their sequence-specific  methylation.   SNARE. A membrane-associated protein that participates in vesicle fusion;  SNAREs from the two fusing membranes form bundles of four helices  that bring the membranes together.   Soap. A salt of a long-chain fatty acid, which contains a polar head group  and a long hydrophobic tail.   Solvation. The state of being surrounded by several layers of ordered  solvent molecules. Hydration is solvation by water.   Somatic hypermutation. The greatly increased rate of mutation that occurs  in the immunoglobulin genes of proliferating B lymphocytes and leads,  over several cell generations, to antibodies with higher antigen affinity.   Somatic recombination. Genetic rearrangement that occurs in cells other  than germline cells.   Sonication. Irradiation with high frequency sound waves. Such treatment is  used to mechanically disrupt cells and subcellular membranous structures.   SOS response. A bacterial system that recognizes damaged DNA, halts its  replication, and repairs the damage, although in an error-prone fashion.   Southern blotting. A procedure for identifying a DNA base sequence after  electrophoresis, through its ability to hybridize with a complementary  single-stranded segment of labeled DNA or RNA. See also Northern  blotting.   Special pair. The set of two closely spaced chlorophyll molecules in a  photosynthetic system that undergo photooxidation.   Spherocytosis. A hereditary abnormality in the erythrocyte cytoskeleton that  renders the cells rigid and spheroidal and which causes hemolytic anemia.    Sphingolipid. A derivative of the C 18 amino alcohol sphingosine.  Sphingolipids include the ceramides, cerebrosides, and gangliosides.  Sphingolipids with phosphate head groups are called  sphingophospholipids.   Sphingomyelin. The most common sphingolipid, consisting of a ceramide  bearing a phosphocholine or phosphoethanolamine head group.   Spliceosome. A 60S particle containing proteins, snRNPs, and pre-mRNA;  it carries out the splicing reactions whereby a pre-mRNA is converted to  a mature mRNA.   Splicing. The usually ribonucleoprotein-catalyzed process by which introns  are removed and exons are joined to produce a mature transcript. Some  RNAs are self-splicing.   Spontaneous process. A thermodynamic process that occurs without the  input of free energy from outside the system. Spontaneity is independent  of the rate of a process.   Squelching. The inhibition of the activity of a transcription factor by  another transcription factor that competes with it for binding to DNA.   SRP receptor (SR). The endoplasmic reticulum protein that serves as a  docking site for the signal recognition particle (SRP) during the synthesis  of a transmembrane or secretory protein.   Stacking interactions. The stabilizing van der Waals interactions between  successive (stacked) bases and base pairs in a polynucleotide.   Standard state. A set of conditions including unit activity of the species of  interest, a temperature of 25°C, a pressure of 1 atm, and a pH of 0.0.   Starch. A mixture of linear and branched glucose polymers that serve as the  principal energy reserves of plants.   energy, whose values depend only on the current state of the system, not  on how they reached that state.   Steady state. A set of conditions in an open system under which the  formation and degradation of individual components are balanced such  that the system does not change over time.   Steady state assumption. A condition for the application of the  Michaelis—Menten model to an enzymatic reaction, in which the  concentration of the ES complex remains unchanged over the course of  the reaction.   Stem-loop. A secondary structural element in a single-stranded nucleic  acid, in which two complementary segments form a base-paired stem  whose strands are connected by a loop of unpaired bases.   Stereoisomers. Chiral molecules with different configurations about at least  one of their asymmetric centers but which are otherwise identical.   Steroid. Any of numerous naturally occurring lipids composed of four  fused rings; many are hormones that are derived from cholesterol.   Sticky end. The single-stranded extension of a DNA fragment that has  been cleaved at a specific sequence (often by a restriction endonuclease)  in a staggered cut such that the single-stranded extension is  complementary to those of similarly cleaved DNAs.   Stop codon. A sequence of three nucleotides that does not specify an  amino acid but instead causes the termination of translation. Also called  a nonsense codon.   Striated muscle. The voluntary or skeletal muscles, which have a striped  microscopic appearance.   Stroma. The concentrated solution of enzymes, small molecules, and ions  in the interior of a chloroplast; the site of carbohydrate synthesis.   Strong acid. An acid that is essentially completely ionized in aqueous  solution. A strong acid has a dissociation constant much greater than  unity (p K < 0).   molecules as guided by knowledge of the target protein’s structure. Also  called rational drug design.   Substrate cycle. Two opposing sets of metabolic reactions that, in many  cases, function together to hydrolyze ATP, but provide a control point  for regulating metabolic flux. Also called a futile cycle.   Substrate-level phosphorylation. The direct transfer of a phosphoryl group  to ADP to generate ATP.   Subunit. One of several polymer chains that make up a macromolecule.   Sulfhydryl group. A portion of a molecule with the formula —SH.   Supercoiling. The topological state of covalently closed circular double  helical DNA in which the double helix is twisted around itself. It arises  through the over- or underwinding of the double helix. Also called  superhelicity.   Superoxide radical. 0 2 ’, a partially reduced oxygen species that can  damage biomolecules through free radical reactions.   Supersecondary structure. A common grouping of secondary structural  elements. Also called a motif.   Suppressor mutation. A mutation that cancels the effect of another  mutation.   Surface labeling. A technique in which a lipid-insoluble protein-labeling  reagent is used to identify the portion of a membrane protein that is  exposed to solvent.   Surroundings. In thermodynamics, the universe other than the particular  system that is of interest.   Svedberg (S). Equivalent to 10 13 s; used to describe a particle’s   Symmetry model of allosterism. A model for allosteric behavior in which  all the subunits of an oligomeric protein are constrained to change  conformation in a concerted manner so as to maintain the symmetry of  the oligomer.   Symport. A transmembrane channel that simultaneously transports two  different molecules or ions in the same direction. See also antiport and  uniport.   Syn conformation. A purine nucleotide conformation in which the ribose  and the base are eclipsed. See also anti conformation.   Syncytium. A single cell containing multiple nuclei that results from  repeated nuclear division without the formation of new plasma  membranes.   System. In thermodynamics, the part of the universe that is of interest; the  rest of the universe is the surroundings. See also closed system, isolated  system, and open system.   Systems biology. The computer-based collection and analysis of data sets  for the purpose of discerning relationships between dynamic or  multifactorial biological entities.   T state. One of two conformations of an allosteric protein; the other is the  R state. The T state is customarily the catalytically less active state.   T —> R transition. A shift in conformation of an allosteric protein induced  by ligand binding.   TAFs. TBP-associated factors, which, along with TBP, constitute the general  transcription factor TFIID required for the transcription of eukaryotic  structural genes.   TATA box. A eukaryotic promoter element with the consensus sequence  TATAtAt located 10 to 27 nucleotides upstream from the transcription  start site.   Tautomers. Isomers that differ only in the positions of their hydrogen  atoms and double bonds.   deficiency of hexosaminidase A, the lysosomal enzyme that breaks down  ganglioside G M2 .   TBP. TATA-binding protein, a DNA-binding protein that is required for  transcription of all eukaryotic genes.   Telomerase. An RNA-containing DNA polymerase that, using the RNA  as a template, catalyzes the repeated addition of a specific G-rich  sequence to the 3' end of a eukaryotic DNA molecule to form a  telomere.   Telomere. The end of a linear eukaryotic chromosome, which consists of  tandem repeats of a short G-rich sequence on the 3 '-ending strand and  its complementary sequence on the 5 '-ending strand.   Tertiary structure. The entire three-dimensional structure of a single-chain  polymer, including that of its side chains.   Tetrahedral intermediate. An intermediate of peptide bond hydrolysis in  which the carbonyl carbon of the scissile bond has undergone  nucleophilic attack so that it has four substituents.   Therapeutic index. The ratio of the dose of a drug that produces toxicity  to the dose that produces the desired effect.   Thermodynamics. The study of the relationships among various forms  of energy.   Thermogenesis. The generation of heat by muscle contraction (shivering)  or by fuel oxidation without ATP synthesis (also called diet-induced  thermogenesis).   0 structure. The appearance of a circular DNA molecule undergoing  replication by the progressive separation of its two strands.   THF. Tetrahydrofolate, a cofactor for reactions that transfer one-carbon  units in various oxidation states.   Thick filament. The sarcomere element that is composed primarily of  several hundred myosin molecules.   Thin filament. The sarcomere element that is composed primarily of actin,  together with tropomyosin and troponin.   30-nm fiber. A condensed chromatin structure in which nucleosomes fold  in a zigzag manner to form a fiber with a diameter of ~30 nm.   3' end. The terminus of a polynucleotide whose C3' is not esterified to  another nucleotide residue.   Thylakoid. The innermost compartment in chloroplasts, which is formed  by invaginations of the chloroplasts inner membrane. The thylakoid  membrane is the site of the light reactions of photosynthesis.   Titration curve. The graphical presentation of the relationship between the  pH of an acid- or base-containing solution and the degree of proton  dissociation (roughly equal to the number of equivalents of strong base  or strong acid that have been added to the solution).   Topoisomerase. An enzyme that alters DNA supercoiling by catalyzing  breaks in one or both strands, passing DNA through the break, and  resealing the break.   Topology. The study of the geometric properties of an object that are not  altered by deformations such as bending and stretching.   Torsion angle. The dihedral angle described by the bonds between four  successive atoms. The torsion angles cj) and i|/ indicate the backbone  conformation of a peptide group in a polypeptide.   TPP. Thiamine pyrophosphate, a cofactor for reactions in which a-keto  acids undergo decarboxylation.   Ti|/C arm. A conserved stem-loop structure in a tRNA molecule that  usually contains the sequence Tv|/C, where vjt is pseudouridine.   Trans conformation. An arrangement of the peptide group in which  successive C a atoms are on opposite sides of the peptide bond.    Trans peptide. A conformation in which successive C a atoms are on  opposite sides of the peptide bond.   Transamination. The transfer of an amino group from an amino acid to  an a-keto acid to yield a new a-keto acid and a new amino acid.   Transcription. The process by which RNA is synthesized under the  direction of a DNA template, thereby transferring genetic information  from the DNA to the RNA. Transcription is catalyzed by RNA  polymerase as facilitated by numerous other proteins.   Transcription factor. A protein that promotes the transcription of a gene  by binding to DNA sequences at or near the gene or by interacting with  other proteins that do so.   Transcriptomics. The study of all the mRNA molecules that a cell  transcribes.   Transfer RNA (tRNA). The small L-shaped RNAs that deliver specific  amino acids, which have been esterified to the tRNA’s 3' ends, to  ribosomes according to the sequence of a bound mRNA. The proper  tRNA is selected through the complementary base pairing of its three-  nucleotide anticodon with the mRNA’s codon, and the growing  polypeptide is transferred to its aminoacyl group.   Transferase. An enzyme that catalyzes the transfer of a functional group  from one molecule to another.   Transformation. (1) The permanent alteration of a bacterial cell’s genetic  message through the introduction of foreign DNA. (2) The genetic  changes that convert a normal cell to a cancerous cell.   Transgene. A foreign gene that is stably expressed in a host organism.   Transgenic organism. An organism that stably expresses a foreign gene  (transgene).   Transition. A mutation in which one purine (or pyrimidine) replaces  another.   Transition state. A molecular assembly at the point of maximal free energy  in the reaction coordinate diagram of a chemical reaction.   Transition state analog. A stable substance that geometrically and  electronically resembles the transition state of a reaction.   Transition temperature. The temperature at which a lipid bilayer shifts  from a gel-like solid to a more fluid liquid crystal form.   Translation. The process by which a polypeptide is synthesized under the  direction of the sequence information contained in a messenger RNA as  specified by the genetic code. Translation is catalyzed by ribosomes and  requires the additional participation of messenger RNA, transfer RNA,  and a variety of protein factors.   Translocation. (1) The movement of a polypeptide through a membrane  during the synthesis of a secreted protein. (2) The movement, by one  codon, of the ribosome relative to the mRNA after peptide bond  synthesis.   Translocon. A multisubunit protein that forms an aqueous pore across the  endoplasmic reticulum membrane that functions to translocate a nascent  polypeptide across the membrane or to install it in the membrane as part  of the secretory pathway.   Transmembrane (TM) protein. An integral protein that completely spans  the membrane.   Transmissible spongiform encephalopathy (TSE). An invariably fatal  neurodegenerative disease resulting from a prion infection, such as  scrapie in sheep.   Transpeptidation. The ribosomal process in which a tRNA-bound nascent  polypeptide is transferred to a tRNA-bound aminoacyl group so as to  form a new peptide bond, thereby lengthening the polypeptide by one  residue at its C-terminus.   Transposition. The movement (copying) of genetic material from one part  of the genome to another or, in some cases, from one organism to  another.   Transposon. A genetic unit that can move (be copied) from one position  to another in a genome; some transposons carry genes. Also called a  transposable element.   Transverse diffusion. The movement of a lipid from one leaflet of a bilayer  to the other. Also called flip-flop.    Transversion. A mutation in which a purine is replaced by a pyrimidine or  vice versa.   Treadmilling. The addition of monomeric units to one end of a linear  assembly, such as an actin fdament, and their simultaneous removal from  the opposite end such that the length of the assembly remains  unchanged.   Triacylglycerol. A lipid in which three fatty acids are esterified to a glycerol  backbone. Also called a triglyceride.   Tumor suppressor. A protein whose loss or inactivation may lead to  cancer.   Twist (T). The number of complete revolutions that one strand of a  covalently closed circular double helical DNA makes around the duplex  axis. It is positive for right-handed superhelical coils and negative for  left-handed superhelical coils.   Two-dimensional (2D) gel electrophoresis. A technique in which proteins  are first subjected to isoelectric focusing, which separates them by net  charge, and then to SDS-PAGE in a perpendicular direction, which  separates them by size.   Tyrosine kinase. An enzyme that catalyzes the ATP-dependent  phosphorylation of a Tyr side chain.   Ubiquitin. A small, highly conserved protein that is covalently attached to  a eukaryotic intracellular protein so as to mark it for degradation by a  proteasome.   Ultracentrifugation. A procedure that subjects macromolecules to a strong  centrifugal force (in an ultracentrifuge), thereby separating them by size  and/or density and providing a method for determining their mass and  subunit structure.   Uncompetitive inhibition. A form of enzyme inhibition in which an  inhibitor binds to the enzyme—substrate complex and thereby decreases  its apparent K M and its apparent V msx by the same factor.   Uncoupler. A substance that allows the proton gradient across a membrane  to dissipate without ATP synthesis so that electron transport proceeds  without oxidative phosphorylation.   Uniport. A transmembrane channel that transports a single molecule or  ion. See also antiport and symport.   Unsaturated fatty acid. A fatty acid that contains at least one double bond  in its hydrocarbon chain.   Urea cycle. A catalytic cycle in which amino groups donated by ammonia  and aspartate combine with a carbon atom from HC0 3 to form urea for  excretion and which provides the route for the elimination of nitrogen  from protein degradation.   Uronic acid. A sugar produced by oxidation of an aldose primary alcohol  group to a carboxylic acid group.   v. Reaction velocity, typically measured as the rate of appearance of product  or disappearance of reactant.   van der Waals distance. The distance of closest approach between two  nonbonded atoms.   van der Waals forces. The noncovalent associations between molecules that  arise from the electrostatic interactions among permanent and/or induced  dipoles.   van’t Hoff plot. A graph of i6 eq versus 1 IT that is used to determine A H°  and A5° and therefore AG° for a chemical reaction.   Variable arm. A nonconserved region of a tRNA molecule that contains  3 to 21 nucleotides and that may include a base-paired stem.   Variable region. The N-terminal portions of an antibody molecule, where  antigen binding occurs and which are characterized by high sequence  variability.   Virus. A nonliving entity that co-opts the metabolism of a host cell to  reproduce.   synthesized by an animal and must therefore be obtained from its diet.   ^max. The observed maximal velocity of an enzymatic reaction, which may  differ from the true value due to the presence of an inhibitor.   Voltage-gated channel. A channel whose opening and closing (gating) is  controlled by a change in membrane potential.   W. (1) See writhing number. (2) The number of energetically equivalent  ways of arranging the components of a system.   w. The thermodynamic term for the work done by a system on its  surroundings.   Water of hydration. The shell of relatively immobile water molecules that  surrounds and interacts with (solvates) a dissolved molecule.   Watson-Crick base pair. A stable pairing of nucleotide bases, either  adenine with thymine or guanine with cytosine, that occurs in DNA  and, to a lesser extent, in RNA (in which thymine is replaced by uracil).  See also Hoogsteen base pair.   Weak acid. An acid that is only partially ionized in aqueous solution. A  weak acid has a dissociation constant less than unity (p K > 0).   Wild type. The naturally occurring version of an organism or gene.   Wobble hypothesis. An explanation for the permissive tRNA—mRNA  pairing at the third anticodon position that includes non-Watson-Crick  base pairs. This allows many tRNAs to recognize two or three different  (degenerate) codons.   Writhing number ( W ). The number of turns that the duplex axis of a  covalently closed circular double helical DNA makes around the  superhelix axis. It is a measure of the DNA’s superhelicity.   molecular structures from the diffraction pattern produced by exposing a  crystal of a molecule to a beam of X-rays.   Xenobiotic. A molecule that is not normally present in an organism.   Yeast artificial chromosome (YAC). A linear DNA molecule that contains  the chromosomal structures required for normal replication and  segregation in a yeast cell. YACs are commonly used as cloning vectors.   Z-scheme. A Z-shaped diagram indicating the sequence of events and their  reduction potentials in the two-center photosynthetic electron-transport  system of plants and cyanobacteria.   Zero-order reaction. A reaction whose rate does not vary with the  concentration of any of its reactants.   Zinc finger. A protein structural motif, often involved in DNA binding,  that consists of 23 to 60 residues that include His and/or Cys residues to  which one or two Zn 2+ ions are each tetrahedrally coordinated.    Page numbers in bold face refer to a major discussion of the entry. F after a page number refers to  a table. Positional and configurational designations in chemical names (e.g., 3-, a, N-, ptrans , D-)  Greek letters are otherwise alphabetized as if they were spelled out.   A, see Adenine  A (angstrom), 12  AAA+ domain, 877, 878  A antigens, 238, 238T  A bands, 197  Abasic sites, 897, 898   ABCA1 (ATP-cassette binding protein Al), 729  ABC transporters, 304, 308-309  Abdominal cavity, adipose tissue in, 772  A|3 precursor protein (pPP), 170  Ap protein (amyloid-P protein), 170  Ab initio protein design, 165  Abl protein, 414   ABO blood group antigens, 237-238  Abortive initiation, 925, 926  Absolute configuration, 85  Absorbance, 97   Accepted (recommended) name, 324  Acceptor stem (tRNA), 968, 969F  Accessory BChl, 631  Accessory pigments, 627—628  Acesulfame, 225   Acetal, acid-catalyzed hydrolysis of, 335F  Acetaldehyde, 318, 319, 491  Acetamide, 313   from tryptophan breakdown, 738, 738F  Acetoacetyl-ACP, in fatty acid synthesis, 685F  Acetoacetyl-CoA, 678, 679F, 680F  Aceto-a-hydroxybutyrate, 748  Acetolactate, 748  Acetolactate synthase, 748F  Acetone, 318, 327F   Acetyl-ACP, in fatty acid synthesis, 685F  Acetylaldehyde, 495  Acetylcholine, hydrolysis, 340  Acetylcholinesterase, 340, 362T   in amino acid degradation, 727F, 736F  in cholesterol biosynthesis from, 701-704  in citric acid cycle, 553F, 555-561, 568F,   in pyruvate dehydrogenase complex, 555-560  in triacyclglycerol biosynthesis, 691  Acetyl-CoA-ACP transacylase, 685F  Acetyl-CoA carboxylase (ACC), 781  7V-Acetyl-D-galactosamine (GalNAC), 229  7V-Acetylglucosamine (NAG, GlcNAc), 228, 333,  333F, 335F, 337, 338  MAcetyl-D-glucosamine, 233, 233F  7V-Acetylglutamate, 726  7V-Acetylglutamate synthase, 726-727  7V-Acetyllactosamine, 545  Acetyl-Lys binding sites, 1039F  8-AkAcetyllysine, 88F  M-Acetylmannosamine, 222   Af-Acetylneuraminic acid (NANA), 222, 222F, 225F  Acetyl phosphate, 448T, 452, 484  Acids, 32-34   conjugate bases, 32  general, 481  as proton donors, 32  Acid-base catalysis (enzymes), 323-327  Acid-base chemistry, 32-34  and ionization of water, 30-31  standard state conventions, 17  Acidic solutions, 31  Aconitase:   in citric acid cycle, 553F, 563-564, 571  free energy of reactions with, 570T  in glyoxylate cycle, 575F  stereospecificity of, 318  Aconitate, 552, 563, 564  ACP (acyl-carrier protein), 683-684  Acquired immunodeficiency syndrome (AIDS), 71,  368, 370   Acridine orange, as intercalating agent, 847, 893  Acromegaly, 401  Actin, 199, 201-203, 206F  and ATPase, 612  myosin-actin interaction, 202F  structure, 20IF  a-Actinin, 203  P-Actinin, 203   Actinomycin D, 930  Action potentials, 295  Action potential time course, 295F  Activation domains, 1044  Activation energy, 320-323  Activation-induced deaminase (AID), 1059  Activators (regulatory proteins), 1043  Active membrane transport, 271, 290, 304—314  ABC transporters, 308-309  ATP-driven, 304-309  calcium ATPase, 306-308  endergonic process, 304  ion-gradient-driven, 309-312  sodium and potassium, 305F  sodium and potassium ATPase, 304—306  Active site:   of serine proteases, 339, 340, 343F  of TIM, 481-482  of zymogens, 351  Active transporters, 304-312  Activity, 17   Acyl-CoA dehydrogenase (AD), 667-668  Acyl-CoA oxidase, 678  Acyl-CoA synthetases, 665, 666F  Acyl-CoA transferase, 698F  Acyl-dihydroxyacetone phosphate, 690F  Acyl-dihydroxyacetone phosphate reductase, 690F  Acyl-enzyme intermediate:   chymotrypsin, 345, 348-349, 348F  serine protease, 344F  2-Acylglycerol, 658  l-Acylglycerol-3-phosphate, 247  1 -Acylglycerol-3-phosphate acyl transferase,   AD (acyl-CoA dehydrogenase), 667-668  ADA, see Adenosine deaminase  Adaptive thermogenesis, 783-784  Adaptor, 408  ADAR2, 953   Addison’s disease, 213T, 251  Addition site (A), RNAP II, 934, 934F  Adenine (A), 41, 42T   as common nucleotide, 42  modified forms in tRNA, 969F  oxidative deamination, 892F  stacking interactions, 840F  Adenine nucleotide translocase, 584-586  Adenine phosphoribosyltransferase (APRT), 799  Adenine ribonucleotide synthesis, 797-798  Adenosine, 42T, 448, 814F  Adenosine-3 , ,5 , -cyclic monophosphate, see cAMP  Adenosine-5 / (P,7-imido)triphosphate (AMPPNP,  ADPNP), 405, 412F  Adenosine deaminase (ADA), 369-370,   Adenosine diphosphate, see ADP  Adenosine monophosphate, see AMP  Adenosine triphosphate, see ATP  S-Adenosylhomocysteine, 732, 1040  S-Adenosylmethionine (SAM, AdoMet), 732, 733F,  1032, 1040  Adenovirus-2, 945   Adenylate cyclase pathway, 779, 779F  Adenylate kinase, 453, 454F, 500  Adenylation, 45IF  Adenylosuccinate, 797, 797F  Adenylosuccinate lyase, 795F, 797, 797F  Adenylosuccinate synthetase, 797F  Adenylylation, 744  Adenylyltransferase, 744  ADH, see Alcohol dehydrogenase  Adhesive mechanism, for CAP binding, 1028  Adipocytes, 244, 244F, 772, 781-783  Adiponectin, 115F, 782, 782F  Adiponectin receptors, 782  Adipose tissue, 244, 442  brown, 615, 784   insulin and fuel storage in, 776, 777  mammalian metabolism in, 769, 770F, 772  A-DNA, 823-827, 825T, 826F-827F  AdoCbl, 672   as citric acid cycle regulator, 570, 572  in glycolysis, 474F, 475  ADP-ATP translocator, 584-586, 586F  ADP-glucose, 43, 43F, 546  ADP-glucose pyrophosphorylase, 648  ADPNP (adenosine-5'(|3,y-imido)triphosphate;   Adrenal glands, 250, 397F, 398, 400-401, 776  Adrenaline, see Epinephrine  Adrenergic receptors (adrenoreceptors), 398, 537  a-, 398-400, 537  P-, 398-400, 419F, 537  Adrenocortical steroids, 400  Adsorption, 97   Adult respiratory distress syndrome, 246  Adverse drug reactions, 389-391  Aequorea victoria, 89  Aerobic heterotrophs, 437  Aerobic metabolism, 491   physiological implications, 617—618  and reactive oxygen species, 618  Affinity chromatography, 102, 102F, 846-847   AICAR transformylase, in IMP synthesis, 795F  AID (activation-induced deaminase), 1059  AIDS, see Acquired immunodeficiency syndrome  AIR (5-aminoimidazole ribotide), 795F, 796  AIR carboxylase, in IMP synthesis, 795F, 796  AIR synthetase, in IMP synthesis, 795F  Akt protein, 432F   a helix/(3 sheet propensities, 14IT  biosynthesis, 742-743, 742F  breakdown, 727-730, 727F, 728F  as common amino acid, 95  genetic code specification, 967T  glucose-alanine cycle, 775, 775F  ionizable groups, 78T  nonpolar side chain, 80, 81F  from phenylalanine breakdown, 739, 739F  side chain hydropathy, 157T  structure, 78T  transamination, 544   from tryptophan breakdown, 738, 738F  Alanine aminotransferase, 728F  Alanine transaminase (ALT), 720  Alanine tRNA (tRNA* 11 ), 968  (3-Alanine, in pyrimidine catabolism, 818, 818F  Alber, T., 856  Alberts, B., 926  Albumin, 664  Alcaptonuria, 464, 740  Alcohols, functional group and linkages, 4T  Alcohol dehydrogenase (ADH), 317-319,   in Calvin cycle, 645F  enzymatic mechanism of, 480F  in glycolysis, 474F, 478-479, 479F  Aldol cleavage, 479, 479F  Aldonic acids, 221  Aldopentoses, 218F  Aldoses, 218, 218F  Aldosterone, 250, 25IF, 400  Aldotetroses, 218F  Aldotrioses, 218F  Alitame, 225   Alkylacylglycerophospholipids, 697  Alkylating agents, mutagenic effects, 893  0 6 -Alkylguanine-DNA alkyltransferase, 896  Alkyltransferases, 896  Allantoic acid, 816, 816F  Allantoin, 816, 816F    Allis, D., 1036  1,6-Allolactase, 1024  Allopurinol, 817  D-Allose, 218F  Allosteric control:   of enzymes, 317, 377-381, 445  of glycogen phosphorylase, 529, 530  of glycogen synthase, 528—530  and metabolic flux, 445  of phosphofructokinase, 498, 499, 499F  phosphorylation and dephosphorylation vs.,  383-384   Allosteric effectors, 377, 544-545  Allosteric modifiers, 380-381  Allosteric proteins, 191-193  Allosterism:   interactions, allosteric, 191-193  sequential model of, 192-193, 193F  symmetry model of, 192-193, 192F  Alloxanthine, 817  Allysine, 140, 14 IF  Almassy, R., 796  a anomers, 220, 22IF  a band (Soret band), 596  a/(3 barrel (TIM barrel), 150, 482  a/(3 proteins, 149  a carbon, 77, 85   Amides, functional group and linkages, 4T  Amidophosphoribosyl transferase, in IMP synthesis,  794, 795F  Amines, 324   functional group and linkages, 4T  physiologically active, 756-757  reaction with carboxylic acids, 3F  Amino acids, 76-90   biologically active, 88-90, 89F  charged polar side chains, 79T, 82, 82F  derivatives in proteins, 87-90, 88F  as dipolar ions, 80   general properties, 77-79, 78T-79T  genetic code specification, 967T  glucogenic, 727  hydrophobic effect, 82  ketogenic, 727  nomenclature, 83-84, 84F  nonessential, 74IT  nonpolar side chains, 78T, 80, 81F  nutritive importance, 76  occurrence, 78T-79T    peptide bonds, 80  pK values, 78T-79T, 82-83  polar side chains, 82, 82F  side chains of, 80-82   specificity of aminoacyl-tRNA synthetases, 971  specifying by codon, 963  stereochemistry, 84-87  structure, 77 - 84 , 78T-79T  translation, 5IF  transport between tissues, 775  uncharged polar side chains, 81, 8IF  a-Amino acids, 77, 77F, 85  D-Amino acids, 85-87  L-a- Amino acids, 85  l- Amino acids, 85-87  Amino acid biosynthesis, 740-751  citric acid cycle intermediates, 573  essential amino acids, 740, 741, 746-751  mammalian metabolism, 768F, 769  nonessential amino acids, 741-746  Amino acid breakdown, 727 - 740 , 727F   arginine, glutamate, glutamine, histidine, and  proline, 731-732  asparagine and aspartate, 730-731  branched-chain amino acids, 736-737, 736F  common intermediates, 727F  leucine and lysine, 737-738  mammalian metabolism, 768F, 769, 773-774  methionine, threonine, isoleucine, and valine,  732-737   phenylalanine and tyrosine, 739 , 740  tryptophan, 727F, 738  Amino acid catabolism, 719F  Amino acid deamination, 718-722  oxidative, 722  transamination, 719-721  Amino acid derivatives, 87-90  Amino acid metabolism, 88, 712-764  biosynthesis of amino acids, 740-751  deamination, 718-722  degradation of amino acids, 727-740  heme from, 752-756  nitric oxide from, 757-758  nitrogen fixation, 758-764  physiologically active amines from, 756—757  protein degradation, 713-718  urea cycle, 722-727  Amino acid residues, 80   a helix/(3 sheet propensities, 14IT  conservative substitution, 118  hydropathies, 157T  hypervariable, 118  invariant, 118  pH effects, 325  polypeptide chain interior, 82  Amino acid stem (tRNA), 968, 969F  Aminoacrylate, 729, 729F  Aminoacyl-adenylate, 971  Aminoacylation site (aminoacyl-tRNA  synthetases), 971   Aminoacyl site (A), 981, 982, 1000, 1003-1004  Aminoacyl-tRNA (aa-tRNA), 971, 97IF,   Aminoacyl-tRNA synthetases (aaRSs), 970-974  amino acid attachment to, 970  Class I vs. Class II, 971  organisms lacking, 974   5-Aminoimidazole-4-(Af-succinylocarboxamide)  ribotide (SACAIR), 795F, 796  5-Aminoimidazole ribotide (AIR), 795F, 796  (3-Aminoisobutyrate, 818, 818F  a-Amino-p-ketobutyrate, 729  a-Amino-p-ketobutyrate lyase, 728F, 729  8-Aminolevulinic acid (ALA), 752, 753F  8-Aminolevulinic acid synthase, 753F  Aminopterin, 813  Amino sugars, 222  Amino terminus, see AHerminus  Aminotransferase, 719, 818F  Ammonia, 722-723, 816F  Ammonification, 764  Ammonium sulfate, 99  Amoeboid motion, 207  AMP (adenosine monophosphate), 42T  animal catabolism pathway, 814F  from IMP, 797-798, 797F  and inhibition of PFK, 499-500  synthesis, 795F, 797-798  AMP deaminase, 814   AMP-dependent protein kinase (AMPK), 683, 692,  704, 780 - 782 , 78IF  amp gene, 912  Amphibolic pathways, 572  Amphipathic molecules, 27-28  Amphiphiles, 259-260  Amphiphilic molecules, 27-28  AMPK, see AMP-dependent protein kinase  AMP nucleosidase, catalytic power, 316T  AMPPNP (adenosine-5XP>74mido)triphosphate;   Amyloid-P protein (Ap), 170  Amyloid fibrils, 172, 172F  Amyloidoses, 169  Amylopectin, 227, 228, 531  a-Amylose, 227, 227F, 531  Amylo-( 1,4 —>1,6)-transglycosylase (branching  enzyme), 524, 529-530  Amylo-( 1,4—>1,6)-transglycosylase deficiency  (Andersen’s disease), 524, 525  Amytal, 589  Anabolic steroids, 401  Anabolism, 436, 444  Anacystis nidulans, 896, 896F  Anaerobes, 437  Anaerobic glycolysis, 492, 618  Anaerobic metabolism, 7, 491, 771  Anaerobiosis, 473  Analbuminemia, 664  Anaplerotic reactions, 574  Andersen’s disease, 524, 525  Androgens, 250, 25IF, 400   sickle-cell, 71, 194 - 196 , 196F, 513  Anencephaly, 732  Anfinsen, C., 164  Angelman syndrome (AS), 1043  Angina pectoris, 757  Angstrom (A), 12  Animal cells, 8F  Anion channels, 298   Ankyrin repeats, 268, 269F  Annealing, 64, 842  a Anomers, 220, 22IF  Anomeric carbon, 220, 223  Anomeric forms, 220  Antagonists, 398, 399  ANT-C (antennapedia complex), 1072  Antenna chlorophyll, 626, 626F  Antennapedia complex {ANT-C), 1072  Antennapedia mutant {antp), 1070  Anteriobithorax mutant {abx), 1070  Anterograde transport, 276  Anthranilate, 749, 749F  Antibiotics:   effects on protein synthesis, 1000-1001  peptidoglycan-specific, 234  transcription inhibitors, 930, 931  as transition state analogs, 332  type II topoisomerase inhibitors as, 838  Antibiotic-resistant transposons, 912-914  Antibodies, 208-213   antigen cross-linking, 21 IF  antigen recognition by, 210-213  constant and variable regions of, 208-210  defined, 208   diversity of, 211 - 213 , 1056-1059  for immunoassays, 97  interaction of antigen and, 21 IF  monoclonal, 210-212  X-ray structure, 209F  Anticodons, 958F, 963   recognition by aminoacyl-tRNA synthetases, 971  Anticodon arm (tRNA), 968, 969F  Anti conformation, 829, 829F  Antifolates, 813  Antigens, 97, 208, 21 IF   antigen-antibody binding, 210 - 213 , 21 IF  oligosaccharides as antigenic determinants, 237,  238, 238T  Antimycin A, 589  Antioxidants, 619-620  Antiparallel P sheet, 134, 134F  Antiparallel strands, DNA, 46  Antiport, 302F, 303  Antiprism, 294  Antisense RNA, 1011, 1051  Antisense (noncoding) strand, 922, 922F  antp {antennapedia) mutant, 1070  Ap 5 A, 454   ApoA-I (apolipoprotein A-I), 661  ApoB (apolipoprotein B), 952  ApoB-48 (apolipoprotein B-48), 952  ApoB-100 (apolipoprotein B-100), 661, 952  Apoenzymes, 320  Apolipoproteins, 661, 952  Apolipoprotein A-I (apo A-I), 661  Apolipoprotein B (apoB), 952  Apolipoprotein B-48 (apoB-48), 952  Apolipoprotein B-100 (apoB-100), 661, 952  Apomyoglobin, 112F  Apoproteins, 661   in development of hands and feet, 1064, 1064F  as essential process, 1064-1065  triggers for, 1066-1068  Apoptosis protease-activating factor-1 (Apaf-1),  1066, 1068   Apoptosome, 1068, 1068F  Apoptotic bodies, 1065  Appetite, regulation of, 783  APRT (Adenine phosphoribosyltransferase), 799  APS/Cbl complex, 433  Aptamers, 1031  Apurinic (AP) sites, 897, 898  Apyrimidinic (AP) sites, 897, 898  AQP0 (aquaporin-0), 258F  AQP1, 299-300, 299F, 300F  Aquaporins, 299 - 300 , 299F, 300F  kidneys, 299  lacrimal glands, 299  salivary glands, 299  water molecules, 299, 300  Aquaporin-0 (AQP0), 258F  Arabidopsis thaliana, 59T, 60, 577  D-Arabinose, 218F  Arachidic acid, 243T  Arachidonate, 699  Arachidonic acid, 243T, 254, 254F  Archaea, 9F   Architectural proteins, 1034-1036, 1044  AREs (AU-rich elements), 1050  Arginase, 724F, 726  Arginine (Arg):   a helix/(3 sheet propensities, 14IT  biosynthesis, 745, 745F, 746  breakdown, 727F, 731-732, 73IF  charged polar side chain, 82  genetic code specification, 966, 967T  ionizable groups, 79T  as NO precursor, 757-758, 757F  side chain hydropathy, 157T  structure, 79T  in urea cycle, 724F, 726  Argininosuccinase, 724F, 726  Argininosuccinate, 724F, 726F  Argininosuccinate synthetase, 724F, 726  Argonaute (AGO), 1053, 1054  Arg residues, 300, 312  Aromatic amino acid decarboxylase, 757F  ARS (autonomously replicating sequences), 888  Arsenicals, 561  Arsenic poisoning, 561  Arsenite, 561   a helix/(3 sheet propensities, 14IT  biosynthesis, 742-743, 742F  breakdown, 727F, 730-731  genetic code specification, 967T  ionizable groups, 79T  and AGinked oligosaccharides, 234-236  side chain hydropathy, 157T  structure, 79T   uncharged polar side chain, 81  Asparagine synthetase, 742F, 743  Aspartame, 225, 740  Aspartate, 82   amino acid biosynthesis from, 746-748, 747F  from amino acid degradation, 719, 727F  in ATCase reaction, 377F  biosynthesis, 742-743, 742F  breakdown, 730-731  charged polar side chain, 82F  reaction with carbamoyl phosphate, 377  in urea cycle, 723, 724F, 726F  Aspartate aminotransferase, 542  Aspartate transaminase (AST), 720  Aspartate transcarbamoylase (ATCase), 377-381  binding sites, 378-380  conformational changes, 380F  feedback inhibition, 377-378  and hemoglobin vs. other enzymes, 381  regulation of nucleotide biosynthesis, 802, 803  structural changes in, 380-381  in UMP synthesis, 800, 80IF  v Q and aspartate concentration, 377F  X-ray structure, 379F  Aspartic acid (Asp):   acid-base catalysis by, 325  a helix/(3 sheet propensities, 14IT  charged polar side chain, 82  covalent catalysis by, 328  genetic code specification, 967T  ionizable groups, 79T  side chain hydropathy, 157T  structure, 79T  Aspartic protease, 370  Aspartokinase, 747  Aspartyl phosphate residue, 305  Aspartyl-(3-phosphate, 747, 747F  Aspartyl-tRNA synthetase (AspRS), 972,   973, 973F  Aspirin, 698, 699  Assays, protein, 97-98  Assimilation, 758, 762-764  Association studies, genome-wide, 1017  AST (aspartate transaminase), 720  Asthma, 400, 425  Asturias, F., 1045  Asx, 83    Atherosclerosis, 707-708  ATM (ataxia telangiectasia), 6IT, 1062  Atomic force microscopy (AFM), 232  Atorvastatin (Lipitor), 706F  ATP (adenosine triphosphate), 43  AMP-to-ATP ratio, 780  ATCase inhibition by, 377, 380-381  biological importance of, 447, 448  in Calvin cycle, 644, 645F, 646  chemical potential energy from, 43  as citric acid cycle regulator, 572  coupled reactions involving, 449, 450F  dissipation of, in dark reactions, 624, 646,   DNA ligase activation by, 881-882  and electron transport, 604-607, 604F  and free energy of phosphate hydrolysis, 448T  and G-actin, 20IF  in galactose metabolism, 505F  in gluconeogenesis, 539F, 540F, 543  in glycolysis, 474F, 475-476, 478, 485-486,  488-491   and GroEL/GroES, 167-169  in group transfer, 448-449  and high-/low-energy phosphate  compounds, 452F  inhibition of PFK by, 499-500  K y for segments of, 372  in mannose metabolism, 506F  metabolic role overview, 439F  and muscle contraction, 496, 771, 77IF  PGK and 3PG in complex with, 485F  production control, 614, 616  production in light reactions, 624, 630, 642-643  in purine biosynthesis regulation, 798  regeneration, 452-453  and standard free energy, 449  structure, 448F  ATPase(s), 166-167, 201  A-type, 304  Ca 2+ -, 306-308  F-type, 304  P-type, 304   (Na + -K + )-, 304-306, 304F, 309, 310  V-type, 304   ATP-cassette binding protein A1 (ABCA1), 729  ATP-citrate lyase, 573, 681  ATP-driven active transport, 304-309  ATP driven complexes, 1033  ATP hydrolysis, 304, 306   and nitrogenase conformation, 760-761  by type II topoisomerases, 837  ATP mass action ratio, 616  ATP synthase (F 1 F 0 -ATPase), 603-604, 607-612  binding change mechanism, 609—610, 61 OF  F 0 component, 608-609  F x component, 607-608, 608F  of mitochondria vs. chloroplasts, 642-643  as rotary engine, 610-612  Atractyloside, 584, 585  ATR protein kinase, 1063-1064  Attenuators, 1028-1030  otto- (prefix), 12  Attmctylis gummifera, 585  A-type ATPases, 304  AU-rich elements (AREs), 1050  Autocatalytic trypsinogen activation, 349  Autoimmune diseases, 213 , 213T  Autoinhibitor segments, 423   3'-Azido-3'-deoxy thymidine (AZT, zidovudine), 370  Azotobacter vinelandii, 759, 759F, 965  AZT (3'-Azido-3'-deoxythymidine, zidovudine), 370  Azurin, 142F   BACs (bacterial artificial chromosomes), 63, 66  Bacteria, 9F. See also specific species  cell walls, 231 - 233 , 232F, 233F  evolutionary studies, 9-10  fatty acids, 243   lipid bilayer fluidity modification, 257  lipid biosynthesis, 27IF  lysozyme action, 333  microfossil, 2F  tetracycline-resistant, 1001  transformed pneumococci, 49F  Bacterial artificial chromosomes (BACs), 63, 66  Bacterial biofilms, 230  Bacterial oxalate transporter, 303  Bacteriochlorophyll a (BChl a), 626, 626F  Bacteriochlorophyll b (BChl b), 626, 626F  Bacteriophages, 52  Bacteriophage 434, 852, 852F  Bacteriophage X, 63-65, 63F, 65F  Bacteriophage SP01, 924  Bacteriophage T4, 884, 963  Bacteriopheophytin (BPheo), 630  Bacteriorhodopsin, 260-262   proton pumping in, 594-595, 594F  structure, 260-261, 260F  Baculoviruses, 63  Bad cholesterol (LDL), 707  Baker, D., 165   Base excision repair (BER), 897 , 898  Base flipping, 896  Base pairs (bp), 47, 824F, 839  Basic helix-loop-helix (bHLH), 706, 857  Basic region leucine zipper (bZIP) proteins, 856,  857, 857F    Basic solutions, 31  Basolateral domain, 272  Bassham, J., 644  bed (bicoid ) gene, 1070, 107IF  B cells (B lymphocytes), 208, 1032F  BChl a (bacteriochlorophyll a), 626, 626F  BChl b (bacteriochlorophyll b), 626, 626F  BCKDH (branched-chain a-keto acid  dehydrogenase), 565, 757  Bcl-2 family, 1066  Bcr protein, 414  B-DNA, 822 - 827 , 825T   conformation, 822, 824, 826F-827F  dimer stacking energies, 840T  Beadle, G., 49   BER (base excision repair), 897 , 898  Berg, P, 973  Berger, J., 836, 837, 928  Beriberi, 494, 495  Berman, FT, 138, 1027  Bernal, J. D., 127, 182  Berson, S. A., 399  Berzelius, J., 76  Best, C., 462, 787, 788  0a 0 motif, 148  (3 anomers, 220, 22IF  (3 band (Soret band), 596  (3 barrels, 149 - 150 , 150F  of porins, 291, 292  in transmembrane protein, 262-263  (3 bends, 136  (3 bulge, 140  (3-clamp, 880, 880F, 881  (3 hairpin motif, 148  (3 keratins, 136, 138  (3 oxidation, 665, 667 - 669 , 671 F  in mammalian metabolism, 768F, 769  in mitochondria vs. peroxisomes, 677—678  (3PP (A(3 precursor protein), 170  (3 propeller, 421  (3 proteins, 149  (3-secretases, 170   147F, 172  (3 subunit, 880  Beta thalassemia, 6IT  bn (cytochrome b^fi, 148, 598  BHA, 284-285, 284F, 285F  bHLH (basic helix-loop-helix), 706, 857  bHLH/Z protein, 857  Bibliome, 465  Bicarbonate, 189  Bicoid {bed) gene, 1070, 107IF  Bicoid protein, 1070  Bidirectional replication, 869  Bilayers, 28   Bile acids, 658, 658F, 704  Bile salts, 658, 659F  Bilirubin, 754-756, 755F  Biliverdin, 754, 755, 755F   Bimolecular nucleophilic substitution reactions, 330  Bimolecular reactions, 329-330, 356  Binding affininty, protein, 99   Biochemical signaling, 396-434. See also Hormones  and heterotrimeric G proteins, 417-426  and metabolic regulation, 779-780  and phosphoinositide pathway, 426-433  and receptor tyrosine kinases, 402-417  Biochemistry, 1, 12  Biocytin, 540  Bioethics, 71  Biofilms, 230, 230F  Bioinformatics, 152-155  Biomolecule bond energies, 25T  Biopterin, 739, 739F  Biosphere, energy flow in, 18F  Biosynthetic pathways, 439  Biotin, 467, 540, 540F  Biotinyllysine, 540, 540F   free energy of phosphate hydrolysis, 448T  in gluconeogenesis, 539F  in glycolysis, 474F, 483-485   and blood oxygen carrying capacity, 488  deoxyhemoglobin binding by, 190, 190F, 191  D-2,3-Bisphosphoglycerate (BPG), 190, 191  Bisphosphoglycerate mutase, 488   2,3-Bisphosphoglycerate phosphatase, 488  Bisubstrate reactions, 366 - 367 , 366F  Bithorax complex ( BX-C ), 1072  Bithorax mutant {bx), 1070  (cytochrome b 598  Black, J., 817  Blackburn, E., 889  Blastocyst, 1042  Blastoderm, 1068  Blobel, G., 273  Bloch, K., 680, 701  Blood:   allowable levels, 770  and diabetes mellitus, 786-789, 787F  and glucagon secretion, 773  and gluconeogenesis, 544F  and liver glucose conversion to G6P, 772, 772F  and starvation, 785  Bloodstream, 769, 772, 773  Blood types, 237-238  Blow, D., 341  Blue copper proteins, 639F  Blunt ends, 54  Blunt end ligation, 882  B lymphocytes (B cells), 208, 1032F  BMD (Becker muscular dystrophy), 203  Bohr, C., 189   Bohr effect, 188-190, 189F  Boltzmann constant {k B ), 12, 13, 363  Bond energies, 25T   Bond rotation, DNA, 828-830, 828F, 829F  Bongkrekic acid, 585  BoNT/A, 282  BoNT/G, 282   Bovine cytochrome c oxidase, 60IF  Bovine F^ATPase, 608F  Bovine growth hormone, 69T  Bovine insulin, 94F, 106  Bovine pancreatic RNase A, 325-327, 327F  Bovine pancreatic RNase S, 325F  Bovine pancreatic trypsin inhibitor (BPTI), 347, 347F  Bovine papillomavirus, 877, 877F  Bovine protein kinase A, 423F  Bovine spongiform encephalopathy (BSE, mad cow  disease), 170, 171  Boyer, H., 64  Boyer, P., 609  bp, see Base pairs   BPG (D-2,3-bisphosphoglycerate), 190, 191  BPheo (bacteriopheophytin), 630  B proteins, 948, 948F  BPTI (bovine pancreatic trypsin inhibitor),   blood-brain barrier, 387  mammalian metabolism in, 769 - 770 , 770F  Branched-chain amino acid degradation,   Branched-chain a-keto acid dehydrogenase  (BCKDH), 565, 757  Branching enzyme (amylo-(l,4—>1,6)-  transglycosylase), 524, 529-530  Branch migration, 903  Branden, C.-I., 647   Butyryl-ACP, in fatty acid synthesis, 685F  BX-C (bithorax complex), 1072  hx {bithorax) mutant, 1070  bZIP proteins, see Basic region leucine zipper  proteins   and calmodulin, 429, 430  citric acid cycle control, 570, 572  and G-actin, 20IF   and glycogen breakdown control, 532, 533  with metal-activated enzymes, 328  with proteins, 95  and tropomyosin, 206F  vitamin D control of, 251—252  Ca 2+ -ATPase (Ca 2+ pump), 306-308  Cadmium (Cd 2+ ) ion, Zn 2+ ion replacement, 319  Caenorhabditis elegans, 59T, 60, 408, 1051,   Cahn-Ingold-Prelog (RS) system, 86  CAIR (Carboxyaminoimidazole ribotide), in IMP  synthesis, 795 F, 796  Cairns, J., 869, 893  CAK (Cdk-activating kinase), 1060  Calcineurin (PP2B), 417  Calcium, muscle contraction and, 204—206,  204F-205F   644 - 651 , 644F-645F  carbon dioxide fixation in, 644-648  control, 649-651   free energy changes for reactions, 650T  starch, sucrose, and cellulose from, 648-649  CaM (calmodulin), 428 - 430 , 533  CAM (crassulacean acid metabolism), 654  Cambillau, C., 658  Camels, 657   cAMP-phosphodiesterases (cAMP-PDEs), 425  cAMP receptor protein (CRP), 1026, 1027F  Camptothecin, 838  Cancer, 473    L-asparaginase as anticancer agent, 731  and fatty acid synthase expression, 687  and gangliosides, 249  and gene expression, 1061—1064  and oncogenes, 410  and telomerase, 891  thymidylate synthesis inhibition, 813  and tumor suppressors, 1061-1064  type II topoisomerases as anticancer agent, 838  Candida albicans, 578  Cantor, C., 847   Cap structure, eukaryotic mRNA, 942, 943F  CapZ, 203  Carazolol, 418  Carbamate, 190, 725, 725F  Carbamoyl aspartate, 377, 800, 80IF  Carbamoyl phosphate, 447  reaction with aspartate, 377  in UMP synthesis, 800, 80IF  in urea cycle, 723, 724F, 725 F  Carbamoyl phosphate synthetase (CPS), 723-725,  724F, 725F   Carbamoyl phosphate synthetase II (CPS II):  regulation of pyrimidine biosynthesis, 802, 803  in UMP synthesis, 800, 80IF  Carbinolamine, 324, 72IF  Carbohydrates, 217-238  defined, 217   energy recovery by citric acid cycle, 552  Fischer convention: D and L sugars, 219  in gangliosides, 249F  glycoproteins, 230-238  metabolism of, in liver, 773  monosaccharides, 218-223  and obesity, 790   from photosynthesis, 623, 644-654  polysaccharides, 224-230  and recognition events, 237  starvation and degradation of, 785  Carbohydrate recognition markers, 280  Carbon:  a, 77, 85   from C 4 cycle, 653-654  from citric acid cycle, 555  fixation of, 644-648  permeability of, 584  from photorespiration, 651-654  in photosynthesis, 623, 624, 644-648  transport of, 189-190, 189F  Carbon fixation, 623  Carbonic anhydrase, 189  catalytic power, 316T  Michaelis-Menten parameters, 362T  Zn 2+ role in, 329F  Carbon monoxide dehydrogenase, 673  Carbonyl carbon atom, 328F   2-Carboxyarabinitol-l,5-bisphosphate, 647F  2-Carboxyarabinitol-l-phosphate (CA1P), 650  Carboxyatractyloside (CATR), 585, 586F  y-Carboxyglutamate, 88 F  Carboxyl group, 26F  Carboxylic acids, 3 F, 4T  Carboxyl terminus, see C-terminus  Carboxymethylcysteine, 483   Caspases (cysteinyl aspartate-specific proteases),  1065-1066  Caspase- 1 , 514  Caspase-3, 1066  Caspase-7, 1065-1066, 1065F  Caspase- 8 , 1066  Caspase-9, 1068  Caspase-10, 1066  Caspase-activated DNase, 1066  Cassette exons, 951   purine ribonucleotides, 814 - 816 , 814F  thermodynamics, 444  Catabolite gene activator protein (CAP),   1026 - 1028 , 1027F  Catabolite repression, 1026-1028  Catalase, 317, 362T, 620  Catalysis, 329-331  Catalysts, 6 , 19   chymotrypsin, 344 - 349 , 344F  of DNA polymerase, 875  enzymes, 323-332  lysozyme, 335-338  Catalytic perfection, 482   CATH (computer program), 153T, 154  Cathepsins, 713  Cathepsin D, 280  Cations, 840    Cationic imines, 328F  CATR (carboxyatractyloside), 585, 586F  Cbl-associated protein (CAP), 432F  CCA-adding polymerase, 959  CCAAT box, 936  C a —C p bond, 729-730  CCVs, see Clathrin-coated vesicles  Cd 2+ ion, Zn 2+ ion replacement, 319  Cdks (cyclin-dependent protein kinases),  1060-1061, 1061F  Cdk2, 1060, 106 IF  Cdk4/6—cyclin D complexes, 1064  Cdk 7 , 1060   Cdk-activating kinase (CAK), 1060  cDNA (complementary DNA), 66 , 886  cDNA libraries, 66  CDP-diacylglycerol, 696, 697F  CDP-glucose, 546   CE (capillary electrophoresis), 103-104  CE (computer program), 153T, 154  Cech, T., 955, 956  Celecoxib (Celebrex), 386, 389, 699  Cells:   lysing of, in protein purification, 96  membranes and shape of, 267-269  metabolic pathways in, 441-443  metabolic reactions of, 5-7  microfilaments in, 206-207  prokaryotic and eukaryotic, 7-8  Cell-cell recognition, 249  Cell cycle, 715, 858, 1059-1061  Cell membranes, 8 F  Cell nucleus, 8 F  Cellular immunity, 208  Cellulase, 226   Central pores, of voltage-gated cation channels,  298-299  Centrioles, 8 F  Centromeres, 1019  Ceramides, 248, 694, 698F, 700  Cerebrosides, 248, 697, 698  Cer-phosphocholine, 700  CFI (cleavage factor I), 943  CF^Fq complex, 643  CFII (cleavage factor II), 943  CFTR (cystic fibrosis transmembrane conductance  regulator), 309   C H 1, 210  C h 2, 210  C h 3, 210   in polypeptide synthesis, 985  ribosomal binding sites for, 999F  RNA polymerase in, 924 - 927 , 925F  Chain initiation, in polypeptide synthesis,  986 - 991 , 989F   Chair conformation, 334F  Chamberlin, M., 927  Chang, G., 308  Changeux, J.-P., 192  Channels, 290   Complex II, 595, 596  muscle troponin, 203F  Chimera, 63  Chiral centers, 85  Chirality, 85   Chk2 (activated protein kinase), 1062  Chi a (chlorophyll a), 626, 626F  Chi b (chlorophyll b ), 626, 626F  Chloramphenicol, 1001   Chloramphenicol acetyltransferase (CAT), 1043  Chloride ions (Cl - ), 298  Chlorocruorins, 181  Chlorophylls, 625-626, 625F, 626F  absorption spectra, 626F  antenna, 626F  electronic states, 629F  Chlorophyll a (Chi a), 626, 626F  Chlorophyll b (Chi b), 626, 626F  Chloroplasts, 8 , 624-628, 624F  evolution, 10   light-absorbing pigments, 625-628  oxygen generation, 637F  oxygen yield per flash, 636F  thylakoid membranes, 624-625, 633F  Chloroquine, 38 5F  Chocolate, 425   Cholecalciferol (Vitamin D 3 ), 252  Cholera toxin, 249, 425  Cholesterol, 249, 250F  biosynthesis, 700-706  isotopic tracer studies, 463  LDL and HDE, 729F  in lipid rafts, 272   ATP-driven complexes, 1033  bromodomains, 1038-1039  chromatin-remodeling complexes, 1033-1034  and genomic imprinting, 1043  and histone acetyltransferases, 1036-1038  and histones, 859, 1036, 1039-1040  and HMG proteins, 1034-1036  and methyl transferases, 1040-1042  nucleosomes in, 859F   self-perpetuating DNA methylation, 1042-1043  structure of, 861 - 863 , 1032-1043  Chromatin map, 1037F   Chromatin-remodeling complexes, 1033 - 1035 , 1035F  Chromatography, 99-102  affinity, 102 , 102F, 846-847  gel filtration, 101   high-performance liquid (HPLC), 99  hydrophobic interaction, 100  immunoaffinity, 102  ion exchange, 99 - 100 , 100F  metal chelate affinity, 102  molecular sieve, 101  nucleic acids, 846-847  proteins, 99  size exclusion, 101  Chromodomains, 1040  Chromophore, 98  Chromosomes, 47, 858F  end-to-end fusion of, 891  eukaryotic, 858-863   histone-depleted metaphase, 861-863, 862F, 863F  histones, 858 - 861 , 858T  insertions and deletions, 913  nucleosomes, 859-861  rearrangements, 913, 915F  recombination, 915F  replication of linear, 889, 889F  Chromosome inactivation, 1033  Chronic myelogenous leukemia (CML), 414  Chylomicrons, 680-681, 785  Chylomicron remnants, 661  Chymotrypsin, 1043   active site, 339, 340, 343F  active site residues, 342F  catalytic mechanism, 344 - 349 , 344F  function of, 339  geometric specificity, 318-319  Michaelis-Menten parameters, 362T  polypeptide degradation, 718  specificity, 111T, 342, 342F, 343  tosyl-L-phenylalanine chloromethyl-ketone  binding, 340, 34IF  X-ray structure, 340-345  Chymotrypsinogen, 157F, 342, 350  Ciechanover, A., 714  Cigarette smoking, 348   in amino acid degradation, 727F  in citric acid cycle, 553F, 563-564, 568F  in glyoxylate cycle, 575 F  Citrate synthase:   in citric acid cycle, 553F, 561 - 563 , 563F,  570-572   conformational changes, 562F  in glyoxylate cycle, 575 F  Citric acid cycle (Krebs cycle; tricarboxylic acid  cycle; TCA cycle), 440  acetyl-CoA synthesis, 555-561  aconitase in, 553F, 563 - 564 , 571  amino acid degradation, 727F  amphibolic functions, 572, 573F  citrate synthase, 553F, 561 - 563 , 570-572  coordinated control, 617F  electron transport sites, 582F  energy-producing capacity, 568  enzymes of, 561 - 568 , 570-572  evolution, 576-577  free energy of reactions with, 570T  fumarase in, 553F, 567  and gluconeogenesis, 540, 541  glyoxylate cycle, 574-578  and glyoxylate cycle, 575F, 576  isocitrate dehydrogenase in, 553F, 564 , 565F,   malate dehydrogenase in, 553F, 567-568  in mammalian metabolism, 768F, 769, 773  net reaction, 554  overview, 552-555   pathways using citric acid cycle intermediates,  572-573  products of, 568F  and purine nucleotide cycle, 815  and pyruvate dehydrogenase, 553F, 555 - 560 ,  569-570   reactions of, 553F, 572-578  regulation, 57IF  regulation of, 568-572  related reactions, 572-578   replenishment of citric acid cycle intermediates, 574  succinate dehydrogenase in, 553F, 567 , 567F  succinyl-CoA synthetase in, 553F, 565 - 566 , 566F  Citrulline, 724F, 725-726, 726F, 757F  Citrullyl-AMP, 726F  Citryl-CoA, 562   Cl (chloride ions), 292  C L (constant region), 210  C x , 1057   Clathrin-coated vesicles (CCVs), 278 - 280 , 279F, 663  Clay, 3   Cleavage factor I (CFI), 943  Cleavage factor II (CFII), 943  Cleland, W. W., 367  Clinical trials, 387-389  Clones, 62, 63  Cloned DNA, 62-67  creating, 62-64  in DNA libraries, 66-67  and selection, 64-66  Cloning, 52   shotgun, 66  techniques, 62-66  Cloning vectors, 63  Closed complex, 921  Closed systems, 18  Clostridium botulinim, 892  Clostridium species, 282  Clp, 717  ClpA, 718  ClpP, 718, 718F  ClpX, 718   CML (chronic myelogenous leukemia), 414  CMP (cytidine monophosphate), 42T, 818F  Cn3D (computer program), 153T, 155  Co 2+ , as cofactor, 328  CoA, see Coenzyme A  Coactivators, 1038  Coagulation cascade, 350-351  CoASH, see Coenzyme A  Coated vesicles, 277, 277F  clathrin, 278 - 280 , 279F, 663  fusion with plasma membrane, 278F  protein transportation, 277-278  Coatomer, 278  Cobalamin, 672  Cobalamin coenzymes, 673  Cobra venom enzyme, 247F  Cochaperones, 166  Cockayne syndrome (CS), 899  Coding (sense) strand, 922, 922F, 923, 923F  Codons, 50F, 963, 967T   993-994, 994F  evolution, 966  frequently used, 975  nonsense, 966  phenotypically silent, 966  Start, 966, 967   Stop, 951, 966, 967T, 1003, 1004  synonyms, 899  Coenzymes, 319, 438-439  Coenzyme A (CoA; CoASH), 440, 447, 455,   590F, 592F  Coenzyme Q 6 , 592  Coenzyme Q 8 , 592  Coenzyme Q 10 , 592  Coenzyme QH*, 592F  Coenzyme QH 2 , 592F, 606  Cofactors, 319 - 320 , 319F, 328-329  Cohen, P., 723  Cohen, S., 64  Coils, polypeptides, 140  Coiled coil structure, 137, 137F  Cointegrate, 913, 914F  Colipase, 658   Colony-stimulating factors, genetically engineered, 69T  Combinatorial chemistry, 386-387, 387F  Committed step (metabolic pathways), 444  Compartmentation, cells, 6  Competition, 303   Competitive enzyme inhibition, 368 - 373 ,  372F-373F, 374T  degree of, 371, 372  HIV enzyme inhibitors, 370-371  measuring K Y , 372-373  Competitive inhibitors, 368, 370-372  Complementarity, 3F, 5, 5F, 317  Complementary base pairing, DNA, 47F  Complementary DNA (cDNA), 66, 886  Complex I (NADH-coenzyme Q oxidoreductase),  588, 590 - 595 , 591F  and bacteriorhodopsin, 594-595  coenzymes of, 591, 592  electrons in, 592, 593  reduction potentials, 590T  translocation of protons by, 593-594  X-ray structure, 607F   Complex II (succinate—coenzyme Q oxidoreductase),  588-590, 590T, 595 , 595F, 596  Complex III (coenzyme Q-cytochrome c   oxidoreductase; cytochrome bci), 596 - 600 , 597F  and cytochrome c, 600  electrons from, 598-600  return of electrons by, 632  sequence of operation, 588, 589, 590T  Complex IV (cytochrome c oxidase), 600 - 603 ,   reduction potentials, 590T  sequence of operation, 588, 589  Complex systems, emergent properties of, 431-433  Complex V (ATP synthase; proton pumping ATP  synthase; F^Q-ATPase), 603-604, 607-612  Composite transposons, 912-913, 913F  Concanavalin A, 135F   Concerted acid-base catalyzed reactions, 323  Condensation reactions, 3, 80, 80F  Congenital erythropoietic porphyria, 754  Conjugate acids, 32  Conjugate bases, 32  Conjugate redox pair, 458  Connexins, 300, 301  Connexin 26, 301    Connexons, 301  Consensus sequence, 923, 946F  Conservative replication, 867, 868  Conservative substitution, 118  Constant region, 208-210  Constitutive enzymes, 926  Constriction region (aquaporin), 300  Contiguous genes, transcription of, 926, 927F  Contours (in electron density maps), 143  Contour length, 858   Controlled rotation mechanism, 834, 835  Convergent evolution, 345  Cooperative oxygen binding, 184-193  Cooperativity:   hemoglobin oxygen binding, 186-193  protein denaturation, 159  of transcription factors, 1044-1045  Cooperman, B., 807  COPI, 278  COPII, 278   Copper ions, see Cu ions  Coproporphyrinogen III, 753F  Coproporphyrinogen oxidase, 753, 753F  CoQ, see Coenzyme Q  Cordycepin, 960  Core, lac repressor, 1025  Core enzyme, 920, 924  Core particles:   of nucleosomes, 859, 859F, 860F  posttranslational modifications of, 1036F  Corepressors, 1028  Core promoter element, 935, 936  Core proteins, 231  Corey, R., 129, 132, 134  Cori, C., 520, 774  Cori, G., 520, 774  Cori cycle, 774 , 774F, 775  Cori’s disease, 524, 525  Corn:   chloroplasts of, 624F  evolution of, 61, 61F  transgenic, 71  transposition in, 911  Correlation spectroscopy (COSY), 145  Corrin ring, 672  Cortex, adrenal gland, 398  Cortisol, 250, 25IF, 400  Cosubstrates, 319   of glycogen phosphorylase, 520, 532  of glycogen synthase, 530 , 531 , 535-536  for metabolic flux control, 445  and protein phosphorylation, 381-385  of proteins, 95, 1006-1009  of pyruvate dehydrogenase, 569-570  COX, 698  Cox, M., 909  COX-1, 699  COX-2, 699  COX-2 inhibitors, 699    COX-3, 699  Coxibs, 699  CP43 (PsbC), 634  CP47 (PsbB), 634  CpG islands, 894, 935, 1016, 1042  CPS, see Carbamoyl phosphate synthetase  CPSF (cleavage and polyadenylation specificity  factor), 943  Cramer, W., 638  c -ras gene, 410   Crassulacean acid metabolism (CAM), 654  Crawling cell, 207, 207F  Creatine kinase, 453, 499-500  Creatine phosphate, 447  CREB (cyclic AMP response element-binding  protein), 161-162, 162F  Crick, F., 45, 50, 127, 199, 822, 823, 867, 919,  963, 966   CTD, see C-terminal domain  CTD kinases, 932  CTD phosphatases, 932  C-terminal a helix, lac repressor, 1025  C-terminal cytoplasmic domain, 296  C-terminal domain (CTD), 932, 933, 941-943, 947  C-terminus (carboxyl terminus), 80  dynamics, 161   CTP (cytidine triphosphate), 378F  ATCase inhibition, 377-378, 380-381  synthesis, 802 , 802F  CTP synthetase, 802, 802F  c-type cytochromes, see specific cytochromes  Cu A center, cytochrome c oxidase, 601  Cu B , cytochrome c oxidase, 601  Cu ions:   as cofactors, 319, 328  Cu A center, 601  Cu B , 601  Curie, E., 399  Curie, M., 399   CybL (Complex II subunit), 595  CybS (Complex II subunit), 595  Cyclic AMP response element-binding protein  (CREB), 161-162, 162F  Cyclic symmetry, 156, 156F  Cyclins, 715, 1060   Cys 2 -His 2 zinc finger, 854 - 856 , 854F   acid-base catalysis by, 325  a helix/(3 sheet propensities, 14IT  biosynthesis of, 746  breakdown of, 727-730, 727F, 728F  covalent catalysis by, 328  disulfide bonds, 8 IF   genetic code specification for, 965, 967T  ionizable groups of, 79 T  in methionine breakdown, 732, 733F  as rare amino acid, 95  side chain hydropathy, 157T  structure of, 79 T  Cysteine proteases, 1065   Cystic fibrosis transmembrane conductance regulator  (CFTR), 309  Cystine, 81  Cyt, see Cytosine  Cytidine, 42T, 818F  Cytidine deaminase, 818F, 952  Cytidine monophosphate (CMP), 42T, 818F  Cytidine triphosphate, see CTP  Cytochromes, 320, 596-597  Cytochrome a, 601  Cytochrome 601  Cytochrome b, 596, 598  Cytochrome b 6 , 638   Cytochrome ^/complex, 633, 637-639, 638F  Cytochrome £ 559 , 634  Cytochrome b 560 , 595  Cytochrome b^ 2 (^h)> 148, 598  Cytochrome b 566 (£ L ), 598  Cytochrome bc x , see Complex III  Cytochrome c, 151 - 152 , 152F, 588, 597F, 1066  evolution of, 117-118, 121, 121F  isoelectric point, 99 T  occurrence, 151   as peripheral membrane protein, 264  phylogenetic tree for, 118 - 119 , 119F  reduction potentials, 590F  ribbon diagram, 600F  of selected species, 116T-117T, 117-118  side chain location, 147F  as soluble electron carrier, 600  Cytochrome c Y , 596  Cytochrome c 2 , 632  Cytochrome d, 606  Cytochrome f, 638  Cytochrome P450, 389 - 391 , 389F  Cytoglobin, 179    Cytokines, 411, 1046-1048  Cytokine receptors, 1046, 1047  Cytoplasm, 7, 953  Cytoplasmic domains, 308  Cytosine (Cyt), 41, 42T, 1041  base pairing, 47F, 824F, 839, 839F  and Chargaff’s rules, 45  as common nucleotide, 42  and hydroxylamine, 917  modified forms in tRNA, 969F  oxidative deamination, 892F  Cytoskeleton, 8  Cytosol, 8   acetyl-CoA transport from mitochondria, for fatty  acid biosynthesis, 681 , 682  heme biosynthesis in, 753F  metabolic functions, 442F   metabolite transport between mitochondria and,  in gluconeogenesis, 542, 542F, 543  Cytosolic (outer) faces, 277  Cytosolic palmitoyl-CoA, 781  Cytosolic reducing equivalents, 584   D (dalton), 12  D (Fischer convention), 85  D1 (PsbA), 634  Di proteins, 948  D2 (PsbD), 634  D 2 proteins, 948  D 3 proteins, 948, 948F  DAG, see 1,2-Diacylglycerol  Dalgarno, L., 987  Dalton (D), 12  Dam methyltransferase, 894  Danio rerio, 59T  Dansyl chloride, 106, 108, 109F  Dark reactions (photosynthesis), 624, 644-654  Calvin cycle, 644-651  carbon dioxide fixation, 644-648  control of Calvin cycle, 649-651  overview, 644F   and photo respiration, 651-654  starch, sucrose, and cellulose from, 648-649  D Arm (tRNA), 968, 969F  Darnell, J., 1046  Darst, S., 921  Darwin, C., 10, 61, 561  Databases:   of nucleotide sequences, 57 , 58  of protein and nucleic acid sequences,   114-115, 115T  Davies, D., 750  Davis, R., 64  Dawkins, R., 10  Dayhoff, M., 119  Dbp5, 953   ddC ( 2 ', 3 '-dideoxycytidine, Zalcitabine), 370  ddl (2',3'-dideoxyinosine, Didanosine), 370  ddNTP (2',3'-Dideoxynucleoside triphosphate), 56  Deadenylases, 1050   5- Deazatetrahydrofolate (5dTHF), 796  Debranching enzyme, 228, 519, 522  DEBS (deoxyerthronolide B synthase), 688  Decapping enzyme, 1050   Deep View (computer program), 154  Degeneracy, codon, 975  Degenerate code, 963, 966, 975  Degradative enzymes, 97  Degradative pathways, 440  7-Dehydrocholesterol, 252  Deletion mutations, 892, 893, 913  Demethylases, 1041  Denaturation:   DNA, 840 - 842 , 84IF  proteins, 94, 140, 159-160  Denaturing conditions, 258  Denitrification, 764  Density gradients, 105  5 , -Deoxyadenosylcobalamin, 672, 673F   6 - Deoxyerythronolide B ( 6 dEB), 688  Deoxyerythronolide B synthase (DEBS), 688  Deoxyhemoglobin, 182 - 183 , 183F   binding of BPG to, 190, 190F, 191  bisphosphoglycerate (BPG) binding to, 190, 191  ion pairs and hydrogen bonds, 188F  Deoxyhemoglobin S, 195F  Deoxyribonucleic acid, see DNA  Deoxyribonucleotides, 41, 4IF, 803-812  ribonucleotide conversion to, 804-808  thymine formation, 808-812  (3-D-2'-Deoxyribose, 41, 222, 804-808  Deoxy sugars, 222  Deoxythymidine, 42T, 818F  Deoxythymidine monophosphate, see Thymidylate  (dTMP)   Depolarization, 295  Depsipeptides, 931  Dermatan sulfate, 229, 229F  Desaturases, 689  Desensitization, 420  Detergents, 159, 258  Development:   of body parts, 1072-1073  embryo segments, 1068-1069  genetically mediated patterns, 1069-1071  Hox genes, 1072-1074  molecular basis of, 1068-1074  Dextrotatory molecules, 85  DHA (fatty acid), 243T  DHAP, see Dihydroxyacetone phosphate  DHF, see Dihydrofolate  DHFR, see Dihydrofolate reductase  Diabetes mellitus, 462, 786-789  1,2-Diacylglycerol (DAG), 247, 426, 658  activation of protein kinase C by, 430-431  in fatty acid synthesis, 694  in triacylglycerol biosynthesis, 69OF  Diacylglycerol acyl transferase, 690, 690F  Diacylglycerophospholipids, synthesis of, 694-697  Dialysis, 29, 29F  Diarrhea, 310  Diazotrophs, 758-759  Dicer, 1053, 1053F   Dicyclohexylcarbodiimide (DCCD), 621  2',3'-Dideoxycytidine (ddC, Zalcitabine), 370  2',3'-Dideoxyinosine (ddl, Didanosine), 370  Dideoxy method, 55 . See also Chain-terminator  method   Diet-induced thermogenesis, 783-784  Diffraction patterns, 142  Diffusion, 28-30  facilitated, 270, 290  lipid bilayers, 256, 256F, 265-267  thermodynamics, 289-290  Diffusion-controlled limit, 363  Digestion, 76  Digitalin, 306  Digitalis, 306, 385  Dihedral angles, polypeptides, 129  Dihedral symmetry, 156, 156F  7,8-Dihydrobiopterin, 740, 74IF  Dihydroceramide, 698, 698F  Dihydroceramide dehydrogenase, 698, 698F  Dihydrofolate (DHF), 734F, 809, 810, 81 OF  Dihydrofolate reductase (DHFR), 734,   Dihydrolipoamide, in citric acid cycle, 558, 558F  Dihydrolipoyl dehydrogenase (E3), 556, 556F,  559-560, 560F, 565   Dihydrolipoyl transacetylase (E2), 556, 556F, 559  Dihydrolipoyl transsuccinylase (E2), 565  Dihydroorotase, 800, 80IF  Dihydroorotate, 800, 80IF   Dihydropteridine reductase, 740  Dihydrosphingosine, 698  Dihydrouracil, 818F  Dihydrouridine, 968, 969F  Dihydroxyacetone, 219, 219F  Dihydroxyacetone phosphate (DHAP):  in Calvin cycle, 645F, 646  in fructose metabolism, 503F  in glycolysis, 474F, 478, 479  in triacylglycerol biosynthesis, 690F  Dihydroxyacetone phosphate acyltransferase,   Dihydroxythymine, in pyrimidine catabolism, 818F  Dihydropyrimidine dehydrogenase, in pyrimidine  catabolism, 818F  Diimine, 761   7V' 6 ,7V' 6 _Dimethyladenine, 954  Dimethylallyl pyrophosphate, 702, 703F  5-Dimethylamino-l-naphthalenesulfonyl chloride,  106, 108, 109F   Dinucleotide-binding fold, proteins, 151  Dipalmitoyl phosphatidylcholine (DPPC), 246, 256F  Dipeptides, 80   position determination, 114F  and protein folding, 158  Divergent evolution, 343, 482  Diversity, antibody, 1056-1059  Diversity segment, 1057  DMB (5,6-dimethylbenzimidazole), 672  DMD (Duchenne muscular dystrophy), 203  DMS (dimethyl sulfate), 924  DNA (deoxyribonucleic acid), 40-42.    UV absorbance, 841, 84IF  Watson-Crick base pairs, 824F  X-ray diffraction photograph, 45F  Z-DNA, 824-827, 825T, 826F-827F, 829  DnaA protein, 877  DnaB protein, 877  DNA chips, 66, 465-467, 466F  DNA circles:   DNA-directed DNA polymerases, 868. See also  DNA polymerases  DNA fingerprinting, 68, 69  DNA glycosylases, 897, 897F, 898  DnaG protein, 878  DNA gyrase, 835, 837, 877  DnaJ, 1005  DnaK, 1005  DNA libraries, 66-67  DNA ligase, 64, 870, 881-882, 882F  DNA loops, 861-863, 862F, 863F  DNA manipulation, 62-72  applications, 68-71  genomic libraries, 66-67  polymerase chain reaction, 67 - 68 , 68 F  DNA methylation, 1042-1043  DNA methyl transferases (DNA MTases), 894, 943,  1041, 1041F, 1042   DNA microarrays, 66, 465 - 467 , 467F  DNA photolyases, 896, 896F  DNA polymerases, 868F, 926  catalytic mechanism, 875  defined, 868   sensing of Watson-Crick base pairs, 874-875  DNA polymerase I, see Pol I  DNA polymerase II (Pol II), 875, 875T  DNA polymerase III, see Pol III  DNA polymerase IV, 901  DNA polymerase V, 901  DNA polymerase a, see Pol a  DNA polymerase y (pol y), 886  DNA polymerase r\ (pol T|), 900  DNA-protein interactions, 849-857   eukaryotic transcription factors, 854-857  prokaryotic repressors, 851-853  restriction endonucleases, 850 - 851 , 850F, 851F  DNA repair, 895-901   base excision repair, 897 , 898  direct damage reversal, 895-896  introduction of errors, 900-901  mismatch repair, 899-900  nucleotide excision repair, 898-899  and Pol I, 873   by recombination, 908 - 911 , 909F, 91 OF  SOS response, 901   DNA replication, 49, 49F, 822, 868-895  and complementarity, 5  and DNA damage, 891-895  eukaryotic, 884-891  overview, 868-870  prokaryotic, 870-884   recombination repair, 908 - 911 , 909F, 91 OF  replication forks, 869  RNA primers, 870, 872F  semiconservative, 867, 868  semidiscontinuous, 869-870  DNase, caspase-activated, 1066  DNase I, 873  dNDP, 804   DNFB (2,4-Dinitrofluorobenzene), 108  DNMTl protein, 1042  DNMT3a (DNA MTase), 1042  DNMT3b (DNA MTase), 1042  DNP (2,4-dinitrophenol), 614, 614F  dNTPs, 55, 67, 68F, 808, 884  Docking protein, 274   Dolichol-PP-oligosaccharide synthesis pathway, 547F  Dolichol pyrophosphate, 546-547F  Donohue, J., 45, 823   l-DOPA (dihydroxyphenylalanine), synthesis from  tyrosine, 756-757, 757F  Dopamine, 89, 89F, 756, 757F  Dopamine (3-hydroxylase, 757F  Dosage compensation, 1033  Double blind tests, 388  Double bonds, in fatty acids, 243  Double-displacement reactions, 367  Double helix, 45 - 48 , 45F, 822-838  Double-reciprocal plot, see Lineweaver-Burk plot  Double-stranded breaks (DSBs), 900-901,   alternative splicing, 951-952, 95IF  antenna and legs, 1072F  development, 1068 - 1074 , 1069F-1073F  differentiation of embryonic segments, 1073F  histone genes reiteration, 1018  insulators, 1046  micro RNAs, 1054  replication forks, 888F  RNA interference, 1052  Drosophila melanogaster:  calmodulin, 429F  chromatin, 859F  genome sequencing, 59T, 60  26S proteasome, 715F  Drug design, 385-391   adverse drug reactions, 389-391  bioavailability and toxicity, 387  clinical trials, 387-389  techniques for, 386-387  Drug-drug interactions, 389-390  Drug resistance, 308-309  Druker, B., 414   DSBs, see Double-stranded breaks  DSCAM protein, 951  dsDNA, see Double-stranded DNA  D segment, 1057   808-809, 808F  Dutzler, R., 298  Dwarfism, 401  Dystrophin, 203, 268, 945   E (exit site), 982, 1000  E. coli, see Escherichia coli  A.% (electrical potential difference), 458  (standard reduction potential), 
FundamentalsBiochemistry4e_201802 FundamentalsBiochemistry4e_djvu.txt