A mutation uncouples the tubulin conformational and GTPase cycles, revealing allosteric control of microtubule dynamics
Microtubules are dynamic polymers of αβ-tubulin that undergo GTPase-dependent switching between phases of growing and rapid shrinking ( reviewed in Desai and Mitchison , 1997 ) . Microtubule growing and shrinking rates , and switching frequencies , are necessary for proper function , highly regulated by cellular factors ( Howard and Hyman , 2003 ) , and targeted by widely used anti-mitotic anticancer drugs ( Jordan and Wilson , 2004; Amos , 2011 ) . αβ-tubulin also undergoes a conformational cycle during polymerization and depolymerization ( Mandelkow et al . , 1991a; Chrétien et al . , 1995 ) Unpolymerized αβ-tubulin adopts a 'curved' conformation ( Ravelli et al . , 2004; Rice et al . , 2008; Buey et al . , 2006 ) that is not compatible with the 'straight' microtubule lattice ( Nogales et al . , 1998 ) . In the body of the microtubule , αβ-tubulin can adopt distinct ‘expanded’ or ‘compacted’ straight conformations depending on nucleotide state ( Alushin et al . , 2014; Hyman et al . , 1995 ) . Partially straightened conformations of αβ-tubulin are thought to occur as polymerization intermediates on the microtubule end . Recent studies have begun to reveal how microtubule regulatory proteins take advantage of these different conformations . In one well-characterized example , end-binding proteins in the EB1 family have been shown to mark the plus-end cap of microtubules by binding preferentially to the ‘expanded’ , GTP-like conformation of αβ-tubulin that only occurs near the end of the growing microtubule lattice ( Maurer et al . , 2011; 2012; Zanic et al . , 2009; Alushin et al . , 2014 ) . Another recent example of selective interactions with specific conformations of αβ-tubulin comes from work in our laboratory showing that TOG domains from the microtubule polymerase Stu2p bind preferentially to the curved conformation of αβ-tubulin ( Ayaz et al . , 2012; 2014 ) , and that this preference of TOG domains for curved αβ-tubulin is probably what allows the polymerase to localize to the growing tip of the microtubule ( Ayaz et al . , 2014 ) . These and numerous studies of other regulatory proteins ( Desai et al . , 1999; Gigant et al . , 2000; Peters et al . , 2010; Alushin et al . , 2010; Bechstedt et al . , 2014 ) contribute to the emerging view that selective binding to distinct conformations of αβ-tubulin represents a common and important strategy for recognizing and controlling microtubules . How the αβ-tubulin conformational cycle contributes to microtubule dynamics is less understood . Indeed , even though structural rearrangements in αβ-tubulin have long been known to occur during and after microtubule incorporation ( Mandelkow et al . , 1991b; Chrétien et al . , 1995; Hyman et al . , 1995 ) , actually showing that these rearrangements contribute to and are required for the dynamic properties of microtubules remains challenging ( Rice et al . , 2008; Wang and Nogales , 2005; Buey et al . , 2006; Jánosi et al . , 2002; Molodtsov et al . , 2005 ) . For example , GTP-bound αβ-tubulin ‘straightens’ during the formation of longitudinal and lateral lattice contacts ( Buey et al . , 2006; Rice et al . , 2008; Nawrotek et al . , 2011 ) , but how ( or even if ) the energetic cost of these assembly-dependent conformational changes affects polymerization dynamics remains debated . A similar argument can be made about the αβ-tubulin conformational changes that occur as a consequence of GTP hydrolysis deeper in the microtubule lattice . One reason these questions are still debated is that to date in vitro microtubule dynamics have only been measured for a small number of αβ-tubulin mutants ( Gupta et al . , 2002; Dougherty et al . , 2001; Sage et al . , 1995 ) , and it has not yet been possible to link altered dynamics to a change in the αβ-tubulin conformational cycle . In the present study we sought to discover how multiple αβ-tubulin conformations affect microtubule dynamics by identifying mutations that selectively perturb some aspect of the conformational cycle . We reasoned that mutating buried residues with different packing environments in the curved and straight conformations might provide a way to perturb the conformational cycle . We focused on helix H7 because it is an element that is positioned differently in the ‘curved’ ( unpolymerized ) and ‘straight’ ( polymerized ) conformations of αβ-tubulin ( Nogales et al . , 1998; Ravelli et al . , 2004 ) ( Figure 1A ) , and it may act as a structural relay connecting the ‘top’ and ‘bottom’ of the tubulin subunits ( Amos , 2004 ) . The equivalent helix in α-tubulin was also recently observed to move in response to GTP hydrolysis in the lattice ( Alushin et al . , 2014 ) . We chose to study the T238A mutation in β-tubulin because this sidechain resides on helix H7 and is buried in the core of the protein ( Figure 1A ) , and because prior studies ( Thomas et al . , 1985; Machin et al . , 1995 ) showed that the β:T238A phenotype was consistent with hyperstable microtubules . The molecular mechanism by which the β:T238A mutation affects microtubule dynamics is unknown , in part because the polymerization dynamics of β:T238A αβ-tubulin have not been studied in vivo or in vitro . Our studies now reveal that the β:T238A mutation affects microtubule dynamics by dramatically reducing the frequency of catastrophe and the rate of shrinking , and that the mutation affects microtubule structure by attenuating post-GTPase conformational changes in the lattice . The effects of the mutation are reminiscent of the way the microtubule stabilizing drug taxol affects microtubule structure ( Alushin et al . , 2014 ) . By showing that the β:T238A mutation weakens the coupling between the conformational and nucleotide cycles , our results demonstrate that catastrophe can occur despite diminished post-GTPase conformational changes in the microtubule lattice , and that the strength of the allosteric response to GDP dictates the rate of microtubule shrinking and the frequency of microtubule catastrophe . 10 . 7554/eLife . 10113 . 003Figure 1 . αβ-tubulin containing a buried mutation in β-tubulin gives hyperstable microtubules in vivo and in vitro . ( A ) Superposition of polymerized ( ‘straight’ , green , PDB 1JFF ) and unpolymerized ( ‘curved’ , blue , PDB 4FFB ) conformations of β-tubulin . T238 is solvent inaccessible and resides on a helix ( H7 ) that undergoes a piston-like movement between the two conformations . The view of β-tubulin shown here is as if from the center of the microtubule looking out , with the plus end at the top . ( B ) Time-lapse imaging of live yeast shows that cells expressing T238A αβ-tubulin make longer , less dynamic microtubules than their wild-type ( WT ) counterparts . N = 13 , 18 for WT and T238A microtubules respectively , with t = 35 , 72 min total time observed . See also Videos 1 , 2 . Bar = 2 µm ( C ) Negative stain electron micrographs ( magnification: 23 , 000x ) of wild-type ( left ) and β:T238A ( right ) microtubules . Mutant microtubules have normal structure . Bar = 100 nm . ( D ) In vitro , T238A and wild-type microtubules show similar concentration-dependent growth rates ( slopes: 29 . 6 +/- 1 . 8 µm/hr/µM for wild-type , 25 . 5 +/- 0 . 8 µm/hr/µM for β:T238A , the differences between the slopes are not statistically significant; x-intercepts: 0 . 12 µM for wild-type , 0 . 033 µM for β:T238A , this difference is statistically significant; significance of differences in regression parameters was evaluated using GraphPad Prism ) . Representative kymographs are shown above the plots . ( N = 12 for all points except 0 . 2 µM where N = 20; bars show s . e . m . ) ( E ) T238A microtubules catastrophe less frequently than wild-type ( top; N = 99 , 115 for wild-type and T238A respectively , bars show s . d . ) and show a ∼hundredfold slower rate of post-catastrophe shrinking ( bottom; N = 16 for wild-type and T238A , bars show s . e . m . ) . DOI: http://dx . doi . org/10 . 7554/eLife . 10113 . 003
The β:T238A mutation was previously identified by virtue of drug- and temperature-sensitive phenotypes that were consistent with hyperstable microtubules ( Thomas et al . , 1985 ) . However , prior studies of this mutant have not directly examined its polymerization dynamics in vivo or in vitro ( Machin et al . , 1995; Dorn et al . , 2005 ) . To obtain insight into how the mutation affects polymerization dynamics in cells , we used time lapse imaging in GFP-Tub1p expressing strains to measure microtubule dynamics of wild-type and β:T238A-containing yeast . These measurements revealed that β:T238A-tubulin forms static microtubules ( neither growing nor shrinking ) that are on average over 50% longer than the dynamic microtubules that form in a strain with wild-type β-tubulin ( Figure 1B and Videos 1 , 2 ) . Because the mutated site is solvent inaccessible , this striking change in microtubule dynamics cannot be the result of a direct perturbation of a polymerization interface or of an interaction with one or more regulatory proteins . 10 . 7554/eLife . 10113 . 004Video 1 . Microtubule dynamics in wild type yeast . Time-lapse images of Tub1-GFP in TUB2 ( β-tubulin ) cells were taken at 15-s intervals; video plays at 4 frames/s . A frame from this video is shown in Figure 1B . DOI: http://dx . doi . org/10 . 7554/eLife . 10113 . 00410 . 7554/eLife . 10113 . 005Video 2 . Microtubule dynamics in tub2-150 ( T238A mutation in Tub2p ) yeast . Time-lapse images of Tub1-GFP in tub2-150 cells were taken at 15-s intervals; video plays at 4 frames/s . A frame from this video is shown in Figure 1B . DOI: http://dx . doi . org/10 . 7554/eLife . 10113 . 005 To determine how the buried β:T238A mutation affected microtubule dynamics in vitro , we purified β:T238A αβ-tubulin from an overexpressing strain of yeast ( Johnson et al . , 2011 ) and used time-lapse differential interference contrast microscopy to measure its polymerization dynamics . We were unable to measure mutant and wild-type microtubule dynamics at equivalent concentrations , because β:T238A αβ-tubulin showed abundant spontaneous nucleation at the higher concentrations where we measured wild-type , and wild-type αβ-tubulin does not elongate measurably at the low concentrations where we were able to measure β:T238A dynamics without excessive nucleation . Mutant and wild-type microtubules nevertheless show similar concentration-dependent elongation rates: fitting lines to mutant and wild-type data reveals that the x-intercepts of the two datasets ( 0 . 12 and 0 . 033 µM for wild-type and β:T238A , respectively ) differ by a factor of ∼3 . 5 and that the difference in slope ( 29 . 6 and 25 . 5 µm/hr/µM for wild-type and β:T238A , respectively ) is not statistically significant ( Figure 1D ) . Because the x-intercept and slope respectively relate to the apparent affinity and association rate constant for elongation , our data indicate that the mutation has little effect on the apparent biochemistry of microtubule elongation . Consistent with this biochemical similarity , negative stain electron microscopy revealed that mutant and wild-type microtubules show similar structure ( Figure 1C ) . In striking contrast to the shared elongation behavior , after catastrophe β:T238A microtubules shrink roughly hundredfold more slowly than wild-type ( 1 . 1 µm/min for β:T238A compared to 96 µm/min for wild-type , Figure 1E , bottom ) . Thus , the mutation significantly strengthens the lattice contacts that dictate the rate of microtubule shrinking . Finally , β:T238A microtubules also undergo catastrophe much less frequently than wild-type . The lower catastrophe frequency we observed is especially notable when considering that in these assays the T238A microtubules were growing much slower than wild-type because of the ∼threefold lower concentration of αβ-tubulin used for the mutant ( Figure 1E , top ) . The β:T238A mutation stimulated spontaneous nucleation and reduced the frequency of catastrophe and the rate of shrinking , all without substantially affecting elongation . It seemed possible that a defective GTPase cycle might explain these observations . We reasoned that if the increased spontaneous nucleation of the β:T238A mutant resulted from slower/defective GTPase activity , then both mutant and wild-type should nucleate with similar efficiency when GTP hydrolysis cannot occur . We initially attempted to use GMPCPP , the hydrolysis-resistant nucleotide of choice for vertebrate microtubules ( Hyman et al . , 1992 ) , but GMPCPP did not support elongation of yeast microtubules in our dynamics assays . Yeast microtubules polymerized readily in the presence of GTPγS , however , indicating that GTPγS better mimics GTP for yeast microtubules . We observed that even in the presence of GTPγS , wild-type microtubules show substantially less nucleation than T238A microtubules ( Figure 2A , B ) . Thus , the abundant nucleation from the mutant cannot be ascribed to a defect in GTPase activity . Instead , the mutation must be affecting some other property that limits spontaneous nucleation in wild-type αβ-tubulin . 10 . 7554/eLife . 10113 . 006Figure 2 . T238A αβ-tubulin undergoes spontaneous nucleation more readily than WT , even in the presence of a nonhydrolyzable GTP analog , GTPγS . ( A ) Fluorescent images of crosslinked microtubules from spontaneous nucleation reactions . Even at low concentrations and in the presence of GTPγS , T238A tubulin shows increased spontaneous nucleation compared to WT . GTPγS reactions are presented next to each other to facilitate a side-by-side comparison . Scale bar in top left is 5 µm . ( B ) Microtubule spindown reactions show that under the same concentration range , in the presence of GTPγS , T238A tubulin produces a greater proportion of microtubules which sediment into the pellet . Gel images of supernatant and pellet fractions ( top ) . ( C ) T238A microtubules do not accumulate GTP or GDP . Pi compared to wild-type . Images show TLC analysis of exchangeable nucleotide content of microtubules grown with GTP or GTPγS . Microtubules were spontaneously assembled using higher concentrations than for the dynamics assays: wild-type microtubules were prepared at 2 µM with GTP and 1 µM with GTPγS . β:T238A microtubules were prepared at 1 µM with either nucleotide . ( D ) Quantification of TLC data , n = 3 and errors shown represent s . d . *indicates one condition where we could not detect any Pi in one of the replicates; instead of using 0 we used the lower of the two other trials . DOI: http://dx . doi . org/10 . 7554/eLife . 10113 . 006 To support the idea that the mutation-induced changes in dynamics result from something other than a defect in GTPase activity , we assayed the nucleotide content of wild-type and β:T238A microtubules . We allowed wild-type and β:T238A αβ-tubulin to spontaneously polymerize at 1 or 2 µM concentration ( within the range of concentrations tested in Figure 2A ) in the presence of 32P-GTP . We harvested the microtubules by centrifugation , denatured them to release bound nucleotides , and analyzed the nucleotide content at the exchangeable site using thin layer chromatography ( TLC ) ( Figure 2C , D ) . We probed for a GTPase defect using α-labeled nucleotide to measure the amounts of GTP and GDP in microtubules , and for a phosphate release defect using γ-labeled nucleotide to measure the amounts of GTP and Pi in microtubules . To avoid possible complications that might arise from the very different catastrophe frequencies and shrinking rates , we initially performed these assays under ‘catastrophe free’ and slow shrinking conditions by using ( unlabeled ) GTPγS to support assembly . These experiments revealed that wild-type and mutant microtubules contain similarly low amounts of GTP ( 2–4% of total exchangeable nucleotide ) and Pi ( fewer than 3% of exchangeable sites ) ( Figure 2C , D ) . When we performed these assays using ( unlabeled ) GTP to support microtubule assembly , β:T238A microtubules contained similarly low amounts of GTP and Pi as we observed with GTPγS but wild-type microtubules appeared to contain a substantially greater amount ( ∼50% ) of GTP ( Figure 2C , D ) . Yeast microtubules have previously been reported to contain more GTP than vertebrate microtubules ( Dougherty et al . , 1998 ) . In control experiments ( not shown ) we confirmed that vertebrate microtubules contain very little GTP and that individual heterodimers were not pelleting . Because we observed very fast shrinking ( Figure 1 ) , little 32P-GTP in microtubules grown with GTPγS ( Figure 2C ) , and normal plus-end recognition by an EB1 family protein ( see below ) , it seems unlikely that wild-type yeast microtubules contain substantial amounts of GTP . The GTP containing material in our assay might instead reflect oligomers that form more readily for yeast αβ-tubulin . Whatever the mechanism , these experiments demonstrate that β:T238A αβ-tubulin remains competent to hydrolyze GTP , and that the altered nucleation and shrinking behavior cannot be ascribed to an accumulation of GTP or GDP . Pi that might result from a defect in assembly-dependent GTPase activity or phosphate release . Some other , nonenzymatic mechanism must therefore account for the observed changes in polymerization dynamics . Unoligomerized GTP-bound αβ-tubulin adopts a curved conformation that is not compatible with the straight microtubule lattice ( Nawrotek et al . , 2011; Pecqueur et al . , 2012; Ayaz et al . , 2012; 2014 ) . It seemed possible that the β:T238A mutation might stabilize microtubules by shifting the conformational preference of unpolymerized αβ-tubulin to favor a straight ( er ) conformation that is more compatible with the microtubule lattice . We used quantitative TOG binding assays ( Ayaz et al . , 2014 ) to determine if the β:T238A mutation changed the ‘curvature’ of unpolymerized αβ-tubulin . The rationale for this approach is based on the fact that TOG domains bind tightly to curved αβ-tubulin but very weakly to straight ( Figure 3A ) ( Ayaz et al . , 2012; 2014 ) . Accordingly , β:T238A αβ-tubulin should bind less tightly to a TOG domain if the β:T238A mutation appreciably changes the conformation of αβ-tubulin , for example by straightening it . TOG1 and TOG2 , two different TOG domains from the yeast microtubule regulatory factor Stu2p ( Wang and Huffaker , 1997 ) , show very similar affinity ( within a factor of two ) for unpolymerized T238A αβ-tubulin as they do for wild-type ( Figure 3B ) . In energetic terms , this modest difference in affinity is roughly equivalent to one hydrogen bond . While we cannot state that the mutant and wild-type αβ-tubulin adopt identical conformations , their very similar TOG binding properties indicate that any differences in conformation must be small . Thus , we conclude that the β:T238A mutation does not significantly change the conformation of unpolymerized αβ-tubulin . 10 . 7554/eLife . 10113 . 007Figure 3 . The buried β:T238A does not appreciably straighten unpolymerized αβ-tubulin . ( A ) Cartoon schematics illustrating that TOG domains ( blue ) bind tightly to curved αβ-tubulin ( pink and green; left ) but weakly to straight αβ-tubulin ( right ) . ( B ) Isotherms for individual Stu2 TOG domains binding to T238A αβ-tubulin . Both TOG domains bind with comparable affinity to T238A and WT tubulin . 1 Σσ confidence intervals for fitted affinities are provided in parentheses . DOI: http://dx . doi . org/10 . 7554/eLife . 10113 . 007 Recent cryo-EM studies of microtubule structure provided atomic models for distinct microtubule lattices containing GTP , GTPγS , or GDP , with GTP favoring an ‘expanded’ form and GTPγS and GDP favoring a ‘compacted’ form ( Alushin et al . , 2014; Zhang et al . , 2015 ) . Plus-end tracking proteins in the EB1 family have been shown to discriminate between these GTP-like and GDP-like lattices ( Maurer et al . , 2011; 2012; 2014; Zanic et al . , 2009 ) . We used Bim1p ( Schwartz et al . , 1997 ) , the yeast EB1 protein , to investigate if the β:T238A mutation affected αβ-tubulin conformation in the lattice . Control experiments revealed that a Bim1-GFP fusion protein tracked the growing end of yeast microtubules , and that Bim1-GFP coated the entire length of GTPγS-containing yeast microtubules with cap-like intensity ( Figure 4 ) . We therefore infer that like other EB1 proteins on vertebrate microtubules ( Zanic et al . , 2009; Maurer et al . , 2011 ) , Bim1 discriminates between GTP and GDP forms of the yeast microtubule lattice ( presumably , expanded and compacted ) . In marked contrast to its behavior on wild-type microtubules , at 50 nM concentration Bim1-GFP coated the entire length of ‘dynamic’ β:T238A microtubules with cap-like intensity . At lower concentrations ( 5 nM ) , the Bim1-GFP coat on the body of β:T238A microtubules is slightly weaker than at the cap but still substantially more intense than on wild-type ( Figure 4B ) . This stronger coating reflects tighter Bim1 binding to mutant microtubules , and it occurs in spite of the fact that the mutant microtubules contain very little GTP or GDP . Pi in the exchangeable site ( Figure 2 ) . Thus , the β:T238A mutation attenuates the conformational response to GTP hydrolysis in the microtubule lattice . The structural consequence of the mutation resembles the effect of taxol binding , which also promotes an expanded conformation of αβ-tubulin in a GDP lattice ( Alushin et al . , 2014 ) . Simply put , the mutation appears to have substantially uncoupled the conformational cycle from the nucleotide cycle in the lattice , allowing αβ-tubulin to retain GTP-lattice-like character even in a GDP lattice . 10 . 7554/eLife . 10113 . 008Figure 4 . The buried β:T238A attenuates the conformational response to GDP in the lattice . ( A ) Cartoon schematics illustrating the basis of the assay: EB1 proteins ( blue outlined ovals ) bind tightly to the GTP-lattice conformation of αβ-tubulin ( left ) but weakly to the GDP-like conformation ( right ) . ( B ) Images of the distribution of Bim1-GFP ( an EB1 family protein ) on wild-type ( left and center ) or mutant microtubules ( right ) . 4 microtubules are shown for each condition , and Bim1-GFP was present at 50 nM and 5 nM concentration for all microtubules . Plots represent average Bim1-GFP intensity on n = 9 microtubules for each condition . Error bars represent s . e . m . Cartoons illustrate the likely lattice conformation inferred from the Bim1-GFP binding . β:T238A microtubules appear GTP-like even though they contain GDP . Scale bars: 1 μm . DOI: http://dx . doi . org/10 . 7554/eLife . 10113 . 008 The Bim1 experiments described above do not report on the conformation of αβ-tubulin at the very microtubule end because EB1 proteins do not bind there ( Maurer et al . , 2012; 2014 ) . After fortuitously discovering that isolated TOG domains from the microtubule polymerase Stu2p stimulate the depolymerization of stabilized microtubules in a dose-dependent way , it seemed that TOG domains might provide an alternative way to probe the conformation of αβ-tubulin on the microtubule end . In light of the fact these TOG domains bind preferentially to the curved conformation of αβ-tubulin , we reasoned that the underlying cause of this induced depolymerization was TOG-mediated stabilization of the curved and faster dissociating conformation of αβ-tubulin on the microtubule end ( Figure 5 ) . According to this view , TOG-induced depolymerization thus provides an assay that probes the linkage ( or lack thereof ) between changes in αβ-tubulin conformation on the polymer end and microtubule shrinking rate . Using drug-stabilized microtubules as a substrate , the rate of TOG-induced depolymerization increases linearly over at least a 20-fold range of TOG concentration . This linear concentration-dependence is consistent with a collisional mechanism in which the rate-limiting step is a TOG domain arriving at the microtubule end . It is not consistent with a mechanism in which the rate-limiting step is some kind of slow αβ-tubulin conformational change . Drug-stabilized β:T238A microtubules depolymerize substantially slower than wild-type at the same concentration of TOG domain ( Figure 5 ) . Only at higher TOG concentrations do β:T238A microtubules show appreciable dose-dependent TOG-induced depolymerization ( Figure 5 ) . Similar results were obtained using MTs stabilized with GTPγS ( Figure 5 ) . 10 . 7554/eLife . 10113 . 009Figure 5 . β:T238A microtubules are resistant to TOG-induced depolymerization . ( A ) A time series of images of stabilized fluorescent microtubules show that under the same concentration of TOG1 , T238A microtubules depolymerize substantially slower than WT microtubules . Scale bars: 1 µm . ( B ) Quantification of the dose-dependence of the rate of TOG-induced depolymerization . Inset: a plot showing that under higher TOG1 concentrations , T238A microtubules also undergo induced depolymerization . We observed similar rates of TOG-induced depolymerization using DIC instead of fluorescence to monitor microtubule length ( black triangles ) , as well as using GTPγS instead of epothilone as a stabilizing reagent ( orange diamonds , purple squares ) . Additional control experiments demonstrate that the TOG-induced depolymerization is greatly reduced when a weakly-binding TOG1 mutant ( R200A ) is used ( green triangles ) . N = 30 for fluorescence measurements of wild-type TOG1-induced depolymerization of wild-type or β:T238A microtubules , N = 20 for the rest . Error bars represent s . e . m . ( C ) Cartoon model illustrating the mechanism of TOG-induced depolymerization , resulting from TOG-stabilization of tubulin subunits sampling curved conformation at the end of the microtubule . T238A microtubules depolymerize slower due to a decrease in tubulin subunits sampling the curved conformation once incorporated into the polymer . DOI: http://dx . doi . org/10 . 7554/eLife . 10113 . 009 TOG domains bind with comparable affinity to curved wild-type and mutant αβ-tubulin , so differences in TOG affinity cannot explain the markedly slower TOG-induced shrinking of β:T238A microtubules . Differences in the amount of curved αβ-tubulin at wild-type and mutant microtubule ends could explain the observed differences in TOG-induced shrinking . In principle there might be other mechanisms that strengthen association without affecting the propensity to be curved on the microtubule end . However , while such ‘curvature invariant’ mechanisms in αβ-tubulin might explain the slow post-catastrophe shrinking of the mutant , the conformation-selective nature of TOG:tubulin interactions means that this alternative view is not easily reconciled with the observed differences in TOG-induced depolymerization . Thus , a mutant-induced change in the propensity to become/remain straight on the microtubule end is the simplest way to explain both the slower post-catastrophe shrinking and weaker TOG-induced depolymerization for the mutant . That the conformation of end-bound αβ-tubulins is affected by the β:T238A mutation is also consistent with the Bim1 coating , which demonstrated that the mutation affected αβ-tubulin conformation elsewhere in the lattice . To gain insight into the local interactions responsible for the β:T238A effects on microtubule dynamics , we measured benomyl sensitivity/resistance and in vivo microtubule dynamics for serine and valine substitutions at the same position ( Figure 6 ) . Yeast expressing β:T238S αβ-tubulin are comparably benomyl resistant to β:T238A , but significantly less benomyl dependent ( Figure 6A ) . By contrast , yeast expressing β:T238V αβ-tubulin are substantially less resistant to benomyl , and show no benomyl dependence ( Figure 6A ) . Measurements of microtubule dynamics in vivo are consistent with the benomyl phenotypes: β:T238S microtubules are β:T238A-like ( static ) whereas β:T238V microtubules are more wild-type-like ( dynamic but with somewhat slower shrinking ) ( Figure 6B ) . The trend with sidechain size and the strong effect obtained from the Thr to Ser mutation at position 238 suggest that some form of ‘steric overpacking’ , not hydrogen bonding , contributes to destabilize the straight , expanded conformation of αβ-tubulin in microtubules containing GDP . 10 . 7554/eLife . 10113 . 010Figure 6 . Insights into the mechanism underlying the β:T238A effects on the conformational cycle . ( A ) Yeast with different substitutions for T238 ( for each mutant , strains with and without GFP-Tub1p are shown ) show intermediate degrees of benomyl resistance/dependence . The volume of the packing defect may be related to the magnitude of the resulting phenotype: T238S has a phenotype closer to T238A whereas T238V has a phenotype closer to wild type . ( B ) In vivo microtubule dynamics of yeast containing T238S and T238V αβ-tubulin . N = 24 , 14 for T238S and T238V microtubules respectively , with t = 120 , 43 min total time observed . ( C ) The buried β:C354A mutation immediately proximal to T238 ( left ) has also been shown to give slowly shrinking microtubules , and also shows an expanded lattice ( right ) . The view of the structure is as if from the center of the microtubule looking out , with the plus end at the top . Scale bar: 1 µm . DOI: http://dx . doi . org/10 . 7554/eLife . 10113 . 010 Reducing the size of β:C354 , a buried sidechain that packs against β:T238 ( Figure 6C ) , has also been shown to stabilize microtubules ( Gupta et al . , 2001; 2002 ) . Indeed , β:C354A or β:C354S substitutions dramatically reduced the rate of microtubule shrinking and the frequency of catastrophe ( Gupta et al . , 2002 ) . If these volume-reducing mutations also stabilize the expanded conformation of αβ-tubulin in a GDP lattice , it would provide complementary support for our overpacking-based rationale for the β:T238 ( A , S , V ) effects . We purified β:C354A αβ-tubulin and used Bim1-GFP to probe its conformation in microtubules . β:C354A microtubules showed Bim1-GFP coating similar to what we observed for β:T238A , indicating the two mutations stabilize microtubules through a common conformational mechanism ( Figure 6C ) . Positions 238 and 354 in β-tubulin show strong evolutionarily conservation ( yeast numbering; β:238 is 65% T and 34% C whereas β:354 is 99% C ) , consistent with an important functional role for these residues . Together , the β:T238A and β:C354A results suggest that changes in packing volume near helix H7 can dictate the dynamic properties of microtubules by tuning an allosteric response to GDP .
We demonstrate that relatively conservative buried mutations in β-tubulin can diminish the conformational response to GDP in the lattice , and that this altered structural response yields dramatic effects on microtubule dynamics . By discovering that mutations can uncouple the conformational and nucleotide cycles of αβ-tubulin , our findings provide new insights into ways that the αβ-tubulin conformational cycle dictates microtubule polymerization dynamics ( Figure 7 ) . 10 . 7554/eLife . 10113 . 011Figure 7 . The αβ-tubulin conformational cycle and its impact on microtubule dynamics . ( A ) Unoligomerized wild-type and β:T238A αβ-tubulin both adopt the curved conformation . That the straight conformation is strained ( flare marks ) even with the mutation-induced reduction in packing volume ( cartooned by larger and smaller red circles for wild-type and β:T238A , respectively ) suggests that extrinsic factors like interactions with the lattice control straightening . ( B ) Compared to wild-type , β:T238A αβ-tubulin is better able to populate a straight conformation on the end of GDP containing microtubules . ‘D’ indicates GDP underneath the bolded terminal subunit , compaction is represented as in Figure 4 , and the white arrow cartoons trans-acting nucleotide ( see text ) . The increased ability of the mutant to be straight could result from an altered response to GDP on the longitudinal interface , from more favorable interactions with the expanded lattice , or from a combination of both . ( C ) The schematic phase diagram illustrates that the mutation-induced changes to the αβ-tubulin conformational cycle decrease the threshold concentrations for appreciable elongation against catastrophe ( line 1 ) and for spontaneous nucleation ( line 2 ) , and also narrows the gap between them . The normal αβ-tubulin conformational cycle contributes to microtubule dynamics and makes them more amenable to regulation . DOI: http://dx . doi . org/10 . 7554/eLife . 10113 . 011 Our experiments using the EB1 family protein Bim1 showed that in the lattice a major effect of the β:T238A mutation is to substantially suppress conformational changes that normally occur as a consequence of GTPase activity ( Hyman et al . , 1995; Alushin et al . , 2014; Zhang et al . , 2015 ) . The different responses of yeast and vertebrate microtubules to GMPCPP , and the lack of high-resolution structures for wild-type or mutant yeast microtubules , make it difficult to know the specific conformations involved and how they relate to the nucleotide cycle . However , the stronger binding of Bim1 to GDP-containing β:T238A microtubules indicates that the mutant microtubule lattice retains more GTP-like character than does wild-type . This observation about αβ-tubulin in the body of the microtubule cannot by itself explain the slow shrinking , however , because the rate of shrinking is determined by the properties of incompletely surrounded terminal subunits , on which EB1 does not report . Indeed , understanding how different lattice structures relate to dynamic properties of microtubules remains a significant challenge . Our experiments with TOG domains indirectly probed the conformation of terminal subunits and showed that the β:T238A mutation reduces the propensity of terminal αβ-tubulins to be curved . Thus , the slow shrinking and lower frequency of catastrophe in the mutant most likely result from the attenuated allosteric response to GDP; this in turn results in a straighter and more strongly associated GTP-like conformation on the microtubule end , despite having GDP at the longitudinal interface ( Figure 7B ) . It is remarkable that the majority of the mutation-induced effects are confined to post-GTPase conformational changes in the lattice . Not observing significant effects on elongation is somewhat surprising , because it had been anticipated that the conformational changes required to enter the lattice would contribute to polymerization dynamics by opposing incorporation into the lattice . The lack of substantial change in the observed concentration-dependence of elongation rates could indicate that the β:T238A mutations do not affect the propensity for αβ-tubulin to ‘straighten’ on a GTP lattice . However , this is not consistent with increased spontaneous nucleation of β:T238A αβ-tubulin in the presence of GTPγS , which indicated that the mutation did affect conformational transitions that occur in/on a GTP lattice . It could be that we did not observe substantial effects on elongation because the energetics of curved to straight transitions on the microtubule end are not rate-contributing for elongation , perhaps because these transitions occur subsequent to end binding . The significant mutant-induced structural changes in the lattice were not accompanied by equivalent changes in the conformation of unoligomerized αβ-tubulin . The differential response to the mutation inside and outside of microtubules ( Figure 7A , B ) is consistent with the view that the curved conformation represents the ‘ground state’ of αβ-tubulin independent of nucleotide state ( Rice et al . , 2008; Buey et al . , 2006; Nawrotek et al . , 2011; Pecqueur et al . , 2012; Ayaz et al . , 2012 ) , that the nucleotide acts across the longitudinal interface ( Rice et al . , 2008; Nawrotek et al . , 2011 ) , and that nucleotide-dependent interactions with the microtubule lattice are what drive αβ-tubulin conformational transitions ( Rice et al . , 2008; Buey et al . , 2006; Nawrotek et al . , 2011 ) . Indeed , the changes we observed can largely be explained by an impaired allosteric response to GDP in the lattice . It will be interesting to discover in future work if other mutations in β-tubulin can straighten unpolymerized αβ-tubulin or modulate the allosteric response ( s ) to nucleotide in the lattice , and to determine if mutations in α-tubulin can yield similar effects . In summary , we showed that buried mutations of or near β:T238 alter the allosteric response to GDP in the microtubule lattice , with dramatic consequences for catastrophe frequency and shrinking rate . By describing microtubules with identical interface composition that nevertheless undergo strikingly different polymerization dynamics , our data demonstrate that allostery in the lattice dictates functionally important aspects of microtubule polymerization dynamics ( Figure 7C ) . Based on the central role of allostery in controlling the frequency of catastrophe and the rate of shrinking , we speculate that cooperative conformational linkage in the lattice amplifies the individual response to nucleotide . Together , the allosteric response to GDP and the intrinsic bias of αβ-tubulin toward the curved conformation elevate and separate the threshold concentrations for persistent elongation and for spontaneous nucleation ( Figure 7C ) , and underlie the fast shrinking rate that makes catastrophe more decisive . These allosteric contributions to microtubule dynamics make microtubules more amenable to regulation by cellular factors that enhance elongation , trigger catastrophe , and promote nucleation .
Plasmids to express TOG1 , TOG2 and wild-type yeast αβ-tubulin were previously described ( Johnson et al . , 2011; Ayaz et al . , 2012; 2014 ) . A Bim1-GFP construct , in pHAT vector containing C-terminal EGFP-tag followed by a Strep-tag II , was a gift from Dr . Gary Brouhard . A plasmid to express the T238A mutation of Tub2p ( yeast β-tubulin ) was made by QuikChange ( Stratagene ) mutagenesis , using an expression plasmid for wild-type Tub2 as template and with primers designed according to the manufacturer’s instructions . A C-terminal FlAsH ( Griffin et al . , 2000 ) sequence was added to a Tub1 construct using polymerase chain reaction with primers designed to add the sequence WDCCPGCCK ( Griffin et al . , 2000 ) . The integrity of all expression constructs was confirmed by DNA sequencing . Wild-type or mutant yeast αβ-tubulin was purified from inducibly overexpressing strains of S . cerevisiae using Ni-affinity and ion exchange chromatography ( Johnson et al . , 2011; Ayaz et al . , 2012; 2014 ) with the exception that T238A and wild-type FlAsH mutants were eluted from the Ni-affinity column with 200 mM NaCl . Prior to ion exchange chromatography , T238A Ni elution fractions were treated with Universal Nuclease ( Pierce ) at RT for 1 hr . Tubulin samples were stored in storage buffer ( 10 mM PIPES pH 6 . 9 , 1 mM MgCl2 , 1 mM EGTA ) containing 20 or 50 µM GTP depending on the application . The TOG1 ( 1-317 ) and TOG2 ( 318-560 ) were expressed in E . coli with C-terminal His6 tags and purified using Ni-affinity , gel filtration ( TOG1 ) and ion exchange chromatography ( TOG2 ) ( Ayaz et al . , 2012; 2014 ) . TOG domains were stored in RB100 ( 25 mM Tris pH 7 . 5 , 100 mM NaCl , 1 mM MgCl2 , 1 mM EGTA ) . Expression of Bim1-GFP was induced in the BL21 ( DE3 ) strain of E . coli with 0 . 5 mM IPTG for 15 hrs at 16C . Single pellets were resuspended in 50 mM Na2HPO4 , 300 mM NaCl , 40 mM imidazole and sonicated for 30 min in the presence of PMSF . Lysates were clarified by centrifugation . Cleared lysate was loaded onto a His60 Superflow Column ( Clontech ) and the final sample was eluted in 200 mM imidazole . Elution fractions were loaded onto a 3 mL Strep-Tactin Superflow column ( IBA , Germany ) and eluted in RB100 with 5 mM desthiobiotin . For expression: JEL1 MATα leu2 trp1 ura3−52 prb1−1122 pep4−3 Δhis3::PGAL10-GAL4 For analysis of growth phenotypes: CUY 2172 MATa TUB2::URA3 ade2-101 his3-∆200 leu2-1 lys2-801 ura3-52 CUY 2179 MATa tub2-T238A::URA3 ade2-101 his3-∆200 leu2-1 lys2-801 ura3-52 CUY 2198 MATa tub2-T238S::URA3 ade2-101 his3-∆200 leu2-1 lys2-801 ura3-52 CUY 2200 MATa tub2-T238V::URA3 ade2-101 his3-∆200 leu2-1 lys2-801 ura3-52 For microtubule imaging: CUY 2265 MATa TUB2::URA3 GFP-TUB1::LEU2::TUB1 ade2-101 his3-∆200 leu2-1 lys2-801 ura3-52 CUY 2238 MATa tub2-T238A::URA3 GFP-TUB1::LEU2::TUB1 ade2-101 his3-∆200 leu2-1 lys2-801 ura3-52 CUY 2208 MATa tub2-T238S::URA3 GFP-TUB1::LEU2::TUB1 ade2-101 his3-∆200 leu2-1 lys2-801 ura3-52 CUY 2209 MATa tub2-T238V::URA3 GFP-TUB1::LEU2::TUB1 ade2-101 his3-∆200 leu2-1 lys2-801 ura3-52 tub2 alleles were integrated into yeast as described previously ( Reijo et al . , 1994 ) in vivo imaging of microtubule dynamics was performed as described previously ( Huang and Huffaker , 2006 ) . Note that introduction of GFP-TUB1 into strains containing the tub2-T238A mutation lessens their dependence on benomyl for growth ( Figure 6 ) and allowed us to image these cells in the absence of benomyl . Flow chambers were prepared as described previously ( Gell et al . , 2010 ) , with the exception that sea urchin axonemes ( Waterman-Storer , 2001 ) were used to template yeast MT growth . Chambers were rinsed with BRB80 ( 80 mM PIPES pH 6 . 9 , 1 mM MgCl2 , 1 mM EGTA ) , followed by 10 min incubation with sea urchin axonemes . Chambers were then blocked with 1% F-127 Pluronic in BRB80 for 5 min , and washed with 1X PEM ( 100 mM PIPES pH 6 . 9 , 1 mM EGTA , 1 mM MgSO4 ) containing 1 mM GTP . Reaction chambers were sealed with VALAP after addition of αβ-tubulin . Wild type or mutant yeast αβ-tubulin ( in storage buffer containing 50 µM GTP ) was taken from -80˚C , rapidly thawed , and passed through a 0 . 1 µm centrifugal filter at 4˚C to remove aggregates . The concentration of αβ-tubulin was measured by UV absorbance using an extinction coefficient of 115000 M-1cm-1 . Protein was kept on wet ice for no more than 30 min before measuring MT dynamics . MT dynamics were imaged by differential interference contrast microscopy ( DIC ) using an Olympus IX81 microscope with a Plan Apo N 60x/1 . 42 NA objective lens and DIC prisms . Illumination at 550 nm was obtained by inserting a bandpass filter of 550/100 nm ( Olympus ) in the light path . Temperature was maintained at 30˚C using a WeatherStation temperature controller with enclosure fit to the microscope’s body . Micro-Manager 1 . 4 . 16 ( Edelstein et al . , 2010 ) was used to control the microscope and a Hamamatsu ORCA-Flash2 . 8 CMOS camera used to record the reactions . MT dynamics were recorded by taking an image every 500 ms for 1 to 2 hrs . At the end of each movie , a set of 100 out-of-focus background images was taken for background subtraction ( see below ) . To improve signal to noise , batches of 10 raw images were averaged using ImageJ ( Schneider et al . , 2012 ) and intensity normalized before background subtraction . MT length was measured manually using a PointPicker plugin for ImageJ . Rates of MT elongation and catastrophe frequencies were determined as described previously ( Walker et al . , 1988 ) . To prepare microtubules for electron microscopy , samples of wild-type ( 3-–4 µM ) or β:T238A αβ-tubulin ( 2 µM ) were prepared in 100 mM PIPES pH 6 . 9 , 10% glycerol , 2 mM MgSO4 , 0 . 5 mM EGTA , 1 mM GTP and incubated at 30 ˚C for 1 hr . 5 µl of the assembly reactions were spotted onto freshly glow-discharged 400 mesh grids with a carbon coated formvar support ( Ted Pella ) , incubated for 30 s , rinsed with water , and stained with 2% aqueous uranyl acetate . Negatively stained grids were imaged at 23 , 000 x magnification using a Tecnai G2 Spirit electron microscope equipped with a 2Kx2K CCD camera ( Gatan ) . Microtubule sedimentation assays were performed using a range of wild-type or mutant yeast αβ-tubulin concentrations ( Figure 2B , 0 . 25 – 4 μM ) in assembly buffer ( 100 mM PIPES pH 6 . 9 , 10% glycerol , 2 mM MgSO4 , 0 . 5 mM EGTA ) containing the indicated nucleotide . Samples were polymerized for 90 min at 30˚C . For SDS-PAGE analysis , microtubules were pelleted by centrifugation at 60 , 000 rpm ( ∼150000 x g ) at 30˚°C for 30 min in a pre-warmed TLA-100 rotor ( Beckman-Coulter ) , supernatant was carefully removed , and the pellet was re-suspended in an equal volume of assembly buffer such that the pellet and supernatant fractions were of equal volume . To image the products of the nucleation reactions , the reactions were cross-linked by diluting 10-fold into assembly buffer containing 1% glutaraldehyde . Cross-linking was quenched after 3 min by 5-fold dilution into assembly buffer containing 20 mM Tris pH 6 . 8 . 150 μL of the quenched , cross-linked reactions were applied to the top of a glycerol cushion ( 20% glycerol in BRB80 ) and spun through the cushion onto poly-L-lysine coated coverslips . Coverslips were washed with BRB80 , fixed with methanol and stained using FITC-DM1α ( Sigma-Aldrich ) for imaging by epifluorescence , as described previously ( Ayaz et al . , 2012 ) . Wild type ( 2 or 1 µM respectively for GTP or GTPγS ) or β:T238A tubulin ( 1 µM for either nucleotide ) was mixed with 100 µM GTP or GTPγS ( containing 33 nM [α-32P or γ-32P]-GTP ) in assembly buffer ( see above ) on ice . Microtubules were assembled and harvested by centrifugation at 80 , 000 rpm ( ∼268000 x g ) at 30˚C for 10 min in a pre-warmed TLA-120 rotor ( Beckman-Coulter ) . Supernatant was carefully removed , and the pellets were gently washed 4 times with pre-warmed assembly buffer , and then resuspended with 6 M guanidine to denature the protein and release bound nucleotide . After 10-fold dilution into water , samples were loaded onto a Cellulose PEI TLC plate ( Selecto Scientific ) and TLC was performed , first with water followed by buffer containing 0 . 75 M Tris , 0 . 4 M LiCl , and 0 . 45 M HCl . The TLC plate was exposed to X-ray film after air drying . Radiolabelled mixtures of GTP/GDP and GTP/Pi were used as markers . Samples for analytical ultracentrifugation ( TOG1 , TOG2 , yeast αβ-tubulin , and mutant T238A polymerization-competent yeast αβ-tubulin ) were dialyzed into final buffer conditions of RB100 ( 25 mM Tris pH 7 . 5 , 1 mM MgCl2 , 1 mM EGTA , 100 mM NaCl ) containing 20 µM GTP . The samples shown in Figure 3 contain 0 . 15 µM T238A yeast αβ-tubulin and 0 . 015 µM , 0 . 03 µM , 0 . 15 µM , 0 . 3 µM , 0 . 6 µM , 1 . 2 µM , 5 µM TOG1 or 0 . 015 µM , 0 . 025 µM , 0 . 05 µM , 0 . 1 µM , 0 . 2 µM , 0 . 4 µM , 0 . 8 µM , 1 . 6 µM , and 5 µM TOG2 . Samples were mixed and incubated at 4˚C for at least one hr prior to the experiment . All analytical ultracentrifugation experiments were carried out in an Optima XL-I centrifuge using an An50-Ti rotor ( Beckman-Coulter ) . Approximately 390 µL of each sample were placed in charcoal-filled , dual-sector Epon centerpieces . Sedimentation ( rotor speed: 50 , 000 rpm ) was monitored using absorbance at 229 nm and centrifugation was conducted at 20˚°C after the centrifugation rotor and cells had equilibrated at that temperature for at least 2 . 5 hrs . Protein partial-specific volumes , buffer viscosities , and buffer densities were calculated using SEDNTERP ( Laue et al . , 1992 ) . Data were analyzed using SEDFIT and SEDPHAT ( available at http://www . analyticalultracentrifugation . com ) ( Schuck , 2000; Schuck et al . , 2002 ) . Wild-type and mutant yeast αβ-tubulin were labeled with 6-–10 µM ReAsH in DMSO ( Life Technologies ) and 1 mM TCEP , in tubulin storage buffer ( see above ) for 90 min at RT . To remove excess unbound dye , samples were exchanged into assembly buffer ( 100 mM PIPES pH 6 . 9 , 10% glycerol , 2 mM MgSO4 , 0 . 5 mM EGTA ) with 2 mL , 7K MWCO Zeba spin desalting columns ( Thermo Scientific ) . Labeled wild-type and mutant yeast αβ-tubulin were polymerized in the presence of 3–5 µM epothilone-B in assembly buffer ( see above ) with 1 mM GTP or in the presence of 500 µM GTPγS in assembly buffer ( see above ) . The mixture was incubated for 30 min at 30 ˚°C . Flow chambers were prepared as described above . His-Tag Antibody ( 1:200 , Gentech ) was incubated in the chamber for 10 min , followed by incubation with 1% Pluronic F-127 in BRB80 for 5 min , followed by a wash with BRB80 . Pre-formed , epothilone- or GTPγS-stabilized wild-type or T238A yeast MTs were then introduced into the chamber and allowed to incubate for 10 min , followed by a wash with BRB80 to remove unbound MTs . Solutions containing a range of TOG1 ( 0 . 1-–10 µM ) or TOG1 ( R200A ) ( 250 , 750 nM ) concentrations ( Figure 5B ) in imaging buffer ( BRB80 + 200 nM epothilone + 0 . 1 mg/mL BSA + antifade reagents ( glucose , glucose oxidase , catalase ) , without the addition of β-mercaptoethanol [Gell et al . , 2010] ) were introduced into the chamber immediately prior to data collection . To ensure that the TOG-induced depolymerization was not an artifact of fluorescence imaging , unlabeled wild-type microtubule were assembled as described above but without ReAsH labeling and then imaged with two concentrations of TOG1 using DIC microscopy ( black points in Figure 5B ) . MT depolymerization reactions with fluorescent microtubules were imaged by epifluoresence microscopy using an Olympus IX81 microscope with a Plan Apo N 60x/1 . 42 NA objective lens and Hamamatsu ORCA-Flash2 . 8 CMOS camera , a mercury short arc lamp , and a Texas Red filter cube ( Olympus ) . Reactions were temperature controlled and the microscope was controlled as described above . Images of MT shrinking were recorded every 60 s for about 10 min . MT depolymerization reactions with non-fluorescent wild-type microtubules were imaged using differential interference microscopy as described above . MT length was measured manually using ImageJ ( Schneider et al . , 2012 ) . Average lengths of MT’s were taken over the time course of the movie and were used to determine rate of depolymerization over an hour time span . Preparation of flow chambers using sea urchin axonemes was followed as described above . Samples of wild-type , β:T238A , or C354A αβ-tubulin with 1 mM GTP or GTPγS were prepared and incubated in flow-chambers for 30–180 min and observed under DIC to evaluate presence of microtubule growth . After a given time , solutions containing desired tubulin sample and nucleotide of interest , along with 5 or 50 nM Bim1-GFP with imaging buffer described above , were flowed into the chamber . Interactions of Bim1-GFP with MTs were imaged by total internal reflection fluorescence microscopy using an Olympus IX81 microscope with a TIRF ApoN 60x/1 . 49 objective lens , a 491 nm 50 mW solid-state laser and Hamamatsu ORCA-Flash2 . 8 CMOS camera ( Olympus ) . Reactions were temperature controlled at 30°C and the microscope was controlled as described above . Axonemes were tracked under DIC and TIRF conditions . Images of MTs were taken over several frames from 15–30 min . Bim1-GFP fluorescence intensity along microtubules and extending beyond their growing ends was obtained using the PlotProfile function in ImageJ ( Schneider et al . , 2012 ) . These linescans were manually aligned to superimpose the sharp change in intensity at the very microtubule end , and aligned intensity values were averaged . To collect a large number of α- and β-tubulin sequences , we performed psiblast ( Altschul et al . , 1997 ) ( non-redundant ( nr ) database , 3 iterations ) on the amino acid sequences of Tub1p and Tub2p from S . cerevisiae . The top 1000 sequences from both searches were aligned using Clustal Omega ( Sievers et al . , 2011 ) and manually pruned in Jalview ( Waterhouse et al . , 2009 ) to eliminate sequences with large insertions or deletions .
Microtubule dynamic instability depends on the GTPase activity of the polymerizing αβ-tubulin subunits , which cycle through at least three distinct conformations as they move into and out of microtubules . How this conformational cycle contributes to microtubule growing , shrinking , and switching remains unknown . Here , we report that a buried mutation in αβ-tubulin yields microtubules with dramatically reduced shrinking rate and catastrophe frequency . The mutation causes these effects by suppressing a conformational change that normally occurs in response to GTP hydrolysis in the lattice , without detectably changing the conformation of unpolymerized αβ-tubulin . Thus , the mutation weakens the coupling between the conformational and GTPase cycles of αβ-tubulin . By showing that the mutation predominantly affects post-GTPase conformational and dynamic properties of microtubules , our data reveal that the strength of the allosteric response to GDP in the lattice dictates the frequency of catastrophe and the severity of rapid shrinking .
Protein filaments called microtubules help move cargo around inside cells . Chromosomes , which contain the cell’s genetic blueprints , are the microtubule’s most precious cargo . Before a cell divides , microtubules grow from the ends of the dividing cell towards the middle , where they attach to the chromosomes that are lined up along the centerline . Then the microtubules shrink and drag the chromosomes back to the opposite ends of the cell . This allows each of the new cells to get one copy of each chromosome . When the microtubules are growing , a molecule called guanosine triphosphate ( or GTP ) is attached to the proteins at the end of the filament . This acts like a cap and protects the microtubule from shrinking . Later a chemical reaction converts GTP into GDP ( short for guanosine diphosphate ) . Without the protective GTP cap , the microtubule quickly shrinks . At the same time , the proteins that make up the microtubule also change shape . In the microtubule , the proteins adopt a straight shape when GTP is attached . The proteins favor a different shape in the microtubule when GDP is attached . However , it is unclear if or how these shape changes contribute to how a microtubule grows or shrinks . Geyer et al . now show how this shape shifting can influence microtubule shrinking , by first identifying a mutation in yeast microtubule proteins that cause the proteins to remain straight even when GDP is attached . Next , powerful microscopes were used to make time-lapse videos of the mutated microtubules . This allowed Geyer et al . to observe how the mutated microtubules behaved and compare this to the behavior of normal microtubules . The experiments revealed that the mutated microtubules were less likely to begin shrinking than typical microtubules . The mutated microtubules also shrunk more slowly . These findings indicate that the shape changes control the speed of shrinking and frequency of entering the shrinking phase . These new details about the control of microtubule growth and shrinkage may help scientists studying how cell division happens in both healthy and cancerous cells .