extfieldsofstudy: Chemistry Medicine

s2fieldsofstudy: Biology Psychology

text:
IH activity is increased in populations of slow versus fast motor axons of the rat

Much is known about the electrophysiological variation in motoneuron somata across different motor units. However, comparatively less is known about electrophysiological variation in motor axons and how this could impact function or electrodiagnosis in healthy or diseased states. We performed nerve excitability testing on two groups of motor axons in Sprague–Dawley rats that are known to differ significantly in their chronic daily activity patterns and in the relative proportion of motor unit types: one group innervating the soleus (“slow motor axons”) and the other group innervating the tibialis anterior (“fast motor axons”) muscles. We found that slow motor axons have significantly larger accommodation compared to fast motor axons upon application of a 100 ms hyperpolarizing conditioning stimulus that is 40% of axon threshold (Z = 3.24, p = 0.001) or 20% of axon threshold (Z = 2.67, p = 0.008). Slow motor axons had larger accommodation to hyperpolarizing currents in the current-threshold measurement (-80% Z = 3.07, p = 0.002; -90% Z = 2.98, p = 0.003). In addition, we found that slow motor axons have a significantly smaller rheobase than fast motor axons (Z = -1.99, p = 0.047) accompanied by a lower threshold in stimulus-response curves. The results provide evidence that slow motor axons have greater activity of the hyperpolarization-activated inwardly rectifying cation conductance (IH) than fast motor axons. It is possible that this difference between fast and slow axons is caused by an adaptation to their chronic differences in daily activity patterns, and that this adaptation might have a functional effect on the motor unit. Moreover, these findings indicate that slow and fast motor axons may react differently to pathological conditions.

INTRODUCTION
Many studies have demonstrated that changes in chronic daily activity patterns alter the electrophysiological properties of cat and rodent motor neurons measured from the soma: after exercise Gardiner, 2002, 2003;Gardiner et al., 2005), chronic stimulation , and hindlimb unweighting or spinal transections (Hochman and McCrea, 1994;Cormery et al., 2005). Muscle contractile properties also change in response to different activity levels like chronic stimulation and exercise (Kernell et al., 1987;Westgaard and Lomo, 1988;Gordon et al., 1997) or decreased activity levels through spinal transection (Cope et al., 1986;Munson et al., 1986). However, there is comparatively little known about the association between the electrophysiological properties of motor axons and motor unit type, or whether the axons change in response to altered chronic activity levels that induce plasticity of motor unit phenotype. The primary exception is conduction velocity (CV), which is altered in response to chronic changes in daily activity patterns of the motor units (Carp and Wolpaw, 1994;Munson et al., 1997;Beaumont and Gardiner, 2002). However, it has been acknowledgd for some time that conventional nerve CV provides limited physiological insight and additional measures from nerve excitability studies are warranted (Krarup and Moldovan, 2009;Kiernan and Lin, 2012).
Nerve excitability studies in humans have demonstrated that electrophysiology is different between axons innervating different muscles (Kuwabara et al., 2000(Kuwabara et al., , 2001Krishnan et al., 2004;Bae et al., 2009;Jankelowitz and Burke, 2009) and between motor axons innervating the same muscle but activated at different percutaneous stimulation thresholds (Trevillion et al., 2010). Similar studies in rodents suggested that motor axons activated at different stimulation thresholds vary moderately in measures of nerve excitability (Mori et al., 2010;Nodera and Rutkove, 2012). These studies suggest that motor axon conductances such as hyperpolarization-activated inwardly rectifying cation conductance (I H ) and slow potassium conductance (I Ks ) may vary across muscles and axons of different threshold. However, none of these studies have examined populations of axons from muscles with distinct distributions of motor unit types. The previous research suggests that the major differences in motor axons are a result of differences in conductances rather than simply a consequence of different axon diameters.
The purpose of this study is to compare the electrophysiology of axons innervating the soleus muscle (SOL) to those innervating the tibialis anterior (TA) muscle in the rat. SOL axons were considered to represent a population of slow-and TA axons fast-motor axons. About 94% of rat TA motor units are classified as a fast (fast fatigable or fast fatigue-resistant) motor unit type while 6% are slow, based on a modified version of Burke's criteria originally used to distinguish motor units in the cat hindlimb (Totosy de Zepetnek et al., 1992). Note the motor unit distribution in the human TA has a much larger contribution from slow motor units (Johnson et al., 1973). On the other hand, about 80% of the motor units in rat SOL are classified as slow motor unit type while 20% are the fast motor unit type (Gillespie et al., 1987). Rat TA and SOL motor units have dramatically different activity patterns in locomotion (Gorassini et al., 2000) and presumably much different overall daily activity patterns (e.g., Hennig and Lomo, 1985). Thus, by using TA and SOL axons in this study, we analyzed axons of different motor unit types as well as axons with different chronic daily firing patterns.

SURGERY AND RECORDINGS
A total of 14 female Sprague-Dawley rats, weighing 280 ± 50 g (mean ± SD), were used in this study. These weights in female Sprague-Dawley rats correspond to an age of approximately 90 days, which represents young but sexually mature females (Yang et al., 2000;George and Bostock, 2007). Rats were housed in pairs in a 12:12 h light-dark cycle, environmentally controlled (22-24 • C, 40-70% humidity) room. Water and rat chow (Lab Diet 5001, PMI Nutrition, Brentwood, MO, USA) were provided ad libitum. All animal studies were conducted in accordance with the Canadian Council on Animal Care Guidelines and Policies with approval from the Animal Care and Use Committee: Health Sciences for the University of Alberta.
Anesthesia was induced by intraperitoneal injection of a mixed dose of 60 mg/kg ketamine and 7.2 mg/kg xylazine (KX) with additional KX dosages to maintain a surgical plane of anesthesia for the duration of the experiment. A rectal thermometer monitored internal body temperature, which was maintained between 34.5 and 38.5 • C by a heating lamp. Upon completion of all experiments, animals were euthanized by an overdose of KX followed by cervical dislocation.
An electrical clipper was used to initially remove hair over the lower back and legs followed by a depilatory cream to remove the remaining hair on the lower back and hip areas. Percutaneous electrical stimulation was delivered via Ag/AgCl 3M Red Dot electrodes (10 mm diameter), with the active electrode placed over the sciatic notch and the return electrode placed over the lumbar vertebrae at the midline of the back ( Figure 1A). Electromyographic (EMG) recordings were made using Teflon-coated stainless steel monofilament wire (Cooner Wire 765-40; 40G) with approximately 3 mm of Teflon removed from the end of the wire. The wire was threaded through a hypodermic needle (TA 26G, SOL 27G) with ∼2 mm extending from the tip and twisted to form a hook. To record a bipolar intramuscular compound muscle action potential (CMAP) signal in TA the first needle was inserted near the motor endplate on the proximal 1/3 of the muscle, and a second needle was inserted about 2.5 mm distally. Insertion of the hypodermic needles in SOL was similar except they were placed about 5 mm apart because of the orientation of the motor endplate and muscle fibers in SOL. After the stainless steel recording electrodes were securely placed in each muscle, the needles were removed. To minimize EMG from other muscles and contamination of the target CMAP during the threshold tracking procedure, extensor digitorum longus, flexor digitorum longus, medial gastrocnemius, and plantaris were denervated. The distal 3/5 of lateral gastrocnemius (LG) was excised, rather than denervated, because SOL axons are intertwined with LG axons in a shared nerve branch that is not easily separated in the rat ( Figure 1B).
Compound muscle action potentials were amplified (to cover one-half to two-thirds of the A/D range), filtered (10-1000 Hz) and digitized by a 12-bit A/D board (National Instruments DAQ-6062e, Austin, TX, USA) at a rate of 10 kHz. Noise introduced from nearby power sources was removed in real time using a Hum Bug 50/60 Hz Noise Eliminator (Quest Scientific Instruments, North Vancouver, BC, Canada). In cases of excessive levels of noise which occurred in six rats, a 60 Hz notch filter was also implemented. The time required from anesthetic induction to complete all measures in an animal was typically 1 h. In five animals the electrodes were repositioned and the tests repeated as a result of signal contamination. The time required in these cases was approximately 1.5 h. Arterial blood gases and acid base status were not monitored but are well maintained at rest in this preparation over this time period (Jendzjowsky and DeLorey, 2013).

NERVE EXCITABILITY TESTING
Multiple excitability measures of TA and SOL axons were made from the right leg. Stimuli were delivered using QTRAC software (Institute of Neurology, Queen Square, London, UK) and an isolated bipolar constant current stimulator (Digitimer DS5, Digitimer Ltd., Welwyn Garden City, Hertfordshire, UK). The QTRAC protocol "TRONDCMW" consists of five nerve excitability indices: threshold electrotonus (TE), current-threshold (I/V), recovery cycle (RC), rheobase, and strength-duration time constant (SDTC). The primary outcome measure for each of these indices is the change in the amplitude of the test stimulus required to produce a target CMAP: in this case 40% of the maximum with a tolerance of ±7.5%. The duration of the test stimulus was 1.0 ms except during rheobase and SDTC in which the duration varies between 0.2 and 1.0 ms. Krishnan et al. (2009) has a more detailed description of this validated methodology.
For TE, a 100 ms sub-threshold conditioning stimulus with an amplitude expressed as a percentage (±40 and ±20%) of the test stimulus is applied and the change in test stimulus amplitude is measured at multiple delays relative to the start of the conditioning stimulus. The I/V is a similar test with a 200 ms sub-threshold conditioning stimulus and the change in test stimulus amplitude is measured at the end of the conditioning stimulus. The amplitude of the conditioning stimulus for I/V measurements varies from +50 to −100% of the unconditioned test stimulus. In RC, a supramaximal conditioning stimulus is applied to the nerve, and the test stimulus is applied at delays ranging from 2 to 200 ms. Rheobase and SDTC are determined via linear regression of the charge-duration data obtained through five test stimuli that range in duration from 0.2 to 1.0 ms.

ANALYSIS
All statistical analysis was done in SPSS version 21. Normality was assessed by the Shapiro-Wilk test and it was found that most data groups (groups within each of the five nerve excitability indices) violated the normality assumption at an alpha level of 0.05. Specifically, the hypothesis of a normal distribution was rejected for all data groups except TE +20% and depolarizing I/V measures. Therefore the non-parametric Wilcoxon signed-rank test was used for paired comparison of TA and SOL axons within each individual rat. For comparisons involving TE, paired comparison of TA and SOL axons were made at three delays for each of the four conditioning stimulus levels, i.e., ±40 and ±20%. For comparisons involving I/V, there were five data groups analyzed from I/V +50 to +10% and 10 groups analyzed from I/V 10 to −100%, all spaced by increments of 10%. In RC, all 18 delays ranging between 2.0 and 200 ms delay were analyzed. Finally, rheobase and SDTC were also used to compare TA and SOL axons. In order to control for type I statistical errors, we divided the alpha level (0.05) by the number of data groups within each respective nerve excitability measure. Therefore, the adjusted alpha level (α a ) for determining statistical significance was 0.017 in depolarizing and hyperpolarizing TE, 0.01 in depolarizing I/V, 0.005 in hyperpolarizing I/V, 0.006 in hyperpolarizing I/V Slope, 0.003 in RC, and 0.05 in rheobase and SDTC. The whiskers in each box-and-whisker plot represent 5 and 95% confidence intervals, while the box represents the interquartile range and the line the median. Line plots include the median value and error bars are the 95% confidence interval.

THRESHOLD ELECTROTONUS
Previous studies using nerve excitability testing have indicated that particular testing delays within the TE measurement are of greater interest since these are where properties diverge (Schwarz et al., 2006;George and Bostock, 2007;Sittl et al., 2011). These delays are 20-30, 100-109, and 120-150 ms and constitute the planned contrasts for statistical analysis (Figure 2A, gray areas). We averaged all the data within each of these delay ranges and used these values for comparisons between TA and SOL axons. Figure 2A displays the averaged data from TA and SOL axons for depolarizing TE +40% and hyperpolarizing TE −40%.
The initial fast phase, which is proportional to the applied current, was the same for TA and SOL axons for all depolarizing and hyperpolarizing conditioning currents. During depolarizing TE +40% there was no statistical difference between TA and SOL axons. In hyperpolarizing TE −40%, TA axons had a significantly greater threshold increase than SOL axons at 20-30 ms (Z = 3.24, p = 0.001) and 100-109 ms (Z = 3.24, p = 0.001), and a significantly smaller threshold decrease at 120-150 ms (Z = −2.61, p = 0.009; Figure 2B). For the TE −20% condition at 100-109 ms, TA axons had a significantly greater threshold increase than SOL axons (Z = 2.67, p = 0.008; Figure 2C). The data for these measures are given in Table 1.

CURRENT-THRESHOLD
There was no significant difference between TA and SOL axons at any of the five depolarizing conditioning strengths. At hyperpolarizing conditioning strengths, however, we found that TA axons had significantly greater threshold increases at I/V −80% (Z = 3.07, p = 0.002) and −90% (Z = 2.98, p = 0.003; Figure 3).
The slopes of the TA and SOL I/V curves were calculated at hyperpolarizing conditioning strengths, and the slope of the TA axon line was found to be significantly smaller than the SOL axon line at I/V −40% (Z = 2.79, p = 0.045) and −50% (Z = 3.10, p = 0.018). The values for these I/V measurements are given in Table 1.

RECOVERY CYCLE
There were no significant differences between TA and SOL axons in RC (Figure 4; Table 1) using the adjusted alpha value of 0.003 for multiple comparisons. Comparison at delays of 2-5 ms showed that TA axons had smaller increases in threshold compared to SOL axons (p-values less than 0.05), but these differences did not meet criteria for rejection of the null-hypothesis given the multiple comparisons.

Frontiers in Human Neuroscience
www.frontiersin.org There is a statistically significant increase in median threshold change for SOL compared to TA axons, only at the end of the hyperpolarizing condition pulse, i.e., 100-109 ms. * p < 0.017.

STIMULUS-RESPONSE CURVES, RHEOBASE, AND STRENGTH-DURATION TIME CONSTANT
The stimulus-response curves showed a rightward shift for the TA axons indicating a higher threshold ( Figure 5A). The normalized stimulus-response curve for TA was less steep compared to that for the SOL axons ( Figure 5B). Paired comparisons found that TA axon rheobase was modestly but significantly larger than SOL axon rheobase (Z = 1.99, p = 0.047; Figure 5C). The higher rheobase for TA axons is consistent with the rightward shift of the stimulusresponse curve. The SDTC was not different between the two axon groups (Z = 1.36, p = 0.17; Figure 5D).

DISCUSSION
The present findings show that many measures of nerve excitability are indistinguishable for slow soleus and fast TA motor axons in the rat. The exception to this general finding is for measures of accommodation to prolonged hyperpolarization during the TE and I/V tests. The most parsimonious explanation for the differences is that voltage-gated inwardly rectifying current I H is greater in slow compared to fast axons. However, some of the differences can not be explained by differences in I H : stimulus-response curves and rheobase, as well as differences in −40% TE at the early delay of 20-30 ms. The most direct evidence implicating the channel responsible for I H , hyperpolarization-activated cyclic nucleotide-gated (HCN) channel, in TE and I/V nerve excitability measures comes from direct in vitro voltage recording of rat spinal root myelinated axons (Baker et al., 1987). HCN channels are blocked by cesium (Biel et al., 2009) andBaker et al. (1987) showed that the late accommodation in measures similar to TE and I/V was antagonized by cesium. Yang et al. (2000) used cesium chloride to decrease accommodation to hyperpolarizing TE measures using an in vivo rat, tail motor axon preparation similar to the current study. For TE −20 and −40% conditioning pulses the threshold reduction at delays of 100-109 ms was greater after cesium administration. Thus reducing I H in mixed tail motor axons generates responses that are similar to the data from fast TA motor axons reported here FIGURE 4 | The recovery cycle in the rat TA and SOL motor axons did not show a period of superexcitability followed by subexcitability, just a slowly decaying relative refractory period. A Wilcoxon signed-rank test did not find any significant differences between TA and SOL axons at any of the 18 delays. It should be noted that the large number of delays (18) that were compared created a very small adjusted alpha level, 0.003. Therefore there is an increased risk of type 2 error. (Figure 2). This supports our inference that fast motor axons have weaker I H activity compared to slow motor axons. This is not the first study to suggest that I H may vary across a population of motor axons, but it is the first to test two clearly distinct populations. A previous study investigated motor axons in Frontiers in Human Neuroscience www.frontiersin.org human median nerve to conclude that the lowest threshold axons had different electrophysiological properties compared to higher threshold axons (Trevillion et al., 2010). The best explanation for the unique properties of low-threshold motor axons was that they had a greater level of I H activity, and this inference was based on computer modeling. The authors used F-wave latency to characterize ten single motor units (45% of their total sample) as having a faster CV and therefore larger diameter compared to the average motor axon in the population responses. Their conclusion was that axons of a larger diameter and faster CV had greater I H activity. This is the opposite of the findings in the present study that found greater I H activity in the slower soleus motor axons compared to fast TA motor axons. It is not clear at this time why there is a clear discrepancy in the two findings other than the obvious species difference and use of anesthetic.
There are four isoforms of the HCN channel, HCN1-HCN4, all of which allow for passage of a mixed inward cationic current of both K + and Na + ions. HCN channels are found in all three major neuron compartments: the soma, dendritic tree, and the axon proper (Baker et al., 1987;Takigawa et al., 1998;Robinson and Siegelbaum, 2003;Biel et al., 2009). In addition to being voltage dependent, HCN channels have been found to be influenced by acidic lipids (i.e., PIP2), protons, extracellular K + concentration, and cytosolic proteins, and usually most significantly by cAMP (Biel et al., 2009). HCN1 has the fastest time constant of activation which can range anywhere from 30 to 200 ms at −140 to −95 mV. HCN2 is the next fastest isoform with the time constant of activation ranging from 150 ms to 1 s (Biel et al., 2009). Two recent nerve excitability studies comparing nerve excitability properties of motor and sensory axons have discussed the hypothesis that differences in I H between these two axon groups are due to sensory axons having either greater expression of faster HCN isoforms and/or a different availability of HCN ligands that renders greater I H in sensory compared to motor axons (Howells et al., 2012;Nodera and Rutkove, 2012). A higher density of channels on slow axons, the expression of different isoforms, or cAMP dependent phosphorylation status generating a difference in half-activation Frontiers in Human Neuroscience www.frontiersin.org voltages could explain the increased I H in slow motor axons in the present study. There was no a priori hypothesis that I H current would dominate the difference between slow and fast axons. Future studies should use a modified nerve excitability testing protocol with additional stronger and more prolonged hyperpolarizing conditioning stimuli to further characterize the difference in I H between fast and slow axons (Tomlinson et al., 2010).
Thus, it appears I H is a plastic ionic conductance that varies according to the type of motor axon as well as between motor and sensory axons. As the type of activity patterns are different between most motor and sensory axons, as well as for lower and higher threshold axons, this points to the possibility that I H expression depends on impulse activity patterns. This is the interpretation given for the finding of reduced I H in human motor axons on the affected side for individuals experiencing hemiparesis subsequent to stroke (Jankelowitz et al., 2007). In these individuals the motor axons on the affected side are expected to have a chronic reduction in impulse activity, if spasticity is not excessive. The reduced chronic activity leads to an adaptive change in I H that is consistent with our finding of less accommodation to hyperpolarization in the rat TA motor axons. Axons innervating the rat TA are mostly from fast motor units that fire less often but at higher frequencies than the axons of SOL, which are mostly from slow motor units (Gillespie et al., 1987;Totosy de Zepetnek et al., 1992;Gorassini et al., 2000). Activity-dependent hyperpolarization of axons is known to correlate positively with firing frequency as well as the number of impulses in a train (Erlanger and Gasser, 1973;Raymond, 1979;Morita et al., 1993;Kiernan et al., 2004), although the magnitude of hyperpolarization saturates at frequencies around 20-50 Hz in lizards (Morita et al., 1993) and possibly at 20-30 Hz in humans . Although the relative importance of firing frequency and impulse load in creating activity-dependent hyperpolarization in rat TA and SOL axons is unclear, (Raymond, 1979) has shown that in the frog, average firing frequencies as low as 1.25 Hz can cause axon threshold depression and therefore probably activity dependent hyperpolarization (see Figure 9 in Raymond, 1979). Bursts in rat SOL motor units fire at an average frequency of 20 Hz for roughly 30% of the day (Hennig and Lomo, 1985), and so it seems likely that SOL axons would generally experience a greater tendency toward activity-dependent hyperpolarization than TA axons due to greater activation by the Na + -K + pump. It is possible that greater I H expression in SOL axons offsets a tendency for their hyperpolarization caused by higher levels of activity.
While many of the differences in nerve excitability indices are consistent with the interpretation that I H is greater in SOL axons, this mechanism may not explain differences in the stimulusresponse curves, rheobase and early differences in hyperpolarizing TE. The finding that TA axons have a greater rheobase and have a rightward shift in the stimulus-response curve indicates that stronger stimuli were required to activate these axons. The magnitude of the differences was small and should be considered when assessing the importance of this finding. For example, rheobase differed on average by 0.06 mA that was much smaller than the SDs for this measure, which were 0.20 and 0.34 mA for TA and SOL axons respectively. Typically differences in rheobase have not been interpreted as resulting from I H . Similarly, the rate of activation of HCN channels is typically considered too slow to contribute to differences in hyperpolarizing TE at the 20-30 ms delay (Tomlinson et al., 2010). It is important to emphasize that the difference reported in Figure 2B was not a result of different extents of the fast change in threshold at the onset of the conditioning stimuli. The typical delay for overt expression of the effects of I H is closer to 100 ms in human studies but can be much earlier if the membrane potential is depolarized (Bostock et al., 1998). There is some evidence from nerve excitability studies in mice that axons in the sciatic nerve may appear to be depolarized, even if there is no reason to suppose this to be the case (Boerio et al., 2009(Boerio et al., , 2010. In any case, it can reasonably be argued that if there is membrane depolarization, it should affect the TA and SOL axons equally and facilitate the measurement of greater I H in SOL axons at earlier delays. Alternatively, differences in tissue impedance between rat and human experiments may facilitate earlier onset of I H in rat experiments. In conclusion, while for many nerve excitability measures we were not able to distinguish between slow and fast motor axons, they were significantly different during strong 100-200 ms hyperpolarizing conditioning stimuli. The axon threshold in these conditions is strongly affected by hyperpolarization-activated current I H and the findings suggest a greater level of I H in slow motor axons. This finding has implications for the use of nerve excitability testing in neurodegenerative conditions like amyotrophic lateral sclerosis (ALS). In the SOD1-G93A mouse model of ALS there is a clear motor unit type dependent vulnerability with fast-fatiguing motor units being most susceptible (Frey et al., 2000;Pun et al., 2006;Hegedus et al., 2008;Deforges et al., 2009;Kanning et al., 2010). As the vulnerable motor axons become denervated early in the disease the population of axons generating the nerve excitability measures will change. Longitudinal changes in nerve excitability measures (e.g., Kanai et al., 2006) may be confounded by the change in population of heterogeneous motor axons that contribute to the measure.

AUTHOR CONTRIBUTIONS
Chad Lorenz contributed to the acquisition, analysis and interpretation of the data, drafted the initial version of the manuscript, derived from a Master's thesis, and gave final approval for the submitted publication. Kelvin E. Jones conceived the study and contributed to acquisition, analysis, and interpretation of the data. The final manuscript was revised for publication by Kelvin E. Jones.

ACKNOWLEDGMENTS
This work would not have been possible without the expert technical assistance of Neil Tyreman. The research was supported by a Canadian Institutes of Health Research (CIHR) grant from the Neuromuscular Research Partnership (NRP) that includes support from Muscular Dystrophy Canada and the ALS Society of Canada.


extfieldsofstudy: Biology Medicine

text:
Epigenetic Control of the Genome—Lessons from Genomic Imprinting

Epigenetic mechanisms modulate genome function by writing, reading and erasing chromatin structural features. These have an impact on gene expression, contributing to the establishment, maintenance and dynamic changes in cellular properties in normal and abnormal situations. Great effort has recently been undertaken to catalogue the genome-wide patterns of epigenetic marks—creating reference epigenomes—which will deepen our understanding of their contributions to genome regulation and function with the promise of revealing further insights into disease etiology. The foundation for these global studies is the smaller scale experimentally-derived observations and questions that have arisen through the study of epigenetic mechanisms in model systems. One such system is genomic imprinting, a process causing the mono-allelic expression of genes in a parental-origin specific manner controlled by a hierarchy of epigenetic events that have taught us much about the dynamic interplay between key regulators of epigenetic control. Here, we summarize some of the most noteworthy lessons that studies on imprinting have revealed about epigenetic control on a wider scale. Specifically, we will consider what these studies have revealed about: the variety of relationships between DNA methylation and transcriptional control; the regulation of important protein-DNA interactions by DNA methylation; the interplay between DNA methylation and histone modifications; and the regulation and functions of long non-coding RNAs.

A Primer on Epigenetics, DNA Methylation and Histone Modifications
Epigenetic modifications perform three main functions in mammalian cells: they contribute to the control of chromosome architecture ensuring stability and appropriate segregation of chromosomes during mitosis; they contribute to regulation of the silencing and inaccessibility of repetitive elements and endogenous retroelements; and they can initiate and maintain the activity and repression of individual genes or clusters of genes. Here we focus on the role of epigenetic modifications in the control of mammalian transcription and the contribution of genomic imprinting studies to our understanding of epigenetic mechanisms.
In mammals, the different cells that make up an organism generally contain the same DNA yet their cellular morphology and function can vary greatly. This is largely a result of differential gene expression, which is developmentally regulated and can then be maintained after repeated cell divisions. The maintenance of expression states/levels requires heritable information to be passed through cell division to ensure propagation in each daughter cell, and it is this information that has been termed epigenetic. Further, cells are subject to dynamic changes in gene expression, dependent, for example, on intrinsic and extrinsic cues, which can be mediated through epigenetic processes. Epigenetic mechanisms include DNA methylation and post translational modifications to core histones. Other related components have been proposed as epigenetic such as non-coding RNAs (ncRNAs) and nucleosomal positioning, however these might also be considered mediators and/or facilitators of epigenetic states. The characterization and mapping of genome-wide epigenetic modifications represent an ever increasing field of research. These studies are revealing genome-wide patterns of epigenetic regulation that not only have confirmed many of the conclusions suggested from more traditional experimental approaches in model systems but also allow for the generation of new hypotheses that await experimental testing. One model system that contributed a foundation for these studies is the process of genomic imprinting.
DNA methylation is a process whereby a methyl (CH 3 ) group is added most commonly to a cytosine in DNA. In mammals it is generally found at CpG dinucleotides and can be correlated with gene repression in a variety of ways (discussed in more detail below). CpG sites are generally depleted in the genome, apart from stretches of DNA called CpG islands where CpG density is high. CpG islands can be concentrated at gene promoters and are generally unmethylated. CpG sites outside CpG islands are generally methylated (reviewed in [1,2])-resulting in a genome-wide methylation pattern that can be described as roughly bimodal. Acquisition of DNA methylation is catalyzed by a family of DNA methyltransferases (DNMTs, reviewed in [3]). DNMT1 has affinity for hemi-methylated DNA and is responsible for maintaining methylation after DNA replication and DNMT3A and DNMT3B catalyze de novo DNA methylation while the DNA methyltransferase homologue, DNMT3L acts as a cofactor and has no methyltransferase activity.
Waves of DNA methylation loss and acquisition are orchestrated during embryonic development. After fertilization the two parental genomes are mostly stripped of their epigenetic marks, a process that presumably "resets" the genome to a naive state applicable for pluripotency (DNA methylation at certain sequences in imprinted loci are among few genomic regions to "escape" this demethylation, see details in Section 2). Around blastocyst implantation de novo methylation then occurs and, to our knowledge, no further genome wide erasure/acquisition waves occur in somatic cells. Another wave of genome-wide reprogramming occurs in primordial germ cells (this time DNA methylation at imprinted loci is also lost, see details in Section 2); erasure of DNA methylation commences in the embryonic germline after embryonic day 7.5 (E7.5) in the mouse and progressive de novo methylation follows at E12.5 in prospermatogonia of male embryos, but occurs after birth in oocytes of female embryos (reviewed in [4]). This germline epigenetic reprogramming is required for generating functional germ cells and failure to do this appropriately usually results in infertility or developmentally abnormal embryos that die during gestation [5,6].
Covalent post-translational modifications to core histones (histone modifications henceforth) can impact the conformation of the nucleosome-nucleosome architecture within chromatin and influence its function such that some modifications are associated with an active chromatin state and others with a repressive state (for extensive review refer to [7]). The full repertoire of histone modifications is unknown, but is complex, with some specific amino acid residues influencing the ability of others to be modified, and some sites having the potential to be modified in multiple different ways. It is currently unclear whether many of the modifications truly are epigenetically heritable in a replication-dependent manner, like DNA methylation. Lysine methylation and lysine acetylation are among the best characterized histone modifications whose correlations with gene activity and repression have been extensively studied. Furthermore, enzymes involved in "writing" and "erasing" these epigenetic marks have been identified and characterized; histone lysine methyltransferases deposit methyl groups to lysine, and histone lysine demethylases remove them. Histone acetyltransferases (KATs) and histone deacetylases (HDACs) deposit and remove acetyl groups, respectively. Generally, regions with acetylated histones are associated with gene activity and regions devoid of acetylated histones are repressed, while associations between histone methylation and gene transcription are more site specific; histone 3 lysine 4 (H3K4) and H3K36 methylation are for example found on expressed genes while H3K9 methylation is associated with repressed genes. Their distribution in the genome can be associated with certain genomic motifs, e.g., gene regions such as promoters or open reading frames (ORFs), or intergenic regions such as repeats. For example, H3K4me3 (me3 denotes tri-methylated) is found at the promoters of active genes, whilst H3K4me1 is associated with enhancers, H3K20me3 is found at repressed repeat regions, and H3K9me3 at promoters of repressed genes, retroelements, imprinted loci and at pericentromeric repeat regions.

Genomic Imprinting and Targeting DNA Methylation
Genomic imprinting is a process causing the mono-allelic expression of a specific subset of mammalian genes in a parental origin specific manner (reviewed in [8,9])-i.e., genes that are expressed either from the paternally inherited chromosome or from the maternally inherited chromosome (paternal allele and maternal allele henceforth) are imprinted. The non-equivalence of parental genomes in mammals was discovered in 1984 [10,11], and individual imprinted genes were first discovered in 1991 (reviewed in [8]). Today, over 100 imprinted genes have been identified, most of which are organized in clusters and are regulated in a coordinated manner by a single imprinting control region (ICR) [9]. Most clusters contain at least one non coding gene and multiple protein coding genes, whose functions regulate embryonic development, placentation and a range of post-natal processes.
Epigenetic mechanisms allow the transcriptional machinery of the cell to distinguish the two parental chromosomes at imprinted loci and hence provide an important paradigm for understanding epigenetic control of gene activity and repression. Specifically, the discovery of differences in DNA methylation in the same place on the two parental chromosomes suggested the importance of epigenetic mechanisms in regulating imprinting [12,13] and the potential for epigenetic control in a wider context. The identification of imprinting control regions and their validation genetically as functional elements essential for the imprinting of multiple genes in cis, elucidated imprinting control. The loss of imprinting after targeted deletion of DNMT1 proved that DNA methylation was required for imprinting [14]. Importantly, in the absence of DNMT1, some imprinted genes were activated but others became repressed, an indication that methylation could impact activity as well as repression.
The acquisition of methylation at ICRs occurs in the germ line de novo by DNMT3A and DNMT3L with a small number of ICRs becoming methylated in sperm cells, and the majority acquiring methylation in oocytes-paternal and maternal ICRs, respectively. It is of interest that paternal ICRs are always located in intergenic regions while maternal ICRs are located at promoter sequences. Importantly, erasure of imprints occurs in the wave of demethylation that occurs in the primordial germ cells. However, in order to retain the memory of the parental origin that is subsequently established after that reprogramming, imprints must be retained during the post-fertilization epigenetic reprogramming phase [4]. Interestingly, other regions of the genome seem refractory to zygotic reprogramming [15] though these are not necessarily parent-specific or retained like imprints during development. The relationship, if any, of these regions to ICRs remains unclear. In addition to the ICR, other differentially methylated regions (DMRs) are located at some imprinted clusters, but a notable difference between ICRs and these DMRs is that differential methylation of the latter is not germline established, but rather is acquired post-fertilization. In all cases, these so-called secondary DMRs-to distinguish them from regions such as ICRs that acquire differential methylation in the germline-require the ICR for their establishment. The mechanisms through which ICRs control gene expression in their respective clusters are diverse and remain the subject of active research, including analysis of regulation by ncRNAs and of the relationships between DNA methylation and histone and non-histone proteins.
Both in imprinted and non-imprinted contexts, little is known about why certain DNA sequences become methylated and not others, or how this may change dynamically within a sequence such as a particular CpG island at a gene promoter. Most likely, it is a process that must be targeted in some manner. Targeting of the DNA methylation machinery has received much attention and efforts made to identify intrinsic sequence specificities of DNMTs and their cofactors. It has thus generally been assumed that the acquisition of methylation represents the "active" process in establishing differential methylation. However, recent studies on DMRs in the germlines and their propagation after fertilization suggest it might also be protection from DNA methylation and maintenance at methylated regions that determine differential methylation ( Figure 1A, reviewed in [16]): Rather than appearing as discrete methylated sequences in otherwise unmethylated regions, maternal ICRs (which represent the vast majority of ICRs) are surrounded by methylation at both flanks. In contrast, these ICRs are unmethylated in sperm but are also flanked by methylation at surrounding sequences, suggesting that DNA methylation may be the "default" state and that it is protection from methylation at the ICRs, and perhaps other non-imprinted sequences as well, that establishes their differential methylation. Furthermore, in the germline, far more sequences are differentially methylated between oocytes and sperm than the ICRs; recent genome-wide studies suggest they are in the counts of thousands in oocytes and hundreds in sperm [15,17,18]. In contrast to ICRs these sequences generally lose methylation after fertilization, suggesting targeted maintenance of DNA methylation at specific sequences is essential for the germline-derived differential methylation of imprinted loci. Hence perhaps, loss of maintenance, in addition to active removal of DNA methylation at non-imprinted loci, contributes to the mechanism through which demethylation occurs in somatic cells. KRAB zinc finger proteins (ZFP) represent a family of over 350 tetrapod-specific genes whose functions remain poorly understood. They bind DNA and have previously been shown to recruit the repressive chromatin machinery in a site-specific manner. One of these KRAB-ZFPs, ZFP57, has been shown to be required to maintain the DNA methylation memory at imprints during post-fertilization reprogramming when the bulk of the genome is changing its epigenetic state [19]. ZFP57 binds methylated DNA and is thought to recruit methyltransferases to imprinting control regions hence preventing them from loss of their imprints.

DNA Methylation Correlates with Repression
The correlation between DNA methylation and gene repression was noted in several experiments assaying viral and endogenous gene expression in mammalian, frog and sea urchin cells in the late 1970s and early 80s [20][21][22][23][24][25][26][27][28][29][30]. Experiments were conducted to determine whether the observed relationship was purely correlational, or whether DNA methylation functionally regulated gene expression. This was, however, challenging, but the strong evidence in many different contexts, showing that hypomethylated regions were associated with activity and hypermethylated regions refractory to transcription, suggested that absence of DNA methylation may be necessary though not sufficient for transcription. Vardimon et al. injected bacterial plasmids containing in vitro methylated or unmethylated DNA encoding a viral gene into frog oocyte nuclei [31]. They observed maintenance of the respective methylation states over a 24 h period, and expression of the gene in oocytes that were injected with unmethylated DNA but not in those that were injected with methylated DNA [31]. In a similar experiment, Stein et al. transfected in vitro methylated or unmethylated plasmids containing the Aprt (adenine phosphoribosiltransferase) gene into cultured Aprt null mouse cells. They observed maintenance of the respective Aprt methylation states after integration into the endogenous genome over several cell divisions for both unmethylated and methylated plasmids, and that integration of the unmethylated but not the methylated gene rescued the Aprt null phenotype, suggesting methylation of the gene was associated with inhibition of its transcription [32].
Correlations between gene expression and DNA methylation have been assessed at CpG sites across whole chromosomes or the whole genome. Consistent with the earlier studies, DNA methylation of promoter sequences, though rare at CpG island promoters, was observed to correlate with gene repression [33][34][35]. The functional role of DNA methylation in repressing gene expression is further suggested by results from studies in which the genes encoding the DNA methyltransferases are deleted conditionally in various cell lineages. Generally, the loss of DNMTs results in dysregulation of multiple genes, with a trend towards gene activation rather than silencing, again suggesting that DNA methylation represses gene expression (reviewed in [36]). Furthermore, treatment of cells in vivo with the DNA methyltransferase inhibitor 5-Azacytidine was shown to result in gene activation in several experiments in the 1980s, with concomitant loss of DNA methylation (reviewed in [37]). Together all these findings have led to the general assumption that loss and acquisition of DNA methylation at a gene promoter results in gene activation and silencing, respectively, but none actually proved that the acquisition of DNA methylation itself causes the gene silencing in all contexts.

DNA Methylation as a Consequence of Transcriptional Silencing
Studies of the temporal onset of mono-allelic expression of imprinted genes and the acquisition of differential methylation at secondary DMRs during mouse development indicate that DNA methylation can be acquired after gene repression ( Figure 1B). The imprinted genes Gtl2, Cdkn1C, H19 and Igf2r each contain a secondary DMR in their promoters, which become differentially methylated days after their mono-allelic expression is observed (summarized in [38]). Generally, mono-allelic expression of these genes is initiated around the morula or blastocyst stage (E3.5-4.5), while differential methylation of the respective secondary DMR occurs after E6.5 [13,[38][39][40][41][42]. In the most extreme case, Igf2r is mono-allelically expressed from the maternal allele from E6.5 onward but the silent paternal allele only becomes methylated at or after E15.5 [13,42]. It is reasonable to assume that this temporal relationship, where methylation is acquired as a consequence of gene repression, also applies to non-imprinted genes ( Figure 1B). In particular, is has recently been shown that DNA methylation levels are secondary to the binding of transcription factors; Stadler et al. [43] identified multiple clusters of CpG sites that have low to intermediate levels of methylation, 10%-50%, in mouse embryonic stem (ES) cells. These low methylated regions (LMRs) are likely distal regulatory regions, and are bound by various transcription factors. Scrambling binding sites for the insulator protein CTCF or knocking out the transcription factor REST led to increased methylation at the LMRs. Furthermore, reintroduction of REST into the REST í/í cells reverted the methylation status of the LMRs to the normal low levels [43]. These findings suggest DNA methylation may not have a direct role in silencing gene expression in all situations. In such cases DNA methylation might rather be acquired after gene silencing to maintain the repressed state or as a secondary readout of other mechanisms of genome control. Nonetheless, there are situations where acquisition of DNA methylation unquestionably does regulate gene expression, notably at the germline DMRs of imprinted genes [1,8,9,[14][15][16].

Proteins Attracted and Repelled
In situations where DNA methylation does indeed direct gene repression there are currently two model mechanisms that are generally acknowledged [1,44]: First, DNA methylation can attract proteins that bring about gene repression through recruitment of chromatin modifiers. A group of proteins, collectively referred to as methyl binding proteins (MBPs) have been characterized and shown to specifically bind to methylated, but not unmethylated, DNA [44][45][46][47][48][49]. MBPs are known to interact with histone modifiers such as HDACs, e.g., in forming complexes, such as the nucleosome remodeling deacetylase (NuRD) complex, which through their histone deacetylase activity and subsequent chromatin condensation bring about gene repression [50][51][52][53][54][55]. Secondly, certain proteins may interact with DNA in a methylation dependent manner. Here, DNA methylation may be refractory to the binding of proteins, such as transcription factors or other regulatory proteins [56][57][58], that are necessary for gene expression ( Figure 1C). For this latter model, the best characterized example is the regulation of CTCF binding at the imprinted H19/Igf2 cluster via differential DNA methylation on the two parental alleles (reviewed in [1]).

Regulation of CTCF Binding at the H19/Igf2 Imprinted Cluster; the Insulator Mechanism
In the H19/Igf2 imprinted cluster, the protein coding gene Igf2 is expressed from the paternally inherited allele [59]. This expression pattern is dependent on the regional ICR [60], on its differential methylation [12,14,61] and on the insulator protein CTCF binding to the ICR. On the unmethylated maternal allele, CTCF can bind, while its binding is inhibited on the methylated paternally inherited chromosome [62][63][64][65]-thus CTCF binding to DNA is methylation-sensitive ( Figure 1C). Igf2 and a downstream non-coding RNA gene, H19, share enhancers that are located at the 3' end of H19 [66,67] and the parental specific expression of Igf2 and H19 are ultimately determined by interaction with these sequences; on the paternally inherited chromosome, Igf2-enchancer interaction is possible and the gene is expressed. On the maternally inherited chromosome this contact is blocked by CTCF binding to the ICR and this facilitates enhancer interaction with a now active H19 instead, and also results in Igf2 repression.
What is the mechanism of CTCF's enhancer blocking activity? The current model (reviewed in [68]) suggests that in the H19/Igf2 cluster, chromatin loop formation on the maternal allele spatially inhibits enhancer interaction with Igf2. The process appears to depend on three elements; dimerization, CTCF binding to more than one region and physical contact between these neighboring sites via CTCF interaction [69][70][71][72][73][74]. The model suggests that on the unmethylated maternally inherited chromosome, CTCF binds to the ICR and also to an upstream somatic DMR located 5' of Igf2. Binding does not occur at the paternal allele where methylation inhibits the binding. On the maternal allele ICR-DMR contact is made possible by CTCF dimerization bringing together the two distinct loci, and because they flank Igf2, the gene is 'looped out' ( Figure 1D). Further chromatin contacts within the cluster, some facilitated by CTCF, then result in physical separation between the Igf2 loop and the enhancers. Recently cohesins have been shown to bind to over half of CTCF binding sites in the genome, including in the H19/Igf2 cluster [75]. Given the ability of cohesins to tether DNA strands (i.e., sister chromatids after cell's S-phase) it is possible that cohesins contribute mechanistically to these chromatin contacts on the maternal H19/Igf2 locus. On the paternal allele, where CTCF cannot bind, long-range chromatin interactions are not observed within the cluster, suggesting a state that allows interaction between the 90 kb distant enhancers and Igf2 ( Figure 1D) [73]. Similar interactions involving CTCF have been noted at other loci ( Figure 1D).

Relationship between DNA Methylation and Histone Modifications
Similar to DNA methylation, correlation between multiple histone modifications in various genomic elements, including promoters, have been associated with gene activity and repression, and early studies illustrating this indeed investigated the relationship in the context of imprinted loci [76][77][78][79][80][81][82]. A functional relationship may therefore potentially exist between DNA methylation and histone modifications whereby the acquisition of one may be dependent on, or mutually exclusive, with the other. Indeed, as noted above, MBPs can recruit histone modification enzymes. Well-defined examples of histone modifications that regulate de novo DNA methylation are however scarce [83,84]. One very compelling example again comes from the study of genomic imprinting, as discussed below.
DNMT3L lacks a DNA methyltransferase activity, but it is necessary for methylation of DNA in certain situations [85,86] because it forms a complex with DNMT3A and DNMT3B, impacts their activity and contributes to their structural interaction with chromatin [87][88][89][90]. The ability is likely a result of a recently discovered affinity of DNMT3L to histone H3 [90] and this interaction is dependent on the methylation state of H3 at lysine K4-the binding only occurs when the histone is unmethylated hence H3 methylation might shield from DNA methylation [90]. A functional role for H3K4 methylation in modulating DNA methylation came from an imprinting study where Ciccone et al. showed that this interaction has important regulatory implications. The group generated mice deficient for a H3K4 demethylase enzyme, KDM1B, which resulted in increased H3K4 methylation in oocytes, where KDM1B is almost exclusively expressed. Consistent with inhibition of the DNMT3L-DNMT3A complex binding to methylated histone H3, DMRs at four imprinted regions that normally acquire DNA methylation in the female germ line were unmethylated in the Kdm1b null oocytes and imprinted expression of the corresponding genes was lost in embryos from Kdm1b null females ( Figure 1E) [91]. These results strongly suggest a functional link between loss of H3K4 methylation and acquisition of DNA methylation, at least at imprinted regions ( Figure 1E).
Cedar and Bergman take this further proposing a model of how the bimodal methylation pattern of mammalian genomes may be dependent on this same relationship. They suggest that de novo DNA methylation at the blastocyst stage is prevented at particular loci by deposition of H3K4 methylation. They further suggest that H3K4 methyltransferases may be targeted to CpG islands by RNA polymerase II and as a consequence, the DNA methyltransferase machinery containing DNMT3L, cannot access CpG sites in regulatory regions that are CpG islands [84].
H3K9 di-and trimethylation is associated with repressive DNA. DNA methylation is often found at such regions. Furthermore, DNA is globally hypomethylated in mouse ES cells carrying deletion of a H3K9 methyltransferase, G9a [92]. In this case the loss of DNA methylation is not a result of the aberrantly low levels of histone methylation, but rather due to loss of the histone methyltransferase enzyme itself; the DNA methyltransferase machinery interacts with G9a, and this interaction is mediated through a protein domain that is independent of the histone methyltransferase catalytic activity by a SET protein domain. Therefore, in G9a í/í mouse ES cells carrying G9a transgenes that lack histone methyltransferase activity, e.g., due to a point mutation in the SET domain, DNA methylation levels are partially rescued [93,94]. Regulation of DNA methylation through interaction of the DNMTs with histone modifiers, rather than with the histone modifications themselves, seems to be common and is observed for multiple mammalian histone methyltransferases [95][96][97], as well as in plant [98] and fungal systems [99]. Interestingly, in G9a í/í ES cells DNA methylation is lost at some imprinted loci [94,100], but where tested this is not observed in embryos [100,101]. This behaviour at imprints may suggest that ES cell culture is not a faithful model for assessing a requirement for histone modifying enzymes in DNA methylation, but equally might also reflect different properties of imprint-specific maintenance in ES cells compared to in vivo. (A) left: In the male germline (sperm), CpG dense regions are generally unmethylated and less dense regions are methylated. In the female germline (oocytes) CpG rich regions are more frequently methylated. This results in multiple differentially methylated regions between the male and female germlines. After fertilization only a small subset of these regions retain differential methylation. Retention of differential methylation at imprinting control regions (diamonds) post-fertilization may therefore be a targeted protection from either demethylation or de novo methylation; right: Model; Changes in DNA methylation may be mediated through loss and gain of such protection-when protection is lost (e.g., upper-as a result of factor (black triangle) binding or a histone modification that is non-permissive (red circle) for DNMT binding) CpGs become methylated by the methyltransferase machinery. If protection is gained (lower) the machinery cannot access the CpG sites to maintain methylation and after cell divisions methylation is therefore lost; (B) left: The secondary DMR located in the promoter region of the imprinted gene Gtl2 becomes methylated on the paternal allele after expression is silenced; right: Model; DNA methylation at CpG sites in promoter regions of non-imprinted genes may therefore, at least in some cases, occur after gene silencing; (C) left: In the H19/Igf2 imprinted locus, CTCF (red pentagon) binds the H19-ICR, on the unmethylated maternal allele, not the methylated paternal allele; right: Model; Methylation of CpG sites can inhibit protein binding (purple) to DNA; (D) left: In the H19-Igf2 imprinted locus CTCF (red pentagon) binds to regions flanking Igf2 and dimerizes, looping the gene and physically inhibiting its interaction with distal enhancers. On the paternally inherited allele, CTCF does not bind and enhancers are in contact with Igf2 and the gene is expressed; right: Model; Looping of DNA sequences through the action of CTCF (red pentagon) can separate regions or bring them into contact; (E) left: In Kdm1b í/í (histone methyltransferase) mouse oocytes, imprints are not established at multiple ICRs due to the inhibitory effect of H3K4 methylation on DNMT3L. Histone modification states in WT and Kdm1b í/í mice are depicted as green and red circles to signify permissive and non-permissiveness to de novo DNA methylation, respectively. In embryos from Kdm1b í/í mothers, imprinted expression is lost, and genes are biallelically expressed (Mest, depicted) or repressed; right: Model; Histone modifications (red and green circles) can regulate DNA methylation; (F) left: In the Igf2r imprinted locus Igf2r expression is inhibited by transcriptional interference from the Airn lncRNA transcript on the paternal allele. The lncRNA recruits histone modifiers such as G9a (blue) to proximal imprinted genes that contribute to silencing of the imprinted Slc22a3 in a lineage specific manner, e.g., through deposition of histone marks that are non-permissive for transcription (red circle); upper right: Model; lncRNAs may exert their effects in trans at proximal genes. As illustrated, a lncRNA is expressed and silences proximal genes, but not the more distal genes; lower right: Model; In the example provided a lncRNA and a coding gene are expressed from within the same ORF. Transcription of the lncRNA inhibits expression of the coding gene.

lncRNAs, Definition, Characterization and Potential Functions
In recent years the roles of long noncoding RNAs (lncRNAs) in regulating genome function have received considerable attention, and are now emerging as a large group of genes with potential functions of fundamental importance for cell biology (for review see [2,[102][103][104]). lncRNAs are defined as noncoding RNA transcripts of >200 bp [104]. Transcription of lncRNAs resembles that of mRNA genes; they are transcribed by the same transcriptional machinery and by RNA polymerase II, the transcripts are 5' capped and can be spliced and shuttled to the cytoplasm [102]. The lack of an open reading frame and their size are therefore the only criteria that currently define lncRNAs as a group [102,104]. On basis of high-throughput RNA sequencing experiments, the numbers of lncRNA transcripts have been suggested to range between 5000-15,000 [105,106]. With higher sensitivity, targeted capture experiments have identified lncRNAs that are undetectable by high-throughput technology, suggesting that this range is an underestimate [107]. However, as a result of their loosely defined criteria, lncRNAs as a group may be very heterogeneous. Therefore, the functional roles discussed below may only apply to a subset of their estimated numbers.
Despite the current excitement surrounding "new" roles for lncRNAs, they were shown to regulate genomic imprinting over a decade ago. Multiple potential functions of lncRNAs have been proposed whereby lncRNAs either exert their effects by acting in trans or by the act of their transcription in cis (transcriptional interference). Both effects have been shown to act at imprinted loci. Some trans acting lncRNAs, such as HOTAIR, have been suggested to exert their effects throughout the genome [108], while others, including most imprinted lncRNAs defined to date, act over a limited area surrounding or close to their transcriptional origin. Some lncRNAs may utilize both cis and trans acting mechanisms. An example is the imprinted Airn lncRNA whose transcription on the paternal chromosome represses Igf2r expression in cis by transcriptional interference [109], while the Airn RNA molecule itself is also necessary for regulating other genes in the cluster in a trans-targeted manner (see below, and Figure 1F [110]). Transcriptional interference is proposed to occur as a result of a collision between the transcriptional machineries of two adjacent or overlapping transcripts which might result in termination of one or both transcriptional events. Alternatively it may occur by promoter occlusion via inhibition of formation of a transcriptional initiation complex due to existing transcription of one transcript through the promoter of another [111].
Functions of trans acting lncRNAs have been proposed to fall into the following categories [112]: (1) Decoys: lncRNAs that bind to DNA binding proteins and prevent their interaction with DNA; (2) Scaffolds: lncRNA that function to join two or more proteins into an lncRNA-RNP (ribonucleoprotein) complex; (3) Guides: lncRNAs that bind proteins to guide them to certain genomic locations, e.g., by lending them specificity and/or binding capacity to certain DNA sequences or chromatin states.

lncRNAs in the Epigenetic Control of Genome Function-Lessons from Imprinting
Every cluster of imprinted genes contains at least one lncRNA and these lncRNAs are regulated by DNA methylation. This was demonstrated in experiments where the genes encoding DNA methyltransferases were deleted in mice to gauge effects on imprinting regulation. Promoters for the Airn, Nespas/Gnasxl, Snrpn and Kcnq1ot1 lncRNA genes lie within the ICR for their respective region and are differentially methylated on the two parental chromosomes. Upon loss of DNMT1, the maintenance methyltransferase, methylation is lost at these ICRs in E10.5 embryos (the genetic manipulation is lethal at later embryonic stages) and Airn, Nespas/Gnasxl, Snrpn and Kcnq1ot1 are biallelically expressed, with effects on neighboring imprinted protein coding genes, some of which may lose imprinting as a result of the lncRNA dysregulation [14,113,114]. Kcnq1ot1 and Airn promoters are located in the ICRs, exhibit differential methylation, and, importantly, are located within genes running antisense to them, hence these provide examples of critical regulatory DNA methylation at genomic regions considered by some to have little or no consequence, i.e., intragenic. The existence of other epigenetically regulated elements within genes and acting in this way to potentially regulate lncRNAs, may have very widespread effects on genome function. Furthermore, imprinted lncRNAs have been demonstrated to be necessary for epigenetic control of genome function, to guide chromatin modifying enzymes in trans to specific sites in the genome. This is thought to mediate changes in histone modifications and be associated with changes in transcriptional activity. Although challenging to address experimentally, this function for lncRNAs is currently the topic of much attention. It was studies on imprinted gene regulation at the Igf2r/Airn and Kcnq1/Kcnq1ot1 imprinted clusters that provided examples of this type of regulation [104]. Kcnq1ot1 and Airn are estimated as greater than 100 kb lncRNA transcripts, transcribed in an antisense orientation from within protein coding genes; Kcnq1ot1 from Kcnq1 in a 1 Mb imprinted cluster that contains eight maternally expressed protein coding genes, and Airn from Igf2r in a 400 kb long imprinted cluster that contains three maternally expressed protein coding genes. Both transcripts generate unspliced lncRNAs that are localized in the nucleus [103,115,116]. The ICRs of both genes are methylated on the maternal, but not paternally inherited chromosome, and determine monoallelic expression of the lncRNAs from the paternal allele. In mouse genetic mutants, where the promoters of Airn and Kcnq1ot1 are deleted or their transcripts truncated by insertion of premature polyA sequence into the endogenous genes, biallelic expression of the imprinted protein coding genes occurs within their respective clusters [110,[117][118][119]. These results suggested that the lncRNAs or the act of their transcription is necessary for silencing of genes in cis ( Figure 1F). In addition, several lines of evidence further indicate that lncRNAs guide chromatin modifying enzymes in trans to establish repressive histone marks and gene silencing on the paternal allele ( Figure 1F): In the Kcnq1 imprinted cluster Osbpl5, Cd81, Ascl2 and Tscc4 are imprinted exclusively in the placenta [40] and so are Slc22a2 and Slc22a3 in the Igf2r/Airn cluster [120]. The paternal chromosomes are bound by the histone methyltransferases G9a and/or Ezh-Eed2 in the extraembryonic lineage [40,110], and both Airn and Kcnq1ot1 lncRNAs associate with G9a histone methyltransferase in a lineage specific manner-in placenta but not embryo [110,115]. These results showed that lncRNAs may be a contributing factor for targeting epigenetic marks ( Figure 1F) with genetic models being used alongside biochemical approaches to generate a more tractable and comparable experimental paradigm for added robustness. These studies have paved the way for explorations of the roles of multiple other lncRNAs which are found in association with different chromatin modifying enzymes [121,122]. Most recently, the imprinted lncRNA Gtl2/Meg3 has been shown to function in trans to target polycomb regulatory complexes in mouse and human stem cells in culture [123].

Conclusions
The robust genetic approaches applied to the regulation of imprinting have allowed it to be an excellent hypothesis-driven model to investigate and understand the epigenetic control of genome regulation. One of its greatest strengths as a model is that it allows the comparison of differentially expressed alleles of the two inherited copies of a gene with identical sequence within the same cell. Because these two parentally inherited alleles have well-defined different epigenetic states the contributions of these to gene expression can be determined. Since imprinted clusters employ multiple different epigenetic mechanisms, acting through various different mediators (long non-coding RNA, CTCF, etc.), this has enabled investigators to explore their hierarchical interactions and relationships with one another. As evidenced by the examples presented here, imprinting has provided insight into some of the most fundamental aspects of a range of epigenetic phenomena and their mediators.
Nevertheless, many important aspects of imprinting and epigenetic control remain to be elucidated. These include: what allows epigenetic marks to be de novo targeted differently in the male and female germlines; whether they are modulated by extrinsic or intrinsic signals, for example in the context of development and disease; and how DNA methylation is actively removed during reprogramming and perhaps at other times in development. The mechanisms regulating some of these processes are beginning to emerge where the context of imprinting has contributed; the DNA binding proteins ZFP57 and PGC/Stella have been shown to target and maintain DNA methylation at imprinted clusters during postfertilisation epigenetic reprogramming [19,124] and selective loss of imprinting is necessary for stem cell regulation in the neurogenic niche of the developing mouse [125]. Whether we can apply more generally what we learn from these mechanisms-for example about the general targeting of epigenetic states or the dynamic changes in epigenetic state in specific cellular nichesremains to be determined. It is likely that future studies, addressing these and other similarly fundamental questions in the context of imprinting will continue to add new layers to our understanding of genome regulation and the epigenetic control of genome function more widely.