The Effects of Replication Stress on S Phase Histone Management and Epigenetic Memory

Review DCC

YJMBI-65262; No. of pages: 19; 4C:4, 9, 10, 11

The Effects of Replication Stress on S Phase Histone Management and Epigenetic Memory Saša Šviković and Julian E. Sale Division of Protein & Nucleic Acid Chemistry, Medical Research Council Laboratory of Molecular Biology, Francis Crick Avenue, Cambridge, CB2 0QH, UK

Correspondence to Julian E. Sale: [email protected] http://dx.doi.org/10.1016/j.jmb.2016.11.011 Edited by T Kutateladze

Abstract When a cell divides, it must not only accurately duplicate its genome but also recapitulate its programme of gene expression. A significant body of evidence suggests that an important fraction of the information specifying the transcriptional programme of vertebrate cells is carried epigenetically by post-translational modifications of histone proteins. For such a system to operate, propagation of key histone marks must be coupled to replication such that they remain correctly associated with the underlying DNA sequence, despite the huge disruption to chromatin structure generated by unwinding the parental DNA strands. Focusing on vertebrate cells but drawing on experimental evidence from a wide range of systems, we will examine the evidence that histone mark propagation through replication contributes to transcriptional stability. We then discuss the emerging molecular mechanisms that ensure that histone recycling is tightly coupled to DNA replication, focusing on how parental histone proteins are chaperoned around the replication fork, and the strategies that ensure that this process is not disrupted by impediments to replication. © 2016 Elsevier Ltd. All rights reserved.

Introduction Chromatin represents an elegant solution to the complex problem of compacting and protecting extremely long and charged DNA molecules within the eukaryotic nucleus while, at the same time, allowing access to the genetic code. The basic subunit of chromatin, the nucleosome, comprises eight histone proteins, a tetramer of (H3/H4)2 bound by two H2A/H2B dimers. Around this core, approximately 146 bp of double-stranded DNA is wrapped in 1.65 helical turns. Nucleosomes are spaced at approximately 200 bp intervals with the intervening DNA bound by the linker histone, H1 (reviewed in Ref. [1]). Nucleosomes are intrinsically inhibitory to transcription [2,3], and thus, efficient gene expression requires that the DNA is made accessible by nucleosome modification or displacement. Conversely, gene repression can be enforced by the formation of heterochromatin, a compact and higher-order chromatin structure that renders the DNA inaccessible to transcription factors and RNA polymerase. 0022-2836/© 2016 Elsevier Ltd. All rights reserved.

The observation that histone modifications could influence RNA synthesis from chromatin templates [4,5] suggested that histones might provide a basis for regulating gene expression. In turn, this developed into the notion that cells might propagate chromatin structures able to act as an “epigenetic code” determining the transcriptional competence of individual genes [6]. The subsequent elucidation of the general structure of chromatin [7–9] and the nucleosome itself [10,11], coupled with a growing appreciation of the potential role of histone modifications in regulating transcription, led to the suggestion that these modifications might not just be effectors of chromatin accessibility but that they also carry epigenetic information [12]. A vast array of histone post-translational modifications have been described [13]. They are concentrated in the unstructured N-terminal tails of the histone proteins, which protrude through the gyres of the DNA in the nucleosome structure [11] but are also found in the globular core where they can directly influence electrostatic interactions between the histones and the DNA [14]. Furthermore, J Mol Biol (2016) xx, xxx–xxx

Please cite this article as: S. Šviković, J. E. Sale, The Effects of Replication Stress on S Phase Histone Management and Epigenetic Memory, J. Mol. Biol. (2016), http://dx.doi.org/10.1016/j.jmb.2016.11.011

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numerous studies have revealed correlations between these “chromatin marks” and the transcriptional state of the underlying DNA [15–17]. Thus, for example, transcriptionally active chromatin is linked with the trimethylation of lysine 4 and the acetylation of lysines 9 and 14 of histone H3, or H3K4me3 and H3K9/14ac for short. Conversely, transcriptionally inactive regions of the genome feature H3K9me3 or H3K27me3. However, whether any of this myriad of histone modifications are truly vectors of epigenetic memory, rather than simply being effectors of chromatin function, or indeed mere epiphenomena, remains contentious [18–20]. Indeed, there is evidence for multiple mechanisms contributing to the robustness of transcriptional states through replication including transcription factor segregation, RNA interference, and DNA methylation, and we have previously considered how these mechanisms may interact with histone-modification-based epigenetic memory [21]. Here, we will focus on the role of histone modifications and their propagation in the maintenance of epigenetic memory over repeated cell divisions. For a histone modification or group of modifications to be deemed to carry epigenetic information through DNA replication, three criteria must be met: (1) The histone and modification should be sufficiently stable during interphase that any epigenetic information carried is not lost before the next round of replication. (2) During replication, the modification must be propagated from the parental chromatin to that formed on the daughter strands while remaining in register with the underlying DNA sequence. (3) Once present on the daughter chromatin, the modification should be capable of promoting its own installation on nearby, newly synthesised histones that are incorporated into the nascent daughter strands in order to maintain nucleosome density. If these criteria are met, and the propagated histone modifications are necessary to specify a stable transcriptional state, then an important prediction is that the interruption of histone flow during replication should result in failure to maintain the transcriptional state associated with the modification. In the four sections of this review, we will examine the extent to which these criteria are met and then discuss the prediction by exploring the epigenetic consequences of failure to maintain processive replication.

Global Turnover of Histones and Their Modifications The ability of histone post-translational modifications to make a viable contribution to epigenetic memory, that is, to act as epigenetic marks, will be influenced by the overall stability of the modification and the host histones. H2A/B dimers can be removed from the nucleosome without the loss of DNA

wrapping [22] and are turned over rapidly, particularly during transcription [23–25]. In contrast, H3 and H4 are turned over less frequently, making them more attractive as vectors of mitotic epigenetic information. Nonetheless, the idea that H3 and H4 are sufficiently stable to transmit epigenetic information across cell divisions has been challenged by pulse-labelling experiments in Drosophila cells, suggesting that the time for a histone protein to turn over is, on average, less than a cell cycle [26]. However, even the maximum observed turnover rate is less than that caused by S phase when parental nucleosomes are diluted by 50%, a situation from which the levels of many key histone modifications recover during the ensuing G1 phase [27]. Thus, the potential for histone turnover to disrupt the propagation of epigenetic information will also be determined by the ability of adjacent nucleosomes to “reeducate” newly incorporated molecules [28], a point we return to below. In terms of histone modifications, the half-life of histone methylations is generally significantly longer than that of histone acetylations [29–33], consistent with the suggestion that histone acetylations are generally more likely to be short-term effectors of histone function [19]. Thus, the H3/4 tetramer and its methylation modifications will form the focus of our discussion as the most likely histone modifications to participate in epigenetic memory.

Propagation of Marked Parental Histones through DNA Replication The recycling of parental histone proteins displaced ahead of the replication fork not only constitutes a ready supply of histones close to the fork but also provides a potential vector for carrying epigenetic information. The general rules governing histone recycling were largely worked out in classic biochemical experiments during the 1970s and 80s and still provide a framework for understanding how histones could transmit epigenetic information. During S phase, the reestablishment of chromatin is closely coupled to replication in both human and yeast cells [34–38]. Both parental histone proteins (those present prior to replication and displaced ahead of the replicative helicase) and newly synthesised histones contribute to the formation of chromatin on the daughter DNA strands. Behind the replication fork, (H3/4)2 tetramers are deposited on both leading and lagging strands as soon as sufficient double-stranded DNA is synthesised [39–43]. This histone recycling takes places largely without the mixing of old and new H3/4 dimers [44–46], although increased tetramer splitting has been noted in transcribed regions in both yeast [47] and human cells [46]. Nucleosome density is maintained by interspersing newly synthesised H3/4 between the parental tetramers, such that on average the old:new histone ratio on each new strand is 50:50. In contrast, the association of H2A and H2B with the newly deposited

Please cite this article as: S. Šviković, J. E. Sale, The Effects of Replication Stress on S Phase Histone Management and Epigenetic Memory, J. Mol. Biol. (2016), http://dx.doi.org/10.1016/j.jmb.2016.11.011

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H3/4 tetramers is not tightly coupled to DNA replication [34], and when H2A/B is incorporated, the dimers are drawn randomly from pools of parental and newly synthesised protein [23,48], further supporting the idea that H2A/B are not attractive candidates for the carriers of epigenetic information. Strong evidence that this histone recycling mechanism is required for the transfer of parental H3/4 marks to the newly formed chromatin after replication in human cells has been provided by the capture of proteins associated with nascent DNA coupled to sensitive mass spectrometry, which has allowed the analysis of both parental and new histones associated with the replication fork [27]. Broadly similar conclusions have been reached on the transmission of histones and their modifications through replication in budding yeast [38,49,50]. In contrast, evidence has been presented that, in Drosophila, histone marks are globally erased during replication and that the resetting of the parental chromatin state is dependent on specific DNA and histone-modifying enzymes [51,52]. We will next look at a more recent work, from a variety of experimental systems, which has begun to reveal the molecular details underlying the management of histone proteins by the replisome, from their eviction ahead of the replicative CMG helicase [named after its key components in budding yeast, Cdc45, MCM2–7, and the GINS (go-ichi-ni-san) complex comprising Sld5, Psf1, Psf2, and Psf3], through being passed back across the replication fork, to their installation on the nascent daughter strands. Eviction of parental nucleosomes ahead of the replisome The forces generated by the replicative helicase when unwinding DNA are able to destabilise one or two nucleosomes ahead of the replication fork [41,53–55]. Indeed, unwinding a DNA helix in vitro with optical tweezers can displace a histone octamer [56]. Superhelical torsion generated during replication and the accumulation of positive supercoiling ahead of the fork could also contribute to nucleosome disassembly. However, in vitro, positive superhelical stress alone leads to the eviction of only H2A/B dimers, leaving the (H3–H4)2 tetramer associated with DNA [57]. Indeed, during both SV40-dependent chromatin replication [41,53,58] and the replication of the rDNA in budding yeast [59], nucleosomes are not completely evicted, but rather destabilised, remaining loosely attached to DNA. Thus, while the physical forces generated during replication, either the unzipping of nucleosomal DNA or superhelical stress, are an appealing and simple solution to nucleosome eviction, it is unclear that they are, in vivo, sufficient on their own. Intriguingly, the conserved Dpb3 and Dpb4 subunits of DNA polymerase ε form a histone-like fold (similar to H2A/B) that is capable of binding double-stranded DNA [60,61]. In budding yeast, the disruption of Dpb3 or

Dpb4 leads to loss of silencing at mating type loci and telomeric regions [62–64] similar to that observed in mutants defective in replication-coupled chromatin assembly [65], suggesting that these proteins also play a role in histone management during replication. It has been proposed that Dpb3/Dpb4, which are also found in the Chromatin Accessibility Complex (CHRAC) chromatin remodelling complex, may represent a “built-in” mechanism by which the replication machinery could facilitate nucleosome disruption [66]. Such an activity linked to Polε appears counterintuitive when considering the familiar model of DNA replication with the helicase lying ahead of the DNA polymerase and Proliferating Cell Nuclear Antigen (PCNA) sliding clamp (Fig. 1a). While this model is certainly supported for DNA synthesis by Polδ, an interaction with the PCNA is not essential for robust DNA synthesis by Polε, which instead possesses a unique DNA binding “processivity domain” [67]. Furthermore, Polε, which appears to operate on the leading strand [68,69], interacts with the CMG helicase [70–72], and remarkable, recent cryo-electron microscopy data suggest that it may lie adjacent to or ahead of the CMG complex [73,74]. Although Sun et al. were unable to locate Dpb3/ 4 in the structure [73], we speculate that this position of Polε could make Dpb3/4 well placed to help destabilise parental nucleosomes and the coupling of leading strand DNA synthesis with nucleosome displacement (Fig. 1b). Chaperoning parental histones to the daughter DNA strands Since histones are highly basic proteins evolved to interact with DNA, they are generally bound to chaperone proteins when not complexed with DNA (reviewed in Ref. [75]). Histone chaperones play a crucial role in chromatin assembly but are also likely to assist in its disassembly ahead of the replication fork. To date, the most studied chaperones implicated in replication-coupled chromatin formation are antisilencing function 1 (ASF1) and chromatin assembly factor 1 (CAF1; for background, see Ref. [75]), and we next consider how these and other chaperones guide H3/4 around the replication fork. A model is emerging in which histones H3/4 displaced ahead of the CMG helicase are handed off between chaperones as they are passed back to the daughter strands. Exactly how the H3/4 tetramer is captured as it leaves the DNA remains incompletely understood, and a highly debated issue is whether it is split into two H3/4 dimers (Fig. 1c). The N terminus of MCM2, which, as a subunit of the hexameric helicase motor, is well placed to be the first point of contact between the replisome and the parental nucleosomes, interacts with H3 in such a way that it can bind either (H3/4)2 tetramers or H3/4 dimers [76–79]. ASF1 can also handle parental H3/4, but it only does so once the tetramer has split into two H3/4 dimers as it interacts

Please cite this article as: S. Šviković, J. E. Sale, The Effects of Replication Stress on S Phase Histone Management and Epigenetic Memory, J. Mol. Biol. (2016), http://dx.doi.org/10.1016/j.jmb.2016.11.011

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with the tetramerisation interface of the H3/4 dimer [80–82]. Thus, MCM2 and ASF1 may work cooperatively in capturing displaced parental H3/4. In supporting this, a complex between ASF1 and H3/4-MCM2, but not (H3/4)2-MCM2, has been observed leading to the suggestion that the ASF1–H3/4-MCM2 complex represents a transient state during the splitting of the H3/4 dimer [78,79]. The involvement of ASF1 in binding parental H3/4 creates a paradox. Since ASF1 binding to H3/4

requires the splitting of the (H3/4)2 tetramer (Fig. 1c), it might be expected to also allow the mixing of parental and newly synthesised H3/4 dimers. However, this is not observed, at least in the case of tetramers containing canonical H3.1/H3.2 rather than the variant H3.3 [46,83]. This suggests two possibilities. Tetramer splitting and parental dimer binding to ASF1 may be rare events during unperturbed replication with the intact (H3/4)2 tetramer being passed back directly for deposition on the nascent daughter DNA strands.

(a) Classical model of DNA replication in which parental (H3/4)2 tetramers are randomly segregated between the daughter strands

(b) A model of DNA replication in which Polε sits ahead of the MCM helicase. This may allow the Dpb3 & 4 subunits of Polε to contribute to (H3/4)2 eviction

(c) Constitutive ASF1 involvement would require splitting of parental (H3/4)2 tetramers with ASF1 handing the dimers off to CAF1 for reassembly and deposition

(d)

In conditions of replication stress, evicted (H3/4)2 is split and buffered by ASF1.

Key parental (H3/4)2 tetramer parental H3/4 dimer bound to ASF1

DNA Polymerase ε with Dpb3 & 4 in orange

CAF1

DNA polymerase δ

CMG helicase with MCM2 and its histone fold domain in pink

PCNA

Fig. 1. (legend on next page)

Please cite this article as: S. Šviković, J. E. Sale, The Effects of Replication Stress on S Phase Histone Management and Epigenetic Memory, J. Mol. Biol. (2016), http://dx.doi.org/10.1016/j.jmb.2016.11.011

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Alternatively, if ASF1 is routinely involved in the management of parental H3/4, then parental H3/4 dimers must be shepherded in a way that segregates them accurately from newly synthesised histone proteins. ASF1 function is intimately linked to, and indeed modulates, DNA unwinding by the replicative helicase [77], and its ability to ensure a steady supply of H3/4 to the newly synthesised daughter strands is essential for normal fork progression [84]. These results imply that ASF1 is required constitutively during replication. However, whether this is because it plays a crucial role in handling newly synthesised H3/4, parental H3/4, or both remains unclear. The best evidence that ASF1 can indeed buffer histones from both pools is seen under the conditions of replication stress induced by hydroxyurea (HU), an inhibitor of ribonucleotide reductase. In these conditions, impeded DNA synthesis leads to the activity of the replicative helicase becoming uncoupled from the polymerases, leading to the formation of excessive single-stranded DNA (ssDNA) ahead of the polymerases [85,86]. This results in ASF1 becoming loaded with H3/4 dimers that carry post-translational modifications characteristic of both newly synthesised and parental protein [87] (Fig. 1d). It is possible that discrimination between parental and newly synthesised H3/4 could be achieved through the differential marking of molecules before deposition. Recently, unmethylated H4K20 has been identified as a pre-deposition mark in mammalian cells. Significantly, this allows the binding of the histone chaperone TONSL. TONSL, which forms an obligate heterodimer with MMS22L, forms a complex not only with H3/4 but also with ASF1 and MCM2, demonstrating how an ASF1new H3/4 predeposition complex could be distinguished from a complex containing parental H3/4 [88]. Ultimately determining whether or not parental tetramer splitting and ASF1 binding occur during “normal” replication or only in certain contexts will be difficult as even during apparently unperturbed replication, coupling between the helicase and polymerase will be challenged as the polymerases encounter difficult-to-replicate sequences. However,

an ability to genetically separate ASF1-bound H3/4 pools could provide a firm basis for discriminating these two possibilities. We would not be surprised if the final answer is that the handling of parental H3/4 changes dynamically depending on the local conditions in the DNA and chromatin at the replication fork. Indeed, there is evidence that the frequency with which parental nucleosomes are split is locally increased by the presence of the histone H3.3 [46], which could affect the mechanism of mark propagation, as discussed further below. Chromatin assembly and maturation behind the fork Behind the fork, the histone chaperone CAF1 plays a crucial role in coupling DNA synthesis with H3/4 deposition [89,90] through its physical interaction with the DNA sliding clamp PCNA [65,91]. In budding yeast, either the loss of CAF1 [92,93] or the interaction of CAF1 with PCNA [65] results in defective maintenance of silenced chromatin structures at telomeres and at the mating type loci. In vertebrate cells, loss of CAF1 results in a catastrophic failure of chromatin assembly and the death of cycling but not quiescent cells [94,95]. In vitro, CAF1 binds H3–H4 dimers with similar affinity to cross-linked (H3–H4)2 tetramers [96]. However, whether CAF1 can handle intact parental (H3/4)2 tetramers is unclear. Indeed, if (H3/4)2 tetramers are transferred intact to the daughter DNA strands, then little is known about the chaperones involved in this presumably ASF1-independent process. Possible candidates include NAP1 [97] and the facilitates chromatin transcription (FACT) complex. FACT is able to bind H3/4 in addition to H2A/B [98] and several key components of the replisome including Pol α [99], the CMG helicase [100], and Replication Protein A (RPA) [101]. Furthermore, conditional deletion of the SSRP1 subunit of FACT in chicken DT40 cells causes a defect in replication fork progression [102].

Fig. 1. Models for handling parental H3/4 at the replication fork. Only parental H3/4 is shown for clarity. (a) Recycling of H3/4 without tetramer splitting. (H3/4)2 tetramers displaced ahead of the CMG helicase are bound by the histone chaperone domain of MCM2 and then passed intact to one of the nascent daughter DNA strands. The chaperone responsible for the installation of an intact H3/4 behind the fork in this pathway is unclear. (b) Recent work has suggested that Polε, widely thought to be responsible for the majority of leading strand synthesis [68], lies ahead of the CMG helicase [73,74]. Although the positions of the “histone mimic”’ Dpb3 and Dpb4 subunits could not be determined in the structure, this position of Polε could potentially allow it to facilitate the displacement of parental nucleosomes. This model implies an extended path of the single-stranded leading strand template around the helicase. While the exact path of the DNA is unknown, this single-stranded region (shown as a red dashed line) could provide an opportunity for the formation of DNA secondary structures during normal replication. (c) A model involving routine (H3/4)2 tetramer splitting by ASF1. This model would allow parental H3/4 dimers to be handled similarly to newly synthesised H3/4 by being handed off from ASF1 to CAF1 for assembly as a tetramer on the nascent DNA. However, there would have to be a mechanism to prevent the mixing of parental and new H3/4, as this is not generally observed. (d) When progression of the fork is impeded, illustrated here by reduced polymerase activity induced by, for instance, HU or aphidicolin, displaced parental H3/4 is split and buffered by ASF1 [87]. The fate of the modifications on this buffered ASF1 is unclear. While it has been proposed that the reinstallation of these marked histones at ectopic sites may result in epigenetic instability, our data suggest that when the uncoupling of the helicase and polymerase is extensive, parental marks are lost.

Please cite this article as: S. Šviković, J. E. Sale, The Effects of Replication Stress on S Phase Histone Management and Epigenetic Memory, J. Mol. Biol. (2016), http://dx.doi.org/10.1016/j.jmb.2016.11.011

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The fate of ASF1-bound H3/4 dimers is better understood. ASF1 transfers H3/4 dimers to CAF1 [80–82,103–106], likely as two H3–H4 dimers, rather than the tetrameric form [96,107] (Fig. 1c). The ability of CAF1 to bind H3–H4 dimers synchronously and promote the formation of (H3–H4)2 tetramers [96] suggests an appealing model in which ASF1 disrupts the parental (H3–H4)2 tetramer and then transfers the dimers to CAF1, which reassembles them as a tetramer prior to deposition on nascent DNA. However, if this pathway does handle both parental and new H3/4, it must do so without the significant mixing of old and new dimers, as noted above. Furthermore, a significant body of evidence from budding yeast suggests that CAF1 preferentially handles only newly synthesised H3/4 dimers acetylated on H4K5/12 [90,108]. Following (H3–H4)2 tetramer deposition, H2A–H2B dimers are added to form the full nucleosome. During the next 10–20 min, the chromatin begins to “mature”, a process in which the newly formed chromatin takes on the biochemical characteristics of non-replicating chromatin, such as reduced nuclease sensitivity [44,48,109]. Modification of the newly deposited histones with marks characterising the parental chromatin state also begins, although this process can extend well into and beyond the subsequent G1 phase [27,110].

Transmission of Parental Histone Marks to New Histones Following S Phase and the Propagation of Transcription Stability To complete the cycle of information transmission, the marks present on the parental histones must be copied onto the newly synthesised tetramers. Enzymatic systems for achieving this have been described for H3K9me3 and H3K27me3, histone modifications associated with the two major forms of heterochromatin [111–114]. In both cases, a mark “reader”, HP1 in the case of H3K9me2/3, or EED, in the case of H3K27me3, is physically associated with a mark “writer”, a histone methyltransferase (Suv3‐9 and EZH2, respectively), permitting the propagation of marks to adjacent histone molecules. This propagation mechanism can stably define a heterochromatic state over many generations [115]. The mechanisms by which histone modifications associated with active transcription are propagated are less clear, as is the case for such modifications actually contributing to the specification and mitotic stability of the transcriptionally active state. Evidence that histone marks are also able to carry information specifying an active transcriptional state is more controversial. The most extensively characterised H3/4 methylations associated with active transcription are H3K4me3, H3K36me3, and H3K79me3. Although H3K36me3 is strongly associated with transcription elongation and is enriched in the body of

transcribed genes, particularly towards the 3′ end [116], there is little evidence to support it being an epigenetic mark. Key to its maintenance in both yeast and humans is the methyltransferase Set2/SETD2, which is recruited directly by the Ser-2-phosphorylated C terminus of elongating RNAPII (reviewed in Ref. [117]). Furthermore, the mark is rapidly lost following the suppression of transcription [118], suggesting that it is laid down in consequence of transcription. It is thought that a principal function of this modification is to prevent spurious transcription initiation by RNAPII within the body of genes [119]. H3K79me3 is also enriched in transcribed genes and correlates with transcriptional activity [120,121]. In budding yeast, H3K79 trimethylation is catalysed by the methyltransferase DOT1, the activity of which is largely dependent on H2BK123 monoubiquitination, which in turn is localised to the transcription start site (TSS) and body of active genes (reviewed in Ref. [122]). The need for H2B ubiquitination for H3K79me3 appears less strict in mammalian than yeast cells with the recruitment of DOT1L also depending on the phosphorylation of the C terminus of actively transcribing RNAPII [123]. Although the function of H3K79me3 in transcription is not yet fully understood, there is little evidence to support the direct involvement of H3K79me3 in epigenetic memory. Nonetheless, very recent work suggests that this modification is essential to allow stable gene expression following the reactivation of expression by introduction of H3K4me3 [124], discussed further below. H3K4me3 is enriched around TSSs [125–128], and the level of this modification correlates approximately with transcriptional activity [128,129]. However, transcriptionally silent loci, often developmentally regulated, can also accumulate H3K4me3, frequently in association with H3K27me3. This has been suggested to represent a repressed state “poised” for expression [130–132]. In budding yeast, H3K4me3 is catalysed by Set1 [133–136], the catalytic subunit of the Complex Of Proteins Associated with Set1 (COMPASS) methyltransferase complex (reviewed in Ref. [137]). This is recruited by the elongating transcription complex through the Ser-5 phosphorylation of the C terminus of RNAPII [138,139]. As for H3K79me3, the H3K4 methyltransferase activity of Set1 is almost entirely dependent on the ubiquitination of H2B at K123 by the Bre1 ubiquitin ligase and the Rad6 ubiquitin-conjugating enzyme [140]. These mechanisms create a number of problems for models proposing H3K4me3 as a vector of epigenetic memory through replication and mitosis. The requirement for an H2B modification and the link to Ser-5 phosphorylation of RNAPII both suggest that H3K4me3 is installed as a consequence of transcription, correlating with rather than promoting it [18]. Furthermore, the argument that histone turnover during the cell cycle is likely to preclude the

Please cite this article as: S. Šviković, J. E. Sale, The Effects of Replication Stress on S Phase Histone Management and Epigenetic Memory, J. Mol. Biol. (2016), http://dx.doi.org/10.1016/j.jmb.2016.11.011

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transmission of epigenetic memory by histone marks [26] poses a particular difficulty for modifications like H3K4me3, which are found in sharply demarcated domains, a point we discuss further below. Nonetheless, several persuasive lines of evidence from other eukaryotic systems suggest that H3K4me3 may play an important role in the heritability of active transcriptional states. Treatment of cells with histone deacetylase inhibitors such as trichostatin A or valproic acid results in increased histone acetylation. That such induced changes can then be transmitted through cell division in the absence of the drug was first observed in the centromeric DNA of Schizosaccharomyces pombe [141,142]. Likewise, exposure of preimplantation mouse embryos to sodium valproate increases H4 acetylation and H3K4me3 at the Hoxb promoter, which was then passed on to daughter cells, although this heritable change in histone modifications was per se insufficient to induce the transcription of the locus [142]. A further example is seen in the inheritance of the memory of MyoD expression in Xenopus nuclei transplanted from somites. This epigenetic memory depends on K4me3 on the variant histone H3.3 and can persist for 24 cell divisions in non-muscle lineages of the transplanted embryos [143]. Direct evidence that the inheritance of H3K4me3 can be required for maintaining transcriptional activity comes from experiments in the slime mould Dictyostelium in which the transcription bursting of an individual locus, act5, was tracked by the direct visualisation of transcript production [144]. Replacement of the single H3 gene of Dictyostelium with an H3K4A mutant resulted in the loss of correlation in transcriptional activity of the act5 locus in daughter cells, suggesting that H3K4 methylation is required to maintain the probability of transcription firing, interestingly without determining that probability in the first place [144]. Very recently, the application of an “epigenetic editing” approach, tethering the H3K4 methyltransferase PDRM9 to specific genomic locations using either a catalytically dead Cas9 nuclease or zinc finger domains, has revealed that “writing” H3K4me3 can induce the expression of silenced genes, providing further evidence that this modification can actually specify an active transcriptional state in vertebrate cells [124]. So, if H3K4me3 can, at least in certain circumstances, act as an epigenetic mark of active transcription in some eukaryotes, then how is it propagated? While the stimulation of H3K4me3 by H2B ubiquitination appears to be conserved in mammalian cells [145,146], the mechanisms determining H3K4me3 also appear to be considerably more complex than in budding yeast. There are at least six homologues of yeast Set1 in mammalian cells (SETDB1A and B, and MLL1–4), which form distinct COMPASS complexes [137]. Within these mammalian COMPASS complexes, a number of components exhibit H3K4me3 recognition, including the PHD3 zinc finger of MLL1

[147–150], the WDR82 subunit of SET1A/B complexes [151], and ASH2, a subunit common to all COMPASS complexes [152]. Thus, within some COMPASS complexes, there exists the potential for coupling H3K4me3 recognition with the installation of the same mark by the catalytic methyltransferase of the complex in a manner analogous to that demonstrated for HP1/ SUV3‐9 and EED/EZH2 in propagating H3K9me3 and H3K27me3, respectively [111–114]. Alternatively, H3K4me3 segregation during replication at the promoters of transcribed genes could promote the reinforcement of the mark indirectly. One straightforward mechanism would be that the H3K4me3-marked parental H3 deposited following replication is sufficient to allow the reestablishment of transcription, which in turn would reinforce the modification through the transcription-coupled incorporation of H3K4me3 discussed above. Another possible mechanism arises from the experiments discussed above in which H3K4me3 is induced at the Hoxb promoter of early mouse embryos by treating them with the histone deacetylase inhibitor valproic acid [142]. In this model, H3 tail acetylation promotes H3K4me3 through the recruitment of the SET methyltransferase MLL4 [153]. In turn, H3K4me3 is recognised by the CHD1, a component of the SLIK and SAGA histone acetyltransferase complexes, which acetylates the tail of H3 [154], suggesting a cooperative model for the maintenance of these marks [142]. Accuracy of histone mark propagation and epigenetic memory All of these models raise an important question about the accuracy with which histone modifications must be propagated, relative to the underlying DNA sequence. How short domains of histone modification, of which H3K4me3 at promoters is an important example, might avoid the destabilising effects of stochastic fluctuation in the segregation of parental (H3/4)2 tetramers from the parental strands? In both budding yeast and mammalian cells, parental nucleosomes are deposited within 400 bp of their original position on both leading and lagging strands [38,41]. Nucleosomes are then repositioned and remodelled within minutes of their assembly [37] by a combination of nucleosome remodelling complexes and transcription [38,155–158]. During this “chromatin maturation” process, it is also likely that nucleosomes are repositioned to comply with constraints imposed by the underlying sequence or by DNA-binding proteins [159–161], which help promote similar nucleosome positioning to that seen in the parental epigenome. Nonetheless, the semiconservative nature of H3/4 partitioning between the daughter DNA strands has the statistical potential to cause locally asymmetric mark segregation through the exclusion of parental

Please cite this article as: S. Šviković, J. E. Sale, The Effects of Replication Stress on S Phase Histone Management and Epigenetic Memory, J. Mol. Biol. (2016), http://dx.doi.org/10.1016/j.jmb.2016.11.011

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Review: Replication Stress and Epigenetic Memory

histones. Large chromatin domains are likely to be robust to such events: modelling suggests that parental nucleosomes should be able to promote the reinstatement of a histone mark in a nucleosome array if they are deposited at a frequency of at least 1 in 3 [162,163], suggesting that there is a degree of leeway in the patterns of deposition that are nonetheless able to lead to the restoration of the chromatin state prior to replication. Furthermore, for the histone lysine methylations associated with gene repression (H3K9me3 and H3K27me3), newly incorporated H3 does not achieve the level of modification of the parental H3 even within a single cell cycle. Thus, it is clear that the inheritance of these modifications at single-nucleotide resolution is not required for the accurate inheritance of the repressed state [27,164]. In contrast, asymmetric segregation of parental nucleosomes does pose a problem for recyclingbased models of epigenetic memory for small domains [28], for instance, TSS-associated H3K4me3 [126,128]. In this regard, it is potentially important that there is evidence for the significant splitting of tetramers containing the variant histone H3.3 in contrast to canonical H3.1/H4 tetramers, which largely remain intact during replication [46]. H3.3 is enriched in transcribed genes, particularly around the TSS (reviewed in Ref. [165]). Thus, in these regions, the mixing of parental and new dimers could mitigate the potential for asymmetric tetramer segregation, thereby increasing the robustness with which H3K4me3 is maintained. As noted above, the inheritance of MyoD expression in transplanted Xenopus nuclei required K4 trimethylation specifically on H3.3 [143]. We next discuss experiments that directly test the histone recycling model by asking whether the propagation of epigenetically encoded transcriptional memory is interrupted by DNA replication impediments that break the coupling between DNA synthesis and histone recycling.

Sensitivity of Epigenetic States to Replication Impediments A clear prediction of the models we have discussed in which epigenetic memory is linked to histone recycling and propagation of a pattern of histone post-translational modifications is that the stability of gene expression states should be sensitive to impediments to replication fork progression. Fork progression can be impeded both by both local problems on the DNA template, such as damage or secondary structures, and by global factors that affect replication, such as nucleotide pool imbalance or mutations in key replication proteins [166]. Such issues are frequently grouped under the term “replication stress”, a rather imprecise phrase that can obscure important differences in the precise mechanism by which replication is impeded. We prefer to use the term to refer specifically

to situations, such as those just highlighted, that lead to the uncoupling of the replicative helicase and polymerases and the exposure of excessive ssDNA between them [167], and we discuss below instances in which this is likely to take place. Significant uncoupling of the helicase and polymerase would be expected to interrupt the flow of parental histones to the blocked daughter strand and thus also be expected to disrupt the flow of epigenetic information to that strand. Evidence that this can indeed happen and that replication stress can lead to stochastic changes in gene expression has emerged over the past 6 years. The first evidence came from our work in chicken DT40 cells lacking the specialised DNA polymerase REV1 [168,169]. Both DT40 and mouse cells lacking REV1 are unable to maintain normal replication fork velocities following DNA damage [170,171], suggesting that REV1 participates in a mode of DNA damage tolerance that acts at, or very close to, the replication fork and that minimises the effect that template DNA damage has on replication fork progression. In the absence of REV1, the replication of DNA lesions is channelled into a post-replicative gap-filling pathway dependent on the monoubiquitination of the DNA sliding clamp PCNA [170]. Crucially, this form of post-replicative bypass can take place remotely from the replication fork and, at least in budding yeast, in G2 phase of the cell cycle [172]. This suggested that the post-replicative DNA synthesis that ultimately fills the gap opposite of the lesion would be uncoupled from a supply of parental, marked histones leading to localised interruption in the flow of epigenetic information from the parental to daughter DNA strands. The interesting twist that made it possible to test this idea was that vertebrate REV1 turns out to be required not only for the replication of damaged DNA but also for the efficient replication of DNA secondary structures known as G quadruplexes (G4s) [168,173]. G4s (Fig. 2) form in single-stranded G-rich DNA as a result of the propensity of dG to form Hoogsteen base-paired G quartets [174]. G quartets can then stack to form a secondary structure, the G4 [175,176], which presents a potent block to DNA polymerases [177]. A general sequence for G4 motifs (Gx-Ny-Gx-Ny-Gx-Ny-Gx with N being any base, x is 3 to 5 bp, and y is 1 to 7 bp) has been proposed [178], but there is now good evidence of biologically significant G4s outside of this consensus [179], with estimates of over 700,000 potential sites in the human genome [180] being supported by a recently devised massive sequencing technique for detecting G4 formation in genomic DNA [181]. A key advantage of studying G4s rather than DNA damage as replication impediments is the ease with which potential sites of replication pausing or arrest can be identified and manipulated based simply on DNA sequence. Thus, in chicken DT40 cells lacking REV1, a G4 formed on the leading strand template can lead to

Please cite this article as: S. Šviković, J. E. Sale, The Effects of Replication Stress on S Phase Histone Management and Epigenetic Memory, J. Mol. Biol. (2016), http://dx.doi.org/10.1016/j.jmb.2016.11.011

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Review: Replication Stress and Epigenetic Memory

(a)

(b)

G4 motif G quartet

G4 quadruplex

Fig. 2. The G4. (a) Guanylic acid is able to form planar Hoogsteen base-paired rings, called G quartets, which are stabilised by a monovalent metal ion [174]. (b) Runs of dG in linear DNA can fold to form stacks of G quartets, a secondary structure known as a G4 [175,176].

localised and stochastic loss of parental histone modifications downstream relative to the direction of replication with associated loss of stability of the parental expression state [168,169,179]. We have developed a model in which this is explained by the delayed replication of the single-stranded region formed when the polymerase is stuck at a G4, while the helicase continues to unwind the parental duplex. The gap is replicated either by the release of the blocked fork or by a fork arriving from the other direction, resulting in a tract of biased new histone incorporation and the loss of the epigenetic information carried by parental histones, a chromatin “scar” [168,169,179] (Fig. 3). Interestingly, this mechanism can destabilise both repressed (H3K9me2/3-rich) and expressed (H3K4me3-rich) loci, resulting in derepression and loss of activation, respectively. This suggests that maintaining both transcriptional states requires epigenetic information that is sensitive to the disruption of replication and histone recycling. Analysis of affected loci reveals the loss of parental histone modifications around the TSS of these genes. For example, H3K9me2 is lost from the G4-harbouring, repressed ρ-globin locus, and H3K4me3 and H3K9/14ac are lost from the active CD72 and BU-1 loci of DT40 cells lacking REV1 [169]. In both cases, the loss of the parental marks can result in an expression state that is altered and, within a population of cells, also more diverse or “noisy” [169]. We speculate that this apparent increase in transcriptional noise, which is seen most clearly in the BU-1 locus following the loss

of H3K4me3 at the promoter [169], may reflect the proposal by Muramoto et al. that this modification is needed to ensure robust expression by maintaining the probability of transcription firing rates as cells divide [144], a hypothesis that merits further investigation. Genetic manipulation of the position of the G4 motif in the BU-1 locus of DT40 cells revealed that the “active” marks H3K4me3 and H3K9/14ac could be lost up to 4.5 kb downstream of the G4 (relative to the direction of replication) [179]. In the BU-1 locus, the window in which H3K4me3 is lost moves with the position of the G4 (Fig. 4), demonstrating that the maintenance of this mark is sensitive to the uncoupling of DNA synthesis and histone recycling. Interestingly, full expression of the gene is only lost if this sliding window reaches the promoter region, but not if it only affects marks within the body of the gene (Fig. 4). Aside from REV1 deficiency, which other situations lead to this form of epigenetic instability? The DT40 BU-1 locus has proved to be a very sensitive reporter that allows G4-dependent transcriptional instability to be monitored on a cell-by-cell basis [169,179]. It has revealed evidence of G4-dependent expression instability in mutants of three other helicases that have been previously implicated in G4 processing in vitro and in vivo FANCJ, WRN, and BLM [169] (Fig. 3f), supporting the idea that G4 persistence is likely to contribute to this effect. Supporting this idea, epigenetic instability of BU-1 can also be induced by stabilising G4 formation with specific ligands [182] (Fig. 3g).

Please cite this article as: S. Šviković, J. E. Sale, The Effects of Replication Stress on S Phase Histone Management and Epigenetic Memory, J. Mol. Biol. (2016), http://dx.doi.org/10.1016/j.jmb.2016.11.011

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Review: Replication Stress and Epigenetic Memory

An alternative approach to mitigating the exposure of ssDNA following replication arrest, and hence the effective degree of uncoupling of the helicase from DNA synthesis, is repriming. Priming involves de novo synthesis of either RNA or DNA on a single-

(a)

strand template, and until recently, the only known primase in vertebrates was PRIM1, the catalytic primase subunit of DNA polymerase α. Pol α is responsible for the initiation of leading strand synthesis and for the repeated priming of Okazaki fragments

ON Unperturbed replication

(b)

• structure resolved / ssDNA formation limited • histone recycling and DNA synthesis remain coupled • successful mark propagation

Leading strand polymerase paused at G4

• replication of DNA around the G4 is delayed

(e)

(i)

Loss of REV1

Nucleotide pool depletion

(h)

(f)

Loss of ‘close coupled’ repriming by PrimPol

Loss of G4 helicases e.g. FANCJ, WRN, BLM

(g)

G4 stabilisation

(c)

ON

(j)

• DNA synthesis and parental histone recycling uncoupled • biased incorporation of new histones lacking parental marks • formation of a ‘chromatin scar’

LOW

Mark propagation preseved

‘Neutral’ state transcription reduced and noisier H3K9 methylation / DNA methylation

(d) ON

(k) Level of transcription retained

OFF me

me

me

me

me

Locus heterochromatinised me

me

me

me

me

transcription shut off

Key

me

parental (H3/4)2 tetramer with ‘active’ marks

CMG helicase with MCM2 and its histone fold domain in pink

new (H3/4)2 tetramer

PCNA

(H3/4)2 tetramer with ‘repressive’ marks

Polymerase ε

new H3/4 dimer bound to ASF1

Polymerase δ

parental H3/4 dimer bound to ASF1

REV1

DNA methylation

G4 helicase G4 ligand

Fig. 3. (legend on next page)

Please cite this article as: S. Šviković, J. E. Sale, The Effects of Replication Stress on S Phase Histone Management and Epigenetic Memory, J. Mol. Biol. (2016), http://dx.doi.org/10.1016/j.jmb.2016.11.011

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Review: Replication Stress and Epigenetic Memory

G4 @ TSS +6.0 kb

H3K4me3

(a)

High Zone of interrupted histone recycling

G4 @ TSS +4.5 kb

H3K4me3

(b)

ori

High Zone of interrupted histone recycling

G4 @ TSS +3.5 kb

H3K4me3

(c)

Low Zone of interrupted histone recycling

1 kb

Fig. 4. G4 and replication-dependent erasure of H3K4me3 as a function of G4 position relative to the TSS of a model gene in REV1deficient cells. The first three exons of the BU-1 gene in chicken DT40 cells are shown as red boxes. A G4 motif is located in the second intron, orientated such that it will stall the leading strand of a replication fork entering from the 3′′’ end of the locus. (a) When the second intron is expanded to place this G4 6 kb from the TSS, the expression of the gene is as stable as in wild type and the distribution of H3K4me3 (and H3K9/14ac) is the same as in wild type with the characteristic enrichment at the promoter and at 5′′’ end of the body of the gene. (b) At 4.5 kb from the TSS, the G4 causes the loss of H3K4me3 within the body of the gene. In this situation, expression of the gene is stable. (c) At 3.5 kb from the TSS, H3K4me3 is also lost from the promoter region of the gene. Expression of BU-1 in these circumstances is unstable, with high level expression being lost stochastically as cells divide [179]. Removing, mutating, or inverting the G4 motif also stabilises the expression of the

locus. This experiment provides evidence that the maintenance of H3K4me3 requires processive replication and suggests that it is sensitive to the loss of histone recycling coupled to DNA replication [179].

needed for discontinuous lagging strand replication. Recently, a second vertebrate primase was described and named PrimPol, as it also possesses significant DNA polymerase activity [183–186]. PrimPol is a member of the archaeo-eukaryotic primase family and is capable of priming with both RNA and DNA. It is able to bind, but not replicate, G4 DNA, and importantly, this binding promotes repriming very

closely coupled to the structure, limiting the extent of ssDNA exposure following replication arrest [187]. This PrimPol-dependent repriming mechanism also prevents G4-dependent epigenetic instability [187] (Fig. 3h). Together, these observations all support a model in which a delay in replicating G4s, or failure to reprime beyond them, leads to localised epigenetic instability caused by the interruption of the recycling of

Fig. 3. Mechanisms promoting and suppressing replication-stress-induced epigenetic instability. (a) Unperturbed replication with the recycling of parental H3/4 and installation of new H3/4 coupled to DNA synthesis. (b) The leading strand polymerase stuck at a G4 structure. The helicase has run ahead, displacing nucleosomes that cannot be recycled. These (H3/4)2 tetramers are split and buffered by ASF1. (c and d) If the impediment can be rapidly negotiated, histone recycling is not interrupted and mark propagation can restore the parental chromatin state. (e–i) Mechanisms promoting replication-stress-induced epigenetic instability. (e) The Y-family polymerase REV1 contributes to G4 unwinding and facilitates prompt G4 processing at the fork, preventing helicase–polymerase uncoupling at the G4 [168,179]. (f) The specialised DNA helicases FANCJ, WRN, and BLM also contribute to prompt G4 destabilisation. FANCJ appears to be able to collaborate with REV1 and with WRN and BLM [168]. (g) Stabilisation of G4 structures can delay their replication and induce epigenetic instability [182]. (h) PrimPol limits the formation of ssDNA at G4 structures by initiating repriming closely coupled to the G4 to which it can bind [187]. (i) Global replication stress induced by HU or aphidicolin leads to widespread helicase–polymerase uncoupling and epigenetic instability that is exacerbated by the presence of a G4 [182]. (j) Extensive helicase–polymerase uncoupling disrupts histone recycling and marked H3/4, specifying that the parental transcriptional state is lost, either because the replication of the ssDNA containing the G4 is delayed or because it is replicated by a fork arriving from the other direction. The result is a tract of biased new histone incorporation, a “chromatin scar”. In the case illustrated, an actively expressed gene enters a “neutral” state, in which neither “active” nor “repressive” marks are present [21]. (k) In some cases, the neutral state can progress to heterochromatinisation with H3K9me3 and DNA methylation [182,187].

Please cite this article as: S. Šviković, J. E. Sale, The Effects of Replication Stress on S Phase Histone Management and Epigenetic Memory, J. Mol. Biol. (2016), http://dx.doi.org/10.1016/j.jmb.2016.11.011

12 parental histone marks. If this occurs in the vicinity of the regulatory region of a gene, it can, in turn, lead to transcriptional instability. The data show that the expression of at least a subset of loci in DT40 cells is sensitive to the loss of processive replication and suggest that normal expression homeostasis is dependent on parental histone recycling. The instances of epigenetic instability discussed above all result from a local replication impediment. What happens in the case of a global replication defect, such as that imposed by an imbalanced or depleted nucleotide supply? In such circumstances, the rate of DNA synthesis is reduced with evidence of uncoupling of the helicase and polymerase throughout the genome [188,189]. We modelled such global replication stress by treating cells with low dose HU, which slows replication by depleting nucleotide pools by inhibiting ribonucleotide reductase [190], or aphidicolin, which is a polymerase inhibitor, particularly of Pol α [191]. Reducing global fork rates by approximately 40% resulted in the destabilisation of BU-1 expression [182] (Fig. 3i). Significantly, this was largely, but not entirely, dependent on the presence of the G4 motif within the body of the gene. Additionally, there was a significant overlap in the G4-containing genes deregulated with HU and by the disruption of the FANCJ helicase, providing further evidence that a global replication defect is also able to induce epigenetic instability and that this effect is exacerbated and focused by the presence of DNA secondary structures [182]. Such a mechanism could explain at least some of the extensive epigenetic changes observed in cancer. Many cancer cells exhibit signs of replication stress, particularly in the early stages of their transformation, when replication stress can be induced by oncogene expression, which has been linked to genetic instability [192–194]. Supporting the idea that global replication stress during oncogene-mediated transformation can give rise to both genetic and epigenetic changes, De and Michor [195] have reported an association between G4 motifs and sites of abnormal DNA hypomethylation in cancers, sites that also correlate with the positions of somatic copy-number alterations, thus supporting a link between G4s and both genetic and epigenetic changes in cancer genomes.

Summary The propagation of epigenetic memory through histone recycling is an efficient approach to maintaining a transcriptional state. The use of such epigenetic information for maintaining heterochromatin is now well established, but an increasing body of evidence from studies in higher eukaryotes suggests that similar mechanisms may also confer robustness to active transcriptional states, which would not be achievable with the simple segregation of often limiting numbers of transcription factor molecules alone.

Review: Replication Stress and Epigenetic Memory

The precise molecular mechanisms that allow histone recycling to contribute to the stability of transcriptional states, and how cells protect these mechanisms from replication stress, require much further work. The ongoing elucidation of the structure of the eukaryotic replisome will be crucial to understand how histone recycling is coupled to DNA replication. A significant challenge will be to understand how the response of the replisome to local impediments dynamically influences the handling of parental histones and the transfer of their marks to the newly synthesised daughter strands. It is likely that further replisome components involved in maintaining processive replication and in the coupling of DNA unwinding and synthesis will emerge as contributions in maintaining epigenetic memory through replication.

Acknowledgments We would like to thank the members of the lab for helpful discussions. Work in the lab is funded by the Medical Research Council (U105178808). S.Š. is supported by an LMB International Scholarship and the Cambridge Commonwealth, European and International Trust. Received 19 August 2016; Received in revised form 10 November 2016; Accepted 11 November 2016 Available online xxxx Keywords: DNA replication; histone modifications; epigenetic memory; G quadruplex; replication stress Abbreviations used: ASF1, anti-silencing function 1; CAF1, chromatin assembly factor 1; HU, hydroxyurea; ssDNA, single-stranded DNA; FACT, facilitates chromatin transcription; TSS, transcription start site; G4, G quadruplex.

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Review: Replication Stress and Epigenetic Memory

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