Temporal and spatial regulation of eukaryotic DNA replication: from regulated initiation to genome-scale timing program.

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Review

Temporal and spatial regulation of eukaryotic DNA replication: From regulated initiation to genome-scale timing program Claire Renard-Guillet a , Yutaka Kanoh b , Katsuhiko Shirahige a , Hisao Masai b,∗ a Laboratory of Genome Structure and Function, Research Center for Epigenetic Disease, Institute of Molecular and Cellular Biosciences, The University of Tokyo, Tokyo 113-0032, Japan b Department of Genome Medicine, Tokyo Metropolitan Institute of Medical Science, Tokyo 156-8506, Japan

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Article history: Available online xxx Keywords: Replication origins Replication timing Pre-replicative complex Rif1 Chromatin loop

a b s t r a c t Replication origins are where pre-replication complexes are assembled during G1 phase. However, only a subset of the origins is actually “fired” to initiate DNA synthesis during S phase. Whereas factors involved in these steps are relatively well understood now, the mechanisms behind the origin specification, the choice of origins to be fired and determination of their timing are still under active investigation. Recent data show that the origin positions as well as the selection of those to be fired may be determined by multiple factors including sequences, chromatin context, epigenetic information, and some specific genomic features, but that the choice is surprisingly plastic and opportunistic. Timing regulation of firing, on the other hand, appears to be related to cell type-specific intrinsic chromatin architecture in nuclei. The conserved Rif1 protein appears to be a major global regulator of the genome-wide replication timing. Replication timing is regulated also by other factors including checkpoint signals, local chromatin structures, timing and quantity of pre-RC formation, and availability of limiting initiation factors. © 2014 Published by Elsevier Ltd.

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Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Replication start sites: origins of DNA replication . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1. Binding to specific DNA sequence . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2. Specific chromatin patterns . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.3. Specific genomic landscape . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.4. Evolution of origins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Replication timing of genomes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1. Establishment of replication timing in G1 phase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.2. Global genomic timing . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.3. Conservation of timing . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.4. Selection of origins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Regulation of origin firing time . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.1. Replication timing at centromeres and telomeres . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.2. Chromatin state . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.3. Genomic features and replication timing . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.4. Replication timing regulators . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.5. Firing regulation by competition for initiation factors loading . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Abbreviations: ARS, autonomously replicating sequence; ACS, ARS-consensus sequence; BAH, bromo-adjacent domain; CTR, constant timing region; CGI, CpG islands; NDR, nucleosome depleted region; ORC, origin recognition complex; OGRE, origin G-rich repeat element; pre-RC, pre-replicative complex; TSS, transcription start sites; TTR, timing transition regions. ∗ Corresponding author at: Department of Genome Medicine, Tokyo Metropolitan Institute of Medical Science, 2-1-6 Kamikitazawa, Setagaya-ku, Tokyo 156-8506, Japan. Tel.: +81 35316 3231; fax: +81 35316 3145. E-mail address: [email protected] (H. Masai). http://dx.doi.org/10.1016/j.semcdb.2014.04.014 1084-9521/© 2014 Published by Elsevier Ltd.

Please cite this article in press as: Renard-Guillet C, et al. Temporal and spatial regulation of eukaryotic DNA replication: From regulated initiation to genome-scale timing program. Semin Cell Dev Biol (2014), http://dx.doi.org/10.1016/j.semcdb.2014.04.014

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4.6. Timing regulation through 3D structure of chromatin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Concluding remarks . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Acknowledgments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

1. Introduction In both prokaryotes and eukaryotes, DNA replication generally initiates at various specified positions, called replication origins. In eukaryotes, the origins are distributed along the whole genome, ranging in number from several hundred to one thousand in yeasts to over 50,000 on the mouse and human genomes. For replication initiation, a highly regulated assembly and disassembly of protein complexes are required, as in many other chromosome transactions. It begins with the assembly of the Origin Recognition Complex (ORC) at potential origins, followed by the association of CDC6 and CDT1, which enables the recruitment of two MCM2∼7 complexes. This process occurs during G1 phase and generates the so-called pre-Replicative Complex (pre-RC). In the next step, origins are activated by the loading of CDC45 and GINS, which is facilitated by the Dbf4-dependent Cdc7 and Cyclindependent kinases, followed by the loading of the DNA polymerases and other accessory factors, leading to the initiation of DNA synthesis [1]. In contrast to the accumulating knowledge on factors and mechanisms of initiation, the nature of replication origins in higher eukaryotes and how their firing is regulated throughout the S phase have long been under debate. However, recent studies started to shed some novel insights into temporal and spatial regulation of origin firing and how it can be integrated into chromatin regulation. In this review, we would like to summarize the latest information on the long-standing questions regarding how the eukaryotic genomes are coordinately and timely replicated.

2. Replication start sites: origins of DNA replication 2.1. Binding to specific DNA sequence Studies on prokaryotic replicons have demonstrated that the replication initiation depends on specific sequences (reviewed in [2]). These studies naturally led to the search for specific sequences required for replication initiation in eukaryotes as well (Fig. 1). In the extensively studied model organism, Saccharomyces cerevisiae, autonomous replication of a plasmid was found to be driven by a 200-bp sequence, later called ARS for Autonomously Replicating Sequence [3,4]. ARSs contain a 11 to 17-bp consensus sequence (ACS), which was later found to be recognized by ORC. Moreover, B elements, located adjacent to ACS, facilitate the binding of ORC and ultimately the generation of pre-RC [5,6]. However, subsequent studies showed that not all ACS sequences are associated with origin activity [7,8]. Out of 1200 ACSs on the entire genome, about 400 of them were found to be associated with origins activity [9]. ACS, which can be similar or unsimilar to the consensus sequence of S. cerevisiae ARS, as well as to the B element for some species, are found in other yeasts, such as Lachancae or Kluyveromycces genus [10–12]. In the fission yeast Schyzosaccharomyces pombe, origins are generally extremely AT-rich [13–15]. This may be consistent with the presence of AT-hook motifs in SpOrc4 which are required for specific recognition of origins in fission yeast [16]. However, such sequence composition might not be a property of all the Schizosaccharomyces species, since origins from S. japonicus are rich in GC [14].

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No specific conserved sequences have been reported for origins in metazoan genomes. However, recently, the presence of G-rich motifs (Origin G-rich Repeat Elements, OGRE) was reported in the regions upstream to most of the origins that were identified in mouse and Drosophila melanogaster [17]. It was proposed that OGRE could form G-quadruplexes; an assumption consistent with a recent report that human replication origins are found widely associated with G-quadruplexes [18]. Thus, contrary to prokaryotic replication initiation, eukaryotes origins are not necessarily characterized by specific sequences; indeed, the purified human ORC shows sequence-non-specific DNA binding in vitro [19]. Even the budding yeast ORC exhibits nonspecific DNA binding under some conditions [20]. This might indicate that ORC may recognize specific DNA structure or conformation rather than sequence per-se. This speculation echoes with the finding that Drosophila ORC has higher affinity for negatively supercoiled DNA than for linear DNA [21]. Alternatively, chromatin structures, histone modification, transcription, and others may determine the ORC specificity (see below).

2.2. Specific chromatin patterns Early studies showed that, in S. cerevisiae, ORC binds to a region devoid of nucleosome [22,23]. Two elegant studies recently published have deepened our understanding of nucleosome positions at and around origins on a genome-wide scale. They showed the presence of asymmetric nucleosome depleted region (NDR) at these sites, flanked by well positioned nucleosome. DNA sequences surrounding these ACS were shown to be sufficient to induce a NDR because of specific sequence composition; B elements and ORC are necessary for a proper positioning of the flanking nucleosomes [24,25].

Fig. 1. Genomic characteristics of origins. Origins can be characterized by specific DNA sequence, nucleosome depletion, epigenetic marks, or specific genomic features (see Section 1 for details). Red bars, replication origins; yellow stars, fragile sites; gray vertical bars, TSS (transcription start sites); green box, CGI.


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In S. pombe, relationship between ORC positioning and nucleosome depletion may be less obvious. Some groups have found that origins are localized in NDR [14,26], and it was proposed that origins may be any sequences which ORC can bind to and are devoid of nucleosomes [14]. On the other hand, other groups failed to see any correlation between ORC binding and nucleosomes [15,27]. NDRs are associated with origins also in Lachancae waltii [12]. In metazoans, although origins and nucleosome data are less abundant, some evidence indicates that origins may co-localize with nucleosome-depleted segments. Association of origins with NDR is observed in D. melanogaster [28], and predicted probability of nucleosome binding decreased around the center of 283 human origins [29]. The genome-wide pre-RC locations in the Chinese hamster also correlated with NDR [30]. Thus, ORC binding regions are generally devoid of nucleosome in almost all the species. However, a study in yeast showed that the interaction of ORC with surrounding nucleosomes is required for proper origin firing [31]. The Orc1 bromo-adjacent domain (BAH), a chromatin-binding module, is necessary for the strong binding of ORC to some origins in vivo [32]. A recent study, using in vitro assays and atomic force microscopy, showed that S. cerevisiae ORC directly interacts with DNA in a rather sequence-non-specific manner, and that this binding is stabilized through interaction with the surrounding nucleosomes [20]. However, this interaction is not dependent on ORC BAH, indicating that BAH may not be required for interaction with core histones in S. cerevisiae (but might be required for interaction with modified histones). Thus, these studies led to a model that ORC binds to NDR, and further interacts with flanking nucleosomes to stabilize its binding and reposition the nucleosomes at the same time. In metazoans, ORC might interact with surrounding chromatin, or chromatin with a special epigenetic state, since human ORC1 BAH was shown to recognize specifically histone H4 with di-methylation on lysine 20 (H4K20me2) [33]. However, so far, there is no report showing re-positioning of flanking nucleosomes or stabilization of ORC binding via interaction with surrounding chromatin in metazoan. Besides the nucleosome positioning, specific modifications of the surrounding chromatin may be determinants of origin specification. Interactions of ORC and other pre-RC components with specific histone signature and histone modifiers may facilitate the recruitment of the pre-RC in specific contexts (reviewed in [34]). Furthermore, methylation status of CpG may also regulate the position of replication initiation [35]. 2.3. Specific genomic landscape In spite of lack of significant sequence specificity of ORC binding, origins do not appear to be randomly located on genomes. Indeed, in both S. cerevisiae and S. pombe, the genomes of which are compact and mostly made of ORF, origins are strongly biased toward intergenic regions [9,27,36]. Moreover, genome comparative studies have shown that, in S. cerevisiae, origins are significantly associated with fragile sites of the genomes, i.e. the sites prone to get broken during the course of genome rearrangement [37,38], but it is not the case in L. kluyveri and L. lactii [39]. In S. pombe as well, some origins were shown to associate with loci prone to undergo doublestrand breaks [40]. However, the mechanisms linking origins to such fragile sites and whether there exist any cause to consequence relationships are still unclear. Although organization of origins in mammalian genomes is still quite far from being elucidated, several recent genome-wide studies revealed some features potentially associated with mammalian replication origins [17,41–45]. Interestingly, all these analyses report association of origins with promoter regions and/or active transcription, though the datasets are only very partially overlapping [18,46]. These datasets rely on enrichment of nascent

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single-stranded DNA or replication bubble trap, and thus most likely represent early or efficiently firing origins. Mapping of ORC or MCM binding sites would provide genomic view on the potential replication origins; i.e. the locations of pre-RC assembly sites. However, the sizes of mammalian genomes have made it technically challenging to precisely determine the genome-wide binding sites of pre-RC. Only very recently, a group managed to map ORC1 binding sites on the human genome using chromatin fractions enriched in pre-RC bound DNA [47]. ORC1 binding sites mapped in this study were generally associated with transcription start sites (TSSs) of coding or non-coding RNAs (ncRNAs). The authors proposed the existence of two classes of origins; one that is mapped close to the TSSs of coding RNAs, associated with moderate/high transcriptional activity and fires in early S and the other that is mapped close to TSSs of ncRNAs, associated with low transcription activity, and fires throughout the entire S phase. However, it should be noted that regions close to (some) highly expressed genes were found to be ‘hyper ChIP-able’ in budding yeast, suggesting that signals are often found at these locations without any biological significance [48]. In light of these observations, strong association of ORC and TSS in metazoans may need to be viewed cautiously. Nevertheless, these findings cause the revisit of argument over whether replication is associated with active transcription. Although some origins are associated with active transcription, this is not the case for all the origins and transcription level is not necessarily high at some origins. For instance, it was proposed that replication initiation events may be associated with moderately transcribed regions but not with highly transcribed regions [35]. Active transcription could be a feature of early origins [47], but other origins can be present in regions devoid of active transcription. A relationship between CpG islands (CGIs) and origins active in germ-line was proposed fifteen years ago based on a limited dataset [49]. Recent large-scale analyses of replication origins have revealed significant overlap between CGIs and origins in various cell lines of human and mouse [17,41,42]. Moreover, putative human replication origins, predicted in silico and assumed to represent highly active origins in germ-line or in early embryos, strongly co-localized with CGIs [2,50]. Finally, the presence of the G-Rich motif, OGRE, is highly coincident with CGIs in mouse embryonic stem cells. CGI-associated OGREs are often flanked by two origins, while most of other OGREs are associated with one origin [17,51]. However, it is still an open question whether CGIs are markers of origins active in germ-line and early embryonic cells. On the other hand, in deep sequencing of short-nascent strands in four human cell lines which significantly increased the number of detected origins, less than half of them are associated with CGIs, TSS and/or gene body [18], although annotations are different from those used in reference [47]. Thus, recent genome-wide analyses clearly indicated a bias of origins toward genes, promoters and/or CGIs in metazoan genomes. However, mechanistic insight into the roles of these sequences in activation of origins is lacking. It would be an interesting possibility that origins may be recognized through multiple mechanisms and those preferentially utilized in germ-line or undifferentiated cells may be distinct from those preferentially utilized in normal, differentiated cells. 2.4. Evolution of origins Recent comparative analyses of replication origins in yeasts indicated that the sequence and structures of origins as well as the dynamics (replication timing) of chromosome replication are well conserved in different Saccharomyces species [52] and even between Lachancea waltii and S. cerevisiae that are




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separated by about 150 million year divergence [12]. However, genomic locations of origins are generally not conserved except for some early origins [12,52]. These works suggest that, in spite of prevalence of global replication timing regulation, genomic locations of replication origins are not under strong evolutionary constraints, consistent with fairly high level of the plasticity and stochasticity in origin usage for completion of replication of the entire genome. Nevertheless, the positions of origins have left prints in the genome over time. Indeed, a strong skew in sequence (i.e. change in relative proportion of G toward C or A toward T) was observed at replication start point in a large part of bacterial genomes [53,54]. This was ascribed to differential mutation mechanisms occurring on the leading and the lagging strands. In eukaryotes, such bias would be more difficult to identify since replication initiates at multiple points. The presence of a replication-related bias in S. cerevisiae and K. lactis genomes, though weaker than in bacteria, was reported, but could not be identified in S. pombe [55]. In metazoans, the search for similar bias is highly complicated by the fact that that only the consequence of mutational biases in germ lines and early embryonic cells can be observed (for transmission to offspring). Cayrou et al. observed some bias around the origins they detected in mouse embryonic cells, but suggested later that it was a mere consequence of the presence of OGRE sequences [17]. Additionally, Nesculea et al. failed to detect bias around origins identified in HeLa cells [41,56]. However, another group, using sophisticated mathematical tools, emphasized the presence in the human genome of N-shaped jumps of cumulative GC and AT skew (later called Ndomains). They suggested that these N-domain-associated origins correspond to very active and efficient origins in germ line [2,57].

Fig. 2. Two models for describing the alternating early and late replication domains in mammalian genomes. The first model (upper) proposes that two constant replication timing regions (CTRs) with different timing (early and late) are separated by transition regions (TTRs) which are replicated with a single replication fork, emanating from the earlier-replicating domain. Another model (lower) proposes that replication timing forms ‘U-domains’; two early ‘edges’ are separated from the ‘center’, a mid- or late-replicating domain, by a transition region which is replicated by ‘domino-cascade’ activation of multiple origins. The vertical black arrow represents the time transition, from early S phase (top) to late S phase (bottom).

3.2. Global genomic timing 3. Replication timing of genomes Firing of single origins in S phase can be monitored by various methods such as two-dimensional gel electrophoresis, dense isotope substitution or DNA combing (reviewed in [58,59]). Such studies highlighted that only a subset of the origins fire at each cell cycle, at various time throughout the S phase. Origin firing can thus be described with two parameters, efficiency and timing [58,60]. The efficiency of an origin refers to the proportion of the cells within the population in which a given origin is activated. On the other hand, origin timing refers to the time within the S phase at which a given origin is duplicated, either by initiation or passive replication. However, in the following sections, we will mainly focus on firing timing rather than replication timing. The origins firing at the beginning of S phase will be referred to as early-firing origins (or early origins), and those firing late S phase as late-firing origins (or late origins).

3.1. Establishment of replication timing in G1 phase Although origin firing occurs only during the S phase, the firing timing appeared to be determined well ahead of S phase. Indeed, using a site-specific recombination, the ability of a telomere to enforce late replication onto a neighboring origin was shown to be established between mitosis and START in the subsequent G1 phase in budding yeast [61]. This idea was further validated by the identification of a timing decision point (TDP) in early G1, which determines the timing of replication in mammalian cells [62]. TDP may be coincident with the chromatin repositioning event [62,63], indicating a possibility that the replication timing of origins is regulated, at least partially, by some factors that may reset the chromatin architecture at early G1 to subdivide the chromosomes into differentially replicating domains.

Recent genomic techniques (ChIP-on-chip and ChIP-seq) coupled with count of DNA copy numbers or BrdU incorporation enabled a visualization of genome-wide profiles of replication timing ([64] and ref. therein). In yeasts, this profile is globally correlated with average origin activity: regions around early-firing origins are replicated early and those around late-firing (or inefficient) origins late. On the other hand, in mammalian genomes, replication profiles revealed the presence of Mb-scale regions with similar timing, called replication domains or constant–timing regions (CTRs; sizes from 0.5 to 2 Mb), separated by so-called timing transition regions (TTR; sizes from 0.1 to 0.6 Mb; reviewed in [2,64]). Comparisons of replication timing in different mammalian cell lines have revealed that although some replication domains are constitutively early (or late), the timing of around half of the genome can change between cell types [64–68]. Replication domains in embryonic stem cells display a specific timing profile with shorter domains (400–800 kb), but these domains merge into larger domains during differentiation [65]. These smaller domains were proposed to correspond to ‘units of replication’ which may be independently regulated and consecutive units with similar timing may form a CTR [69–71]. How TTRs are formed is also a debated question and two different, but non mutually-exclusive models have been proposed. One view is that a TTR, located between an early and a late CTR, is replicated unidirectionally by a fork coming from the early domain (Fig. 2). However, the replication rate of TTRs is not always compatible with a replication by a single fork. Thus, Guilbaud et al. proposed another model in which around a half of the genome may be organized in segments with Ushaped replication timing, sizing in average from 1 to 1.6 Mb. They were called U-domains ([72,73], reviewed in [2]), some of which overlap with CTRs/TTRs. These U-domains are proposed to be bordered by two early peaks, and the transition




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between the earlier-replicating edges and the later-replicating center would be generated by a ‘domino-cascade’ activation of several origins [72] (Fig. 2). In this model, although more than one origins are involved, the replication is also almost unidirectional, because a ‘up-bound’ fork from a newly fired origin is almost immediately merging with a fork coming from the upper origin, while a ‘down-bound fork’ will travel longer [2]. Interestingly, the previously mentioned N domains, proposed on the basis of skew observations, are partly overlapping with the U domains detected.

4. Regulation of origin firing time

3.3. Conservation of timing

4.1. Replication timing at centromeres and telomeres

Recent genomic tools also enabled comparison of replication profiles between closely related species. These analyses revealed that the position and the activity of early origins are conserved amongst Saccharomyces species, while dormant origins are poorly conserved; leading to conservation of global replication profile amongst these species [52]. Similarly, comparison of replication timing between human and mouse showed strong conservation of replication timing domains in similar cell lines [67,68]. All these data thus suggest that replication timing is at least partially retained over evolution, although whether conservation of replication timing itself is under selective pressure or it is an indirect consequence of other evolutionary constraint is still an open question. The physiological significance of having replication timing regulation is not understood at the moment, but several hypotheses have been proposed. Those include protection of early-replicating regions from mutation (mutation rate is known to be higher in late replicating regions), genes dosage effect of early-replicating regions, or avoiding too much fork collision due to over-initiation [64].

Specific replication patterns at two special regions of the chromosomes, centromeres and telomeres, should be noted. Centromeres, in spite of their heterochromatic nature, were found to replicate early in both S. cerevisiae and S. pombe (reviewed in [77]). In budding yeasts, the presence of a centromere itself was shown to promote early firing of neighboring origins, albeit on a local scale (∼20 kb) [78,79], probably via accumulation of DDK at kinetochores which promotes the prompt recruitment of Sld3-Sld7 [80]. The mechanism for early firing of S. pombe centromeres is different, and appears to involve the recruitment of Dfp1 (Dbf4) to centromere via the heterochromatic protein Swi6 (HP1) [81]. Telomeres, on the other hand, were found to fire late. In S. cerevisiae, this feature was shown to be governed by the silencing protein Sir3 [82] and the Ku complex, which appears to regulate the replication timing by affecting the length of telomeres [83]. A short telomere is replicated abnormally early [84,85] and this effect is seen over ∼80 kb beyond the affected telomere. Thus, segments surrounding the centromeres are replicated early, whereas those near the telomeres are replicated late.

More than twenty years ago, the firing timing of an early origin was shown to be delayed when moved close to a late-firing origin, and conversely the firing timing of a late origin was advanced when moved onto a plasmid [75]. These pioneering experiments highlighted the importance of chromosomal position and context in determination of the origin firing time and suggested the presence of a cis-acting element that affects the firing timing of a nearby origin. Subsequently, small fragments of chrXIV were shown to be capable of delaying the firing time of adjacent origins [76].

4.2. Chromatin state 3.4. Selection of origins Not all individual origins fire at each cell cycle; the question of how the ones that fire are selected has long been debated. In between a totally deterministic replication program (each origin fire sequentially in an immutable order) and a totally stochastic one (each origin has the same probability of firing), data led to a ‘controlled stochastic’ model: each origin fires stochastically, but not with the same probability of firing, due to genomic context and global factors [58]. As explained above, global, genome-wide replication timing is established in G1, and firing is executed during S phase at individual origins, each of them firing stochastically but with a different firing probability [64]. Several models for this stochastic firing have been proposed. Firing of a single origin, while regulated by genomic context, may be totally or largely independent of neighboring origins, as proposed and reviewed [74]. On the other hand, another group showed that some mouse replication data were more consistent with a model of ‘flexible replicon’, in which 4–5 origins are organized into a replicon cluster, amongst which only one origin fires stochastically [17,34]. Moreover, as mentioned earlier, replication data in several genomic segments in human cells are compatible with ‘domino-cascade’ activation of origins (i.e. the firing of one origin increases the firing probability of the neighboring one [72]). Thus, the firing timing for a single origin might be predicted on the basis of combined effects of its intrinsic probability of firing and its chromosomal context, which may be related to the global regulation of genome-wide replication program. Recent studies have provided some insight into the mechanisms by which the probability of origin firing is controlled.

Chromatin state around origins can influence their firing time by affecting either the assembly of pre-RC during G1 phase or the recruitment of initiation factors during S phase. Indeed, recent reports indicate that both assembly of pre-RC itself and its activation step can be determinants of the firing time. As stated earlier, the interaction of ORC1 bromo-adjacent homology (BAH) domain with specific nucleosome structures was shown to be necessary for efficient origin firing [32]. Furthermore, two classes of origins were identified by comparing the strength of DNA-ORC interaction in vitro (with naked DNA) and in vivo (in the cellular context; [86]). Origins exhibiting similar binding strength in vitro and in vivo and those showing higher affinity in vivo than in vitro were defined as ‘DNA-dependent’ origins and ‘chromatin-dependent’ origins, respectively (here, ‘chromatin’ indicates DNA associated with histones or with any other factors). The interaction between ORC and ‘chromatin’ seems to be independent of the BAH domain, suggesting that other domains of ORC might contribute to its interaction with surrounding nucleosomes or factors. Unexpectedly, DNAdependent origins were found to be late firing, and the authors suggested that a stronger interaction between ORC and DNA may be inhibitory for recruitment of MCM helicases and other factors. Conversely, a weaker but more dynamic interaction of ORC with surrounding chromatin context would allow efficient pre-RC assembly and possibly loading of multiple MCMs. This hypothesis relies on the in vitro observations that multiple MCM complexes can be loaded at a single origin, which may potentially increase their ability to recruit initiation factors and allow origins to fire earlier and/or more efficiently [87,88]. However, no correlations between the amount of ORC or MCM complex loaded at each origin and its activity to fire was reported in a recent study [89], leaving it open


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whether the amount of pre-RC loaded onto an origin influences its firing timing or efficiency. Furthermore, several epigenetic modifications were found associated with the firing timing of neighboring origins. Mainly, histone acetylation was found to be strongly correlated with origin firing activity in S. cerevisiae. It can affect either pre-RC establishment or recruitment of initiation factors or both. Indeed, the activity of some origins may be inhibited by unfavorable nucleosome positioning via a mechanism dependent on the histone deacetylase Sir2p [90]. On the other hand, deletion of Rpd3, another histone deacetylase, caused late origins to fire early, presumably due to increased histone acetylation [91–93]. Tethering the histone acetyltranferase Gcn5 near a late origin advanced its initiation timing [91]. However, another histone deacetylase complex (Hst1-Sum1-Rfm1) was shown to promote initiation at several origins [94], suggesting that the role of histone acetylation in origin firing regulation could be more complex than previously thought. Other histone marks were also found to be associated with origin activation in budding yeast: di-methylation of lysine 4 of histone 3 (H3K4me2) with proper recruitment of pre-RC [95] and monomethylation of lysine 36 of histone 3 (H3K36me) with recruitment of Cdc45 [96]. Mammalian replication domains are also associated with specific chromatin states: early CTRs tend to be enriched in regions with open chromatin marks and late CTRs in those with close chromatin marks [64]. Similarly, U-domain borders (early replicating) are also associated with open chromatin marks [2]. Recent analyses indicated that epigenetic marks can be clustered into four states, ranging from open and transcriptionally active chromatin to heterochromatin, which are associated with different average replication time (from early to late, respectively). These states correlated nicely with the arrangements of U-domains (from the open and early-replicating U-border to the closed and latereplicating central area) as well as with the profiles of CTRs [97]. 4.3. Genomic features and replication timing In addition to chromatin states, several genomic features were shown to be correlated with replication timing domains in mammalian genomes. Gene-rich regions with high GC-content and low LINE density are associated with constitutively early CTRs, while regions with opposite characteristics are associated with constitutively late CTRs; and replication domains whose timing varies amongst cell lines have usually intermediate features [64]. Although a general correlation between active transcription or silenced transcription with early or late replication, respectively, has been observed, whether transcription (and more generally chromatin structure) regulates replication timing or vice versa has long been debated [98]. However, the recent genomic data indicated that genes in the early-replicating domains are transcribed with relatively constant probability, and that a correlation between fractions of transcribed genes or expression level and replication timing can be observed only in the mid- or late-replicating domains [64,99]. Thus, transcription is also more likely to be indirectly associated with replication timing rather than having a direct cause-consequence relationship. Similarly, in yeasts, available evidence shows no relationship between transcription and timing regulation [77]. 4.4. Replication timing regulators Apart from chromatin states, other factors were shown to bind to or near origins and affect their firing timing. It has been known that, in S. cerevisiae, late origin firing is under S phase checkpoint regulation. Replicative stress such as HU or MMS can suppress the

firing of late origins in a manner dependent on mec1 or rad53, key regulators of the DNA replication checkpoint [100–104]. Suppression of firing of late origins through intra-S checkpoint was shown to operate also in human cells [105]. Recently, the transcription factors Fkh1 and Fkh2 (Fkh1/2) were found to regulate the firing timing of a large fractions of the origins in S. cerevisiae. They were shown to bind near early-firing origins, and to tether them to facilitate their firing by increasing the local concentration of initiators [77,106]. It was also shown that B3 and B4 elements in S. cerevisiae are needed for timing regulation at some origins [107,108]. However, later analysis suggested that the Fkh1/2 binding sites may be responsible for this regulation [109]. Furthermore, some origins fire constitutively early while the chromatin context affects the firing timing of others. The constitutively early-firing origins are generally associated with two Fkh1/2 binding sites and a constitutive NDR [109]. A recent study identified another sequence required for the early activation of ARS1 (in S. cerevisiae), although the factor(s) involved and mechanisms are still unclear [110]. Early firing origins might also be marked by other factors such as Mrc1, as suggested in S. pombe [111]. On the other hand, factors that delay or suppress the firing of origins have also been reported. The Rif1 protein was identified as a timing regulator in S. pombe on the basis of a screening for mutants rescuing the hsk1 (Cdc7) deletion. Rif1 deletion causes many late origins in both sub-telomeric and arm regions to fire earlier [112]. Interestingly, some early origins fire later in rif1 cells. Thus, Rif1 may regulate origin firing both positively and negatively. Alternatively, hyper-firing at dormant origins may titrate out the limiting initiation factors available to those origins normally fired early. Rif1 binding sites, mostly located in intergenic regions, are distributed along the S. pombe genome, but around half of them are close to late origins that are deregulated in the rif1-deletion mutant. Interestingly, a conserved motif was discovered in most of those Rif1 binding sites, that is similar to a sequence previously reported to regulate the late firing of ARS727, the so-called late-consensus sequence (LCS; [113]). It would be of interest to determine whether chromatin binding of Rif1 depends on these conserved sequences. Rif1 binding sites do not precisely coincide with origins, and the number of Rif1 binding sites (about 80 excluding the telomere) are much lower than that of origins affected by its deletion (about 400 including both upregulated and downregulated ones), suggesting that a single bound Rif1 protein might affect the firing of more than one origins. Another telomere factor, Taz1, was reported to suppress some dormant origins in S. pombe [114]. It was shown to bind to the telomeric repeat motifs present near the affected origins. Since all origins regulated by Taz1 are regulated by Rif1 (but not vice versa), some late/dormant origins may need Taz1 to be regulated by Rif1.

4.5. Firing regulation by competition for initiation factors loading It has been shown that the initiation factors are present in a limited amount and the competition for these limiting factors is responsible for replication timing regulation. Indeed, overexpression of Cdc7-Dbf4, Cdc45 or other initiation proteins were shown to activate dormant origins, first in S. pombe [115,116]. Later studies in S. cerevisiae showed more clearly that overexpression of CDK, DDK and its substrates, Sld3 and Sld2 [117], or that of Sld3-Sld7-Cdc45 led to early firing of late/dormant origins [118]. Similar observations were made in mammalian cells [119]. Interestingly, increase in the number of firing origins due to the overexpression of these initiation factors strongly depletes dNTP pools [117], suggesting that this regulatory process may contribute to adjusting the fork number to the dNTP pools, avoiding problems during the replication process.




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Fig. 3. Combination of different mechanisms ensuring ordered firing of replication origins in yeasts. ORC binding at early origins is relatively weak, and ORC interact with ‘chromatin’ (nucleosomes or other factors) to recruit MCM efficiently, possibly loading multiple MCM. Furthermore, acetylation of surrounding nucleosomes (stars) and the binding of Fkh1/2 factors permits efficient recruitment of initiation factors, present in a limiting amount (represented by Cdc45 shown as red circles with “45”). On the other hand, the strong binding of ORC at late origins may delay MCM loading, and Rif1 and condensed heterochromatin also delays the loading of initiation factors to the nearby origins. Rif1 binds to chromatin at early G1 near mid- or late-firing origins, and suppresses their firing until mid-S when the inhibition is somehow relieved, as indicated by the change of the color for Rif1. Although Rif1 is shown as being released from the chromatin after mid-S, this is not yet experimentally confirmed. Rif1 has been shown to dissociate from mitotic chromosomes. Throughout S phase, the limiting initiation factors may become available to mid- or late-firing origins, only after they are released from the replisome in which they have been engaged.

4.6. Timing regulation through 3D structure of chromatin Increasing evidence shows that nuclear organization and subnuclear localization of origins influence their firing time as well. Heun et al. showed that in budding yeast late-firing origins are enriched close to the nuclear periphery in G1 (but not necessarily in S phase), while early-firing origins are dispersed within the nucleus [120]. They proposed that late-firing origins are targeted to nuclear membrane via the sequences distinct from origins but present nearby, where some chromatin modification may occur. However, the nuclear periphery localization alone may not be sufficient to define late replication, since artificial tethering of an early replication origin to the nuclear membrane in S. cerevisiae did not cause late replication of this origin [121]. On the other hand, 4C analyses have shown that early origins tend to contact with each other in S. cerevisiae [122]. This sort of 3D clustering of early origins could facilitate early firing and may be mediated by factors such as Fkh1/2 which can form a dimer [77]. A current model for replication timing regulation is presented in Fig. 3. The chromatin architecture in nucleus that may be determined in early G1 (generation of specific chromatin loop structures and tethering of late replicating segments at nuclear periphery) as well as local chromatin structures (histone modification, binding of transcription factors etc.) would determine the global timing of DNA replication and firing efficiency and timing of each origin. These regulatory mechanisms may facilitate the creation of a local gradient in the concentration of limiting factors, including Cdc45, and coordinate the firing of origins throughout the entire S phase.

Similarly, replication timing in mammalian cells appears to be regulated by subnuclear organization and 3D structure of DNA (Fig. 4). Indeed, genomic profiles of CTRs are best correlated with 3D structures of DNA as assessed by Hi-C experiment ([64,67], unpublished data from Pope and Gilbert). Moreover, a strong association between lamina nuclear domains [123], attached to nuclear membrane, and replicated regions was shown [66]. Additionally, 4C experiments showed frequent interaction between early replicating domains even derived from distant chromosome segments [124]; and genomic rearrangements between mouse and human were shown to occur more frequently between two early replicating domains [68]. This suggests that early replicating domains are clustered together inside the nucleus. On the other hand, borders of replication domains correlate with ‘insulator marks’ such as CTCF binding sites [69]. All these observations suggest that replication domains with similar timing may be spatially clustered, with early replicating domains spreading throughout the whole nucleus, while mid- and late-replicating ones attached to nucleus membrane or targeted to heterochromatin foci. At the onset of S phase, limiting initiation factors would be readily accessible to early CTR but not to mid or late CTR. As S phase proceeds, the latter domains may undergo conformational change so that the initiation factors are now accessible to mid- or late-replicating domains (Fig. 4). Although molecular mechanisms linking 3D organization of DNA inside the nucleus and replication timing are still not clear, recent analyses suggested that Rif1 may play a key role in this regulation. Human Rif1 was found to be localized at DNaseI-insoluble


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Fig. 4. Models for 3D organization of replication timing domains in mammals. Early replicating domains are associated with open chromatin and spread throughout the interior of the whole nucleus, allowing the limiting initiation factors to easily get access to them. They may be bordered by ‘insulators’, specific chromatin marks or binding segment of a specific factor (e.g. CTCF). On the other hand, mid- or late-firing domains are associated with nuclear lamina at nuclear periphery or heterochromatin regions, respectively.

nuclear periphery as well as at the nucleoli boundary, where mid-S replication foci are observed, and mid-S replication foci are specifically lost when Rif1 is depleted in human cells. Thus, human Rif1 may tether the mid-S replication origins at particular nuclear compartments and this will suppress them until mid-S phase by generating a special chromatin compartments that are refractory for the actions of initiation factors (Fig. 5). It is tempting to assume that Rif1 plays a similar role in S. pombe, where late origins are present near nuclear periphery. Indeed, it was reported that budding yeast Rif1 is associated with nuclear membrane through

Fig. 5. Two non-exclusive models for actions of Rif1 in regulation of replication program. (A) Through DNA looping, Rif1 can confine clusters of origins and sequester them from the action of initiation factor, preventing their firing until mid-S when this chromatin configuration is somehow disrupted. (B) Rif1 can recruit PP1 (phosphatase) which in turn can inhibit the firing of surrounding origins by counteracting the phosphorylation events catalyzed by Cdc7. Red bars, replication origins.

C-terminal lipid modification [125]. Recently, Rif1 in budding yeast was reported to recruit PPase1 to counteract the Cdc7-mediated phosphorylation of MCM components in pre-RC, thus inhibiting the initiation [126] (Fig. 5). This mechanism may not be mutually exclusive with the “compartment model” above. 5. Concluding remarks The specification of ORC binding can be affected by many factors including specific sequences, transcription, nucleosome depletion, epigenetic traits, and other yet-to-know genomic features. There may be different modes of origin recognition; for instance, in the AT-rich genome of S. pombe, ORC recognizes the origins generally present in the intergenic AT-rich regions through the AT-hook motif contained in the Orc4 subunit. However, in the related fission yeast, Scizosaccharomyces japonicus, replication origin peaks are GC-rich, and it is not clear how ORC recognizes the replication origins in this species [127]. S. japonicus Orc4 contains five repeats of AT-hook, compared to nine in S. pombe Orc4, although it is not known whether the origins are recognized through this motif [128]. On the other hand, close association of G-quadruplex (G4) with replication origins mapped in Drosophila, mouse and human cells suggests a possibility that ORC could recognize G4 structures. Indeed, recent biochemical characterization of purified ORC showed that ORC binds to G4 [129]. ORC can also bind to RNA [130–132] and involvement of RNA in origin recognition cannot be excluded. Furthermore, ORC binding is affected by nucleosome positioning, and NDRs are preferred locations for ORC binding. Bound ORC could then be stabilized through its interaction with neighboring nucleosomes. Association of ORC binding sites with TSS underscores the close relationship between replication and transcription, at least in metazoan, but it is not clear whether transcription has a direct role in initiation of replication or whether it is the transcriptionpermissive chromatin structure that is favored also by replication. We also want to point out the possibility that different modes of




origin recognition may operate with different species and even under different external conditions in a single species, which may help living organisms to better adapt to the changing environment. The number of the pre-RC exceeds that of the origins that are actually fired in S phase. The selection of origins to be fired appears to be stochastic, although the firing efficiency and its timing in S phase may be affected by local chromatin structures, checkpoint signals as well as by other factors. Special proteins such as Fkh1/2 may facilitate the firing of a set of origins. Furthermore, early and efficient generation of pre-RC may facilitate early firing. Competition for limiting factors may also be responsible for firing of some origins earlier than others origins that “lose” the initial competition. Moreover, histone acetylation generally facilitates both pre-RC formation and firing, thus contributing to early replication. In higher eukaryotes with larger genomes, the genome would be divided into different domains (replication domains), each of which is replicated at a specific timing within the S phase. The structure of replication domains may be an intrinsic chromatin trait specific to each cell type, and within each domain, selection of origins to be fired and its timing would be regulated by local factors described above and may be even stochastic. Rif1, bound to nuclear periphery or nucleoli boundaries, plays a key role in defining the replication domain structures. Rif1 binds to chromatin at the nuclear structures in early G1 and this may be a part of the “replication timing decision” process that is known to involve chromosome repositioning. The future challenges include understanding the molecular basis of origin recognition by ORC, that is characterized by its low sequence specificity but is affected by local transcription and chromatin structures, and elucidating how higher order chromosome structures that define the replication domains are generated, how they are spatially arranged within the nuclei, and how they suppress the firing of origins sequestered in these structures. Acknowledgments We thank T. Sutani for critical reading of the manuscript, and all the colleagues from our laboratories for collaboration and useful discussion. References [1] Méchali M. Eukaryotic DNA replication origins: many choices for appropriate answers. Nat Rev Mol Cell Biol 2010;11(10):728–38, http://dx.doi.org/10.1038/nrm2976. [2] Hyrien O, Rappailles A, Guilbaud G, Baker A, Chen CL, Goldar A, et al. From simple bacterial and archaeal replicons to replication N/U-domains. J Mol Biol 2013, http://dx.doi.org/10.1016/j.jmb.2013.09.021. [3] Stinchcomb DT, Struhl K, Davis RW. Isolation and characterisation of a yeast chromosomal replicator. Nature 1979;282(5734):39–43. [4] Theis JF, Newlon CS. The ARS309 chromosomal replicator of Saccharomyces cerevisiae depends on an exceptional ARS consensus sequence. Proc Natl Acad Sci USA 1997;94(20):10786–91. [5] Marahrens Y, Stillman B. A yeast chromosomal origin of DNA replication defined by multiple functional elements. Science 1992;255(5046):817–23. [6] Leonard AC, Méchali M. DNA replication origins. Cold Spring Harb Perspect Biol 2013;5(10), http://dx.doi.org/10.1101/cshperspect.a010116. [7] Linskens MH, Huberman JA. Organization of replication of ribosomal DNA in Saccharomyces cerevisiae. Mol Cell Biol 1988;8(11):4927–35. [8] Newlon CS, Collins I, Dershowitz A, Deshpande AM, Greenfeder SA, Ong LY, et al. Analysis of replication origin function on chromosome III of Saccharomyces cerevisiae. Cold Spring Harb Symp Quant Biol 1993;58:415–23. [9] Nieduszynski CA, Knox Y, Donaldson AD. Genome-wide identification of replication origins in yeast by comparative genomics. Genes Dev 2006;20(14):1874–9, http://dx.doi.org/10.1101/gad.385306. [10] Liachko I, Bhaskar A, Lee C, Chung SCC, Tye BK, Keich U. A comprehensive genome-wide map of autonomously replicating sequences in a naive genome. PLoS Genet 2010;6(5):e1000946, http://dx.doi.org/10.1371/journal.pgen.1000946.

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Insights into the Initiation of Eukaryotic DNA Replication.

Regulated eukaryotic DNA replication origin firing with purified proteins.

Eukaryotic DNA replication.

Eukaryotic DNA replication complex.

Regulation of DNA Replication Initiation by Chromosome Structure.

How MCM loading and spreading specify eukaryotic DNA replication initiation sites.

Spatial-temporal structure of coincident-timing responses.

Nutritional Control of DNA Replication Initiation through the Proteolysis and Regulated Translation of DnaA.

Processing ribonucleotides incorporated during eukaryotic DNA replication.

Structural insights into eukaryotic DNA replication.

Regulation of DNA Replication through Natural Impediments in the Eukaryotic Genome.

Biological asymmetries and the fidelity of eukaryotic DNA replication.

DNA replication timing: Coordinating genome stability with genome regulation on the X chromosome and beyond.

The replication timing program in the hands of two HDACs.

Initiation and regulation of paramyxovirus transcription and replication.

Integrin β6 can be translationally regulated by eukaryotic initiation factor 4E: Contributing to colonic tumor malignancy.

Inferring the spatiotemporal DNA replication program from noisy data.

Rapid turnover of DnaA at replication origin regions contributes to initiation control of DNA replication.

Eukaryotic replication origins as promoters of bidirectional DNA synthesis.

The two faces of higher eukaryotic DNA replication origins.

The Many Roles of PCNA in Eukaryotic DNA Replication.

Eukaryotic genome instability in light of asymmetric DNA replication.

Control of eukaryotic DNA replication at the chromosomal level.

DNA replication initiator Cdc6 also regulates ribosomal DNA transcription initiation.