Search tips
Search criteria 


Logo of nihpaAbout Author manuscriptsSubmit a manuscriptHHS Public Access; Author Manuscript; Accepted for publication in peer reviewed journal;
Curr Opin Genet Dev. Author manuscript; available in PMC 2010 April 1.
Published in final edited form as:
PMCID: PMC2677117

Replication timing and transcriptional control: beyond cause and effect. Part II


Replication timing is frequently discussed superficially in terms of its relationship to transcriptional activity via chromatin structure. However, so little is known about what regulates where and when replication initiates that it has been impossible to identify mechanistic and causal relationships. Moreover, much of our knowledge base has been anecdotal, derived from analyses of a few genes in unrelated cell lines. Recent studies have revisited longstanding hypotheses using genome-wide approaches. In particular, the foundation of this field was recently shored up with incontrovertible evidence that cellular differentiation is accompanied by coordinated changes in replication timing and transcription. These changes accompany subnuclear repositioning, and take place at the level of megabase-sized domains that transcend localized changes in chromatin structure or transcription. Inferring from these results, we propose that there exists a key transition during the middle of S-phase and that changes in replication timing traversing this period are associated with subnuclear repositioning and changes in the activity of certain classes of promoters.


In every multi-cellular system examined, early replication and transcription are strongly correlated. This longstanding correlation has recently been confirmed statistically in Drosophila, human and mouse cells [18]. Moreover, an extensive developmental regulation of replication timing was rigorously demonstrated, associated with changes in transcription [8]. However, progress in understanding the underlying mechanisms remains sluggish. The relationship is clearly indirect, since 10–20% of late-replicating genes are expressed and some genes change transcription without changes in replication timing and vice versa [8]. The simplest explanation is that replication timing is related to features of chromatin and nuclear architecture rather than transcription per se.

Two genres of models, in which either chromatin dictates replication time or vice versa, were illustrated in a prior review (Figure 1 of [9]) and need no revision. Advances in the intervening years have underscored the complexity and variety of mechanisms by which chromatin can influence replication timing, with most effects surprisingly modest and some paradoxical. Ironically, some of our deepest mechanistic insights have come from budding and fission yeasts, yet there is no evidence for an association between replication timing and transcription in these unicellular organisms [10]. On the other hand, evidence that different types of chromatin are assembled at different times during S-phase remains indirect. A third genre of models suggests that replication timing is intimately linked to the 3-dimensional (3D) organization of the genome [1114]. Recent findings have strengthened this association [1517], but the relationship of 3D chromosome organization to chromatin states and transcription remains as elusive as replication timing [18].

Here, we summarize recent progress toward understanding this complex liaison between copying and reading genetic information. We begin with some basic facts about replication control that necessitate framing any discussion of the significance of replication to gene expression in terms of large chromosomal domains. Next, developmental changes in replication timing are discussed, which involve large chromosome segments and accompany spatial repositioning and transcriptional changes for certain classes of promoters. We will then discuss recent experiments that address mechanisms by which chromatin influences replication time and vice versa. Finally, we will discuss how these multiple mechanisms might be related to 3D genome organization.

Replication timing is regulated at the level of large chromosomal domains

In order to discuss how replication timing might influence transcription, it is important to appreciate that replication is regulated at the level of large chromosomal domains. In mammals, rates of fork elongation are on average 1–2 kb/minute and replication is bidirectional [19]. Hence, a single replicon (chromosomal DNA replicated from a single origin) will duplicate 100–200 kb within one hour of a 10 hour S-phase. Moreover, replication frequently proceeds via the nearly synchronous firing of several adjacent origins, resulting in the rapid coordinate duplication of multi-replicon, megabase-sized segments of chromosomes [19]. These segments replicate reproducibly at characteristic times during S-phase, punctuated by origin-less regions through which forks move unidirectionally until they encounter a fork from a neighboring replication domain [4,68,1922]. Hence, any influence of replication timing over chromatin structure must extend over several hundred kilobases.

Mechanisms by which chromatin influences replication time are frequently discussed in terms of the effects of altered histone modifications on the accessibility of specific replication origins to the initiation machinery. However, such simplified models are difficult to reconcile with the concept of origin efficiency [23,24]. Any particular origin is utilized in only a fraction of cell cycles with each cell using a different collection of origins [21,24,25]. Hence, the inactivation of any particular origin may be of little consequence to replication time and frequently will increase the efficiency with which an adjacent origin is used [24]. On the other hand, a localized chromatin change that results in the earlier firing of an origin could potentially cause a replicon-sized shift in replication timing, but if confined to a single origin it would likely fire in only a fraction of cells. In fact, several observations suggest that replication timing is independent of where replication initiates. The human β-globin locus frequently replicates from one of two closely spaced origins while the mouse locus uses many widely dispersed origins yet replication timing is conserved [24]. Moreover, the replication time of chromosomal domains is re-established in each cell cycle prior to and independent of origin site specification [12,26]. Together, these results suggest that any working model for the influence of chromatin over replication timing must account for stable replication timing despite variable origin selection between individual cells.

Replication timing is regulated during development

If replication timing is related to transcription, then it should be subject to developmental control. The last several years witnessed first a challenge to and then a confirmation of the generality of replication timing changes during development. Until a few years ago, evidence other than during mammalian X-chromosome inactivation was restricted to a small number of genes whose replication time had been compared primarily between established, non-isogenic, transformed cell lines [9]. Moreover, many genes replicated at similar times in different cell types [9]. More recently, comprehensive surveys found that replication timing correlates quite strongly with static sequence features of mammalian chromosomes such as isochore GC content and gene density [3,4,27], raising legitimate questions regarding the extent to which replication timing changes during development [16,27]. In fact, a microarray-based comparison of human chromosome 22 between fibroblast and lymphoblastoid cells revealed that only 1% of this chromosome differed in replication time [4]. It then became painfully obvious that there was not a single documented case of an autosomal replication timing change in response to differentiation cues analogous to what was seen during X-chromosome inactivation.

Dynamic changes in replication timing were first confirmed for several individual gene loci during mouse embryonic stem cell (ESC) differentiation to neural precursor cells (NPCs) [28,29]. Approximately one fourth of genes queried in these studies showed measurable changes in replication time and these genes all resided within AT-rich isochores, providing a potential explanation for why the GC-rich human chromosome 22 showed so few differences [4]. However, these studies were not sufficiently comprehensive to infer the frequency of such changes. The question was finally put to rest with a genome-wide study of replication timing during mouse ESC differentiation to NPCs in three independent cell lines using two differentiation schemes [8]. Replication timing changes were highly reproducible and affected up to 20% of the genome with changes at the level of large domains averaging 600 kb. As expected, replication-timing changes were coordinated with changes in transcription and furthermore accompanied subnuclear repositioning [8], supporting an earlier observation for the Mash1 locus [30]. Most recently, extensive replication timing differences were reported between Drosophila Kc (embryonic origin) and Cl8 (from wing imaginal discs) cell lines [31].

Developmental changes: unique isochores facing opposing forces?

The mouse genome-wide study showed that domains that changed replication timing were AT-rich and above a certain threshold of LINE-1 transposon density [8], consistent with an earlier study [28]. Unexpectedly, it further revealed that these domains showed inverse correlation between GC content and gene density, which generally correlate with each other (Figure 1a). In general, isochore AT content is strongly associated with LINE-1 density, proximity to the nuclear periphery and late replication ([8] and references therein), which may involve chromatin association with the nuclear lamina [3234]. In contrast, high gene density is associated with transcription, proximity toward the nuclear interior (where RNA polymerase II transcription factories are enriched [35,36]) and early replication. Therefore, depending on their transcriptional activity, chromosomal domains with inverse correlation between GC content and gene density (e.g. AT-rich/gene-rich isochores) may experience two opposing physical forces that influence their radial positioning and replication timing: an as-yet-undefined isochore sequence-based force (toward the nuclear periphery and late replication) and a transcriptional activity-based force (toward the interior and early replication). Moreover, mouse ESCs have substantially more smaller replication domains and do not show the strong relationship between replication timing and isochore GC content as compared to differentiated cells. This unusual replication domain organization is re-established when adult fibroblasts are induced to the pluripotent state (induced pluripotent stem [iPS] cells; [37]), suggesting that it is characteristic of the pluripotent state [8]. After ESC differentiation, early replication timing correlates better with GC content and radial subnuclear positioning, while maintaining its correlation to transcription and gene density. This suggests that an isochore sequence-based force becomes increasingly influential in shaping the functional and spatial organization of the genome upon differentiation.

Figure 1
Relationship between isochore properties and replication timing regulation, subnuclear position, and transcription

Distinct classes of genes differ in their relationship to replication timing

While correlative, recent microarray studies have allowed us to sharpen hypotheses regarding the relationship between replication timing and transcription. It is now clear that the statistical relationship is similar across cell types and species and is confined to certain classes of genes. In both Drosophila and mouse, most genes replicate in the first third of S-phase and have an equally high probability of being expressed independent of their replication time within this period; a strong relationship between earlier replication timing and transcription is restricted to the ~25% of genes that replicate later during S-phase [1,8] (Figure 1b). In mammalian cells, early-replicating genes are enriched for high CpG-density promoters, while late-replicating genes are enriched for low CpG-density promoters [8] that maintain their repressed state even upon loss of DNA methylation or treatment with TSA [38]. Analyses of mESC differentiation revealed that high and low CpG-density promoters, which generally possess strong and weak promoter activity, respectively, showed distinct behaviors upon switching to a late-replicating environment, only CpG-poor promoters showing higher tendency toward transcriptional down-regulation [8]. Thus, the occasional strong promoter that finds itself located in a replication timing “switching” domain may “come along for the ride” but be unaffected by the replication timing change. This is consistent with reports in several systems that strong promoters can overcome heterochromatin silencing ([8] and references therein) and the observation that tethering a chromosomal region to the nuclear periphery represses some genes but not others [3941]. Overall, these results reinforce the notion that a strong association exists between replication time and transcription for specific classes of genes.

Alterations in chromatin structure induce modest changes in replication timing

Several studies suggest that chromatin modifications directly regulate replication timing, but the effects of any particular modification are relatively minor. Chemical inhibition of histone deacetylases (HDAC) can partially advance replication timing of several mammalian genes [42] as well as the Epstein Barr Virus mini-chromosome [43], while over-expression of a chromatin remodeling complex NoRC delays replication of rRNA genes [44]. In budding yeast, silent chromatin proteins SIR3 [45] and the HDAC Rpd3 [46,47] delay the firing of specific origins, while tethering histone acetyltransferase (HAT) partially advances origin firing [46]. Recently, HAT/HDAC-tethering to the human β-globin origin in mouse cells caused similar partial changes (~20% of S-phase) [48]. Furthermore, modest changes in replication timing were detected for pericentric heterochromatin in Dnmt1, G9a, Eed, and Dicer mutant mouse ESCs, but rather surprisingly, 20 gene loci analyzed were unaffected [49]. In addition, a Suv39h1/h2 lysine methyltransferase (KMTase) mutation showed cell-type specific effects, slightly advancing replication timing of pericentric heterochromatin in embryonic fibroblasts, while slightly delaying it in ESCs [49,50]. Indeed, the role of specific histone modifications in regulating replication timing is still difficult to fathom, since even origins that fire synchronously within a replication domain have different histone modifications [51].

An elegant study in fission yeast provides our first glimpse into the mechanisms by which heterochromatin regulates replication timing in different contexts [52]. Heterochromatic pericentromeres and the mat locus are early-replicating [53] in a Swi6-dependent manner [Swi6: an HP1 (heterochromatin protein 1) ortholog] [52]. In a Swi6 mutant, late replication of the pericentromere but not the mat locus is dependent on the KMTase Clr4 [a Su(var)3–9 ortholog], whereas neither Swi6 nor Clr4 contributed to late replication of subtelomeric heterochromatin [52]. Importantly, Swi6 stimulated loading of the replication factor Sld3 onto origins at the pericentromere and the mat locus in a manner dependent on Dfp1 (Dbf4)-dependent kinase (DDK), an essential kinase required for the initiation of replication. Despite their varied regulation, targeting of Dfp1 advanced replication timing of all three loci.

Together, these results support the model (Figure 1 of [9]) that chromatin can control the accessibility of initiation factors to pre-replication complexes, but also underscore the complexity of the underlying mechanisms. To date, no epigenetic mark has been shown to correlate with replication timing significantly better than transcription itself [8,31]. Notably, however, the effects of most chromatin manipulations are modest and context-dependent, with a few exceptions [43,44,54,55].

Does replication timing affect chromatin structure?

Since chromatin is assembled at the replication fork, an appealing scenario is that replication timing dictates chromatin states that in turn regulate replication timing in the subsequent cell cycle, providing a means of epigenetic inheritance during somatic development. Moreover, since replication is regulated at the level of replicons, a change in replication timing could rapidly transmit a change in chromatin state to many genes simultaneously. Clearly this attractive model merits a definitive test, but it is currently impossible to manipulate replication timing without affecting other properties of a chromatin domain. The most compelling, albeit indirect, evidence that different chromatin structures are assembled at different times during S-phase is that reporter plasmids injected into early or late S-phase mammalian nuclei assembled into hyper- or hypo-acetylated chromatin, respectively [56]. This result prompted speculation that the replication fork scaffold protein PCNA might recruit different chromatin modifiers at different times during S-phase to assemble different types of chromatin [57]. Unfortunately, although dozens of proteins localize to replication forks, only HDAC2 [58] and MBD2-MBD3 [59] have been found to localize specifically to sites of late-replicating chromatin and both of these studies examined overexpressed epitope-tagged proteins. Hence, the existence of temporally regulated chromatin assembly mechanisms remains an attractive but unsubstantiated hypothesis.

Large replication timing changes accompany radial subnuclear repositioning

In every case examined, dynamic developmental changes in replication timing accompany subnuclear repositioning [8,30]. Although a genome-wide survey of such relationship is currently impractical, spatial patterns of DNA replication in the nucleus change dramatically as cells move through S-phase, demonstrating a global coupling of subnuclear repositioning with replication-timing changes [19,57] (Figure 1b). Moreover, replication timing is re-established during early G1-phase at the timing decision point (TDP), coincident with the repositioning of chromosomal domains in the nucleus after mitosis [12,13]. Consistently, chromatin mobility is relatively high during the first 1–2 hours of G1-phase, after which it is locally constrained through the remainder of interphase [60,61]. Moreover, inducible targeting of loci to the nuclear lamina requires passage through mitosis and takes place during late telophase to early G1-phase [39,40]. Together, these results predict that, during differentiation, subnuclear repositioning takes place at the TDP within the G1-phase preceding changes in replication timing. Testing this prediction will require an evaluation of the sequence of events during intermediate steps of differentiation. Notably, G1-phase length is highly variable between cell types, which may influence the extent to which nuclei are reorganized before replication initiates. In particular, the extremely short G1-phase in ESCs could result in replication starting before isochore sequence-based chromosome reorganization mechanisms (discussed earlier) are completed, explaining the larger number of smaller replication domains that do not replicate according to their isochore sequence properties.

The middle of S-phase represents a period of dramatic change, including: (1) a dramatic change in the spatial distribution of replication sites (Figure 1b, pattern II to III) [19,57]; (2) a transition from R to G band replication [17,62]; (3) a temporary reduction in replication fork movement [62]; (4) a sharp decline in the density of genes being replicated [5]; and (5) the onset of a relationship between replication timing and transcription (Figure 1b) [8]. Thus, changes in replication timing that straddle mid S-phase are likely to accompany movements between subnuclear compartments (Figure 1b) or even formation of compartments that are more difficult to reverse once established (e.g. formation of a Barr body during X-chromosome inactivation [63]). In contrast, even large changes in replication timing that are restricted to the first half of S-phase may be less consequential (Figure 1b). Consistently, several early-replicating loci such as Oct-4 become slightly later replicating upon ESC differentiation but do not straddle mid S-phase [29,64] nor accompany radial subnuclear repositioning [8,65]. In addition, asynchronously replicated homologs of imprinted or monoallelically-expressed genes can show relatively small replication timing differences during mid S-phase but may exhibit significant radial position differences [42,66,67].

Conclusions and future directions

Our understanding of replication timing remains a fragmented set of half-truths that are currently impossible to integrate into absolutes. In fact, among the many experimental manipulations performed over the years, it is arguably only G1 nuclei before the TDP that display a globally disturbed replication-timing program [12,13]. In contrast, the majority of chromatin manipulations have resulted in relatively minor effects. Moreover, modifications of chromatin proteins generally persist through mitosis or are reinstated onto chromatin during mitotic exit, prior to the TDP [50,6870]. This suggests that various chromatin structures analyzed are not sufficient to dictate the global timing program but their effects on replication timing may represent a secondary, fine-tuning role. Our current view is that replication-timing changes through the middle of S-phase are qualitatively distinct and more likely to involve subnuclear compartment changes (Figure 1b). Testing this hypothesis will require deciphering the principles of how nuclear genome reorganization occurs in early G1-phase, particularly the influence of isochore sequences and transcription. Since the molecular nature of these principles is still a mystery, comparative genome-wide analyses will continue to provide important insights to sharpen hypotheses. Studies of the effects of chromatin structures on replication timing regulation will continue to be important, particularly those that can tease out relationships to replication initiation mechanisms [52]. Also important are experiments to determine the extent to which replication time can influence chromatin assembly, for which at present evidence is scant. Finally, there are clearly non-transcriptional roles for replication timing such as the maintenance of genome stability [43,71,72]. At present, it appears that we are each collecting separate views of complex mechanisms linking genome structure and function, waiting for our various half-truths to intersect and reveal a more complete picture.


We would like to thank J Huberman, H Masukata and M Schwaiger for helpful discussions, and P Norio for helpful comments. Research in the Gilbert lab is supported by NIH grant GM83337. We apologize to those who could not be cited due to space limitation.


Publisher's Disclaimer: This is a PDF file of an unedited manuscript that has been accepted for publication. As a service to our customers we are providing this early version of the manuscript. The manuscript will undergo copyediting, typesetting, and review of the resulting proof before it is published in its final citable form. Please note that during the production process errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain.


1. Schubeler D, Scalzo D, Kooperberg C, Van Steensel B, Delrow J, Groudine M. Genome-wide DNA replication profile for Drosophila melanogaster: a link between transcription and replication timing. Nat Genet. 2002;32:438–442. [PubMed]
2. MacAlpine DM, Rodriguez HK, Bell SP. Coordination of replication and transcription along a Drosophila chromosome. Genes Dev. 2004;18:3094–3105. [PubMed]
3. Woodfine K, Fiegler H, Beare DM, Collins JE, McCann OT, Young BD, Debernardi S, Mott R, Dunham I, Carter NP. Replication timing of the human genome. Hum Mol Genet. 2004;13:191–202. [PubMed]
4. White EJ, Emanuelsson O, Scalzo D, Royce T, Kosak S, Oakeley EJ, Weissman S, Gerstein M, Groudine M, Snyder M, et al. DNA replication-timing analysis of human chromosome 22 at high resolution and different developmental states. Proc Natl Acad Sci U S A. 2004;101:17771–17776. [PubMed]
5. Jeon Y, Bekiranov S, Karnani N, Kapranov P, Ghosh S, MacAlpine D, Lee C, Hwang DS, Gingeras TR, Dutta A. Temporal profile of replication of human chromosomes. Proc Natl Acad Sci U S A. 2005;102:6419–6424. [PubMed]
6. Karnani N, Taylor C, Malhotra A, Dutta A. Pan-S replication patterns and chromosomal domains defined by genome-tiling arrays of ENCODE genomic areas. Genome Res. 2007;17:865–876. [PubMed]
• 7. Farkash-Amar S, Lipson D, Polten A, Goren A, Helmstetter C, Yakhini Z, Simon I. Global organization of replication time zones of the mouse genome. Genome Res. 2008;18:1562–1570. A recent genome-wide study of DNA replication timing in a mouse lymphocytic leukemia cell line confirmed correlations of replication timing with GC content, gene density, and transcription previously reported for human lymphoblasts. Although the genome-wide analysis was performed at relatively low resolution (probes unevenly spaced averaging 45 kb), the unique synchrony method allowed the authors to conclude that 9% of the genome replicates asynchronously, as compared to 20% concluded from an earlier study in human HeLa cells [6] [PubMed]
•• 8. Hiratani I, Ryba T, Itoh M, Yokochi T, Schwaiger M, Chang CW, Lyou Y, Townes TM, Schubeler D, Gilbert DM. Global reorganization of replication domains during embryonic stem cell differentiation. PLoS Biol. 2008;6:e245. This is the first and most comprehensive metazoan genome-wide study of DNA replication timing during differentiation, providing incontrovertible evidence for an extensive regulation of replication timing during development. Differentiating mouse ESCs to NPCs, this study shows that: (1) chromosomes consist of multi-megabase domains of coordinate replication, separated by apparently origin-less transition regions, (2) replication domain organization is highly conserved within a cell type but roughly 20% of the genome changed upon ESC differentiation to NPCs, (3) differentiation was accompanied by the consolidation of smaller differentially replicating domains into larger coordinately replicated units whose replication time was more aligned to isochore GC content and LINE-1 density but not gene density, (4) replication-timing changes were coordinated with transcription changes for weak CpG-poor promoters more than strong CpG-rich promoters, and were accompanied by rearrangements in subnuclear position, and (5) more smaller replication domains and a higher density of timing transition regions that interrupt isochore replication timing define a novel characteristic of the pluripotent state common to ESCs and iPS cells. [PubMed]
9. Gilbert DM. Replication timing and transcriptional control: beyond cause and effect. Curr Opin Cell Biol. 2002;14:377–383. [PubMed]
10. Donaldson AD. Shaping time: chromatin structure and the DNA replication programme. Trends Genet. 2005;21:444–449. [PubMed]
11. Gilbert DM. Nuclear position leaves its mark on replication timing. J Cell Biol. 2001;152:F11–16. [PMC free article] [PubMed]
12. Dimitrova DS, Gilbert DM. The spatial position and replication timing of chromosomal domains are both established in early G1-phase. Mol Cell. 1999;4:983–993. [PubMed]
13. Li F, Chen J, Izumi M, Butler MC, Keezer SM, Gilbert DM. The replication timing program of the Chinese hamster beta-globin locus is established coincident with its repositioning near peripheral heterochromatin in early G1 phase. J Cell Biol. 2001;154:283–292. [PMC free article] [PubMed]
14. Heun P, Laroche T, Raghuraman MK, Gasser SM. The positioning and dynamics of origins of replication in the budding yeast nucleus. J Cell Biol. 2001;152:385–400. [PMC free article] [PubMed]
15. Englmann A, Clarke LA, Christan S, Amaral MD, Schindelhauer D, Zink D. The replication timing of CFTR and adjacent genes. Chromosome Res. 2005;13:183–194. [PubMed]
• 16. Grasser F, Neusser M, Fiegler H, Thormeyer T, Cremer M, Carter NP, Cremer T, Muller S. Replication-timing-correlated spatial chromatin arrangements in cancer and in primate interphase nuclei. J Cell Sci. 2008;121:1876–1886. High throughput fluorescence in situ hybridization is an oxymoron, but this paper comes as close as one can get, analyzing 64 extremely early or extremely late replicating BACs for their subnuclear position. The results confirm conclusions from global labeling of DNA synthesis that late replication takes place close to the nuclear periphery, while early replication takes place more internally and identified gene density as an important parameter of three-dimensional genome organization, particularly with respect to position within a particular chromosome territory. [PMC free article] [PubMed]
17. Federico C, Cantarella CD, Di Mare P, Tosi S, Saccone S. The radial arrangement of the human chromosome 7 in the lymphocyte cell nucleus is associated with chromosomal band gene density. Chromosoma. 2008;117:399–410. [PubMed]
18. Fraser P, Bickmore W. Nuclear organization of the genome and the potential for gene regulation. Nature. 2007;447:413–417. [PubMed]
19. Berezney R, Dubey DD, Huberman JA. Heterogeneity of eukaryotic replicons, replicon clusters, and replication foci. Chromosoma. 2000;108:471–484. [PubMed]
20. Raghuraman MK, Winzeler EA, Collingwood D, Hunt S, Wodicka L, Conway A, Lockhart DJ, Davis RW, Brewer BJ, Fangman WL. Replication dynamics of the yeast genome. Science. 2001;294:115–121. [PubMed]
21. Norio P, Kosiyatrakul S, Yang Q, Guan Z, Brown NM, Thomas S, Riblet R, Schildkraut CL. Progressive activation of DNA replication initiation in large domains of the immunoglobulin heavy chain locus during B cell development. Mol Cell. 2005;20:575–587. [PubMed]
• 22. McCune HJ, Danielson LS, Alvino GM, Collingwood D, Delrow JJ, Fangman WL, Brewer BJ, Raghuraman MK. The Temporal Program of Chromosome Replication: Genomewide Replication in clb5{Delta} Saccharomyces cerevisiae. Genetics. 2008;180:1833–1847. This paper provides a clear demonstration that domains of early and late replication in budding yeast frequently span multiple origins and can encompass hundreds of kilobases as in mammals, revealing existence of replication domains in this single-celled, genetically tractable organism. [PubMed]
23. Rhind N. DNA replication timing: random thoughts about origin firing. Nat Cell Biol. 2006;8:1313–1316. [PMC free article] [PubMed]
24. Aladjem MI. Replication in context: dynamic regulation of DNA replication patterns in metazoans. Nat Rev Genet. 2007;8:588–600. [PubMed]
25. Takebayashi SI, Manders EM, Kimura H, Taguchi H, Okumura K. Mapping sites where replication initiates in mammalian cells using DNA fibers. Exp Cell Res. 2001;271:263–268. [PubMed]
26. Li F, Chen J, Solessio E, Gilbert DM. Spatial distribution and specification of mammalian replication origins during G1 phase. J Cell Biol. 2003;161:257–266. [PMC free article] [PubMed]
27. Costantini M, Bernardi G. Replication timing, chromosomal bands, and isochores. Proc Natl Acad Sci U S A. 2008;105:3433–3437. [PubMed]
28. Hiratani I, Leskovar A, Gilbert DM. Differentiation-induced replication-timing changes are restricted to AT-rich/long interspersed nuclear element (LINE)-rich isochores. Proc Natl Acad Sci U S A. 2004;101:16861–16866. [PubMed]
29. Perry P, Sauer S, Billon N, Richardson WD, Spivakov M, Warnes G, Livesey FJ, Merkenschlager M, Fisher AG, Azuara V. A dynamic switch in the replication timing of key regulator genes in embryonic stem cells upon neural induction. Cell Cycle. 2004;3:1645–1650. [PubMed]
30. Williams RR, Azuara V, Perry P, Sauer S, Dvorkina M, Jorgensen H, Roix J, McQueen P, Misteli T, Merkenschlager M, et al. Neural induction promotes large-scale chromatin reorganisation of the Mash1 locus. J Cell Sci. 2006;119:132–140. [PubMed]
•• 31. Schwaiger M, Stadler MB, Bell O, Kohler H, Oakeley EJ, Schubeler D. Chromatin state marks cell-type and gender specific replication of the Drosophila genome. Genes Dev. 2009 in press. This recent Drosophila genome-wide study (35 bp spacing between probes) shows ~20% replication timing differences between Kc cells (embryonic origin) and Cl8 cells (from wing imaginal discs), which generally correlated with differential transcriptional regulation. Furthermore, a transcription-independent correlation of replication timing with histone H4 lysine-16 acetylation was identified. Finally, unlike female cells (Kc), male cells (Cl8) were found to replicate the X chromosome uniformly early in S-phase regardless of transcriptional up-regulation due to dosage compensation. [PubMed]
32. Chubb JR, Boyle S, Perry P, Bickmore WA. Chromatin motion is constrained by association with nuclear compartments in human cells. Curr Biol. 2002;12:439–445. [PubMed]
33. Pickersgill H, Kalverda B, de Wit E, Talhout W, Fornerod M, van Steensel B. Characterization of the Drosophila melanogaster genome at the nuclear lamina. Nat Genet. 2006;38:1005–1014. [PubMed]
• 34. Guelen L, Pagie L, Brasset E, Meuleman W, Faza MB, Talhout W, Eussen BH, de Klein A, Wessels L, de Laat W, et al. Domain organization of human chromosomes revealed by mapping of nuclear lamina interactions. Nature. 2008;453:948–951. This study in a human cell line confirms conclusions from an earlier study in Drosophila tissue culture cells [33], demonstrating that genome-wide interactions of the nuclear lamina occur primarily for late-replicating sequences and that ~90% of lamina-interacting genes are silenced. [PubMed]
35. Sadoni N, Langer S, Fauth C, Bernardi G, Cremer T, Turner BM, Zink D. Nuclear organization of mammalian genomes. Polar chromosome territories build up functionally distinct higher order compartments. J Cell Biol. 1999;146:1211–1226. [PMC free article] [PubMed]
36. Xie SQ, Pombo A. Distribution of different phosphorylated forms of RNA polymerase II in relation to Cajal and PML bodies in human cells: an ultrastructural study. Histochem Cell Biol. 2006;125:21–31. [PubMed]
37. Takahashi K, Yamanaka S. Induction of pluripotent stem cells from mouse embryonic and adult fibroblast cultures by defined factors. Cell. 2006;126:663–676. [PubMed]
• 38. Lande-Diner L, Zhang J, Ben-Porath I, Amariglio N, Keshet I, Hecht M, Azuara V, Fisher AG, Rechavi G, Cedar H. Role of DNA methylation in stable gene repression. J Biol Chem. 2007;282:12194–12200. The roles of different gene repression mechanisms were assessed by taking advantage of wild-type and Dnmt1−/− fibroblast cells treated with or without TSA and analyzing gene expression profiles by microarray analysis. Group-A genes were de-repressed in Dnmt1−/− cells, with no further activation upon TSA treatment. Group-B genes were not expressed in Dnmt1−/− cells and were only slightly induced by TSA. All 8 group-A genes analyzed were early replicating while half of the 18 group-B genes tested were late replicating. Thus, late-replicating genes appear to be enriched for certain classes of promoters whose regulation is more tightly controlled. [PubMed]
• 39. Reddy KL, Zullo JM, Bertolino E, Singh H. Transcriptional repression mediated by repositioning of genes to the nuclear lamina. Nature. 2008;452:243–247. [PubMed]
• 40. Kumaran RI, Spector DL. A genetic locus targeted to the nuclear periphery in living cells maintains its transcriptional competence. J Cell Biol. 2008;180:51–65. [PMC free article] [PubMed]
• 41. Finlan LE, Sproul D, Thomson I, Boyle S, Kerr E, Perry P, Ylstra B, Chubb JR, Bickmore WA. Recruitment to the nuclear periphery can alter expression of genes in human cells. PLoS Genet. 2008;4:e1000039. These three reports analyzed the effect of targeting genomic loci to the nuclear periphery. Targeting required passage through mitosis [39,40], and Kumaran et al [40] demonstrated that the repositioning took place during the transition from late telophase to early G1-phase (coincident with TDP). Peripheral targeting repressed some endogenous genes surrounding the targeted site but not others [41], implying gene promoter-specific effects similar to those seen within domains that change replication timing in [8] [PMC free article] [PubMed]
42. Bickmore WA, Carothers AD. Factors affecting the timing and imprinting of replication on a mammalian chromosome. J Cell Sci. 1995;108:2801–2809. [PubMed]
• 43. Zhou J, Snyder A, Lieberman PM. Epstein-Barr Virus Episome Stability Is Coupled to a Delay in Replication Timing. J Virol. 2009;83:2154–2162. Epstein-Barr Virus (EBV) is a good model for mammalian chromosomes because its autonomous replication as a mini-chromosome relies on host pre-replication complex (pre-RC) proteins. These authors provide evidence that proteins located at the EBV replication origin recruit a histone de-acetylase that is required for the late replication of EBV. Preventing de-acetylation significantly advanced replication timing of EBV and reduced the stable maintenance of the viral mini-chromosome. This work suggests an intriguing model in which positioned nucleosomes near origins are first acetylated by HBO1 during pre-RC assembly and then a subset of origins is de-acetylated to delay their firing. [PMC free article] [PubMed]
44. Li J, Santoro R, Koberna K, Grummt I. The chromatin remodeling complex NoRC controls replication timing of rRNA genes. Embo J. 2004;24:120–127. [PubMed]
45. Stevenson JB, Gottschling DE. Telomeric chromatin modulates replication timing near chromosome ends. Genes Dev. 1999;13:146–151. [PubMed]
46. Vogelauer M, Rubbi L, Lucas I, Brewer BJ, Grunstein M. Histone acetylation regulates the time of replication origin firing. Mol Cell. 2002;10:1223–1233. [PubMed]
47. Aparicio JG, Viggiani CJ, Gibson DG, Aparicio OM. The Rpd3-Sin3 histone deacetylase regulates replication timing and enables intra-S origin control in Saccharomyces cerevisiae. Mol Cell Biol. 2004;24:4769–4780. [PMC free article] [PubMed]
• 48. Goren A, Tabib A, Hecht M, Cedar H. DNA replication timing of the human beta-globin domain is controlled by histone modification at the origin. Genes Dev. 2008;22:1319–1324. This paper shows, as was previously shown by similar experiments in budding yeast [46,47], that targeting a strong acetylase or deacetylase to a replication origin near the human β-globin locus can advance or retard its replication timing. The changes in replication timing did not affect transcription, although the manipulations only altered replication timing by about 20% of the length of S-phase and the size of the affected chromosomal regions was not determined. Subnuclear position changes were not evaluated. [PubMed]
• 49. Jorgensen HF, Azuara V, Amoils S, Spivakov M, Terry A, Nesterova T, Cobb BS, Ramsahoye B, Merkenschlager M, Fisher AG. The impact of chromatin modifiers on the timing of locus replication in mouse embryonic stem cells. Genome Biol. 2007;8:R169. These authors analyzed replication timing of 20 different gene loci and several types of repetitive elements in mouse ESCs with different chromatin modifier mutations. Surprisingly, no changes were detected in gene loci while modest changes were found for some of the repetitive elements analyzed, including pericentric heterochromatin in Eed, Dnmt1, G9a, Suv39h1/h2, and Dicer mutants. Interestingly, while Suv39h1/h2 KMTase mutation slightly delayed replication timing of pericentric heterochromatin in ESCs, the same mutation slightly advanced its replication timing in mouse embryonic fibroblasts [50], consistent with the complex, context-dependent effects of chromatin mutations on replication timing seen in [52] [PMC free article] [PubMed]
50. Wu R, Singh PB, Gilbert DM. Uncoupling global and fine-tuning replication timing determinants for mouse pericentric heterochromatin. J Cell Biol. 2006;174:185–194. [PMC free article] [PubMed]
51. Norio P. DNA replication: the unbearable lightness of origins. EMBO Rep. 2006;7:779–781. [PubMed]
•• 52. Hayashi MT, Takahashi TS, Nakagawa T, Nakayama JI, Masukata H. The heterochromatin protein Swi6/HP1 activates replication origins at the pericentromeric region and silent mating-type locus. Nat Cell Biol. 2009 In fission yeast, the pericentromeres, mating type loci and telomeres are all assembled into silent chromatin organized by heterochromatin proteins Clr4 and Swi6, yet telomeres replicate late while pericentromeric and mating type loci heterochromatin replicates early [53]. These authors show that the replication timing of each of these blocks of heterochromatin is altered differently upon mutations in their chromatin components, but timing at all three loci appears to be regulated by the accessibility of the Dfp1 (Dbf4)-dependent kinase (DDK). Specifically, they show that Swi6 recruits Dfp1 to the pericentromeric and mating type loci to advance their replication, but not in the context of late replicating telomeric heterochromatin. These results underscore the context dependency of chromatin influences on replication. [PubMed]
53. Kim SM, Dubey DD, Huberman JA. Early-replicating heterochromatin. Genes Dev. 2003;17:330–335. [PubMed]
54. Lin CM, Fu H, Martinovsky M, Bouhassira E, Aladjem MI. Dynamic alterations of replication timing in mammalian cells. Curr Biol. 2003;13:1019–1028. [PubMed]
55. Bergstrom R, Whitehead J, Kurukuti S, Ohlsson R. CTCF regulates asynchronous replication of the imprinted H19/Igf2 domain. Cell Cycle. 2007;6:450–454. [PubMed]
56. Zhang J, Xu F, Hashimshony T, Keshet I, Cedar H. Establishment of transcriptional competence in early and late S phase. Nature. 2002;420:198–202. [PubMed]
57. McNairn AJ, Gilbert DM. Epigenomic replication: linking epigenetics to DNA replication. Bioessays. 2003;25:647–656. [PubMed]
58. Rountree MR, Bachman KE, Baylin SB. DNMT1 binds HDAC2 and a new co-repressor, DMAP1, to form a complex at replication foci. Nat Genet. 2000;25:269–277. [PubMed]
59. Tatematsu KI, Yamazaki T, Ishikawa F. MBD2-MBD3 complex binds to hemi-methylated DNA and forms a complex containing DNMT1 at the replication foci in late S phase. Genes Cells. 2000;5:677–688. [PubMed]
60. Walter J, Schermelleh L, Cremer M, Tashiro S, Cremer T. Chromosome order in HeLa cells changes during mitosis and early G1, but is stably maintained during subsequent interphase stages. J Cell Biol. 2003;160:685–697. [PMC free article] [PubMed]
61. Thomson I, Gilchrist S, Bickmore WA, Chubb JR. The radial positioning of chromatin is not inherited through mitosis but is established de novo in early G1. Curr Biol. 2004;14:166–172. [PubMed]
62. Takebayashi S, Sugimura K, Saito T, Sato C, Fukushima Y, Taguchi H, Okumura K. Regulation of replication at the R/G chromosomal band boundary and pericentromeric heterochromatin of mammalian cells. Exp Cell Res. 2005;304:162–174. [PubMed]
63. Heard E, Bickmore W. The ins and outs of gene regulation and chromosome territory organisation. Curr Opin Cell Biol. 2007;19:311–316. [PubMed]
64. Azuara V, Perry P, Sauer S, Spivakov M, Jorgensen HF, John RM, Gouti M, Casanova M, Warnes G, Merkenschlager M, et al. Chromatin signatures of pluripotent cell lines. Nat Cell Biol. 2006;8:532–538. [PubMed]
65. Wiblin AE, Cui W, Clark AJ, Bickmore WA. Distinctive nuclear organisation of centromeres and regions involved in pluripotency in human embryonic stem cells. J Cell Sci. 2005;118:3861–3868. [PubMed]
66. Gribnau J, Hochedlinger K, Hata K, Li E, Jaenisch R. Asynchronous replication timing of imprinted loci is independent of DNA methylation, but consistent with differential subnuclear localization. Genes Dev. 2003;17:759–773. [PubMed]
67. Takizawa T, Gudla PR, Guo L, Lockett S, Misteli T. Allele-specific nuclear positioning of the monoallelically expressed astrocyte marker GFAP. Genes Dev. 2008;22:489–498. [PubMed]
68. Belyaev ND, Keohane AM, Turner BM. Histone H4 acetylation and replication timing in Chinese hamster chromosomes. Exp Cell Res. 1996;225:277–285. [PubMed]
69. Kruhlak MJ, Hendzel MJ, Fischle W, Bertos NR, Hameed S, Yang XJ, Verdin E, Bazett-Jones DP. Regulation of global acetylation in mitosis through loss of histone acetyltransferases and deacetylases from chromatin. J Biol Chem. 2001;276:38307–38319. [PubMed]
70. Cowell IG, Aucott R, Mahadevaiah SK, Burgoyne PS, Huskisson N, Bongiorni S, Prantera G, Fanti L, Pimpinelli S, Wu R, et al. Heterochromatin, HP1 and methylation at lysine 9 of histone H3 in animals. Chromosoma. 2002;111:22–36. [PubMed]
• 71. Chang BH, Smith L, Huang J, Thayer M. Chromosomes with delayed replication timing lead to checkpoint activation, delayed recruitment of Aurora B and chromosome instability. Oncogene. 2007;26:1852–1861. This paper, along with a series of earlier papers by the same group, identifies a cancer-related chromosomal phenotype that is associated with a significant delay in mitotic chromosome condensation (DMC), a delay in the mitosis-specific phosphorylation of histone H3, and a 2–3 h delay in the replication timing (DRT) of the entire chromosome. The delay appears to be controlled by a cis-regulatory element, similar in principle to X-chromosome inactivation, which can globally delay replication of the chromosome while maintaining the relative differences in replication timing along the length of the chromosome. [PMC free article] [PubMed]
• 72. Bianchi A, Shore D. Early replication of short telomeres in budding yeast. Cell. 2007;128:1051–1062. Telomeric heterochromatin replicates late in budding yeast, and mutations in telomeric heterochromatin proteins Sir3 or yKu can advance telomere replication timing. Here, the authors artificially shorten telomeres and demonstrate that this shortening of telomeric heterochromatin results in earlier replication and increased telomerase elongation activity at the early replicating telomeres. This suggests the existence of a feedback loop between replication timing and telomere length in which shorter telomeres advance replication timing to re-lengthen the telomere, revealing a novel function for replication timing in maintaining chromosome integrity. [PubMed]