ACFrOgABPC6r7ZS13kpZG7xm7yFM17c88c_kSsCqodzRHc2VSN8SG5EdOC3v-P4r_dh_FTurvJrSapzfCoS27xnVpa5Vs1VNWIcecEP_V5PFRK__HIM-xectZ5IIg7xXskttRJPn6ckkAFMGa7NA

Prévia do material em texto

Before each cell division, eukaryotic chromosomes 
must be entirely duplicated. Duplication involves the 
replication not only of DNA but also of all chromatin 
components, appropriate epigenetic information and 3D 
organization. Almost 60 years ago, the first insights into 
this process revealed that large segments of mammal­
ian chromosomes replicate at distinct times during the 
S phase of the cell cycle1. Modern genomics approaches 
have revealed that multi­ megabase segments of chromo­
somes, referred to as constant timing regions (CTRs), repli­
cate via the coordinate activation of adjacent replicons at 
characteristic times during the S phase. These CTRs are 
delimited by large timing transition regions (TTRs) along 
which DNA synthesis progressively advances2–8 (Fig. 1a).
All known eukaryotes have such a ‘DNA replication 
timing programme’, but the size of CTRs and TTRs var­
ies, being scaled to the size of their genome9. This pro­
gramme is extremely robust — almost every attempt to 
disrupt it, using chemical inhibitors, gene knockdown 
or gene knockout, has been unsuccessful (Table 1). 
The programme is also highly conserved between 
closely related eukaryotic species9–16. Altogether, these 
observations suggest that the timing of replication of 
discrete regions of chromosomes has an important bio­
logical function. Replicating different segments of the 
genome at different times ensures that the number of 
replication forks does not exceed the availability of limit­
ing factors such as nucleo tides and proteins required for 
DNA replication17, consolidates forks for rapid response 
to replication stress18 and ensures that the genome is 
fully replicated18. However, these constraints that lead 
to segmental replication of the genome do not explain 
why the genome should be replicated in a defined and 
evolutionarily conserved temporal order. Another pro­
posed function of controlling DNA replication timing 
is to regulate the gene dosage19. However, most genes 
are subject to mechanisms that reduce the expression of 
post­ replicated genes two­ fold20,21 so it is not clear how 
extensively such a mecha nism could drive the positive 
evolutionary selection of a specific genome­ wide timing 
programme. Of note, mutation frequencies vary strongly 
during the S phase22. The frequency of point mutations 
is much higher for late­ replicating DNA, possibly 
owing to the downregu lation of mismatch repair (MMR) 
proteins during mid S phase23, and different types of 
structural mutation are correlated with early or late 
replication22–27. Thus, one hypothesis for the biological 
function of a defined replication timing programme is 
that it concentrates genomic variation to particular parts 
of the genome.
In mammalian cells, considerable attention has 
been given to the relationship of replication timing to 
chromatin architecture and cell differentiation. The 
replication timing programme remains highly stable 
within a cell type and even between individual cells 
of the same type28,29, unlike transcription and many 
epigenetic marks30, but the timing of replication of at 
least 50% of the genome changes during cell fate tran­
sitions. These developmental switches occur in smaller 
units of 400–800 kb that are referred to as ‘replication 
Constant timing regions
(CTRs). Regions of the 
chromatin containing one 
or more adjacent replication 
domains that are replicating 
at nearly the same time.
Replicons
Units of replication comprising 
a replication origin and the 
DNa being replicated.
Timing transition regions
(TTRs). Regions of the 
chromatin between two 
replications domains 
replicating at different times.
Replication forks
Complexes comprising 
template DNa, oligonucleotide 
primers and proteins necessary 
for DNa replication. Two ‘sister’ 
replication forks are assembled 
at each replication origin 
and may be associated in 
3D space.
Control of DNA replication timing 
in the 3D genome
Claire Marchal, Jiao Sima and David M. Gilbert *
Abstract | The 3D organization of mammalian chromatin was described more than 30 years 
ago by visualizing sites of DNA synthesis at different times during the S phase of the cell cycle. 
These early cytogenetic studies revealed structurally stable chromosome domains organized 
into subnuclear compartments. Active- gene-rich domains in the nuclear interior replicate early, 
whereas more condensed chromatin domains that are largely at the nuclear and nucleolar 
periphery replicate later. During the past decade, this spatiotemporal DNA replication programme 
has been mapped along the genome and found to correlate with epigenetic marks, transcriptional 
activity and features of 3D genome architecture such as chromosome compartments and 
topologically associated domains. But the causal relationship between these features and DNA 
replication timing and the regulatory mechanisms involved have remained an enigma. The recent 
identification of cis- acting elements regulating the replication time and 3D architecture of 
individual replication domains and of long non- coding RNAs that coordinate whole chromosome 
replication provide insights into such mechanisms.
Department of Biological 
Science, Florida State 
University, Tallahassee, 
FL, USA.
*e- mail: gilbert@bio.fsu.edu
https://doi.org/10.1038/ 
s41580-019-0162-y
REvIEWS
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domains’4,13,31–33. Replication domains consist of 1–4 
replicons that synchronously initiate DNA replication, 
forming ‘replicon clusters’ that can be observed on iso­
lated DNA fibres34,35. The larger CTRs are believed to 
consist of multiple adjacent replication domains that 
replicate at similar times4 (Fig. 1a). The replication timing 
of specific replication domains becomes stably altered in 
several diseases. Certain diseases, as well as individual 
patients, can therefore be identified by ‘replication timing 
signatures’36–38. As not just DNA but the entire structure 
of the chromosome must be replicated, a second logical 
hypo thesis for the biological function for robust cell­ type­ 
specific spatial and temporal control of DNA replication 
is that where and when chromatin replicates is impor­
tant for the maintenance of cell­ type­specific epigenetic 
states and provides a mechanism to alter them during 
cell fate transitions. Testing this hypothesis will require 
an understanding of the mechanisms regulating the 
coordinated firing of replicons at defined times and their 
organization into replication domains. Thus, decrypting 
the relationship between replication timing, chromatin 
architecture, transcription and cell differentiation is an 
important challenge for the future.
The regulation of DNA replication timing in mam­
malian cells also occurs at the level of entire chromo­
somes39. Asynchronous replication of chromosomes was 
also first observed nearly 60 years ago, but it was origi­
nally thought to be unique to X chromosomes in female 
mammals. However, it was later discovered that most 
cancer cells have at least one chromosome for which 
replication is severely delayed and often continues when 
cells enter the G2 phase or even mitosis40–45. The mecha­
nisms that control entire chromosome replication, and 
their relationship to the mechanisms that regulate the 
Chromosomea
b
Early
Late
TADs
LADs
RT
Multiple cell
divisionsFoci ≈ domains
Late-replicating ≈
B compartment
Early-replicating ≈
A compartment
Single chromosome stained
≈ chromosome territory
Nucleolus
Nuclear envelope
Lamina
RD RD RD RD RD RD
TTR TTR TTR
CTR CTR CTR CTR
A compartment B compartment
Fig. 1 | Replication timing relationship to 3D chromatin structure. a | Current model of the relationship between 
replication timing (RT) and chromatin structure. Early and late constant timing regions (CTRs) are 1–5 Mb regions 
separated by timing transition regions (TTRs) as demarcated within the red shaded area. These CTRs consist of one to 
severalreplication domains (RDs), defined as chromatin segments that coordinately switch RT during cell fate changes 
(that is, between different cell types; see Fig. 2a). RDs share the properties and approximate boundaries of a subset of 
topologically associated domains (TADs), aligning most closely with TADs that are at compartment boundaries. Early CTRs 
correspond to the A compartment, but late CTRs correspond to the B compartment and TTRs correspond to the transitions 
between compartments. Both TTRs and late CTRs correspond to lamina- associated domains (LADs). b | RT illuminates 
genome architecture: nuclei after an early S pulse label (green) followed by several hours of a chase period and then a late 
S pulse label (red). In this model, observable foci of DNA synthesis correspond to the replication domains in panel a and 
early/late- replicating chromatin corresponds to A/B compartments. After multiple passages, only one chromosome 
per cell remains labelled, marking the chromosome territory47,202. However, the foci retain their label intensity46–49 and 
genetic continuity203, demonstrating that the DNA that is synthesized during one cell cycle remains clustered together 
as a structural unit of chromosomes for many generations.
Replication stress
Stalling of replication forks, 
which can be caused by 
chemical interference or 
radiation, or can be the result 
of a lack of nucleotides or 
proteins necessary for DNa 
replication.
Mismatch repair
(MMR). DNa repair mechanism 
occurring at the replication 
fork involved in the repair of 
base mismatch and small 
insertions/deletions occurring 
during DNa replication or 
recombination.
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722 | DeCemBeR 2019 | volume 20 
temporal order of replication of individual chromosomal 
domains, are poorly understood.
In this Review, we discuss our current understand­
ing of the relationships between DNA replication timing 
and the 3D architecture of the genome in mammalian 
cells. First, we describe how replication timing is related 
to the spatial organization of the genome. Second, we 
discuss the dynamics of replication timing and 3D chro­
matin interactions during cell differentiation and dur­
ing progression through the cell cycle. We then present 
a general overview of the regulation of the replication 
timing, followed by a discussion of recent discoveries of 
cis elements and trans­ acting factors that regulate repli­
cation timing. Finally, we discuss the interplay between 
replication timing and transcription.
Links to spatial genome organization
Until recently, studying large­ scale chromatin architec­
ture was restricted to cytogenetic methods, with genome 
organization often highlighted by labelling sites of DNA 
replication. Pulse labelling of cells with nucleotide ana­
logues for a short period of time enabled the visualization 
of ‘replication foci’ (Fig. 1b), which were sites of coordi­
nated replicon activation. After chasing for multiple cell 
divisions, these foci retained their size, shape and inten­
sity, suggesting that they are stable chromosomal units 
of structural organization46–49. When chased for longer 
periods of time, these pulse–chase experiments resulted 
in cells in which only one or a few chromosomes retained 
the label, revealing the nuclear organization of chromo­
somes into discrete territories (Fig. 1b), which could also 
be visualized by fluorescence in situ hybridization (FISH) 
with chromosome­ specific probes50. Finally, the spatial 
distribution of the labelled foci was seen to vary during 
the S phase, most dramatically when comparing early 
with late S phase, suggesting the existence of two organi­
zational compartments of chromatin in the nucleus that 
replicate in the first or second half of the S phase51–53 
(Fig. 1b). This spatial organization of late­ replicating chro­
matin at the nuclear periphery and around the nucleolus, 
and of early­ replicating chromatin in more central 
regions of the nucleus, is remarkably conserved from 
single­ cell ciliates to humans54. Once these patterns are 
established during early G1 phase49, they are maintained 
throughout interphase owing to the immobile nature of 
mammalian interphase chromatin55. Over the past dec­
ade, several genomics technologies to study the molecular 
organization of chromatin in the nucleus were devel­
oped, which enabled these cyto genetic observations to 
be related to 1D molecular maps of chromosomal DNA 
and its epigenetic marks, as well as to 3D genome archi­
tecture revealed by Hi- C (high­ throughput chromosome 
conformation capture) methods (box 1). Here we discuss 
how these different scales of genome organization are 
related to different aspects of DNA replication.
Early- and late- replicating chromatin compartments. 
From its initial inception, Hi­ C identified the first prin­
cipal component of large­ scale chromatin folding in 
the nucleus that separates chromatin into two nuclear 
compartments, defined as A and B56 (box 1). These 
A and B compartments roughly correlate with actively 
transcribed open chromatin and more silent compact 
chromatin, respectively, and very closely correlate with 
early­ and late­ replicating DNA13,57, as predicted by the 
cytogenetic studies summarized above (Fig. 1). At higher 
resolution, Hi­ C can resolve these compartments into 
several sub­ compartments (box 1), which continue to 
correlate with DNA regions that replicate at different 
times58. Interestingly, when compartments are defined 
at very high resolution, they overlap with active and 
inactive transcription units and no longer correlate with 
DNA replication timing59, indicating that the resolution 
of analysis of genomics data must be scale­ appropriate 
for the biological process being investigated.
Chromosome territories. In the nucleus, chromosomes 
occupy limited areas defined as chromosome territories. 
These territories can be observed by FISH60, but they can 
also be visualized by labelling replicating DNA during 
Hi- C
a genome- wide chromatin 
conformation capture 
technology that uses 
chromatin digestion, re- ligation 
of sequences in close proximity 
and sequencing to identify 
chromatin pairwise 3D 
chromatin interactions in 
the nucleus.
Table 1 | Gene deletion effects on global replication timing and chromatin architecture in mammals
Gene deletion Effect on replication timing Effect on 3D structure
Rif1 (KO and KD) Genome- wide changes (MEF, mESC, 
HeLa)164–166
Localized changes (mESC)166
Cohesin (KD and KO of 
factors from the complex)
No change (HCT116)77 Loss of a part of TAD boundaries, no compartment 
change (HCT116, mouse hepatocytes)73,80
CTCF (KD and KO) No change (KO: D.M.G., unpublished 
data, mESC)75
Loss of CTCF loops and local interactions, no 
compartment change (mESC, mNPC, HEK293T)151,205
Suz12 (KO) No change (mESC)66 N/A
MeCP2 (KO) No change (D.M.G., unpublished data) N/A
G9a (KO) Few local changes (mESC, mNPC)112 No change (mESC)71
H1a–H1d–H1e (triple KO) No change (D.M.G., unpublished data) Local changes (mESC)206
cMyc–nMyc (double KO) No change (D.M.G., unpublished data) N/A
BAF53a (KO) No change (mESC)207 N/A
BAF250a (KO) Local changes (mESC)207 N/A
Brg1 (KO) Local changes (mESC)207 N/A
CTCF, CCCTC- binding factor; KD, knockdown; KO, knockout; MEF, mouse embryonic fibroblast; mESC, mouse embryonic stem 
cell; mNPC, mouse neural precursor cell (derived from ESC); N/A, data not available; TAD, topologically associated domain.
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one cell cycle and then letting cells divide multiple times47. 
Indeed, after each division, labelled chromosomes will 
be randomly distributed among the daughter cells, even­
tually resulting in nuclei containing only one labelled 
chromo some, for which the signal will remain undiluted 
and will highlight the entire chromosome. Thus, the 
observed labelled region of the nucleus corresponds to 
the territory of that chromosome(Fig. 1b).
It is possible that the temporal window within which 
each whole chromosome replicates is determined by 
long non­ coding RNAs (lncRNAs) coating the chromo­
some from which they are expressed. Three lncRNAs 
have been shown to be essential for the regulation of 
the replication time window of the chromosome from 
which they are transcribed. First, xist is necessary for 
late replication of the whole inactive X chromosome 
in mice61, but also its deletion in differentiated human 
cells results in the inactive X being replicated even later 
in S phase (that is, the entire window of time during which 
the chromosome replicates is shifted)61,62. Secondly, 
it was recently discovered that deletion of two lnc­
RNAs (ASAR6 and ASARA15) known as asynchronous 
replication and autosomal RNas (ASARs) drastically 
delay the replication of the chromosomes they are 
coating40–42.
Lamina- associated domains. A large subnuclear 
compartment is the chromatin near the nuclear 
periphery63. This compartment, consisting of lamina- 
associated domains (LADs), can be identified either 
cytogenetically64,65 or using genomics methods66,67 as 
chromatin that is bound by proteins of the inner nuclear 
envelope. As much of the chromatin that is near the 
nuclear lamina can also be in proximity to the nucleolar 
Principal component 
analysis
(PCa). Mathematical 
transformation applied 
on complex (with multiple 
variables) datasets, to extract 
values describing the data 
called PC1, PC2 and so forth, 
with PC1 the value best 
describing the data.
Xist
long non- coding RNa that is 
expressed in female cells from 
only one x chromosome. xist 
inactivates the x chromosome 
from which it is expressed, 
enabling dosage compensation 
(similar x- linked gene 
expression levels in male 
and female cells).
Asynchronous replication 
and autosomal RNAs
(aSaRs). long non- coding 
RNas that are essential for 
the timely replication and 
condensation of the entire 
chromosome from which they 
are expressed.
Lamina- associated domains
(laDs). Domains of chromatin 
that come in close proximity 
to the nuclear lamina.
Box 1 | Compartments and topologically associated domains in cell population and single cells
In the past decade, the development of chromatin conformation capture methods, particularly Hi- C (high- throughput 
chromosome conformation capture) and its derivatives, has enabled exquisite quantification of the frequency with which 
pairs of chromosomal sites interact genome- wide187, allowing the molecular identification of 3D chromatin architecture. 
These experiments consist of cleaving DNA within immobilized chromatin and re- ligating it to create chimeric DNA 
fragments composed of two or more interacting regions. These chimeric DNA fragments are then analysed by quantitative 
PCR or sequencing (see the figure, part a). The first component (PC1) of the principal component analysis (PCA), also 
named the eigenvector, of the pairwise interactions identifies the two major folding compartments termed A and B56, 
enriched for interactions within each group (see the figure, part b)56. This division of the chromatin correlates well with 
different chromatin features such as histone marks, gene transcription and DNase hypersensitivity, and particularly well 
with replication timing, likely representing the euchromatin and heterochromatin compartments illuminated by observing 
DNA synthesis cytologically13,56,57. These compartments identified through PCA can be further sub- stratified via a 
clustering analysis into several sub- compartments called A1, A2, B1, B2 and B3 (ReF.58). Similar compartments can be 
observed when Hi- C is done in single cells where only a snapshot of interactions can be observed per cell28,188. on a 
finer scale, chromosomes can be seen to consist of a series of sub- megabase-sized self- interacting units70,71 termed 
topologically associated domains (TADs), which appear to be punctuated by boundaries that are stable in different cell 
types (see the figure, part c). These units may correspond to the stable units of chromosome structure also illuminated by 
labelling DNA synthesis cytologically, termed replication foci35. Within TADs, even finer clusters of interactions, or sub- 
TADs, can be identified58,189 that are less conserved among cell types83,190,191 and may represent cell- type-specific chromatin 
hubs. TADs can also be observed at the single- cell level97, but this overlap is not perfect and both single- cell Hi- C188 and 
Oligo-​STORM​(oligo-​fluorescence​in situ​hybridization-​mediated​tracing​of​large​chromosome​segments​through​
individual cell nuclei assisted by super- resolution microscopy) show that the positions of TAD boundaries vary from cell to 
cell78. CCCTC- binding factor (CTCF) and the cohesin complex are central to TAD structure. Depletion of CTCF or a subunit 
of the cohesin complex leads to an important perturbation of TADs without affecting chromatin compartments73,151 or 
replication timing75,77, highlighting the fact that compartments and TADs are being maintained by different mechanisms.
TAD TAD
Chromosome
A
B
cb
a
Chromatin
digestion
Chromatin
ligation
DNA
purification
and
sequencing
Read 1 Read 2
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724 | DeCemBeR 2019 | volume 20 
periphery or at other internal heterochromatic sites in 
alternating cell cycles65,68, LADs comprise most of the 
late­ replicating compartment observed by cytogenetic 
labelling (Fig. 1b), also defined as the B compartment 
by Hi­ C69. However, LADs do not contain only late­ 
replicating chromatin and do not correlate as well with 
replication timing as do the A/B compartments. This 
puzzle was resolved by the finding that the borders of 
the earliest replication domains overlap strongly with the 
borders of LADs. Therefore, replication forks originat­
ing from early­ replicating domains move rapidly into 
the adjacent LADs, causing them to replicate early even 
though they lack active origins of replication66. Moreover, 
the transitions from LADs (lamina­ associated) to inter- 
 laDs (not lamina­ associated) are sharp, whereas tran­
sitions from the A to B compartments are gradual 
and correspond to the gradual TTRs between early­ and 
late­replicating CTRs. Thus LADs, which comprise periph­
eral, nucleolar and other heterochromatic foci, consist of 
TTRs and late CTRs (Fig. 1a).
Topologically associated domains. Along the length 
of chromosomes, 3D chromatin interactions are clus­
tered within chromatin domains known as topologically 
associated domains (TADs)70,71 (box 1). The boundaries 
of TADs, which can be identified as the sites where 
chromatin interactions change in directionality, align 
with the boundaries of replication domains, suggesting 
that TADs correspond to the stable units of chromo­
some structure identified cytogenetically as repli cation 
foci66,72 (Fig. 1a). Indeed, replication foci labelled in 
living cells and tracked by correlative live and super- 
resolution microscopy displayed biophysical param eters 
consistent with TADs35. However, the relationship 
between TAD structure and replication domains remains 
obscure and, similar to compartments, TADs also lose 
their alignment with replication domains at high resolu­
tion as they become further divided into sub­ TADs or 
individual chromatin loops. An important challenge is to 
understand the biological significance of TADs defined 
at different scales, and their relationship to chromosome 
functions, including DNA replication.
TADs are delimited by boundaries and are defined 
as regions within which chromatin interactions occur 
with higher frequency (that is, within the TAD) than 
in other regions (outside the TAD) (box 1). The CCCTC- 
binding factor (CTCF) zinc finger protein and the cohesin 
complex are the best­ studied factors that play a role in 
determining TAD structure. These factors bind to some 
TAD boundaries and form chromatin loops through the 
extrusion of chromatin throughthe doughnut­ shaped 
cohesin molecule until a second CTCF site of the oppo­
site polarity is encountered73. Although their binding at 
these boundaries can restrict enhancer–promoter inter­
actions to within TADs74, there is no evidence that the 
TAD boundaries provide a mechanism for confining 
coordinated replicon activation to within the domain. 
Indeed, deletion of chromatin sites localized at the TAD 
boundaries did not affect replication timing of several 
domains75. Furthermore, although depleting CTCF or 
cohesin can severely diminish insulation of chromatin 
interactions between adjacent TADs measured by Hi­ C, 
CTCF75,76 and cohesin77 depletion has no effect on repli­
cation timing. Moreover, high­ resolution oligopaint 
tracing of individual chromosome segments revealed 
that there is a high degree of cell­ to­cell heterogeneity 
in the positioning of TAD boundaries. These usually 
coincide with CTCF and cohesin binding sites but, in 
the absence of cohesin, TAD boundaries persist but no 
longer form at specific sites78.
In principle, TAD boundaries do not need to be 
fixed, but could instead simply be the site of transi­
tion between two chromosomal domains enriched 
for self­ interaction79. In terms of replication timing, 
cis elements required for early replication, called early- 
replication control elements (ERCEs), were identified 
within TADs, located far from the boundaries. ERCEs 
interact with each other independently of CTCF/cohesin, 
and their interactions persist in the absence of CTCF75. 
Moreover, ERCEs are sites of master transcription regu­
latory factor binding and resemble transcriptional super- 
enhancers that have been shown to maintain their inter­
actions in the absence of cohesin73,80. In the case of the 
analysed TAD, deletion of its boundaries had no effect 
on its replication timing, but deletion of ERCEs affected 
the local TAD architecture. Thus, ERCEs promote early 
replication within the confines of a TAD, and interact 
to form loops that contribute to TAD structure, inde­
pendently of any specific boundary elements. Together, 
these observations suggest that repli cation timing may 
be an additional mechanism to link genome architec­
ture to function, and interfering with ERCEs should 
provide a means to test this hypothesis and identify the 
underlying mechanisms.
Replication timing in cell fate transitions
Whereas TAD boundaries have been claimed to be 
largely conserved between cell types58,70,81,82, the A/B 
compartments and the regions associating with the 
nuclear lamina frequently differ between cell types67,83. 
Similarly, replication domains retain their boundaries in 
different cell types, but differ in their replication timing, 
correlating with A/B compartments and lamina associa­
tion66. These observations support a ‘replication domain 
model’84, in which replication domains are invariant 
structural units, but the time at which they replicate, and 
their higher order organization and positioning within 
the nucleus, is developmentally regulated.
Spatiotemporal domain consolidation and compartment 
switching during differentiation. Stem cells provide a 
system to track changes in properties (for example, 
repli cation timing and subnuclear localization of replica­
tion domains) relative to each other. Early observations 
demonstrated that when replication timing changes 
during a switch in cell fate, it does so in units of approxi­
mately 0.5 Mb (replication domains, Fig. 2a), and as 
stem cells become progressively more restricted in their 
differentiation potential, an increasing number of adja­
cent replication domains replicate at similar times. This 
results in fewer and larger early and late CTRs through 
a process termed ‘domain consolidation’4. Replication 
domains also consolidate spatially, which was initially 
visualized by FISH — as the physical compaction and 
Inter- LADs
Domains between two adjacent 
lamina- associated domains.
Topologically associated 
domains
(TaDs). Domains of chromatin 
enriched for 3D interactions 
within the domain as 
compared with between 
domains.
Correlative live and super- 
resolution microscopy
Combines the temporal 
resolution of time- lapse 
fluorescence microscopy 
with the spatial resolution of 
super- resolution microscopy.
CCCTC-binding factor
(CTCF). Protein highly 
conserved in eukaryotes that 
associates with chromatin to 
mediate transcription and 
chromatin insulation.
Cohesin
Protein complex composed 
of SMC1 and SMC3 that forms 
a large ring structure that 
accommodates two strands 
of chromatin. Cohesin has 
a key architectural role, 
forming chromatin loops and 
maintaining sister chromatids 
tied together after DNa 
replication.
Early replication control 
elements
(eRCes). DNa sequences 
necessary for the early 
replication of entire replication 
domains.
Super- enhancers
Chromatin region rich in 
enhancers, with a high level of 
transcription associated factors 
and acetylated histones.
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movement of domains closer to their neighbours — and 
by targeted chromatin conformation capture (4C) — as 
increased interactions with neighbouring domains85. 
Compartment consolidation is coincident with changes 
in the subnuclear position of replication domains (inte­
rior to periphery or vice versa), also originally visualized 
by FISH85,86. Later, it was confirmed genome­ wide by 
Hi­ C that adjacent TADs consolidate spatially into A and 
B compartments, correlated with changes in replication 
timing that consolidate temporally83. These findings, 
together with the cytogenetic studies from the 1990s, 
have suggested an intimate link between subnuclear 
compartmentalization and replication timing.
Interestingly, it has been found that the correlation 
between replication timing and chromatin A/B compart­
ments is weaker in human embryonic stem cells (ESCs) 
than in differentiated cells and that during lineage com­
mitment the first cell cycles are accompanied by uncoor­
dinated changes in compartments and replication timing 
that become resolved in later cell cycles87. Thus, either 
the compartments or the replication timing can switch 
prior to the other during the early period of germ layer 
commitment, demonstrating that the two can be uncou­
pled, at least during periods of high plasticity. To under­
stand the mechanisms underlying the close but indirect 
connection between compartments and replication 
timing, it will be necessary to be able to independently 
manipulate the compartment in which a domain resides 
and the compartment’s replication timing. The identi­
fication of cis elements that control replication timing 
and compartments provides the first molecular tool with 
the potential to enable such manipulation75.
Chromatin conformation 
capture
Technologies that assess 3D 
interactions within chromatin. 
These technologies are based 
on the cutting and re- ligation 
of chromatin immobilized in 
intact nuclei and identification 
of pairwise interactions 
by quantitative PCR or 
sequencing.
Early
Late
Early
Late
Early
Late
RT
RT
RT
H9 ESC
Chromosome 8a
H9 smooth
muscle
H9
mesothelial
130 Mb 140 Mb
G1b S G2 MitosisTDP
DNA replicationNo
competency
for RT
Competency
 for RT
Loss of competency for RT
TADs
 A/B compartments
Expected early (green)/
late (red) patterns after
forced replication
Early/late replication
patterns (compartments)
labelled in a normal
S phase and chased to
the next cell cycle
Segregation of chromatin
interactions into
A/B compartments
Segregation of chromatin
interactions into TADs
Fig. 2 | 3D chromatin structure and replication timing are dynamic during cell differentiation and during the cell 
cycle. a | Replication timing (RT) is regulated during differentiation. Here, RT is shown for a region of chromosome 8, 
in human embryonic stem cells H9 (H9 ESC) and two differentiated H9 cells. Some regions switch RT during differentiation(black to grey, or grey to black), whereas others remain constant. RT data are available at www.replicationdomain.com. 
b | Both a defined RT programme and interphase chromatin architecture are set up coincidently at the timing decision 
point (TDP) during the G1 phase49,100. The information that defines RT is lost during the G2 phase99. In nuclei that are 
artificially forced to replicate their DNA before the TDP or after the S phase, DNA replication does not follow any specific 
RT49,99. The early- and late- replicating 3D compartments illuminated by replication labelling in the prior S phase are 
re- established at the TDP and persist through the remainder of interphase into the G2 phase, demonstrating that this 
spatial organization is not sufficient to dictate an RT programme. 3D chromatin interactions — both the separation 
between large- scale spatial compartments and the distinction between topologically associated domains (TADs) — are 
dismantled during mitosis and re- formed at the TDP, coincident with the establishment of RT100. Whereas compartments 
and TADs become slightly more or less distinct, respectively, during the course of the S phase101, the major architectural 
changes in genome architecture occur during entry into and exit from mitosis.
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Uncoupling between replication timing and 3D chroma-
tin interactions during early embryogenesis. Replication 
timing can be uncoupled from TAD structures. A recent 
study revealed that TADs form after the four­ cell stage of 
mouse embryonic development, in a DNA replication­ 
dependent manner88. However, spatiotemporal patterns 
of replication can already be observed in one­ cell mouse 
embryos89, suggesting that A/B compartments and a 
replication timing programme are present already in the 
mammalian zygote, in the absence of TAD structures or 
zygotic transcription. Similarly, in early embryos under­
going rapid cleavage divisions, distinct patterns of repli­
cation timing and chromatin architecture are observed 
before (in zebrafish) or concomitant with (in Drosophila 
melanogaster) zygotic genome activation (ZGA), but it is 
not yet known when they are established during these 
early cleavage stages90–95. The study of genome archi­
tecture and DNA replication timing in cleavage­ stage 
embryos will be facilitated by the recent development 
of single­ cell Hi­ C, single­ cell RNA sequencing and 
single­ cell replication timing28,96,97.
Replication timing during the cell cycle
The replication programme is established early during the 
G1 phase of the cell cycle, at the timing decision point 
(TDP)49,98, and is dismantled during the S phase99. If DNA 
replication is forced to initiate very early during G1 (before 
the TDP) or later during G2, this results in replication 
taking place in a random temporal order49,99.
The TDP: replication timing and chromatin architecture 
are established during early G1 phase. The TDP is a short 
window of time during which chromatin domains move 
to the positions where they will reside for the remainder 
of interphase, which can be identified cytogenetically49. 
Moreover, targeted chromatin conformation capture (4C) 
showed that the re­ establishment of 3D chromatin inter­
actions to form both TADs and A/B compartments fol­
lowing cell division occurs within the same time window 
as the TDP100 (Fig. 2b), and single­ cell Hi­ C technology 
confirmed that chromatin interactions are re­ established 
during the early G1 phase, which is the time window of 
the TDP101. Interestingly, this time window precedes the 
time when replication origin sites are selected, known 
as the origin decision point102. Forcing replication to 
initiate between the TDP and the origin decision point 
results in a correct programme of DNA replication — 
that is, early­ and late­ replication regions replicate at 
the expected time — but replication initiation (box 2) 
occurs at random sites103. This demonstrates that repli­
cation timing and genome architecture are established 
prior to and independently of mechanisms that control 
the sites where replication initiates. Consistent with 
independent mechanisms, replication timing is highly 
determini stic from cell to cell28,29, while initiation sites are 
chosen in a highly stochastic fashion with many sites used 
in less than 2% of cell cycles and no two chromosomes 
replicated from same cohort of initiation sites104 (box 2).
Replication timing but not chromatin architecture is 
lost after S phase. The position of subnuclear domains 
observed cytogenetically and the chromatin interactions 
observed by Hi­ C are preserved during the G2 phase, 
whereas replication timing is lost99,100, indicating that 
the basic scaffold of interphase genome architecture 
is not sufficient to dictate replication timing (Fig. 2b). 
Thus, during the cell cycle, replication timing becomes 
established at the moment of compartmentalization 
and TAD formation, but this temporal programme is 
lost prior to the dismantling of 3D structure that occurs 
in mitosis99,100. It is worth noting that single­ cell Hi­ C 
studies found that 3D chromatin interactions are modi­
fied during the S phase, as TAD boundaries are weaker 
whereas compartments are more strongly separated in 
the G2 phase compared with G1 phase101. The finding 
that the partitioning of compartments becomes stronger 
when the replication timing programme becomes erased 
further emphasizes the indirect relationship between 
nuclear compartments and replication timing.
These results suggest a model in which 3D chromatin 
interactions create a scaffold during early G1 phase, with 
which additional cell cycle­ regulated proteins interact to 
establish the DNA replication timing programme and 
are then cleared from chromatin during DNA replication. 
Possible candidates for these factors, the nature of which 
remains to be determined, are suggested below.
Regulation of replication timing
Identifying the regulators of DNA replication timing is 
challenging. First, almost all gene knockouts and knock­
downs assessed in the literature do not cause any major 
alterations in the timing of replication, which suggests 
that a robust maintenance system is in place (Table 1). 
Second, studying origins of replication is technically 
difficult in mammals as the pool of origins that are used 
for replication differs from cell to cell (box 2), obscuring 
genome­ wide studies on cell populations.
Epigenetic mechanisms as replication timing regula-
tors. Replication timing is extremely robust, remaining 
largely unaltered in a given cell type following genetic 
knockouts and drug perturbations, but it is extensively 
modi fied during cell fate transitions. It is also erased 
during the S phase, but restored during the following 
G1 phase of the cell cycle99. Altogether, this suggests the 
involvement of epigenetic mechanisms. Several obser­
vations support this hypothesis. First, replication timing 
has been cytologically observed to differ between the two 
alleles of imprinted genes in a cell­ type­dependent man­
ner105,106. Allele­ specific replication timing can also be 
observed on non­ imprinted loci107,108. In some cases, the 
asynchrony observed between alleles could be explained 
either by epigenetic mechanisms or by single­ nucleotide 
polymorphisms or structural variation that influence the 
replication timing (see below), but in cases of random 
allelic differences such as X­ chromosome inactivation39 
or allele switching109 it presumably must occur via speci­
fic epigenetic marks. Second, some copies of the ribo­
somal RNA (rRNA) genes replicate at different times 
than other copies, and this difference is dependent on 
their DNA methyl ation state (Fig. 3a). rRNA genes are 
present in multiple copies in eukaryotes. In mammals 
they are regulated by the chromatin remodeller complex 
NoRC, which establishes silenced chromatin to repress 
Zygotic genome activation
(Zga).Stage of embryonic 
development at which the 
transcription of the zygotic 
genome becomes activated.
Imprinted genes
genes that are expressed 
only from one chromosome 
homologue, the choice of 
which is dependent on its 
parental origin. The mechanism 
behind genomic imprinting 
relies on DNa methylation.
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transcription. Each rRNA gene can be present in either of 
two states: either not associated with NoRC, presenting a 
low level of methylation at the promoter and replicating 
early; or bound by NoRC, highly methylated and repli­
cating late. Overexpression of NoRC leads to an increase 
in the proportion of rRNA gene copies per cell that are 
highly methylated and replicating late, indicating that 
DNA methylation and chromatin silencing can delay DNA 
replication110. It has been shown that for each rRNA 
gene, the activity is different on each allele: one allele is 
repressed and replicates late, whereas the other allele 
is active and replicates early111. Third, differential repli­
cation timing of the inactive (Xi) and active X chromo­
somes is a clear example where epigenetic mechanisms 
regulate replication timing. Delayed replication of the 
inactive X chromosome seems to be dependent on 
the expression of the lncRNA Xist, which coats the entire 
chromosome. However, Xist is also required to silence 
the Xi, raising the possibility that the effect on replication 
timing might be indirect. Indeed, direct effects of Xist on 
replication timing remain to be confirmed. Moreover, 
deletion of Xist in mouse fibroblasts causes the Xi to rep­
licate even later in the S phase61, highlighting the complex 
role of Xist in controlling DNA replication. Finally, it has 
recently been shown that random allelic asynchrony of 
replication is common and regulated by lncRNAs43.
The genomics era has enabled correlation of many 
features of chromatin and DNA sequence to replication 
Box 2 | Initiation of DNA replication; from origin licensing to origin firing
In eukaryotes, origin recognition complex (oRC), Cdc6 and Cdt1 cooperate to load two copies of the ring- shaped 
heterohexameric mCm replicative helicase around DNA in a process called origin licensing, and the fully loaded double 
hexamer is then referred to as a pre- replication complex (pre- RC). Pre- RCs are assembled during telophase but can continue 
to assemble throughout most of the G1 phase. A subset of origins is then activated (origin activation) during the G1/S 
transition whereas others remain dormant and are removed during passage of the replication fork (see the figure). origin 
firing occurs at different times during S phase in a defined temporal order. Pre- RCs assembled as early as telophase49,192 are 
fully functional to initiate replication193,194 but remain inactive until they are phosphorylated by the Dbf4-dependent kinase 
(DDK)155, and are activated when cyclin- dependent kinase (CDK), mCm10, Treslin and TopBP1 participate in the assembly 
of Cdc45 and the GINS complex with the mCm hexamer to form the active CmG helicase157,195,196. Restricting initiation to 
only once per cell cycle is elegantly accomplished with a two- cycle engine; pre- RCs assemble under conditions that do not 
permit initiation, and initiation does not occur until conditions are no longer permissive for pre- RC assembly197. In mammals, 
the features of DNA or chromatin that dictate where pre- RCs assemble is unknown, but assembly sites appear to be highly 
flexible as many more pre- RCs are assembled than are activated and sites of initiation are chosen in a highly stochastic 
fashion; indeed, the same chromosomes in different cells do not use the same cohort of replication origins104,198. It is also 
not known how certain pre- RCs are chosen for initiation199. The excess pre- RCs are called ‘dormant origins’200,201 and they 
are used to ensure that all DNA is replicated in a timely manner, by serving as a backup when encountering conditions 
that slow replication forks or to complete fortuitous large regions that have not initiated as cells approach the G2 phase. 
Although replication origins do not have an intrinsic firing time, elements that regulate replication timing promote or 
inhibit the activation of pre- RCs in their chromatin domain.
TopBP1
ORC
Cdc6 Cdt1
MCM10
GINS
Cdc45
Treslin
MCM2–7
Active origin
Origin
licencing 
Origin
licensing
Dormant origin
Origin
activation
Origin
firing
G1
S
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timing. These studies have suggested many intriguing 
hypotheses, but also highlight that even very strong cor­
relations do not imply causal relationships (for exam­
ple112). Several histone post­ translational modifications 
are associated with replication timing. Marks of active 
transcription such as histone H3 lysine 4 monomethyl­
ation (H3K4me1), dimethylation (H3K4me2) and tri­
methylation (H3K4me3), H3K20me1 and H3K36me3, 
and H3K9 and H3K27 acetylation, correlate with early 
replication113. Interestingly, the only mark strongly 
associated with late replication is H3K9me2 (ReF.13), but 
removing detectable H3K9me2 by knocking out the 
methyltransferase responsible for the deposition of this 
mark has no effect on replication timing112, despite 
the fact that it may dissociate LADs from the nuclear 
periphery68 and can cause spreading of phosphorylated 
H3S10, which is normally restricted to early­ replicating 
regions, into TTRs and late­ replicating regions114. 
However, histone methylation on H4K20 is required for 
the proper replication of some late­ replicating domains: 
impairing H4K20 methylation leads to a further delay 
in the replication of several late­ replicating regions115.
A simple explanation for some marks being corre­
lated with early DNA replication is that if these marks 
are rapidly established during chromatin maturation, the 
density of such marks detected by chromatin immuno­
precipitation (ChIP) would be directly proportional to 
the copy number of each locus, and thus their profile 
along the genome would exactly match replication tim­
ing. For example, monomethylation is deposited rapidly, 
whereas dimethylation and trimethylation occur much 
later, sometimes early in the following cell cycle116. 
This could be the reason why H3K9me1 is predicted to 
correlate very closely with early DNA replication, but 
H3K9me2 and H3K9me3 do not. This hypothesis does 
not explain the H3K9me2 correlation, but does pro­
vide a clear example of the logical fallacy of concluding 
causality from correlation.
Early studies on the β-globin domain in human cells 
pointed to a role for histone acetylation in the control 
of replication timing117. This chromosomal domain 
is replicated early in erythroid cells that express the 
β­ globin gene, but replicates late in cell types that do not 
express β­ globin118 (Fig. 3b). In HeLa cells, inhibition of 
histone deacetylation by trichostatin A (TSA) advanced 
the time of origin firing in this locus119, whereas for cing 
deacetylation of this locus on a transgene integrated into 
the genomes of erythroid cells by targeting a histone 
deacetylase to the transgene delayed its replication120.
In mouse fibroblasts, TSA treatment of mouse cells 
slightly advanced the timing of replication of the nor­
mally late­ replicating pericentromeric heterochromatin 
measured by immunofluorescence121. Also, in yeast, 
two histone deacetylases, Sir2 and Rpd3, regulate the 
replication timing of ribosomal DNa (rDNA)122. These 
studies indicate that DNA acetylation levels have an 
impact on the timing of its replication. Furthermore, 
the reverse may also be true; replication timing can 
influence the acetylation levels of chromatin123,124. Thus, 
histone acetylation seems to be a bona fide player in the 
regulation of early replication, which is now also sup­
ported by the discovery of ERCEs, which are strongly 
enriched in H3K27Ac marks75.Importantly, however, 
histone acetylation, especially H3K27Ac, is commonly 
found at all enhancer regions, whether or not they are 
ERCEs75, so histone acetylation alone is not sufficient to 
advance replication timing.
Interestingly, using mathematical modelling it is pos­
sible to predict cell­ type­specific replication timing on 
the basis of DNase hypersensitivity profiles125. Although 
there is a general correlation between early replica­
tion and chromatin nuclease accessibility, domains 
whose replication timing is developmentally regulated 
are nuclease inaccessible, similar to late­ replicating 
domains, and have many epigenetic features of late­ 
replicating domains, even in cell types where they are 
replicated early (and thus predicted to have features of 
early­ replicating domains)85,100,126.
Sequence- dependent mechanisms regulating replica-
tion timing. At the genome­ wide level, different tech­
niques have tried to establish maps of initiation sites, 
or active origins of replication in mammalian cells, 
by sequencing different elements — purified small 
nascent DNA strands (SNS­ seq; RNA­ primed single­ 
stranded DNA fragments of 800–1200 bp)127, replica­
tion bubbles (Bubble­ seq; DNA fragments containing 
a Ribosomal gene cluster
b β-Globin gene locus
c
Non-erythroid cells Erythroid cells
Late-replication timing Early-replication timing
Late-replication timing Early-replication timing
Methylated CpG
Non-acetylated histones Acetylated histones
Unmethylated CpG
TSA
(histone acetylation)
HDAC
(histone deacetylation)
Early
Late
RTRep 1
Early
Late
RTRep 2
60 MbChr10 70 Mb 80 Mb
Castaneus allele 129 alleleSequence-specificreplication timing
Fig. 3 | Epigenetic versus sequence- specific regulation of replication timing. 
a | The cluster of ribosomal DNA consists of repeated genes coding for ribosomal 
RNAs. Correlated with the methylation status of each copy of these genes, they will 
replicate late (highly methylated) or early (low methylation level)110,111. b | The β- globin 
locus is an example of developmental regulation of replication timing (RT): in erythroid 
cells, the locus is activated, rich in histone acetylation and early replicating. In non- 
erythroid cells, the locus is silenced, without histone acetylation and late replicating. 
Inducing histone acetylation in non- erythroid cells with trichostatin A (TSA) treatment 
promotes early replication of the locus, whereas depleting the locus for histone 
acetylation by targeting histone deacetylases (HDACs) to a β- globin transgene in 
erythroid cells induces late replication of the locus117–120. c | Sequence- specific RT can 
be observed using subspecies F1 hybrid mouse embryonic stem cells (ESCs). Here, some 
non- imprinted regions have different RT, depending on the allele. Data from GEO series 
GSE95091 (ReF.204).
β- Globin domain
a megabase- sized locus 
containing the β- globin gene, 
which is expressed only in 
erythroid cells. DNa replication 
properties of this domain have 
been extensively studied as it 
replicates early in erythroid 
cells and late in other 
cell types.
Ribosomal DNA
(rDNa). Part of the genome 
coding for ribosomal RNa. 
rDNa is constituted of several 
copies of the same genes, the 
expression of each varying 
depending on cell type.
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replication bubble structures)128, Okazaki fragments 
(OK­ seq; nascent, EdU­ labelled, single­ stranded DNA 
fragments smaller than 250 bp)129 or initiation sites 
(ini­ seq; first nucleotides to replicate at the onset of 
S phase)130 — or by chromatin immunoprecipitation 
of pre­ replication complex proteins (ChIP­ seq)131,132. 
The main observations are that the mapped sites are 
often associated with G­ quadruplexes (secondary 
structures formed by guanines)127,130,133, high GC den­
sity134,135 or a nucleotide distribution asymmetry, with 
a bias in G/C or in A/T content, with different features 
often found at different origins136. Despite these differ­
ences in nucleotide content, no origin­ specific sequence 
has been identified in eukaryotes other than in budding 
yeast137. There is great flexibility in the sites at which 
replication can initiate (box 2) and any DNA sequence 
can function as an origin in the right context138.
The relationship of origins to replication timing in 
mammalian cells remains elusive. The density of replica­
tion origins is generally higher in open early­ replicating 
chromatin128–132. However, replication domains that 
switch replication timing during development (approxi­
mately 50% of the genome) have the same origin density 
as constitutively late­ replicating regions, even when they 
replicate very early during the S phase100,127. Moreover, 
there is ample evidence to indicate that regu lation of 
DNA replication timing acts upstream of the machin­
ery that regulates where replication initiates18. This is 
also true in budding yeast, where factors binding to 
sites nearby but separable from origins influence repli­
cation timing139,140. However, in mammalian cells, the 
regulation is likely at the level of entire domains rather 
than individual origins and, even before the discovery of 
ERCEs, there was evidence that specific DNA sequences 
do play a role in replication timing. For example, ectopic 
insertion of the β­ globin locus control region (LCR) 
advanced local replication timing141,142, demonstrating 
that exogenous DNA sequences can modify replication 
timing. In a mouse strain carrying a copy of human 
chromosome 21, the exogenous human chromosome 
conserved its species­ specific replication timing in vari­
ous mouse tissues, suggesting that the DNA sequence 
of the human chromosome dictates its characteris­
tic pattern of replication timing143. To identify DNA 
sequences that potentially control replication timing, 
asynchronously replicating regions were identified by 
constructing separate replication timing profiles for each 
chromosome homologue in human erythroblasts, and 
these regions were then linked to large structural varia­
tions107. Consistent with the independent regulation of 
origin sites and timing of origin firing, these regions 
of asynchrony, although rare, could be linked to changes 
in origin efficiency, but not to the use of different ori­
gins136. A similar study identified rare replication timing 
quanti tative trait loci as clusters of single­ nucleotide poly­
morphisms and short indels linked to replication timing 
differences in human individuals144. These observations 
suggest a role for specific short DNA sequence alter­
ations in the regulation of replication timing, in addition 
to epigenetic mechanisms. One prediction from these 
results is that crosses between more distantly related 
individuals should enhance asynchronous replication 
in the F1 hybrid offspring. However, an extensive study 
of cells derived from crosses between Mus musculus 
castaneus and Mus musculus musculus in different paren­
tal configurations, although clearly identifying replica­
tion timing asynchronies linked to subspecies genome 
(Fig. 3c), did not detect an enhanced degree of replica­
tion asynchrony, which was cell­ type specific and varied 
from 1% to 12% in different cell types108. These results 
support a role for DNA sequences that is modulated by 
epigenetic events occurring during development.
Altogether, there is substantial evidence in support 
for both genetic and epigenetic having roles in regulat­
ing DNA replication timing. Interestingly, as replication 
timing can affect histone acetylation and genetic muta­
tion frequency22,23, there is also evidence for replication 
timing affecting the epigenetic status and DNA sequence 
composition.
Cis and trans regulation of replication timing
Several elements are known to contribute to the regula­
tion of DNA replication timing, acting either in cis, such 
as ASARs and ERCEs, or in trans, such as factors that 
promoteor inhibit replication initiation (box 2). First, 
at the chromosome level, replication timing could require 
the presence of single long interspersed nuclear element 1 
(LINE1), in the antisense orientation, within long non­ 
coding ASARs, which are present in multiple copies 
on each chromosome, as observed on human chromo­
some 6 (ReFS41–43). Although expressed from only one 
transcription unit on one of the two homologous chro­
mosomes, ASARs identified on human chromosomes 6 
and 15 associate in cis with the entire chromosome, 
forming an ‘RNA cloud’ that coats the entire chromo­
some (Fig. 4a), but if the LINE1 element within the 
ASAR is deleted or inverted, replication of the entire 
chromosome is severely delayed. Second, early repli­
cation of individual domains requires the presence of 
multiple ERCEs that interact in 3D space and, when all 
ERCEs within a domain are deleted, the entire domain 
replicates very late in the S phase75. Finally, trans regu­
latory elements comprise the factors directly involved in 
the regulation of origin firing, allowing or inhibiting the 
activation of the replicative helicase (box 2).
lncRNAs that coordinate autosomal DNA synthesis. 
The inactive Xi chromosome in female mammals is 
replicated with considerable delay compared with the 
active X chromosome. This delay is due to the expres­
sion of the Xist gene from the Xi chromosome, which 
then coats the Xi in cis. However, Xist gene deletion in 
a differentiated cell line leads to an increase in the delay 
of the replication of the Xi chromosome61. This obser­
vation has been largely ignored by the research commu­
nity until recently, when lncRNAs with similar deletion 
phenotypes to Xist — named ASARs — were found 
to be expressed from human autosomes 6 and 15 and 
coat the chromosome from which they are expressed in 
cis41–43. Like Xist RNA145, ASARs also coat the chromo­
some from which they are expressed (Fig. 4a) when they 
are ectopically expressed and can delay the replication 
timing of the chromosome into which they are inserted. 
Also similar to Xist, loss of ASAR expression from the 
Indels
insertions or deletions in 
the genome of a cell or an 
individual.
Long interspersed nuclear 
element 1
(liNe1). Class of transposable 
elements estimated to be 
present at around 500,000 
copies in the human and 
mouse genomes.
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endogenous chromosome leads to a drastic delay in 
replication of the entire chromosome42,43. In some cases, 
the chromosome continues to be replicated into mito­
sis and fails to condense properly, probably because 
condensation requires completion of replication41,42. 
On human chromosome 6, maintenance of a normal 
replication timing programme was found to be depen­
dent on a single LINE1 element within the ASAR6 gene. 
Normal timely chromosome 6 replication required anti­
sense expression of this LINE1; deletion or inversion of 
the LINE1 resulted in delayed replication of the entire 
chromosome. When inserted ectopically into a differ­
ent chromosome, ASAR6 suppressed the expression of 
endogenous LINE1 across the entire chromosome, sug­
gesting that ASAR6 suppresses expression of the endog­
enous ASARs43. Interestingly, ASARs reside in regions of 
chromosomes that are subject to random asynchronous 
replication and are expressed mono­ allelically from the 
allele that replicates later41,42. The molecular mechanisms 
by which ASARs function remain to be elucidated, but 
a working hypothesis is that homologous chromosomes 
express different ASARs, which coat the chromosome 
they are expressed from to ensure the synchronous repli­
cation of homologous chromosomes43. Thus, in this 
model, ectopic expression suppresses the resident ASAR, 
explaining why both deletion of the resident ASAR and 
ectopic expression of a second ASAR give rise to the 
same phenotype.
The search for cis elements that regulate origin firing. 
The discovery of cis­ acting lncRNAs that coordinate the 
synthesis of entire autosomes during the cell cycle was 
entirely unexpected; however, cis elements that regu­
late the firing of replication origins have been sought 
for decades. Early yeast studies indicated that origins 
replicated at times dictated by their chromosomal con­
text146,147. However, the mechanisms of action of the yeast 
elements remained elusive and attempts to identify simi­
lar elements in higher eukaryotes have failed for a long 
period of time. In 1990, much excitement followed the 
discovery that cells from patients with Hispanic thal­
assaemia harboured a naturally occurring 35­kb dele­
tion removing most of the human β­ globin LCR. This 
chromosomal deletion was associated with the loss of 
β­ globin gene transcription, the gain of closed chro­
matin structure and a shift to late replication across the 
β­ globin locus in erythroid cells148,149. Ten years later, 
targeted deletion of the human LCR was achieved. This 
deletion led to the loss of β­ globin gene transcription, 
but the domain still switched to early replication and 
open chromatin during erythroid cell differentiation150. 
The lack of effect of this deletion on replication timing 
at the native locus was never resolved and this setback, 
coupled with the failure to identify other cis elements 
regulating replication timing, led many to speculate that 
replication timing may be regulated exclusively by epi­
genetic mechanisms, as suggested by the observed ran­
dom and imprinted allelic differences (discussed above), 
or by complex sequence features that are very diffi­
cult to identify by chromosome engineering methods 
(for example, AT content and G4 quadruplex density).
This longstanding open question was resolved by 
recent studies using CRISPR–Cas9 to generate a large 
series of deletions and inversions in a replication domain 
in mouse ESCs, which revealed that replication timing, 
transcription and the 3D architecture of the domain 
are under the control of defined cis­ acting elements 
termed ERCEs (discussed above)75 (Fig. 4b). A single 
ERCE has partial activity to advance replication timing, 
but at least two ERCEs are required for replication early 
in the S phase.
ERCEs resemble super­ enhancers as they contain 
broad domains of acetylated histones (mainly H3K27ac) 
and multiple sites that are co­ occupied by master tran­
scription factors Oct4, Sox2 and Nanog (OSN) as well 
as by the histone acetyltransferase p300. Interestingly, 
ERCEs interact with each other in 3D but are not bound 
by CTCF and cohesin, which are the major known play­
ers in mediating chromatin interactions73,151. On the basis 
of these properties, ERCE loci were predicted genome­ 
wide, and deletions of several predicted ERCEs further 
validated their existence75. Moreover, deletion of ERCEs 
Chromosomea
b Hub of ERCEs
c
ASAR RNA ASAR locus
MCM2–7
Early-replicating chromatin
H3K27Ac
hub
Master transcription factors
Late-
replicating
chromatin
ORC
Cdc6 Cdt1
TopBP1
MCM10
GINS
Cdc45
Treslin
Treslin
S
DDK, CDK
Sirt1 PP1 Rif1
Fig. 4 | Cis- and trans- acting elements regulating replication timing. a | Long non- 
coding RNA asynchronous replication and autosomal RNAs (ASARs) are expressed from 
one locus specific to each homologous chromosome, and coat the whole chromosome 
from which they are expressed. This coating is necessary to define the temporal window 
during which DNA replication can occur on each chromosome43. b | Early- replication 
control elements (ERCEs) are sequences that are found to interact with each other within 
a megabase- sized domain, forming a cluster or ‘hub’ of ERCEs. They are large domains of 
acetylated histones and are bound by master transcription factors. ERCEs are necessary 
for the early replication of the chromosome domain in which they reside75. c | Replication 
origins are activated by recruitment of S- phase promoting factors (SPFs) such as the 
kinases Dbf4-dependent kinase (DDK) and cyclin- dependent kinase(CDK), and the 
protein Treslin, and inhibited by factors such as Rif1 and the deacetylase Sirt1 (see also 
box 2) that inhibit SPFs. All known trans- acting regulators of replication timing act on 
DDK, CDK or Treslin but potentially other such regulators may exist that act upon other 
SPFs such as TopBP1, Cdc45, Mcm10 and GINS.
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results in a domain­ wide switch from A to B Hi­ C com­
partments, disruption of local TAD architecture and 
loss of all transcription throughout the domain75. These 
results show that replication timing, spatial chromatin 
organization and transcription are intimately linked and 
co­ regulated. They also provide an explanation for the 
enigmatic results of the past: the β­ globin LCR may be 
one of several redundant erythroid­ activated ERCEs in 
the region of the β­ globin domain that are missing in the 
Hispanic thalassaemia deletion.
Trans- acting regulators of replication timing recruit 
or antagonize essential initiation factors. Replication 
is initiated when the cyclin­ dependent kinases152,153, 
the Dbf4­dependent kinase Cdc7­Dbf4 (DDK)154,155, the 
protein complex GINS156 and the initiation factors 
Treslin157,158, Mcm10 and Cdc45 (ReF.156) cooperate to 
activate a subset of pre­ loaded MCM helicases, known 
as pre­ replication complexes (pre­ RCs) (box 2, Fig. 4c). 
The remainder of the pre­ RCs are either activated later 
in S phase or remain dormant and are removed when 
a replication fork passes through them (in vertebrates, 
the vast majority remain dormant).
Most of our knowledge of trans­ acting factors that 
can control replication timing comes from studies in 
yeasts, where the common function is to promote or 
inhibit pre­ RC activation. For example, the fission yeast 
telomere binding factor TAZ1 impairs DDK activation 
during early S phase at a set of late origins by blocking 
the recruitment of Sld3 (the homologue of mammalian 
Treslin) to the pre­ RC, thus preventing origin firing 
until late S phase159. In budding yeast and fission yeast, 
pericentromeric heterochromatin is replicated early, 
following the recruitment of Dbf4 by Swi6 (the homo­
logue of mammalian HP1) in fission yeast160 or Ctf19 
(the homologue of CENP­ P) in budding yeast161. In bud­
ding yeast, the forkhead 1 (Fkh1) and Fkh2 transcrip­
tion factors recruit Dbf4 to a set of early­ firing origins139. 
In mammalian cells, phosphorylated SIRT1 prevents 
dormant origin firing162, possibly through its ability to 
deacetylate and destabilize MCM10 (ReF.163). Finally, the 
chromatin­ binding factor RIF1 has a conserved role in 
replication timing control from yeast to humans164–166. 
In yeast, D. melanogaster, mouse and human, RIF1 has 
been shown to interact with the PP1 phosphatase167–170 
(Fig. 4c), which has the ability to dephosphorylate the 
MCM complex at its Cdc7­dependent phosphoryla­
tion sites168, suggesting that RIF1 may delay DNA repli­
cation initiation by antagonizing phosphorylation by 
Cdc7 (ReF.168). However, in budding yeast, RIF1 binds 
to DNA in the proximity of early­ and late­ replicating 
origins and may affect the activity of both171. Moreover, 
in mammalian cells, PP1 recruitment by RIF1 is also 
required for pre­ RC formation167. In mouse ESCs, Rif1 is 
broadly enriched in late­ replicating chromatin, but Rif1 
depletion has effects on both early and late replication166. 
Overall, it seems that, in many cases, regulation of repli­
cation timing involves either promoting or antagonizing 
the function of proteins that activate pre­ RCs.
In yeast, the ability of trans­ acting factors to advance 
or delay replication is often linked to the 3D coales­
cence of commonly regulated origins. TAZ1­regulated 
origins localize near telomeres during G1 and early 
S phase172, and various genetic manipulations in this study 
indicated that the delayed initiation of DNA repli cation 
from these origins is dependent on their spatial prox­
imity in the 3D nuclear space172. In addition, Fkh1 and 
Fkh2 transcription factors mediate the 3D clustering of 
a set of early origins173, and the ability of Fkh1 and Fkh2 
to dimerize is essential for origin clustering and early 
replication independent of its ability to activate tran­
scription140. The fact that mammalian ERCEs were 
identified by virtue of their 3D interactions could sug­
gest that they function similarly. It is possible that spa­
tial clustering promotes the formation of subnuclear 
domains that concentrate replication initiation factors. 
Human Treslin was recently shown to interact with Brd4 
(ReF.174), a bromodomain protein with high affinity for 
acetylated histones. As ERCEs are characterized by the 
presence of large stretches of H3K27Ac, this suggests a 
clear link between ERCE clustering and the formation 
of Treslin­ rich domains that promote pre­ RC activation.
Replication timing and transcription
The positive correlation between early replication and 
transcriptional activity in higher eukaryotes is well 
known4–8,31,175–179. However, several studies indicate that 
the two processes can be uncoupled in many contexts 
and are almost certainly indirectly related. Indeed, most 
genes that switch replication timing in mammals can be 
expressed when late replicating32. ASAR lncRNA genes 
are asynchronously replicated and expressed from the 
late­ replicating allele41. In yeast, no correlation between 
the level of transcription and the timing of replication 
has been observed180. Moreover, although budding yeast 
Fkh1 and Fkh2 transcription factors are required for 
early replication of a large number of origins, mutations 
that impair their ability to dimerize without affecting 
their transcription regulatory function delay early origin 
firing without affecting transcription140,173.
In mammals, several studies have attempted to 
establish a causality between these two processes, but 
the results have been difficult to reconcile. A study in 
ESCs showed that targeting a strong transactivator to a 
late­ replicating site led to transcription activation and 
re­ localization of that chromosomal region towards 
the interior of the nucleus, whereas targeting a related 
peptide that could cause re­ localization but not tran­
scription did not anticipate DNA replication178. This 
led the authors to conclude that transcription activation 
is sufficient to advance replication timing178. However, 
studies deleting the β­ globin LCR have shown that dis­
rupting transcription at this locus does not prevent the 
switch to early replication of the β­ globin locus during 
erythropoiesis117,150,181. Moreover, targeting a histone 
acetyltransferase to the β­ globin locus in non­ erythroid 
cells advanced replication timing whereas transcription 
of the β­ globin gene remained low120. There is some 
evidence in support of the idea that transcriptional 
activity and the transcript length might have to reach a 
minimal threshold for it to have an effect on replication 
timing182. At the DppA2/4 locus in mouse ESCs, deletion 
of all ERCEs within a domain had an impact on both 
replication timing and transcription, but other deletions 
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732 | DeCemBeR 2019 | volume 20 
inactivated transcription with no changes in replication 
timing, or substantially delayed replication timing with 
no changes in transcription75. Overall, these studies 
show that transcription activation can be associated 
with earlier replication, but not systematically, reinfor­
cing the hypothesis of an indirect relationship between 
these two processes.
Genome­ wide analyses reporting widespread cor­
relation between transcription and replication timing 
in the context of cell differentiation have also provided 
abundant examples of uncoupling between transcription 
and replication timing. In human and mouse cells, it is 
the constitutively early­ replicating genes — which are 
early replicating in every cell type andconstitute two­ 
thirds of all genes — that drive the genome­ wide cor­
relation of early replication with transcription because 
they are, by definition, always early replicating when 
expressed. Very few genes are constitutively late repli­
cating. By contrast, the majority of genes for which the 
timing of replication changes during differentiation 
(approximately one­ third of genes) can be found to 
be transcribed in one or more cell types in which they 
are late replicating31,32. Moreover, the global correlation 
between replication timing and gene expression tends to 
decrease during the early stages of human stem cell fate 
commitment and differentiation32. Recently, taking close 
time points during the first few cell divisions of human 
ESCs undergoing a cell fate transition, many replication 
timing changes were found to be independent of changes 
in gene expression87, albeit often being only a temporary 
uncoupling. Finally, in early animal embryos undergoing 
rapid cleavage divisions, a DNA replication timing pro­
gramme is established prior to the onset of transcription 
at the mid­ blastula transition90,91,183,184.
A recent study assembling novel ‘replication timing 
networks’ provides a potential resolution to some of 
these complex observations. This study linked tran­
scriptional changes that are coordinated with replication 
timing changes across many human ESC lines differen­
tiating towards different lineages. Sets of genes were 
identified whose expression predicted early replication 
of a separate, unlinked, set of replication domains185. 
These genes almost exclusively encoded transcription 
factors. When ChIP data for these transcription factors 
were analysed, remarkably, all of them were found to 
co­ occupy specific sites in the affected domains. This is 
reminiscent of the observation that transcription factors 
of the major pluripotency transcription­ related network 
all bind together to ERCEs75. From these observations, 
we suggest that the imperfect correlation between repli­
cation timing and transcription could result from early 
replication being regulated by combinations of transcrip­
tion factors, independently of their roles in transcription. 
Perhaps replication is influenced by transcription­ related 
network transcription factors that, like Fkh1 and Fkh2 
in budding yeast140, have separable roles in transcription 
and replication timing. Altogether, the most plausible 
conclusion from these studies is that transcription and 
replication timing are regulated by closely related but 
separable mechanisms.
Conclusion and perspectives
Studies using multiple model systems in the past few 
years have provided numerous new insights into the 
regulation of DNA replication timing and its relationship 
to 3D genome architecture that promise to fuel rapid 
future progress in this research field. We have discussed 
here how the timing of replication reflects chromosome 
architecture: early­ and late­ replicating chromatin coin­
cides with A and B compartments, respectively, whereas 
replication domains are stable units of replication cor­
responding to a certain category of TADs. Coordinated 
firing of DNA replication origins within defined repli­
cation domains occurs at characteristic times that are 
conserved between homologous chromosomes and 
between individual cells. Replication timing is regulated 
at the level of these replication domains, independently 
of known 3D genome architectural factors or bound­
aries. Instead, association of cis­ acting elements with 
transcriptional regulatory factors and their interaction 
in 3D space may establish subnuclear environments that 
recruit replication initiation factors. The independence 
of their roles in transcription versus 3D clustering (Fig. 5) 
could explain, in part, the imperfect correlation between 
transcription and replication timing. For example, the 
super­ enhancer nature of ERCEs could create platforms 
for their self­ association and recruitment of replication 
initiation proteins such as Treslin and DDK. A single 
ERCE may create a weak platform that can advance 
replication timing slightly, whereas additional ERCEs 
A compartment
(attracts SPFs)
B compartment
(antagonizes SPFs)
Cytosol Nuclear envelope
Nucleolus
Late-replication
associated factor
(Rif1)
Early-replication
associated factors
(Brd2–Brd4)
Lamina
Late-replicating chromatin
ERCEs
Early-replicating chromatin
ASAR
H3K27Ac
Fig. 5 | Proposed model of organization within the nucleus. Model for the compart-
mentalization of the nucleus into and early- and late- replicating chromatin regions. In 
this model, early- replicating regions, within the A compartment, contain elements rich in 
H3K27Ac, bound by factors such as Brd2 and Brd4 that recruit S- phase-promoting factors 
(SPFs; see Fig. 4) such as Treslin. These elements, called early- replication control elements 
(ERCEs), form a platform for recruiting SPFs within the domain in which they belong. 
In this model, the replication of late- replicating regions, within the B compartment and 
associated with the nuclear lamina and the nucleolus, would be delayed by factors 
inhibiting origin firing (that is, antagonizing SPFs), such as Rif1 (which antagonizes DDK). 
At the chromosome level, coating of each chromosome by long non- coding RNAs known 
as asynchronous replication and autosomal RNAs (ASARs) could ensure the presence 
of replication machinery factors within each chromosome territory.
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synergize by interacting to create a strong platform for 
recruitment, potentially through phase separation186. 
Likewise, factors that create transcriptionally repressed 
subnuclear environments may recruit factors that antag­
onize initiation proteins and delay replication. At the 
chromosome level, ASARs coat the entire chromosome 
to set the overall temporal window of replication dur­
ing the S phase. Their mechanism of action and their 
relationship with ERCEs remain to be elucidated.
The recent discovery of ERCEs and ASARs and the 
recently available tools to dissect their structure and 
underpinning molecular mechanisms will fuel many 
future discoveries. First, both ERCEs and ASARs are 
expected to be widespread elements, and the systematic 
identification of such elements, at the genome­ wide level, 
is an important and achievable goal. Second, ERCEs will 
be dissected molecularly to identify critical sequences 
and, ultimately, trans­ acting binding partners, while 
proteins and RNAs that interact with ASARs will be 
identified. The question of whether ASARs delay entire 
chromosomes without disrupting the temporal order of 
replication will be addressed, and ultimately whether 
ASARs and ERCEs cooperate or whether ASARs are a 
global chromosome­ wide threshold to ERCE activity. 
Third, our model predicts that there are different ERCEs 
in different tissues and that deletion of ERCEs specific 
to one cell type will have no effect on early replication 
in other cell types. Ultimately, artificially creating the 
individual components of ERCEs by targeting proteins 
that can induce chromatin loops, histone acetylation 
as well as manipulations that separate the functions of 
transcription versus chromatin replication of transcrip­
tion factors, such as those recently initiated in budding 
yeast, will dissect the mechanisms regulating repli­
cation timing and how it is related to transcription and 
3D organization.
Published online 2 September 2019
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