Download Regulation of DNA replication during development

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Development 139, 455-464 (2012) doi:10.1242/dev.061838
© 2012. Published by The Company of Biologists Ltd
Regulation of DNA replication during development
Jared Nordman and Terry L. Orr-Weaver*
As development unfolds, DNA replication is not only
coordinated with cell proliferation, but is regulated uniquely in
specific cell types and organs. This differential regulation of
DNA synthesis requires crosstalk between DNA replication and
differentiation. This dynamic aspect of DNA replication is
highlighted by the finding that the distribution of replication
origins varies between differentiated cell types and changes
with differentiation. Moreover, differential DNA replication in
some cell types can lead to increases or decreases in gene copy
number along chromosomes. This review highlights the recent
advances and technologies that have provided us with new
insights into the developmental regulation of DNA replication.
Key words: Amplification, Drosophila, Endo cycle, Mammalian
differentiation, ORC, Replication origins
Introduction
Every cell division cycle requires the faithful duplication of the
entire genome. In multicellular organisms, the rate of cell division
of cycling cells can vary greatly, ranging from minutes to hours,
depending on the developmental state of a particular cell within the
organism. Further compounding the challenges of genome
duplication is the epigenetic state of the genome, which is
constantly changing during development. Although these
requirements are daunting, the DNA replication machinery is
precisely regulated to ensure that genome duplication occurs within
the proper time frame without compromising genome stability.
It has been known for decades that the properties of DNA
replication are exquisitely sensitive to developmental states. For
example, in early Drosophila and Xenopus embryogenesis, the
duration of S phase is in the order of minutes, with replication
origins spaced <10 kb apart (Blumenthal et al., 1974; Hyrien and
Mechali, 1993). This is in stark contrast to fully differentiated cell
types in which S phase can last longer than ten hours, with origins
of replication >100 kb apart (Spradling and Orr-Weaver, 1987). The
developmental signals and underlying molecular mechanisms that
impact the parameters of DNA replication during development are
poorly understood. Recent technological advances combined with
genome-wide approaches have given a more precise view of the
changes that development imparts on the DNA replication program,
providing a foundation from which to unravel the regulatory
signals.
Here, we briefly summarize the mechanism of initiation of DNA
replication and detail recent discoveries showing that mutations in
key replication initiation proteins cause developmental defects in
humans. We then examine how differentiation affects DNA
replication, both the sites of replication origins and the timing of
when origins are activated during genomic DNA replication.
Whitehead Institute and Department of Biology, Massachusetts Institute of
Technology, Cambridge, MA 02142, USA.
*Author for correspondence ([email protected])
Finally, we present recent studies on the mechanism by which
DNA copy number is altered through differential DNA replication
during development.
Assembly and activation of the pre-replicative
complex
DNA replication initiates at cis-acting sites in the genome termed
origins of replication (see Glossary, Box 1). For every round of
DNA replication, thousands of replication origins are utilized in a
coordinated manner. In metazoans, origins of replication are
sequence-independent, highly influenced by chromatin structure
and often change during the course of cell differentiation. The
proteins involved in initiating DNA synthesis are conserved from
yeast to humans (Table 1). The formation of replication complexes
Box 1. Glossary
Amplicon. A defined region of increased gene copy number, either
in a somatic cell or stably inherited in the germline.
Chorion. The eggshell surrounding an egg; composed of chorion
proteins.
Endo cycle. A cell cycle in which cells undergo repeated S and G
phases with no intervening mitoses.
Gene amplification. Increased gene copy number relative to
overall ploidy.
Imaginal disc. A group of cells present in larvae that will form an
adult organ such as eye, leg or wing; contains groups of
determined diploid cells that divide during larval stages and
differentiate into adult tissues during pupation.
Meirer-Grolin syndrome (MGS). A form of primordial dwarfism
characterized by several developmental abnormalities, such as short
stature, small ears and absent or underdeveloped patellae.
Origin of replication. Cis-acting DNA sequence where replication
initiates. Origins of replication are sequence independent in
metazoans.
Origin recognition complex (ORC). Six-subunit complex (Orc1Orc6) that binds to origins of replication.
p19 cell line. A mouse pluripotent cell line derived from an
embryonic carcinoma; can be induced to differentiate into multiple
cell types.
Polyploidy. Having increased genomic content, i.e. more than two
paired sets of chromosomes; occurs as a result of repeated S-G
cycles with no intervening mitoses.
Polytene. Polyploid cells in which the sister chromatids are held in
tight association.
Pre-replicative complex (pre-RC). Complex of Orc1-Orc6, Cdc6,
Cdt1 and a double hexamer of the Mcm2-7 complex. The pre-RC
refers to the loaded but inactive form of the replicative helicase.
Replication timing. The relative time during S phase when a
specific genomic region is replicated.
Rothmund-Thomson syndrome (RTS). A rare autosomal
recessive disorder characterized by several abnormalities including
a facial rash, short stature, skeletal abnormalities, premature aging,
chromosome fragility and a predisposition to tumor formation.
Under-replication. Reduced gene copy number relative to overall
ploidy of a cell.
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Summary
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Development 139 (3)
Table 1. Components necessary for helicase loading and helicase activation
Budding yeast
Orc1
Orc2
Orc3
Orc4
Orc5
Orc6
Cdc6
Cdt1
Mmc2
Mcm3
Mcm4
Mcm5
Mcm6
Mcm7
Mcm10
Cdc45
Sld5
Psf1
Psf2
Dpb11
Sld2
Sld3
Metazoan homolog
Function
ORC1
ORC2
ORC3 (Latheo in Drosophila)
ORC4
ORC5
ORC6
CDC6
Cdt1 (DUP in Drosophila)
MCM2
MCM3
MCM4 (DPA in Drosophila)
MCM5
MCM6
MCM7
MCM10
CDC45
SLD5
PSF1
PSF2
TopBP1 (MUS101 in Drosophila)
RECQL4
Treslin/ticcr (only in vertebrates)
Helicase loading
Helicase loading
Helicase loading
Helicase loading
Helicase loading
Helicase loading
Helicase loading
Helicase loading
Helicase
Helicase
Helicase
Helicase
Helicase
Helicase
Helicase activation
Helicase activation/Helicase
Helicase activation/Helicase
Helicase activation/Helicase
Helicase activation/Helicase
Helicase activation/Helicase
Helicase activation
Helicase activation
Nomenclature is based on S. cerevisiae or Drosophila melanogaster unless otherwise indicated. For further information regarding the function of replication subunits involved
in helicase loading see Bell and Dutta (Bell and Dutta, 2002). For more information on the replication subunits involved in helicase activation see Araki (Araki, 2010).
loading, but also origin firing by the activation of the Mcm27/Cdc45/GINS complex, which is the active form of the replicative
helicase (Ilves et al., 2010; Moyer et al., 2006).
Once replication has initiated, multiple mechanisms exist to
ensure that DNA replication occurs once per cell cycle. As S phase
progresses, S-CDK activity steadily increases and CDK-dependent
phosphorylation of ORC subunits inhibits the further assembly and
activation of pre-RCs (Arias and Walter, 2007; Chen and Bell,
2011). In metazoans, Cdt1 is degraded and the Geminin protein
binds to and inhibits Cdt1, thereby preventing further recruitment
of the Mcm complex to sites of DNA replication (Havens and
Walter, 2009; McGarry and Kirschner, 1998; Wohlschlegel et al.,
2000). Failure to inhibit re-replication can result in gene
amplification and have deleterious effects on genome stability
(Davidson et al., 2006; Green et al., 2010). Surprisingly, Geminin
not only controls DNA replication through Cdt1 but also plays
crucial developmental roles in Xenopus and mouse embryos by
inhibiting lineage commitment or differentiation (Lim et al., 2011;
Yang et al., 2011).
Mutations in pre-RC components cause
developmental abnormalities
The differentiation of particular cell types and the development of
organs are uniquely dependent on the proper regulation of DNA
replication, and a number of studies have shown that mutations in
pre-RC subunits can give rise to specific developmental phenotypes.
In Drosophila, latheo was initially identified in a genetic screen for
olfactory learning and memory mutants (Boynton and Tully, 1992).
Surprisingly, the latheo gene was shown to encode ORC3 (Pinto et
al., 1999). Subsequent analysis showed that latheo mutants exhibit
disrupted cell proliferation within the imaginal discs (see Glossary,
Box 1) and the central nervous system (CNS), with the strongest
alleles affecting all imaginal discs and the late larval CNS, whereas
hypomorphic mutants affect the adult brain. Remarkably,
hypomorphic latheo mutants display a reduction in overall volume
of a specific anatomical region of the adult brain known to be
DEVELOPMENT
at origins of replication occurs in two phases: helicase loading or
pre-replicative complex (pre-RC, see Glossary, Box 1) assembly;
and helicase activation/replisome assembly. Helicase loading starts
when the origin recognition complex (ORC, see Glossary, Box 1)
binds to an origin of replication in late M and G1 phases of the cell
cycle (Fig. 1A). ORC consists of six subunits (Orc1-Orc6), five of
which are related to a large family of ATPases, the AAA+ ATPases
(Bell and Dutta, 2002). Together with two additional replication
factors, cell division cycle 6 (Cdc6) and cdc10-dependent transcript
1 (Cdt1), ORC loads a double hexamer of the minichromosome
maintenance proteins 2-7 (Mcm2-Mcm7) onto DNA (Remus et al.,
2009) (Fig. 1A).
Pre-RC complexes are ‘licensed’, or poised for replication, but
are incapable of initiating DNA synthesis because the helicase is
loaded in an inactive state. Activation of the helicase requires both
S phase-specific cyclin-dependent and Dbf4-dependent kinases (SCDK and DDK, respectively) (Fig. 1B). ORC binding and pre-RC
formation occur in late M and G1 phases of the cell cycle when SCDK activity is low. For helicase activation, S-CDK and DDK
phosphorylation events must occur (Arias and Walter, 2007). In
budding yeast, DDK phosphorylates multiple subunits of the
Mcm2-Mcm7 complex (Randell et al., 2010; Sheu and Stillman,
2010). Also in budding yeast, two proteins required for activation
of the helicase, Sld2 and Sld3, are the only S-CDK targets required
for the initiation of DNA replication (Tanaka et al., 2007;
Zegerman and Diffley, 2007). Once phosphorylated, Sld2 and Sld3
recruit the BRCT repeat-containing protein Dpb11 (TopBP1 in
metazoans) to potential origins of replication. Homologs of both
Sld2 (RECQL4) and Sld3 (Treslin, ticrr) have been identified in
metazoans and recent work indicates that the S-CDK-dependent
interaction between Sld3/Treslin and Dbp11/TopBP1 is conserved
(Boos et al., 2011). Dpb11/TopBP1 is necessary for the recruitment
of Mcm10, Cdc45 and the GINS complex [Sld5 (Gins4), Psf1
(Gins1), Psf2 (Gins2) and Psf3 (Gins3)], which result in CMG
complex formation and activation of the helicase (Fig. 1C).
Together, DDK and S-CDK activity regulate not only helicase
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A Helicase loading (pre-RC assembly)
Open/active
chromatin
Repressed
chromatin
B Kinase activation
Repressed
chromatin
DDK
Phosphorylation
Activation
S-CDK
Recruitment of
additional subunits
C Helicase activation
Key
ORC
Cdc6
Sld3
Dpb11
Mcm2-7
Cdt1
Sld2
Cdc45-GINS
Fig. 1. Helicase loading (pre-RC assembly) and activation.
(A)Helicase loading (pre-RC assembly) at a potential origin of
replication coinciding with a transcription start site (arrow). Although
origins are sequence independent in metazoans, histone modifications
are associated with transcriptional activation, and open/active
chromatin correlates with ORC binding. Histone modifications
associated with repressive chromatin are negatively associated with
ORC binding. During helicase loading, binding of the ORC complex
promotes recruitment of Cdc6, which in turn promotes the loading of
a Cdt1–Mcm2-7 double hexamer. (B)Kinase activation. DDK promotes
helicase activation by phosphorylating multiple Mcm subunits. S-CDK
promotes helicase activation by phosphorylating Sld2 and Sld3, which
in turn allow the recruitment of additional factors necessary for
helicase activation. (C)Helicase activation is dependent on the
recruitment of additional factors necessary to activate the replicative
Mcm2-7 helicase, which is a complex of Mcm2-7, Cdc45 and GINS
proteins (Sld5, Psf1, Psf2 and Psf3). Arrows indicate the direction of
helicase movement.
involved in olfactory learning, the mushroom body, potentially
explaining how a subunit of the ORC complex could have such a
specific function in learning and memory (Pinto et al., 1999). It is
still unknown, however, why cells within distinct regions of the adult
brain are susceptible to a reduction in the amount of ORC3. One
possibility is that the rate of cell division is not uniform throughout
the adult brain. Regions of the brain that require a faster cell division
cycle would, therefore, place a higher demand on replication
components and be more susceptible to reduced levels of ORC3.
This is consistent with the observation that developmental control of
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S phase length is modulated by the number of origins used, rather
than changes in the rate of replication fork progression (Blumenthal
et al., 1974).
Recent studies on Meirer-Gorlin syndrome (MGS; see Glossary,
Box 1) highlight further the importance of proper DNA replication
during human development (Bicknell et al., 2011a; Bicknell et al.,
2011b; Guernsey et al., 2011). MGS is a form of primordial
dwarfism characterized by several developmental abnormalities
(Gorlin, 1992). MGS follows an autosomal recessive mode of
inheritance with differing degrees of severity and until recently no
genes or loci had been found to be associated with this syndrome.
A series of elegant studies, however, has now identified mutations
in several pre-RC components as the cause of MGS.
Through single nucleotide polymorphism (SNP) mapping and
direct sequencing of candidate genes in affected individuals,
mutations in ORC1, ORC4, ORC6, CDT1 or CDC6 have been
identified as the cause of MGS (Bicknell et al., 2011a; Bicknell et
al., 2011b; Guernsey et al., 2011). The identification of mutations
within multiple pre-RC subunits strongly implies that a defect in
DNA replication, rather than DNA replication-independent
functions of ORC (Hemerly et al., 2009; Sasaki and Gilbert, 2007),
is responsible for MGS. Consistent with this hypothesis, cell lines
derived from MGS patients with a mutation in ORC1 display a
prolonged G1 phase and slower progression through S phase,
probably owing to a failure in proper activation of replication
origins (Bicknell et al., 2011b). Furthermore, using a zebrafish
model system, injection of a morpholino oligonucleotide that
targets orc1 resulted in a reduction in body size that correlated with
the degree of orc1 transcript level depletion. Similar results were
also obtained with a morpholino oligonucleotide that targets mcm5,
supporting the notion that failure to activate replication origins
properly can cause developmental abnormalities associated with
body size (Bicknell et al., 2011b).
Mutations potentially affecting helicase activation can also give
rise to developmental abnormalities in humans, emphasizing the
importance of proper DNA replication for organismal development.
Rothmund-Thomson syndrome (RTS; see Glossary, Box 1) is an
autosomal recessive disorder characterized by several abnormalities
including short stature, skeletal abnormalities, premature aging and
chromosome fragility (Larizza et al., 2010). Mutations in RECQL4,
the metazoan homolog of yeast Sld2, are responsible for ~66% of
RTS cases (Kitao et al., 1999; Larizza et al., 2010). RECQL4
functions in initiation of DNA replication and DNA repair.
Therefore, it is not possible to assess which symptoms of RTS
result from a failure to properly initiate DNA replication versus
those that are due to a defect in DNA repair.
Understanding how DNA replication and cell proliferation are
coordinated during development is a fascinating and challenging
area of developmental biology. Many highly proliferative cell and
tissue types are essential for proper organismal development, but
mutations affecting origin licensing must only affect a subset of
those cells and tissues in order to give rise to a specific set of
phenotypes.
The effects of differentiation on DNA replication
Potential origins of replication are subject to
developmental control
In metazoan cells, ORC binding is sequence independent, therefore
it is a challenge to identify origins of replication. Given the
plasticity of chromatin modifications and gene expression during
differentiation, origins have to be flexible to accommodate the
ever-changing chromatin landscape while faithfully replicating the
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Development 139 (3)
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genome within a precise developmental timeframe. One approach
to determining the location and properties of metazoan replication
origins has been to establish the location of all potential origins by
identifying the genome-wide binding profile of ORC. In
Drosophila, genome-wide ORC binding profiles (see Box 2) have
revealed that promoters of active genes, sites of open and active
chromatin, are enriched with ORC binding sites (Eaton et al., 2011;
MacAlpine et al., 2010). Furthermore, the frequency of ORC
binding sites is nearly three times greater in regions of the genome
that replicate early in S phase compared with those that replicate
late in S phase (MacAlpine et al., 2010). A more comprehensive
analysis of ORC binding sites in multiple Drosophila cell lines has
uncovered cell-type differences in ORC binding (Eaton et al.,
2011). Additionally, ORC binding has now been analyzed in a
primary tissue, the salivary gland of larvae (Sher et al., 2012). This
analysis revealed that ~30% of ORC binding sites are unique to the
salivary gland. Taken together, these data indicate that potential
origins of replication are subject to developmental regulation.
Defining origins of replication has been a rate-limiting factor in
studying the properties of origin firing in mammalian cells.
Recently, multiple techniques for mapping replication initiation
sites have been developed and adapted for genome-wide
identification (see Box 2); this is no small feat given that
replication products are short lived and extremely low in
abundance (Gilbert, 2010). Short nascent-strand analysis, for
example, relies on the isolation and purification of small DNA
fragments derived from leading strand replication products. Using
this approach, profiles of initiation sites have been generated for
0.4% of the mouse genome and for 1% of the human genome from
Box 2. Techniques used to study DNA replication
ORC binding profiles. Generated using chromatin
immunoprecipitation (ChIP) of ORC subunits followed by
hybridization of precipitated DNA to microarrays or highthroughput sequencing. ORC binding profiles can be used to
identify all potential replication origins on a genome-wide scale.
Short nascent strand analysis. Utilizes the characteristics of
newly synthesized DNA (i.e. size and presence of an RNA-DNA
hybrid at the 5⬘ end because of the RNA primer) to isolate small
leading strand fragments. Fragments can then be hybridized to a
microarray or directly sequenced to identify origins of replication.
Bubble-trap analysis. Restriction enzyme-digested genomic DNA
is mixed and slowly cooled in an agarose matrix, which selectively
traps replication bubbles as a result of the polymerization of
agarose through a replication bubble. Following electrophoresis,
bubbles recovered from the gel can be hybridized to a microarray
or directly sequenced to identify origins of replication.
Replication timing profiles. DNA is isolated at different times
during S phase (e.g. early S phase versus late S phase) followed by
copy number analysis to determine when a genomic region
replicates as a function of S phase progression.
Comparative genomic hybridization (aCGH). Hybridization to
microarray slides of genomic DNA derived from an experimental
tissue or cell type together with control DNA. Allows for the
analysis of copy number variation as well as genomic structural
aberrations, such as insertions and deletions.
Single molecule analysis of replicated DNA (SMARD). Direct
visualization of newly replicated single DNA molecules pulsed with
thymidine analogs. In vivo pulse-labeled DNA molecules are
extracted and stretched onto a glass slide for subsequent
fluorescence in situ hybridization (FISH) analysis of specific DNA loci.
This technique can be used to analyze both origin firing and
replication fork progression at a single molecule level.
Development 139 (3)
cell lines (Cadoret et al., 2008; Karnani et al., 2010; SequeiraMendes et al., 2009). Together, these studies have confirmed the
positive relationship between origins, transcriptional activity, and
markers of open chromatin. One caveat to this approach is that
<14% of origins identified by small nascent-strand analysis in
HeLa cells from two independent studies show overlap (Cadoret et
al., 2008; Karnani et al., 2010). The reasons for such low
concordance could be biological or technical, making future studies
necessary to address these disparities.
More recently, two genome-scale studies have used nascentstrand analysis (see Box 2) to map replication origins in two human
cancer cell lines, three mouse cell lines and a Drosophila cell line
(Cayrou et al., 2011; Martin et al., 2011). In all lines, the majority
of initiation events were associated with active transcription units.
But, in contrast to ORC-binding profiles in Drosophila (MacAlpine
et al., 2010), gene bodies rather than promoter regions of active
genes were enriched with initiation events (Cayrou et al., 2011).
Furthermore, both studies found that initiation events were underrepresented at the actual transcription start sites (TSS) and that
CpG-rich sequences were enriched with initiation events (Cayrou
et al., 2011; Martin et al., 2011).
An independent method to identify initiation sites entails
isolating early replication structures, or bubble structures, by the
bubble-trap method (see Box 2) (Mesner et al., 2006). Genomewide analysis of replication origins in mammalian cells by this
method demonstrated that replication origins are clustered within
zones and are associated with active and open chromatin (Mesner
et al., 2011). Furthermore, analysis of replication origins by the
bubble-trap method revealed cell-type specificity to origin usage.
Perhaps not surprisingly, analysis of origin usage by this method
also revealed that cell synchronization alters origin usage.
Numerous replication origins were identified in synchronized cells
that were not identified in unperturbed log-phase cells,
demonstrating the need for limited cell manipulation for accurate
assessment of replication properties (Mesner et al., 2011).
These studies in Drosophila and mammalian cell lines
demonstrate that there is tissue- and cell type-specific regulation
for which genomic sites serve as replication origins. The
association between active transcription and replication origins
suggests that the replication program is heavily influenced by the
transcription program that accompanies differentiation.
The timing of replication is developmentally controlled
Replication timing (see Glossary, Box 1) is a fundamental property
of genome duplication; some regions of the genome are replicated
early in S phase, whereas other regions replicate later in S phase
(Fig. 2). The significance of these timing differences remains
obscure. For example, replication timing could passively reflect the
chromatin state and, therefore, accessibility of replication factors
to chromosomal regions or domains. Alternatively, replication of
particular chromatin domains, such as heterochromatin, could
require the coordination of chromatin-modifying enzymes with
late-acting replication forks to propagate epigenetic information.
The activity of replication fork-associated chromatin-modifying
enzymes could be confined to the time in S phase when they are
needed to establish and propagate particular epigenetic states. This
might provide a mechanism to maintain the chromatin state
characteristic of each genomic region.
Recent advances in microarray and DNA sequencing
technologies have permitted rapid and cost effective identification
of genome-wide replication timing profiles (see Box 2) for
numerous cell types (Farkash-Amar et al., 2008; Hansen et al.,
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A Analyzing replication domains
S
G1
G2
Late
S phase
Cell number
Early
S phase
DNA Content
B A replication domain
Early
Late
Copy number
3 Mb
Late
replicating
Late
replicating
Early
replicating
Replication domain
Fig. 2. Replication domains encompass numerous potential
origins of replication. (A)Replication timing experiments monitor
changes in gene copy number or new DNA synthesis during multiple
intervals of S phase, defining zones of early or late replication. Typically,
cells can be separated into early (green box) and late (pink box)
replication fractions based on their DNA content. DNA isolated from
each fraction can be labeled and hybridized to a microarray, or directly
sequenced to monitor copy number changes during S phase.
(B)Replication domains typically extend from 200 kb to 2 Mb in size,
encompassing numerous individual origins of replication (diagram
shows the pre-RC as illustrated in Fig. 1). Consequently, populationscale replication timing experiments lack the resolution to monitor
changes in individual origin usage during differentiation.
2010; Hiratani et al., 2008; Macalpine et al., 2004; Pope et al.,
2010; Schübeler et al., 2002; Schwaiger et al., 2009). These studies
rely on the isolation of DNA from defined times during S phase
and the subsequent analysis of the twofold changes in copy number
diagnostic of a region having been replicated. Additionally, a more
recent method for monitoring the temporal order of replication
during S phase has been developed that relies on monitoring newly
synthesized DNA during S phase progression (Hansen et al., 2010).
Replication timing experiments have revealed that large domains,
typically 200-2000 kb in mammalian cells, are each replicated at
distinct times in S phase (Desprat et al., 2009; Hansen et al., 2010;
Hiratani et al., 2008; Pope et al., 2010). Generally, early-replicating
regions of the genome are correlated with active transcription and
chromatin marks associated with open forms of chromatin, whereas
late-replicating regions are transcriptionally silent and enriched in
chromatin marks associated with repressive chromatin. This is in
agreement with previous correlations of replication timing and
transcription from individual replication origins. These studies have
been extensively reviewed, as has been the link between replication
timing and imprinting, the process in which gene expression occurs
from either the maternal or paternal allele (for details, see Gilbert,
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2002; Göndör and Ohlsson, 2009). Consequently, we will focus on
how differentiation affects not only replication timing, but also
origin selection and usage.
Comparison of genome-wide replication timing profiles from
established Drosophila cell lines revealed that ~20% of autosomal
sequences have different times of replication in different cell types
(Schwaiger et al., 2009). Additionally, comparing regions of the
genome that display cell-type differences with respect to replication
timing confirms correlations between replication timing and gene
expression. It still remains unclear whether increased gene
expression actively promotes early replication, or transcription and
replication are governed by a common chromatin state.
In Drosophila, males have only one copy of the X chromosome.
This necessitates the need to increase gene expression from the
single X chromosome, a process termed dosage compensation.
Early experiments analyzing the replication patterns of Drosophila
salivary gland polytene chromosomes by tritium labeling revealed
that the X chromosomes of males completed replication earlier than
the female X chromosomes, whereas autosomes replicated with
similar kinetics (Berendes, 1966). Consistent with these
observations, analysis of the dosage-compensated X chromosome
from a male-derived cell line revealed that the male X replicates
almost entirely in early S phase, whereas the X chromosomes from
a female-derived cell line have early and late replication profiles
similar to those of autosomes (Schwaiger et al., 2009).
Interestingly, however, dosage compensation-dependent
upregulation of transcription cannot explain the shift to early
replication of the male X chromosome, because non-transcribed
regions are also early replicating. Furthermore, histone H4 lysine16
(H4K16) acetylation profiles, which reflect transcriptionally active
chromatin, were found to correlate very well with early replication
of the male X and early replicating regions of autosomes even in
regions of no transcription (Schwaiger et al., 2009). Therefore,
these results favor a model in which chromatin accessibility, not
transcription per se, dictates replication timing.
Comparison of multiple transformed human cell lines has
demonstrated that independent cell lines of the same lineage
display a remarkable similarity in replication timing profiles
(Hansen et al., 2010). By contrast, ~50% of the genome displays
cell type-specific differences in replication timing profiles (Hansen
et al., 2010). Similar to observations in human cell lines, analysis
of 22 cell lines derived from different stages of mouse development
indicated that ~50% of the genome displays differences in
replication timing (Hiratani et al., 2010). Although useful,
comparing transformed non-isogenic cell lines that have been
independently cultured for many years has limitations when
assessing how differentiation affects processes such as replication
timing. However, replication timing profiles of independently
derived mouse embryonic stem cell (mESC) lines, separated for
over 15 years in culture were shown to be nearly identical,
suggesting that replication timing is an extremely stable property,
at least in mESCs (Hiratani et al., 2008).
Cells in culture that can be triggered to differentiate provide
useful models for understanding how replication is affected
during development. Multiple independent studies using both
mouse and human ESCs and various cell types derived from
them have begun to define the effect of differentiation on
replication timing. In both human and mouse cells, the size of
replication domains ranges from ~200 kb to many megabytes,
and these are consolidated into fewer and larger domains during
differentiation. The molecular events responsible for these
changes are yet to be established (Desprat et al., 2009; Hiratani
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et al., 2008; Pope et al., 2010). Replication of many early and
late domains appears to be coordinated, meaning multiple
domains fire at very precise times during S phase. Monitoring
changes in replication timing as a function of differentiation in
a mouse ESC line and in a single differentiated neural precursor
cell type derived from that line revealed that ~20% of the
genome is subject to changes in replication timing (Hiratani et
al., 2008). This might be a more accurate percentage than that
derived from comparing independently derived differentiated cell
lines.
Given the vast array of cell types present in a metazoan
organism, more cell types will have to be examined to determine
the full impact that differentiation has on the replication program.
Most importantly, future studies need to focus on individual
replication origins. This is because multiple potential origins of
replication have been shown to cluster within zones (Vaughn et al.,
1990), and replication-timing experiments lack the resolution to
distinguish individual origin-firing events (Fig. 2). For example,
insertion of a single transgene into a region of late replication is
sufficient to change the replication timing of that region from late
to early (Lin et al., 2003). Therefore, relatively small changes on a
genetic and/or epigenetic level can dramatically change the
properties of a much larger replication domain.
Origin firing is modulated by differentiation
Characterization of individual replication origins within a
chromosomal domain has revealed that origin firing is itself
regulated during differentiation. This was first observed at the II/9A
locus of the fly Sciara coprophila, which undergoes developmental
programmed changes in gene copy number (Lunyak et al., 2002).
Mapping of replication origins by nascent-strand analysis within
the 120 kb HoxB locus in undifferentiated mouse P19 cells (see
Glossary, Box 1) identified at least five origins of replication
(Grégoire et al., 2006). Retinoic acid treatment causes P19 cells to
differentiate and induce gene expression from the HoxB locus.
Origins of replication are silenced during differentiation with the
exception of the Hoxb1 origin, which becomes the single dominant
replication origin within this domain. Furthermore, by assaying
transcript levels at various time points during retinoic acid
treatment, kinetic analysis revealed that the Hoxb1 origin is not
affected by transcription and that restriction of origin usage in this
locus upon differentiation is correlated with histone acetylation
(Grégoire et al., 2006). These results contrast with the positive
correlations between origin activation, transcription and histone
acetylation seen in genome-wide studies using cell culture. They
reveal the importance of studying individual replication origins to
address the mechanisms that regulate origin firing and the potential
for diverse regulatory schemes for modulation of individual origin
usage.
Powerful single-molecule analysis of replicated DNA (SMARD;
Box 2) has also been used to study the properties of individual
replication origins upon differentiation (Norio et al., 2005). The
benefit of SMARD analysis is that replication initiation and fork
elongation can be monitored on individual chromatin fibers. In
primary mouse ESCs and non-B cell lines, the Igh locus is silent
and replication initiates from a zone ~80 kb downstream of the
locus. A single unidirectional replication fork travels ~400 kb and
is responsible for replicating the entire Igh locus. However, in both
pro- and pre-B cell lines in which the locus is expressed, multiple
origins of replication become active within the Igh locus,
demonstrating that individual replication origins are regulated
during differentiation (Norio et al., 2005).
Development 139 (3)
Regions in which a unidirectional replication fork emanating
from an early domain is responsible for replicating a large originless region often separate early and late replicated domains and are
termed temporal transition regions (Ermakova et al., 1999). Elegant
single molecule analysis of the Igh locus confirmed the existence
of such domains, which have the potential to be regions of genomic
instability (Norio et al., 2005; Watanabe et al., 2004). Furthermore,
because temporal transition regions are a product of differences in
replication timing, particularly replication timing influenced by
development, temporal transition regions have the potential to be
developmentally augmented regions of genomic instability.
SMARD analysis of the replication program of the POU5F1
locus in human cells also revealed a change in both origin firing
and replication fork progression as a function of differentiation
(Schultz et al., 2010). In multiple ESC lines, the pattern of origin
usage and replication fork progression is similar within the
POU5F1 locus. By contrast, analysis of a differentiated cell line as
well as a multipotent neuronal rosette cell line derived from one of
the ESC lines used in this study revealed a change in origin usage
and directionality of fork progression within the POU5F1 region
(Schultz et al., 2010).
Modulation of origin firing during differentiation has recently
been shown to have a profound effect on genomic stability.
Common fragile sites are chromosomal regions susceptible to
breakage upon replication stress (Durkin and Glover, 2007). Recent
work has demonstrated that the fragility at these sites results from
the combined lack of initiation sites within these regions and their
late-replicating nature (Letessier et al., 2011). Furthermore, changes
in origin usage in multiple cell types can result in cell type-specific
differences in chromosomal fragility (Letessier et al., 2011). These
results suggest that changes in the DNA replication program, which
occur during the course of differentiation, could ultimately be
responsible for the tissue and/or cell type-specific genomic
instability associated with tumor cell progression.
Differential replication in development
Alterations in the cell cycle and DNA replication programs are often
utilized during organismal development to ensure proper tissue
function. One variant cell cycle is the endo cycle (see Glossary, Box
1) in which repeated S-G cycles occur in the absence of mitosis. This
results in polyploid cells, or cells with increased genome content
(Fig. 3) (Edgar and Orr-Weaver, 2001; Lee et al., 2009).
Polyploidy (see Glossary, Box 1) is common in plant and animal
development and is generally thought to be a developmental
strategy to produce large, highly metabolically active cells. In
mammals, polyploidy is essential for the development of the
placenta. In the mouse, genetic ablation of the S phase cyclin,
cyclin E, results in embryonic lethality (Geng et al., 2003; Parisi et
al., 2003) due to a block in the endo cycle of the placental giant
trophoblast cells, resulting in their failure to become polyploid. The
giant trophoblast cells attain ploidy levels up to 1000C in rodents,
and their large size might facilitate a maternal-fetal barrier.
Megakaryocytes, the large precursor cells from which blood
platelets are derived, also are polyploid (Nguyen and Ravid, 2010).
The large size of polyploid megakaryocytes is required for
sufficient cytoplasm to bud off adequate numbers of platelets. In
Drosophila, nearly all larval tissues and numerous adult tissues
have increased ploidy. Most of these cell types are polytene (see
Glossary, Box 1), meaning that the replicated sister chromatid
copies are held in tight association with one another. Blocking
polyploidization in Drosophila inhibits cell and larval growth and
perturbs proper tissue function (Edgar and Orr-Weaver, 2001).
DEVELOPMENT
460
REVIEW
A Mitotic cycle
B Endo cycle
S phase
S phase
G1 phase
Endo
cycle
Mitotic
cycle
G2 phase
M phase
G phase
C Gene amplification
16
100 kb
8 4
D Under-replication
2
16 8
1
4
2
1
300 kb
Fig. 3. Differential DNA replication. (A)A mitotic cycle with four
distinct phases: G1, S, G2 and M phase. (B)By contrast, the endo cycle
consists of repeated rounds of S and G phases with no intervening
mitoses, resulting in polyploid cells with increased genome content per
cell. (C)Developmentally programmed gene amplification can occur
through repeated rounds of origin firing followed by bidirectional
replication fork progression, resulting in gradients of copy number over
a 100 kb domain. The example shown depicts four rounds of origin
firing. Arrows at top indicate direction of fork progression.
(D)Schematic of the copy number differences associated with underreplication. Arrows at top indicate direction of fork progression relative
to the maximally under-replicated region. Under-replicated domains
within euchromatin range from 100 to 400 kb.
Although polyploid cells have increased genome content per
cell, gene copy number is not always uniform throughout the
genome (Fig. 3). For example, programmed gene amplification
(see Glossary, Box 1) in Drosophila ovarian follicle cells and the
Sciara salivary glands results in an increase in genomic copy
number by repeated rounds of origin firing at specific loci
(Claycomb and Orr-Weaver, 2005). This process is essential to the
function of these cells during development. Polyploid cells can also
have regions of reduced copy number relative to overall ploidy, a
phenomenon termed under-replication (see Glossary, Box 1). The
function of under-replication during development still remains
unclear. However, the identification of under-replicated regions by
genome-wide techniques has demonstrated tissue-specific control
of under-replication and has provided insight into the mechanisms
that result in the repression of DNA replication in these regions.
The developmental control of under-replication
In Drosophila, it has long been appreciated that heterochromatic
regions of the genome are under-replicated with respect to
overall ploidy (Spradling and Orr-Weaver, 1987). Additionally,
a small number of sites distributed along the euchromatic arms
in the larval salivary gland and larval fat body have been shown
at the molecular level to be under-replicated (Belyaeva et al.,
1998; Marchetti et al., 2003). Using genome-wide array-based
comparative genomic hybridization (aCGH; see Box 2) it has
now been demonstrated that under-replication is prevalent
throughout the euchromatic arms of the Drosophila genome
(Nordman et al., 2011). Comparing the copy number profiles of
DNA isolated from larval salivary gland, larval midgut and the
larval fat body tissues revealed that differential replication
displays tissue specificity. The mechanisms responsible for these
461
tissue-specific differences in DNA copy number are not clear,
but differences in ploidy cannot account for them (Nordman et
al., 2011). This would suggest that an active, developmentally
controlled mechanism exists to regulate differential replication.
Whether this is at the level of chromatin structure or regulation
of DNA replication factors remains to be determined as does the
role played by differential replication during organismal
development. Additionally, regions of under-replication provide
a powerful tool for investigating the molecular mechanisms that
control DNA copy number during development.
DNA under-replication in Drosophila is dependent on the
Suppressor of Under-Replicaton gene, SuUR (Belyaeva et al.,
1998). Genome-wide aCGH experiments have demonstrated that
all regions of under-replication become fully replicated if SuUR
is not functional (Nordman et al., 2011). Little is known about
how SUUR functions to repress DNA replication, but its N
terminus shows homology to the SWI/SNF family of chromatin
remodelers and, therefore, it could act by influencing chromatin
structure (Makunin et al., 2002). SUUR has been shown to be a
component of a newly identified repressive form of chromatin in
Drosophila termed BLACK chromatin (different chromatin
states were named with color names for simplicity), which in cell
culture is generally late replicating and repressed for
transcription (Filion et al., 2010).
To determine whether SUUR prevents replication by inhibiting
origin function within regions of under-replication, ORC binding
was mapped genome-wide in wild-type and SuUR mutant larval
salivary glands. The frequency of ORC binding sites was reduced
significantly within regions of under-replication compared with
fully replicated regions of the genome (Sher et al., 2012).
Remarkably, in the SuUR mutant, although copy number is
completely restored to the normally under-replicated regions, they
remain devoid of ORC binding, showing a distribution of ORC
binding virtually identical to that in wild type. This result indicates
that SUUR does not affect ORC binding, but rather could inhibit
replication forks from entering regions of under-replication. Using
a model system to study replication fork progression, it was shown
that replication forks travel a greater distance in the same
developmental time frame in SuUR mutant cells compared with
wild-type cells (Sher et al., 2012). Therefore, these results strongly
indicate that developmentally programmed changes in gene copy
number can be achieved by regulation of fork progression.
During S phase of a mitotic cell cycle, multiple mechanisms
exist to ensure that the genome is replicated once per cell cycle
(Arias and Walter, 2007). However, during the S phases of the
endo cycle, both heterochromatic and euchromatic sequences are
under-replicated, suggesting that the checkpoints that regulate
proper copy number in mitotic cells are inactive in endo cycling
cells (Lilly and Spradling, 1996). In support of this idea, it has
been demonstrated that the boundary between fully replicated
euchromatin and under-replicated heterochromatin accumulates
DNA damage (Hong et al., 2007). Furthermore, endo cycling
cells are unable to induce apoptosis following genotoxic stress,
owing to the inability of these cells to induce the apoptotic genes
that would be expressed in mitotic cells following genotoxic
stress (Mehrotra et al., 2008). The mechanism(s) responsible for
silencing the checkpoint activity in endo cycling cells is
currently unknown. Additionally, these studies raise the
possibility that the response of a cell to DNA damage and
replication stress can be influenced by its developmental state,
which has broad implications for cell-type specific genomic
instability and tumor formation.
DEVELOPMENT
Development 139 (3)
REVIEW
Developmentally regulated gene amplification
Developmentally programmed gene amplification is another form
of differential DNA replication necessary for proper tissue function
(Claycomb and Orr-Weaver, 2005). The follicle cells of the
Drosophila ovary form an epithelial layer around the developing
oocyte. After a series of mitotic divisions, follicle cells enter the
endo cycle and increase their overall ploidy to 16C. Follicle cells
then exit the endo cycle and enter into a developmentally regulated
gene amplification program, in which six sites are repeatedly
amplified, forming so-called amplicons (see Glossary, Box 1), by
a ‘re-replication’-based mechanism resulting in ~100 kb gradients
of copy number (Fig. 3) (Claycomb et al., 2004; Kim et al., 2011).
The mechanism by which most replication origins are silenced but
six origins are selected as amplification sites is unknown. Follicle
cell amplicons provide a powerful tool to study metazoan origin
regulation, because changes in gene copy number are precisely
timed during development. Thus, both initiation of replication and
replication fork progression can be individually assessed during
development.
The identification of all the amplicons in follicle cells has
permitted the integrative analysis of genome-wide ORC binding,
histone acetylation, and transcription levels (Kim et al., 2011).
Amplification was previously shown to be dependent on histone
acetylation, a property correlated with ORC binding and origin
activation in many genome-wide studies (Aggarwal and Calvi,
2004; Hartl et al., 2007). Histone acetylation was assayed both
genome wide and specifically within amplification regions
revealing that histone acetylation is found at some sites of
amplification, but not all. These results demonstrate that histone
acetylation is not a prerequisite for origin firing (Kim et al., 2011).
Follicle cell gene amplification is a developmental strategy that
increases the amount of DNA template to support the high level of
transcription necessary to produce sufficient chorion (see Glossary,
Box 1) proteins in a narrow developmental window (Spradling and
Mahowald, 1980). Although this strategy has been well
characterized for a small number of genes involved in chorion
synthesis, the relationship between gene amplification and
transcription had not been analyzed genome wide. To understand
this relationship better, whole-transcriptome profiling was
performed on amplifying follicle cells and gene expression levels
were compared with copy number changes (Kim et al., 2011). As
expected, many of the most highly expressed genes are located
within the two main chorion gene clusters. Many robustly
transcribed genes lie outside the amplicons, however. In addition,
genes located within amplified regions are not always highly
expressed (Kim et al., 2011). This could be important for
understanding the significance of amplified genomic regions that
are prevalent in cancer cells. The discovery that gene expression is
not always correlated with copy number in Drosophila amplicons
is an important consideration when assessing how copy number
variation might affect gene expression in cancer cells.
Conclusions
It is now clear that the DNA replication program is modulated as a
function of development. Numerous genome-wide studies and
studies of individual replication origins have established a
correlation between origin efficiency and chromatin state (Ding and
MacAlpine, 2011). It remains to be determined whether the
changes in the replication program that occur during development
reflect an active mechanism to maximize the efficiency of DNA
replication and/or genomic stability during development, or if
changes in the DNA replication program are a passive reflection of
Development 139 (3)
developmentally regulated changes in chromatin state. Research in
model systems such as Drosophila has clearly demonstrated that
developmentally programmed changes in DNA replication are
necessary for proper cell function. How common programmed
changes in DNA replication are in development remain to be
determined.
Recent studies addressing the cause of MGS demonstrated that
failure to initiate properly DNA replication can have specific
effects on human development (Bicknell et al., 2011a; Bicknell et
al., 2011b; Guernsey et al., 2011). But, why are certain tissues
and/or cell types so susceptible to defects in DNA replication? A
simple hypothesis is that the most highly proliferative cell types
would be the most vulnerable to alterations in the DNA replication
program. The phenotypes associated with MGS suggest that this is
unlikely to be the case. Therefore, a major focus in addressing the
relationship between DNA replication and development will be to
understand why certain cell types and tissues are affected by
alterations in the DNA replication program. Only then will we be
able to gauge the impact programmed DNA replication has on
proper organismal development.
Defects in proper DNA replication have long been associated
with cancer. For example, gene amplification and polyploidy are
associated with numerous tumor types, but the primary defects
responsible for generating gene amplification have been difficult to
identify (Albertson, 2006; Beroukhim et al., 2010). Recent work
has demonstrated directly that loss of DNA replication control can
initiate the amplification process (Green et al., 2010). Furthermore,
defects in proper replication licensing result in increased levels of
chromosomal instability and spontaneous tumor formation
(Beroukhim et al., 2010; Kawabata et al., 2011). Thus, proper DNA
replication is not only essential for proper organismal development
but also to prevent the onset of cancer.
Acknowledgements
We thank Mary Gehring, Jane Kim, Gerry Fink, Angelika Amon and Steve Bell
for constructive comments on the manuscript. We are grateful to Tom
DiCesare for technical assistance in figure preparation.
Funding
J.N. is a Howard Hughes Medical Institute (HHMI) Fellow of the Damon
Runyon Cancer Research Foundation and has received support from a
Margaret and Herman Sokol postdoctoral award. This work was supported by
the National Institutes of Health (NIH) and an American Cancer Society
Research Professorship to T.L.O.-W. Deposited in PMC for release after 12
months.
Competing interests statement
The authors declare no competing financial interests.
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