Abstract

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. 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 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).

Table 1.

Components necessary for helicase loading and helicase activation

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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.

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 (S-CDK and DDK, respectively) (Fig. 1B). ORC binding and pre-RC formation occur in late M and G1 phases of the cell cycle when S-CDK 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 loading, but also origin firing by the activation of the Mcm2-7/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 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 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

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.

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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, population-scale replication timing experiments lack the resolution to monitor changes in individual origin usage during differentiation.

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., 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, 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 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.

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).

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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 under-replication. 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.

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).

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.

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 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.

Acknowledgments

Footnotes

References