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Keywords:

  • Checkpoint;
  • C-value;
  • heterochromatin;
  • mutation rate;
  • ribonucleotide reductase;
  • transposable elements

Abstract

  1. Top of page
  2. Abstract
  3. Nucleotypic Effect and Asynchrony of DNA Replication Program in Eukaryotes
  4. Asynchronous Replication and Mutation Rates in Vertebrates
  5. Mutation Rates and Gene Location Effects in Fungi and Invertebrates
  6. DNA Repair Systems in Heterochromatin
  7. Checkpoint Function and Mechanisms Underlying Replication Asynchrony
  8. Epigenetic Regulation of Replication Timing
  9. Role of Ribonucleotide Reductase in Replication Asynchrony
  10. Genome Size, Checkpoint Proficiency, and Mutation Rate Heterogeneity
  11. Genome Size and Rates of Speciation
  12. Transposons and DNA Repair Systems
  13. ACKNOWLEDGMENTS
  14. LITERATURE CITED

Mutation rates vary significantly within the genome and across species. Recent studies revealed a long suspected replication-timing effect on mutation rate, but the mechanisms that regulate the increase in mutation rate as the genome is replicated remain unclear. Evidence is emerging, however, that DNA repair systems, in general, are less efficient in late replicating heterochromatic regions compared to early replicating euchromatic regions of the genome. At the same time, mutation rates in both vertebrates and invertebrates have been shown to vary with generation time (GT). GT is correlated with genome size, which suggests a possible nucleotypic effect on species-specific mutation rates. These and other observations all converge on a role for DNA replication checkpoints in modulating generation times and mutation rates during the DNA synthetic phase (S phase) of the cell cycle. The following will examine the potential role of the intra-S checkpoint in regulating cell cycle times (GT) and mutation rates in eukaryotes. This article was published online on August 5, 2011. An error was subsequently identified. This notice is included in the online and print versions to indicate that both have been corrected October 4, 2011.

Eukaryotic genomes are organized into two major spatially separate chromatin compartments: highly condensed gene-poor heterochromatin and decondensed gene-rich euchromatin (Dillon 2004; Bassett et al. 2009). These spatially separate compartments are also separated temporally with respect to replication timing, or when in the DNA synthetic phase (S-phase) of the cell cycle a region of the genome is replicated (Woodfine et al. 2004; Karnani et al. 2007; Gilbert 2010; Shermoen et al. 2010). Although the relationship between chromatin structure and replication timing is complex, euchromatin in general replicates earlier in S-phase, while heterochromatin replicates later (Gilbert et al. 2004; Lee et al. 2010; Fig. 1). Initial studies on two unrelated species revealed that heterochromatin is located in distinct regions at the periphery of the nucleus and contains three times more DNA than euchromatin (Lima-de-Faria 1959). A more recent study found that replication origins—the sites where DNA synthesis begins—appear to be more clustered (3X) at mid-to-late S-phase compared to early S-phase, which is enriched in single bidirectional origins (Frum et al. 2009). In addition, a third study has demonstrated that in all eukaryotes examined so far the activation of replication origins increases during the first half of S-phase and then decreases toward the end of the phase (Goldar et al. 2009).

image

Figure 1. The eukaryotic DNA replication program (Huberman and Riggs 1968). Early replicating replication domains (boxes) are interspersed between late replicating domains (lines). Replication origins (black dots) are less clustered in gene expressing euchromatin that replicates early in S-phase (Frum et al. 2009). Replication bubbles (ovals) form when replication origins fire stochastically. Probabilistic origin firing results in a different replication pattern each S-phase (S1, S2, S3). Some origins, however, fire preferentially and more frequently (second black dot in E2). Replication domains, in contrast, are activated nonrandomly according to a deterministic replication-timing program (E1-L6) (Jackson and Pombo 1998; Labit et al. 2008).

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The differences in the timing and dynamics of genome duplication in metazoa are mediated in part by the ATR-Chk1 checkpoint pathway, which coordinates DNA replication and DNA repair. At mid S-phase in a murine cell line, for example, a checkpoint-associated transition takes place between the S-phase regulatory factors Cdk2/cyclinE, which govern early-replication origin activities, and Cdk1/cyclinA2, which govern late-replication origin activity (Katsuno et al. 2009). Similarly in yeast, late replication is governed by the S-phase regulatory cyclin Clb5, while early replication is governed by the cyclin Clb6 (Donaldson et al. 1998; McCune et al. 2008). DNA synthesis in euchromatin and heterochromatin therefore corresponds to different replication regimes with distinct replication kinetics in both lower and higher eukaryotes.

The evolutionarily conserved spatial and temporal compartmentalization of the eukaryote genome indicates that heterochromatin plays an important role in genome dynamics and architecture. Genome size, for example, varies in proportion to the fraction of heterochromatin it contains (Bennetzen et al. 2005; Gregory 2005a; Table 1). A substantial fraction of heterochromatin is composed of repetitive DNA sequences and transposable elements (TE), which include DNA transposons, retro-transposons, long interspersed elements (LINE), and short interspersed elements (SINE) (Richard et al. 2008). The nature and composition of these sequences vary in a lineage-specific manner, and the sequences themselves frequently exhibit high mutation rates (Crouau-Roy and Clisson 2000). Alu elements, for example, are found in the primate lineage and are enriched in mutation-prone methylated CpG sites (Yoder et al. 1997; Dimitri et al. 2003; Lippman et al. 2004; Zamudio and Bourc’his 2010). CpG methylation is believed to have evolved as a means to silence gene expression and repress TE transposition events (Meunier et al. 2005), which can damage the genome and result in genetic disease and sterility (Charlesworth et al. 1994). Because TE insertions can be deleterious, the host has evolved a variety of strategies to limit, but not eradicate, TEs and transposition events (Biémont and Vieira 2005). In response, TEs have evolved counter strategies to guarantee their persistence in the host germline (Cam et al. 2008; Blumenstiel 2011).

Table 1.  Species genome size and composition compared to preferred DSB repair pathway (NHEJ:HR) and mutation rates (μ)
SpeciesGenome sizea (pg)Genic densityb (Ge/G)Intronsc (per gene)TEd (%)NHEJ: HRcμeμ (per genome per replication)
  1. bEffective genome size, Ge, is the approximate number of genes × 103 base pairs. G is estimated genome size in base pairs.

  2. dsee refs: Kim JM et al. (1998), Consortium IHGS (2001), Mural RJ et al. (2002).

  3. eMutation per base pair per replication. Data are in units of 10−10. See Lynch et al. (2008) and Drake et al. (1998).

S. cerevisiae 0.017 0.4680 0.07 3.1 1:100 2.2 0.0027
C. elegans 0.10.19744.76.5NA2.30.018
D. melanogaster 0.12–0.21 0.0756 4 3.86–9 3:2 3.4 0.058
Mus musculus 3.280.00768363:11.80.49
Homo sapiens 3.5 0.0070 9 45 9:1 0.5 0.16

Genome size and organization are clearly consequences of adaptive measures operating through the molecular machinery that governs genome stability and progression through S-phase of the cell cycle (Wyngaard and Gregory 2001). Preference for DNA double-strand break (DSB) repair pathways, for example, varies in a genome size-dependent manner, indicating an unambiguous effect of heterochromatin not only on the DNA replication program, but also on the choice between DNA repair systems (Farlow et al. 2011; Table 1). How the DNA replication program and DNA repair systems are coordinated, however, remains to be fully elucidated. One aspect of that coordination concerns the role of checkpoint function in regulating the S-phase in species with different genome sizes. The following will examine the relationship between the organization of the DNA replication program and corresponding differences in DNA repair efficiencies, both within the genome and across species.

Nucleotypic Effect and Asynchrony of DNA Replication Program in Eukaryotes

  1. Top of page
  2. Abstract
  3. Nucleotypic Effect and Asynchrony of DNA Replication Program in Eukaryotes
  4. Asynchronous Replication and Mutation Rates in Vertebrates
  5. Mutation Rates and Gene Location Effects in Fungi and Invertebrates
  6. DNA Repair Systems in Heterochromatin
  7. Checkpoint Function and Mechanisms Underlying Replication Asynchrony
  8. Epigenetic Regulation of Replication Timing
  9. Role of Ribonucleotide Reductase in Replication Asynchrony
  10. Genome Size, Checkpoint Proficiency, and Mutation Rate Heterogeneity
  11. Genome Size and Rates of Speciation
  12. Transposons and DNA Repair Systems
  13. ACKNOWLEDGMENTS
  14. LITERATURE CITED

Early investigations in plants demonstrated a strong nucleotypic effect on cell cycle times: cell cycle duration correlates strongly with genome size (Bennett 1972; Francis et al. 2008). The extended division cycle in cells with larger genomes was found to be due to an increase in replication asynchrony (Kidd et al. 1987). Asynchrony refers to the time in S-phase at which different regions of the genome are replicated, and is roughly defined as the average time to duplicate a replicon (Rs) divided by the time required to duplicate the entire genome (Ds): Rs/Ds (Fig. 2A). Replicon size here refers to the length of DNA replicated from a single origin of replication. Importantly, the correlation between genome size and S-phase duration can have significant effects on life-history traits such as developmental and metabolic rates (Gregory 2004; 2005b). Genome size thus acts as a molecular pacemaker that mediates the durations of S-phase and the cell cycle, and in some species has higher order physiological effects on the organism (Cavalier-Smith 1978, 1980; Gregory 2004; Bromham 2009).

image

Figure 2. Scaling of replication asynchrony with genome size. (A) The degree of replication asynchrony is measured as the ratio of the time required to duplicate a replicon (Rs) divided by the time required to duplicate the genome (Ds): Rs/Ds (Van't Hof and Bjerknes 1981; Kidd et al. 1987). As the size of a genome increases, the degree of asynchrony increases. Asynchrony could reflect the different times that individual replicons are activated during S-phase, or the different times that replication domains, which contain multiple replicons, are replicated, or both. Replication domain size scales with genome size, so the degree of asynchrony would depend on the time required to replicate both the replication domain and, at mid S-phase, the large origin-suppressed regions of the genome (not shown). (B) Increase in replication asynchrony during Drosophila development. In embryos, cell division cycles are rapid, devoid of checkpoint function, gene transcription, and G1 and G2 phases. At the MBT the Drosophila grapes/Chk1 intra-S checkpoint is activated, chromatin is remodeled, and a gene transcription program implemented. In somatic cells, the division cycle is extended to 600 min (adapted from Shermoen et al. 2010).

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The same plant studies revealed a direct correlation between replicon size and replication fork rate, defined as the rate at which DNA synthesis elongates the growing DNA daughter molecule. Larger replicons tend to be replicated by faster replication forks, whereas smaller replicons tend to be replicated by slower forks. Hence, the average replicon duplication rate is remarkably constant in plants: 1–2 h in each species examined (Kidd et al. 1989). Variation in asynchrony in different plants therefore depends principally on Ds and genome size. Replicon sizes and replication fork rates, however, are highly heterogeneous, both within a given eukaryotic genome and between different tissue types and organisms (Van't Hof and Bjerknes 1981; Berezney et al. 2000). Consequently, average replicon duplication times can vary considerably depending on species, tissue type, and location in the genome.

Nevertheless, a similar correlation between replicon size and fork rate has been repeatedly observed across a broad range of species (Hand 1975; Kidd et al. 1989; Conti et al. 2007b), although these observations have been questioned as possible artifacts (Stimac et al. 1977; Liapunova 1994). Moreover, in almost every species examined so far, including bacteria, experimentally increasing (decreasing) the number of replication initiation events per division cycle simultaneously results in a corresponding decrease (increase) in replication fork rates (Herrick and Sclavi 2007; Herrick and Bensimon 2008). No such correlations have been observed, however, between either replicon size or replication fork rate and genome size (Kidd et al. 1987). Clearly, the regulation of replicon duplication rates (Rs) and the asynchronous nature of DNA replication (Rs/Ds) in eukaryotes play a significant role in determining the pace of progression through the S-phase of the cell cycle (Fig. 2A,B).

How replication asynchrony is regulated during S-phase is unclear, but recent genome-wide studies on the DNA replication program in metazoa have revealed that megabase-sized blocks of the genome (1 to 2 Mb on average) replicate at reproducibly different times in S-phase (Woodfine et al. 2004; Farkash-Amar et al. 2008; Hiratani et al. 2008; Hansen et al. 2010). These “replication-timing domains” contain multiple replicons (1–10), and the average size of the domains apparently scales with genome size (Gilbert 2010). When one domain completes replication, a genetically adjacent domain begins replication in sequence (Maya-Mendoza et al. 2010). In addition to replication-timing domains, replication asynchrony might also be affected by large (up to 1.5 Mb) regions of DNA referred to as Timing Transition Regions (TTR) (Méndez 2009). TTRs, which are associated with the transition between early and late S-phase, do not contain replication origins, and are often replicated by a single replication fork. Similarly, a recent report demonstrates that common fragile sites, such as human FRA3B, are devoid of replication origins and consequently complete replication very late in S-phase (Letessier et al. 2011). Replication asynchrony thus operates on at least two levels in the genome: between individual replicons and between replication-timing domains. Remarkably, the replication-timing program is evolutionarily conserved between mouse and man (Ryba et al. 2010; Yaffe et al. 2010).

Asynchronous Replication and Mutation Rates in Vertebrates

  1. Top of page
  2. Abstract
  3. Nucleotypic Effect and Asynchrony of DNA Replication Program in Eukaryotes
  4. Asynchronous Replication and Mutation Rates in Vertebrates
  5. Mutation Rates and Gene Location Effects in Fungi and Invertebrates
  6. DNA Repair Systems in Heterochromatin
  7. Checkpoint Function and Mechanisms Underlying Replication Asynchrony
  8. Epigenetic Regulation of Replication Timing
  9. Role of Ribonucleotide Reductase in Replication Asynchrony
  10. Genome Size, Checkpoint Proficiency, and Mutation Rate Heterogeneity
  11. Genome Size and Rates of Speciation
  12. Transposons and DNA Repair Systems
  13. ACKNOWLEDGMENTS
  14. LITERATURE CITED

Mutation rates within and between genomes in vertebrate and invertebrate species are heterogeneous (Drake et al. 1998) and display variable patterns that depend on genomic location (Baer et al. 2007), as well as nucleotide context (flanking nucleotides) and lineage (Hwang and Green 2004). Changes in DNA replication dynamics, although not the unique basis of nucleotide substitution rates, play a significant role in the heterogeneity of mutation rates in the genome (Hwang and Green 2004). In all organisms examined so far, for example, mutation rates increase with the distance from replication origins (Mira and Ochman 2002; Flynn et al. 2010; Mugal et al. 2010). Likewise, DNA replication in mammalians slows down significantly at mid S-phase when TTRs are replicated, and these regions frequently coincide with DNA fragile sites (Takebayashi et al. 2005; Durkin and Glover 2007). The compartmentalization of mutation rate has long been suspected to reflect the effects of DNA replication timing and gene transcription (Britten 1986; Wolfe et al. 1989; Widrow et al. 1998).

Recent studies on replication timing in vertebrates have revealed a monotonic increase in mutation rates as S-phase advances. In two investigations, mutation rates in late replicating DNA were found to increase with replication timing by 10.5% in rodents and 22% in primates (Stamatoyannopoulos et al. 2009; Pink and Hurst 2010). The third investigation reported a 30% increase in diversity in primates and a 29% increase with replication timing in rodents (Chen et al. 2010). The latter study revealed an important contribution from CpG methylation status to the increase in mutation rate, which is consistent with region-specific variability in mutation rates due to CpG substitutions (Elango et al. 2008; Walser and Furano 2010). A negative correlation, for example, has been observed between CpG substitution rates and G + C content (Elango et al. 2008). In metazoa, early replicating gene-rich regions associated with open chromatin generally have a high GC base composition (Gilbert et al. 2004) and hence are expected to have lower CpG substitution rates.

These findings are also consistent with earlier reports on the genomic location of mutation “hot spots” and “cold spots” in the human genome (Chuang and Li 2004). Early replicating genes, such as essential housekeeping genes and genes involved in apoptosis, are more conserved than later replicating genes (Cohen et al. 2007; Kaufman et al. 2011), which tend to be devoted to more environmentally adaptive and species-specific functions such as immune response genes and the olfactory gene cluster (Chuang and Li 2004). In addition, gene duplication clusters in primates occur with higher frequency in late replicating regions of the genome. Clusters of duplicated genes arise from gene amplification and recombination events (Hahn et al. 1986; Hoy et al. 1987; Lugo et al. 1989; Reams and Neidle 2004; Conti et al. 2007a; Green et al. 2010), both of which are consistent with higher mutation rates in late S-phase. Together, these observations suggest that later replicating DNA is more genetically labile than early replicating DNA. Not surprisingly, then, the later replicating genes located in mutational hot spots correspond to the type of gene most likely involved in speciation events and adaptive radiations.

Mutation Rates and Gene Location Effects in Fungi and Invertebrates

  1. Top of page
  2. Abstract
  3. Nucleotypic Effect and Asynchrony of DNA Replication Program in Eukaryotes
  4. Asynchronous Replication and Mutation Rates in Vertebrates
  5. Mutation Rates and Gene Location Effects in Fungi and Invertebrates
  6. DNA Repair Systems in Heterochromatin
  7. Checkpoint Function and Mechanisms Underlying Replication Asynchrony
  8. Epigenetic Regulation of Replication Timing
  9. Role of Ribonucleotide Reductase in Replication Asynchrony
  10. Genome Size, Checkpoint Proficiency, and Mutation Rate Heterogeneity
  11. Genome Size and Rates of Speciation
  12. Transposons and DNA Repair Systems
  13. ACKNOWLEDGMENTS
  14. LITERATURE CITED

Similar observations have been reported in invertebrates and yeast. In budding yeast, subtelomeric DNA and DNA flanking Sir-silenced heterochromatin are late replicating and have a higher rate of diversification compared to the genome-wide average (Teytelman et al. 2008). Single nucleotide polymorphisms in these regions have an elevated frequency: 7% versus 4.4% in the rest of the genome. Other studies on chromosome VI in yeast likewise revealed a replication-timing effect on mutation rate (Lang and Murray 2011). Interestingly, many of the genes located in the hypervariable regions play a role in adapting to changes in the environment. In eukaryotic microbial parasites such as Giardia lamblia, for example, genes involved in evading the host immune response also have subtelomeric locations (Adam et al. 2010), suggesting that a genomic war of attrition has mutually driven the organization and evolution of host and pathogen genomes.

Studies in Drosophila also revealed similar patterns of nucleotide diversity between early and late replicating DNA (Díaz-Castillo and Golic 2007; Anderson et al. 2008). Genes located near Drosophila heterochromatin display higher levels of diversity than in euchromatin, and per site sequence divergence between Drosophila melanogaster and D. simulans was threefold higher in subtelomeric heterochromatin than in neighboring euchromatin. Consistent with the studies on mouse and human, the higher mutation rates in late replicating DNA are in part attributable to CpG methylation status (Díaz-Castillo and Golic 2007; Prendergast et al. 2007; Walser and Furano 2010), suggesting that TE-enriched heterochromatin positively influences mutation rates in late replicating DNA independently of DNA replication itself. With the apparent exception of Caenorhabditis elegans (Denver et al. 2009), these findings, given the wide range of species, indicate that higher nucleotide substitution rates are likely due to reduced replication fidelity (elevated mutation rates) and lower levels of DNA repair efficiencies (elevated substitution rates) in late replicating heterochromatin.

DNA Repair Systems in Heterochromatin

  1. Top of page
  2. Abstract
  3. Nucleotypic Effect and Asynchrony of DNA Replication Program in Eukaryotes
  4. Asynchronous Replication and Mutation Rates in Vertebrates
  5. Mutation Rates and Gene Location Effects in Fungi and Invertebrates
  6. DNA Repair Systems in Heterochromatin
  7. Checkpoint Function and Mechanisms Underlying Replication Asynchrony
  8. Epigenetic Regulation of Replication Timing
  9. Role of Ribonucleotide Reductase in Replication Asynchrony
  10. Genome Size, Checkpoint Proficiency, and Mutation Rate Heterogeneity
  11. Genome Size and Rates of Speciation
  12. Transposons and DNA Repair Systems
  13. ACKNOWLEDGMENTS
  14. LITERATURE CITED

Early studies demonstrated preferential and faster rates of DNA repair in actively transcribed euchromatin (fast repair component) compared to heterochromatin (slow repair component; Zolan et al. 1982; Surrallés et al. 1997a,b). More recent studies have provided additional evidence that DNA repair systems are less efficient in late replicating heterochromatin (Prendergast et al. 2007; Walser and Furano 2010). Missmatch (MMR) and nucleotide excision repair (NER) systems, for example, are directly associated with DNA synthesis and are more active in open euchromatin than heterochromatin (Masih et al. 2008; Edelbrock et al. 2009; Solimando et al. 2009). The DNA replication processivity factor, PCNA, recruits MMR proteins to replication forks and co-localizes with the NER apparatus (Katsumi et al. 2001). In rodents and humans, it was found that NER efficiency depends on chromatin compartment, being most efficient in gene-rich R bands, less efficient in gene-poor G bands, and least efficient in heterochromatic C bands (Sanders et al. 2004). In yeast, NER is likewise significantly less efficient in subtelomeric heterochromatin compared to unrepressed transcriptionally active regions (Irizar et al. 2010) and has been found to be functionally impaired in Sir-associated heterochromatin (Chaudhuri et al. 2009). Similarly, base excision repair (BER) is substantially more efficient in euchromatin than in heterochromatin (Amouroux et al. 2010).

DNA replication repair foci have also been found to form preferentially in euchromatin compared to heterochromatin (Cowell et al. 2007; Karagiannis et al. 2007). Euchromatin has long been believed to be more sensitive to ionizing radiation than more compact heterochromatin (Kaufman et al. 2011); however, evidence suggests that heterochromatin is more prone to mutation (Falk et al. 2010), consistent with a preferential repair of euchromatin. These observations point to different modes of mutation and repair in heterochromatin compared to euchromatin (Chiolo et al. 2011). Indeed, heterochromatin differs from euchromatin in its greater reliance on ATM activation to decondense heterochromatin to repair DSBs (Goodarzi et al. 2008; Shibata et al. 2010).

Although DSBs induced in heterochromatin are efficiently repaired by homologous recombination (HR; Branzei and Foiani 2008; Beucher et al. 2009), the error-prone nonhomologous end joining (NHEJ) recombination repair pathway repairs 80–85% of DSBs in G2 (Beucher et al. 2009), and has been shown to increase as the cell cycle advances (Mao et al. 2008b). NHEJ activity is low in G1, increases in S-phase and peaks in G2. In contrast, error-free HR is most active in S-phase, but is significantly slower than NHEJ. HR-mediated repair can take up to 7 h, whereas NHEJ is considerably faster and completes repair in 30 min (Mao et al. 2008a). The different kinetics of repair might explain in part the differential reliance on the respective repair systems as the cell cycle advances (Shibata et al. 2011). HR, for example, requires the presence of a sister chromatid, and depends on intra S-phase checkpoint factors such as Chk1 (Sørensen et al. 2005). Consequently, more open and accessible euchromatin would favor the more faithful but slower HR repair system when euchromatin is being replicated earlier in S-phase. In contrast, more compact and less-accessible heterochromatin might act to spatially stabilize and constrain a DSB, and so favor the more rapid NHEJ repair system as the cell cycle advances into G2. HR, however, is required to repair DSBs in heterochromatin, whereas NHEJ is not (Chiolo et al. 2011).

HR:NHEJ ratios also vary across species (Farlow et al. 2011). Species with smaller genomes, such as yeast, rely predominantly on HR, whereas vertebrates rely predominantly on NHEJ. Invertebrates fall between the two extremes (Table 1). Recently, the proposal was made that the preference for NHEJ over HR can account for the observed increase in the average number of introns per gene in different species (Farlow et al. 2011). The number of introns increases in proportion to the ratio of NHEJ to HR activity in the cell, indicating that a clear trend exists in the relationship between the mode of DSB repair and the fraction of noncoding DNA in both euchromatin and heterochromatin (Table 1). The growing preference for NHEJ repair as genome size increases reflects the fact that HR is used to repair DSBs caused by blocked replication forks, whereas NHEJ is used to repair “accidental” DNA double-strand breaks (DSBs) induced by genotoxins in more compact chromatin compartments (Sonoda et al. 2006). Thus, HR is under direct control of the ATR Chk1 intra-S checkpoint during S-phase (Sørensen et al. 2005), whereas in G2, HR is primarily dependent on the ATM checkpoint pathway (Goodarzi et al. 2008, 2009, 2010).

Checkpoint Function and Mechanisms Underlying Replication Asynchrony

  1. Top of page
  2. Abstract
  3. Nucleotypic Effect and Asynchrony of DNA Replication Program in Eukaryotes
  4. Asynchronous Replication and Mutation Rates in Vertebrates
  5. Mutation Rates and Gene Location Effects in Fungi and Invertebrates
  6. DNA Repair Systems in Heterochromatin
  7. Checkpoint Function and Mechanisms Underlying Replication Asynchrony
  8. Epigenetic Regulation of Replication Timing
  9. Role of Ribonucleotide Reductase in Replication Asynchrony
  10. Genome Size, Checkpoint Proficiency, and Mutation Rate Heterogeneity
  11. Genome Size and Rates of Speciation
  12. Transposons and DNA Repair Systems
  13. ACKNOWLEDGMENTS
  14. LITERATURE CITED

The regulation of replication timing and asynchrony during the cell cycle is a complicated and poorly understood process that involves multiple factors, including cis-acting DNA sequences and chromatin modifiers (Yompakdee and Huberman 2004; Göndör and Ohlsson 2009; Pope et al. 2010). In yeast, for example, centromeric heterochromatin replicates early, whereas subtelomeric heterochromatin replicates late (Hayashi et al. 2009). In contrast, different telomers in human cells replicate at different times in S-phase (Arnoult et al. 2010). One model to account for replication timing in euchromatin and heterochromatin proposes that MCM proteins, which are essential for replication initiation, are more abundant at more efficient, earlier-firing replication origins, whereas heterchromatin represses MCM activity in late-replicating DNA (Fox and Weinreich 2008; Rhind et al. 2010; Yang et al. 2010). Alternatively, the abundance, or “occupancy,” at replication origins in yeast by the DNA replication initiation factor Cdc45 regulates origin efficiency and DNA replication timing (Wu and Nurse 2009); whereas histone methylation and acetylation determine the time of Cdc45 association with replication origins, and hence when they fire in the S-phase (Pryde et al. 2009).

In support of the proposal that replication timing depends on origin efficiencies, the Chk1/Rad53 intra-S checkpoint effectors repress replication origins in late replicating DNA during a normal S-phase (Santocanale and Diffley 1998; Miao et al. 2003), indicating that checkpoint function plays an important role in regulating replication asynchrony and S-phase progression (Maya-Mendoza et al. 2007; Niida et al. 2007). Chk1 negatively regulates Cdc45 (Liu et al. 2006); and Cdc45 depletion suppresses apoptosis in Chk1-depleted cells (Rodriguez et al. 2008), indicating that Chk1 mediates replication initiation via Cdc45 (Meuth 2010). Moreover, Chk1 is known to interact negatively with the initiation factor Cdc7, which activates MCM proteins during replication initiation (Montagnoli et al. 2008). Cdc7 levels, for example, increase when Chk1 is downregulated (Petermann et al. 2010; Fig. 3). Whether Chk1 directly inhibits late-firing replication origins via Cdc7 is unclear; but a direct interaction between Rad53 and Cdc7 in yeast at replication origins has been established (Dohrmann and Sclafani 2006). These observations are consistent with the proposal that origin efficiency also depends on Hsk1 (Cdc7) in both lower and higher eukaryotes (Patel et al. 2008).

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Figure 3. The role of checkpoint effectors and ribonucleotide reductase (RNR) in regulating Rs. Chk1 promotes replication fork movement at two levels: by inhibiting late firing origins and by stimulating RNR. The molecular details underlying checkpoint-mediated inhibition of replication initiation and simultaneous stimulation of replication elongation are not fully understood; but evidence is emerging that Chk1 interacts either directly or indirectly with the replication initiation factor Cdc7:Dbf4, which is required for activating factors involved in initiating DNA synthesis (Cdc45, MCM, etc.). Other factors such as the global S-phase transcription factor E2F and the Skip-Cullin-F box proteasome also play critical roles in the pathway regulating Rs. For simplicity, regulation of replication initiation is represented by Chk1 in metazoa (A) and Rad53 in yeast (B). Early origin activation imposes a block on late origin activation through a checkpoint-mediated function (ATR/Mec1) in normal S-phase. Origin initiation results in RPA-associated ssDNA, which stimulates the ATR/Mec1 pathways. Chk1/Rad53 inhibit Cdk2 and other SPKs via Cdc25A phosphorylation and degradation. The SPKs positively regulate Cdc45 and MCM activity. Loading of Cdc45 in complex with MCMs and other proteins (not shown) activates replication origins. Both Rad53 and Chk1 at the same time upregulate RNR and control its activity at multiple levels, including transcription regulation (Rfx and Crt1) and RNR enzymatic activity (Dun1 and Sml1). In this manner, the two main parameters governing the kinetics of DNA replication, frequency of replication initiation, and replication elongation rates are carefully balanced and coordinated during S-phase. Rad53/Chk1 also coordinate homologous recombination and DNA replication, and consequently act to limit the mutation rate and enhance error-free DNA repair efficiency in euchromatin (see Herrick and Bensimon 2008 for further details).

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Several recent experiments in metazoa confirm a role for Chk1 in modulating replication asynchrony. Inhibiting Chk1 induces a subset of late-firing replication origins to fire earlier in S-phase, indicating that the replication-timing program at these replication origins is disrupted when Chk1 is inhibited (Costanzo et al. 2003; Miao et al. 2003; Marheineke and Hyrien 2004; Shechter et al. 2004; Syljuåsen et al. 2005; Karnani and Dutta 2011). Conversely, mutations in UbcH7, a ubiquitin conjugating enzyme associated with the SCF (Skp1-Cul1-F-box) proteasome that controls S-phase progression, result in concomitant upregulation of Chk1 and delayed S-phase progression (Whitcomb et al. 2009). Mutations in a gene involved in the circadian cycle in humans (HCLK) likewise display a prolonged S-phase that is related to Chk1 stabilization (Collis et al. 2007). Finally, the nutrient pathway transducer Akt/PKB negatively regulates Chk1 at the G2/M transition (King et al. 2004; Xu et al. 2010), indicating that the ATR-Chk1 pathway is affected by multiple factors at different stages of a normal cell cycle.

Epigenetic Regulation of Replication Timing

  1. Top of page
  2. Abstract
  3. Nucleotypic Effect and Asynchrony of DNA Replication Program in Eukaryotes
  4. Asynchronous Replication and Mutation Rates in Vertebrates
  5. Mutation Rates and Gene Location Effects in Fungi and Invertebrates
  6. DNA Repair Systems in Heterochromatin
  7. Checkpoint Function and Mechanisms Underlying Replication Asynchrony
  8. Epigenetic Regulation of Replication Timing
  9. Role of Ribonucleotide Reductase in Replication Asynchrony
  10. Genome Size, Checkpoint Proficiency, and Mutation Rate Heterogeneity
  11. Genome Size and Rates of Speciation
  12. Transposons and DNA Repair Systems
  13. ACKNOWLEDGMENTS
  14. LITERATURE CITED

Consistent with a complex network of regulators converging on the intra-S checkpoint to control replication asynchrony, a number of studies have shown that both histone and DNA methylation affect replication timing in a manner related to the ATR-Chk1 pathway (Unterberger et al. 2006; Peng and Karpen 2009). A branch of the ATR checkpoint pathway, for example, activates the human MLL methyltransferase, which negatively regulates Cdc45 occupancy at late-firing replication origins (Liu et al. 2010). In addition, a mutant of the DNA methyltransferase DNMT3 results in earlier replication of normally late replicating DNA (Hassan et al. 2001). Other studies revealed that in Xenopus laevis, CpG methylation inhibits DNA replication (Harvey and Newport 2003). Finally, overexpression of the JMJD2A/KDM4A de-methylase in cultured human cells and in vivo in C elegans antagonizes heterochromatin binding protein 1 (HP1), and results in an increase in the number of active replication forks, faster S-phase progression and a change from normally late to early replication of the satellite 2 DNA locus on chromosome 1 in human cells (Black et al. 2010).

These effects are remarkably similar to abrogation of Chk1 and its impact on replication timing (Miao et al. 2003; Marheineke and Hyrien 2004; Syljuåsen et al. 2005), suggesting that modification of either DNA or histone methylation status, in coordination with histone acetylation (Unnikrishnan et al. 2010), promotes a more open chromatin configuration and consequently a change in the density of replication origin initiation events and/or a change in the rate of replication fork movement (Housman and Huberman 1975; Takebayashi et al. 2005). Consistent with that proposal, depleting cells of the JMJD2A/KDM4A de-methylase slowed DNA replication, and resulted in elevated rates of DNA damage-induced apoptosis in an ATR-dependent, but ATM-independent, manner (Black et al. 2010). Likewise, Chk1 function appears to depend on DNMT1 maintenance methylation of DNA (Ha et al. 2011), suggesting a complex and as yet to be elucidated interplay between checkpoint function and both DNA and histone methylation status in the regulation of the timing of origin activation during S-phase (Liu et al. 2010; Karnani and Dutta 2011; Liang et al. 2011).

Chk1 is also involved in the developmental regulation of replication asynchrony. Embryonic development is associated with epigenetic changes to chromatin that result in alterations in gene transcription programs (Li et al. 2010). In Drosophila, for example, the lengthening of S-phase in embryos coincides with global changes in gene transcription, and depends on the increasing shift to later, and hence more asynchronous, replication of heterochromatic satellite DNA (Fig. 2B; Shermoen et al. 2010). The imposition of a late-replicating regime occurs as embryos enter the mid-blastula transition (MBT), a developmental phase that is associated with a transient upregulation of the intra-S checkpoint pathway Mei1/grapes (ATR/Chk1 in vertebrates) (Sibon et al. 1997; McCleland et al. 2009). Developmental slowing of S-phase in X. laevis has also been shown to depend on the activity of Chk1 (Shimuta et al. 2002; Carter and Sible 2003; Conn et al. 2004; Adjerid et al. 2008), indicating that vertebrates and invertebrates both rely on the intra-S checkpoint to monitor and control replication timing during normal S-phase and during development.

Role of Ribonucleotide Reductase in Replication Asynchrony

  1. Top of page
  2. Abstract
  3. Nucleotypic Effect and Asynchrony of DNA Replication Program in Eukaryotes
  4. Asynchronous Replication and Mutation Rates in Vertebrates
  5. Mutation Rates and Gene Location Effects in Fungi and Invertebrates
  6. DNA Repair Systems in Heterochromatin
  7. Checkpoint Function and Mechanisms Underlying Replication Asynchrony
  8. Epigenetic Regulation of Replication Timing
  9. Role of Ribonucleotide Reductase in Replication Asynchrony
  10. Genome Size, Checkpoint Proficiency, and Mutation Rate Heterogeneity
  11. Genome Size and Rates of Speciation
  12. Transposons and DNA Repair Systems
  13. ACKNOWLEDGMENTS
  14. LITERATURE CITED

In yeast, replication asynchrony also increases when ribonucleotide reductase, which is universally essential for dNTP synthesis, is inhibited; but the overall replication program remains largely unaffected (Alvino et al. 2007). The slowing of S-phase without alteration of the replication program suggests that the level of RNR activity restricts the pace of progression through S-phase of the cell cycle. In a normal S-phase in metazoans, Chk1 is responsible for delaying replication initiation in late replicating DNA, whereas ribonucleotide reductase is rate limiting for DNA synthesis after replication origins fire (Nordlund and Reichard 2006). Chk1 promotes replication fork progression by inhibiting origin firing and by upregulating RNR (Lubelsky et al. 2005; Petermann et al. 2010), suggesting that Chk1 and RNR interact to coordinate replication origin firing and replication fork rate in a manner that regulates the observed constancy of replicon duplication rates (Lubelsky et al. 2005; Herrick and Bensimon 2008; Naruyama et al. 2008; Zhang et al. 2009b). Chk1 (Rad53 in Saccharomyces cerevisiae) and RNR thus play an important and coordinated role in determining the degree of replication asynchrony during a normal cell cycle in eukaryotes (Fig. 3).

Evidence in support of a role for the checkpoint and RNR in regulating replication asynchrony comes from a recent study in yeast. A screen for mutants with altered S-phases revealed three classes of factors involved in determining the length of S-phase (Koren et al. 2010). The three classes of mutant corresponded to mutations in genes involved in cell-cycle regulation (Sic1, Clb5, and Dia2), DNA replication (Mrc1, DNA polymerases, etc.), and nucleotide metabolism (RNR, etc.). The lengthening of S-phase in these mutants did not result from an altered replication program, indicating that replication asynchrony alone was affected. In the Mrc1 mutant, in contrast, the replication program was affected and more replication origins fired than normal. Mrc1 operates at the replication fork and signals DNA damage to activate the Rad53 intra-S checkpoint pathway that upregulates RNR. Hence, RNR activity, in association with checkpoint functions and cell cycle regulators, controls the pace of replicon activation in yeast, and consequently, the overall length of S-phase. What then is the relationship between genome size and checkpoint function?

Genome Size, Checkpoint Proficiency, and Mutation Rate Heterogeneity

  1. Top of page
  2. Abstract
  3. Nucleotypic Effect and Asynchrony of DNA Replication Program in Eukaryotes
  4. Asynchronous Replication and Mutation Rates in Vertebrates
  5. Mutation Rates and Gene Location Effects in Fungi and Invertebrates
  6. DNA Repair Systems in Heterochromatin
  7. Checkpoint Function and Mechanisms Underlying Replication Asynchrony
  8. Epigenetic Regulation of Replication Timing
  9. Role of Ribonucleotide Reductase in Replication Asynchrony
  10. Genome Size, Checkpoint Proficiency, and Mutation Rate Heterogeneity
  11. Genome Size and Rates of Speciation
  12. Transposons and DNA Repair Systems
  13. ACKNOWLEDGMENTS
  14. LITERATURE CITED

As the amount of DNA increases in the cell, the number of replication origins must also increase, and so must the total demand on the precursors of DNA synthesis. dNTP pool imbalances, however, are highly mutagenic (Lis et al. 2008), and abnormally high levels of dNTPs inhibit cell cycle progression (Mathews 2006; Chabes and Stillman 2007; Xu et al. 2008). Consequently, dNTP pools are sufficient to support only a few minutes of DNA synthesis during a 3- to 15-h S-phase (Mathews 2006; Nordlund and Reichard 2006). The number of active replicons, corresponding to about 15% of the genome (Shaw et al. 2010), must therefore likewise be restricted at any given time in S-phase (Hand 1978; Berezney et al. 2000; Woodfine et al. 2004). Larger genomes will be expected then to rely on more proficient intra-S checkpoint systems to inhibit the greater number of later firing origins that potentially can disrupt S-phase with catastrophic consequences (Syljuåsen et al. 2005).

Currently, it is unknown if checkpoint functions vary significantly with genome size, although circumstantial evidence supports such a proposal. Emerging evidence suggests that the temporal order of origin activation in yeast, for example, is more relaxed than in higher eukaryotes with larger genomes. Late firing origins in S cerevisiae frequently fire in early S-phase; but early firing origins rarely fire in late S-phase, again indicating different replication regimes (Yang et al. 2010). In the yeast Schizosaccharomyces pombe, abrogating the checkpoint affects only a few late replication origins (3%; Mickle et al. 2007), which suggests weaker checkpoint-mediated inhibition of late firing origins. Interestingly, the checkpoint-constrained replication origins locate to telomeric heterochromatin regions that are enriched in histone H3K9 methylation [Correction made here after initial online publication]. A relaxation of replication timing control in late replicating heterochromatin in Physarum polycephalum (C-value 0.3 pg) has also been suggested (Cunningham and Dove 1993; Diller and Sauer 1993; Bénard et al. 1996), which would be consistent with a weaker inhibition of late-firing replication origins in organisms with small genomes.

A weaker checkpoint in species with smaller genomes might explain the paradox that mutation rates in yeast and C. elegans are more uniform than in other eukaryotes, yet yeast, as do metazoa, display higher rates of diversification in late replicating compared to early replicating DNA (Fox et al. 2008). How can yeast exhibit similar replication timing effects, and yet have more uniform mutation rates than vertebrates? The proposal was made that the yeast replication-timing program is more mutable, and thus allows regions of the genome to switch from late to early replication faster than a polymorphism fixes in the population (Fox et al. 2008). The result will be more uniform mutation rates in yeast that blur the correlation between mutation rate and replication timing (Fig. 4A,B). In contrast, species with more static, or stable, replication programs, and hence stronger checkpoint functions, will display more heterogeneous mutation rates within the genome. Whether a more relaxed intra-S checkpoint can explain the more uniform mutation rates in fungi and worms remains to be demonstrated.

image

Figure 4. Hypothetical effect of genome size on DNA repair efficiency and mutation rate. (A) Cells with larger genomes, or higher fractions of non-coding DNA (ncDNA, or heterochromatin), are expected to have more proficient checkpoint function than cells with smaller genomes, and consequently will have more efficient DNA repair systems. Although the mutation rate per genome is represented here as the same (total number of diamonds), the larger amounts of heterochromatin have a stronger “buffering” effect on the mutation rate in gene rich euchromatin, because of a proportionally stronger checkpoint function and more efficient DNA repair systems. In yeast, for example, inefficient late firing origins can sometimes fire early, possibly because the lower fraction of ncDNA results in a relatively weaker checkpoint response and shorter relative repair times. Consequently, fungi will have more uniform mutation rates than vertebrates and invertebrates. (B) Replication-timing “partitions” mutation rates into different spatio-temporal compartments of the genome. Early replicating euchromatin initiates DNA replication (ovals) at early firing replication origins, and exposes mutation-sensitive ssDNA. In response to initiation and the formation of ssDNA, the checkpoint effector Chk1 inhibits late firing origins (black dots) in heterochromatic regions. As S phase advances, mutations (diamonds) accumulate in early replicating DNA (euchromatin). Asynchrony of DNA replication, however, allows more time and hence more efficient repair. The formation during DNA replication of a sister chromatid (ovals) is also essential for homologous recombination (HR) repair. Late replicating DNA in contrast is refractory to HR and other DNA repair systems. Consequently, mutation rates in the genome will increase monotonously with replication timing, and late replicating genes will experience higher mutation rates than earlier replicating genes.

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Genome Size and Rates of Speciation

  1. Top of page
  2. Abstract
  3. Nucleotypic Effect and Asynchrony of DNA Replication Program in Eukaryotes
  4. Asynchronous Replication and Mutation Rates in Vertebrates
  5. Mutation Rates and Gene Location Effects in Fungi and Invertebrates
  6. DNA Repair Systems in Heterochromatin
  7. Checkpoint Function and Mechanisms Underlying Replication Asynchrony
  8. Epigenetic Regulation of Replication Timing
  9. Role of Ribonucleotide Reductase in Replication Asynchrony
  10. Genome Size, Checkpoint Proficiency, and Mutation Rate Heterogeneity
  11. Genome Size and Rates of Speciation
  12. Transposons and DNA Repair Systems
  13. ACKNOWLEDGMENTS
  14. LITERATURE CITED

Genome size varies over 200,000-fold in eukaryotes (Gregory et al. 2007). A genome size effect on speciation rates has been considered for some time (Kraaijeveld 2010), but the significance of the correlations between genome size, mutation rate, and speciation rate remains unclear. Genome size often correlates negatively with species richness in different taxa (Vinogradov 2004; Knight et al. 2005; Olmo 2006; Fig. 5A–C), whereas rates of molecular evolution frequently correlate positively with taxonomic diversity (Lancaster 2010; Venditti and Pagel 2010). More species-rich taxa therefore reflect smaller genomes and faster rates of diversification, suggesting a possible generation time (GT) effect on mutation rates (Thomas et al. 2010) The negative correlation between species richness and genome size can be attributed to the fact that larger genomes are more prone to mutation (Lynch 2008, 2010), and hence are inherently more unstable and have diminished fitness. Alternatively, organisms with larger genomes tend to have larger body size and consequently reduced effective population size, which results in genetic drift having a greater impact than selection on the evolution of larger organisms (Lynch and Conery 2003).

image

Figure 5. Correlation between genome size and cell cycle times, and frequency distributions of plant species according to different genome sizes (Knight et al. 2005; Francis et al. 2008). (A) Cell cycle durations correlate with genome size in plants. A clear nonlinear effect of genome size on cell cycle time is apparent above a genome size of 25 pg (from: Francis et al. 2008; reprinted with permission from the publisher). (B) Zoom in on eudicots, revealing the correlation for C-values between 1 and 14 pg. (C) Large genomes are rarer and have fewer species (from: Knight et al. 2005; reprinted with permission from the publisher). Most plant species have genomes less than 5 pg in size, indicating that large genomes above 5 pg are maladaptive and tend to be less species rich.

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Consistent with a nucleotypic effect, species with faster GTs, or the number of duplications a genome undergoes, also tend to have smaller genomes (Knight et al. 2005; Francis et al. 2008) (Fig. 5A,B), whereas reductions in genome size have been found to coincide with adaptive radiations (Kraaijeveld 2010). A GT effect has been proposed, for example, to explain higher rates of mutation found in zebra finch compared to chicken (Nam et al. 2010). The same study also showed that the songbird lineage, compared to the chicken lineage, has undergone more frequent adaptive evolution in genes involved in cognitive processes. In agreement with a nucleotypic effect on GT and mutation rate, an increase in GT during primate evolution has been proposed to result in a decrease in mutation rates corresponding to DNA replication-dependent errors (Hwang and Green 2004).

Perhaps more intriguingly, several studies report a coincidence between massive bursts of transposition and adaptive radiations (Feschotte and Pritham 2007; Rebollo et al. 2010). Abrupt changes in environmental resistance and stress might then positively influence speciation rates through genome-level molecular adaptations driven by transposons periodically invading euchromatin (Turner 2009). This appears to be the case for the human lineage, which displays higher than expected rates of transposition and correspondingly faster rates of evolution, in particular with respect to the brain (Britten 2010). What might be the relationship, if any, between GT- dependent variations in mutation rate and transposon-driven changes in evolutionary rates?

Transposons and DNA Repair Systems

  1. Top of page
  2. Abstract
  3. Nucleotypic Effect and Asynchrony of DNA Replication Program in Eukaryotes
  4. Asynchronous Replication and Mutation Rates in Vertebrates
  5. Mutation Rates and Gene Location Effects in Fungi and Invertebrates
  6. DNA Repair Systems in Heterochromatin
  7. Checkpoint Function and Mechanisms Underlying Replication Asynchrony
  8. Epigenetic Regulation of Replication Timing
  9. Role of Ribonucleotide Reductase in Replication Asynchrony
  10. Genome Size, Checkpoint Proficiency, and Mutation Rate Heterogeneity
  11. Genome Size and Rates of Speciation
  12. Transposons and DNA Repair Systems
  13. ACKNOWLEDGMENTS
  14. LITERATURE CITED

Genome size increases with the amount of TE and repetitive DNA contained in the genome. Transposons interact positively and negatively with DNA repair systems, and can influence the type of DSB repair system employed when damage occurs (Izsvak et al. 2009). The DNA repair hypothesis explains different rates of molecular evolution in terms of varying efficiencies of DNA repair (Britten 1986; Baer et al. 2007). Accordingly, genome size can affect mutation rates and speciation rates at two levels: (1) negatively, by enhancing DNA repair efficiencies due to genome size-dependent checkpoint-mediated extension of S-phase (GT), as proposed here; and (2) positively, by increasing the probability of either adaptive or deleterious transposition events. The balance between these two forces might adjust the molecular clock in a species-specific manner. The first proposal can be easily tested in yeast, for example; while the second is garnering a growing consensus as the hypothesis of transposon-driven evolution (Kazazian 2004; Feschotte and Pritham 2007).

In yeast, Ty transposition, although independent of the checkpoint, is stimulated by checkpoint function (Curcio et al. 2007). Mutants defective in the S-phase regulatory factor Rtt101 recover poorly from checkpoint function (Zaidi et al. 2008), and experience bursts of transposition. Moreover, transposition events depend on RNR and increase when RNR is up-regulated during DNA damage and repair (O’Donnell et al. 2010). Indeed, Ty transposons can participate in NHEJ-directed DNA repair (Teng et al. 1996); and can mediate gene amplification when unscheduled DNA replication occurs (Green et al. 2010), indicating that a disrupted S-phase can result in elevated adaptive, or maladaptive, mutation rates. A corollary to the transposon hypothesis of evolution then is that many, if not all, adaptive radiations in eukaryotes reflect DNA damage events, and are ultimately consequences of adaptations in checkpoint-mediated DNA repair. The evolution of DNA repair systems underlies, in that sense, the evolution of species (Britten 1986).

CONCLUSION

The increase in mutation rate with replication timing in eukaryotes correlates with two well-established phenomena: (1) reduced efficiency in DNA repair systems in heterochromatin; and (2) transition to a distinct late-S replication program after euchromatin replication is completed. The shift to a late replicating regime in both higher and lower eukaryotes is likely due to the change in chromatin status between early and late replicating DNA. Cdk1/cyclinA2, for example, might be required in addition to Cdk2/cyclinE to mediate histone modifications such as acetylation and demethylation that result in HP1 redistribution and relaxation of late replicating heterochromatin. Relaxation of heterochromatin is expected to result in an increase in the sites available for replication initiation, because later replicating heterochromatin contains more DNA (3X), and replicons are more clustered at mid S-phase (3X; Lima-de-Faria 1959; Frum et al. 2009). Consequently, higher levels of mutation-prone ssDNA at mid-to-late S-phase can contribute to an increase in mutations that will be repaired less efficiently (Prendergast et al. 2007; Stamatoyannopoulos et al. 2009; Herrick 2010).

In mice, the transition to a late S-phase replication program is associated with the intra S-phase checkpoint (Katsuno et al. 2009), suggesting that Chk1 activation at mid S-phase is coupled to its inactivation in late S/G2 phases (Herrick 2010). Evidence in support of that proposal comes from a recent study showing that Chk1 phosphorylation simultaneously activates Chk1 and promotes its degradation in a DDB1/Cul4A ubiquitin ligase and proteasome-dependent manner (Zhang et al. 2009a; Guervilly et al. 2011). Downregulation of Chk1 at mid-to-late-S/G2 provides an additional potential explanation for why DNA repair systems are less efficient in repairing DNA in heterochromatin. Downregulation of Chk1 during late S-phase in a normal cell cycle remains, however, to be fully demonstrated.

Importantly, the error-prone DNA repair processes NHEJ and DNA translesion synthesis (TLS) can both occur independently of the ATR/Chk1 checkpoint response (Bi et al. 2006; Andersen et al. 2008), and TLS mediates checkpoint downregulation and resumption of DNA synthesis (Bi et al. 2006; Pillaire et al. 2007; Despras et al. 2010). TLS, which replicates through damaged DNA, takes place predominantly during and after S-phase, and is not dependent on genome duplication (Daigaku et al. 2010). In yeast, the Rev1 TLS factor increases slowly from early to mid S-phase, and then increases rapidly in late S-phase until it peaks in G2 (Waters and Walker 2006). Cell cycle regulation of Rev1 in higher eukaryotes, however, has not been observed, and the exact nature of the kinetics of TLS remains obscure (Shaheen et al. 2010). TLS has also recently been proposed as a possible explanation for the increase in non-CpG mutation rates associated with CpG mutations, the so-called “CpG effect” (Walser and Furano 2010). The observed changes in mutation rate within the genome are thus consistent with a growing reliance on Chk1-independent NHEJ (and possibly TLS (Lis et al. 2008)) in late S/G2 phases, whereas differences across species are consistent with greater reliance on NHEJ as genome size (TE-associated heterochromatin) increases (Table 1).

Together, the observations discussed above suggest that genome size has an impact on mutation rates that operates through checkpoint regulation of replication asynchrony and S-phase progression. Larger amounts of heterochromatin are associated with larger chromosomes and genomes, and larger chromosomes exhibit lower levels of recombination and higher rates of mutation in late replicating compared to early replicating DNA (3% increase in yeast versus 10% in rodents and 22% in primates). The unexpectedly low rates of mutation, for example, in the very late replicating Y chromosome can be attributed to lower recombination rates due to a higher relative fraction of heterochromatin (Pink and Hurst 2010). A negative correlation between chromosome size and sequence divergence has also been observed in birds (Nam et al. 2010). An unambiguous correlation between genome size and mutation rate, however, has not been observed to date (Lancaster 2010).

Nevertheless, it will be interesting to investigate if mutation rates are lower in genes embedded in larger genomes that are, paradoxically, more prone to mutation because of their sizes (Herrick 2011). The proopiomelanocortin gene in lungfish (C-value 74.8 pg), for example, is evolving at a much slower rate than in rodents or primates (C value 2.8 and 3.2 pg; Lee et al. 2006). Rates of diversity at other loci are also exceptionally low (Frentiu et al. 2001). Moreover, mutation rates in nuclear genes decrease up to threefold ranging from yeast, C elegans, D melanogaster to humans, whereas mutations in repetitive DNA increase several fold, respectively (Lynch et al. 2008). Interestingly, C. elegans has a relatively higher nuclear mutation rate compared to the other organisms; but a correspondingly lower repeat DNA mutation rate, suggesting an inverse relationship between the two rates. Together, these observations suggest a differential effect of chromatin status on gene location and mutation rate between early and late replicating DNA within the genomes of different species (Fig. 4B).

Here it has been argued that the late replicating status of heterochromatin and degree of replication asynchrony constitute important components—among many other factors—that affect DNA repair efficiencies, and can explain, in part, why eukaryotes have played host to so much junk in their genomes. Late-replicating heterochromatin acts as a substrate for checkpoint function that depends on Chk1/Rad53-mediated inhibition of late-firing replication origins during a normal S-phase and checkpoint-mediated upregulation of DNA repair systems during a perturbed cell cycle. This raises the intriguing possibility that checkpoint function and TE-associated heterochromatin coevolved symbiotically in a way that mutually benefited both host and parasite, a functional relationship made energetically feasible by the emergence of mitochondria (Lane and Martin 2010). Downregulation of DNA repair systems in late S/G2 phases, for example, might have been an adaptation that erodes TE genetic integrity, while at the same time a growing reliance on NHEJ repair incidentally retained TE sequences in the genome (Table 1). Far from being parasites, TEs and other forms of junk DNA, in competing with gene-rich euchromatin, had the fortuitous consequences of promoting the DNA damage response and repair systems that maintain genome integrity, while increasing species diversity through the adaptations that so-called “junk DNA” made possible.

Associate Editor: D. Fairbairn

LITERATURE CITED

  1. Top of page
  2. Abstract
  3. Nucleotypic Effect and Asynchrony of DNA Replication Program in Eukaryotes
  4. Asynchronous Replication and Mutation Rates in Vertebrates
  5. Mutation Rates and Gene Location Effects in Fungi and Invertebrates
  6. DNA Repair Systems in Heterochromatin
  7. Checkpoint Function and Mechanisms Underlying Replication Asynchrony
  8. Epigenetic Regulation of Replication Timing
  9. Role of Ribonucleotide Reductase in Replication Asynchrony
  10. Genome Size, Checkpoint Proficiency, and Mutation Rate Heterogeneity
  11. Genome Size and Rates of Speciation
  12. Transposons and DNA Repair Systems
  13. ACKNOWLEDGMENTS
  14. LITERATURE CITED