Chromosoma DOI 10.1007/s00412-013-0411-3
REVIEW ARTICLE
DNA replication and homologous recombination factors: acting together to maintain genome stability Antoine Aze & Jin Chuan Zhou & Alessandro Costa & Vincenzo Costanzo
Received: 30 November 2012 / Revised: 27 March 2013 / Accepted: 27 March 2013 # Springer-Verlag Berlin Heidelberg 2013
Abstract Genome duplication requires the coordinated action of multiple proteins to ensure a fast replication with high fidelity. These factors form a complex called the Replisome, which is assembled onto the DNA duplex to promote its unwinding and to catalyze the polymerization of two new strands. Key constituents of the Replisome are the Cdc45Mcm2-7-GINS helicase and the And1-Claspin-Tipin-Tim1 complex, which coordinate DNA unwinding with polymerase alpha-, delta-, and epsilon- dependent DNA polymerization. These factors encounter numerous obstacles, such as endogenous DNA lesions leading to template breakage and complex structures arising from intrinsic features of specific DNA sequences. To overcome these roadblocks, homologous recombination DNA repair factors, such as Rad51 and the Mre11-Rad50-Nbs1 complex, are required to ensure complete and faithful replication. Consistent with this notion, many of the genes involved in this process result in lethal phenotypes when inactivated in organisms with complex and large genomes. Here, we summarize the architectural and functional properties of the Replisome and propose a unified view of DNA replication and repair processes.
Introduction Genome duplication is a key event in the life cycle of all proliferating organisms and its careful control is essential to preserve the physical integrity of chromosomes (Arias and Walter 2007). The main player in this process is the Replisome, an assembly of macromolecular machines that serve A. Aze : J. C. Zhou : A. Costa (*) : V. Costanzo (*) Clare Hall Laboratories, London Research Institute, South Mimms, Herts EN63LD, UK e-mail:
[email protected] e-mail:
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two essential functions: (1) coupling parental duplex–DNA unwinding with daughter strand synthesis (Macneill 2012) and (2) integrating DNA damage response signals to modulate fork progression, pausing, and restart (Errico and Costanzo 2012). Eukaryotic Replisome assembly occurs in multiple steps, which are timed in accordance with cell cycle cues. During the G1 phase, the origin recognition complex transiently associates with the Cdc6 initiator to recruit a Cdt1•Mcm2-7 heptamer to DNA replication start sites (“origins”) (Boos et al. 2012). The end result of this reaction is the formation of a topological link between duplex DNA and two copies of the hexameric Mcm2-7 helicase, which are found tethered via their N-terminal ends. In this configuration, origins are “licensed” for activation; however, the unwinding function of the Mcm2-7 enzyme remains dormant (Remus et al. 2009). Upon entry into S phase, multiple factors are recruited to activate the replication origins by either associating with, or chemically modifying, the Mcm2-7 helicase (reviewed in Labib 2010). The events that lead to the opening of duplex DNA are still poorly understood at a molecular level. According to the current consensus model, the two Mcm particles are thought to move apart following DNA melting, to travel at the front of the Replisome (Botchan and Berger 2010; Yardimci et al. 2010). Helicase activation depends on the association of the Replisome component, Cdc45 (Tercero et al. 2000), with Mcm2-7 and the concomitant recruitment of the GINS assembly (Gambus et al. 2006) (together forming the CMG) (Moyer et al. 2006). Multiple factors contribute to this event. For example, a phospho-protein assembly acts as a GINS•Cdc45 chaperone (called Sld2•Sld3•Dpb11 in yeast) (Zegerman and Diffley 2007), while also promoting origin deposition of the leading strand polymerase Pol ε (Muramatsu et al. 2010). Another key player is Mcm10, which transiently associates with the CMG to promote polymerase α/Primase origin
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association and possibly to aid in DNA opening (Kanke et al. 2012; van Deursen et al. 2012). Additional factors travel with the Replisome at the fork. One example is the replication pausing complex, composed of Tipin, Tim1, and And1 (Errico et al. 2009) and Claspin (Nedelcheva et al. 2005), structural proteins that tether the CMG helicase to the replicative polymerases and couple their activities (Fig. 1). These factors play a primary role in maintaining chromosomal integrity under replication stress conditions, as they keep the CMG from translocating when replicative polymerases stall (Errico and Costanzo 2012). Other Replisome-associated factors are Topoisomerase IB (Gambus et al. 2006) that relieves the positive supercoils accumulating ahead of the replication fork (Vos et al. 2011) and the FACT histone chaperone complex (Gambus et al. 2006), which has been implicated in parental nucleosome disassembly ahead of the fork or in daughter strand nucleosome reassembly at the back of the Replisome (Abe et al. 2011; Winkler and Luger 2011). In this review, we aim at building an architectural framework to help describe the function of the Replisome unperturbed or engaged in the interaction with the DNA repair machineries. We discuss the known structural features of the isolated Replisome components as well as their assemblies, with a focus on the mechanisms for helicase activation and inactivation, and the roles of the Replisome pausing complex in modulating helicase/polymerase crosstalk. We then describe the known relationships between the Replisome and the DNA repair machinery focusing on the role of DNA damage response proteins and homologous recombination factors in unchallenged and perturbed DNA replication.
Mcm2-7 Primase
AAA+
GINS
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Pol α
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Tipin Tim1 Pοl δ
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Fig. 1 Interactions involving some key Replisome components. The Ctf4/And1-Tipin-Tim1-Mrc1/Claspin complex plays an important role in bridging between the Cdc45-Mcm2-7-GINS helicase and the replicative polymerases alpha, delta, and epsilon
The building of a Replisome Mcm2-7 activation requires large structural rearrangements The engine of the replicative helicase is formed by six distinct, however, related polypeptides (Mcm2, 3, 4, 5, 6, 7) (Vijayraghavan and Schwacha 2012) that contain two domains: an N-terminal DNA interacting collar (Fletcher et al. 2003) and a C-terminal motor domain that belongs to the superfamily of AAA+ ATPases (Brewster et al. 2008; Neuwald et al. 1999). These modules form two stacked rings that spool DNA through their aligned central cavity (Fletcher et al. 2003) with a 3’→5’ polarity (Chong et al. 2000; Kelman et al. 1999) via a steric exclusion mechanism (Fu et al. 2011) and through an intricate allosteric network involving four pore loops per protomer (Barry et al. 2009). Indeed, limited DNA unwinding activity by the isolated Mcm2-7 complex has been observed in vitro, only in a small number of species and in a narrow window of buffer conditions (Bochman and Schwacha 2008), consistent with the idea that the recruitment of activators is required to stimulate the helicase function (Botchan and Berger 2010). In agreement with this notion, studies on recombinant Drosophila (Ilves et al. 2010; Moyer et al. 2006) or human proteins (Kang et al. 2012) indicate that both the ATPase and DNA unwinding functions are greatly enhanced when the Mcm2-7 is co-expressed with GINS and Cdc45, whose association must induce activating structural rearrangements within the helicase subunits (Ilves et al. 2010). Three-dimensional electron microscopy explains the nature of this conformational transition. The isolated Mcm2-7 complex was shown to form an open lock-washer ring containing a gap in between Mcm2 and Mcm5 (Costa et al. 2011; Lyubimov et al. 2012), also predicted in earlier biochemical studies (Bochman and Schwacha 2008, 2010). In the context of the CMG holoenzyme instead, the Mcm2-7 helicase is closed, with Cdc45 and GINS binding across the Mcm2/5 gate, effectively working as a latch that compresses the hexameric ring, sealing its gap (Costa et al. 2011). Unique features of the Mcm2-7 ATPase assembly explain the mechanistic implications of activator binding. Hexameric, ring-shaped helicases all contain bipartite active sites, with catalytic residues contributed by neighbouring, closely packed subunits, and usually need a set of six functional ATPases for unwinding (Lyubimov et al. 2011). The Mcm2-7 is peculiar in that it tolerates inactivating changes in many of its subunits, while still working as a helicase; however, it requires a functional Mcm5/2 active site for unwinding (Bochman and Schwacha 2008, 2010; Ilves et al. 2010). So, as they force the motor ring into a closed configuration and lock the 2/5 gate, GINS and Cdc45 turn on the unwinding function of the Mcm2-7 helicase by acting to reconfigure the most critical ATPase active site in the hexamer, located at the 2/5 interface (Costa et al. 2011). This mechanism can be employed during
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origin activation but could also be used to modulate fork progression when the Replisome encounters a DNA lesion (Hashimoto et al. 2011; Ilves et al. 2012). In particular, the encounter of the helicase with a nick or gap in the template induces a dissociation of the GINS subunit, leaving Cdc45 still bound to the Mcm2-7 helicase. This process might lead to a slowing down/halting of the helicase progression in the presence of DNA damage. The mechanism by which this takes place is still unclear and might involve the participation of the DNA damage checkpoint at local level (Hashimoto et al. 2011). GINS, the central nexus in the eukaryotic replication fork The GINS hetero-tetramer is composed of four gene products (Sld5 and Psf1/2/3) evolved from one common archaeal ancestor through two sequential gene duplication events (Kamada 2012). The architecture of the GINS assembly reflects its evolutionary history, as the four polypeptides form a pseudo-twofold symmetric assembly, with an elongated structure containing two small, coaxial apertures of unknown function (Chang et al. 2007; Choi et al. 2007; Kamada et al. 2007). Accumulating evidence indicates that GINS not only works to activate the Mcm2-7 helicase but also has a structural role in connecting multiple key Replisome components (Gambus et al. 2009; Muramatsu et al. 2010). Combined structural and genetic studies provide important insights into the architectural role of GINS. The preservation of three surface-exposed residue patches have been identified as vital for yeast survival, these include: (1) the Psf2 α helical domain, contacting the Mcm DNA interacting collar, (2) the Psf2 C-terminal beta domain interfacing with Cdc45, and (3) the water-accessible face of Sld5 (Choi et al. 2007), which does not engage in any CMG contact but rather appears to project from the CMG core, as the apex of a lateral protuberance, poised to interact with other Replisome factors (Costa et al. 2011). Indeed, GINS has been shown to associate with multiple components involved in both leading and lagging strand synthesis. For example, studies in yeast indicate that GINS is recruited onto origins together with Pol ε (Muramatsu et al. 2010) by directly contacting the essential, non-catalytic subunit B, a configuration that is likely kept in the context of the moving Replisome. A direct interaction between GINS and Pol α/Primase has been detected in vitro by surface plasmon resonance (De Falco et al. 2007), while Ctf4 (the yeast ortholog of the Replisome pausing factor, And1—also see below) has been implicated in bridging the CMG and Pol α/Primase by binding the GINS subunits, Sld5 and Psf2 (Gambus et al. 2009). It is remarkable that GINS, a small ~100-kDa assembly, can interface with two distinct replicative polymerases, Pol α and ε (although whether these interactions are concomitant is unknown). Multiple lines of evidence support the notion of a
role for GINS as a polymerase-bridging factor. For example, a biochemical study on human proteins indicates that all three replicative polymerases co-elute with GST-tagged GINS, although this interaction is weak and fails to survive a glycerol gradient sedimentation step (Bermudez et al. 2011). Further support for direct leading and lagging strand coordination by GINS comes from the ancestral archaeal replication system. For example, GINS purified from Sulfolobus solfataricus can be pulled down by recombinant Primase (Marinsek et al. 2006), while endogenous, HIS-tagged Thermococcus kodakarensis GINS can be co-purified with the replicative DNA polymerase Pol D (Li et al. 2010) (an enzyme in part related to the catalytic subunit of Pol ε, see below) (Johansson and Macneill 2010). Overall, biochemical data on the archaeal Replisome mirror the observations in the eukaryotic system, supporting the notion that GINS is a key architectural factor that tethers the Mcm2-7 helicase to the replicative polymerases. Cdc45, a catalytically dead exonuclease Despite its key role during DNA replication initiation, fork elongation, and pausing, progress in our understanding of the molecular function of Cdc45 has been slow. Some structural insights have been derived from a small angle X-ray scattering study (Krastanova et al. 2012) on the isolated human protein and from the EM structure of the full CMG complex (Costa et al. 2011). According to both studies, Cdc45 contains a globular core with protruding arms. In the context of the CMG holoenzyme, these arms interface either with the GINS assembly or the N-terminal domain of Mcm2 and 5, engaging in intimate contacts that keep Cdc45 well anchored to the helicase. Like GINS, the Cdc45 protomer plays multiple roles within the Replisome, which go beyond a structural function during the activation of the replicative helicase. For example, Cdc45 has been implicated in directly interacting with a number of replication factors including Sld3 (Kamimura et al. 2001) during a key step in the cascade of events that lead to origin activation or Mrc1/Claspin (Lee et al. 2003) that coordinates helicase and polymerase activities during fork elongation. Interestingly, the N-terminal domain of Cdc45 is homologous to the phosphodiesterase domain of the prokaryotic RecJ factor (Sanchez-Pulido and Ponting 2011), a single-stranded specific 5’→3’ exonuclease (Makarova et al. 2012), which is involved in the RecF double-strand break (DSB) repair pathway in bacteria (Lovett and Clark 1984). Close inspection of the catalytic site sequence of Cdc45 highlights the presence of conserved inactivating mutations that target key residues involved in catalytic metal coordination in RecJ (Krastanova et al. 2012; Makarova et al. 2012). In agreement with this observation, recent biochemical studies on recombinant human Cdc45, reported single-stranded DNA binding but failed
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to detect any nuclease or duplex–DNA binding activity (Krastanova et al. 2012), suggesting that this factor might employ a defunct exonuclease domain as a scaffold for tracking on one strand at the replication fork. The Cdc45/RecJ homology provides important insights into the evolution of the eukaryotic DNA replication machinery. Indeed, many archaea contain a RecJ homolog that can be purified in a complex with endogenous GINS (Marinsek et al. 2006). Although in some archaea, RecJ contains inactivating amino acid changes as in Cdc45 (Makarova et al. 2012) (or lacks the phosphodiesterase active site altogether) (Marinsek et al. 2006), other species contain a catalytically active enzyme (Li et al. 2011). These data suggest that a prokaryotic DSB repair system was hijacked (independently, more than once) in archaea to become part of the DNA replication machinery, a role maintained through evolution. Taken together with the dissociation of the GINS factor following an encounter with a nick in the template, which forms a DSB at replication forks, this finding suggest that the DSB repair machinery is intimately linked to the replication process. The role of the single-stranded DNA-binding function of Cdc45 during DNA replication remains unclear. Given the marginal localization of Cdc45 within the CMG helicase, offset of the Mcm2-7 DNA interacting channel (Costa et al. 2011), it can be postulated that Cdc45 might be involved in tracking on one of the tails of the moving replication fork. Alternatively, Cdc45 might act as a brake for the helicase during fork pausing or collapse. For example, detection of a DNA lesion could trigger the disassembly of the GINS activator factor, removing the latch from Mcm and causing the Mcm2-5 gate to open. This event would cause the disruption of a topological link between the helicase and the leading strand. To prevent release of the helicase from the stalled replication fork, Cdc45 could employ its dead exonuclease scaffold to clamp onto the DNA, in a “parked” configuration, waiting for Replisome re-assembly and ready for fork restart (Fig. 2). Two studies support the notion of a helicase break role for Cdc45: a recent work on yeast proteins (Bruck and Kaplan 2013), showing helicase polymerase uncoupling for a DNA-binding-deficient Cdc45, and the studies on collapsed forks in Xenopus egg extracts (Hashimoto et al. 2011). Polymerase α/Primase The leading and lagging strand polymerases, Pol ε and δ, elongate DNA polymers starting from RNA–DNA primers synthetized by the Pol α–Primase complex. De novo synthesis can only be catalyzed by the Primase that produces short ~7– 12 nucleotide RNA segments subject to limited extension by the DNA polymerase α (Frick and Richardson 2001). This highly coordinated process occurs in the context of a tetrameric complex containing two Primase subunits (Pri1 and Pri2) and
two Pol α subunits (Pol 1 and Pol 2). A combination of X-ray crystallography and electron microscopy using yeast or the orthologous archaeal proteins provide a good architectural view of this tetrameric assembly (also known as primosome) (Pellegrini 2012). In particular, the archaeal Primase serves as a model for the architecture of the eukaryotic enzyme. Here, the two subunits, named PriS and PriL, form a curved complex, with PriS containing the active site and PriL located distally and not directly involved in catalysis, but rather poised to control the length of the nascent RNA primer (Augustin et al. 2001; Lao-Sirieix et al. 2005a, b). Pol α contains a B family-type catalytic subunit 1 connected to a regulatory subunit 2, also found in Pol δ and ε (Johansson and Macneill 2010). The C-terminal domain of Pol1 interacts with Pol2 forming a stable heterodimeric complex (Klinge et al. 2009) while the very C-terminal tail of Pol1 directly tethers the Primase dimer (Kilkenny et al. 2012). Contacts between the globular catalytic core domain, CTD-Pol1•Pol2 and the Pri1•2 appear to be tenuous, resulting in a highly flexible structure (Nunez-Ramirez et al. 2011), representing a major challenge to high-resolution structural characterization. And1/Ctf4 Ctf4 (And1 in higher eukaryotes) was initially isolated in Saccharomyces cerevisiae via a protein affinity screen as a DNA polymerase α interactor (Miles and Formosa 1992). Although non-essential for cell viability, deletion of Ctf4 leads to defects in DNA replication with cells displaying abnormal morphology and a marked reduction in the rate of DNA replication. Recent evidence indicates that Ctf4/And1 might have a dual function, bridging between the helicase and polymerases within the unperturbed Replisome, but also being part of the replication pausing complex, including Claspin, Tim1 and Tipin, the protein assembly responsible for modulating helicase/polymerase crosstalk during replication fork stalling, pausing, and restart (Errico and Costanzo 2012). Studies in yeast indicate that Ctf4 bridges between the Primase-associated DNA polymerase α and the replicative helicase component GINS (Gambus et al. 2009; Tanaka et al. 2009). In particular, pull-down experiments performed on recombinant proteins indicate that the C-terminal portion of Ctf4 directly interfaces the GINS subunits Psf2 and Sld5 while also interacting with the catalytic subunit of Pol α (Gambus et al. 2009). The yeast Ctf4 interaction network appears recapitulated in metazoan systems. For example in human cells, the Ctf4 ortholog And1 is required for the association of GINS with the Mcm2-7 helicase in human cells (Im et al. 2009) while in Xenopus And1 is found to directly interact with Pol α and bind Tim/Tipin (Errico et al. 2009). In summary, Ctf4/And1
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Cdc45
Mcm2-7 7
Parental Mcm2-7 7 Lagging strand
GINS Cdc45
Pol ε Leading g strand
GINS Pol ε
Fig. 2 A speculative mechanism for helicase halting in response to fork collapse. When the Cdc45-Mcm2-7-GINS helicase encounters a single-stranded DNA lesion, GINS and Pol ε disengage from the Replisome, while Cdc45 and Mcm2-7 remain bound to the replication fork. GINS disengagement likely causes the opening of the Mcm2-7
DNA gate, which in turn could promote extrusion of the leading strand template. The single-stranded DNA binding function of Cdc45 could help maintain the helicase tethered to the collapsed replication fork. According to our model, Cdc45 likely catches the leading strand template, released upon the Mcm2-7 ring opening
appears to not only provide an important architectural link between the replicative helicase and polymerases, but also bridge the Replisome with the Replisome pausing complex that controls Replisome pausing and restart (Errico et al. 2007). The molecular mechanism of fork progression modulation remains to be elucidated.
fork, as a single-stranded DNA tracking element or whether it engages DNA in other contexts, for example during replication initiation (Muramatsu et al. 2010) and/or fork pausing. Surprisingly, studies in yeast indicate that the C-terminal, catalytically dead half of Pol ε is the only domain essential for viability (although cells bearing a truncation of the catalytic domain grow slower) (Dua et al. 1998; Dua et al. 1999; Feng and D'Urso 2001; Kesti et al. 1999). This notion is coherent with the idea that the role of the two tandem polymerase repeats can be uncoupled. Equally complex domain architecture can be found in subunit 2, which contains three recognizable modules. Remarkably, the structure of the N-terminal region resembles the lid of an AAA+ ATPase (Nuutinen et al. 2008). This observation is particularly tantalizing, as Pol ε subunit 2 is a known interactor of the GINS complex (Muramatsu et al. 2010), which in turn works to modulate the opening/closure of the Mcm2-7 DNA gate (Costa et al. 2011). When inactive, Mcm2-7 exists in an open-end configuration, which exposes one AAA+ active site surface, a potential interactor for the Pol ε AAA+ lid-like domain (however, it remains to be tested whether a direct contact between Mcm2-7 and Pol ε occurs). Following a central predicted oligosaccharide/oligonucleotide binding fold, the C-terminal region of subunit 2 contains a calcineurin-like phosphodiesterase domain (yet another dead nuclease domain), a feature common to all eukaryotic replicative DNA polymerases (Johansson and Macneill 2010). Altogether, the complex evolutionary history of the Pol ε multicomponent enzyme is mirrored by an intricate network
Polymerase ε The leading strand polymerase, Pol ε, is a four-member enzyme containing a large catalytic subunit (1), an essential, non-catalytic subunit (2), and two non-essential subunits (3 and 4), characterized by a histone fold motif (Hogg and Johansson 2012). The catalytic subunit forms a large, globular head domain followed by an extended, flexible tail composed of subunits 2, 3, and 4, as shown by cryo-EM studies (Asturias et al. 2006). Although the catalytic subunit belongs to the same B family of polymerases (as do the two other eukaryotic replicative polymerases), this protomer is peculiar in that it contains a C-terminal zinc finger appendix homologous to the archaeal DNA polymerase D, preceded by a tandem repeat of two whole polymerase domains (Tahirov et al. 2009). Whereas the N-terminal repeat of the catalytic subunit contains canonical DNA polymerization and editing functions found in other B-family polymerases, the C-terminal repeat bears inactivating mutations that make it a catalytically dead polymerase module. Similar to Cdc45, it is unclear whether this defunct enzyme is employed in the context of the moving
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of interactions with other replication factors. Two Pol ε protomers contain a catalytically dead DNA processing domain, whose function to date is only partially understood, but most likely involves single-stranded DNA engagement (Tahirov et al. 2009). Claspin/Mrc1 In similar fashion to Ctf4 on the lagging strand, the Claspin factor (mediator of replication checkpoint, Mrc1, in yeast) has been shown to provide a physical link between the DNA helicase and Pol ε on the leading strand (Lou et al. 2008). Indeed, domain mapping experiments have shown that both the N-terminal and C-terminal regions of Mrc1 have a direct role in engaging Pol ε. Interestingly, the N-terminal domain of Mrc1 becomes phosphorylated in response to DNA damage (Alcasabas et al. 2001; Osborn and Elledge 2003), leading to dissociation of this domain from Pol ε while the C-terminal region remains anchored to the polymerase (Lou et al. 2008). The functional consequence of this structural rearrangement remains to be elucidated. Like Ctf4, Mrc1-depleted cells are viable but lead to DNA damage accumulation (Liu et al. 2006) and show a greatly reduced fork progression rate possibly due to the higher frequency of Replisome dissociation from chromatin (Szyjka et al. 2005; Tourriere et al. 2005). Further studies will likely elucidate how cells can survive in the absence of architectural factors such as Claspin or Ctf4, which link the CMG helicase and the replicative polymerases.
3´-5´ proofreading capacity (Meng et al. 2010; Meng et al. 2009). Indeed, upon DNA damage, ATR signaling-induced degradation of the D subunit leads to an enhanced 3´-5´ exonuclease activity at the expense of the nucleotide extension rate, which is lower compared to the four subunit polymerase. The crystal structures of the human Pol δ subunit 2 in complex with the N-terminal region of the C-subunit and the budding yeast catalytic core are available (Baranovskiy et al. 2008; Swan et al. 2009). The small D subunit remains the only structurally poorly characterized region of the polymerase although pull-down assays indicate that it binds to the catalytic subunit (Li et al. 2006). The conserved subunit 2 in Pol δ acts as a central hub in bringing the catalytic and the regulatory elements of the polymerase together into a complete holoenzyme. Interestingly, all four subunits of Pol δ interact with the PCNA sliding clamp, although each via distinct binding domains. For example, biochemical studies have revealed that the zinc-coordinating C-terminal region of the catalytic subunit is important for PCNA binding (Netz et al. 2012). Pol δ subunits B, C, and D have also been reported to contain different PCNA-interacting motifs (Bruning and Shamoo 2004; Li et al. 2006; Lu et al. 2002). The reason for multiple distinct interactions between Pol δ and PCNA is unclear although it may contribute to the processivity of the polymerase during translocation along the DNA.
Emerging roles of DNA damage repair and response factors in unchallenged and perturbed replication
DNA polymerase δ Mammalian DNA polymerase δ was initially characterised using the in vitro SV40 DNA replication system where, together with Pol α/Primase, it mediates DNA synthesis on both leading and lagging strands at the replication fork (Waga and Stillman 1994). Pol δ was later found to preferentially act on the lagging strand, as shown by mutation rate analysis in yeast strains that carry an exonuclease-deficient Pol δ and a wildtype Pol ε (Nick McElhinny et al. 2008; Pursell et al. 2007). Nonetheless, deletion of the catalytic domain of Pol ε in yeast does not affect cell viability (Kesti et al. 1999), suggesting that Pol δ can be found structurally associated with both replication strands, in the absence of a dedicated leading strand polymerase. Together with the other two replicative DNA polymerases, Pol δ shares the evolutionarily conserved catalytic domain and the accessory subunit B. Pol δ contains two additional, however, non-ubiquitous (Gerik et al. 1998), members: the C and D subunits, evolutionarily unrelated to Pol ε’s subunits 3 and 4 (Liu et al. 2000). While subunit D is not required for mitotic growth in fission yeast or DNA synthesis in vitro (Podust et al. 2002; Reynolds et al. 1998), it contains an important finetuning function that balances the polymerizing activity and the
Protecting replication forks and promoting Replisome stability DNA replication progression is frequently impaired by secondary DNA structures, covalent adducts, and DNA lesions. Several systems ensure correct duplication of genomic DNA in prokaryotic and eukaryotic organisms. In organisms where replication starts from a single origin, restarting mechanisms assist fork progression by exploiting the homologous recombination DNA repair machinery (Errico and Costanzo 2012). In eukaryotes the mechanisms underlying the function of DNA repair and DNA damage response proteins in DNA replication are less clear. Moreover, the links between DNA damage response and repair factors with Replisome components and their contribution to the maintenance of genome stability DNA replication are largely unknown. This is an aspect of DNA replication of higher eukaryotes, which has been difficult to address due to the fact that many DNA repair genes are essential in metazoan cells. Experiments performed in yeast have greatly contributed to understand the role of some of the DNA damage response genes at stalled and collapsed replication forks. Replisome components are maintained stably associated to
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DNA to ensure rapid replication resumption when replication forks stall. To this end, numerous proteins, which are not essential for DNA synthesis, are recruited to the replication fork via interaction with members of the Replisome. Among these proteins, Timeless/Tim1/Tof1, Tipin/Csm3 together with Claspin/Mrc1 have been identified, both in yeast and higher eukaryotes, as members of the replication pausing complex that contributes both to fork stabilization and to checkpoint activation. Tim1, Tipin, and Claspin are considered mediators of the ATR signalling cascade, being important for the efficient phosphorylation of the effector kinase Chk1. This pathway is essential to prevent the loss of Replisome components from stalled forks. The ATR-Chk1 pathway coordinates several mechanisms that contribute to maintain replication fork stability. The ATRChk1 signalling cascade is required to maintain replicative polymerases bound to forks to regulate branch migrating helicases such as Blm and to fine tune homologous recombination (HR), either positively or negatively (Cimprich and Cortez 2008). The absence of a functional checkpoint leads to Replisome dissociation and formation of aberrant DNA structures that are processed by recombination proteins and exonucleases (Cotta-Ramusino et al. 2005; Segurado and Diffley 2008). The Tipin/Tim1 complex also has a direct role in preserving Replisome integrity by preventing excessive unwinding of DNA at stalled replication forks. It is possible that the complex directly promotes the coupling between helicase and polymerases. The interaction of these proteins with several Replisome components, including polymerases and the Cdc45–MCM–GINS (CMG) helicase complex indicates that Tipin, Tim1, and Claspin play a major structural role at stalled forks (Errico and Costanzo 2012). In contrast to yeast cells, the absence of Tipin/Tim1 is lethal in mammalian organisms and leads to the accumulation of chromosomal breakage even in the absence of apparent DNA damage (Chou and Elledge 2006). It is possible that Tipin/Tim1 assists continuous fork restart by promoting Pol α re-priming on the leading strand (Errico et al. 2009). These events might be frequent due to the formation of endogenous DNA lesions even in the absence of exogenous DNA damaging insults (Lindahl and Barnes 2000). In line with the proposed role, recent evidence showed that Tim1 is able to increase the activity of all major DNA polymerases and to decrease the activity of MCM helicase (Cho et al. 2013). Homologous recombination factors and replication fork progression Genetic inactivation of many DNA recombination genes is lethal at very early stages of development in higher eukaryotes. This suggests that DNA recombination genes are probably required to assist DNA replication or to correct problems
encountered by the replication machinery more frequently in higher eukaryotes than in simpler organisms. Although the function of many DNA repair factors following DNA damage is known the biochemical mechanisms underlying their function during unchallenged DNA replication in vertebrate cells are poorly understood. For example, inactivation of important homologous recombination genes such as Mre11, Rad50, or Nbs1 is lethal in mice, indicating that these are required for cell survival in complex organisms (Errico and Costanzo 2012). Moreover, replacement of Mre11 with an allele that does not have nuclease activity causes phenotypes that are indistinguishable from those of mice null for Mre11 (Buis et al. 2008). In contrast, mutations in the nuclease domain of Mre11 in S. cerevisiae have a limited effect and Mre11 null cell are mostly viable (D'Amours and Jackson 2002). Therefore, while yeast mutants in many of the key DNA repair genes are viable, loss of the same proteins in higher eukaryotes results in cell or embryonic lethality. The reasons behind this discrepancy are unclear. It is possible that other pathways in lower eukaryotes compensate for the absence of HR genes in unchallenged conditions. An important role attributed to HR genes in DNA replication is the restart of collapsed replication forks formed when Replisomes encounter an obstacle or a discontinuity in the template leading to the formation of a broken end. The mechanism by which HR promotes the rebuilding of a Replisome has been largely characterized in yeast cells. This pathway is also known as break-induced replication (BIR), and it is defined as the restart of DNA replication from a DSB (Llorente et al. 2008). The mechanisms underlying BIR have been clarified in S. cerevisiae genomes in which an artificial DSB is induced a mitotically arrested cell to initiate long stretches of newly synthesized DNA. This system provides a unique model of fork repair by HR (Lydeard et al. 2010). This system has been useful to establish that most of the HR genes are required for efficient fork restart through BIR. Extensive DNA synthesis associated with BIR requires both the leading and lagging strand polymerases and all the components of the replicative helicase, whereas the replication factors necessary to assemble a pre-replication complex at a replication origin are not required (Lydeard et al. 2010). An important difference with normal DNA replication forks is that the replication apparatus built through HR process is mutagenic. DNA–polymerases α and δ are required for the initial steps of DNA synthesis, whereas DNA–polymerase ε becomes involved only later (Lydeard et al. 2007). Pol 32, the accessory subunit of polymerase δ, is essential for BIR but not to the progression of the normal replication fork. The mutagenic Replisome that is formed during BIR results in large frame shifts, leading to genomic instability. Therefore, although BIR is important to rescue the lethality arising from a DSB formed at collapsed forks, it
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compromises the genetic stability of the rescued cells (Deem et al. 2011; Hicks et al. 2010). Intriguingly, recent investigations in fission yeast show that HR can restart forks arrested by a replication fork barrier independently of a DSB (Lambert et al. 2010; Mizuno et al. 2009). This mechanism is also potentially mutagenic as shown by the observation that recombination-restarted forks have a considerably high propensity to execute a U-turn at small inverted repeats contributing to the generation of gross chromosomal rearrangements (Mizuno et al. 2013). BIR, which requires coordinated repair and replication events might have a central role in vertebrate cells. The presence of repetitive sequences might facilitate this type of repair allowing homologous pairing of the broken arm of the replication fork with DNA segments downstream or upstream of the lesion (Costanzo et al. 2009). This type of repair might be essential for cell survival in the presence of collapsed forks at the expense of genome stability. Although the involvement of HR has been clearly established in genetic systems, the detailed biochemical analysis of the role of HR factors in DNA replication is still less clear. Experiments in the vertebrate Xenopus laevis egg extract cell free system have been helpful to study the biochemistry of DNA damage response and DNA repair factors in eukaryotic DNA replication, overcoming survival issues related to the inactivation of these factors. These cell free systems allow extensive biochemical analysis and can reproduce basic cell cycle events such as chromatin formation, nuclear assembly, and semi-conservative DNA replication. Egg extracts have proven a powerful tool for the in vitro study of both DNA replication and cell cycle progression (Costanzo et al. 2009; Costanzo and Gautier 2003, 2004). Using specific antibodies to deplete specific proteins, it has been shown that Mre11 is required during unchallenged DNA replication to prevent accumulation of DSBs during a single round of DNA replication (Costanzo et al. 2001). More recently, this system has been useful to dissect the role in DNA replication of another DNA repair gene involved in homologous recombination such as Rad51, which is the eukaryotic ortholog of RecA in Escherichia coli, and plays a central role during meiosis as well as in DSB repair. Rad51 is not essential in S. cerevisiae, and yeast cells deficient for Rad51, Rad52, and Rad54 are viable under unchallenged conditions, whereas Rad51 depletion results in cellular lethality in vertebrates (San Filippo et al. 2008). This suggests that Rad51 plays indispensable roles not only in meiotic chromosomal recombination but also in normal cell cycle in higher organisms. Chicken DT40 cells arrest at G2 phase even without exogenous DNA damage upon conditional knockdown of Rad51, which leads to the accumulation of single-stranded DNA lesions activating the G2/M checkpoint (Su et al. 2008). However, it was not clear how
these single-stranded DNA (ssDNA) regions were formed. Using Xenopus egg extracts to examine the role of Rad51 during DNA replication, it was found that Rad51 binds to chromatin during DNA replication and that its binding is partially suppressed by inhibition of replication origin assembly. This indicated that a fraction of Rad51 binding to chromatin takes place after replication forks have been established and that in addition to its well-known role in DSB repair, Rad51 might be required for DNA replication (Hashimoto et al. 2010). To gain insight into the function of Rad51 at replication forks, electron microscopic analyses (EM) of genomic replication intermediates (RIs), recovered after psoralencrosslinking of nuclei replicated extracts, was performed. EM samples showed a high frequency of RIs in Rad51depleted extracts showing at least one ssDNA gap behind the replication fork and the presence of ssDNA regions directly at the fork (Hashimoto et al. 2010). These data showed that Rad51 is directly required at DNA replication forks for uninterrupted and accurate replication of undamaged templates. Rad51 could have a protective role towards nascent DNA chains and the observed extended ssDNA stretches at the fork could result from increased susceptibility to exonucleolytic degradation. This hypothesis was confirmed by evidence that nascent DNA strands are actually degraded by Mre11 nuclease in the absence of Rad51 leading to the formation of ssDNA gaps (Hashimoto et al. 2010). Mre11dependent degradation of nascent DNA at stalled forks probably reflects a physiological role of Mre11 nuclease at forks. One hypothesis is that Mre11-dependent cleavage of the 3´ end of the nascent DNA is required to free the stalled polymerase and promote replication fork restart downstream of the stalling site. This would explain the essential role of Mre11 nuclease observed in mouse cells. It is possible that Rad51 limits the extent of the resection, which progresses to pathological levels in its absence. Regulating Rad51 and Mre11 function How Rad51 and Mre11 are regulated on replication forks is still poorly understood. As Rad51 is mainly in complex with BRCA2, it is likely that this large protein coordinates the different roles of Rad51 in replication and DSB repair (Pellegrini and Venkitaraman 2004). BRCA2 contains a number of repeats that are critical for binding to Rad51 called the BRC repeat. There is also a helical domain, which adopts an alpha helical structure, consisting of a four-helix cluster core (alpha 1, alpha 8, alpha 9, alpha 10) and two successive beta-hairpins. The alpha 9 and alpha 10 helices pack with the BRCA2 OB1 domain, which consists of a five-stranded beta-sheet that closes on itself. An intriguing region is the tower domain, which adopts a secondary structure consisting of a pair of
Chromosoma
long, antiparallel alpha-helices (the stem) that support a three-helix bundle at their end, called the 3HB domain, which is similar to the DNA binding domains of the bacterial site-specific recombinases. The Tower domain has an important role in the tumour suppressor function of BRCA2, and is essential for BRCA2 binding to DNA. However, how the different domains work together is poorly understood (Pellegrini and Venkitaraman 2004; Pellegrini et al. 2002). As recently shown, BRCA2 loads Rad51 onto replication forks to prevent the nuclease activity of Mre11 from degrading stalled replication forks (Schlacher et al. 2011). It is likely that BRCA2-dependent assembly of Rad51 onto stalled replication forks is required to prevent Mre11mediated degradation of nascent DNA. BRCA2 role in DNA replication might be even more important for chromosome integrity than its role in DSB repair. However, there are some unresolved questions arising form these studies. It is unclear what makes forks stall so frequently in the absence of DNA damaging agents. The mechanism by which BRCA2 loads Rad51 is also unclear. BRCA2 might load Rad51 onto regressed arms formed at reversed forks arising in conditions that halt Replisome progression. These reversed forks would have DSB-like ends available for resection and Rad51 binding (Fig. 3a). This process might facilitate DNA damage bypass in the presence of fork stalling lesion (Fig. 3b). Alternatively, BRCA2 might directly promote Rad51 loading onto ssDNA gaps that might form frequently during DNA replication (Jensen et al. 2010). This role for BRCA2 might explain the wider requirement for HR-mediated Fig. 3 a A speculative mechanism for Rad51/BRCA2 function at replication forks. During DNA replication, Rad51 might be loaded in BRCA2 dependent fashion onto ssDNA gaps that accumulate behind replication forks and on regressed arms formed at reversed forks. Rad51 bound to DNA might prevent Mre11 dependent degradation of nascent DNA. b From replication fork collapse to restart. Transition from a chicken foot DNA structure to a restarted replication fork. It remains unknown whether this process requires nucleolytic activity or rather damage bypass. Damaged DNA is indicated by a red triangle
a
processes during unchallenged DNA replication. The clarification of these alternative models awaits further studies. Responding to template breakage during DNA replication Although we are beginning to understand how the Replisomes deal with DNA lesions that halt the progression of replication forks, we have limited knowledge of the molecular events occurring during fork restart, especially in higher eukaryotes. The behavior of Replisome components and DNA repair factors on unrepaired nicks at the passage of the forks has been recently addressed at the biochemical level. In particular, the behavior of the CMG complex subunits Mcm2-7, Cdc45, and GINS was analyzed and it was found that the GINS subunit and Pol ε are specifically lost upon induction of ssDNA lesions in the template (Hashimoto et al. 2011). Intriguingly, it was found that the Mcm2-7 helicase is maintained on DNA and replication forks are then restored in a Rad51 and Mre11-dependent fashion. In this process, the GINS and Pol ε are reloaded onto forks to restart replication. The uncoupling of GINS from the CMG complex was unexpected, considering that Cdc45 and GINS are recruited onto replication forks interdependently during the initiation of DNA replication. The release of GINS at the passage of the fork across a discontinuous template might be due to the structural configuration that the GINS factor adopts within the CMG complex (Fig. 2). Importantly, a consequence of GINS detachment would be the slowing of helicase progression owing to the loss of a
Parental DNA Mcm2-7 7
??
Mre11 Rad 51
Rad 51 BRCA2
Mre11 or other nucleases
b
??
BRCA2
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major activator of the complex. This would also limit the extent of ssDNA accumulation potentially arising from DNA unwinding in the absence of DNA synthesis. The reloading of GINS onto the Mcm2-7–Cdc45 complex during fork restart could then reactivate the stalled helicase. In these studies, it was also found that inhibition of Mre11 activity impairs replication fork restart and Replisome integrity after fork collapse. These findings suggest that Mre11 functions are coordinated with DNA replication factors to ensure efficient DNA replication under stressful conditions. Consistent with this, cells lacking Mre11 nuclease were shown to be sensitive to replication fork-stalling agents, indicating that Mre11 is involved in the repair of these structures (Buis et al. 2008). Therefore, Mre11- and Rad51-dependent fork repair leading to reloading of the GINS onto the Mcm27–Cdc45 complex still engaged with the DNA could be sufficient to restore a functional CMG helicase complex and promote replication fork restart following template breakage in higher eukaryotes.
essential for cell duplication and survival, the study of their integrated function will increasingly rely on biochemical model systems such as the X. laevis egg extract coupled to advanced imaging tools to uncover the unknown links between these processes. These studies, combined with the structural/enzymatic characterization of in vitro reconstituted protein assemblies, will provide a framework to describe the link between DNA replication and repair transactions at a molecular level. These studies will be crucial to understand how DNA structure is maintained and propagated.
Conclusions
Abe T, Sugimura K, Hosono Y, Takami Y, Akita M, Yoshimura A, Tada S, Nakayama T, Murofushi H, Okumura K, Takeda S, Horikoshi M, Seki M, Enomoto T (2011) The histone chaperone facilitates chromatin transcription (FACT) protein maintains normal replication fork rates. J Biol Chem 286:30504–30512 Alcasabas AA, Osborn AJ, Bachant J, Hu F, Werler PJ, Bousset K, Furuya K, Diffley JF, Carr AM, Elledge SJ (2001) Mrc1 transduces signals of DNA replication stress to activate Rad53. Nat Cell Biol 3:958–965 Arias EE, Walter JC (2007) Strength in numbers: preventing rereplication via multiple mechanisms in eukaryotic cells. Genes Dev 21:497–518 Asturias FJ, Cheung IK, Sabouri N, Chilkova O, Wepplo D, Johansson E (2006) Structure of Saccharomyces cerevisiae DNA polymerase epsilon by cryo-electron microscopy. Nat Struct Mol Biol 13:35–43 Augustin MA, Huber R, Kaiser JT (2001) Crystal structure of a DNAdependent RNA polymerase (DNA primase). Nat Struct Biol 8:57–61 Baranovskiy AG, Babayeva ND, Liston VG, Rogozin IB, Koonin EV, Pavlov YI, Vassylyev DG, Tahirov TH (2008) X-ray structure of the complex of regulatory subunits of human DNA polymerase delta. Cell Cycle 7:3026–3036 Barry ER, Lovett JE, Costa A, Lea SM, Bell SD (2009) Intersubunit allosteric communication mediated by a conserved loop in the MCM helicase. Proc Natl Acad Sci U S A 106:1051–1056 Bermudez VP, Farina A, Raghavan V, Tappin I, Hurwitz J (2011) Studies on human DNA polymerase epsilon and GINS complex and their role in DNA replication. J Biol Chem 286:28963–28977 Bochman ML, Schwacha A (2008) The Mcm2-7 complex has in vitro helicase activity. Mol Cell 31:287–293 Bochman ML, Schwacha A (2010) The Saccharomyces cerevisiae Mcm6/2 and Mcm5/3 ATPase active sites contribute to the function of the putative Mcm2-7 'gate'. Nucleic Acids Res 38:6078– 6088 Boos D, Frigola J, Diffley JF (2012) Activation of the replicative DNA helicase:breaking up is hard to do. Curr Opin Cell Biol 24:423–430 Botchan M, Berger J (2010) DNA replication: making two forks from one prereplication complex. Mol Cell 40:860–861 Brewster AS, Wang G, Yu X, Greenleaf WB, Carazo JM, Tjajadia M, Klein MG, Chen XS (2008) Crystal structure of a near-full-length
Recent progress indicates that the DNA recombination/breakage repair machinery and the DNA replication apparatus might be more intimately linked than anticipated. The evolutionary history of various eukaryotic Replisome components is coherent with this emerging theme. In fact, multiple subunits have been identified both within the replicative helicase and polymerases, which are catalytically dead enzymes, derived from nucleases, originally involved for example in the prokaryotic double-strand break repair pathway. Likewise, multiple Replisome components, including Ctf4/And1 and Mrc1 have been in turn described as having an architectural function within the unperturbed, moving Replisome, or playing a key role in modulating fork halting/restart in the context of the Replisome pausing complex. Use of the same players during both elongation and DNA damage response suggests that perturbations to Replisome progression are more frequent than expected. The requirement of homologous recombination factors during DNA replication suggests that DNA templates are highly vulnerable to breakage that might irreversibly halt replication progression. The frequent accumulation of disruptive DNA lesions might explain the hijack of the DSB repair factors by the Replisome machinery. Alternatively, the evolutionary solutions found to replicate and process the DNA might have been subsequently adopted by the DNA damage repair machinery to ensure maintenance of DNA integrity as shown by the conserved domain used by both apparatus. In both cases, the two functions need to run in parallel and chromosome replication cannot be completed in the absence of DNA repair even in the absence of apparent stress. As these functions are
Acknowledgments The authors should like to thank Adelina Davies for the critical reading of the manuscript. This work was funded by Cancer Research UK. V.C. is also supported by the European Research Council (ERC) start-up grant (206281), the Lister Institute of Preventive Medicine and the European Molecular Biology Organization (EMBO) Young Investigator Program (YIP).
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