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: [email protected]
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
Chromosoma
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
Non-catalytic
NTD
Pol α
Cdc45
Pol ε
Ctf4/And1
Catalytic
?
?
?
Tipin
Tim1
Pοl δ
Mrc1/
Claspin
Non-catalytic
Non-catalytic
Catalytic
Catalytic
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
Chromosoma
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
Chromosoma
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
Chromosoma
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
Chromosoma
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
Chromosoma
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
Chromosoma
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
Chromosoma
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).
References
Chromosoma
archaeal MCM: functional insights for an AAA + hexameric
helicase. Proc Natl Acad Sci U S A 105:20191–20196
Bruck I, Kaplan DL (2013) Cdc45 Protein-single-stranded DNA interaction is important for stalling the helicase during replication
stress. J Biol Chem 288:7550–7563
Bruning JB, Shamoo Y (2004) Structural and thermodynamic analysis of
human PCNA with peptides derived from DNA polymerase-delta
p66 subunit and flap endonuclease-1. Structure 12:2209–2219
Buis J, Wu Y, Deng Y, Leddon J, Westfield G, Eckersdorff M,
Sekiguchi JM, Chang S, Ferguson DO (2008) Mre11 nuclease
activity has essential roles in DNA repair and genomic stability
distinct from ATM activation. Cell 135:85–96
Chang YP, Wang G, Bermudez V, Hurwitz J, Chen XS (2007) Crystal
structure of the GINS complex and functional insights into its role
in DNA replication. Proc Natl Acad Sci U S A 104:12685–12690
Cho WH, Kang YH, An YY, Tappin I, Hurwitz J, Lee JK (2013)
Human Tim-Tipin complex affects the biochemical properties of
the replicative DNA helicase and DNA polymerases. Proc Natl
Acad Sci U S A 110:2523–2527
Choi JM, Lim HS, Kim JJ, Song OK, Cho Y (2007) Crystal structure
of the human GINS complex. Genes Dev 21:1316–1321
Chong JP, Hayashi MK, Simon MN, Xu RM, Stillman B (2000) A doublehexamer archaeal minichromosome maintenance protein is an ATPdependent DNA helicase. Proc Natl Acad Sci U S A 97:1530–1535
Chou DM, Elledge SJ (2006) Tipin and Timeless form a mutually
protective complex required for genotoxic stress resistance and
checkpoint function. Proc Natl Acad Sci U S A 103:18143–18147
Cimprich KA, Cortez D (2008) ATR: an essential regulator of genome
integrity. Nat Rev Mol Cell Biol 9:616–627
Costa A, Ilves I, Tamberg N, Petojevic T, Nogales E, Botchan MR,
Berger JM (2011) The structural basis for MCM2-7 helicase
activation by GINS and Cdc45. Nat Struct Mol Biol 18:471–477
Costanzo V, Gautier J (2003) Single-strand DNA gaps trigger an ATRand Cdc7-dependent checkpoint. Cell Cycle 2:17
Costanzo V, Gautier J (2004) Xenopus cell-free extracts to study DNA
damage checkpoints. Methods Mol Biol 241:255–267
Costanzo V, Robertson K, Bibikova M, Kim E, Grieco D, Gottesman
M, Carroll D, Gautier J (2001) Mre11 protein complex prevents
double-strand break accumulation during chromosomal DNA
replication. Mol Cell 8:137–147
Costanzo V, Chaudhuri J, Fung JC, Moran JV (2009) Dealing with
dangerous accidents: DNA double-strand breaks take centre stage.
Symposium on Genome Instability and DNA Repair. EMBO Rep
10:837–842
Cotta-Ramusino C, Fachinetti D, Lucca C, Doksani Y, Lopes M, Sogo
J, Foiani M (2005) Exo1 processes stalled replication forks and
counteracts fork reversal in checkpoint-defective cells. Mol Cell
17:153–159
D'Amours D, Jackson SP (2002) The Mre11 complex: at the crossroads
of dna repair and checkpoint signalling. Nat Rev Mol Cell Biol
3:317–327
De Falco M, Ferrari E, De Felice M, Rossi M, Hubscher U, Pisani FM
(2007) The human GINS complex binds to and specifically stimulates human DNA polymerase alpha-primase. EMBO Rep 8:99–103
Deem A, Keszthelyi A, Blackgrove T, Vayl A, Coffey B, Mathur R,
Chabes A, Malkova A (2011) Break-induced replication is highly
inaccurate. PLoS Biology 9:e1000594
Dua R, Levy DL, Campbell JL (1998) Role of the putative zinc finger
domain of Saccharomyces cerevisiae DNA polymerase epsilon in
DNA replication and the S/M checkpoint pathway. J Biol Chem
273:30046–30055
Dua R, Levy DL, Campbell JL (1999) Analysis of the essential
functions of the C-terminal protein/protein interaction domain of
Saccharomyces cerevisiae pol epsilon and its unexpected ability
to support growth in the absence of the DNA polymerase domain.
J Biol Chem 274:22283–22288
Errico A, Costanzo V (2012) Mechanisms of replication fork protection: a safeguard for genome stability. Crit Rev Biochem Mol Biol
47:222–235
Errico A, Costanzo V, Hunt T (2007) Tipin is required for stalled
replication forks to resume DNA replication after removal of
aphidicolin in Xenopus egg extracts. Proc Natl Acad Sci U S A
104:14929–14934
Errico A, Cosentino C, Rivera T, Losada A, Schwob E, Hunt T,
Costanzo V (2009) Tipin/Tim1/And1 protein complex promotes
Pol alpha chromatin binding and sister chromatid cohesion.
EMBO J 28:3681–3692
Feng W, D'Urso G (2001) Schizosaccharomyces pombe cells lacking
the amino-terminal catalytic domains of DNA polymerase epsilon
are viable but require the DNA damage checkpoint control. Mol
Cell Biol 21:4495–4504
Fletcher RJ, Bishop BE, Leon RP, Sclafani RA, Ogata CM, Chen XS
(2003) The structure and function of MCM from archaeal M.
thermoautotrophicum. Nat Struct Biol 10:160–167
Frick DN, Richardson CC (2001) DNA primases. Annu Rev Biochem
70:39–80
Fu YV, Yardimci H, Long DT, Ho TV, Guainazzi A, Bermudez VP,
Hurwitz J, van Oijen A, Scharer OD, Walter JC (2011) Selective
bypass of a lagging strand roadblock by the eukaryotic replicative
DNA helicase. Cell 146:931–941
Gambus A, Jones RC, Sanchez-Diaz A, Kanemaki M, van Deursen F,
Edmondson RD, Labib K (2006) GINS maintains association of
Cdc45 with MCM in replisome progression complexes at eukaryotic DNA replication forks. Nat Cell Biol 8:358–366
Gambus A, van Deursen F, Polychronopoulos D, Foltman M, Jones
RC, Edmondson RD, Calzada A, Labib K (2009) A key role for
Ctf4 in coupling the MCM2-7 helicase to DNA polymerase alpha
within the eukaryotic replisome. EMBO J 28:2992–3004
Gerik KJ, Li X, Pautz A, Burgers PM (1998) Characterization of the
two small subunits of Saccharomyces cerevisiae DNA polymerase delta. J Biol Chem 273:19747–19755
Hashimoto Y, Ray Chaudhuri A, Lopes M, Costanzo V (2010) Rad51
protects nascent DNA from Mre11-dependent degradation and promotes continuous DNA synthesis. Nat Struct Mol Biol 17:1305–1311
Hashimoto Y, Puddu F, Costanzo V (2011) RAD51- and MRE11dependent reassembly of uncoupled CMG helicase complex at
collapsed replication forks. Nat Struct Mol Biol 19:17–24
Hicks WM, Kim M, Haber JE (2010) Increased mutagenesis and
unique mutation signature associated with mitotic gene conversion. Science 329:82–85
Hogg M, Johansson E (2012) DNA Polymerase epsilon. Subcell
Biochem 62:237–257
Ilves I, Petojevic T, Pesavento JJ, Botchan MR (2010) Activation of
the MCM2-7 helicase by association with Cdc45 and GINS proteins. Mol Cell 37:247–258
Ilves I, Tamberg N, Botchan MR (2012) Checkpoint kinase 2 (Chk2)
inhibits the activity of the Cdc45/MCM2-7/GINS (CMG) replicative helicase complex. Proc Natl Acad Sci U S A 109:13163–13170
Im JS, Ki SH, Farina A, Jung DS, Hurwitz J, Lee JK (2009) Assembly
of the Cdc45-Mcm2-7-GINS complex in human cells requires the
Ctf4/And-1, RecQL4, and Mcm10 proteins. Proc Natl Acad Sci U
S A 106:15628–15632
Jensen RB, Carreira A, Kowalczykowski SC (2010) Purified human
BRCA2 stimulates RAD51-mediated recombination. Nature
467:678–683
Johansson E, Macneill SA (2010) The eukaryotic replicative DNA
polymerases take shape. Trends Biochem Sci 35:339–347
Kamada K (2012) The GINS Complex: structure and function. Subcell
Biochem 62:135–156
Kamada K, Kubota Y, Arata T, Shindo Y, Hanaoka F (2007) Structure
of the human GINS complex and its assembly and functional
interface in replication initiation. Nat Struct Mol Biol 14:388–396
Chromosoma
Kamimura Y, Tak YS, Sugino A, Araki H (2001) Sld3, which interacts
with Cdc45 (Sld4), functions for chromosomal DNA replication
in Saccharomyces cerevisiae. EMBO J 20:2097–2107
Kang YH, Galal WC, Farina A, Tappin I, Hurwitz J (2012) Properties
of the human Cdc45/Mcm2-7/GINS helicase complex and its
action with DNA polymerase epsilon in rolling circle DNA synthesis. Proc Natl Acad Sci U S A 109:6042–6047
Kanke M, Kodama Y, Takahashi TS, Nakagawa T, Masukata H (2012)
Mcm10 plays an essential role in origin DNA unwinding after
loading of the CMG components. EMBO J 31:2182–2194
Kelman Z, Lee JK, Hurwitz J (1999) The single minichromosome
maintenance protein of Methanobacterium thermoautotrophicum
DeltaH contains DNA helicase activity. Proc Natl Acad Sci U S A
96:14783–14788
Kesti T, Flick K, Keranen S, Syvaoja JE, Wittenberg C (1999) DNA
polymerase epsilon catalytic domains are dispensable for DNA
replication, DNA repair, and cell viability. Mol Cell 3:679–685
Kilkenny ML, De Piccoli G, Perera RL, Labib K, Pellegrini L (2012) A
conserved motif in the C-terminal tail of DNA polymerase alpha
tethers primase to the eukaryotic replisome. J Biol Chem
287:23740–23747
Klinge S, Nunez-Ramirez R, Llorca O, Pellegrini L (2009) 3D architecture of DNA Pol alpha reveals the functional core of multisubunit replicative polymerases. EMBO J 28:1978–1987
Krastanova I, Sannino V, Amenitsch H, Gileadi O, Pisani FM, Onesti S
(2012) Structural and functional insights into the DNA replication
factor Cdc45 reveal an evolutionary relationship to the DHH
family of phosphoesterases. J Biol Chem 287:4121–4128
Labib K (2010) How do Cdc7 and cyclin-dependent kinases trigger the
initiation of chromosome replication in eukaryotic cells? Genes
Dev 24:1208–1219
Lambert S, Mizuno K, Blaisonneau J, Martineau S, Chanet R, Freon K,
Murray JM, Carr AM, Baldacci G (2010) Homologous recombination restarts blocked replication forks at the expense of genome
rearrangements by template exchange. Mol Cell 39:346–359
Lao-Sirieix SH, Nookala RK, Roversi P, Bell SD, Pellegrini L (2005a)
Structure of the heterodimeric core primase. Nat Struct Mol Biol
12:1137–1144
Lao-Sirieix SH, Pellegrini L, Bell SD (2005b) The promiscuous
primase. Trends Genet: TIG 21:568–572
Lee J, Kumagai A, Dunphy WG (2003) Claspin, a Chk1-regulatory
protein, monitors DNA replication on chromatin independently of
RPA, ATR, and Rad17. Mol Cell 11:329–340
Li H, Xie B, Zhou Y, Rahmeh A, Trusa S, Zhang S, Gao Y, Lee EY, Lee
MY (2006) Functional roles of p12, the fourth subunit of human
DNA polymerase delta. J Biol Chem 281:14748–14755
Li Z, Santangelo TJ, Cubonova L, Reeve JN, Kelman Z (2010)
Affinity purification of an archaeal DNA replication protein network. mBio 1
Li Z, Pan M, Santangelo TJ, Chemnitz W, Yuan W, Edwards JL,
Hurwitz J, Reeve JN, Kelman Z (2011) A novel DNA nuclease
is stimulated by association with the GINS complex. Nucleic
Acids Res 39:6114–6123
Lindahl T, Barnes DE (2000) Repair of endogenous DNA damage.
Cold Spring Harb Symp Quant Biol 65:127–133
Liu L, Mo J, Rodriguez-Belmonte EM, Lee MY (2000) Identification
of a fourth subunit of mammalian DNA polymerase delta. J Biol
Chem 275:18739–18744
Liu S, Bekker-Jensen S, Mailand N, Lukas C, Bartek J, Lukas J (2006)
Claspin operates downstream of TopBP1 to direct ATR signaling
towards Chk1 activation. Mol Cell Biol 26:6056–6064
Llorente B, Smith CE, Symington LS (2008) Break-induced replication: what is it and what is it for? Cell Cycle 7:859–864
Lou H, Komata M, Katou Y, Guan Z, Reis CC, Budd M, Shirahige K,
Campbell JL (2008) Mrc1 and DNA polymerase epsilon function
together in linking DNA replication and the S phase checkpoint.
Mol Cell 32:106–117
Lovett ST, Clark AJ (1984) Genetic analysis of the recJ gene of
Escherichia coli K-12. J Bacteriol 157:190–196
Lu X, Tan CK, Zhou JQ, You M, Carastro LM, Downey KM, So AG
(2002) Direct interaction of proliferating cell nuclear antigen with
the small subunit of DNA polymerase delta. J Biol Chem
277:24340–24345
Lydeard JR, Jain S, Yamaguchi M, Haber JE (2007) Break-induced
replication and telomerase-independent telomere maintenance require Pol32. Nature 448:820–823
Lydeard JR, Lipkin-Moore Z, Sheu YJ, Stillman B, Burgers PM, Haber
JE (2010) Break-induced replication requires all essential DNA
replication factors except those specific for pre-RC assembly.
Genes Dev 24:1133–1144
Lyubimov AY, Strycharska M, Berger JM (2011) The nuts and bolts of
ring-translocase structure and mechanism. Curr Opin Struct Biol
21:240–248
Lyubimov AY, Costa A, Bleichert F, Botchan MR, Berger JM (2012)
ATP-dependent conformational dynamics underlie the functional
asymmetry of the replicative helicase from a minimalist eukaryote. Proc Natl Acad Sci U S A 109:11999–12004
Macneill S (2012) Composition and dynamics of the eukaryotic
replisome: a brief overview. Subcell Biochem 62:1–17
Makarova KS, Koonin EV, Kelman Z (2012) The CMG (CDC45/RecJ,
MCM, GINS) complex is a conserved component of the
DNA replication system in all archaea and eukaryotes. Biol
Direct 7:7
Marinsek N, Barry ER, Makarova KS, Dionne I, Koonin EV, Bell SD
(2006) GINS, a central nexus in the archaeal DNA replication
fork. EMBO Rep 7:539–545
Meng X, Zhou Y, Zhang S, Lee EY, Frick DN, Lee MY (2009) DNA
damage alters DNA polymerase delta to a form that exhibits
increased discrimination against modified template bases and
mismatched primers. Nucleic Acids Res 37:647–657
Meng X, Zhou Y, Lee EY, Lee MY, Frick DN (2010) The p12 subunit
of human polymerase delta modulates the rate and fidelity of
DNA synthesis. Biochemistry 49:3545–3554
Miles J, Formosa T (1992) Evidence that POB1, a Saccharomyces
cerevisiae protein that binds to DNA polymerase alpha, acts in
DNA metabolism in vivo. Mol Cell Biol 12:5724–5735
Mizuno K, Lambert S, Baldacci G, Murray JM, Carr AM (2009)
Nearby inverted repeats fuse to generate acentric and dicentric
palindromic chromosomes by a replication template exchange
mechanism. Genes Dev 23:2876–2886
Mizuno K, Miyabe I, Schalbetter SA, Carr AM, Murray JM (2013)
Recombination-restarted replication makes inverted chromosome
fusions at inverted repeats. Nature 493:246–249
Moyer SE, Lewis PW, Botchan MR (2006) Isolation of the Cdc45/
Mcm2-7/GINS (CMG) complex, a candidate for the eukaryotic
DNA replication fork helicase. Proc Natl Acad Sci U S A
103:10236–10241
Muramatsu S, Hirai K, Tak YS, Kamimura Y, Araki H (2010) CDKdependent complex formation between replication proteins
Dpb11, Sld2, Pol (epsilon}, and GINS in budding yeast. Genes
Dev 24:602–612
Nedelcheva MN, Roguev A, Dolapchiev LB, Shevchenko A, Taskov
HB, Shevchenko A, Stewart AF, Stoynov SS (2005) Uncoupling
of unwinding from DNA synthesis implies regulation of MCM
helicase by Tof1/Mrc1/Csm3 checkpoint complex. J Mol Biol
347:509–521
Netz DJ, Stith CM, Stumpfig M, Kopf G, Vogel D, Genau HM, Stodola
JL, Lill R, Burgers PM, Pierik AJ (2012) Eukaryotic DNA polymerases require an iron-sulfur cluster for the formation of active
complexes. Nat Chem Biol 8:125–132
Chromosoma
Neuwald AF, Aravind L, Spouge JL, Koonin EV (1999) AAA+: A class
of chaperone-like ATPases associated with the assembly, operation,
and disassembly of protein complexes. Genome Res 9:27–43
Nick McElhinny SA, Gordenin DA, Stith CM, Burgers PM, Kunkel
TA (2008) Division of labor at the eukaryotic replication fork.
Mol Cell 30:137–144
Nunez-Ramirez R, Klinge S, Sauguet L, Melero R, Recuero-Checa
MA, Kilkenny M, Perera RL, Garcia-Alvarez B, Hall RJ, Nogales
E, Pellegrini L, Llorca O (2011) Flexible tethering of primase and
DNA Pol alpha in the eukaryotic primosome. Nucleic Acids Res
39:8187–8199
Nuutinen T, Tossavainen H, Fredriksson K, Pirila P, Permi P, Pospiech
H, Syvaoja JE (2008) The solution structure of the amino-terminal
domain of human DNA polymerase epsilon subunit B is homologous to C-domains of AAA+ proteins. Nucleic Acids Res
36:5102–5110
Osborn AJ, Elledge SJ (2003) Mrc1 is a replication fork component
whose phosphorylation in response to DNA replication stress
activates Rad53. Genes Dev 17:1755–1767
Pellegrini L (2012) The Pol alpha-Primase complex. Subcell Biochem
62:157–169
Pellegrini L, Venkitaraman A (2004) Emerging functions of BRCA2 in
DNA recombination. Trends Biochem Sci 29:310–316
Pellegrini L, Yu DS, Lo T, Anand S, Lee M, Blundell TL, Venkitaraman
AR (2002) Insights into DNA recombination from the structure of a
RAD51-BRCA2 complex. Nature 420:287–293
Podust VN, Chang LS, Ott R, Dianov GL, Fanning E (2002)
Reconstitution of human DNA polymerase delta using recombinant
baculoviruses: the p12 subunit potentiates DNA polymerizing activity of the four-subunit enzyme. J Biol Chem 277:3894–3901
Pursell ZF, Isoz I, Lundstrom EB, Johansson E, Kunkel TA (2007)
Yeast DNA polymerase epsilon participates in leading-strand
DNA replication. Science 317:127–130
Remus D, Beuron F, Tolun G, Griffith JD, Morris EP, Diffley JF (2009)
Concerted loading of Mcm2-7 double hexamers around DNA
during DNA replication origin licensing. Cell 139:719–730
Reynolds N, Watt A, Fantes PA, MacNeill SA (1998) Cdm1, the
smallest subunit of DNA polymerase d in the fission yeast
Schizosaccharomyces pombe, is non-essential for growth and
division. Curr Genet 34:250–258
San Filippo J, Sung P, Klein H (2008) Mechanism of eukaryotic
homologous recombination. Annu Rev Biochem 77:229–257
Sanchez-Pulido L, Ponting CP (2011) Cdc45: the missing RecJ
ortholog in eukaryotes? Bioinformatics 27:1885–1888
Schlacher K, Christ N, Siaud N, Egashira A, Wu H, Jasin M (2011)
Double-strand break repair-independent role for BRCA2 in
blocking stalled replication fork degradation by MRE11. Cell
145:529–542
Segurado M, Diffley JF (2008) Separate roles for the DNA damage
checkpoint protein kinases in stabilizing DNA replication forks.
Genes Dev 22:1816–1827
Su X, Bernal JA, Venkitaraman AR (2008) Cell-cycle coordination
between DNA replication and recombination revealed by a vertebrate N-end rule degron-Rad51. Nat Struct Mol Biol 15:1049–1058
Swan MK, Johnson RE, Prakash L, Prakash S, Aggarwal AK (2009)
Structural basis of high-fidelity DNA synthesis by yeast DNA
polymerase delta. Nat Struct Mol Biol 16:979–986
Szyjka SJ, Viggiani CJ, Aparicio OM (2005) Mrc1 is required for
normal progression of replication forks throughout chromatin in
S. cerevisiae. Mol Cell 19:691–697
Tahirov TH, Makarova KS, Rogozin IB, Pavlov YI, Koonin EV (2009)
Evolution of DNA polymerases: an inactivated polymeraseexonuclease module in Pol epsilon and a chimeric origin of
eukaryotic polymerases from two classes of archaeal ancestors.
Biol Direct 4:11
Tanaka H, Katou Y, Yagura M, Saitoh K, Itoh T, Araki H, Bando M,
Shirahige K (2009) Ctf4 coordinates the progression of helicase
and DNA polymerase alpha. Genes Cells: Devoted Mol Cell
Mech 14:807–820
Tercero JA, Labib K, Diffley JF (2000) DNA synthesis at individual
replication forks requires the essential initiation factor Cdc45p.
EMBO J 19:2082–2093
Tourriere H, Versini G, Cordon-Preciado V, Alabert C, Pasero P (2005)
Mrc1 and Tof1 promote replication fork progression and recovery
independently of Rad53. Mol Cell 19:699–706
van Deursen F, Sengupta S, De Piccoli G, Sanchez-Diaz A, Labib K
(2012) Mcm10 associates with the loaded DNA helicase at replication origins and defines a novel step in its activation. EMBO J
31:2195–2206
Vijayraghavan S, Schwacha A (2012) The eukaryotic mcm2-7 replicative helicase. Subcell Biochem 62:113–134
Vos SM, Tretter EM, Schmidt BH, Berger JM (2011) All tangled up:
how cells direct, manage and exploit topoisomerase function. Nat
Rev Mol Cell Biol 12:827–841
Waga S, Stillman B (1994) Anatomy of a DNA replication fork revealed
by reconstitution of SV40 DNA replication in vitro. Nature
369:207–212
Winkler DD, Luger K (2011) The histone chaperone FACT: structural
insights and mechanisms for nucleosome reorganization. J Biol
Chem 286:18369–18374
Yardimci H, Loveland AB, Habuchi S, van Oijen AM, Walter JC
(2010) Uncoupling of sister replisomes during eukaryotic DNA
replication. Mol Cell 40:834–840
Zegerman P, Diffley JF (2007) Phosphorylation of Sld2 and Sld3 by
cyclin-dependent kinases promotes DNA replication in budding
yeast. Nature 445:281–285