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TRENDS in Cell Biology Vol.12 No.11 November 2002
509
Checking on the fork:
the DNA-replication stress-response
pathway
Alexander J. Osborn, Stephen J. Elledge and Lee Zou
To ensure the fidelity of DNA replication, cells activate a stress-response
pathway when DNA replication is perturbed. This pathway regulates not only
progress through the cell cycle but also transcription, apoptosis, DNA
repair/recombination and DNA replication itself. Mounting evidence has
suggested that this pathway is important for the maintenance of genomic
integrity. Here, we discuss recent findings about how this pathway is activated
by replication stress and how it regulates the DNA-replication machinery to
alleviate the stress.
Published online: 26 September 2002
Lee Zou
Verna and Marrs McLean
Department of
Biochemistry and
Molecular Biology,
Howard Hughes Medical
Institute, Baylor College
of Medicine, One Baylor
Plaza, TX 77030, USA.
Alexander J. Osborn
Stephen J. Elledge*
Verna and Marrs McLean
Department of
Biochemistry and
Molecular Biology, and
Department of Molecular
and Human Genetics,
Howard Hughes Medical
Institute, Baylor College
of Medicine, One Baylor
Plaza, TX 77030, USA.
*e-mail:
[email protected]
Dividing cells are vulnerable to genotoxic insults and
other stochastic events that impede the proper
replication and segregation of their genomes to daughter
cells. To respond to these potentially life-threatening
insults, cells have evolved a DNA-replication
stress-response pathway, also referred to as the
DNA-replication checkpoint or S-phase checkpoint.
This pathway responds to replicational interference
by slowing down DNA replication to allow the damage
to be repaired before polymerases encounter more
damage. In addition, in response to stress, this
pathway can activate gene expression, activate
specific repair pathways and prevent entry into
mitosis, allowing cells to maintain a high degree of
genomic integrity [1].
The pathways that respond to replication stress
are signal-transduction pathways that are conserved
across evolution [2]. There are two parallel pathways
that respond to different types of stress (Fig. 1).
The first pathway is the ATM (mutated in ataxia
telangiectasia) pathway, which responds to the
presence of double-strand breaks (DSBs) (Fig. 1a).
This pathway acts during all phases of the cell cycle
and can activate many of the downstream components
of the second pathway, the ATR (ATM–Rad3-related)
pathway. The ATR pathway also responds to DSBs, but
more slowly than ATM. In addition, the ATR pathway
can respond to agents that interfere with the function
of replication forks (Fig. 1b), such as hydroxyurea
(HU), ultraviolet (UV) light and DNA-alkylating
agents such as methyl methane sulfonate (MMS).
The ATR pathway is the main focus of this article.
The ATM and ATR proteins are conserved in all
eukaryotes and belong to a protein kinase family
related to phosphoinositide 3-kinases (PIKKs). Among
the members of the PIKK family, the Saccharomyces
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cerevisiae Mec1, Schizosaccharomyces pombe Rad3
and human ATR kinases are essential for the response
to replication stress. Each of these PIKKs stably
associates with a partner that is likely to function as a
regulatory subunit for the kinase (Table 1).
Functionally downstream of these PIKKs and
dependent upon them for activation are members of
the CHK kinase family (Table 1). In response to
replication stress, a group of proteins are important
for mediating the checkpoint signal from the PIKKs to
the CHKs and other substrates (Table 1, Fig. 1). Some
of these proteins might be involved in sensing the
replication stress and regulating the activity of PIKKs,
whereas others might organize a checkpoint-responsive
complex in which the PIKKs can phosphorylate
the CHKs and other substrates. Together, the
PIKKs and CHKs form the core module of the
replication-stress-response pathway, into which signals
from sensory components of the pathway flow and from
which the activation of effector components emanates.
Ultimately, signals that elicit the S-phase checkpoint
arise at replication forks and effect actions at those forks.
However, the best-understood aspects of the S-phase
checkpoint are the intermediary ones that allow the
signals to be translated into effects. Little is known
about which signal is detected at a stalled replication
fork or which targets are ultimately affected at the fork.
What is sensed during S phase?
Although it is still unclear how the S-phase checkpoint
is activated, several types of replication block are
known to elicit checkpoint responses during S phase.
The first type of replication block is imposed by direct
inhibition of DNA synthesis. For example, HU
(a well-studied activator of the S-phase checkpoint)
stalls replication forks by depleting the deoxynucleotide
triphosphate (dNTP) pool. Aphidicolin, by contrast,
activates the checkpoint by inhibiting DNA synthesis
via polymerases. Certain mutations in the replication
machinery itself can also trigger checkpoint responses
during S phase [3,4], suggesting that a signal for
checkpoint activation can be generated when DNA
synthesis is perturbed.
The second type of replication block results from
DNA adducts induced by DNA-modifying agents.
Although these types of DNA damage could elicit the
DNA-damage checkpoint, they clearly impose stress on
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Review
510
(a)
TRENDS in Cell Biology Vol.12 No.11 November 2002
Double-strand breaks
ATM
Mre11 Rad50
Nbs1
?
P
Mdm2
P
P
53BP1
P
P
p53
P
P
SMC1
BRCA1
CHK2
P
P
FANC-D2
BLM
P
H2AX
P
CDC25s
MUS81
p21
Cyclin–CDKs
Apoptosis
(b)
DNA replication Repair/recombination
Replication stress or double-strand breaks
Polα-primase, Polε?,
TopBP1? RPA
P
P
Rad17
ATR ATRIP
Rad1–Rad9–Hus1
P
CLASPIN
P
H2AX
P
BLM
P
P
P
CHK2
p53
BRCA1
P
CHK1
P
CDC25s
P
p21
CDC25s
MUS81
Cyclin–CDKs
Apoptosis
Repair/recombination
CyclinB–Cdc2
DNA replication
Mitosis entry
TRENDS in Cell Biology
Fig. 1. The two parallel damage-response pathways in mammalian cells. The ATM-dependent pathway
is primarily involved in the response to double-strand breaks (a), whereas the ATR-mediated pathway
responses to both replication stress and double-strand breaks (b). ATM and ATR phosphorylate the
CHKs and several other substrates (indicated by encircled P). The phosphorylation of these effectors
collectively downregulates DNA replication, promotes DNA repair and recombination (or apoptosis),
and delays cell-cycle transitions. It must be realized that the ATM and ATR kinases are also likely to
function through novel effectors that have yet to be discovered.
progressing replication forks. In budding yeast, MMS
profoundly reduces the rate of DNA-replication-fork
progression [5]. UV-induced DNA lesions can also slow
down S phase if they are left unrepaired in G1 [6]. In
Xenopus egg extracts, UV-irradiated sperm chromatin
induces phosphorylation of Chk1 [7], a hallmark of
S-phase-checkpoint activation.
Another type of replication block might be
associated with the DNA breaks generated during
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DNA replication. In theory, DSBs could arise if
replication forks pass through nicked DNA or
certain repair or recombination intermediates.
Replication-associated DSBs could also be induced
by agents such as topoisomerase-I poisons [8].
Currently, it is not clear whether different types of
replication block are sensed by the same mechanism.
It is possible that a common DNA structure generated
at stalled replication forks is recognized by the
checkpoint sensors. It is also possible that different
DNA lesions are processed differently by replication
forks and sensed by different groups of sensor.
Furthermore, some types of DNA damage could be
sensed in both replication-dependent and
replication-independent manners, depending on the
cell-cycle status of the cells when they encounter
the damage.
In addition to the response to replication blocks, the
S-phase checkpoint might also have a function during
an unperturbed S phase. Consistent with this, Mec1 and
Rad53 are essential for cellular viability in budding
yeast, and ATR and Chk1 are essential in mammalian
cells. Even in the absence of replication-blocking agents,
the budding yeast Ddc1, Ddc2 and Rpa2 proteins are
phosphorylated during S phase in a Mec1-dependent
manner [3,9,10]. Furthermore, during S phase, the
Sml1 protein is phosphorylated by Dun1 (a kinase
activated by Mec1 and the Rad53 CHK) and degraded
in a Dun1-dependent manner [11]. These findings
suggest that Mec1, Rad53 and Dun1 are at least
partially activated during S phase, perhaps by certain
replication intermediates. The S-phase checkpoint
might generate a signal of ongoing replication and
delay the onset of mitosis. It might also facilitate the
ability of replication forks to go through spontaneous
DNA lesions and various chromosomal structures.
Indeed, certain regions on chromosomes are replicated
slowly in mec1 mutants and are prone to DSBs [12].
Sensors of DNA replication stress
The processes by which replication blocks or ongoing
DNA synthesis are sensed by checkpoint sensors and
by which the checkpoint signaling is initiated are still
largely unknown. However, recent studies have shed
new light on the sensors and the DNA structures that
they might recognize.
ATR–ATRIP complex
One candidate for a sensor of replication stress is
ATR. In humans and Xenopus, ATR is required for the
Chk1 phosphorylation induced by DNA-replication
blocks [13,14]. Human ATR exists in a stable complex
with a protein called ATRIP [15]. The fission yeast
Rad3 and the budding yeast Mec1 also form similar
complexes with the ATRIP-related factors Rad26 and
Ddc2/Lcd2/Pie1, respectively [9,16–18]. In response
to ionizing radiation, Rad26 is phosphorylated by
Rad3 independently of any other known
checkpoint protein, suggesting that the Rad3–Rad26
complex might directly recognize certain types of
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511
Table 1. Proteins involved in the replication stress response in various
organisms
Replication factors and
associated proteins
PIKKs
PIKK-associated factors
RFC-like factors
PCNA-like factors
Mediators
CHK kinases
Vertebrates
Fission yeast
Budding yeast
Polα-primase
a
Polε
RFCs 2–5
RPA
a
TopBP1
ATR
ATM
ATRIP
Rad17
Polα-primase
a
Polε
a
RFCs 2–5
a
RPA
Cut5
Drc1
Rad3
a
Tel1
Rad26
Rad17
Rad9
Rad1
Hus1
Claspin
a
BRCA1, 53BP1
Chk2
Chk1
Rad9
Rad1
Hus1
Mrc1
a
Rhp1/Crb2
Cds1
Chk1
Polα-primase
Polε
RFCs 2–5
RPA
Dpb11
Drc1/Sld2
Mec1
a
Tel1
Ddc2/Pie1/Lcd1
Rad24
Ctf18/Chl12
a
Ddc1
a
Rad17
a
Mec3
Mrc1
a
Rad9
Rad53
a
Chk1
MCMs
Pre-RC
Mcm10
Unwinding
Cdc45–Sld3
RPA
ATR–ATRIP
Priming
Polα-primase and others
a
Indicated proteins or complexes have not been demonstrated to be important for these
organisms in their replication stress response, but are included for the sake of completeness.
DNA damage [16]. Recently, studies using the
chromatin-immunoprecipitation assay and
green-fluorescent-protein-tagged proteins confirmed
that the budding yeast Mec1–Ddc2 complex is indeed
recruited to the DSBs induced by HO endonuclease
cleavage and to telomeric single-stranded DNA
(ssDNA) caused by a mutation in the telomere-binding
protein Cdc13 [19–21]. In human cells, ATR localizes
with ATRIP in nuclear foci after damage, indicating
that the ATR–ATRIP complex might also be recruited
to the sites of DNA damage [15].
How does ATR–ATRIP recognize various types of
DNA damage? Ddc2 is required for Mec1 to localize to
the HO-induced DSBs and the sites of cdc13-induced
DNA damage [21]. Furthermore, Ddc2 has affinity for
ends of double-stranded DNA (dsDNA). Based on these
data, it was proposed that Ddc2 binds to DNA lesions
and recruits Mec1. Nonetheless, the association of
Ddc2 with DNA damage in vivo without Mec1 remains
controversial [20,21]. In contrast to the above
hypothesis, human ATR itself binds to dsDNA [22].
Moreover, ATR has a slightly higher affinity to
UV-damaged DNA [22]. Although these studies provide
attractive models for damage recognition by ATR, they
do not address how its function is coupled to S phase.
Complementary to these studies, two studies using
Xenopus revealed a link between ATR and DNA
replication. First, Xenopus ATR associates with
chromatin in a replication-dependent manner, even
in the absence of a replication-blocking agent [23].
Also, ATR can no longer associate with chromatin
when the initiation of DNA replication is blocked by
the depletion of the replication factor RPA [24].
Furthermore, DNA polymerase α (Polα) is not
required for the chromatin association of ATR.
These data suggest that, at least during the initiation
stage of DNA replication, a partially assembled
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?
Rad17 complex
Rad1–Rad9–Hus1 complex
TRENDS in Cell Biology
Fig. 2. The association of checkpoint complexes with chromatin during
the initiation of DNA replication in Xenopus. During the initiation of
DNA replication, initiation factors such as Mcm10, Cdc45–Sld3 and RPA
bind to the pre-replicative complex (pre-RC) on chromatin and trigger
the unwinding of DNA. The ATR–ATRIP complex is recruited onto
chromatin after DNA unwinding. The Polα–primase complex and
several other replication proteins are also recruited to the unwound
DNA. The Rad1–Rad9–Hus1 complex binds to chromatin after the
Polα–primase complex. The chromatin association of the
Rad1–Rad9–Hus1 complex requires the Rad17 complex, but when the
Rad17 complex is recruited onto chromatin is unclear. It is not known
whether the ATR–ATRIP and the Rad1–Rad9–Hus1 complexes recruited
during this process are actively signaling.
DNA-replication fork that is generated after origin
unwinding and before the loading of Polα can recruit
ATR (Fig. 2). Nonetheless, it is not clear whether the
ATR recruited during the initiation of DNA replication
is actively signaling in the absence of stress.
Furthermore, it remains untested whether the same
replication intermediate is also responsible for
recruiting ATR at stalled replication forks. It is possible
that the initial loading of ATR allows it subsequently
to scan the replicating chromatin for problems, where
it is then activated to initiate the stress response.
RFC- and PCNA-like complexes
Although the ATR–ATRIP complex might associate
directly with certain DNA structures, it cannot
fully activate the replication-stress response
without the replication-factor-C (RFC) and
proliferating-cell-nuclear-antigen (PCNA)-like
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TRENDS in Cell Biology Vol.12 No.11 November 2002
proteins. During DNA replication, RFC recognizes
the primer–template junction and recruits PCNA
onto DNA, where PCNA functions as a ‘sliding clamp’
to tether DNA polymerases. In fission yeast, the
RFC-like factor Rad17 and PCNA-like factors Rad1,
Rad9 and Hus1 are required not only for the S-phase
checkpoint but also for the DNA-damage checkpoint
outside S phase [25]. Furthermore, Rad17 forms an
RFC-like complex with the four small RFC subunits
in yeast and humans [26], whereas Rad1, Rad9 and
Hus1 form a heterotrimeric ring-shaped complex like
PCNA [27,28]. In budding yeast, the PCNA-like
complex (Rad17–Mec3–Ddc1) is recruited to
HO-induced DSBs and the sites of cdc13-induced
DNA damage in a Rad24-dependent manner [19,20].
Similarly, the human PCNA-like complex
(Rad1–Rad9–Hus1) is also recruited onto chromatin
after damage in a Rad17-dependent manner [29].
The recruitment of the PCNA-like complex to sites of
DNA damage is independent of the ATR kinase
complex in yeast and humans, indicating that the
Rad17 complex can respond to damage independently
of ATR–ATRIP [19,20,29]. Although all of these
findings are consistent with a model in which the
Rad17 complex recognizes DNA damage and loads
the PCNA-like complex onto DNA, a direct
biochemical proof of this model is still lacking.
The DNA structure that is recognized by the
Rad17 complex during the stress response remains a
mystery. The Rad17 complex can bind to ssDNA, dsDNA
and DNA with both single- and double-stranded
regions in vitro, like RFC [26]. However, purified
Rad17 complexes are incapable of loading the
PCNA-like complex onto these DNA structures.
Interestingly, it was shown in Xenopus that the
Polα–primase complex (which synthesizes the
RNA–DNA hybrid primer during DNA replication) is
required for the recruitment of Hus1 onto chromatin
during the initiation of DNA replication [24] (Fig. 2).
Whether the PCNA-like complex recruited has a
signaling role in the absence of stress is unknown.
The RNA portion of the primer synthesized by
primase was thought to be the activator of the
checkpoint, partly because actinomycin D, an
inhibitor of primase, blocks the checkpoint response
to aphidicolin [30]. However, this interpretation has
to be re-evaluated because actinomycin D was
recently found to prevent the chromatin binding of
RPA, Polα and possibly other factors [24]. Whether
the RNA primer is directly involved in checkpoint
activation is still unclear. Nonetheless, recombinant
wild-type human primase, but not a primase mutant,
can restore the checkpoint response in primase-depleted
Xenopus extract [30], suggesting that primase is
required for the checkpoint activation. If the RNA
primer was indeed an activator of the checkpoint, the
Rad17 complex might recognize the RNA-primed
DNA template and recruit the PCNA-like complex.
Nonetheless, it is unclear whether the RNA primer is
accessible to the Rad17 complex in vivo, because the
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RNA and DNA syntheses by the Polα–primase
complex are coupled in eukaryotic cells.
The damage-induced recruitment of the
PCNA-like complex onto DNA appears to be a crucial
step for checkpoint signaling. As in fission yeast,
mammalian Rad17 and Hus1 are required for the
phosphorylation of Chk1 by ATR [29,31]. Rad17 itself
is also a substrate of ATR [29,32,33]. Interestingly,
although both ATR and Rad17 associate with
chromatin in undamaged cells, the phosphorylation of
Rad17 by ATR is significantly stimulated by the
increased amounts of PCNA-like complexes recruited
onto chromatin after damage [29]. A two-step model
for S-phase-checkpoint activation has emerged from
these findings. The first step is the independent
localization of ATR–ATRIP and the Rad17 complex to
the sites of DNA damage, and the second is the
Rad17-dependent loading of the PCNA-like complex
onto DNA (Fig. 3). Once loaded onto DNA, the
PCNA-like complex enables ATR to phosphorylate its
chromatin-associated substrates such as Chk1,
Rad17 and Rad9. Thus, the phosphorylation of Chk1,
a key step towards the activation of the S-phase
checkpoint, is controlled by the interaction of two
parallel sensory pathways. The damage-induced
phosphorylation of Rad17 and Rad9 is also important
for the G1 and G2–M checkpoints [32–34]. Whether
and how these phosphorylation events contribute to
the S-phase checkpoint remains to be tested.
Replication fork
In addition to the proteins that function specifically in
checkpoint signaling, several proteins essential for
DNA replication are also implicated in the activation
of the S-phase checkpoint. In budding yeast, besides
the small RFC subunits that complex with Rad24,
DNA polymerase ε and its interacting partners
Dpb11 and Drc1/Sld2 are also required for efficient
checkpoint activation [35–37]. Notably, the checkpoint
functions of these proteins are not entirely separable
from their replication functions, suggesting that they
might contribute to damage detection, at least in part,
by supporting efficient DNA replication [38].
Interestingly, Dpb11 associates with the PCNA-like
protein Ddc1 [38]. TopBP1, the human homologue of
Dpb11, also associates with human Rad9 [39]. Moreover,
deletion of the RFC- or PCNA-like proteins from
dpb11 mutants renders them more sensitive to HU,
suggesting that these proteins might collaborate to
monitor the progression of replication forks [38].
As described above, the Polα–primase complex
and RPA are also implicated in the response to
replication blocks. In budding and fission yeast,
certain mutations in Polα–primase or RPA can alter
these responses [40–42]. In Xenopus, Polα–primase
and RPA are required to recruit ATR and Hus1
onto chromatin during the initiation of DNA
replication [24]. A Xenopus study suggested that the
RNA-polymerase activity of primase, rather than
the DNA-polymerase activity of Polα, is needed for
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UV
Step 1: Localization of the
ATR and Rad17 complexes
to damage
Stalled replication fork
Rad17 complex
ATR–ATRIP
Step 2: Loading of the
Rad1–Rad9–Hus1 complex
Rad1–Rad9–Hus1 complex
Phosphorylation of
ATR substrates
P
P
P
Chk1
TRENDS in Cell Biology
Fig. 3. A two-step model for S-phase checkpoint activation. The first
step of S-phase-checkpoint activation is the independent localization of
the ATR–ATRIP complex and the Rad17 complex to the sites of damage
or stalled replication forks. The second step is the damage-induced and
Rad17-dependent loading of the Rad1–Rad9–Hus1 complex onto
chromatin. The damage-induced loading of Rad1–Rad9–Hus1 complexes
on chromatin enables ATR to phosphorylate Chk1 and other substrates.
the checkpoint response to replication blocks [30].
However, mutational analysis of the fission yeast
Polα demonstrated that DNA synthesis by Polα is
required for the S-phase checkpoint [4]. Thus, how
these proteins contribute to checkpoint signaling at
stalled replication forks remains to be elucidated.
513
suggest a role for Claspin in the S-phase checkpoint of
vertebrate cells. Claspin in both Xenopus and humans
contains many SQ or TQ motifs, which are potential
phosphorylation sites for PIKKs [44]. Based on these
findings, it was proposed that the activation of Chk1
by ATR might be regulated by Claspin in a similar way
to the activation of Rad53 via Rad9 in S. cerevisiae –
Rad9 is phosphorylated by Mec1 in response to
DNA damage and is thought subsequently to serve
as a scaffold protein upon which Rad53 might
autophosphorylate and self-activate [45].
Phosphorylated Rad9 associates directly with the
C-terminal FHA domain of Rad53 [46] and mutant
versions of Rad9 that lack key phosphorylatable SQ
and TQ motifs fail to bind Rad53 or to stimulate
Rad53 activity [47].
Recently, a protein named Mrc1 has been
discovered in both budding and fission yeast [48,49],
and the fission-yeast Mrc1 bears homology to Claspin.
In budding and fission yeast, Mrc1 has been shown to
be important for the activation of Rad53 and Cds1,
respectively, in response to HU. Like Claspin, both
budding-yeast and fission-yeast Mrc1 contain
multiple SQ and TQ motifs, and both proteins are
phosphorylated in response to treatment with HU.
In addition, the fission-yeast Mrc1 has been shown to
interact physically with Cds1. The parallels between
Mrc1 and Claspin suggest that Mrc1 might mediate
the checkpoint response to replication blocks in a
similar manner to Claspin. A major difference between
Mrc1 and Claspin is that Mrc1 controls activation of
the Chk2 homologs Rad53 and Cds1, whereas Claspin
controls Chk1 activation. Thus, both proteins respond
to replication blocks but have different effectors.
Interestingly, S. cerevisiae mrc1 mutants display a
slow, abnormal S phase [48]. One possible explanation
for this phenotype is that the S-phase checkpoint is
required for normal DNA replication [12,50]. Another
is that this protein, like Drc1, Dpb11 and the other
S-phase-checkpoint proteins mentioned above, is a
component of the replication machinery. If PIKK
family members do in fact localize to stalled replication
forks, the association of Mrc1 with the replication
machinery would place it in a unique position to
transduce the signal of stalled replication forks from
the PIKKs to the CHK kinases.
Claspin and Mrc1
The replication fork and the checkpoint sensors that
might associate with stalled forks are required for the
activation of CHKs, but how the CHKs are physically
linked to these proteins is still elusive. The recent
discovery of a group of proteins that includes Xenopus
laevis Claspin, S. cerevisiae Mrc1 and S. pombe Mrc1
might be an important step in the determination of
this mechanism.
Claspin was first identified by Kumagai and
Dunphy as a Chk1-interacting protein in Xenopus [43],
and was shown to be important for the Chk1 activation
induced by synthetic oligonucleotides and for the
checkpoint response to aphidicolin. These results
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Checkpoint function at replication forks
Maintenance of fork integrity
To complete S phase, cells need to maintain the ability
to synthesize DNA when DNA replication is stressed
and, furthermore, to resume DNA synthesis when the
stress is removed or overcome. Desany et al. made
the important discovery that the budding-yeast mec1
and rad53 mutants cannot complete chromosomal
replication after a transient replication block [50].
This finding suggested that the checkpoint plays a
vital role in maintaining stalled replication forks,
restarting stalled forks or both. Recently,
characterization of replication forks in checkpoint
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TRENDS in Cell Biology Vol.12 No.11 November 2002
mutants has uncovered further details of the
checkpoint’s function at forks.
Using a density-transfer approach, Tercero and
Diffley examined the progression of replication forks on
damaged DNA [5]. They found that Mec1 and Rad53
allow cells to replicate slowly, yet processively and to
completion, when treated with MMS. Cells lacking
Mec1 or Rad53 also replicate slowly in MMS, like the
wild-type cells, but cannot complete replication.
Importantly, this study directly demonstrated that
the processivity of replication forks through damaged
DNA is compromised in the absence of the checkpoint.
Complementary to this study, Foiani and colleagues
used two-dimensional gel electrophoresis to show that,
in the presence of HU, cells require Rad53 to undergo
a slow yet processive replication [51]. Furthermore, the
Rad53 pathway maintains the competence of stalled
replication forks to reestablish replication after release
from HU. Cells with a rad53 mutation accumulate
aberrant DNA structures in HU and cannot restart
replication after release from HU. Accumulation of
similar aberrant DNA structures also occurs when
the replisome is destabilized by a mutation in Polα.
These results support the idea that the checkpoint
maintains the integrity of the replisome in HU.
Interestingly, when the replication forks in the
HU-treated rad53 cells are examined by electron
microscopy, there are two classes of abnormal DNA
structures that are not seen in wild-type cells: long
single-stranded stretches of DNA and Holliday
junctions caused by fork reversal (also known as the
‘chickenfoot’ structure) [52]. The accumulation of
single-stranded DNA might result from unstable
replication forks caused by uncoordinated replication
of the leading and lagging strands in HU. In response
to fork stalling, Rad53 might modify the properties of
the replication machinery, allowing it to tolerate a
reduction in the rate of synthesis, maintain its
processivity and prevent its dissociation from DNA.
Restarting stalled forks
The observation of reversed forks in HU-treated rad53
cells is particularly interesting because it implicates
the checkpoint in the process of restarting stalled forks.
Two processes have been proposed to participate in
restarting stalled forks: fork reversal leading to lesion
bypass and recombination-mediated reinitiation of
replication. Each of these processes has been studied
extensively in the bacterium Escherichia coli.
Given a lesion or stress that stalls a replication
fork, rather than inducing a double-strand break, the
Y-branched structure of the fork can undergo a
process known as fork reversal, which leads to the
formation of an X-shaped Holliday junction. During
fork reversal in E. coli, the RecG helicase separates
the nascent and parental strands, and promotes the
annealing of nascent strands with each other [53].
Fork reversal might allow the fork to be stabilized
while the lesion is removed (Fig. 4a). Alternatively,
when the leading-strand DNA polymerase encounters
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a DNA lesion, fork reversal might allow it to continue
synthesis beyond the site of the lesion by switching to
the newly synthesized lagging strand as a template
(Fig. 4b, dashed line). Subsequent action by a second
helicase (the RuvAB complex or possibly RecG) causes
migration of the Holliday junction past the site of the
lesion to re-establish the Y-shaped structure that will
allow the replication fork to continue replication [54]
(Fig. 4b.1). Alternatively, the Holliday junction caused
by fork reversal might be resolved by Holliday-junction
resolvases. In this pathway, RuvC (a Holliday-junction
resolvase that is recruited by RuvAB) nicks
non-complementary strands of the junction (Fig. 4b.2,
black triangles). The DNA ends generated by the
cleavage then lead to resumption of DNA synthesis
through a recombinogenic pathway. These topics are
covered in greater detail elsewhere [55].
Both the non-recombinogenic (helicase-based) and
recombinogenic (resolvase-based) processes of
Holliday-junction resolution are conserved in
eukaryotic cells. The family of RecQ helicases,
including Sgs1 in budding yeast, Rqh1 in fission yeast
and BLM and WRN in humans, might promote the
formation and migration of reversed forks. BLM [56]
and WRN [57] have both demonstrated an in vitro
ability to catalyze branch migration of Holliday-junction
structures. Furthermore, the Mus81–Eme1 complex
(an endonuclease that can cleave the opposite strands
of Holliday-junction structures in vitro [58,59]) has
been found in budding yeast [60], fission yeast [58]
and humans [59]. It has been suggested that the RecQ
and Mus81 pathways act independently at stalled
forks to effect the same end of restarting replication.
The double mutants mus81 sgs1 and mus81 rqh1 are
nonviable, consistent with the hypothesis that these
two pathways each contribute to the resolution of
stalled replication forks [58,61].
Why do HU-treated rad53 cells accumulate
reversed forks? One possibility is that Rad53 is
required for their resolution. Reversed forks might
also occur in wild-type cells but be rapidly resolved
when the checkpoint is active. In support of this idea,
BLM is phosphorylated in response to HU [62].
Furthermore, Mus81–Cds1 and Mus81–Chk2
interactions have been demonstrated [59,63]. In fission
yeast, Mus81 is phosphorylated in a Cds1-dependent
manner in response to HU [63] and, in human cells,
the abundance of Mus81 increases in response to HU
and UV treatments [59]. Another possibility is that
the integrity of stalled forks, which relies on Rad53 in
HU, is required to resolve the reversed forks. A third
possibility is that certain DNA structures that
accumulate in the rad53 cells might promote the
formation of reversed forks. These three explanations
are not mutually exclusive: for example, Rad53 might
act both to prevent the formation of aberrant DNA
structures through its modification of the replisome
and to promote the resolution of such structures,
should they arise, through its modification of Mus81
and perhaps other effectors.
Review
TRENDS in Cell Biology Vol.12 No.11 November 2002
515
(a)
Lesion removal
Fork regression
and reinitiation
Non-recombinogenic
Fork reversal to
‘chicken foot’
(b.1)
or
Lesion is
encountered and
the fork stalls
Continued lagging
strand synthesis
Fork reversal and
template switching
Extension of the
leading strand
Fork regression
and lesion bypass
HJ resolution and double-strand
break end formation
Strand invasion and Reinitiation of
lesion bypass
replication fork
(c)
Strand invasion
Nicked DNA or a stalled fork is
converted to a double-strand break
Recombinogenic
(b.2)
Reinitiation of
replication fork
TRENDS in Cell Biology
Acknowledgements
We thank M. Foiani for
sharing results before
publication. S.J.E. is
supported by a NIH Grant
(GM44664) and is an
Investigator with the
Howard Hughes Medical
Institute and the
Robert E. Welch Professor
of Biochemistry. L.Z. is a
fellow of the Cancer
Research Fund of the
Damon Runyon–Walter
Winchell Foundation.
A.J.O. was supported by
an NIGMS training grant
(#5 T32 GM08307-11).
We apologize to our
colleagues whose work
we could not cover owing
to length constraints.
Fig. 4. Three pathways by which stalled forks might reinitiate replication.
(a) Upon encountering a lesion (shown as a dot), the stalled fork might
undergo fork reversal. This helicase-mediated process might allow the
lesion to become accessible to repair machinery and subsequently to
be removed. (b) In the event of continued lagging-strand synthesis, the
leading strand might be extended by using the nascent lagging strand as
a template. This Holliday junction might undergo fork regression (1), upon
which the replication fork is re-established. Alternatively, the Holliday
junction might be resolved by a resolvase. One possible resolution is
shown in (2). The free double-strand-break (DSB) end subsequently
invades the intact chromosome and acts as a template for
recombination-mediated replication initiation. The successful segregation
of the daughter chromosomes will depend upon the resolution of the
resultant second Holliday junction. (c) Nicked DNA is converted to a DSB
when encountered by the replisome. In addition, the stalled fork might be
processed before fork reversal to yield a DSB end. This DSB end might
invade the intact chromosome to mediate recombination-mediated
reinitiation. Again, a Holliday junction must be resolved.
The function of the stress-response pathway at
stalled forks might vary according to the nature of the
replication stress. The reduction in the rate of DNA
synthesis in response to MMS might be a consequence
of DNA alkylation and its creation of a poor template.
Alternatively, the slowing in replication might be due
to the repair or bypass of DNA lesions. In this case,
the overall slowing observed by Tercero and Diffley [5]
would be a result of short spurts of regularly advancing
replication separated by relatively long periods of
processing stalled intermediates. It is essential to
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