Fork Protection Complex Proteins in Various Species.
Replication fork progression is blocked by a variety impediments including DNA damage, aberrant DNA structures, or nucleotide depletion [1-3]. The response to replication fork stalling varies according the type of replication inhibition, the number of stalled forks and the duration of the treatment [3-7]. Stalled replication forks are at increased risk for DNA damage, which can lead to mutation or cell death [7-13]. The cell relies on the Intra-S phase checkpoint and DNA damage response proteins to preserve fork structure to allow recovery and resumption of the cell cycle [5, 10, 14-19]. Thus, the mechanisms that maintain replication fork structure are crucial for genome maintenance, and form a primary barrier to malignant transformation [20, 21].
The drug hydroxyurea (HU) induces a reversible early S-phase arrest by causing deoxynucleotide triphosphate (dNTP) depletion [22-24]. HU is a venerable chemotherapeutic, used for its ability to inhibit cell proliferation, but also because it predisposes proliferating cells to genome instability. The loss of replication fork stability and its associated DNA damage following HU treatment is loosely termed “replication fork collapse”. Changes in dNTP pool levels through other mechanisms (
Wild type cells recover from HU arrest and complete S-phase once drug is removed from the culture medium. Alternatively, some cultures may recover from HU arrest prior to its removal by up-regulating nucleotide synthesis and overcoming HU replication inhibition to slowly complete S-phase [17, 32, 33]. The ability to recover stalled replication out of an HU arrest requires restoration of replication forks, restart of DNA synthesis and completion of S-phase.
Whether a replication fork successfully recovers, or collapses with DNA damage, depends in part on the Intra-S phase checkpoint pathway. Cells lacking the checkpoint suffer fork collapse and death. Notably, cells that do not trigger the Intra-S phase checkpoint continue to synthesize DNA despite the presence of HU. Continued synthesis in the presence of low dNTP pools leads to reduced replication rates and increased single stranded DNA (ssDNA) [33-37]. This is a fragile state of “open” DNA that is prone to double strand breaks (DSBs) [38-40]. Further, altered dNTP levels during DNA replication enhance point mutations, in which the base inserted shifts towards that of the dominant pool or away from the lowest pool [25, 41-44]. This explains why the replication checkpoint is a crucial barrier to genome instability.
Thus, replication fork collapse in checkpoint mutants does not occur immediately after HU treatment, detection of decreased dNTP levels, or failure to mount a checkpoint response. Instead, replication fork collapse across a population of forks, within a culture of cells, is a consequence of continued fork activity. The signs and symptoms of replication fork collapse represent a new execution point, the Replication Fork Collapse Point. This metric describes the time at which the majority of replication forks in a cell population become non-functional. In this review, we describe the causes and symptoms of the Replication Fork Collapse Point, with particular regard to the Intra S-phase checkpoint.
2. Replication Fork Structure is Maintained During Stalling
The replication fork describes a region of denatured DNA where DNA synthesis is actively occurring, resembling a two-tined fork. The replisome encompasses the forked DNA, and the entire complex is large and dynamic, coupling DNA unwinding and polymerization [45-47]. Unwinding is performed by a conserved hexameric helicase (MCM) and its associated proteins Cdc45 and GINS. The processive helicase produces single strand DNA (ssDNA) which becomes transiently coated with replication protein A (RPA), a ssDNA binding protein homologue. ssDNA is the substrate for leading- (polε) and lagging-strand (polδ and polα-primase) polymerases.
These functions must be linked to facilitate DNA synthesis. Coupling generation of ssDNA with its use in replication is particularly important, because ssDNA is vulnerable to forming secondary structures, which leads to DNA damage [40, 48, 49], and recombination [50-52]. Thus, fork proteins limit the amount of DNA unwinding and ssDNA [39, 53]. In normal conditions, synthesis may occur rapidly and the goal of minimizing ssDNA production (<200bp) is easily accomplished . However, if either the leading or lagging strand polymerases become stalled or arrested in a slow zone, the helicase must also be slowed down to prevent it from generating excessive ssDNA and potentially dissociating entirely from the replisome.
Helicase and polymerases are linked by the replication Fork Protection Complex (FPC), which contributes to replication fidelity and later chromosome segregation. Tim1 (
Because of its role maintaining replisome structure, the FPC promotes replication fork efficiency and speed, particularly during fork stalling or pausing. While not essential for DNA replication [58, 64-66], the FPC contributes to processivity [67-70], and has additional roles in response to replication stalling [55, 71, 72], and facilitating sister chromatid cohesion, which is essential for faithful chromosome segregation [73-75].
|TIMELESS (TIM)||Tim1 (
|[56, 57, 63, 65, 72]|
|TIPIN (TIP)||Tipin (
|[11, 56-58, 63, 75, 75-77]|
|[14, 63, 78-82]|
|[59, 61, 73, 83]|
3. Causes of Replication Fork Stalling
DNA replication occurs in a short period during the cell cycle. In yeasts, replication of the ~12 Mb genome occurs within 20 to 30 minutes out of a 2.5 to 3h cell cycle. Human cells require several hours, a fraction of a full cell cycle, to replicate a substantially larger genome. The rate-limiting factor is replication fork velocity at 1–2 kb/min. This is an astonishing rate, considering secondary and tertiary structure of the genome packaged into higher order chromatin domains. The tight links between helicase, polymerase and FPC promote highly processive replication. Importantly, they also contribute to replication fidelity. Disruption of any one component (if not already lethal) leads to significant disruptions in processivity and/or fidelity. This is particularly true when impediments to replication are encountered.
Replication pausing and stalling is caused by both natural barriers and external factors [3, 84, 85]. Some regions of DNA cause replication fork stalling through sequence elements (
A replication termination sequence (RTS1) at the mating locus of fission yeast also promotes unidirectional fork progression by binding the replication termination factor 1 (Rtf1) [88, 90-95]. Unidirectional DNA replication is required to establish an imprint that directs mating type switching. RTS1 replication fork pausing is polar, meaning that forks approaching the barrier from one direction will be affected; forks from the opposite direction continue replication [93, 96].
Similarly, ribosomal DNA (rDNA) arrays are an example of a natural, repetitive element that is at risk for fork pausing. Each of the rDNA repeats contains a polar replication terminator, which ensures that forks proceed unidirectionally through each element [86, 97-100]. This occurs as a response to the binding of a fork arrest protein. For example, in fission yeast the Reb1 protein binds the replication termination element Ter3, which promotes long-range DNA interactions with other chromosomal Ter sequences [101, 102]. Localized to the nucleolus, this may nucleate a zone for replication termination . Based on similarity to prokaryotic replication terminators, Reb1-Ter binding may stop the MCM helicase from creating more ssDNA leading to fork pausing and stalling. Pausing of the fork at this site also depends on FPC proteins Swi1 and Swi3.
Replication termination at rDNA is also seen in budding yeast and mammals. In
The rDNA elements define one type of genomic sequence that causes replication slowing or pausing sites. Other regions of the genome may also cause fragile sites, which are broadly characterized as replication slow zones that are prone to forming DNA breaks [38, 40, 109, 110]. These may be dependent upon the chromatin context, transcriptional activity, or impairment of the fork by external agents, such as HU .
HU inhibits the activity of ribonucleotide reductase, which causes a reduction of dNTP pools . HU is frequently used as a reversible early-S phase block reagent in cultured cells. In this sense, HU response is similar to excess thymidine treatment, which changes dNTP pools and induces an early S-phase arrest in metazoan cells . The size of dNTP pools is intimately linked to cell cycle and checkpoint responses [24, 32, 113-115]. Critically, checkpoint proficiency allows cells to survive HU arrest, hold forks stable, and efficiently restart during release.
4. Intra-S Checkpoint: keeping things connected
The Intra-S phase checkpoint is a kinase cascade that responds to HU treatment. It serves to stabilize replication forks and arrest replication until dNTP pools recover. The checkpoint also prevents DNA damage from forming, particularly DNA double strand breaks, by restricting endonucleases such as Mus81 that can act on stalled fork structures [9, 10]. In addition, the Intra-S checkpoint regulates recombination enzymes (
The remainder of this review will focus on the effects of the checkpoint on the replisome itself. During checkpoint activation, the helicase is restrained and stabilized, to prevent excessive unwinding and allow the fork to restart when HU is removed or bypassed. DNA synthesis is also restrained, preventing mutations that may occur during replication in the presence of altered dNTP pools. Late replication origins are prohibited from firing, conserving these “second-chance” origins for later replication restart. These activities help to stabilize established forks after HU treatment, later allowing them to restart. Alternatively, new forks may be established from the late origins in restart to rescue collapsed forks and complete DNA synthesis.
Wild type cells are actively inhibited from DNA synthesis during HU block [10, 17, 36, 58, 65, 117, 118]. That is, the forks do not cease synthesis because they run out of nucleotides. Rather, the checkpoint ensures that the forks are slowed or stopped before such starvation occurs, saving them from the mutagenic effects of dNTP imbalance [34, 42, 119]. These observations are consistent with depletion, rather than exhaustion of specific dNTP pools , and extremely slow residual synthesis . This fork arrest is accompanied by inhibition of the helicase [15, 53, 54, 65], which reduces ssDNA accumulation and concomitant RPA binding until HU is removed.
The Intra-S phase checkpoint is a key component of the response to HU and actively restrains forks during replication stress. The initial signal to activate the checkpoint is provided by increased ssDNA created as replication forks stall [39, 53, 54]. ssDNA is bound by RPA and recruits the Rad9-Rad1-Hus1 (9-1-1) complex and ATR family kinases to stalled forks [120, 121]. Thus, the symptom of slow or stalled forks (generation of ssDNA) initiates the checkpoint [4, 120-125]. However, the FPC and checkpoint together ensure that the helicase cannot generate too much ssDNA, which provides one defense against replication fork collapse during HU stalling.
Checkpoint activation is also coupled to the FPC proteins, particularly CLASPIN and its yeast equivalent, Mrc1 [118, 126, 127]. In fission yeast, Mrc1 is phosphorylated by the upstream Rad3/ATR kinase to a checkpoint-active form . This activation recruits the downstream Cds1 kinase to the stalled replication fork and is essential to signal amplification and transmission by activated Cds1. This pathway is conserved: in humans and budding yeast, respectively, Chk1/Rad53 is recruited to stalled forks by CLASPIN/Mrc1 and ATR/Mec1 kinase [6, 14, 16, 124, 129-131].
This S phase checkpoint has a parallel structure to the DNA double strand break (DSB) response: Mrc1 is a replication-specific version of the
5. The Rules of Replisome Restraint and Restart,1: Fork Movement
Considering the phenotype of checkpoint mutants, we infer that an active mechanism restrains the helicase during HU treatment. Genome-wide studies in budding yeast show accumulation of single stranded DNA occurs in checkpoint mutants, adjacent to replication forks upon treatment with HU [39, 58, 133]. Similarly, in fission yeast checkpoint mutants, large masses of RPA can be visualized in whole cells treated with HU, which depend upon the MCM helicase .
A simple interpretation is that the helicase becomes uncoupled from the stalling polymerase and unwinds DNA ahead of it. This excessive unwinding generates ssDNA that is prone to breakage, which generates a characteristic DSB marker, phosphorylated histone H2A(X) [15, 39]. In many cases, the RPA signal is associated with markers of DNA synthesis, such as incorporation of the nucleotide analogue BrdU , or proximity to replication fork proteins . Importantly, this uncoupling and unwinding occurs at the same time as DNA synthesis during both HU block and release. This suggests a more subtle effect in which leading and lagging strand synthesis is uncoupled, which leads to simultaneous accumulation of ssDNA and markers of synthesis, either because they are in the same region or because the ssDNA is a functional template.
6. Rules of Restraint and Restart, 2: Synthesis
The second key to restraint and successful restart is modulating the DNA polymerases. Wild type cells incorporate minimal amounts of nucleoside analogue in the presence of HU. Forks slow but remain stable [7, 17, 34, 54]. The rate of nucleotide analogue incorporation decreases, and DNA content does not increase significantly [36, 117, 134, 135]. In the yeast system, studies suggest that early replication forks extend about 5kb from the origin in the presence of HU before stopping [134, 136]. Decreased dNTP pools slow replication elongation during HU arrest. However, ectopic expansion of dNTPs by expressing ribonucleotide reductase from a plasmid can increase fork velocity even in HU . Upon release from HU, replication rapidly restarts, whether from new origins or reactivation of existing forks, which results in rapid completion of DNA synthesis before cell division.
Budding yeast dNTP metabolism is quite robust and resistant to challenge, sensitive only to high levels of HU or significant NTP imbalance. In contrast, fission yeast  and metazoan cells are sensitive to low levels of HU, or modest dNTP imbalance, both which are sufficient to provoke replication arrest . In all systems, there is an intimate connection to the Intra-S phase checkpoint.
Surprisingly, checkpoint mutants do not block DNA synthesis in HU, indicating that they are not actually starved for nucleotides, but rather lacking the ability to monitor pool levels . Fission yeast
The difference between the two situations is that much of this synthesis occurs
Polymerase ε is coupled to the helicase by Mrc1 and the FPC proteins Tof1 (Swi1) and Csm1 (Swi3) [63, 65, 132, 138]. This is thought to stabilize leading strand components at stalled forks in HU. Asynchronously growing
Mrc1 brings Cds1 and Rad3 together to phosphorylate Cds1 on threonine 11 [128, 142]. Subsequently, Cds1 activation is amplified by dimerization and autophosphorylation, setting in motion the full Intra-S phase checkpoint . HU treatment induces little Cds1-T11 phosphorylation in
7. Rules of Restraint and Restart, 3: the late origins
An additional function of the Intra-S phase checkpoint is to restrain late origins from firing. Upon release from HU, these origins become competent for replication, and establish “rescue forks” that ensure completion of DNA replication [33, 36, 143-145]. Could these origins explain the post-release DNA synthesis observed in the checkpoint mutants? While late origin firing must contribute to some of the synthesis after release, we suggest that much of the post-release DNA synthesis does not occur from late origin firing, for the following reasons.
First, origin firing is de-regulated in HU blocked checkpoint mutants, which suggests that many late origins have already fired at the time of release, and are not available for this further synthesis. Recent work on dNTP pools in budding yeast suggests that >200 additional origins are fired in a
Second, it is likely that late origins that fire in checkpoint mutants after HU release are incapable of synthesizing more than a short tract length, due to lack of nucleotides . More analogue is incorporated in
Together, these observations from multiple systems suggest that wild type cells survive HU block and release through coordination of several mechanisms: control of late origin firing, maintenance of existing replication forks, and later restart of the stabilized forks. Wild type cells do not encounter the Replication Fork Collapse Point because forks are maintained, replication is successfully restarted, and DNA synthesis completed.
8. Converting Stalled Forks to Restart
After HU is removed from culture medium, stabilized replication forks are returned to competence for DNA synthesis. In theory, immediate restart from a stabilized fork may be possible if all components are in place, having been protected from disassembly during HU arrest. In many cases this is likely to involve recombination pathways and the Rad51 recombinase. Rad51 binds to replisome components in HU, and around damaged replication forks [7, 15, 151]. Rad51 binds to ssDNA and promotes homologous recombination by allowing broken DNA to invade a homologous region for repair [52, 152, 153]. Checkpoint mutants have additional ssDNA, and experience “branch migration” of the fork structure [7, 52, 94, 154]. The resulting “chicken foot” structure is at risk for becoming a break or collapsed fork. Alternatively, the cruciform structure can be resolved by exonuclease Exo1, but leads to a partially replicated structure that cannot be replicated without
The amount of time in HU until release has different effects in yeast and metazoan cultures. Both budding and fission yeast begin to arrest in HU within the first hour of HU exposure (e.g. [53, 134, 144, 159]). After a few hours at normal growth temperature, adaptation occurs, probably through changes in ribonucleotide reductase activity. In budding yeast, long-term HU exposure causes normal replication profiles to proceed at a glacial pace . Similarly, human cells show increased sensitivity to HU over time, where fewer forks are observed with extended HU dose . Peterman
9. The Collapse Point: A Metric for Fork Stability
The concept of replication fork “collapse” encompasses the observations that DNA damage and broken forks lead to loss of replication. ssDNA accumulates at susceptible forks and is a marker of increased risk of collapse [39, 54, 133, 137]. The DNA damage created at a stalled fork at or before collapse may not simply be DSBs. In fact, single strand breaks may form an important part in the damage process, converted to DSBs either during fork regression or in a second S-phase [163-165].
We propose the Collapse Point as the time when the balance between replication fork processivity and instability tips toward disaster. The time when the majority of forks in a cell have irreversibly, irrecoverably failed and replication will not be completed. Ongoing synthesis in checkpoint mutants during HU treatment sets the forks on a course to destruction, but actual collapse does not occur until the attempt to recover. We suggest some replication forks retain activity and undergo a shortened replication restart after HU release. This is consistent with data in fission and budding yeasts that fork components are retained and move during HU arrest in checkpoint deficient cells [53, 133].
The Replication Fork Collapse Point has no meaning for an individual fork; instead, it is the emergent property of the sum behavior of forks in a cell. The Collapse Point will generally be
While forks in
These results point to the
In turn, these studies prompt further questions. Do dNTP pools recover after release in
Monitoring replication competency, accumulation of ssDNA and DNA damage signals around replication forks permits modeling to determine how replication forks respond to HU arrest and recovery. This, in turn, indicates what role checkpoint proteins Cds1 and Mrc1 play in fork stability and effective restart. The Replication Fork Collapse Point incorporates the signs and symptoms of fork collapse and attempts to put a time to when the majority of replication forks undergo collapse. This is likely different for different genetic backgrounds missing key components of checkpoint signal, fork stabilization and replication restart. Future work will dissect replication fork proteins in HU and release, and take the genome-wide data from microarray and sequencing, moving into monitoring patterns at individual replication forks. Since replication stability and fidelity is a key barrier to malignancy, defining when and how replication forks collapse in the absence of checkpoint will allow insights into the development and prevention of cancer.
We thank Forsburg and Aparicio lab members (USC) for helpful discussions, and Marc Green and Ruben Petreaca for manuscript comments. This work funded by a National Institutes of Health grant R01 GM059321 to SLF.
Torres-Rosell J. De Piccoli G. Cordon-Preciado V. Farmer S. Jarmuz A. Machin F. et al. 2007 Anaphase onset before complete DNA replication with intact checkpoint responses. 315 5817 1411 1415
Rothstein R. Michel B. Gangloff S. 2000 Replication fork pausing and recombination or "gimme a break" 14 1 1 10
Labib K. Hodgson B. 2007 Replication fork barriers: pausing for a break or stalling for time? 8 4 346 353
Lucca C. Vanoli F. Cotta-Ramusino C. Pellicioli A. Liberi G. Haber J. et al. 2004 Checkpoint-mediated control of replisome-fork association and signalling in response to replication pausing. 23 6 1206 1213
Meister P. Taddei A. Vernis L. Poidevin M. Gasser S. M. Baldacci G. 2005 Temporal separation of replication and recombination requires the intra-S checkpoint. 168 4 537 544
Tourriere H. Pasero P. 2007 Maintenance of fork integrity at damaged DNA and natural pause sites. 6 7 900 913
Petermann E. Orta M. L. Issaeva N. Schultz N. Helleday T. 2010 Hydroxyurea-stalled replication forks become progressively inactivated and require two different RAD51-mediated pathways for restart and repair. 37 4 492 502
Mao N. Kojic M. Holloman W. K. 2009 Role of Blm and collaborating factors in recombination and survival following replication stress in Ustilago maydis. 8 6 752 9
Froget B. Blaisonneau J. Lambert S. Baldacci G. 2008 Cleavage of stalled forks by fission yeast Mus81/Eme1 in absence of DNA replication checkpoint. 19 2 445 456
Kai M. Boddy M. N. Russell P. Wang T. S. 2005 Replication checkpoint kinase Cds1 regulates Mus81 to preserve genome integrity during replication stress. 19 8 919 932
Noguchi E. Noguchi C. Du L. L. Russell P. 2003 Swi1 prevents replication fork collapse and controls checkpoint kinase Cds1. 23 21 7861 7874
Bryant H. E. Petermann E. Schultz N. Jemth A. S. Loseva O. Issaeva N. 2009 PARP is activated at stalled forks to mediate Mre11-dependent replication restart and recombination. 28 17 2601 2615
Bernstein K. A. Shor E. Sunjevaric I. Fumasoni M. Burgess R. C. Foiani M. et al. 2009 Sgs1 function in the repair of DNA replication intermediates is separable from its role in homologous recombinational repair. 28 7 915 925
Alcasabas A. A. Osborn A. J. Bachant J. Hu F. Werler P. J. Bousset K. et al. 2001 Mrc1 transduces signals of DNA replication stress to activate Rad53. 3 11 958 965
Bailis J. M. Luche D. D. Hunter T. Forsburg S. L. 2008 Minichromosome maintenance proteins interact with checkpoint and recombination proteins to promote s-phase genome stability. 28 5 1724 1738
Branzei D. Foiani M. 2007 Interplay of replication checkpoints and repair proteins at stalled replication forks. 6 7 994 1003
Lopes M. Cotta-Ramusino C. Pellicioli A. Liberi G. Plevani P. Muzi-Falconi M. et al. 2001 The DNA replication checkpoint response stabilizes stalled replication forks. 412 6846 557 561
Marchetti M. A. Kumar S. Hartsuiker E. Maftahi M. Carr A. M. Freyer G. A. 2002 A single unbranched S-phase DNA damage and replication fork blockage checkpoint pathway. 99 11 7472 7477
Tsang E. Carr A. M. 2008 Replication fork arrest, recombination and the maintenance of ribosomal DNA stability. 7 10 1613 1623
Bartkova J. Horejsi Z. Koed K. Kramer A. Tort F. Zieger K. 2005 DNA damage response as a candidate anti-cancer barrier in early human tumorigenesis. 434 7035 864 870
Bartkova J. Rezaei N. Liontos M. Karakaidos P. Kletsas D. Issaeva N. et al. 2006 Oncogene-induced senescence is part of the tumorigenesis barrier imposed by DNA damage checkpoints. 444 7119 633 637
Koc A. Wheeler L. J. Mathews C. K. Merrill G. F. 2004 Hydroxyurea arrests DNA replication by a mechanism that preserves basal dNTP pools. 279 1 223 30
Matsumoto M. Rey D. A. Cory J. G. 1990 Effects of cytosine arabinoside and hydroxyurea on the synthesis of deoxyribonucleotides and DNA replication in L1210 cells. 30 47
Bianchi V. Pontis E. Reichard P. 1986 Changes of deoxyribonucleoside triphosphate pools induced by hydroxyurea and their relation to DNA synthesis. 261 34 16037 16042
Goodman M. F. Hopkins R. L. Lasken R. Mhaskar D. N. 1985 The biochemical basis of 5-bromouracil- and 2-aminopurine-induced mutagenesis. 31 409 423
Hakansson P. Dahl L. Chilkova O. Domkin V. Thelander L. 2006 Thelander L. The Schizosaccharomyces pombe replication inhibitor Spd1 regulates ribonucleotide reductase activity and dNTPs by binding to the large Cdc22 subunit. 281 3 1778 1783
Kunz BA Kang X. L. Kohalmi L. 1991 The yeast rad18 mutator specifically increases G.C----T.A transversions without reducing correction of G-A or C-T mismatches to G.C pairs. 11 1 218 225
Kohalmi S. E. Haynes R. H. Kunz B. A. 1988 Instability of a yeast centromere plasmid under conditions of thymine nucleotide stress. 207 1 13 16
Potter C. G. 1971 Induction of polyploidy by concentrated thymidine. 68 2 442 448
Meuth M. 1981 Role of deoxynucleoside triphosphate pools in the cytotoxic and mutagenic effects of DNA alkylating agents. 7 1 89 102
Meuth M. 1983 Deoxycytidine kinase-deficient mutants of Chinese hamster ovary cells are hypersensitive to DNA alkylating agents. 110 2 383 391
Mulder K. W. Winkler G. S. Timmers H. T. 2005 DNA damage and replication stress induced transcription of RNR genes is dependent on the Ccr4-Not complex. 33 19 6384 6392
Alvino G. M. Collingwood D. Murphy J. M. Delrow J. Brewer B. J. Raghuraman M. K. 2007 Replication in hydroxyurea: it’s a matter of time. 27 18 6396 6406
Poli J. Tsaponina O. Crabbe L. Keszthelyi A. Pantesco V. Chabes A. 2012 dNTP pools determine fork progression and origin usage under replication stress. 31 4 883 894
Bolderson E. Scorah J. Helleday T. Smythe C. Meuth M. 2004 ATM is required for the cellular response to thymidine induced replication fork stress. 13 23 2937 2945
Feng W. Collingwood D. Boeck M. E. Fox L. A. Alvino G. M. Fangman W. L. et al. 2006 Genomic mapping of single-stranded DNA in hydroxyurea-challenged yeasts identifies origins of replication. 8 2 148 55
Vassin V. M. Anantha R. W. Sokolova E. Kanner S. Borowiec J. A. 2009 Human RPA phosphorylation by ATR stimulates DNA synthesis and prevents ssDNA accumulation during DNA-replication stress. Pt 22 4070 4080
Letessier A. Millot G. A. Koundrioukoff S. Lachages A. M. Vogt N. Hansen R. S. et al. 2011 Cell-type-specific replication initiation programs set fragility of the FRA3B fragile site. 470 7332 120 123
Feng W. Di Rienzi S. C. Raghuraman M. K. Brewer B. J. 2011 Replication stress-induced chromosome breakage is correlated with replication fork progression and is preceded by single-stranded DNA formation. 1 5 327 35
Durkin S. G. Glover T. W. 2007 Chromosome fragile sites. 41 169
Chabes A. Georgieva B. Domkin V. Zhao X. Rothstein R. Thelander L. 2003 Survival of DNA damage in yeast directly depends on increased dNTP levels allowed by relaxed feedback inhibition of ribonucleotide reductase. 112 3 391 401
Davidson M. B. Katou Y. Keszthelyi A. Sing T. L. Xia T. Ou J. et al. 2012 Endogenous DNA replication stress results in expansion of dNTP pools and a mutator phenotype. 31 4 895 907
Fasullo M. Tsaponina O. Sun M. Chabes A. 2010 Elevated dNTP levels suppress hyper-recombination in Saccharomyces cerevisiae S-phase checkpoint mutants. 38 4 1195 1203
Kumar D. Abdulovic A. L. Viberg J. Nilsson A. K. Kunkel T. A. Chabes A. 2011 Mechanisms of mutagenesis in vivo due to imbalanced dNTP pools. 39 4 1360 1371
Muzi-Falconi M. Giannattasio M. Foiani M. Plevani P. 2003 The DNA polymerase alpha-primase complex: multiple functions and interactions. 3 21
Langston L. D. Indiani C. O’Donnell M. 2009 Whither the replisome: emerging perspectives on the dynamic nature of the DNA replication machinery. 8 17 2686 2691
Hubscher U. 2009 DNA replication fork proteins. 521 19
Lopez-Contreras A. J. Fernandez-Capetillo O. 2010 The ATR barrier to replication-born DNA damage. 9 12 1249 1255
Glover T. W. Arlt M. F. Casper A. M. Durkin S. G. 2005 Mechanisms of common fragile site instability. 2 R197 R205
Wang X. Haber J. E. 2004 Role of Saccharomyces single-stranded DNA-binding protein RPA in the strand invasion step of double-strand break repair. E21
Alabert C. Bianco J. N. Pasero P. 2009 Differential regulation of homologous recombination at DNA breaks and replication forks by the Mrc1 branch of the S-phase checkpoint. 28 8 1131 1141
Sugiyama T. Kantake N. 2009 Dynamic regulatory interactions of rad51, rad52, and replication protein-a in recombination intermediates. 390 1 45 55
Sabatinos S. A. Green M. D. Forsburg S. L. 2012 Continued DNA synthesis in replication checkpoint mutants leads to fork collapse.
Sogo J. M. Lopes M. Foiani M. 2002 Fork reversal and ssDNA accumulation at stalled replication forks owing to checkpoint defects. 297 5581 599 602
Mc Farlane R. J. Mian S. Dalgaard J. Z. 2010 The many facets of the Tim-Tipin protein families’ roles in chromosome biology. 9 4 700 705
Gotter A. L. Suppa C. Emanuel BS 2007 Mammalian TIMELESS and Tipin are evolutionarily conserved replication fork-associated factors. 366 1 36 52
Noguchi E. Noguchi C. Mc Donald W. H. Yates J. R. 3rd Russell P. 2004 Swi1 and Swi3 are components of a replication fork protection complex in fission yeast. 24 19 8342 8355
Katou Y. Kanoh Y. Bando M. Noguchi H. Tanaka H. Ashikari T. et al. 2003 S-phase checkpoint proteins Tof1 and Mrc1 form a stable replication-pausing complex. 424 6952 1078 1083
Williams D. R. McIntosh J. R. 2002 mcl1+, the Schizosaccharomyces pombe homologue of CTF4, is important for chromosome replication, cohesion, and segregation. 1 5 758 773
Miles J. Formosa T. 1992 Evidence that POB1, a Saccharomyces cerevisiae protein that binds to DNA polymerase alpha, acts in DNA metabolism in vivo. 12 12 5724 6735
Gambus A. van Deursen F. Polychronopoulos D. Foltman M. Jones R. C. Edmondson R. D. 2009 A key role for Ctf4 in coupling the MCM2-7 helicase to DNA polymerase alpha within the eukaryotic replisome. 28 19 2992 3004
Im J. S. Ki S. H. Farina A. DS Jung Hurwitz. J. Lee J. K. 2009 Assembly of the Cdc45-Mcm2-7-GINS complex in human cells requires the Ctf4/And-1, RecQL4, and Mcm10 proteins. 106 37 15628 15632
Bando M. Katou Y. Komata M. Tanaka H. Itoh T. Sutani T. et al. 2009 Csm3, Tof1, and Mrc1 form a heterotrimeric mediator complex that associates with DNA replication forks. 284 49 34355 34365
Gambus A. Jones R. C. Sanchez-Diaz A. Kanemaki M. van Deursen F. Edmondson R. D. 2006 GINS maintains association of Cdc45 with MCM in replisome progression complexes at eukaryotic DNA replication forks. 8 4 358 366
Nedelcheva M. N. Roguev A. Dolapchiev L. B. Shevchenko A. Taskov H. B. Shevchenko A. et al. 2005 Uncoupling of unwinding from DNA synthesis implies regulation of MCM helicase by Tof1/Mrc1/Csm3 checkpoint complex. 347 3 509 21
Calzada A Hodgson B Kanemaki M Bueno A Labib K 2005 Molecular anatomy and regulation of a stable replisome at a paused eukaryotic DNA replication fork. 16 19 1905 1919
Hamdan S. M. Johnson D. E. Tanner N. A. Lee J. B. Qimron U. Tabor S. et al. 2007 Dynamic DNA helicase-DNA polymerase interactions assure processive replication fork movement. 27 4 539 549
Kim S. Dallmann H. G. Mc Henry C. S. Marians K. J. 1996 Coupling of a replicative polymerase and helicase: a tau-DnaB interaction mediates rapid replication fork movement. 84 4 643 650
Stano N. M. Jeong Y. J. Donmez I. Tummalapalli P. Levin M. K. Patel S. S. 2005 DNA synthesis provides the driving force to accelerate DNA unwinding by a helicase. 435 7040 370 373
Tougu K. Marians K. J. 1996 The interaction between helicase and primase sets the replication fork clock. 271 35 21398 21405
Unsal-Kacmaz K. Chastain P. D. Qu P. P. Minoo P. Cordeiro-Stone M. Sancar A. et al. 2007 The human Tim/Tipin complex coordinates an Intra-S checkpoint response to UV that slows replication fork displacement. 27 8 3131 3142
Yoshizawa-Sugata N. Masai H. 2007 Human Tim/Timeless-interacting protein, Tipin, is required for efficient progression of S phase and DNA replication checkpoint. 282 4 2729 2740
Errico A. Cosentino C. Rivera T. Losada A. Schwob E. Hunt T. et al. 2009 Tipin/Tim1/And1 protein complex promotes Pol alpha chromatin binding and sister chromatid cohesion. 28 23 3681 3692
Tanaka H. Kubota Y. Tsujimura T. Kumano M. Masai H. Takisawa H. 2009 Replisome progression complex links DNA replication to sister chromatid cohesion in Xenopus egg extracts. 14 8 949 963
Leman A. R. Noguchi C. Lee C. Y. Noguchi E. 2010 Human Timeless and Tipin stabilize replication forks and facilitate sister-chromatid cohesion. Pt 5 660 670
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. 104 38 14929 34
Tanaka T. Yokoyama M. Matsumoto S. Fukatsu R. You Z. Masai H. 2010 Fission yeast Swi1-Swi3 complex facilitates DNA binding of Mrc1. 285 51 39609 22
Kumagai A. Dunphy W. G. 2000 Claspin, a novel protein required for the activation of Chk1 during a DNA replication checkpoint response in Xenopus egg extracts. 6 4 839 849
Kumagai A. Kim S. M. Dunphy W. G. 2004 Claspin and the activated form of ATR-ATRIP collaborate in the activation of Chk1. 279 48 49599 45608
Lee J. Gold D. A. Shevchenko A. Shevchenko A. Dunphy W. G. 2005 Roles of replication fork-interacting and Chk1-activating domains from Claspin in a DNA replication checkpoint response. 16 11 5269 5282
Lou H. Komata M. Katou Y. Guan Z. Reis C. C. Budd M. et al. 2008 Mrc1 and DNA polymerase epsilon function together in linking DNA replication and the S phase checkpoint. 32 1 106 117
Osborn A. J. Elledge S. J. 2003 Mrc1 is a replication fork component whose phosphorylation in response to DNA replication stress activates Rad53. 17 14 1755 1767
Williams D. R. McIntosh J. R. 2005 Mcl1p is a polymerase alpha replication accessory factor important for S-phase DNA damage survival. 4 1 166 177
Arlt M. F. Mulle J. G. Schaibley V. M. Ragland R. L. Durkin S. G. Warren S. T. 2009 Replication stress induces genome-wide copy number changes in human cells that resemble polymorphic and pathogenic variants. 84 3 339 350
Mirkin E. V. Mirkin S. M. 2007 Replication fork stalling at natural impediments. 71 1 13 35
Krings G. Bastia D. 2004 swi1- and swi3-dependent and independent replication fork arrest at the ribosomal DNA of Schizosaccharomyces pombe. 101 39 14085 90
Noguchi C. Noguchi E. 2007 Sap1 promotes the association of the replication fork protection complex with chromatin and is involved in the replication checkpoint in Schizosaccharomyces pombe. 175 2 553 566
Dalgaard J. Z. Klar A. J. 2000 swi1 and swi3 perform imprinting, pausing, and termination of DNA replication in S. pombe. 102 6 745 751
Razidlo D. F. Lahue R. S. 2008 Mrc1, Tof1 and Csm3 inhibit CAG.CTG repeat instability by at least two mechanisms. 7 4 633 640
Ahn J. S. Osman F. Whitby M. C. 2005 Replication fork blockage by RTS1 at an ectopic site promotes recombination in fission yeast. 11 24 2011 2023
Eydmann T. Sommariva E. Inagawa T. Mian S. Klar A. J. Dalgaard J. Z. 2008 Rtf1-mediated eukaryotic site-specific replication termination. 180 1 27 39
Codlin S. Dalgaard J. Z. 2003 Complex mechanism of site-specific DNA replication termination in fission yeast. 22 13 3431 3440
Dalgaard J. Z. Klar A. J. 2001 A DNA replication-arrest site RTS1 regulates imprinting by determining the direction of replication at mat1 in S. pombe. 16 15 2060 2068
Lambert S. Mizuno K. Blaisonneau J. Martineau S. Chanet R. Freon K. et al. 2010 Homologous recombination restarts blocked replication forks at the expense of genome rearrangements by template exchange. 39 3 346 359
Vengrova S. Codlin S. Dalgaard J. Z. 2002 RTS1-an eukaryotic terminator of replication. 34 9 1031 1034
Lee B. S. Grewal S. I. Klar A. J. 2004 Biochemical interactions between proteins and mat1 cis-acting sequences required for imprinting in fission yeast. 24 22 9813 9822
Coulon S Noguchi E Noguchi C Du LL Nakamura TM Russell P 2006 Rad22Rad52-dependent repair of ribosomal DNA repeats cleaved by Slx1-Slx4 endonuclease. 4 17 2081 2090
Kaplan D. L. Bastia D. 2009 Mechanisms of polar arrest of a replication fork. 72 2 279 285
Krings G. Bastia D. 2005 Sap1p binds to Ter1 at the ribosomal DNA of Schizosaccharomyces pombe and causes polar replication fork arrest. 280 47 39135 39142
Maric C. Levacher B. Hyrien O. 1999 Developmental regulation of replication fork pausing in Xenopus laevis ribosomal RNA genes. 291 4 775 788
Biswas S. Bastia D. 2008 Mechanistic insights into replication termination as revealed by investigations of the Reb1-Ter3 complex of Schizosaccharomyces pombe. 28 22 6844 6857
Zhao A. Guo A. Liu Z. Pape L. 1997 Molecular cloning and analysis of Schizosaccharomyces pombe Reb1p: sequence-specific recognition of two sites in the far upstream rDNA intergenic spacer. 25 4 904 910
Singh S. K. Sabatinos S. Forsburg S. Bastia D. 2010 Regulation of replication termination by Reb1 protein-mediated action at a distance. 142 6 868 78
Bochman M. L. Sabouri N. Zakian V. A. 2010 Unwinding the functions of the Pif1 family helicases. 9 3 237 49
Bairwa N. K. Zzaman S. Mohanty B. K. Bastia D. 2010 Replication fork arrest and rDNA silencing are two independent and separable functions of the replication terminator protein Fob1 of Saccharomyces cerevisiae. 285 17 12612 9
Evers R. Grummt I. 1995 Molecular coevolution of mammalian ribosomal gene terminator sequences and the transcription termination factor TTF-I. 92 13 5827 31
Gerber J. K. Gogel E. Berger C. Wallisch M. Muller F. Grummt I. et al. 1997 Termination of mammalian rDNA replication: polar arrest of replication fork movement by transcription termination factor TTF-I. 90 3 559 567
Langst G. Becker P. B. Grummt-I I. 1998 TTF-I determines the chromatin architecture of the active rDNA promoter. 17 11 3135 45
Arlt M. F. Durkin S. G. Ragland R. L. Glover T. W. 2006 Common fragile sites as targets for chromosome rearrangements. 5 9-10 1126 1135
Howlett N. G. Taniguchi T. Durkin S. G. D’Andrea A. D. Glover T. W. 2005 The Fanconi anemia pathway is required for the DNA replication stress response and for the regulation of common fragile site stability. 14 5 693 701
Bermejo R. Capra T. Gonzalez-Huici V. Fachinetti D. Cocito A. Natoli G. et al. 2009 Genome-organizing factors Top2 and Hmo1 prevent chromosome fragility at sites of S phase transcription. 138 5 870 884
Yarbro J. W. 1992 Mechanism of action of hydroxyurea. 9 1 10
Tsaponina O. Barsoum E. Astrom S. U. Chabes A. 2011 Ixr1 is required for the expression of the ribonucleotide reductase Rnr1 and maintenance of dNTP pools. 7 5 e1002061
Zhao X. Chabes A. Domkin V. Thelander L. Rothstein R. 2001 Thelander L, and Rothstein R. The ribonucleotide reductase inhibitor Sml1 is a new target of the Mec1/Rad53 kinase cascade during growth and in response to DNA damage.The EMBO journal 20 13 3544 3553
Huang A. Fan H. Taylor W. R. Wright J. A. 1997 Ribonucleotide reductase R2 gene expression and changes in drug sensitivity and genome stability. 57 21 4876 4881
Miyabe I. Morishita T. Shinagawa H. Carr A. M. 2009 Schizosaccharomyces pombe Cds1Chk2 regulates homologous recombination at stalled replication forks through the phosphorylation of recombination protein Rad60. Pt 20 3638 3643
Lindsay H. D. Griffiths D. J. Edwards R. J. Christensen P. U. Murray J. M. Osman F. et al. 1998 S-phase-specific activation of Cds1 kinase defines a subpathway of the checkpoint response in Schizosaccharomyces pombe. 12 3 382 395
Zhao H. Tanaka K. Nogochi E. Nogochi C. Russell P. 2003 Replication checkpoint protein Mrc1 is regulated by Rad3 and Tel1 in fission yeast. 23 22 8395 8403
Kumar D. Viberg J. Nilsson A. K. Chabes A. 2010 Highly mutagenic and severely imbalanced dNTP pools can escape detection by the S-phase checkpoint. 38 12 3975 3983
Zou L. Liu D. Elledge S. J. 2003 Replication protein A-mediated recruitment and activation of Rad17 complexes. 100 24 13827 13832
Kanoh Y. Tamai K. Shirahige K. 2006 Different requirements for the association of ATR-ATRIP and 9-1-1 to the stalled replication forks. 377 388
Kemp M. Sancar A. 2009 DNA distress: just ring 9-1-1. 19 17 R733 R734
Yan S. Michael W. M. 2009 TopBP1 and DNA polymerase-alpha directly recruit the 9-1-1 complex to stalled DNA replication forks. 184 6 793 804
Branzei D. Foiani M. 2006 The Rad53 signal transduction pathway: Replication fork stabilization, DNA repair, and adaptation. 312 14 2654 2659
Yan S. Michael W. M. 2009 TopBP1 and DNA polymerase alpha-mediated recruitment of the 9-1-1 complex to stalled replication forks: implications for a replication restart-based mechanism for ATR checkpoint activation. 8 18 2877 2884
Tanaka K. Russell P. 2001 Mrc1 channels the DNA replication arrest signal to checkpoint kinase Cds1. 3 11 966 972
Tanaka K. Russell P. 2004 Cds1 phosphorylation by Rad3-Rad26 kinase is mediated by forkhead-associated domain interaction with Mrc1. 279 31 32079 32086
Xu Y. J. Davenport M. Kelly T. J. 2006 Two-stage mechanism for activation of the DNA replication checkpoint kinase Cds1 in fission yeast. 20 8 990 1003
Osborn A. J. Elledge S. J. Zou L. 2002 Checking on the fork: the DNA-replication stress-response pathway. 12 11 509 516
Naylor M. L. Li J. M. Osborn A. J. Elledge S. J. 2009 Mrc1 phosphorylation in response to DNA replication stress is required for Mec1 accumulation at the stalled fork. 106 31 12765 12770
Schleker T. Nagai S. Gasser S. M. 2009 Posttranslational modifications of repair factors and histones in the cellular response to stalled replication forks. 8 9 1089 100
Tourriere H. Versini G. Cordon-Preciado V. Alabert C. Pasero P. 2005 Mrc1 and Tof1 promote replication fork progression and recovery independently of Rad53. 19 5 699 706
De Piccoli G. Katou Y. Itoh T. Nakato R. Shirahige K. Labib K. 2012 Replisome stability at defective DNA replication forks is independent of S phase checkpoint kinases. 45 5 696 704
Kim S. M. Huberman J. A. 2001 Regulation of replication timing in fission yeast. 20 21 6115 6126
Feng W. Bachant J. Collingwood D. Raghuraman M. K. Brewer B. J. 2009 Centromere replication timing determines different forms of genomic instability in Saccharomyces cerevisiae checkpoint mutants during replication stress. 183 4 1249 1260
Lengronne A. Pasero P. Bensimon A. Schwob E. 2001 Monitoring S phase progression globally and locally using BrdU incorporation in TK(+) yeast strains. 29 7 1433 1442
Sabatinos S. A. Mastro T. L. Forsburg S. L. 2012 Nucleoside analogues create DNA damage and sensitivity in fission yeast. submitted.
Chin J. K. Bashkirov V. I. Heyer WD Romesberg FE 2006 Esc4/Rtt107 and the control of recombination during replication. 5 5 618 628
Koren A. Soifer I. Barkai N. 2010 MRC1-dependent scaling of the budding yeast DNA replication timing program. 20 6 781 790
Hayano M Kanoh Y Matsumoto S Masai H 2011 Mrc1 marks early-firing origins and coordinates timing and efficiency of initiation in fission yeast. 12 31 2380 2391
Szyjka S. J. Viggiani C. J. Aparicio O. M. 2005 Mrc1 is required for normal progression of replication forks throughout chromatin in S. cerevisiae. 19 5 691 697
Xu Y. J. Kelly T. J. 2009 Autoinhibition and autoactivation of the DNA replication checkpoint kinase Cds1. 284 23 16016 16027
Hayashi M. Katou Y. Itoh T. Tazumi A. Yamada Y. Takahashi T. et al. 2007 Genome-wide localization of pre-RC sites and identification of replication origins in fission yeast. 26 5 1327 1339
Santocanale C. Diffley J. F. 1998 A Mec1- and Rad53-dependent checkpoint controls late-firing origins of DNA replication. 395 6702 615 618
Heichinger C. Penkett C. J. Bahler J. Nurse P. 2006 Genome-wide characterization of fission yeast DNA replication origins. 25 21 5171 5179
Chabes A. Stillman B. 2007 Constitutively high dNTP concentration inhibits cell cycle progression and the DNA damage checkpoint in yeast Saccharomyces cerevisiae. 104 4 1183 8
Mickle K. L. Ramanathan S. Rosebrock A. Oliva A. Chaudari A. Yompakdee C. et al. 2007 Checkpoint independence of most DNA replication origins in fission yeast. 8 112
Hanada K. Budzowska M. Davies S. L. van Drunen E. Onizawa H. Beverloo H. B. et al. 2007 The structure-specific endonuclease Mus81 contributes to replication restart by generating double-strand DNA breaks. 14 11 1096 1104
Robison J. G. Elliott J. Dixon K. Oakley G. G. 2004 Replication protein A and the Mre11.Rad50.Nbs1 complex co-localize and interact at sites of stalled replication forks. 279 33 34802 34810
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. 145 4 529 42
Ouyang K. J. Woo L. L. Zhu J. Huo D. Matunis M. J. Ellis N. A. 2009 SUMO modification regulates BLM and RAD51 interaction at damaged replication forks. 7 12 e1000252
Kurokawa Y. Murayama Y. Haruta-Takahashi N. Urabe I. Iwasaki H. 2008 Reconstitution of DNA strand exchange mediated by Rhp51 recombinase and two mediators. 6 4 e88
Wray J. Liu J. Nickoloff J. A. Shen Z. 2008 Distinct RAD51 associations with RAD52 and BCCIP in response to DNA damage and replication stress. 68 8 2699 2707
Lambert S. Froget B. Carr A. M. 2007 Arrested replication fork processing: interplay between checkpoints and recombination. 6 7 1042 1061
Aggarwal M. Sommers J. A. Morris C. Brosh R. M. Jr 2010 Delineation of WRN helicase function with EXO1 in the replicational stress response. 9 7 765 776
Tinline-Purvis H. Savory A. P. Cullen J. K. Dave A. Moss J. Bridge W. L. et al. 2009 Failed gene conversion leads to extensive end processing and chromosomal rearrangements in fission yeast. 28 21 3400 3412
Tran P. T. Fey J. P. Erdeniz N. Gellon L. Boiteux S. Liskay R. M. 2007 A mutation in EXO1 defines separable roles in DNA mismatch repair and post-replication repair. 6 11 1572 1583
Lambert S. Watson A. Sheedy D. M. Martin B. Carr A. M. 2005 Gross chromosomal rearrangements and elevated recombination at an inducible site-specific replication fork barrier. 121 5 689 702
Meister P. Taddei A. Ponti A. Baldacci G. Gasser S. M. 2007 Replication foci dynamics: replication patterns are modulated by S-phase checkpoint kinases in fission yeast. 26 5 1315 1326
Willis N. Rhind N. 2010 The fission yeast Rad32(Mre11)-Rad50-Nbs1 complex acts both upstream and downstream of checkpoint signaling in the S-phase DNA damage checkpoint. 184 4 887 897
Hashimoto Y. Puddu F. Costanzo V. 2012 RAD51- and MRE11-dependent reassembly of uncoupled CMG helicase complex at collapsed replication forks. 19 1 17 24
Brugmans L. Verkaik N. S. Kunen M. van Drunen E. Williams B. R. Petrini J. H. 2009 NBS1 cooperates with homologous recombination to counteract chromosome breakage during replication. 8 12 1363 1370
Kuzminov A. 2001 Single-strand interruptions in replicating chromosomes cause double-strand breaks. 98 15 8241 8246
Caldecott K. W. 2007 Mammalian single-strand break repair: mechanisms and links with chromatin. 6 4 443 453
Hutchinson F. 1993 Induction of large DNA deletions by persistent nicks: a new hypothesis. 299 3-4 211 218