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Vol. 20, Issue 3, 819-833, February 1, 2009
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Department of Biochemistry and Molecular Pharmacology, University of Massachusetts Medical School, Worcester, MA 01605
Submitted August 5, 2008;
Revised October 20, 2008;
Accepted November 19, 2008
Monitoring Editor: Orna Cohen-Fix
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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mutants display a complete checkpoint defect. We have identified proteins downstream of Cds1 required for checkpoint-dependant slowing, including the structure-specific endonuclease Mus81 and the helicase Rqh1, which are implicated in replication fork stability and the negative regulation of recombination. Removing Rhp51, the Rad51 recombinase homologue, suppresses the slowing defect of rqh1 mutants, but not that of mus81 mutant, defining an epistatic pathway in which mus81 is epistatic to rhp51 and rhp51 is epistatic to rqh1. We propose that restraining recombination is required for the slowing of replication in response to DNA damage. |
INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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). Checkpoint activation prevents entrance into S phase in the presence of DNA damage in G1, prevents entrance into mitosis in the presence of damaged or unreplicated DNA, and reduces replication rate in the presence of damaged template during S phase. The mechanism of the G1 and G2 DNA damage checkpoints are well described (Lukas et al., 2004). However, mechanisms used to slow replication in response to DNA damage during S phase are less clear. Two mechanisms have been invoked for replication slowing in response to DNA damage: reduced replication fork rate and reduced origin firing (Tercero and Diffley, 2001; Merrick et al., 2004). Prevention of origin firing requires factors to act on origins located far from sites of DNA damage and thus represents an in trans mechanism, whereas reduced fork rate likely represents an in cis mechanism in which factors act locally at the site of DNA damage to slow the affected fork. Slowing in response to DNA damage in higher eukaryotes seems to be a combination of both direct action on replication forks and reduced origin firing (Seiler et al., 2007).
In the fission yeast Schizosaccharomyces pombe, the well-established S-phase DNA damage checkpoint signaling cascade includes Rad3, the central checkpoint kinase and homologue of the metazoan ATR kinase, and Cds1, homologue of the Rad53 and Chk2 effector kinases (Lindsay et al., 1998; Rhind and Russell, 1998). However, the checkpoint targets underlying replication slowing are not well understood (Figure 1F). Replication slowing in metazoans is catalyzed by parallel pathways acting downstream of the central checkpoint kinases (Falck et al., 2002; Henry-Mowatt et al., 2003). One pathway depends upon Chk2/Cds1 negative regulation of Cdc25-dependent origin firing; another is dependent upon the Mre11–Rad50–Nbs1 (MRN) recombinational repair complex, which may reflect direct regulation of replication fork progression (Falck et al., 2002). Only when both MRN and Cdc25 mediated pathways are compromised do mammalian cells display a complete failure of the S-phase DNA damage checkpoint similar to ATM mutants (Falck et al., 2002). In S. pombe, Cdc25 is not required for the S-phase DNA damage checkpoint (Kommajosyula and Rhind, 2006). However, members of the MRN complex are required (Chahwan et al., 2003; Kommajosyula and Rhind, 2006). The lack of Cdc25 involvement in the S phase DNA damage response suggests either that regulation of origin firing is not required for replication slowing or that origins are regulated in a manner independent of Cdc25.
Replication fork response to DNA damage involves fork stalling, recombination and DNA repair (Michel et al., 2004). Recombination allows for error-free repair of damaged DNA through strand exchange between homologous sequences. In fission yeast, this exchange is catalyzed by the central mitotic recombinase Rhp51, homologue of bacterial RecA and budding yeast Rad51 (Muris et al., 1993). Rhp51 is loaded onto the 3'-end of single-stranded DNA by the mediator Rad22, homologue of Rad52. Rhp51 then forms a nucleoprotein filament on single-stranded DNA (ssDNA), which is stabilized and regulated by the additional Rhp51 mediators Rhp54, Rhp55, Rhp57, Sfr1, and Swi5 (Raji and Hartsuiker, 2006). Single-fiber analysis shows the Rad51 recombinase and its paralogue XRCC3 are required for slowing of replication fork progression in response to cisplatin and UV in mammalian cells (Henry-Mowatt et al., 2003).
Although beneficial, recombination must be tightly regulated for cells to properly respond to DNA damage and fork stalling. One mechanism for controlling recombination involves the RecQ helicase Rqh1, which is implicated in negatively regulating recombination. Cells lacking Rqh1 display phenotypes attributed to inappropriate, ectopic recombination including hyper-recombination, sensitivity to replication arrest and sensitivity to UV-induced DNA damage (Doe et al., 2000). The DNA damage sensitivity displayed by rqh1 cells is alleviated by disrupting recombination (Hope et al., 2005). The structure-specific endonuclease Mus81 is also implicated in negatively regulating recombination. Like the helicase mutants, mus81 mutants display a mutator phenotype and sensitivity to hydroxyurea and UV (Boddy et al., 2000; Kai et al., 2005). Consistent with our results summarized below, the DNA damage sensitivity of mus81 and synthetic lethality of the mus81 rqh1 mutant is not rescued by removing recombination (Doe and Whitby, 2004).
The hallmark of the S-phase DNA damage checkpoint in the fission yeast S. pombe is slowing of replication in response to DNA damage (Lindsay et al., 1998; Rhind and Russell, 1998). With this in mind, we used cell-cycle synchronization and flow cytometry to measure bulk replication slowing in response to DNA damage. We show that several proteins in addition to the checkpoint kinases are required for replication slowing in fission yeast: the Rqh1 helicase, the Mus81 endonuclease, and the Sfr1 mediator. We show that the slowing defects displayed by rqh1 and sfr1 strains are suppressed by abrogating recombination, but that the defects displayed by mus81 and cds1 mutants are not. Previous experiments using these mutants have shown the importance of recombinational control in proper repair of DNA lesions after replication (Laursen et al., 2003). Our data illustrate the importance of proper control of recombination in response to DNA damage during replication itself.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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). Yeast strains are listed in Table 1. Strains containing the cds1 overexpression cassette were grown in EMM supplemented with 15 µM thiamine to repress nmt1 transcription. Thiamine was washed out 16 h before G1 elutriation. The rad51::natMX6 allele was constructed using a polymerase chain reaction (PCR)-based targeting method replacing the open reading frame of rhp51 with the nourseothricin resistance gene natMX6 (Bahler et al., 1998; Sato et al., 2005). We targeted rhp51 using PCR product amplified from the pCR2.1-nat plasmid using the following primers, 5'-ctaatctttcttttctttaataatataaaaaactcttttcaattccagaatagtgataatttcgtgcttaacaagttatacggatccccgggttaattaa and 5'- cacatacatatctatccttacaaactcatcccatagaatttgcaaaataataaataaaaatgaaacgatactaaaataatgaattcgagctcgtttaaac.
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Isolated Nuclei Preparation for Flow Cytometry
Whole cells arrested using cdc10 temperature-sensitive alleles or treated with hydroxyurea display significant background fluorescence when analyzed by flow cytometry. The background increases during the arrest because cells continue to elongate. To reduce this background and improve the signal-to-noise ratio in our experiments, we adapted a protocol to analyze isolated nuclei (Carlson et al., 1997). Fixed cells were pelleted and washed with 1 ml of 0.6 M KCl. Cells were spheroplasted by resuspension in 1 ml of 0.6 M KCl containing 0.5 mg/ml Zymolyase 20T and 1.3 mg/ml lysing enzyme (L1412; Sigma-Aldrich) and incubated at 37°C for 30 min. Spheroplasts were pelleted, washed with 1 ml of 0.1% Triton X-100, 0.1 M KCl, and finally washed with and resuspended in 1 ml of 20 mM Tris-HCl, pH 8.0, 1 mM EDTA, pH 8.0. 20 µl of RNase A (10 mg/ml) was added to each preparation followed by overnight incubation at 37°C. Spheroplasts were pelleted, left in buffer, and chilled to 0°C. Spheroplasts were disrupted and nuclei released using a Branson Sonifier 450 set to 0.6 power for 10 s and a chilled microtip. 300 µl of sonicated cells was mixed with 300 µl of 2 µM Sytox Green (S7020; Invitrogen, Carlsbad, CA) in 1x phosphated-buffered saline (PBS) and analyzed on a FACScan flow cytometer (BD Biosciences, San Jose, CA).
S-Phase Progression Analysis
S phase progression was quantitated using CellQuest software version 3.3 (BD Biosciences). The 1C unreplicated value was determined by measuring DNA content of freshly elutriated samples, and the 2C fully replicated value was determined by measuring DNA content from later time points for untreated cultures and asynchronous, prearrested samples. Mean values for S-phase peaks after release were compared with 1C and 2C values. S-phase progression was quantitated using the following equation: % progression = (C – A)/(B – A), where A = 1C, B = 2C, and C = mean histogram value. Error bars represent the SE of the mean for strains in which three or more experiments were performed. For strains for which two experiments were performed, error bars represent the variance. The error bars may be smaller than data symbols used. See Figure 7 for the number of experiments performed for each individual strain.
In Vitro Kinase Assay
G1 cells were synchronized as described above and kinase activity measured as described previously (Lindsay et al., 1998; Kai et al., 2005). 0.5 ODs of cells were fixed for flow cytometry, and 5 ODs of cells were pelleted and frozen in liquid nitrogen. Pellets were thawed on ice in 200 µl of ice-cold lysis buffer (150 mM NaCl, 50 mM Tris-HCl, pH 8.0, 5 mM EDTA, pH 8.0, 10% glycerol, 1% NP-40, 50 mM NaF, 5 µg/ml leupeptin, 1 mM phenylmethylsulfonyl fluoride, and 1 mM Na3VO4) and broken by vortexing with silica bead for 15 min at 4°C. Crude anti-Cds1 antibodies (a kind gift from T. Wang) were conjugated to protein A-Sepharose beads by constant mixing at 4°C for 1 h in lysis buffer, washed in ice-cold lysis buffer, and resuspended as a 1:1 slurry in lysis buffer. Crude cell lysates were cleared by brief centrifugation (3000 x g; 4 min; 4°C); cleared lysate protein concentration determined by BCA Protein Assay kit (23225; Pierce Chemical, Rockford, IL). Protein concentration was normalized within each experiment in a final volume of 200 µl. Twenty microliters of the 1:1 antibody-protein A bead slurry was added to each sample, and mixtures were incubated with constant mixing at 4°C for 2 h. Beads were washed twice with ice-cold lysis buffer, twice with ice-cold 1x kinase buffer (5 mM HEPES, pH 7.5, 37.5 mM KCl, and 2.5 mM MgCl2), and resuspended in 10 µl of 1x kinase buffer.
To each sample, 10 µl of 2x kinase buffer (10 mM HEPES, pH 7.5, 75 mM KCl, and 5 mM MgCl2), 0.5 µl of 10 µCi/µl [
-32P]dATP, 2 µl of 1 mM dATP, and 5 µl of 1 mg/ml myelin basic protein [M1891; Sigma-Aldrich]) were added, mixed and incubated at 30°C for 15 min. Reactions were stopped by addition of 20 µl of 4x protein loading buffer (40% glycerol, 200 mM Tris-HCl, pH 6.8, and 4% SDS) and boiled for 5 min at 95°C. Reactions were allowed to cool to room temperature, and beads were pelleted. Twenty microliters of the supernatants was run on a 15% SDS-polyacrylamide gel, Coomassie stained to visualize myelin basic protein target, dried for 30 min at 90°C under vacuum, and exposed to a phosphorimager screen for 6–72 h. Screens were developed on a FLA-5000 PhosphorImager, and bands were quantitated using FujiFilm ImageGauge software (FujiFilm, Tokyo, Japan). For comparison between strains and independent experiments, kinase activity was normalized to percentage of G1 cells at time 0 (G2 cells treated with 0.03% MMS neither septate nor replicate and do not contribute to measured Cds1 activity), and values were normalized to kinase activity in MMS-treated wild-type cells 120 min after elutriation.
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Data collected by flow cytometry was analyzed as histogram stacks shown in Figure 1A. The untreated wild-type population in the left-hand stack displays a distinct shift from 1C toward 2C around 80 min after release, whereas MMS-treated cells display no such dramatic shift but a slow progression from 1C toward 2C with time. As reported previously, cells lacking Rad3, the central checkpoint kinase in fission yeast, replicate quickly both in the presence and absence of DNA damage, confirming that the checkpoint is required for replication slowing in response to DNA damage (Figure 1A, right; Lindsay et al., 1998; Rhind and Russell, 1998).
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40% in the presence of MMS (Figure 1B, closed vs. open diamonds), whereas the rad3 strain did not (Figure 1B, closed vs. open triangles). To illustrate the difference between wild-type and rad3 strains, the extent of replication at a single time point, 140 min after release, designated Rep140, is shown for both strains under both conditions (Figure 1C). Cds1, the S-phase-specific effector kinase, and Mrc1, the S phase checkpoint mediator protein required for Rad3-dependent Cds1 phosphorylation and activation, are also required for DNA damage-induced replication slowing (Figure 4D and Supplemental S3A; Lindsay et al., 1998; Rhind and Russell, 1998). Moreover, replication slowing observed in all mutants is checkpoint dependent (Figure 7 and Supplemental S3B and C).
Checkpoint Signaling Is Not Sufficient to Produce Slowing
DNA damage during S phase induces checkpoint signaling and activates the effector kinase Cds1 as measured by in vitro kinase assay (Lindsay et al., 1998). Using an in vitro kinase assay to quantitate the activity of Cds1 immunoprecipitated from cell lysates, we confirmed that untreated wild-type cells display slightly increased, periodic kinase activity during S phase and strong activity when exposed to MMS during replication (Figure 1, D and E). rad3 strains failed to slow replication and displayed no increased Cds1 activity in unperturbed S phase or in response to DNA damage during replication (Figure 1, D and E).
Although Cds1 activation correlates with replication slowing, exposure to ionizing radiation (IR) during S phase, which induces Cds1 signaling, does not induce slowing (Rhind and Russell, 1998). Therefore, we suspected that Cds1 signaling was not sufficient for slowing. We tested for sufficiency by comparing S-phase slowing and Cds1 signaling responses to IR, bleomycin, and MMS during S phase. Exposing cells to 200 Gray IR or 1 µg/ml IR-mimetic bleomycin, dosages capable of producing a mitotic delay, induces Cds1 kinase activity, but not slowing (Figure 2, A and B). Bleomycin titration experiments show that 9 µg/ml produced Cds1 kinase activity equivalent to that in MMS-treated cells but this activity was still not sufficient to induce a replication slowing response (Figure 2, D and E). Therefore, simply activating Cds1 is not sufficient to cause slowing of replication.
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; de Bruin et al., 2008). We overexpressed Cds1 from the strong, thiamine-regulated nmt1 promoter in a rad3 background so that Cds1 activity could be ascribed to overexpression and not checkpoint signaling. To prevent a Cds1-induced G2 arrest, we used a genetic background in which the G2 arrest is overridden by deletion of mik1, a Cdc2-inhibitory tyrosine-kinase activated by Cds1, and overexpression of Pyp3, a Cdc2 activating tyrosine-phosphatase that can constitutively drive the G2/M transition (Rhind and Russell, 2001). Cds1 overexpression induces Cds1 kinase activity in the rad3 mutant with no significant increase in activity when cells are treated with MMS (Figure 3A). Overexpressing Cds1 did not cause replication slowing nor did MMS-treatment of the nmt1:cds1 rad3 cells when Cds1 is repressed. However, MMS treatment of Cds1-overexpressing cells caused robust replication slowing (Figure 3B). The slowing displayed is kinase-dependent because overexpression of a kinase-dead allele of cds1 failed to induce MMS-dependent replication slowing (Figure 7). Bleomycin treatment did not induce slowing upon Cds1 overexpression, further suggesting that replication slowing requires a high density of DNA lesions (Figure 3B). Although slowing is normally dependent upon the mediator protein Mrc1 (Supplemental Figure S3A), slowing upon Cds1 overexpression is not, suggesting that Mrc1's main role in the checkpoint is the activation of Cds1 (Figure 3C). These results show that, although checkpoint signaling through Cds1 is necessary for slowing replication in response to DNA damage, Cds1 activity alone is not sufficient.
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), we wished to assay other proteins involved in recombination. We conducted a survey of additional mutants involved in recombination. We found three strains, rqh1, sfr1, and mus81, that do not slow replication in response to DNA damage (Figure 4A). In addition to other mutants discussed below, we found a number of strains that display normal replication slowing in response to DNA damage, including swi6, clr4, swi10, srs2, and tel1. Rqh1 is a DNA helicase implicated in replication fork stability and in negative regulation of recombination in response to replication stress (Murray et al., 1997). Sgs1, the Rqh1 homologue in budding yeast, has been shown previously to be involved in replication slowing in response to MMS, although the replication slowing phenotype of sgs1 cell is much weaker than the of rqh1 cells (Frei and Gasser, 2000). Sfr1 is a recently discovered mediator of the Rhp51 recombinase (Akamatsu et al., 2003). Mus81 is a structure specific endonuclease required for meiotic recombination and regulation of recombination in response to replication stress (Boddy et al., 2000; Gaillard et al., 2003; Kai et al., 2005). Recent work supports a role for Mus81 in replication-dependent recombination between sister-chromatids (Roseaulin et al., 2008). mus81 strains do not slow replication in response to DNA damage; rqh1 and sfr1 strains display a strong defect with minor residual checkpoint-dependent slowing (Figure 4A and Supplemental S3B).
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). We observe no defect in replication slowing in response to DNA damage in mus81-T239A cells, indicating that T239-dependent phosphorylation by Cds1 is not required for Mus81's role in replication slowing (Figure 4A). Mus81 forms a heterodimer with Eme1, which is required for in vitro endonuclease activity (Boddy et al., 2001). Similar to Mus81, Eme1 is required for replication slowing (Figure 7).
Rqh1, Sfr1, and Mus81 Are Not Required for Cds1 Activation
Because Cds1 activity is necessary for replication slowing in response to DNA damage, it was important to determine whether rqh1, sfr1, and mus81 mutations influenced checkpoint signaling. Both rqh1 and sfr1 strains displayed strong MMS-induced Cds1 signaling (Figure 4B). In contrast, mus81 displayed 30–50% reduction in Cds1 kinase activity (Figure 4B). However, mus81-T239A shows a reduction in Cds1 kinase activity similar to mus81 but slows replication normally. Thus, reduced kinase activity is not sufficient to cause the slowing defect observed in the mus81 mutant (Figure 4B). These results suggest that Rqh1, Sfr1 and Mus81 act downstream of, or parallel to, the checkpoint kinase Cds1.
Recombinases Are Not Required for Replication Slowing
In contrast to the sfr1 mutant slowing defect, rhp51 mutants slow like wild-type (Figure 5A). Rhp51, the S. pombe Rad51/RecA homologue, is the central mitotic recombinase required for the majority of homologous recombination events that occur in vegetative cells. We found that the first rhp51 strain we tested failed to slow (Supplemental Figure S1E). However, this phenotype is due to an unlinked modifier (Willis and Rhind, unpublished data). We deleted rhp51 in a wild-type background and found no slowing defect (Figure 5A). We used this allele for all our subsequent assays and crosses.
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and dmc1 rhp51 mutants were tested. Both the single and double mutants slowed normally in response to MMS (Figure 7 and Supplemental S1A). To show that recombination is not required for checkpoint signaling, in vitro kinase assays were performed comparing rhp51 to wild-type strains. rhp51 mutants displayed wild-type Cds1 activity upon exposure to DNA damage during replication (Figure 5C). Thus, the recombinases are not required for replication slowing or to produce DNA structures required to activate the checkpoint.
In addition to the Rhp51 recombinase, we tested whether the central Rhp51 mediator, Rad22, was required for slowing. Rad22 is the fission yeast homologue to budding yeast Rad52, which is responsible for loading Rhp51 onto 3' end of ssDNA (Kim et al., 2002). Some recombination events in fission yeast such as single-strand annealing are Rad22 dependent but Rhp51 independent (Osman et al., 2000). rad22 strains slowed normally indicating that neither Rhp51-dependent nor -independent recombination events are required for slowing (Figure 7 and Supplemental S1A).
The sfr1 and rqh1 Slowing Defects Are Recombinase Dependent
Although Rqh1, Sfr1, and Mus81 are required for replication slowing in response to DNA damage, homologous recombination itself is not. Surprisingly, Rhp51 is required for the sfr1 and rqh1 strains to display a slowing defect, because sfr1 rhp51 and rqh1 rhp51 double mutants slow in response to DNA damage (Figure 5B). Additionally, deletion of dmc1 also rescues the slowing defects of sfr1 and rqh1 mutants (Figure 7 and Supplemental S1B). Dmc1 has been characterized as a meiotic recombinase (Fukushima et al., 2000). However, microarray transcriptional profiling has shown Dmc1 to be expressed in vegetative cells (Rustici et al., 2004). The suppression of the sfr1 and rqh1 slowing defects by rhp51 or dmc1 is checkpoint dependent because triple mutants harboring a deletion of cds1 fail to slow replication (Figure 7 and Supplemental S3C). Unlike the sfr1 and rqh1 slowing defects, recombination is not required for the mus81 defect. mus81 rhp51 and mus81 dmc1 strains still display the mus81 nonslowing phenotype (Figure 5B, and Supplemental S1B). In addition, mus81 rad22 double mutants also display the mus81 nonslowing phenotype, indicating that neither Rhp51-dependent nor -independent recombination events are required for mus81 cells to replicate quickly through damaged DNA (Figure 7 and Supplemental S1D).
The Swi2–Swi5 Complex Is Required for Defects Displayed by sfr1 and rqh1 Mutants
Because the sfr1 mutant displayed a failure to slow, we tested whether its partner Swi5 is also required for slowing. Sfr1 and Swi5 have been shown to act as a heterodimer to promote Rhp51-dependent recombination in vegetative cells. In vitro, this complex is able to load Rhp51 and Dmc1 onto ssDNA to promote strand-exchange reactions (Haruta et al., 2006). Surprisingly, swi5 strains slowed like wild-type in a checkpoint-dependent manner indicating that Swi5 is not required for slowing in response to DNA damage (Figure 5A and Supplemental Figure S3B). Furthermore, swi5 deletion suppressed the sfr1 slowing defect (Figure 5B). This result indicates that, like Rhp51 and Dmc1, Swi5 is required for the quick replication through damaged DNA displayed by sfr1 mutants. Like sfr1, the rqh1 mutant defect is suppressed by swi5 deletion (Figure 5B). Also, the rqh1 sfr1 double mutant failed to slow and the rqh1 swi5 sfr1 triple mutant slowed like wild-type (Figure 7 and Supplemental S1C). Similar to the lack of suppression of the mus81 slowing defect by rhp51 or dmc1, swi5 had no effect on the failure of mus81 to slow (Figure 4B).
The Rhp51 mediator Swi5 forms two independent protein complexes by binding to either Sfr1 or Swi2. The Swi2–Swi5 complex is required for efficient mating-type switching, whereas the Swi5–Sfr1 complex promotes homologous recombinational repair (Akamatsu et al., 2003). With only the sfr1 mutant displaying a slowing defect, and this phenotype being Swi5 dependent, we tested what role Swi2 played in Sfr1-dependent replication slowing. swi2 single mutants slow normally and swi5 swi2 double mutant also slowed, indicating that the Swi5–Swi2 complex is not required for replication slowing in response MMS (Figure 6A). Similar to swi5, deletion of swi2 suppresses the slowing defects of sfr1 and rqh1 but has no effect on the mus81 defect (Figure 6B).
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Slowing Defects). To determine whether fork stability plays a role in slowing in response to DNA damage, we tested the role of Swi1. swi1 cells slow normally, indicating the Fork Protection Complex is not required for the checkpoint (Figure 5A). However, Swi1 is required for sfr1 strains to fail to slow, much as Rhp51 is (Figure 5B). Suppression of the rqh1 and mus81 strains by swi1 could not be determined because mus81 swi1 and rqh1 swi1 are synthetic lethal (Willis, unpublished data; Noguchi et al., 2004).
Suppression of Slowing Defects Does Not Act by Modulation of Checkpoint Signaling
Exposure to lower concentrations of MMS produces less slowing in wild-type strains, suggesting that increased checkpoint signaling may lead to increased replication slowing (Figure 2F). swi5 mutants display slightly increased checkpoint signaling over wild type (Figure 5C). Increased checkpoint signaling strength may be related to increased replication slowing and could be responsible for the slowing displayed in the sfr1 swi5 double mutant. However, suppression of the sfr1 defect does not correlate with increased signaling because the sfr1 and sfr1 swi5 strains displayed a similar level of Cds1 kinase activity in response to MMS (Figure 5C).
The Role of Other Rhp51 Mediators in Replication Slowing
Given the importance of the Rhp51 mediator Sfr1 for replication slowing, we tested whether additional Rhp51 mediator mutants were also required for slowing. rhp54, rhp55, and rhp57 mutants all slow normally (Figure 7 and S2A). To rule out redundant functions among the three mediators, double mutants were constructed and tested. All double mutants slowed replication well (Figure 7 and Supplemental S2A). These data indicates that disruption of Rhp51 mediator activity does not necessarily impact cells' ability to slow replication in response to DNA damage.
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are recombination dependent and alleviated by disruption of rhp51 or the recombinase mediators swi5 and rhp55 (Hope et al., 2005). Individually, mutation of mediators rhp54, rhp55, or rhp57 on an rqh1 background did not suppress the rqh1 slowing defect. However, rqh1 mutants containing deletions for any two of the three recombinase mediators did slow (Figure 7 and Supplemental S2B). These data show that inhibition of recombination, either by deletion of swi5 or the disruption of several other mediators, prevents the rqh1 slowing defect. A similar relationship was established between these mediators and Sfr1 (Supplemental Figure S2B).
Slowing of Replication Does Not Correlate with Resistance to DNA Damage
The genetic interactions between recombinational repair genes and checkpoint-dependent slowing of replication raise the issue of whether slowing directly contributes to resistance to DNA damage. To investigate the relationship between slowing and resistance, we assayed MMS sensitivity in various slowing and nonslowing strains. We found no correlation between the ability to slow replication in response to DNA damage and sensitivity to MMS (Figure 8). Both wild-type and the cds1 mutant, strains that slow and fail to slow, respectively, tolerate acute exposure to MMS. Conversely, the rhp51 and rqh1 mutants are both sensitive to MMS exposure; yet, the rhp51 strain is still able to slow replication in response to MMS. These results show that slowing of replication is not necessary for cell to repair DNA damage and suggest that the choice between slowing and not slowing may instead reflect the choice between alternative repair pathways.
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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mutants and the nonslowing phenotype displayed by rqh1, sfr1 and mus81.
Three Epistasis Groups Regulate Replication Slowing
Our genetic analysis identified three epistasis groups with respect to S-phase DNA damage checkpoint-regulated slowing of replication (Figure 9). Group I includes the cds1 and mus81 mutants, both of which display a nonslowing phenotype not suppressed by removing recombination. Group II includes the rhp51 and dmc1 recombinase mutants, the swi2, swi5, rhp54, rhp55, and rhp57 mediator mutants and the swi1 Fork Protection Complex mutant, all of which slow replication in response to DNA damage. Group III consists of the sfr1 and rqh1 mutants, which display a nonslowing phenotype similar to that of group I except that it is dependent upon recombination. Group I is epistatic to group II because double mutants between members of each group display the group I nonslowing phenotype. Group II is epistatic to group III because double mutants between members of these groups display the group II slowing phenotype. It should be noted that these epistasis groups are simply formal genetic relationships and do not imply any biochemical interactions. For example, Cds1 and Mus81 clearly have different biochemical roles in the checkpoint, but null mutants have the same slowing phenotype and the same epistatic interactions with rhp51; therefore, they are formally in the same epistasis group. Moreover, although the genes in groups I and III have the same phenotypes, they can be placed in separate groups because they have opposite epistatic interactions with the genes in group II.
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We suspect that the difference in response to MMS and bleomycin is due to the quantitative differences between the lesions induced (Figure 2D; Rhind and Russell, 1998). MMS (0.03%) and 200 J/m2 UV are each expected to produce tens of thousands of adducts leading to a lesion density of close to one every 500 base pairs (Sedgwick, 1975; Courcelle et al., 2006). Therefore, replication forks would frequently encounter damage and even minor slowing at each lesion could add up to a significant effect on bulk replication. By contrast, 200-Gray IR and 1 µg/ml bleomycin are each expected to produce only 15 double-strand breaks in a fission yeast genome (Povirk et al., 1977; Prise et al., 1998). Even if each break permanently arrests two replication forks, such treatment would reduce the number of forks by only 5%, a reduction that would be undetectable with currently available techniques.
To further discriminate between cis and trans effects on slowing, we measured slowing upon overexpression of Cds1. Because Cds1 overexpression produces kinase activity capable of inducing a G2/M arrest and checkpoint transcription in the absence of DNA damage (Brondello et al., 1999; de Bruin et al., 2008), we determined whether Cds1 overexpression could produce slowing in the absence of DNA damage. To exclude Rad3-dependent effects on Cds1 activity we overexpressed Cds1 on a rad3 background. Cds1 overexpression results in the accumulation of Cds1 kinase activity but failed to induce damage-independent slowing, suggesting that kinase activity alone is insufficient to induce replication slowing (Figure 3A). However, rad3 strains overexpressing Cds1 do slow in the presence of MMS, showing that overexpressed Cds1 activity is sufficient to produce slowing, but only in the presence of appropriate DNA damage (Figure 3, A and B). Similarly, rad3 mrc1 double mutants overexpressing Cds1 also slow but only in response to DNA damage. Consistent with our in vivo data, high concentration of Cds1 in vitro stimulates Cds1 autophosphorylation and kinase activity (Xu et al., 2006). We propose that upon Cds1 overexpression, active Cds1 is able to interact with replication forks independently of the upstream checkpoint components Rad3 and Mrc1. The overexpression of a kinase-dead allele of Cds1 has no effect, suggesting that Cds1 has no kinase-independent role in the checkpoint (Figure 7). We interpret these results to mean that Cds1 must be catalytically active at individual forks as they encounter damage for the forks to slow. Moreover, because Cds1 is activated in these experiments by overexpression and is not influenced by Rad3-dependent checkpoint signaling, these results cannot be affected by any potential qualitative difference in Cds1 signaling in response to MMS- and bleomycin-induced DNA damage. We believe our results are inconsistent with in trans models of either global fork slowing or global origin inhibition.
Our Cds1 overexpression experiments also confirm the conclusion from previous work that Rad3 acts through Cds1 in the S phase DNA damage checkpoint (Lindsay et al., 1998; Martinho et al., 1998; Brondello et al., 1999). These results show that Cds1 kinase activity is sufficient for replication slowing and suggest that Rad3 has no Cds1-independent role in the checkpoint. Conversely, that normally expressed Cds1 shows no genetic or biochemical activity in the absence of Rad3, suggests that Cds1 normally has no Rad3-independent function.
An alternate model to explain the failure to slow bulk replication displayed by the rqh1, sfr1, and mus81 mutants is that fork slowing is the primary mechanism of the checkpoint in wild-type cells but that the failure to slow in checkpoint mutants is due to an abnormal increase in origin firing, not a decrease in fork slowing. Deregulation of origin firing would allow for more origins to fire than normal, producing more forks in the presence of damage and leading to fast bulk-S-phase progression even though individual forks are slowed. This model is motivated by the observation that in budding yeast, MMS causes a reduction in fork rates independent of checkpoint activity (Tercero and Diffley, 2001). However, several observations suggest that deregulated origin firing is not the cause of the failure to slow seen in fission yeast checkpoint mutants. First, even if all dormant origins were to fire, it would only increase the total number of forks by approximately threefold (Heichinger et al., 2006)—not enough to compensate for the >2-h S phase delay caused by MMS treatment. Second, the requirement for the three recombinational regulators for slowing and the genetic interaction between rqh1, sfr1 and rhp51 suggest that suppression of slowing involves strand exchange between nascent chromatids. Third, if slowing were produced by a combination of origin firing inhibition and slowing of forks (either in a local or global manner) then overexpression and activation of Cds1 would be expected to produce a partial slowing phenotype, which it does not. Together, all of these observations suggest that failure to slow in mutants that disrupt the S-phase DNA damage checkpoint is caused by an increase in the rate of replication fork progression. Nonetheless, because our flow cytometry assay measures only bulk replication, we cannot rule out more complicated possibilities. A rigorous demonstration of fork slowing and its abrogation in checkpoint mutants will require direct measurement of fork rates, which has not yet been reported in fission yeast.
A Model for the Regulation of Replication Slowing in Response to DNA Damage
We propose the following model for checkpoint-dependent replication slowing in which replication fork progression at sites of damage is regulated by a balance between the checkpoint, which promotes replication slowing at lesions, and recombination, which facilitates fast replication through damage (Figure 9). We imagine that leading-strand lesions would have a much greater affect on bulk replication speed, because lesions on the lagging strand can be easily bypassed by repriming. By default, replication forks traverse the genome quickly and bulk replication is completed in 20 min, even in the presence of DNA damage (Figure 1). Replication slowing requires Cds1 checkpoint kinase activation and the Mus81 endonuclease, which promote slowing independent of recombination. However, replication slowing does require negative regulation of recombination by Rqh1 and Sfr1. In particular, recombination interferes with replication fork slowing when Rqh1 or Sfr1 are absent by allowing forks to replicate quickly through damaged DNA (Figure 9). In this model, upon encountering DNA damage, replication forks serve as both the essential substrates for checkpoint activation and as the direct targets of the checkpoint (Tercero et al., 2003). It is possible that the decision to replicate damaged DNA slowly or quickly is an entirely local decision—that checkpoint activation at a single fork is sufficient to cause that fork to slow and may not significantly effect the rest of the cell. Alternatively, it may be necessary to have a threshold level of checkpoint signaling in the cell for any fork to slow, thereby limiting replication slowing to situations in which multiple forks are encountering damage (Shimada et al., 2002). Distinguishing these two possibilities will require techniques that can assay the slowing of individual replication forks.
The biological function of such a model is unclear, especially because we find no correlation between ability to slow replication and resistance to MMS (Figure 8). Several extremely sensitive mutants slow well, whereas others not sensitive fail to slow. We attribute the DNA damage sensitivity in our strains to a DNA damage repair defect in G2 and speculate that any benefit from slowing replication is masked by the greater importance of the G2 repair. This lack of correlation between slowing and resistance has been observed in mammalian cells, as well. However, human patients that lack the S phase DNA damage checkpoint are predisposed chromosome rearrangements and early-onset cancers, suggesting that the checkpoint may prevent genome instability associated with DNA repair (Tauchi et al., 2002).
The Roles of Rqh1 and Recombination in Regulating Replication Slowing
We propose that the Rqh1 helicase promotes replication slowing in response to DNA damage by negatively regulating recombination. In the absence of Rqh1-dependent inhibition, recombination could facilitate quick replication of a damaged template by template switching (Branzei and Foiani, 2007). Recombination could also catalyze polymerase bypass of DNA damage without strand exchange occurring. Recently, the bacterial recombinase RecA has been proposed to allow bypass of DNA damage by repriming on the leading strand after leading and lagging strand polymerases become uncoupled (Heller and Marians, 2006; McInerney and O'Donnell, 2007).
The quick replication through DNA damage also requires Swi1, a member of the Fork Protection Complex, and Swi2, a Rhp51 mediator, both of which are involved in Rhp51-mediated mating-type switching, but not in general Rhp51-mediated recombinational repair (Akamatsu et al., 2003; Noguchi et al., 2004). These results suggest that the role of Rhp51 in the checkpoint is more closely affiliated with its mating-type switching function, which may require fork stabilization, than its general recombination function.
The Rhp51-Mediator Sfr1 and Its Relationship with the Swi2–Swi5 Complex
Disruption of sfr1 does not have the same consequences for replication slowing as deletion of the other recombinase mediators, therefore Sfr1 must serve a unique function, presumably as a heterodimer with Swi5 because it has no characterized role independent of Swi5. Our genetic results suggest that Sfr1 acts to negatively regulate recombination. It is possible that Sfr1–Swi5 can both positively and negatively regulate recombination and that its negative role functions in the checkpoint. Alternatively, Sfr1 may play no direct role in the checkpoint. Instead, it may indirectly inhibit Swi2–Swi5 by simply sequestering Swi5, thereby preventing Swi5–Swi2 complex formation. This model assumes that the Swi2-dependent, mating-type switching function of Swi5 is involved in replication slowing, not the Sfr1-dependent, recombination-repair function (Akamatsu et al., 2003, 2007). This mechanism would provide a simple explanation why only a single Rhp51 mediator, Sfr1, would be required for slowing, whereas others are not.
Mechanisms for Fast Replication through Damaged DNA in the Absence of Recombination
The mechanisms of recombination-dependent DNA damage bypass discussed above do not explain the quick replication displayed by the rhp51 cds1 or rhp51 mus81 double mutants. In these cells, DNA lesions blocking fork progression must be dealt with in an Rhp51-independent manner. One possibility might be replication by translesion synthesis polymerases. However, we see no evidence for involvement of translesion synthesis in the quick replication of cds1 cells. A quadruple mutant lacking Cds1 and the three defined translesion synthesis (TLS) polymerases 
, 
, and 
(Kai and Wang, 2003) still fails to slow in response to MMS treatment, indicating that TLS activity is not required for fast replication in the absence of the checkpoint (Willis, unpublished data). Alternatively, fork collapse and reassembly, termed break-induced replication, could account for quick replisome movement along damaged template and bypass of DNA damage. Break-induced replication can occur in both an Rhp51-dependent and independent manner (Signon et al., 2001; Davis and Symington, 2004; Cortes-Ledesma et al., 2007) and cds1 mutants display increased fork instability upon replication arrest (Noguchi et al., 2003). However, fork stabilization per se, dependent on the Fork Protection Complex protein Swi1, is not required for slowing (Figure 5A). Finally, Rhp51-independent leading-strand repriming, in the absence of fork collapse, could allow rhp51 cds1 and rhp51 mus81 cells to replicate damaged DNA quickly.
Comparison of S-Phase DNA Damage Checkpoints in Fission Yeast, Budding Yeast, and Vertebrates
DNA damage induced slowing of replication is a widely conserved checkpoint response. However, the details of the response differ between fission yeast, budding yeast, and metazoa. The checkpoint in vertebrates seems to involve two parallel mechanisms: one mechanism regulating origin firing and one mechanism involving fork slowing and recombination. Contribution of these two mechanisms to replication slowing seems to differ with differing to damaging agents and doses (Falck et al., 2002; Merrick et al., 2004). In response to cis-platin and UV, the response seems to be largely fork dependent and to require recombination proteins (Henry-Mowatt et al., 2003). In addition, both Mus81 and BLM, a mammalian Rqh1 homolog, are required for the regulation of replication forks in response to aphidicolin-induced replication stress (Shimura et al., 2008). In contrast, low doses of IR trigger an origin-based response (Merrick et al., 2004), and higher doses of IR may affect fork progression in addition to origin firing (Falck et al., 2002). In budding yeast, replication is slowed in response to different DNA damaging agents, including MMS and bleomycin (Andrews and Clarke, 2005). Both reduction in origin firing and checkpoint independent fork slowing contribute to slowing in response to MMS (Tercero and Diffley, 2001). In contrast to budding yeast and metazoa, fission yeast slows replication only in response to MMS and not the double-strand break producing agents IR and bleomycin. We attribute this slowing to direct regulation of replication fork progression. The fact that fission yeast seems to regulate primarily replication forks makes it an excellent model organism for the exploration of replication fork regulation by the S-phase DNA damage checkpoint.
In conclusion, our data show that regulation of recombination plays an important downstream role in the S phase DNA damage checkpoint. Involvement of helicases, endonucleases, recombinases, and recombinase mediators in the S phase DNA damage checkpoint implicates recombination as an important factor in regulating the slowing of replication in response to DNA damage and further strengthens the notion that the replication slowing we observe is primarily produced by a coordinated response at replication forks. Slowing is likely due to a rapid and local response at replication forks encountering damage through the regulation of recombinational exchange between replicating sister chromatids.
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ACKNOWLEDGMENTS |
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Footnotes |
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Address correspondence to: Nicholas Rhind (nick.rhind{at}umassmed.edu)
Abbreviations used: IR, ionizing radiation; MMS, methyl methane sulfonate.
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