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Vol. 18, Issue 10, 4062-4073, October 2007

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Shu Proteins Promote the Formation of Homologous Recombination Intermediates That Are Processed by Sgs1-Rmi1-Top3

Cancer Research UK Laboratories, Weatherall Institute of Molecular Medicine, University of Oxford, John Radcliffe Hospital, Oxford, OX3 9DS, United Kingdom

Submitted May 24, 2007; Revised July 18, 2007; Accepted July 20, 2007
Monitoring Editor: Wendy Bickmore

	ABSTRACT

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
CSM2, PSY3, SHU1, and SHU2 (collectively referred to as the SHU genes) were identified in Saccharomyces cerevisiae as four genes in the same epistasis group that suppress various sgs1 and top3 mutant phenotypes when mutated. Although the SHU genes have been implicated in homologous recombination repair (HRR), their precise role(s) within this pathway remains poorly understood. Here, we have identified a specific role for the Shu proteins in a Rad51/Rad54-dependent HRR pathway(s) to repair MMS-induced lesions during S-phase. We show that, although mutation of RAD51 or RAD54 prevented the formation of MMS-induced HRR intermediates (X-molecules) arising during replication in sgs1 cells, mutation of SHU genes attenuated the level of these structures. Similar findings were also observed in shu1 cells in which Rmi1 or Top3 function was impaired. We propose a model in which the Shu proteins act in HRR to promote the formation of HRR intermediates that are processed by the Sgs1-Rmi1-Top3 complex.

	INTRODUCTION

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
Homologous recombination repair (HRR) is a well-conserved cellular process for the repair of single-strand DNA (ssDNA) gaps and double-strand DNA breaks (DSBs) that can arise during DNA synthesis or as a result of replication fork stalling during S-phase. Although many of the key proteins involved in HRR have been identified (reviewed in Paques and Haber, 1999; Sung et al., 2003; West, 2003; Krogh and Symington, 2004), in some cases their precise function(s) remains to be identified. Moreover, the mechanisms for suppression of inappropriate HRR in S-phase are only poorly defined. It is likely that HRR must be carefully regulated and/or executed in dividing cells, as inappropriate or excessive HR can lead to genome rearrangements and cancer in mammals. This is exemplified by the cancer-predisposition disorder, Bloom's syndrome, which is caused by mutation in the human BLM gene (reviewed in German, 1993). Because the BLM protein, in conjunction with its associated proteins, hTOPOIII and hRMI1 (Johnson et al., 2000; Wu et al., 2000; Yin et al., 2005), can catalyze dissolution of HRR intermediates in vitro (Wu and Hickson, 2003; Plank et al., 2006; Raynard et al., 2006; Wu et al., 2006), it is likely that unprocessed and/or aberrantly processed HRR intermediates at least partly contribute to the cellular defects in Bloom's syndrome. Indeed, Bloom's syndrome cells classically demonstrate elevated levels of sister chromatid exchanges, mitotic recombination, and genome instability. Mutation of the BLM, hTOPOIII, or hRMI1 homologues in Saccharomyces cerevisiae (SGS1, TOP3, or RMI1, respectively) similarly causes sensitivity to genotoxic agents, hyper-recombination, and synthetic lethality with mutations in other genes also implicated in HRR (e.g., MUS81 and SRS2; Gangloff et al., 1994; Watt et al., 1996; Gangloff et al., 2000; Mullen et al., 2001; Fabre et al., 2002; Chang et al., 2005; Mullen et al., 2005). Furthermore, unresolved HRR intermediates have been directly visualized by two-dimensional (2D) gel electrophoresis in cells lacking Sgs1 or in cells with impaired Top3 function (Liberi et al., 2005; Mankouri and Hickson, 2006). Interestingly, many of the cellular/phenotypic defects observed in sgs1, top3, or rmi1 cells can be suppressed by deletion of genes that control the early steps of HRR (e.g., RAD52, RAD51, RAD55, RAD57, and RAD54; Gangloff et al., 2000; Fabre et al., 2002; Oakley et al., 2002; Shor et al., 2002; Chang et al., 2005; Mullen et al., 2005). Taken together, these observations suggest that excessive, unscheduled, or incomplete HRR can create toxic DNA repair intermediates, and highlights the requirement for cells to carefully regulate HRR during S-phase.

CSM2, PSY3, SHU1, and SHU2 (collectively referred to as the SHU genes) were identified recently as four novel genes within the same epistasis group that, when mutated, can suppress various defects in sgs1 or top3 mutants (Shor et al., 2005). Interestingly, the single csm2, psy3, shu1, or shu2 mutants all demonstrate similar phenotypes (a mutator phenotype and moderate sensitivity to methyl methane sulfonate (MMS) and cross-linking agents), and mutation of all four does not cause any additive effects (Shor et al., 2005). Taken together with the demonstration that all four SHU gene products interact in a two-hybrid assay, it has been proposed that these proteins exist in a multimeric complex that fails to function when any one member is missing (Shor et al., 2005). It is likely that the Shu complex is involved in some aspect of HRR, because mutation of RAD52, which abolishes all types of HRR (Symington, 2002), is epistatic to shu mutations for MMS sensitivity and a mutator phenotype (Shor et al., 2005). However, it is unlikely that the Shu proteins are bona fide members of the core HRR machinery. Unlike canonical HRR mutants (e.g., rad52, rad51, rad54, rad55, and rad57), the shu mutants are not appreciably sensitive to hydroxyurea (HU) or gamma-rays (Shor et al., 2005). Therefore, unlike classic HRR proteins, Shu proteins are apparently not essential for the repair of DSBs or collapsed replication forks. Nevertheless, mutation of the SHU genes does affect some aspect of HRR, because DNA damage-induced Rad52 foci persist for longer in the nuclei of shu1 cells exposed to MMS as compared with wild-type cells (Shor et al., 2005).

Further investigation of the SHU genes in S. cerevisiae is likely to be relevant to all eukaryotes, because putative PSY3 and SHU2 homologues have recently been identified in Schizosaccharomyces pombe (rld1⁺ and sws1⁺, respectively) and human cells (RAD51D and SWS1, respectively; Martin et al., 2006). RAD51D is one of the so-called human RAD51 paralogs, based on some shared sequence similarity to RAD51 (Cartwright et al., 1998a; Kawabata and Saeki, 1998; Pittman et al., 1998). Interestingly, like psy3 and shu2 mutations in S. cerevisiae, mutation of rld1⁺ or sws1⁺ in S. pombe also causes sensitivity to MMS (but not to other types of DNA damage) and rescues various cellular defects caused by mutation of the RecQ helicase gene, rqh1⁺, in S. pombe (Martin et al., 2006). Furthermore, Sws1 associates in vivo with Rld1 and a novel protein, Rlp1, which shows sequence similarity to another of the human RAD51 paralogs, XRCC2 (Cartwright et al., 1998b; Liu et al., 1998; Martin et al., 2006). Therefore, Sws1, Rld1, and Rlp1 appear to be part of a multimeric complex in S. pombe, which is similar to the complex proposed to exist in S. cerevisiae comprising Csm2, Psy3, Shu1, and Shu2 (Shor et al., 2005). Interestingly, SWS1 and XRCC2 associate with RAD51D in human cells (Braybrooke et al., 2000; Martin et al., 2006), and RAD51D can bind to BLM (Braybrooke et al., 2003), suggesting that Shu-like complexes also exist in human cells and therefore probably perform an evolutionarily conserved role. Ablation of SWS1 in human cells reduces the number of RAD51 foci in both control and IR-treated human cells (Martin et al., 2006), suggesting that SWS1, like its yeast counterparts, is involved in some aspect of HRR.

It is likely that a modulation of HRR is responsible for the suppression of sgs1/top3 phenotypes by shu mutations (S. cerevisiae) or rqh1 phenotypes by rld1/rlp1/sws1 mutations (S. pombe). In S. cerevisiae, mutation of SHU1 suppresses the increased rate of recombination and elevated (spontaneous) Rad52 foci in sgs1 and top3 cells (Shor et al., 2005). A similar scenario exists in S. pombe, where sws1 and rlp1 mutations reduce the increased recombination rate and elevated number of nuclei containing spontaneous Rad22 foci in rqh1 mutants, without apparently affecting the outcome of HRR (Martin et al., 2006). Taken together, these data are consistent with the Shu complex in S. cerevisiae and the Sws1, Rld1, and Rlp1 complex in S. pombe, somehow modulating HRR in cells lacking RecQ helicases or Topoisomerase III. However, the molecular mechanisms underlying this suppression remain to be clarified.

We sought to identify the mechanism by which the shu mutations suppress sgs1 and top3 defects in S. cerevisiae. We demonstrate that, in addition to suppressing sgs1 or top3 phenotypes (Shor et al., 2005), shu1 mutation also suppresses the poor growth caused by the deletion of RMI1 or the overexpression of a dominant-negative allele of TOP3 (TOP3^Y356F). Consistent with a role for the Shu proteins in HRR (Shor et al., 2005), we demonstrate that Shu1 acts in the Rad51/Rad54-dependent HRR repair of MMS lesions, upstream of the Sgs1-Rmi1-Top3 complex. Interestingly, we demonstrate that, unlike mutation of RAD51 (or RAD54), mutation of SHU genes does not prevent unresolved HRR intermediates from persisting in cells compromised for Sgs1, Rmi1, or Top3. However, the level of unresolved HRR intermediates was attenuated to some extent by shu mutations. We propose that Shu proteins perform a nonessential role in HRR to promote the formation of HRR intermediates that are substrates for Sgs1-Rmi1-Top3.

	MATERIALS AND METHODS

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
S. cerevisiae Strains and Plasmids
All strains were isogenic derivatives of T344 or BY4741. All strains carrying gene deletions were either obtained from the yeast deletion collection (EUROSCARF, University of Frankfurt, Germany) or constructed using a PCR-based gene disruption method (Wach et al., 1994). The rmi1 haploid strain was obtained by sporulating a heterozygous RMI1^+/– diploid strain. All haploid rmi1 strains used for our experimental analyses were confirmed to demonstrate a slow-growth phenotype (Chang et al., 2005; Mullen et al., 2005). The pYES2-TOP3 and pYES2-TOP3^Y356F plasmids have been described previously (Oakley et al., 2002; Mankouri and Hickson, 2006).

Growth Conditions, Cell Synchronization, and Flow Cytometry Analysis
Strains were grown and synchronized with -factor mating pheromone as described previously (Mankouri and Hickson, 2006). After release from -factor arrest, all experiments were performed at 25°C. Release from MMS treatment was achieved by centrifugation, washing, and resuspension of cells in drug-free medium. Cell cycle progression was monitored using flow cytometry (fluorescent-activated cell-sorting [FACS]) as described previously (Mankouri and Hickson, 2006).

2D Gel Electrophoresis
The hexadecyltrimethylammonium bromide (CTAB) method of DNA extraction and 2D gel procedures were described previously (Brewer and Fangman, 1987; Allers and Lichten, 2000; Lopes et al., 2003; Liberi et al., 2006). DNA was digested with NciI and NcoI before running the first-dimension gels. Quantification of X-shaped molecules was performed using Image Quant analysis software (Molecular Dynamics, Sunnyvale, CA). "Object Average" mode of background correction was used and then each X-molecule was normalized with respect to the monomer (1N) spot, to provide an arbitrary value of X-molecule intensity.

	RESULTS

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
SHU Mutations Are Epistatic with rad51 and rad54 for Suppression of TOP3^Y356F-induced Poor Growth
We demonstrated previously that overexpression of a catalytically dead mutant of TOP3, TOP3^Y356F, causes a dominant-negative top3-like phenotype (Oakley et al., 2002; Mankouri and Hickson, 2006). Because the SHU genes (CSM2, PSY3, SHU1, and SHU2) were initially identified as suppressors of top3 (Shor et al., 2005), we investigated if SHU1 mutation could also suppress the poor growth caused by overexpression of TOP3^Y356F in the T344 strain (Hovland et al., 1989). Consistent with the demonstration that shu mutations suppress top3 poor growth (Shor et al., 2005), we found that mutation of SHU1 also partially suppressed the poor growth caused by overexpression of TOP3^Y356F (Figure 1A). Interestingly, we noted that the partial suppression of TOP3^Y356F-induced poor growth was reminiscent of that previously caused by mutation of the HRR gene, RAD51 (Mankouri and Hickson, 2006). We therefore asked if a combination of shu1 and rad51 mutations caused additive suppression of TOP3^Y356F-induced poor growth. For comparison, we also compared the effects of mutating RAD54, which functions in RAD51-dependent HRR (Rattray and Symington, 1995; Zhang et al., 2007) and also partially suppresses top3 poor growth when mutated (Oakley et al., 2002; Shor et al., 2002). Interestingly, we observed that rad51, rad54 and shu1 mutations all demonstrated a similar partial ability to suppress TOP3^Y356F-induced poor growth (Figure 1A). Furthermore, this suppression was not additive in rad51shu1 or rad54shu1 double mutants. Therefore, we propose that SHU1, RAD51, and RAD54 are epistatic for suppression of TOP3^Y356F-induced poor growth. These data are consistent with a proposed role for the Shu proteins in RAD52-dependent HRR (Shor et al., 2005). However, because mutation of RAD52 abolishes all types of HRR repair (Symington, 2002), some of which are Rad51-independent, our data implies that, more specifically, Shu1 is involved in Rad51- and Rad54-dependent branch of HRR.

View larger version (98K): [in this window] [in a new window]   Figure 1. SHU1, RAD51, and RAD54 are epistatic for suppression of TOP3Y356F-induced poor growth and MMS sensitivity. (A) Wild-type, shu1, rad51, rad51shu1, rad54, and rad54shu1 strains transformed with pYES2-TOP3 or pYES2-TOP3Y356F were diluted to equivalent densities and spotted onto control plates or plates containing 2% galactose (to induce expression from the pYES2 GAL1 promoter). Spots from left to right represent serial 1 in 10 dilutions of yeast cultures. Plates were grown at 30°C. The large single colonies that arise in strains overexpressing TOP3Y356F represent suppressors (probably SGS1-linked) as seen previously (Oakley et al., 2002; Mankouri and Hickson, 2006). (B) Wild-type, shu1, rad51, rad51shu1, rad54, and rad54shu1 strains were diluted to equivalent densities and spotted onto control plates (no drug) or plates containing MMS. Spots from left to right represent serial 1 in 10 dilutions of yeast cultures. Plates were grown at 30°C.

Our previous data demonstrated that the mutation of RAD51 causes an impaired ability to traverse S-phase in the presence of MMS (Mankouri and Hickson, 2006). This phenotype is a consequence of a persistent activation of the DNA damage checkpoint, because this phenotype can be overridden by addition of caffeine (Mankouri and Hickson, 2006). Interestingly, mutation of RAD52 also impairs S-phase progression in the presence of MMS (Oakley et al., 2002), and substitution of phosphorylation sites targeted by checkpoint kinases in Rad55 causes defects in cell cycle resumption after MMS treatment (Herzberg et al., 2006). Taken together, these findings suggest that impaired S-phase progression in the presence of MMS may be a general property of HRR-defective mutants. To explore this possibility further, we examined if mutation of RAD54 or SHU1 affects S-phase progression in the presence of MMS. The strains used in Figure 1B were synchronously released from G1 arrest into fresh medium and DNA content was analyzed at regular intervals using flow cytometry (FACS). Under these unperturbed conditions, all strains progressed through S-phase (as measured by a doubling of DNA content) by 60 min (Figure 2A), with only a very modest increase in S-phase duration observed in rad51shu1 and rad54shu1 double mutants (note DNA content at 40 min). In the presence of 0.0167% MMS, wild-type cells successfully traversed S-phase by 2 h (Figure 2B). This prolongation of S-phase by MMS (relative to unperturbed cells) is a consequence of MMS-induced replication fork stalling and subsequent checkpoint activation (Paulovich and Hartwell, 1995; Santocanale and Diffley, 1998; Shirahige et al., 1998; Tercero and Diffley, 2001). In agreement with our previous data (Mankouri and Hickson, 2006), rad51 cells failed to completely traverse S-phase by 4 h in the presence of 0.0167% MMS, demonstrating a mid-S-phase DNA content at 4 h (Figure 2B). Interestingly, we observed that rad54 and shu1 mutants resembled rad51 mutants and also failed to traverse S-phase by 4 h in 0.0167% MMS (Figure 2B). Furthermore, like rad51 cells (Mankouri and Hickson, 2006), the impaired S-phase progression phenotype of MMS-treated rad54 or shu1 mutants could be overridden by addition of caffeine (Figure 2C). Therefore, these data imply that a more persistent or robust checkpoint activation occurs after MMS treatment in cells lacking Rad51, Rad54, or Shu1. Similar results were also observed for the other shu mutant strains (data not shown), consistent with the similar phenotypes caused by mutation of any one of the SHU genes (Figure 3A; Shor et al., 2005). Interestingly, rad51shu1 or rad54shu1 double mutants did not demonstrate additive impaired S-phase progression phenotypes in the presence of MMS, consistent with the proposed epistasis between these genes (Figure 1). Furthermore, the impaired S-phase progression in MMS-treated rad51shu1 or rad54shu1 double mutants could be overridden by caffeine (Figure 2C). We propose that Rad51, Rad54, and Shu1 all act at a similar step in a common pathway for the repair of MMS-induced DNA lesions during S-phase and that perturbation/inactivation of this pathway leads to accumulation of an HRR substrate that causes persistent checkpoint activation.

View larger version (36K): [in this window] [in a new window]   Figure 2. Shu1 functions in a Rad51/Rad54-dependent pathway to repair/tolerate MMS-induced lesions during S-phase. (A and B) Wild-type, shu1, rad51, rad51shu1, rad54, and rad54shu1 strains were released from G1 arrest into fresh medium (A) or medium containing 0.0167% MMS (B). DNA content was analyzed by flow cytometry at the indicated times. The shaded peaks represent experimental data, whereas the unshaded peak is a reference to indicate a normal G2/M peak (at 2 h release from G1 arrest). The positions of the 1n (G1) and 2n (G2/M) peaks are indicated below. (C) After 3 h treatment with 0.0167% MMS, 1-ml aliquots of cells were taken, and caffeine was added to a final concentration of 5 mg/ml, or an equivalent amount of fresh medium was added. Samples were taken 1 h later for analysis of DNA content by flow cytometry.

View larger version (58K): [in this window] [in a new window]   Figure 3. shu1 mutation suppresses sgs1 HU sensitivity and is epistatic to sgs1 for MMS sensitivity. (A) Wild-type, sgs1, csm2, csm2sgs1, psy3, psy3sgs1, shu1, shu1sgs1, shu2, and shu2sgs1 strains were diluted to equivalent densities and spotted onto control (no drug) plates or plates containing indicated doses of MMS or HU. Spots from left to right represent serial 1 in 10 dilutions of yeast cultures. Plates were grown at 30°C. (B) Wild-type, shu1, sgs1, and shu1sgs1 strains were released from G1 arrest into fresh medium containing 0.0167% MMS. DNA content was analyzed by flow cytometry at the indicated times, as in Figure 2. (C) Wild-type, sgs1, shu1, shu1sgs1, rad51, rad51sgs1, rad51shu1, rad51shu1sgs1, rad54, rad54sgs1, rad54shu1, and rad54shusgs1 strains were diluted to equivalent densities and spotted onto control (no drug) plates or plates containing MMS. Spots from left to right represent serial 1 in 10 dilutions of yeast cultures. Plates were grown at 30°C.

To further investigate this apparent discrepancy, and to test for any strain-dependent variations, we investigated the genetic relationship between SHU1 and SGS1 in a third strain background, BY4741. In this background, mutation of SHU1 again suppressed sgs1 HU sensitivity (data not shown). However, in agreement with a recent independent study (St. Onge et al., 2007), we observed that mutation of SHU1 suppressed sgs1 MMS sensitivity in the BY4741 genetic background (data not shown). We propose that this apparent inconsistency between our T344 and BY4741 shu1sgs1 strains can be reconciled by the fact that shu mutants are more sensitive to MMS than are sgs1 strains in the T344 background, whereas sgs1 mutants are more sensitive to MMS than are shu mutants in the BY4741 background. In both of the strain backgrounds that we analyzed, the phenotype of the shu sgs1 double mutant mimics the phenotype of the corresponding shu single mutant. Therefore, the additive sensitivity of shu1sgs1 double mutants to MMS as reported by Shor et al. (2005) likely reflects a strain-specific effect. Taken together, our data implies that, despite apparent strain-dependent differences in MMS sensitivity, shu mutations are epistatic to sgs1 for MMS sensitivity. Moreover, our data are consistent with the Shu proteins acting upstream of Sgs1.

To further examine the relationship between SHU1 and SGS1 in the repair of MMS-induced DNA lesions, we examined S-phase progression in wild-type, sgs1, shu1 and shu1sgs1 cells in the presence of 0.0167% MMS. We observed that all strains demonstrated similar S-phase kinetics in unperturbed cells (data not shown). In the presence of 0.0167% MMS, however, wild-type and sgs1 cells completed DNA replication by 3 h, whereas shu1 cells failed to complete S-phase by 4 h (Figure 3B). Therefore, in agreement with previous data (Mankouri and Hickson, 2006), sgs1 cells do not demonstrate impaired S-phase progression in the presence of 0.0167% MMS. Consistent with the genetic relationship observed between SHU1 and SGS1 (Figure 3A), we observed that shu1sgs1 cells resembled shu1 cells in failing to complete DNA replication by 4 h in 0.0167% MMS (Figure 3B). Similar results were also observed in the BY4741 strain background (data not shown), consistent with our proposal that shu1 is epistatic to sgs1 for MMS sensitivity in both the T344 and BY4741 strain backgrounds. These findings were not limited to shu1, because csm2, csm2sgs1, psy3, psy3sgs1, shu2, and shu2sgs1 mutants all demonstrated a similar degree of impaired S-phase progression in the presence of MMS to that of shu1 or shu1sgs1 mutants (data not shown). Furthermore, in each shu single mutant or shu sgs1 double mutant, the impaired S-phase progression in the presence of MMS could be overridden by addition of caffeine (Figure 2C and data not shown). Therefore, loss of any one of the Shu proteins causes persistent checkpoint activation after treatment with MMS, regardless of whether Sgs1 is present or not. Taken together with the fact that shu mutations are epistatic to sgs1 for MMS sensitivity (Figure 3A), these findings are consistent with a role for the Shu complex upstream of Sgs1 in the same pathway to repair MMS-induced DNA lesions during S-phase.

To further investigate the relationship between Shu1, Sgs1, and the HRR pathway, we examined the effects of deleting RAD51 and RAD54. Interestingly, we observed that, similar to what was observed when SHU1 was mutated (Figure 3B), mutation of RAD51 or RAD54 impaired the S-phase progression of sgs1 mutants (Supplementary Figure 1). Consistent with our proposal that the impaired S-phase progression phenotype of a shu1 mutant is not additive with rad51 or rad54 mutations, we observed that rad51shu1sgs1 and rad54shu1sgs1 triple mutants demonstrated rates of (impaired) S-phase progression in the presence of MMS similar to that of rad51sgs1, rad54sgs1, or shu1sgs1 double mutants (Supplementary Figure 1). Furthermore, rad51shu1sgs1 and rad54shu1sgs1 triple mutants were no more sensitive to MMS than rad51 or rad54 single mutants, respectively (Figure 3C). This apparent lack of additive phenotypes is consistent with the proposal that Rad51/Rad54, Shu1, and Sgs1 are all epsitatic for MMS sensitivity. We conclude that Rad51, Rad54, Shu1, and Sgs1 all function in the same (HRR) pathway to repair MMS lesions during S-phase and that Sgs1 acts at a later stage than Rad51, Rad54, or Shu1.

Formation of MMS-induced X-Molecules in Cells Lacking SGS1 Is Abolished by Mutation of RAD51 or RAD54, and Is Attenuated by Mutation of SHU Genes
Previous studies indicated that sgs1 cells exhibit persistent, Rad51-dependent, HRR intermediates on 2D gels after exposure to MMS (Liberi et al., 2005). On the basis of our proposal that the Shu proteins act upstream of Sgs1 in the same HRR pathway to repair MMS-induced lesions, we examined whether mutation of the SHU genes prevents the accumulation of HRR intermediates in MMS-treated sgs1 cells. To confirm that sgs1 mutants demonstrate Rad51-dependent X-shaped molecules after exposure to MMS in the T344 strain background, we analyzed DNA replication intermediates on 2D gels originating from an early firing origin, ARS305, in wild-type, sgs1, and rad51sgs1 strains. Additionally, we also analyzed whether persistent X-shaped molecules were present in MMS-treated shu1 or rad54sgs1 cells. Strains were released from G1 arrest into fresh medium containing 0.033% MMS, and cells were harvested at early (30 min) or late (180 min) time points. Genomic DNA was prepared using the CTAB method of DNA extraction to restrain branch migration of joint (X-shaped) molecules, as described previously (Lopes et al., 2003). We observed that origin firing at ARS305 was detectable after 30 min in all strains, as evidenced by the appearance of bubbles, Y-molecules and the so-called origin-associated X-structures (Figure 4A). Previous studies have indicated that the origin-associated X-structures are normal DNA replication intermediates that are not dependent on Rad51 or Rad52 for their formation and are not, therefore, HRR intermediates (Lopes et al., 2003). In each strain, the relative ratio between bubbles, Y-molecules, and X-structures was similar at 30 min, suggesting that origin firing occurs normally in shu1, sgs1, rad51sgs1, and rad54sgs1 mutants. After 180 min, all of the ARS305 replication intermediates detectable at 30 min were substantially diminished, or had disappeared, in wild-type cells, consistent with replication fork progression beyond the boundaries of this genomic region by this time. Similar results were observed in shu1 cells, suggesting that mutation of SHU1 does not noticeably affect DNA replication at ARS305 (Figure 4A). In contrast, we observed that, although bubbles and Y-molecules had largely disappeared from the ARS305 region at 180 min, abnormal DNA replication intermediates (X-molecules) persisted in MMS-treated sgs1 cells (Figure 4A). In agreement with previous data (Liberi et al., 2005), we also observed that these X-molecules were not detectable in rad51sgs1 cells (Figure 4A). Interestingly, we also observed a similar effect when RAD54 was mutated, because rad54sgs1 mutants also did not exhibit persistent MMS-induced X-molecules (Figure 4A). Therefore, we conclude that persistent X-molecules in MMS-treated sgs1 cells are both Rad51- and Rad54-dependent. This finding is consistent with Rad54 acting at an early stage in Rad51-dependent HRR upstream of Sgs1 (Rattray and Symington, 1995; Petukhova et al., 1998; Zhang et al., 2007).

View larger version (49K): [in this window] [in a new window]   Figure 4. Mutation of SHU genes attenuate persistent X-molecules in MMS-treated sgs1 cells. (A) Wild-type, shu1, sgs1, rad51sgs1, and rad54sgs1 strains were released from G1 arrest into fresh medium containing 0.033% MMS. DNA replication intermediates were analyzed by 2D gel electrophoresis at the times indicated. DNA samples were analyzed with a probe for the early-firing ARS305 replication origin. The diagrammatic representation on the top right of the panel denotes DNA structures that can be identified by the 2D gel technique. For quantification of replication intermediates (middle and bottom panels on the right), each signal (bubbles, Ys, and Xs) was normalized to its corresponding monomer (1N) spot. The arrowhead indicates the position of MMS-induced persistent X-molecules. (B) DNA replication intermediates at ARS305 were analyzed in sgs1, csm1sgs1, psy3sgs1, shu1sgs1, and shu2sgs1 mutants after 3-h treatment with 0.033% MMS after release from G1 arrest. For quantification of X-molecules (right), each X-molecule signal was normalized to its corresponding monomer (1N) spot. (C) A time course of DNA replication at ARS305 is shown for sgs1 and shu1sgs1 mutants after release from G1 arrest into fresh medium containing 0.033% MMS. (D) DNA replication intermediates at ARS305 were analyzed in sgs1, shu1sgs1, rad51shu1sgs1, and rad54shu1sgs1 mutants after 3-h treatment with 0.033% MMS after release from G1 arrest. The filled arrowhead indicates the position of MMS-induced persistent X-molecules, whereas the white arrowhead indicates the position of attenuated X-molecules.

To determine if the low level of MMS-induced X-molecules in shu1sgs1 cells were nevertheless HRR intermediates, we investigated the effects of deleting RAD51 or RAD54 in shu1sgs1 cells. We found that origin firing at ARS305 and S-phase progression were indistinguishable in MMS-treated shu1sgs1, rad51shu1sgs1, and rad54shu1sgs1 strains (data not shown and Supplementary Figure 1). After 180 min of MMS treatment, we again observed that persistent X-molecules were detectable in MMS-treated sgs1 cells and that these were attenuated in shu1sgs1 cells (Figure 4D). However, we found that the MMS-induced X-molecules were absent from rad51shu1sgs1 or rad54shu1sgs1 mutants, indicating that the attenuated X-molecules in shu1sgs1 cells are RAD51- and RAD54-dependent HRR structures. (Figure 4D). Taken together, the data presented in Figure 4 indicate that, whereas acute formation of MMS-induced persistent X-molecules in sgs1 cells is abolished by mutation of RAD51 or RAD54, these structures are not completely abolished by mutation of SHU genes. However, because the level of MMS-induced X-molecules is attenuated in shu sgs1 double mutants, we propose that mutation of the SHU genes affects either the rate of formation/removal, or the nature, of the X-molecules in sgs1 cells.

Mutation of shu1 Attenuates Persistent MMS-induced X-Molecules in Cells Overexpressing TOP3^Y356F
We demonstrated previously that overexpression of TOP3^Y356F causes Rad51-dependent X-molecules to persist during S-phase after exposure to MMS. To determine if the attenuation of sgs1 MMS-induced X-molecules by shu mutations was specific for sgs1 X-molecules, or general for both sgs1 and TOP3^Y356F-induced X-molecules, we examined if mutation of SHU1 affects MMS-induced X-molecule persistence in cells overexpressing TOP3^Y356F. For this, wild-type and shu1 strains overexpressing TOP3^Y356F were released from G1 arrest into medium containing 0.033% MMS. In agreement with our previous data (Mankouri and Hickson, 2006), wild-type cells overexpressing TOP3^Y356F demonstrated persistent MMS-induced X-molecules at 180 min after G1 release (Figure 5, A and B). These structures were absent in MMS-treated rad51 or rad54 cells overexpressing TOP3^Y356F, verifying that Rad51 and Rad54 are required for their formation (data not shown; Mankouri and Hickson, 2006). Persistent MMS-induced X-molecules were detected in shu1 cells overexpressing TOP3^Y356F, although their level was attenuated relative to those observed in wild-type cells overexpressing TOP3^Y356F (Figure 5, A and B). Furthermore, no MMS-induced X-molecules were detectable in rad51shu1 TOP3^Y356F strains, suggesting that these X-molecules are also Rad51-dependent HRR intermediates (Figure 5B). Because these results are qualitatively very similar to those observed in shu1sgs1 cells (Figure 4, B and C), we propose that mutation of SHU1 attenuates MMS-induced X-molecule formation when Top3 function is impaired by a similar mechanism to that occurring in shu1sgs1 cells.

View larger version (82K): [in this window] [in a new window]   Figure 5. Mutation of SHU1 attenuates Rad51-dependent, MMS-induced X-molecules in rmi1 cells and in cells overexpressing TOP3Y356F. (A) Wild-type and shu1 strains transformed with pYES2-TOP3Y356F were arrested in G1 with -factor and simultaneously treated with 2% galactose (to induce overexpression from the pYES2 GAL1 promoter). Cultures were released into fresh medium containing 0.033% MMS, and DNA replication intermediates were analyzed by 2D gel electrophoresis at the times indicated. DNA samples were analyzed with a probe for the early-firing ARS305 replication origin. The filled arrowhead indicates the position of MMS-induced persistent X-molecules, whereas the white arrowhead indicates the position of attenuated X-molecules. (B) DNA replication intermediates at ARS305 were analyzed in wild-type, shu1, and rad51shu1 cells overexpressing TOP3Y356F after 3-h treatment with 0.033% MMS after release from G1 arrest. (C) Wild-type, shu1, rad51, rmi1, rad51rmi1, and shu1rmi1 strains were diluted to equivalent densities and spotted onto control plates (no drug) or plates containing MMS. Spots from left to right represent serial 1 in 10 dilutions of yeast cultures. Plates were grown at 30°C. (D) Wild-type, rmi1, rad51rmi1, and shu1rmi1 strains were released from G1 arrest into fresh medium containing 0.033% MMS. DNA replication intermediates were analyzed by 2D gel electrophoresis at the times indicated. DNA samples were analyzed with a probe for the early-firing ARS305 replication origin.

On the basis of its interactions with Sgs1 and Top3 (Chang et al., 2005; Mullen et al., 2005), we examined if rmi1 mutants also demonstrate persistent X-molecules in the presence of MMS. We observed that, similar to cells lacking SGS1, or overexpressing TOP3^Y356F (Figures 4 and 5; Liberi et al., 2005; Mankouri and Hickson, 2006), rmi1 cells demonstrated persistent X-molecules at ARS305 after exposure to MMS (Figure 5D). We also observed that, although mutation of RAD51 eliminated the MMS-induced X-molecules at ARS305 in rmi1 cells, mutation of SHU1 again only attenuated their level (Figure 5D). Therefore, the persistent X-molecules detected in rmi1 cells likely represent Rad51-dependent HRR intermediates, similar or identical to those identified in cells lacking Sgs1 or in cells overexpressing TOP3^Y356F (Figures 4 and 5; Liberi et al., 2005; Mankouri and Hickson, 2006). Furthermore, because mutation of SHU1 only attenuated these MMS-induced X-molecules, we propose that the Shu proteins normally promote the formation of Rad51/Rad54-dependent HRR intermediates that are ultimately resolved by the Sgs1-Rmi1-Top3 complex.

	DISCUSSION

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
CSM2, PSY3, SHU1, and SHU2 (collectively referred to as the SHU genes) were recently identified in S. cerevisiae as four genes in the same epistasis group that, when mutated, cause sensitivity to MMS and suppression of various sgs1 and top3 mutant phenotypes (Shor et al., 2005). RMI1 encodes a protein that associates with Sgs1 and Top3 in vivo, and rmi1 mutants phenotypically resemble top3 mutants (Chang et al., 2005; Mullen et al., 2005). This physical interaction is also evolutionarily conserved, because the human Rmi1 homologue, hRMI1, is an integral component of the BLM-hTOPOIII complex in human cells (Yin et al., 2005). In this study, we have demonstrated that mutation of SHU1 suppresses poor growth caused by overexpression of TOP3^Y356F or deletion of RMI1, indicating that the Shu complex actively contributes to the cellular defects seen in cells lacking Sgs1, Top3, or Rmi1.

Role of Shu Proteins in Rad51/Rad54-dependent Homologous Recombination Repair
The ability of shu mutations to suppress rmi1, top3, and TOP3^Y356F-induced poor growth, and to suppress the synthetic lethality of sgs1srs2, sgs1mus81, and sgs1mms4 mutant combinations is very reminiscent of that achieved by deletion of genes that control the early steps of HRR (e.g., RAD52, RAD51, RAD55, RAD57, and RAD54; Shor et al., 2005; St. Onge et al., 2007). Shor et al. (2005) demonstrated that rad52 is epsitatic to shu mutations for MMS sensitivity and a mutator phenotype, suggesting that the Shu proteins are involved in RAD52-dependent HRR. However, because mutation of RAD52 abolishes all types of HRR (Symington, 2002), the precise role(s) of the Shu complex in HRR was not clear. In this study, we identified a specific role for the Shu complex in Rad51/Rad54-dependent pathway(s) to repair MMS-induced lesions. This assertion is based on the following observations: 1) Mutation of SHU1 does not cause additive sensitivity to MMS when combined with rad51 or rad54 mutations. 2) Mutation of RAD51, RAD54, or SHU1 causes a similar, and nonadditive, impairment of S-phase progression in the presence of MMS, due to a more persistent or robust activation of the DNA damage checkpoint. 3) Mutation of SHU1 is epistatic (i.e., nonadditive) with rad51 or rad54 for suppression of TOP3^Y356F-induced poor growth. However, it should be noted that, unlike rad51 and rad54 mutants, the shu mutants do not exhibit sensitivity to HU or ionizing radiation (Shor et al., 2005). Therefore, it is unlikely that the Shu proteins are involved in HRR repair of double-strand breaks. We propose, therefore, that the Shu proteins function at an early stage in a Rad51/Rad54-dependent HRR subpathway specifically to repair certain types of DNA lesions (probably ssDNA gaps; see below) that arise during S-phase.

Suppression of sgs1, rmi1, and top3/TOP3^Y356F Phenotypes by Mutation of SHU Genes
Because mutation of SHU1 or CSM2 suppresses hyper-recombination in sgs1 and top3 cells (Shor et al., 2005), we examined if shu mutations prevent unprocessed HRR intermediates from accumulating in cells compromised for Sgs1, Top3, or Rmi1. Previous data demonstrated that unresolved Rad51-dependent HRR intermediates are detectable in MMS-treated sgs1, top3, and TOP3^Y356F cells (Liberi et al., 2005; Mankouri and Hickson, 2006). In this study, we also demonstrated that rmi1 mutants exhibit persistent, Rad51-dependent, MMS-induced X-molecules, suggesting that the accumulation of unprocessed HRR intermediates is a common property of strains mutated for SGS1, TOP3, or RMI1. It should be noted, however, that we do not know presently if the X-molecules arising in MMS-treated sgs1, rmi1, and TOP3^Y356F strains are identical or if they represent different types of HRR intermediates that cannot readily be distinguished by the 2D gel technique. One possibility, based on the known enzymatic functions of BLM, hTOPOIII and RMI1 (Wu and Hickson, 2003; Plank et al., 2006; Raynard et al., 2006; Wu et al., 2006), is that X-molecules arising in sgs1 mutants are double Holliday junctions, whereas those arising in TOP3^Y356F, or rmi1, mutants are hemicatenanes formed by the Sgs1-dependent convergent branch migration of double Holliday junctions.

Surprisingly, we found that although mutation of RAD51 prevented persistent MMS-induced HRR intermediates in sgs1, rmi1, and TOP3^Y356F cells, mutation of SHU1 did not. However, mutation of SHU1 attenuated the level of MMS-induced HRR intermediates in sgs1, rmi1, and TOP3^Y356F cells. This finding is consistent with the report that shu1 suppresses the elevated recombination rate in sgs1 and top3 cells (Shor et al., 2005). Taken together with the fact that the shu mutations are epistatic to sgs1 for MMS sensitivity and also suppress the poor growth caused by deletion of RMI1 or TOP3, our data are consistent with an early role for the Shu proteins in HRR to promote the formation of HR intermediates that are resolved by the Sgs1-Rmi1-Top3 complex. We therefore propose that, in the absence of the Shu proteins, the maturation of Rad51-filaments into Sgs1-Rmi1-Top3 substrates (e.g., double Holliday junctions) occurs inefficiently (Figure 6). If true, this suggests that the Shu proteins likely function as nonessential HRR accessory factors that facilitate efficient and timely HRR. These findings are consistent with the proposal that Shu proteins may represent so-called "Rad51 paralogs" (Martin et al., 2006), because these have been demonstrated to facilitate the action of Rad51 in HRR (Takata et al., 2001).

View larger version (16K): [in this window] [in a new window]   Figure 6. Proposed role of the Shu complex in promoting the formation of HRR intermediates that are processed by Sgs1-Rmi1-Top3. MMS-induced DNA lesions cause discontinuities in DNA synthesis, leading to the accumulation of ssDNA gaps during S-phase. These ssDNA gaps can be repaired by Rad51-dependent homologous recombination repair (HRR) or postreplicative gap filling by a translesion polymerase. RPA binds to ssDNA gaps and activates the DNA damage checkpoint. If HRR is initiated, then the engagement of the HRR machinery causes removal of RPA and subsequent deactivation of the checkpoint signal. However, the absence of Rad51, Rad54, or any one of the four Shu proteins results in a persistent activation of the DNA damage checkpoint, presumably due to inefficient RPA removal by the HRR machinery. DNA structures that are predicted to (persistently) activate the checkpoint are indicated by an asterisk. Rad51, Rad54, and associated proteins catalyze strand invasion and copying of genetic information from the sister chromatid to create (extended) D-Loops that either are substrates for synthesis-dependent strand annealing or else are converted into double Holliday junctions for processing by Sgs1-Rmi-Top3. In the absence of Sgs1, Rmi1, or Top3, late-stage HRR intermediates persist and are detectable as X-molecules on 2D gels. In the absence of Shu proteins, poor-quality Rad51-filaments are unstable, and maturation of these into Sgs1-Rmi1-Top3 HRR substrates occurs very inefficiently. Thus, X-molecules detectable in sgs1, rmi1, or top3/TOP3Y356F cells are attenuated.

	ACKNOWLEDGMENTS

 
We thank Drs. P. McHugh and L. Wu and various members of the Hickson laboratory for helpful comments and criticisms. We also thank the Cancer Research UK peptide synthesis laboratory for providing -factor. This work was funded by Cancer Research UK and the Bloom's Syndrome Foundation.

	Footnotes

 
This was published online ahead of print in MBC in Press (http://www.molbiolcell.org/cgi/doi/10.1091/mbc.E07-05-0490) on August 1, 2007.

The online version of this article contains supplemental material at MBC Online (http://www.molbiolcell.org).

Address correspondence to: Ian D. Hickson (ian.hickson{at}imm.ox.ac.uk)

Abbreviations used: DSB, double-strand break; FACS, fluorescent-activated cell sorting; HRR, homologous recombination repair; HU, hydroxyurea; MMS, methyl methane sulfonate.

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