Cells repair most double-strand breaks (DSBs) that arise during replication or by environmental insults through homologous recombination, a high-fidelity process critical for maintenance of genomic integrity. However, neither the detailed mechanism of homologous recombination nor the specific roles of critical components of the recombination machinery—such as Bloom and Werner syndrome proteins—have been resolved. We have taken a novel approach to examining the mechanism of homologous recombination by tracking both a DSB and the template from which it is repaired during the repair process in individual yeast cells. The two loci were labeled with arrays of DNA binding sites and visualized in live cells expressing green fluorescent protein–DNA binding protein chimeras. Following induction of an endonuclease that introduces a DSB next to one of the marked loci, live cells were imaged repeatedly to determine the relative positions of the DSB and the template locus. We found a significant increase in persistent associations between donor and recipient loci following formation of the DSB, demonstrating DSB-induced pairing between donor and template. However, such associations were transient and occurred repeatedly in every cell, a result not predicted from previous studies on populations of cells. Moreover, these associations were absent in sgs1 or srs2 mutants, yeast homologs of the Bloom and Werner syndrome genes, but were enhanced in a rad54 mutant, whose protein product promotes efficient strand exchange in vitro. Our results indicate that a DSB makes multiple and reversible contacts with a template during the repair process, suggesting that repair could involve interactions with multiple templates, potentially creating novel combinations of sequences at the repair site. Our results further suggest that both Sgs1 and Srs2 are required for efficient completion of recombination and that Rad54 may serve to dissociate such interactions. Finally, these results demonstrate that mechanistic insights into recombination not accessible from studies of populations of cells emerge from observations of individual cells.
Genetic recombination not only promotes genetic diversity in a population but also insures the integrity of an organism's genome. Inappropriate or inefficient recombination drives tumor formation in cancers and underlies certain premature aging diseases in humans, such as Bloom's and Werner's syndromes. To gain insight into the role of recombination in these diseases, the researchers developed a means of observing recombination in individual cells. They accomplish this by repeatedly monitoring the positions of the two chromosomal segments participating in recombination. Their observations show that interacting strands undergo repeated association and dissociation prior to fully completing recombination. This view contradicts the accepted model of recombination as an uninterrupted, continuous process, but explains, as has been observed, how sequences from different regions of the genome might arrive at a single site after recombination. Moreover, their results show that mutants lacking the Bloom's or Werner's genes have difficulty bringing the interacting strands of DNA together. Such delays could lead to errors in genome structure that could account for the disease characteristics. In short, observing recombination in individual cells reveals unexpected but important features of this process.
Citation: Houston PL, Broach JR (2006) The Dynamics of Homologous Pairing during Mating Type Interconversion in Budding Yeast. PLoS Genet 2(6): e98. https://doi.org/10.1371/journal.pgen.0020098
Editor: Maria Pia Longhese, University of Milan–Bicocca, Italy
Received: February 14, 2006; Accepted: May 12, 2006; Published: June 23, 2006
Copyright: © 2006 Houston and Broach. This is an open-access article distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.
Funding: This work was supported by grant GM48540 from the National Institutes of Health.
Competing interests: The authors have declared that no competing interests exist.
Abbreviations: DSB, double-strand DNA break; GFP, green fluorescent protein
Double-stranded DNA breaks (DSBs) occur spontaneously during replication and by exposure to certain genotoxic chemicals or ionizing radiation. Efficient repair of these DSBs can be accomplished non-conservatively by non-homologous end joining (NHEJ) or with exact fidelity using homologous recombination. Failure to repair these DSBs accurately can perturb the normal cell cycle progression and increase genomic instability, thereby promoting tumorigenesis.
The timing, choreography, and genetic dependencies of steps during DSB repair by homologous recombination in the budding yeast have been examined on populations of cells following synchronous initiation of a DSB [1–3]. Moreover, microscopic observation of live cells expressing specific green fluorescent protein (GFP)-tagged proteins has provided information on the temporal sequence and dependencies in recruitment of repair and recombination proteins to nuclear repair foci that form following initiation of DSBs . These studies demonstrated that the repair of DSBs by homologous recombination occurs in several steps. First, shortly after formation of a DSB, signal transduction and nuclease modules recognize and process the DSB. The Mre11, Rad50, Xrs2 (MRX) module in conjunction with the Tel1 kinase tether the ends of the newly formed DSB and promote efficient nucleolytic processing of the break to generate 3′ single-stranded ends [4–6]. The single-strand binding complex, RPA, binds to the newly exposed single-stranded region. Rad51 then forms a nucleoprotein filament on the exposed single strands at the ends of the DSB. Rad52, Rad54, Rad55, and Rad57 also bind to the exposed single-strand region and promote formation and/or stabilization of the Rad51 nucleoprotein filament, in part by alleviating the inhibitory effect of RPA on Rad51 binding [2,3].
In the second step in DSB repair, the nucleoprotein filament performs a search for homology and associates with a donor sequence. Initial association likely occurs through a triple-strand association between the nucleoprotein filament with the duplex donor DNA, followed by isomerization to form a D-loop with the single-strand end of the DSB invading the duplex to pair with the template strand of the duplex . Rad54 plays a role in strand invasion along with DNA polymerase holoenzyme , which extends the invading 3′ strand. In yeast, the joint between invading and template strands is resolved 2 to 4 h after DSB formation. However, most of these studies report only on the average behavior of populations of cells. Our results described in this report indicate that the discrete dynamics of the recombination process in individual cells are quite different.
A variety of helicases participate in recombination. One such helicase in Saccharomyces, Sgs1, is a member of the RecQ helicase family that resembles the WRN and BLM helicases defective in Werner and Bloom genome instability syndromes in humans . Deletion of SGS1 increases the frequency of gross chromosomal rearrangements and, in conjunction with deletion of a second related helicase gene, SRS2, causes a severe growth defect in yeast. This growth defect can be suppressed by mutations that inhibit initiation of homologous recombination, suggesting that Sgs1 and Srs2 prevent formation of aberrant, irreparable complexes that could arise during the process of homologous recombination . Other studies have indicated that, despite the synthetic lethality of sgs1 and srs2, the two corresponding proteins act at different steps in recombination, with Sgs1 helping to resolve recombination intermediates and Srs2 functioning to counteract initiation of recombination . Moreover, these proteins appear to play critical roles during replication fork stalling, with Sgs1 promoting resolution of recombination-dependent structures that form at damaged replication forks and Srs2 assisting to prevent the formation of such structures . Finally, while these studies suggest that Sgs1 and Srs2 suppress recombination, sgs1 mutants are defective in DNA damage–induced heteroallelic recombination, suggesting that Sgs1 may promote recombination in certain contexts [9,12]. Consistent with this positive role of the yeast RecQ helicase in recombination, WRN protein is required to complete mitotic recombinants efficiently in unperturbed conditions in fibroblast cell lines . Thus, we still do not have a clear conception of the function of RecQ helicases in recombination and a full appreciation of the mechanism underlying the pathological consequences of losing RecQ function in mammalian cells.
In order to address unresolved aspects of recombination, we have examined mating type switching in individual yeast cells. Mating type switching in yeast has been extensively used as a model system for homologous recombination [2,3,14,15]. Haploid Saccharomyces cells exist in one of two mating types, a or α, dependent solely on the particular allele present at the MAT locus on Chromosome III. Haploid cells can change mating type as often as every generation (reviewed in [16,17]) through recombination initiated by introduction of a DSB at the MAT locus, catalyzed by an endonuclease encoded by HO . Switching then occurs by a gene conversion event, without associated crossover, that replaces the mating information at the MAT locus with the opposite mating information, a or α, present at either of two repository mating loci, HML and HMR, located at the opposite ends of Chromosome III. Mating type switching follows a precise developmental pattern that results from the intricate mode of transcriptional regulation of the HO gene and from a highly regulated interaction between donor and recipient loci [19–21]. Specifically, cell type dictates which donor locus is selected for participation in the gene conversion event. MATa cells predominantly select HML, which normally contains α mating information, whereas MATα cells select HMR, which normally contains a mating information. This selection process insures efficient mating type switching.
We have created strains for observing the interaction of MAT and its preferred donor locus during mating type switching in single cells in order to assess the discrete dynamics of the pairing process. The results of this analysis, described below, indicate that the dynamics of gene conversion during mating type interconversion are quite different, and much more rapid, in individual cells than would be predicted from previous studies on populations of cells. Furthermore, the dynamics of interaction of donor and recipient loci in various mutant backgrounds suggest unexpected roles for several components of the recombination machinery.
Method for Observing Homologous Pairing in Live Yeast Cells
We developed strains and a method to observe the pairing of homologous DNA loci during recombination in live yeast cells by adopting previously described methods to visualize discrete chromosomal loci and applying them to the process of mating type switching [22,23]. As diagrammed in Figure 1A, we inserted an array of lac repressor binding sites adjacent to a donor locus, HML, and an array of tet repressor sites adjacent to the recipient locus, MAT, and then expressed both GFP-lacI and GFP-tetR fusions in the strain [23,24]. We performed these experiments in a cdc15–2 strain, which arrests at the non-permissive temperature just prior to exit from telophase. By shifting the strain from non-permissive to permissive temperature, cells synchronously enter G1 phase . This insures that, at the beginning of our experiments, all cells have only one marked chromosome and are at the stage of the cell cycle in which switching normally occurs. Finally, the strain also carries the HO gene under control of the GAL10 promoter to allow regulated induction of a DSB at the MAT locus.
(A) Diagram of yeast cells used in this study, indicating the relevant structure of Chromosome III, which carries arrays of LacO sites at MAT and TetO sites at HML. The strain also constitutively expresses LacI-GFP and TetR-GFP protein fusions and contains a plasmid that carries a galactose-inducible HO gene and CFP under control of an α-cell type–specific promoter.
(B) Fluorescence micrograph of a MATa strain, Y3343, freshly transformed with plasmid B2609, 6 h after induction of HO. Green dots mark HML and MAT loci in each cell, and uniform blue staining indicates that the cell has undergone a mating type switch to MATα. Cells are demarked by the diffuse cyan fluorescence, and the diffuse yellow fluorescence defines the nuclei.
(C) Deconvolved and compressed images showing GFP foci in isogenic strains carrying or lacking PGAL1-HO, taken 90 min following induction with galactose.
To monitor cells under controlled conditions, we affixed yeast cells to the coverslip of a thermally regulated flow cell in which we could control both temperature and medium while repeatedly interrogating by fluorescence microscopy multiple fields of cells on the coverslip. Cells freshly transformed with the PGAL1-HO plasmid were pregrown in non-inducing medium at 23 °C and then shifted to 35 °C for 3 h. This arrests all the cells just prior to entry into G1 phase of the cell cycle. After attaching cells to the coverslip and centering the flow cell on the microscope, we initiated each experiment by simultaneously inducing the DSB at MAT and releasing cells from cell cycle arrest. This was accomplished by perfusing the flow cell with medium containing 2% galactose to induce HO and shifting the temperature of the flow cell to 23 °C. After DSBs were induced, we shut off HO expression to prevent subsequent rounds of recombination, by changing the medium flowing over the cells to one lacking galactose and containing 2% glucose. Since HO mRNA and HO protein have very short half-lives, no further DSB formation occurs much beyond this medium switch .
We confirmed that cells attached to the coverslip and treated as above switch efficiently and select the appropriate donor. We can determine by fluorescence microscopy at the single-cell level whether a cell has switched from MATa to MATα by including in the strain a PMFα1-CFP construct, which consists of the CFP coding region under control of the MFα1 promoter. The MFα1 promoter is active only in MATα cells. Accordingly, a MATa cell prior to switching does not express CFP, but after it switches to MATα, it begins to accumulate CFP. We found that less than 1% of MATa PMFα1-CFP cells without HO induction accumulated CFP at the end of the experiment, whereas approximately 50% of the cells that were induced for HO expression accumulated CFP (Figure 1B). The switching rate of 50% agrees well with that determined by genetic assay with the same strain induced for 1 h in liquid culture, indicating that manipulations in the flow cell and observations by fluorescence microscopy did not affect the process of switching. The rate is somewhat less than the 85% switching rate observed with wild-type HO cells and reflects a slight reduction in the initiation of switching under our induction conditions, rather than any alteration in donor preference (unpublished data). This agrees with Southern analysis and plate-based mating assays of our test strain, from which we concluded that under our induction conditions, approximately half of our test cells sustain a DSB at MAT.
Pairing between Donor and Recipient Loci during Recombination Is Short and Reversible
To monitor pairing of MAT and HML following initiation of a DSB at MAT, we treated MATa cells as described above and then interrogated multiple cells every minute for 3 h. We determined in each cell, at each time point, whether the nucleus contained two GFP dots or a single GFP dot. In the former case, we assume that the MAT and HML loci are unpaired. In the latter case, we assume that the two loci are either paired or simply too close to be resolved. Representative images from experiments in which HO was induced or not induced are shown in Figure 1C, demonstrating the resolution of the two loci in the test strains and indicating that all cells can be readily scored. In Figure 2A, we represent the histories of association of MAT and HML as determined by this method following HO induction for 11 wild-type MATa cells, which normally use HML as the preferred donor. In addition, we examined 11 wild-type MATα cells following HO induction. MATα cells normally use the untagged HMR locus as donor and thus MAT would not be expected to pair with HML in this strain. Finally, we examined 11 wild-type MATa cells in the absence of HO induction, in which no recombination should occur. In these representations, each line describes the history of a single cell; a white bar represents the presence of two dots in the cell's nucleus at that time point and a black bar represents the appearance of only a single dot.
(A) Shown are cell histories for 11 cells each from three strains, configured as diagrammed in Figure 1, indicating along the horizontal time line for each cell whether at each time point the cell presented a single dot (black bar) or two dots (white bar). Cells were interrogated at 1-min intervals for 3 h with galactose present (HO induced) for the first hour. Upper group: Y3343 (MATa) transformed with B2609; middle group: Y3343 (MATa) transformed with pRS415 ; and lower group: Y3342 (MATα) transformed with B2608.
(B) At each time point, the fraction of cells exhibiting a single dot was determined from the data underlying Figure 2A and plotted as a three-point running average versus time following addition of galactose. Legends are indicated to the left of the cell histories in (A). Data were calculated either including all 11 Y3343 (MATa) cells transformed with B2609 (dark blue diamonds) or for only those six cells presumed to have sustained a DSB at MAT (light blue rectangles). The horizontal axis is aligned with the cell histories in (A).
(C) The duration of periods of apparent continuous association between MAT and HML. The number of continuous, uninterrupted time points in which a cell shows only a single dot, were determined from the cell histories in Figure 2A. The number of periods of a given duration divided by the total number of periods (probability density) is plotted versus the duration of apparent association for each strain shown in 2A. MATa + HO (light blue circles); MATa − HO (green diamonds); and MATα + HO (red squares). Error bars are estimated by dividing the data successively into random halves and computing the standard deviation among 20 random halves. The dashed line shows a fit of the combined values for MATa − HO and MATα + HO cells to a normalized, exponential distribution. Under this assumption, the p-value for the likelihood that the data for MATa + HO would arise by chance is less than 10−5.
(D) Data from Figure 2A for MATa + HO cells is replotted to show only those pairing events of duration longer than 6 min.
These interrogations clearly show that, relative to the behavior of MATa cells in which HO was not induced, MATa cells in which HO was induced exhibited a statistically significant increase in the overall proportion of time MAT and HML were in close proximity. Moreover, cells from the induced culture divided into two discrete populations. In one group, cells exhibited a pattern of association that was indistinguishable from those of uninduced cells, whereas in the other group, cells exhibited a succession of extended associations between MAT and HML lasting from 7 to 15 min, starting approximately 80 min after initiation of HO expression. The fraction of cells exhibiting the extended association matched the fraction of cells that ultimately undergo gene conversion, and we believe that cells showing extended association were those in which cleavage at MAT had occurred.
A distinction between the association patterns in induced and uninduced cells is also evident by calculating the fraction of cells containing a single dot at each time point. As shown in Figure 2B, cells in which HO was not induced show a uniform, low level of association between HML and MAT throughout the experiment. On the other hand, MATa cells in which HO was induced exhibit increased association of HML and MAT starting from 80 min after initiation of induction of HO and lasting for approximately 1 1/2 h.
We also noted that, despite the fact that more than half of the cells expressing HO undergo gene conversion of MAT using HML as donor, continuous pairing between MAT and HML occurs for no longer than 17 min in any cell. We evaluated the number and duration of events in which HML and MAT appear to be in continuous association, i.e., cases in which a cell presents a single dot in an uninterrupted series of time points. As shown in Figure 2C, in uninduced MATa cells or induced MATα cells, the log of the number of association events of a given duration plotted versus the duration of association exhibits essentially a linear relation, consistent with a stochastic process in which the likelihood that HML and MAT are in close proximity at any time is independent of whether they were in close association at the previous or subsequent time point. Only one event longer than 6 min was observed in uninduced cells. In contrast, induced MATa cells exhibit a statistically significant excess of events of duration longer than 6 min. These extended associations occur for an average of 10 min, predominantly between 80 and 160 min postinduction (Figure 2D). Unexpectedly, each cell undergoes more than one prolonged association event. Moreover, during the period between two extended associations, the average distance between the two spots (0.49 ± 0.25 μm) is equivalent to that in uninduced cells (0.51 ± 0.24 μm), which do not undergo switching. Thus, we conclude that the multiple associations of donor and recipient in a single cell are independent events, suggesting that the pairing of donor and recipient loci is readily reversible during recombination.
Finally, we note from the data in Figure 2 that, in contrast to the pattern seen with MATa cells, MATα cells, which do not use HML as the preferred donor, do not exhibit an enhanced association between MAT and HML over the course of the experiment. Rather, induced MATα cells present a pattern indistinguishable from that obtained in uninduced cells. This suggests that the cleaved MAT locus does not spend significant time in association with the non-preferred donor locus during mating type switching.
Sgs1 and Srs2 Are Required for Stable Pairing between Donor and Recipient Loci during Switching
We anticipate that assembly and resolution of recombination intermediates during switching would require the activity of DNA helicases, and several genes encoding helicases have been implicated in recombination and repair. In an attempt to clarify the role of various helicases in recombination, we examined the extent of association between MAT and HML in strains deleted for individual helicase genes. As shown in Figure 3, the level of pairing between HML and MAT in a MATa sgs1Δ (Figure 3A) or a MATa srs2Δ (Figure 3B) mutant following HO induction was significantly reduced compared to that observed in wild-type cells. The level of pairing in the induced srs2Δ strain was slightly higher than that in the same strain lacking HO but was certainly significantly lower than that observed in the wild-type strain. More dramatically, the level of apparent pairing in the induced sgs1Δ strain was not significantly different than that in the same strain lacking HO. Thus, association of the DSB with the donor locus appears to be delayed in both mutants. These results would predict that sgs1 or srs2 mutants should be defective in mating type switching.
Data were obtained for 17 cells under each condition and presented as described in Figure 2 for sgs1 (A) and srs2 (B) mutant strains, with the difference that cells were interrogated every 2 min starting 30 min after addition of galactose.
To confirm the roles of Sgs1 and Srs2 in repair of DSBs during mating type switching, we examined the effect of introducing these mutants into wild-type HO strains. A diploid HO/HO strain, which we used previously to determine switching frequencies under normal conditions , was rendered heterozygous for deletion of SGS1 or SRS2. After sporulation and dissection, we examined the behavior of cells and the switching pattern during outgrowth of spore clones. As seen in Table 1, HO SGS1 and HO SRS2 clones showed normal growth and switching patterns. Similarly, ho sgs1 and ho srs2 control strains grew indistinguishably from ho SGS1 and ho SRS2 strains (unpublished data and ). In contrast, HO srs2 spore clones were smaller than their sister HO SRS2 spore clones, and pedigree analysis of cells following germination indicated that mother cells in the lineage exhibited delayed cell cycle progression or failed to divide at all. This growth pattern is similar to, but less severe than, the pedigree of death that occurs in HO yeast strains incapable of completing HO-induced switching , which we observe with rad52 HO and rad54 HO spores (Table 1). Spore clones carrying an sgs1 deletion did not show as extensive delays in growth as did srs2 spore clones, but the viability of mother cells in pedigrees was clearly less than that seen with sister SGS1 spore clones (unpublished data). These results are consistent with the assumption that srs2 mutants, and to a lesser extent, sgs1 mutants, fail to efficiently heal DSB generated in lineages of HO haploid cells.
Switching Efficiency in Mutant Strains
We also find that, although both srs2 and sgs1 mutants exhibited normal donor selection in a MATa background, both mutants showed essentially random selection of donor loci following switching in a MATα background (Table 1). This observation confirms that the mutants exhibit a delay in repairing the DSB during mating type switching. In cells in which repair of the HO-induced DSB at MAT is delayed, resection of the DSB into the adjacent coding region results in deletion of the MAT locus, rendering the cell temporarily mat-null. Since mat-null cells exhibit a MATa phenotype, donor selection in MATa cells would be unaffected by this delay-induced resection. However, the delay-induced resection in a MATα would render the cell phenotypically MATa, changing the donor preference from HMR to HML. Thus, the extent to which MATα cells select HML as donor reflects the extent of delay in the repair of the DSB. Accordingly, our results confirm that Srs2 and Sgs1 are both required for timely repair of DSBs.
Rad54 Is Not Required for Stable Pairing between Donor and Recipient Loci during Switching
RAD54 encodes a member of the Swi2/Snf2 family of DNA-stimulated ATPases, and loss of RAD54 function substantially diminishes homologous recombination and renders cells sensitive to ionizing radiation. Consistent with previous reports that Rad54 is required for completion of DSB repair , HO rad54 spore clones yield microcolonies in our donor preference assay, resulting from extensive death of mother cells during growth of the clone (Table 1). Previous studies on the precise role Rad54 plays in recombination have been inconsistent, with some evidence pointing to its involvement in initial formation of the Rad51 presynaptic filament and other evidence suggesting that it functions after formation of the presynaptic complex [2,3]. To help resolve this issue, we examined pairing of MAT and HML following HO induction in a rad54Δ mutant.
As evident in Figure 4A, the two loci paired as efficiently in the rad54 mutant as in wild-type cells. Surprisingly, MAT and HML also paired efficiently in a MATα rad54 mutant strain (Figure 4B), despite HML not being the preferred donor for switching in MATα cells. This result is quite distinct from that observed with HO-induced RAD54 MATα cells, which, as shown in Figure 2, show no prolonged association between MAT and HML. Therefore, association between donor and recipient loci proceeds efficiently in the absence of Rad54 function and, furthermore, association with the inappropriate donor occurs as frequently as does association with the preferred donor.
To study the dynamics of recombination, we have developed a real-time assay for homologous pairing in individual living cells. The results from this assay correlate well with previous observations on the time course of the recombination reaction obtained with populations of cells, but also reveal unanticipated aspects of recombination not previously accessible through population studies. The system we examined—mating type interconversion following synchronous cleavage of the MAT locus by HO induction in Saccharomyces—has been used extensively as a model system for homologous recombination [2,3,14,15]. Previous studies examining products and intermediates of recombination in vivo by PCR or Southern analysis indicate that strand invasion begins approximately 90 min following HO induction and ends approximately 90 min later [2,14]. Furthermore, by using chromatin immunoprecipitation to quantify in vivo association of recombination proteins with regions of the genome, several studies determined that Rad51 initially associates with the DSB at MAT approximately 30 min following HO induction—coincident with DSB formation—and with the donor locus, HML, approximately 30 min later [2,3]. This association continues for up to 3 h. These results suggest that synapsis between the DSB and the donor locus begins approximately 60 min following HO induction and that the two loci remain physically associated for up to several hours. Our data are consistent with these population studies in that we observe an overall increase in the average association of MAT and HML beginning 80–90 min following HO induction and lasting for 60–90 min.
Monitoring the association of HML and MAT over time in individual cells revealed aspects of synapsis not evident from these population studies. First, we observed a statistically significant increase in the number of periods of extended association of MAT and HML in induced versus uninduced cells, although the durations of these associations were in general no longer than 17 min and averaged approximately 10 min. Thus, distinct periods of synapsis in individual cells are much shorter than would be expected from population studies. Second, individual cells exhibit several distinct periods of extended association following HO induction. Since HO protein is not present at these later times and since the same pattern occurs in rad54 mutants unable to complete recombination, these sequential associations do not represent sequential rounds of resolution and re-initiation of recombination. Also, these multiple associations are unlikely to be a single synaptic junction punctuated by dynamic stretching, since the distributions of distances between the two loci in the interval between two such extended associations are identical to those in uninduced cells. Rather, we would interpret these observations to suggest that the Rad51-mediated complex formed between the DSB and the template is reversible. For example, the initial Rad51-mediated triple-strand complex consisting of the Rad51-coated single-strand DNA bound to the homologous double-strand region of the donor template could be resolved in either two ways: isomerization to yield a single-strand invasion creating a D-loop on the template, or disassociation to return to the pre-complex state. Alternatively, D-loop formation could be the reversible step, either before or after extension of the single-strand primer. Our results suggest that either or both of these initial interactions between the DSB and the template can occur several times before completion of recombination. Such cycles for strand invasion have been suggested to occur during repair of DSBs in Drosophila . The results from those studies with Drosophila would suggest that dissociation could occur not only after D-loop formation but also after partial extension of the primer, as depicted in Figure 5. Finally, we note that the multiple, stochastic associations between the DSB and the donor locus in individual cells account for the apparent extended association between the two loci as measured on a population basis.
Following HO-induced double-strand cleavage of MAT, Rad51 (ellipses) binds to the resected single-strand ends of the DSB and promotes their association with both HML and HMR via homology present at all three loci. Blue- and pink-colored regions at the mating type loci represent allele-specific DNA sequences, which are bracketed by sequences in common among all three loci (shown in brown). Dissociation of these initial complexes, suggested to be paranemic joints in this illustration, is stimulated by Rad54-promoted removal of Rad51. Since these associations are seen only in rad54 cells, we assume that they are too short-lived to be evident in wild-type cells, and thus are indicated in brackets. We propose that Rad54 also stimulates conversion of the initial joint complex into an initial D-loop structure, also by promoting removal of Rad51, allowing subsequent extension of the heteroduplex between the invading strand and donor locus, and elongation of the invading strand. These subsequent steps are promoted or stabilized by Srs2 and Sgs1. Our results suggest that one or more of the later-stage intermediates in recombination, perhaps even after partial extension of the invading strand, are also reversible prior to completion of the gene conversion event. The dissociated, but incompletely healed, MAT locus then would recycle through Rad51 filament formation and reassociation with the donor locus, ultimately yielding a completely reconstituted MAT locus.
We explored some of the genetic requirements for homologous pairing in our system. We first examined the role of the helicases encoded by SGS1 and SRS2 and found that inactivation of either of these genes delayed sustained association between MAT and HML. We confirmed this result obtained from the physical pairing assay by showing that repair of DSBs during mating type switching is delayed in HO spore clones carrying sgs1 or srs2 mutations. Thus, each of these genes appears to be independently required for efficient homologous pairing in our system, an observation consistent with previous studies suggesting that, despite their synthetic lethality, Sgs1 and Srs2 play independent roles in recombination [9,12].
Our results are somewhat unexpected since previous genetic and biochemical studies have implicated Srs2 in inhibiting initiation of recombination by promoting disassembly of Rad51 filament complexes [31–33], whereas Sgs1 and other RecQ helicases are likely involved in resolution of recombination intermediates [10,34–38]. Thus, we might have anticipated that sgs1 or srs2 mutants would exhibit prolonged association of MAT and HML in our assay rather than reduced association. However, previous studies have shown that, although sgs1 mutants exhibit a hyperrecombination phenotype under some conditions, they are hyporecombinogenic for DNA damage–induced heteroallelic recombination [9,12], a situation similar to DSB-induced mating type switching. In addition, Sgs1 is required for telomere elongation in the absence of telomerase, apparently through recombination via a break-induced repair process that is mechanistically similar to the DSB-induced gene conversion we have studied here . Thus, Sgs1 could be required to stabilize the initial joint formed between a DSB and the donor by promoting formation of an extended D-loop or initiation of synthesis to extend the invading 3′ end. In a similar vein, the effect of inactivation of SRS2 on recombination depends on the particular assay used. Perhaps most relevant to our observations, Aylon et al.  examined gene conversion of a ura3 allele from an ectopic site of homology following introduction of an HO-induced DSB within the ura3 allele, an assay quite similar to mating type switching. They found that inactivation of SRS2 substantially reduced the level of strand invasion and subsequent extension of the invading strand and concluded that only a small subpopulation of srs2 cells are able to complete recombination, whereas the majority of cells behaved as if no homology to the DSB were present in the cell. This is consistent with results from Pâques and Haber suggesting that Srs2 functions to extend and stabilize initial joints between the ssDNA-Rad51 nucleofilament and the donor locus . Thus, both Sgs1 and Srs2 have been implicated independently in stabilizing initial joint formation following DSB-induced recombination, consistent with our current observations.
We also used our assay to examine the role of RAD54 in pairing in vivo. Rad54, a dsDNA-dependent ATPase of the Swi2/Snf2 family of chromatin remodeling enzymes, is required for resistance to ionizing radiation and repair of DSBs by homologous recombination [7,29]. Rad54 interacts with Rad51 in vivo, and addition of Rad54 to Rad51-mediated recombination reactions dramatically stimulates homologous pairing in vitro. In addition, Rad54 is essential for DNA strand invasion reactions using chromatin-packaged substrates . Two studies recently used chromatin immunoprecipitation to examine the in vivo requirement of Rad54 for association of Rad51 with MAT and HML following initiation of a DSB at MAT. Wolner et al.  found that Rad51 association with either locus was essentially eliminated in a rad54Δ mutant background and provided evidence that Rad54 promotes formation of the Rad51 presynaptic filament. On the other hand, Sugawara et al.  found that Rad51 could, in fact, associate with MAT and HML in the absence of Rad54, but the synaptic complex could not proceed to a mature strand invasion complex in the absence of Rad54.
We find that MAT and HML associate as well in a rad54 mutant as they do in a wild-type strain. Moreover, we find that MAT associates frequently with HML even in a MATα strain, in which HML is the non-preferred donor. One possible explanation for these associations between MATα and HML is phenotypic switching of mating due to delay in resolution of the MAT DSB, as we proposed for sgs1 and srs2 mutants. However, since the timing of the observed associations in rad54 mutants is the same as that seen in wild-type cells, in which no phenotypic switching occurs, we conclude that the inappropriate associations are not the result of phenotypic change in mating type. Rather, our results suggest that Rad54 plays a critical role in recombination at a step beyond initial synapsis. This conclusion is consistent with the observations above from Sugawara et al.  and also with subsequent work by Wolner and Peterson . Although Rad54 may also have a role in vivo in promoting Rad51 nucleofilament formation, our results would suggest that this is not the rate-limiting step in recombination stimulated by Rad54. Our results can be explained by the proposal, based on the observation that Rad54 stimulates dissociation of Rad51 from dsDNA, that Rad54-stimulated removal of Rad51 promotes either dissociation of the paired loci or conversion of the initial synaptic intermediate into a D-loop and the subsequent extension of the heteroduplex region . Thus, in the absence of Rad54, pairing between the DSB and either donor locus should still occur, but the formation of a D-loop and subsequent elongation of the invading strand would not. This model would account for the fact that we observe pairing of MAT and HML in the non-preferred mating background, reflecting formation of Rad51-mediated pairing of MAT with either donor locus via the common homology at all three loci. We would propose that these initial pairing events would be eliminated in a Rad54-dependent fashion, either reversing the initial pairing or promoting progression in the appropriate partner to productive recombination.
A model for recombination during mating type switching is presented in Figure 5. Our results suggest that following HO-induced cleavage of MAT, the locus associates readily and reversibly with both HML and HMR. Donor selection would then occur at a subsequent step, for instance in the isomerization from the initial synaptic intermediate to the D-loop, upon recruitment of DNA polymerase or with extension of the invading strand. Cell type–specific loading of factors promoting D-loop formation and strand extension at the preferred donor locus would then fix donor preference. Our results also suggest that this step is stimulated by both Sgs1 and Srs2. Our results further indicate that, even in the presence of Rad54, the association between donor and template are reversible. This would suggest that even after D-loop formation, the donor and recipient loci can dissociate. Moreover, given recent results suggesting that a double-strand gap requires multiple cycles of strand invasion, synthesis, and dissociation of the nascent strand during repair of a DSB in Drosophila , this reversibility could occur after partial extension of the invading strand, as represented in Figure 5. Further analysis should resolve precisely at what steps recombination is reversible. However, these results clearly indicate that the recombination process is reversible at a much later stage than previously anticipated and raises a novel mechanism for generating genetic diversity.
Materials and Methods
Plasmids and strain construction.
Plasmid pPJS218 (HIS3 TetR-GFP LacI-GFP) has been previously described . Plasmid B2609 (LEU2 CEN4 ARS1 PGAL1-HO PMFα1-CFP) was constructed by in vivo recombination by co-transforming a leu2 trp1 yeast strain with plasmid B2686 (LEU2 TRP1 CEN4 ARS1 PGAL1-HO) and two PCR fragments, one spanning the MFα1 promoter and one spanning the coding sequence of CFP from plasmid pDH3 (http://depts.washington.edu/~yeastrc). Both PCR fragments carried 50 base pair ends with homology to direct recombination with the appropriate adjacent fragments. Leu+ FAA-resistant transformants were selected and plasmid was recovered by transformation into Escherichia coli. Plasmid B2608 (LEU2 CEN4 ARS1 PGAL1-HO PMFa1-CFP) was constructed in an analogous fashion using the MFa1 promoter. Plasmid structures were confirmed by functional assays and restriction analysis.
Strains used in this study are listed in Table 2 and were all obtained from the S288C-derived haploid strain S150-2B. Construction of strains carrying DNA binding site arrays adjacent to MAT and HML have been described previously . GFP fusions were introduced by integrative transformation of plasmid pPJS218, selecting His+ transformants. The cdc15–2 allele was introduced into S150-2B by first integrating plasmid B2400 (pRS406-cdc15–2) into the CDC15 locus and then selecting FOA-resistant revertants of selected transformants. FOA-resistant, temperature-sensitive isolates were scored microscopically for terminal arrest phenotype to confirm the presence of the cdc15–2 allele. The cdc15–2 allele was then introduced into the tagged strains by genetic crosses. Deletion alleles were introduced as needed by selecting transformants resistant to G418 (200 μg/ml, CalBiochem, San Diego, California, United States) following transformation with PCR products using DNA from the appropriate strain from the kanMX deletion collection, and primers 500 base pairs upstream, 5′, and downstream, 3′, of the gene.
Strains for donor preference were obtained by transformation of strain Y2902 using PCR products from the deletion collection as described above. Donor preference in spore clones of mutant strains were determined as described previously .
The yeast strains were freshly transformed with plasmid B2609 or B2608 and grown 3 d at 23 °C on SC-leu media. Three transformants of each strain were inoculated to 20-ml SC-leu broth + 2% glucose and grown overnight at 23 °C. Cultures were then diluted into SC-leu broth + 2% raffinose and lacking glucose to a density of 106 cells/ml and incubated for 3 h at 37 °C to arrest growth and render GAL1-HO readily inducible. Cells from each culture were examined by fluorescence microscopy to confirm that most exhibited two nuclear GFP dots and none expressed CFP. Cells from cultures with these properties were applied to a microscope coverslip (No.1 22×60 mm, Corning, Corning, New York, United States) precoated with Concanavalin A (Sigma, St. Louis, Missouri, United States) as described (http://www.cgr.harvard.edu/thornlab/protocols/ConA.htm). Samples of test and control cultures were applied to the same coverslip but separated by a hydrophobic Pap pen (Daido Sangyo Co. Ltd., Tokyo, Japan) to preclude mixing. The cells were mounted on a flow cell (BioSurface Technologies, Bozeman, Montana, United States) and perfused with SC-leu medium + 2% raffinose as the sole carbon source, maintained at 37 °C with a thermocouple detector and a feedback-modulated heated stage. Cells from several fields of each strain were repeatedly interrogated every 1 or 2 min using a Nikon Eclipse TE200 microscope with 100× objective (1.4 aperture) and a planapochromatic light source (Nikon, Tokyo, Japan). Approximately 5 min after mounting the coverslip on the flow cell, the temperature was shifted to 23 °C to release the cdc15–2–imposed cell cycle arrest, and medium perfusing the flow cell was changed to SC-leu + 2% galactose. This time was set as the zero time point, and galactose perfusion was maintained for 60 min, at which time medium was switched to SC-leu + 2% glucose. At each interrogation, we captured 30 0.4-μm Z-sections of a 512 × 512 pixel image (0.1 μm per pixel), and prior to analysis, the images were processed by five cycles of deconvolution using Deltavision SoftWoRx v. 2.5 (Applied Precision, Issaquah, Washington, United States). The locations of GFP dots in each cell in a field were determined from the identification of the maximum intensity in the three-dimensional images. Distance between the separable dots was determined by applying the Pythagorean triangulation to the Cartesian coordinates of the maxima using Softworx pick points function and processed with the timedist and yeastparser programs (John Houston, Multimedia Gaming, Austin, Texas, United States). Those cells in which only a single intensity maximum was observed were defined as having a single dot, i.e., the two loci were closer than could be resolved by our microscopy.
We thank Dr. Mark Rose for helpful discussions and assistance with the Deltavision microscope, and Dr. David Tank and Don Peoples for advice and technical help in design and construction of the flow cell apparatus. John Houston provided methods for working with the dataset parsing, and Erin Smith and Dr. William Bialek provided invaluable assistance with statistical analysis of the data.
PLH and JRB conceived and designed the experiments. PLH performed the experiments. PLH analyzed the data. PLH and JRB wrote the paper.
- 1. Holmes A, Haber JE (1999) Physical monitoring of HO-induced homologous recombination. Methods Mol Biol 113: 403–415.
- 2. Sugawara N, Wang X, Haber JE (2003) In vivo roles of Rad52, Rad54, and Rad55 proteins in Rad51-mediated recombination. Mol Cell 12: 209–219.
- 3. Wolner B, van Komen S, Sung P, Peterson CL (2003) Recruitment of the recombinational repair machinery to a DNA double-strand break in yeast. Mol Cell 12: 221–232.
- 4. Lisby M, Barlow JH, Burgess RC, Rothstein R (2004) Choreography of the DNA damage response: spatiotemporal relationships among checkpoint and repair proteins. Cell 118: 699–713.
- 5. Lobachev K, Vitriol E, Stemple J, Resnick MA, Bloom K (2004) Chromosome fragmentation after induction of a double-strand break is an active process prevented by the RMX repair complex. Curr Biol 14: 2107–2112.
- 6. Kaye JA, Melo JA, Cheung SK, Vaze MB, Haber JE, et al. (2004) DNA breaks promote genomic instability by impeding proper chromosome segregation. Curr Biol 14: 2096–2106.
- 7. Sung P, Trujillo KM, Van Komen S (2000) Recombination factors of Saccharomyces cerevisiae. Mutat Res 451: 257–275.
- 8. Ellis NA, Groden J, Ye TZ, Straughen J, Lennon DJ, et al. (1995) The Bloom's syndrome gene product is homologous to RecQ helicases. Cell 83: 655–666.
- 9. Gangloff S, Soustelle C, Fabre F (2000) Homologous recombination is responsible for cell death in the absence of the Sgs1 and Srs2 helicases. Nat Genet 25: 192–194.
- 10. Fabre F, Chan A, Heyer WD, Gangloff S (2002) Alternate pathways involving Sgs1/Top3, Mus81/ Mms4, and Srs2 prevent formation of toxic recombination intermediates from single-stranded gaps created by DNA replication. Proc Natl Acad Sci U S A 99: 16887–16892.
- 11. Liberi G, Maffioletti G, Lucca C, Chiolo I, Baryshnikova A, et al. (2005) Rad51-dependent DNA structures accumulate at damaged replication forks in sgs1 mutants defective in the yeast ortholog of BLM RecQ helicase. Genes Dev 19: 339–350.
- 12. Onoda F, Seki M, Miyajima A, Enomoto T (2001) Involvement of SGS1 in DNA damage-induced heteroallelic recombination that requires RAD52 in Saccharomyces cerevisiae. Mol Gen Genet 264: 702–708.
- 13. Prince PR, Emond MJ, Monnat RJ Jr. (2001) Loss of Werner syndrome protein function promotes aberrant mitotic recombination. Genes Dev 15: 933–938.
- 14. White CI, Haber JE (1990) Intermediates of recombination during mating type switching in Saccharomyces cerevisiae. EMBO J 9: 663–673.
- 15. Haber JE (2000) Lucky breaks: Analysis of recombination in Saccharomyces. Mutat Res 451: 53–69.
- 16. Bi X, Broach JR (1999) Cell type determination in yeast. In: Russo VEA, Cove DJ, Edgar LG, Jaenisch R, Salamini R, editors. Development: Genetics, epigenetics and environmental regulation. Heidelberg: Springer-Verlag. pp. 49–66. pp.
- 17. Haber JE (1998) Mating-type gene switching in Saccharomyces cerevisiae. Annu Rev Genet 32: 561–599.
- 18. Strathern JN, Klar AJS, Hicks JB, Abraham JA, Ivy JM, et al. (1982) Homothallic switching of yeast mating type cassettes is initiated by a double-stranded cut in the MAT locus. Cell 31: 183–192.
- 19. Strathern JN, Herskowitz I (1979) Asymmetry and directionality in production of new cell types during clonal growth: The switching pattern of homothallic yeast. Cell 17: 371–381.
- 20. Klar AJ, Hicks JB, Strathern JN (1982) Directionality of yeast mating type interconversion. Cell 28: 551–561.
- 21. Nasmyth K (1993) Regulating the HO endonuclease in yeast. Curr Opin Genet Dev 3: 286–294.
- 22. Robinett CC, Straight A, Li G, Willhelm C, Sudlow G, et al. (1996) In vivo localization of DNA sequences and visualization of large-scale chromatin organization using lac operator/repressor recognition. J Cell Biol 135: 1685–1700.
- 23. Straight AF, Belmont AS, Robinett CC, Murray AW (1996) GFP tagging of budding yeast chromosomes reveals that protein-protein interactions can mediate sister chromatid cohesion. Curr Biol 6: 1599–1608.
- 24. Michaelis C, Ciosk R, Nasmyth K (1997) Cohesins: Chromosomal proteins that prevent premature separation of sister chromatids. Cell 91: 35–45.
- 25. Spellman PT, Sherlock G, Zhang MQ, Iyer VR, Anders K, et al. (1998) Comprehensive identification of cell cycle-regulated genes of the yeast Saccharomyces cerevisiae by microarray hybridization. Mol Biol Cell 9: 3273–3297.
- 26. Kaplun L, Ivantsiv Y, Kornitzer D, Raveh D (2000) Functions of the DNA damage response pathway target Ho endonuclease of yeast for degradation via the ubiquitin-26S proteasome system. Proc Natl Acad Sci U S A 97: 10077–10082.
- 27. Houston P, Simon PJ, Broach JR (2004) The Saccharomyces cerevisiae recombination enhancer biases recombination during interchromosomal mating-type switching but not in interchromosomal homologous recombination. Genetics 166: 1187–1197.
- 28. Klar AJS, Strathern JN, Abraham JA (1984) Involvement of double-strand chromosomal breaks for mating-type switching in Saccharomyces cerevisiae. Cold Spring Harbor Symp Quant Biol 49: 77–88.
- 29. Krogh BO, Symington LS (2004) Recombination proteins in yeast. Annu Rev Genet 38: 233–271.
- 30. McVey M, Adams M, Staeva-Vieira E, Sekelsky JJ (2004) Evidence for multiple cycles of strand invasion during repair of double-strand gaps in Drosophila. Genetics 167: 699–705.
- 31. Ira G, Pellicioli A, Balijja A, Wang X, Fiorani S, et al. (2004) DNA end resection, homologous recombination and DNA damage checkpoint activation require CDK1. Nature 431: 1011–1017.
- 32. Krejci L, Van Komen S, Li Y, Villemain J, Reddy MS, et al. (2003) DNA helicase Srs2 disrupts the Rad51 presynaptic filament. Nature 423: 305–309.
- 33. Veaute X, Jeusset J, Soustelle C, Kowalczykowski SC, Le Cam E, et al. (2003) The Srs2 helicase prevents recombination by disrupting Rad51 nucleoprotein filaments. Nature 423: 309–312.
- 34. Constantinou A, Tarsounas M, Karow JK, Brosh RM, Bohr VA, et al. (2000) Werner's syndrome protein (WRN) migrates Holliday junctions and co-localizes with RPA upon replication arrest. EMBO Rep 1: 80–84.
- 35. Harmon FG, Kowalczykowski SC (1998) RecQ helicase, in concert with RecA and SSB proteins, initiates and disrupts DNA recombination. Genes Dev 12: 1134–1144.
- 36. Ira G, Malkova A, Liberi G, Foiani M, Haber JE (2003) Srs2 and Sgs1-Top3 suppress crossovers during double-strand break repair in yeast. Cell 115: 401–411.
- 37. Karow JK, Constantinou A, Li JL, West SC, Hickson ID (2000) The Bloom's syndrome gene product promotes branch migration of Holliday junctions. Proc Natl Acad Sci U S A 97: 6504–6508.
- 38. Wu L, Hickson ID (2003) The Bloom's syndrome helicase suppresses crossing over during homologous recombination. Nature 426: 870–874.
- 39. Huang P, Pryde FE, Lester D, Maddison RL, Borts RH, et al. (2001) SGS1 is required for telomere elongation in the absence of telomerase. Curr Biol 11: 125–129.
- 40. Aylon Y, Liefshitz B, Bitan-Banin G, Kupiec M (2003) Molecular dissection of mitotic recombination in the yeast Saccharomyces cerevisiae. Mol Cell Biol 23: 1403–1417.
- 41. Pâques F, Haber JE (1997) Two pathways for removal of nonhomologous DNA ends during double-strand break repair in Saccharomyces cerevisiae. Mol Cell Biol 17: 6765–6771.
- 42. Sugawara N, Ivanov EL, Fishman-Lobell J, Ray BL, Wu X, et al. (1995) DNA structure-dependent requirements for yeast RAD genes in gene conversion. Nature 373: 84–86.
- 43. Wolner B, Peterson CL (2005) ATP-dependent and ATP-independent roles for the Rad54 chromatin remodeling enzyme during recombinational repair of a DNA double strand break. J Biol Chem 280: 10855–10860.
- 44. Solinger JA, Kiianitsa K, Heyer WD (2002) Rad54, a Swi2/Snf2-like recombinational repair protein, disassembles Rad51:dsDNA filaments. Mol Cell 10: 1175–1188.
- 45. Simon P, Houston P, Broach J (2002) Directional bias during mating type switching in Saccharomyces is independent of chromosomal architecture. EMBO J 21: 2282–2291.
- 46. Sikorski RS, Hieter P (1989) A system of shuttle vectors and yeast host strains designed for efficient manipulation of DNA in Saccharomyces cerevisiae. Genetics 122: 19–27.