Rad51 determines pathway usage in post-replication repair

Domain: chemistry
Published: 2026-01-10
Editorial outcome: major_revision

Abstract
Stalled replication forks are processed in post-replication repair by homologous recombination, fork regression, and translesion DNA synthesis. However, the regulation of pathway usage is not fully understood. Rad51 protein maintains genomic stability through its roles in recombination and in protecting stalled replication forks. We report isolation of mutations in Saccharomyces cerevisiae Rad51 that shift post-replication repair from recombination to alternate pathways including mutagenic translesion synthesis. Rad51-E135D and Rad51-K305N show near normal in vitro recombination despite changes in their DNA binding profiles, in particular to dsDNA. The mutants lead to a defect in Rad51 recruitment to stalled forks in vivo as well as a defect in the protection of dsDNA from degradation by Dna2-Sgs1 and Exo1 in vitro. Together, the evidence suggests that Rad51 binding to duplex DNA is critical to control pathway usage at stalled replication forks.

Introduction
Homologous recombination (HR) provides a central pathway to maintain genomic stability1. In the context of double-strand break (DSB) repair in mitotically growing cells, HR uses the intact sister chromatid as a template for DNA repair synthesis2. Initially, the DSB is resected in a 5’−3’ direction to generate 3’-OH ending single-stranded DNA tails3. Resection is initiated by the Mre11 nuclease in the Mre11-Rad50-Xrs2 complex, which, in conjunction with Sae2, can also process chemically complex DSB including those covalently bound to proteins. After an initial endonucleolytic incision and 3’−5’ resection by the Mre11 nuclease, two long-range nuclease pathways operate in the 5’−3’ direction including the dsDNA-specific exonuclease Exo1 and the ssDNA nuclease Dna2, which functions in conjunction with the helicase-topoisomerase complex Sgs1-Top3-Rmi1 that generates the ssDNA substrate for Dna2. Resection involves 100s if not 1000s of nucleotides of ssDNA which is bound by the ssDNA-binding protein RPA. Nucleation of Rad51 filaments on RPA-coated ssDNA is achieved by mediator proteins, mainly Rad52 in budding yeast, a role that is usurped by the BRCA2 tumor suppressor protein in mammalian cells4. The resulting Rad51-ssDNA filament catalyzes the signature HR steps of homology search and DNA strand invasion to generate displacement loops (D-loops) and prime repair DNA synthesis in conjunction with the dsDNA motor protein Rad544. In budding yeast, Rad54 is essential to D-loop formation in vivo and in vitro5,6. The HR pathway, including Rad51, Rad52, and Rad54, also supports DNA replication by recovering broken replication forks in a process called break-induced replication and through the repair of replication-associated single-stranded gaps that occur after repriming7–10.

Rad51 is the eukaryotic homolog of bacterial RecA with highly similar functions in HR4,11,12. RecA, however, has at least two additional functions in bacteria. It acts as a cofactor during translesion synthesis (TLS) by DNA Pol V and it triggers the SOS response, a bacterial DNA damage response signaling pathway initiated by RecA-bound ssDNA13,14. Currently, no data suggests that Rad51 plays a similar role in eukaryotic TLS. A function of Rad51 bound to ssDNA in adaption to DNA damage has been postulated, but the mechanism of this potential role in DNA damage signaling has not been defined15. Eukaryotic Rad51 proteins bind ssDNA and dsDNA with similar affinity and only a slight preference for ssDNA16–19. This is in strong contrast to bacterial RecA, which strongly prefers binding to ssDNA20,21. Binding to ssDNA by Rad51 is essential for HR, whereas Rad51 binding to duplex DNA inhibits HR, providing the rationale for proteins that mediate the assembly of Rad51 filaments on ssDNA and the dissociation of Rad51 from dsDNA16,22–26. Rad51 binding to chromosomal duplex DNA may also be limited by chromatin and accessibility. This poses the question, why did high-affinity dsDNA binding evolve in eukaryotic Rad51 and what is its function?

An independent role of some HR proteins in response to replication stress has been identified in vertebrate cells in protecting nascent DNA at replication forks from pathological degradation by MRE11, DNA2 or EXO127–29. Blockage of the replicative helicase or DNA polymerases and regressed forks generate substrates for these nucleases in the form of ssDNA/dsDNA junctions in gaps. In addition, fork reversal generates a DSB at the spur of the chicken foot structure. The HR proteins required for fork protection are BRCA2 and RAD51 but not RAD5427,29, suggesting that RAD51 filament formation but not the entire HR process is required for fork protection. This is consistent with separation-of-function mutants in human RAD51 and BRCA2 that affect differentially the HR and fork protection functions27,30. The mechanisms involved and the exact binding sites for BRCA2 and RAD51 to protect stalled replication forks are not fully understood and recent work suggested an involvement of RAD51 binding to dsDNA31,32. It is unclear how fork protection relates to the response to replication stress in yeast.

Replication stress and fork stalling can be elicited by unusual DNA structures, protein-DNA complexes, limiting nucleotide pools, interference from transcription as well as endogenously and exogenously induced DNA damage29. Stalling at DNA damage sites on the leading and lagging DNA strands can be overcome by direct translesion synthesis (TLS) or downstream repriming leaving a post-replicative gap9,29,33. Such a gap can either be closed by TLS or a homologous recombination (HR)-based mechanism of template switching (TS) using the sister chromatid as a donor. These processes may be spatially and temporally uncoupled from the replication fork34–38. TLS skips the lesion through the engagement of specialized DNA polymerases39. In yeast, a key TLS polymerase is Pol zeta (Polζ), a four-subunit assembly of Pol31, Pol32, Rev7 and the catalytic subunit Rev340. TLS is supported by monoubiquitylation of PCNA at lysine 164 by the Rad6 E2 ubiquitin conjugase in concert with the Rad18 E3 ubiquitin ligase41,42. HR is critical for TS and involves the formation of Rad51-ssDNA filaments that perform homology search and DNA strand exchange in conjunction with its co-factors the Rad51 paralogs Rad55-Rad57 and the dsDNA motor protein Rad549,10,43.

Unique to fork stalling at leading strand lesion is the process of fork regression, another mechanism of TS, where the fork regresses into a four-stranded structure called a chicken foot. Yeast Rad5 was shown to promote fork regression in vitro44. However, in vivo, DNA damage checkpoint signaling suppresses fork regression in yeast and direct electron microscopic analysis detected few regressed forks in wild type cells after the addition of hydroxyurea (HU), an inhibitor of ribonucleotide reductase that leads to fork stalling45. In contrast, in mammalian cells, fork regression is frequently observed in response to various fork stalling agents including HU, methyl methanesulfonate (MMS) and UV, and this process is dependent on the central HR protein RAD5146 as well as the human RAD5 homolog HLTF47. Contradirectional migration of the regressed fork can reestablish a replication fork with the original lesion being avoided. In human cells, fork regression and restart of regressed forks are controlled by a complex ensemble of dsDNA translocases (ZRANB3, SMARCAL1, HLTF) and DNA helicases (FBH1, RECQ1) regulated by poly-ubiquitylation of PCNA29. Alternatively, the chicken foot structure which creates a DSB could be processed by 5’−3’ exonucleases to generate a 3’-ending ssDNA strand available for Rad51 filament formation and DNA strand invasion in front of the fork stall point to restart DNA synthesis48. In sum, TS comprises several mechanistically diverse processes, including fork regression and several forms of gap repair, which can occur at the stalled fork or behind the stalled fork49. How the usage of these different pathways is managed is currently not fully understood. One aspect is PCNA poly-ubiquitylation by the ubiquitin-conjugating complex Ubc13/Mms2 and Rad549, but it is not clear whether all TS pathways are affected. Consistently, inactivation of PCNA poly-ubiquitylation by a ubc13 mutation inhibits TS leading to a strong increase in the use of the TLS pathway50. Interestingly, Rad5 also provides an additional pathway to recruit TLS polymerases to stalled forks51–55. Another regulator in budding yeast appears to be the Srs2 helicase, which has the ability to strip Rad51 from ssDNA, an activity that is counteracted by Rad55-Rad5756–58. Interestingly, Srs2 is recruited to stalled forks by sumoylation of PCNA on lysines 127 and 164 thanks to its C-terminal SUMO binding motif59,60. The genetic observations that sensitivity to fork stalling agents caused by mutations in Rad5, Rad6 and Rad18 that abolish TLS and TS are largely suppressed by mutations in Srs2 are consistent with a model that Srs2 suppresses activation of an HR-dependent TS pathway independent of PCNA ubiquitylation (also termed salvage pathway) when recruited by SUMO-PCNA61–63. In sum, HR-dependent and independent TS as well as TLS pathways operate at regressed forks in yeast, but it remains unclear what regulates their usage64,65.

To define what controls pathway usage at stalled replication forks in the budding yeast Saccharomyces cerevisiae, we isolated mutants in RAD51, which severely affect pathway usage at replication forks stalled by the alkylating agent MMS. The mutants display sensitivity to MMS, and their survival strongly depends on other post-replication repair pathways, namely TLS and fork regression. We directly show that pathway usage at fork stalling lesions is dramatically changed. The Rad51 mutant proteins are poorly recruited to stalled forks in vivo and cannot protect dsDNA from exonucleolytic attack in vitro. Biochemical analysis identified a strong defect in dsDNA binding as the major characteristic that correlated between the in vitro and in vivo phenotypes of the mutants. We suggest that Rad51 binding to dsDNA is a major determinant in post-replication repair and that the properties of the Rad51 filament are critical for pathway usage at stalled forks. While the mutants were proficient in spontaneous and DSB-induced sister chromatid recombination, they displayed reduced interhomolog recombination likely caused by a defect in forming extended filaments on DNA, suggesting different requirements for inter-sister recombination versus a genome-wide homology search.

Results
RAD51 mutant isolation
Rad51 protein functions as the central homology search and DNA strand invasion factor during HR, which involves filament formation on ssDNA. In an independent role, Rad51 protects stalled replication forks, but the mechanisms involved are less well defined. Rad54 is a dsDNA motor protein that is critical for HR but not required for fork protection in mammalian cells27. Using sensitivity to the fork stalling agent MMS, we isolated two mutations in RAD51, each with a single amino acid change, rad51-E135D (referred to as rad51-ED) and rad51-K305N (referred to as rad51-KN), that suppress the MMS-sensitivity of rad51 rad54 double mutants (Fig. 1A). In high copy when borne on a 2µ plasmid, both rad51 mutants substantially suppressed the MMS sensitivity of a rad51Δ rad54ts double mutant at the restrictive temperature. There was also significant but less extensive suppression of the MMS sensitivity of a rad51Δ rad54Δ double mutant (Fig. 1B). We surmise that the RAD51 mutants change Rad51 filament properties that allows use of alternate post-replication repair pathways at stalled forks to bypass the block in HR imparted by the RAD54 mutation.

rad51-ED and rad51-KN cause sensitivity to fork stalling
To study these mutations in a more controlled fashion, we introduced them into the chromosomal RAD51 gene and conducted survival assays through colony formation (Fig. 1C–E). We found the expected high sensitivity of rad51Δ and rad54Δ mutant cells to MMS and the expected epistasis between both mutations (Fig. 1C). The rad51-ED mutant was as sensitive to MMS as the rad51Δ strain, whereas the rad51-KN was only mildly more sensitive than wild type cells, notable only at higher MMS doses (Fig. 1D). Neither chromosomal mutant was able to suppress the MMS sensitivity of a rad54Δ mutation in the colony formation assay (Fig. 1E), suggesting that the overexpression during mutant isolation was a factor. We also note that the colony formation assay (Fig. 1C–E) measures survival whereas the serial dilution assay used in the screen and validation (Fig. 1B) measures a sum of growth and survival. The steady-state levels of the Rad51-ED and Rad51-KN mutant proteins were somewhat elevated compared to wild type Rad51 protein (Supplementary Fig. 1), eliminating the possibility that low protein levels are the cause of the observed phenotypes. We conclude that Rad51-ED and Rad51-KN are mutant proteins with altered functions in the cellular response to MMS.

rad51-KN is proficient in sister chromatid recombination while rad51-ED has a subtle defect
To assess the recombination properties of the newly isolated rad51 mutants, we employed a well-established sister chromatid recombination assay that allows monitoring of spontaneous and DSB-induced recombination between two directly repeated copies of mutant leu2 genes that can give rise to Leu+ recombinants (Fig. 2A). The system uses URA3 as a marker between the repeated leu2 genes to distinguish gene conversion events (Leu+ Ura+) from other events that include single-strand annealing, intra-chromatid crossing over, unequal sister-chromatid exchange, unequal sister-chromatid conversion, and replication slippage and are summarily called pop-out events (Leu+, Ura-). Spontaneous pop-out events in this system are largely mediated by Rad52-dependent strand annealing and are independent of Rad51 (ref. 66 and Fig. 2B left). Neither rad51Δ and rad54Δ nor rad51-ED and rad51-KN mutations affected these events. The Leu+ Ura+ gene conversion events strongly depended on Rad51, Rad52, and Rad54, as expected (Fig. 2B right). The rate for spontaneous Leu+ Ura+ events in rad51-ED was slightly but statistically significantly reduced (p=0.017), while rad51-KN (p=0.084) was not. The mutants did not suppress the recombination defect caused by the rad54Δ mutation (Fig. 2B right). The frequency of both events (Leu+ Ura+, Leu+ Ura-) can be dramatically stimulated by the induction of a DNA double-stranded break using a galactose-controlled version of the HO-endonuclease which cleaves a HO recognition site engineered between the repeated mutant leu2 genes (Fig. 2A). In determining the frequencies of DSB-induced pop-out (Leu+ Ura-) and gene conversion (Leu+ Ura+) events, the results showed that the rad51-ED and rad51-KN mutants exhibited no interference with pop-out events, no defect in DSB-induced gene conversion, and no suppression of the rad54Δ gene conversion defect (Fig. 2C). We conclude that rad51-KN is proficient for mitotic sister chromatid recombination, while rad51-ED has a subtle defect in spontaneous gene conversion but is fully proficient for DSB-induced events.

While rad52Δ showed a significant survival defect after DSB induction by HO, rad51Δ and rad54Δ showed a smaller survival defect (Supplementary Fig. 2), as expected. This reflects the involvement of Rad52 in both pop-out and gene conversion events, whereas pop-out events are unaffected in rad51Δ and rad54Δ cells, allowing substantial DSB repair. The rad51-ED mutant shows slightly better survival after DSB induction than rad51Δ cells which was within the error of the experiment, whereas rad51-KN cells survived significantly better than rad51Δ (Supplementary Fig. 2). This difference in response to a single DSB between both mutants was also recapitulated by their differential sensitivity to 100 Gy of ionizing radiation in serial dilution plate assays, where rad51-ED was IR sensitive but somewhat less so than rad51Δ (Supplementary Fig. 3A, compare rows 1 and 5). rad51-KN showed resistance to IR that was equivalent to wild type cells (Supplementary Fig. 3A, compare rows 5 and 9). We conclude that rad51-ED has a survival defect in response to a single DSB or IR despite being fully proficient in DSB-induced inter-sister HR. This may be related to the signaling role of DNA bound Rad51 affecting adaptation from DNA damage15. IR also induces base damages that may elicit replication fork stalling leading to a possible alternate interpretation that the IR-sensitivity of the mutants is related to fork protection (see below), although we did not further explore the IR phenotype.

rad51-ED and rad51-KN exhibit a defect in inter-homolog recombination
To further define the HR phenotypes of the rad51-ED and rad51-KN mutants we conducted a series of physical and genetic HR assays that require genome-wide homology search. To directly test the ability of the mutant protein to form D-loops in vivo, we first employed the D-loop capture (DLC) assay6. In short, this assay physically monitors D-loop formation between an HO-induced DSB on chromosome V with a homologous donor on chromosome II (Fig. 3A, B) by capturing the D-loop through psoralene-crosslinking and using proximity ligation between two EcoRI sites adjacent to the DSB and donor site to quantify by qPCR the amounts of D-loops. The DLC signal completely depends on Rad51, with an over 100-fold reduction in signal in the rad51Δ mutant6. The rad51-ED and rad51-KN displayed a strong defect in D-loop formation, with rad51-ED being significantly more affected than rad51-KN (Fig. 3C, D). We confirmed this observation in an independent assay measuring BIR between a DSB on chromosome V and a donor site on chromosome XI, which also showed a strong defect in both mutants with rad51-ED being more affected than rad51-KN in BIR efficiency measured through the genetic endpoint (LYS+) and the physical product (Fig. 3E–I). We conclude that rad51-ED and rad51-KN lead to substantial defect in recombination requiring genome-wide homology search, despite their substantial proficiency in sister chromatid recombination.

Rad51-ED and Rad51-KN proteins exhibit a strongly reduced Kd for dsDNA
Rad51-E135 is an invariable residue in a highly conserved region of the eukaryotic Rad51 proteins which is not conserved in the bacterial RecA homologs. Rad51-K305 is conserved in most Rad51 proteins in its charge (K or R), although some Rad51 proteins carry hydrophobic amino acids (L, A) at this position (Supplementary Fig. 5A). Both residues are not directly involved in DNA or nucleotide co-factor binding in the Rad51-ssDNA-ADP-AlF structure67 (Supplementary Fig. 5B), although K305 is part of an alpha helix that ends in R293 which intercalates between bases to enforce the triplet spacing of DNA bound to Rad51 (Supplementary Fig. 5C). To better understand the underlying biochemical defects and mechanisms involved, we purified the native mutant proteins to apparent homogeneity for biochemical analysis (Supplementary Fig. 5D). The mutant proteins purified like the wild type protein and were free of contaminating activities (endo/exonuclease, topoisomerase, phosphatase) that would interfere with the in vitro experiments and their interpretation.

The key biochemical activities of Rad51 are binding and filament formation on DNA, DNA-stimulated ATP hydrolysis, homology search and DNA strand invasion, which underlie its functions in HR and at stalled forks. First, we measured DNA binding of 100 nt and 100 bp substrates using electrophoretic mobility shift assays (EMSA) (Fig. 4). The binding to ssDNA by wild type Rad51 showed a Kd of 497±39 nM for 50 mM NaCl and 578±27 nM for 100 mM NaCl consistent with previous measurements16. Rad51-ED showed identical behavior (Kd=486±12 nM for 50 mM NaCl, 602±11 nM for 100 mM NaCl), whereas Rad51-KN had slightly higher affinity than wild type Rad51 with a Kd of 339±10 nM for 50 mM NaCl and 402±3.3 nM for 100 mM NaCl (Fig. 4A). The mutant complexes showed more of a smear, which is likely related to subtly reduced stability of the protein:ssDNA complexes. Importantly, in contrast to ssDNA, both mutants showed a very significant defect in binding to dsDNA, such that a Kd could not be calculated (Fig. 4B). The defects of both proteins were stronger at elevated salt concentration (100 vs. 50 mM NaCl). The Rad51-ED mutant protein showed consistently a stronger defect than the RAD51-KN protein. We extended these results using longer DNA substrate (ssDNA: ΦX174 viral DNA 5386 nt; dsDNA: ΦX174 RF1 5386 bp) by conducting salt midpoint titrations (Supplementary Fig. S6). The salt titration curves allowed the calculation of salt titration midpoints (STM; Supplementary Fig. 6E) as the salt concentration, at which 50% of the protein-DNA complexes are formed. A lower STM signals lower affinity for DNA. Under no salt conditions (0 mM NaCl), both mutant proteins showed no defect in binding to ssDNA and a small but significant defect for dsDNA (Supplementary Fig. 6C, D). The defect for Rad51-ED was again significantly stronger than with Rad51-KN for both substrates. In fact, at physiological ionic strength (~120 mM NaCl68), dsDNA binding was largely suppressed, especially for Rad51-ED. In contrast to the 100 nt substrate, the mutants showed a defect in ssDNA binding with the longer substrate, which is explained by the formation of some duplex regions in the more complex ssDNA substrate and possibly by reduced stability of ssDNA binding.

To further explore the DNA binding defect, we examined the Rad51 filaments on 1000 bp dsDNA and 700 nt ssDNA substrates by electron microscopy (EM). Unexpectedly, we saw a significant defect of both mutants to form extended nucleoprotein filaments on both substrates (Supplementary Fig. 7). As in the EMSA and STM assays, the defect in dsDNA binding was more pronounced than with ssDNA (compare Supplementary Fig. 7D with H). The defect with rad51-KN was more pronounced than with Rad51-ED. This defect correlated with severely diminished Rad51 focus formation in response to MMS as observed by immunofluorescence (Supplementary Fig. 8). It is unclear whether the MMS-induced Rad51 foci represent Rad51 bound to ssDNA, dsDNA or both, but the results suggest that the filament length of both mutants in cells is below the detection threshold in our experiment.

Rad51-ED and Rad51-KN proteins have altered ATPase profiles
The ATPase of Rad51 is strongly dependent on DNA binding, and an ATPase defect could be a consequence of an underlying DNA binding defect. We measured the ATPase activity of the mutant proteins along with wild type Rad51 at 200 mM NaCl to accentuate potential differences. The ATPase activities of the mutant proteins were diminished with both ssDNA and dsDNA co-factors. The defect of the Rad51-ED mutant protein was significantly more pronounced than that of the Rad51-KN mutant (Fig. 5A, B).

Rad51-ED and Rad51-KN proteins are proficient in D-loop formation
The in vitro recombination activity of the mutant proteins was assessed in the classic D-loop assay using a tailed 100mer with a 5’ 25 bp duplex region and a 2686 bp negatively supercoiled duplex target DNA in reconstituted reactions with Rad51, Rad54 and RPA conducted as a time course at the same salt concentrations (50 mM Supplementary Fig. 9A–C; 100 mM NaCl Fig. 5C–E) as the EMSA assays of Fig. 4. Rad51 wild type protein showed the expected behavior of forming D-loops peaking at 5 min that were processed over time (Fig. 5E). The mutant Rad51-ED and Rad51-KN proteins were as effective as wild type Rad51 in forming D-loop in vitro and the D-loops formed showed somewhat higher D-loop instability over time (Fig. 5E). This is consistent with a functional defect in binding dsDNA, as after D-loop formation Rad51 is bound to dsDNA, from where it is displaced for D-reversal by Rad5469. D-loop formation by the Rad51-ED and Rad51-KN proteins was fully dependent on Rad54, identical to wild type Rad51 (Supplementary Fig. 9D), as expected from the genetic experiments (Fig. 2). We conclude that the Rad51-ED and Rad51-KN mutant proteins are proficient in D-loop formation, consistent with the in vivo inter-sister recombination data (Fig. 2).

In sum, the biochemical analysis of the Rad51-ED and Rad51-KN proteins showed that they are proficient in forming D-loops in vitro and largely proficient in binding ssDNA. The major defect is in binding to dsDNA, which we further explored below.

Rad51-ED and Rad51-KN fail to protect dsDNA from Exo1 or Dna2
Besides its key function in HR by forming the presynaptic filament on ssDNA for homology search and DNA strand invasion, Rad51 protects nascent DNA at stalled DNA replication forks from unscheduled degradation by nucleases including Exo1 and Dna2. The fork protection function of certain HR proteins was first noticed in human cells, particularly in BRCA1/2-deficient backgrounds27–29. The DNA protection function of RAD51 is distinct from its role in HR, as indicated by the absence of a requirement for RAD5427. However, the fork protection function of RAD51 involves its binding to dsDNA31,32.

To define the capacity of the yeast Rad51 variants in DNA protection, we employed in vitro assays with 100 bp-long dsDNA (Fig. 6A). We first used yeast Exo1, one of the nucleases involved in long-range end resection. Without Rad51, Exo1 degraded ~90% of the DNA substrate. Wild type Rad51 inhibited DNA degradation by Exo1, while the two Rad51 mutants, Rad51-KN and Rad51-ED, did not protect DNA (Fig. 6B, C). Similar results were obtained when we used the Sgs1-Dna2 helicase-nuclease pair. While wild type Rad51 efficiently prevented DNA degradation, Rad51-KN and Rad51-ED were much less effective (Fig. 6C, D). Mechanistically, Rad51 prevents DNA unwinding by Sgs1, which prevents the generation of ssDNA for the Dna2 nuclease (Supplementary Fig. 10). Rad51-KN and Rad51-ED did not prevent unwinding (Supplementary Fig. 10B, C). Together, our reconstitution experiments revealed that Rad51-KN and Rad51-ED are impaired in dsDNA protection.

Synergy with defect in post-replication repair
The biochemical analysis revealed a significant dsDNA binding defect in the mutants and a defect in protecting dsDNA from nuclease degradation. These properties are associated with replication fork stabilization by Rad51 and prompted us to explore the genetic interactions of rad51-ED and rad51-KN with mutations affecting various aspects of post-replication repair. We first tested a deletion mutation of REV3, which by itself shows no sensitivity to MMS at the low levels (0.0033%) used for the serial dilution assays in this experiment (Fig. 7A row 3). This contrasts with the significant sensitivity of HR-defective mutants (rad54Δ row 2, rad51Δ row 5, rad51Δ rad54Δ row 6). An HR defect displayed strong synergy with a defect in REV3 leading to extreme MMS sensitivity and also affecting growth/survival on YPD growth media without MMS (rad54Δ rev3Δ row 4, rad51Δ rev3Δ row 7, rad51Δ rad54Δ rev3Δ row 8). These results were expected and show that HR and TLS represent parallel competing pathways to process MMS-induced DNA damage, and the single mutant phenotype suggests that HR plays a more prominent role in wild type cells than TLS.

rad51-ED showed the MMS sensitivity expected from the previous quantitative survival assays, we confirmed the relative mild MMS survival defect of mms2Δ cells compared to HR-defective cells (rad51Δ, rad54Δ). Analysis of the rad51Δ mms2Δ and rad54Δ mms2Δ double mutants revealed a non-epistatic relationship between an HR defect and a loss of Mms2 function (Fig. 7C). These results show that in wild type cells HR makes a stronger contribution to MMS survival than pathway(s) dependent on PCNA poly-ubiquitylation. Moreover, these results suggest that Mms2 controls a strand exchange-independent template switching mechanism, in analogy to mammalian cells70. This mechanism is unlikely to be HR-mediated gap repair that is Rad51 dependent; alternatively, it could be fork regression operating in yeast in a Rad51-independent manner. rad51-ED showed the MMS sensitivity expected from the previous experiments (Fig. 1D) that rivals that of a complete HR defect (Fig. 7A compare rad54Δ row 2, rad51Δ row 5, rad51Δ rad54Δ row 6 with rad51-ED row 9). The rad51-ED rev3Δ double mutant showed extreme MMS sensitivity and a growth/survival defect on YPD, like a complete HR defect (Fig. 7A compare rad51-ED row 9 with rad51-ED rev3Δ row 11). The addition of a rad54Δ mutation to these strains (rows 10, 12) did not further increase MMS sensitivity or the growth/survival defect on YPD. The low MMS concentration (0.0033%) was needed to compare the widely varying sensitivity of all strains on a single plate. rad51-KN showed little to no MMS sensitivity in the serial dilution assays (Fig. 7B row 9), as expected from Fig. 1, where this mutant showed MMS sensitivity only at high MMS concentrations. Remarkably, rad51-KN showed synergy with rev3Δ (Fig. 7 row 11), although none of the single mutants were measurably sensitive at the concentration used (rows 3, 9). These results show that MMS-stalled forks in rad51-ED cells are preferentially processed by TLS similar to the situation in cells with a complete HR defect, although this mutant is capable of normal HR. The rad51-KN mutant shows similar but less pronounced phenotypes.

Next, we tested the role of Mms2, which is required for poly-ubiquitylation of PCNA at K16441 and template switching. In quantitative survival assays, we confirmed the relative mild MMS survival defect of mms2Δ cells compared to HR-defective cells (rad51Δ, rad54Δ). Analysis of the rad51Δ mms2Δ and rad54Δ mms2Δ double mutants revealed a non-epistatic relationship between an HR defect and a loss of Mms2 function (Fig. 7C). These results show that in wild type cells HR makes a stronger contribution to MMS survival than pathway(s) dependent on PCNA poly-ubiquitylation. Moreover, these results suggest that Mms2 controls a strand exchange-independent template switching mechanism, in analogy to mammalian cells70. This mechanism is unlikely to be HR-mediated gap repair that is Rad51 dependent; alternatively, it could be fork regression operating in yeast in a Rad51-independent manner. rad51-ED showed the MMS sensitivity expected from the previous experiments (Fig. 1D) that rivals that of a complete HR defect. rad51-ED showed the same increase in MMS sensitivity in the double mutant with mms2Δ as the rad51Δ strain that remained unchanged by adding a rad54Δ mutation (Fig. 7D). rad51-KN showed little MMS sensitivity as expected from Fig. 1, which was similar to the MMS sensitivity of the mms2Δ single mutant (Fig. 7E). Remarkably, rad51-KN showed strong synergy with mms2Δ that was further enhanced by fully eliminating HR through a rad54Δ mutation (Fig. 7F). We conclude that MMS-stalled forks in rad51-ED cells have a higher propensity to be processed by a Mms2-dependent pathway (hypothetically fork regression) similar to the situation in cells with a complete HR defect. The rad51-KN mutant shows a less accentuated phenotype.

Furthermore, we tested a PCNA mutant (encoded by the POL30 gene in budding yeast), which eliminated the K127 and K164 acceptor sites (pol30-K127R, K169R abbreviated as pol30-KKRR) for ubiquitylation and sumoylation to obstruct Srs2-mediated anti-recombination, TLS, and TS (likely fork regression) pathways. This double mutant displayed only very mild MMS sensitivity at the low (0.0033%) concentration used (Fig. 7F row 3), as expected41. This result suggests that under these conditions the salvage HR is the predominant pathway. This notion is confirmed by the extreme synergy in MMS sensitivity with an HR defect as well as the strong growth/survival defect on YPD (Fig. 7F, compare pol30-KKRR row 3 with rad54Δ pol30-KKRR row 4 or rad51Δ pol30-KKRR row 7 or rad51Δ rad54Δ pol30-KKRR row 8). rad51-ED mimicked again the behavior of a complete HR defect (Fig. 7F; compare rad51-ED row 9 with rad51-ED pol30-KKRR row 11) including the remarkable growth/survival defect on YPD lacking MMS. Consistent with the genetic interaction analysis involving rev3Δ and mms2Δ, the rad51-KN mutant shows a less pronounced phenotype which was enhanced by a complete HR defect (Fig. 7G, compare rad51-KN row 9 with rad51-KN pol30-KKRR row 11 and with rad51-KN rad54Δ pol30-KKRR row 12). We conclude that in rad51-ED and rad51-KN cells, HR-independent template switching and TLS process a larger fraction of MMS-stalled forks than in wild type cells.

Finally, we used a subtle mutation in RAD5, rad5-G535R, which by itself has little to no phenotype71 (Supplementary Fig. 3C). Rad5 promotes PCNA polyubiquitylation and template-switching and also plays a role in recruiting TLS polymerases, affecting two major post-replication repair pathways. In consequence, the deletion of the RAD5 gene causes one of the most pronounced MMS sensitivities caused by a single mutant in budding yeast, requiring a rad5 allele with a more subtle phenotype. In serial dilution assays, we noticed some suppression of the MMS sensitivity of rad54Δ by rad51-ED and rad51-KN which depended on the wild type RAD5 gene (Supplementary Fig. 3B compare rad51-ED rad54Δ RAD5 row 4 with rad51-ED rad54Δ rad5-G535R row 3 and rad51-KN rad54Δ RAD5 row 10 with rad51-KN rad54Δ rad5-G535R row 11). The discrepancy between the results from the serial dilution (Supplementary Fig. 3) and colony formation assays regarding rad54 suppression (Fig. 1) is explained by the fact that serial dilution assays measure a sum of growth and survival, whereas colony formation measures survival only. These results reinforce the conclusion that rad51-ED and rad51-KN rely more heavily on TLS and HR-independent template switching to process stalled replication forks. The rad5-G535R mutation did not affect the IR sensitivity of rad51-ED or rad51-KN (Supplementary Fig. 3A) and has been described as wild type for IR resistance (https://wiki.yeastgenome.org/index.php/CommunityW303.html).

From this genetic analysis with four genes affecting single or multiple post-replication repair pathways, we conclude that the balance between post-replication repair pathways is significantly altered in the rad51-ED and rad51-KN mutants.

rad51-ED and rad51-KN display a strong mutator phenotype
HR-defective cells such as rad51Δ or rad54Δ mutants display a considerable mutator phenotype likely caused by spontaneous DNA damage that, instead of being repaired or tolerated by high fidelity HR, is processed by the error-prone TLS pathway, causing an increase in spontaneous mutagenesis. We used the forward mutagenesis CAN1 assay to determine the rate of spontaneous mutations and saw the expected 9-10-fold increase in rad51Δ, rad54Δ single and double mutants (Fig. 8). It was previously reported that this increase largely depends on the central TLS polymerase component Rev372. To our surprise, rad51-ED and rad51-KN showed a significant increase of 9-fold and 6-fold, respectively, in their mutation rate. These values are not significantly different from those of the rad51Δ, rad54Δ single and double mutants. Both mutants did not suppress the mutator phenotype of the rad54Δ mutant (Fig. 8). Instead, rad51 ED slightly elevated the mutation rate in rad54Δ cells from 9-fold in the single mutant to 24-fold in the rad51-ED rad54Δ double mutant, which may suggest that the rad51-ED mutant favors mutagenic events that are suppressed by RAD54. We conclude that rad51-ED and rad51-KN show a significant mutator phenotype due increased engagement of TLS at stalled replication forks.

Dramatic shift from template switching to TLS
To investigate the effect of rad51-ED and rad51-KN mutants on lesion bypass, we took advantage of the iDamage approach (Fig. 9A)50. In short, a plasmid containing the single lesion of interest is stably inserted at a specific locus of the yeast genome by means of the Cre recombinase and modified lox sites. Cre-mediated recombination between the mutant lox71 (on the vector) and the mutant lox66 (in the genome) lead to a double mutant lox site that is no longer recognized by the Cre recombinase, avoiding excision of the lesion-containing vector. Lesion bypass is monitored by counting blue and white colonies, as the lesion is inserted into the lacZ reporter gene (Fig. 9A). While this system does not allow to study MMS-induced DNA damage, we investigated the bypass of two common lesions: the thymine-thymine pyrimidine (6–4) pyrimidone photoproduct [TT(6–4)], and the N2-dG-Acetylaminofluorene (N2dG-AAF). All experiments were performed in a parental strain where NER and mismatch repair are inactivated to prevent the repair of the lesion and the DNA loop used as a strand marker, respectively. We measured a level of TLS of ~5% and ~17% for the TT(6–4) and the G-AAF lesions in this parental strain (Fig. 9B). The inactivation of rad51 led to a decrease in the level of the template-switching pathways (HR-mediated gap repair, fork regression) and a concomitant increase in the level of TLS (Fig. 9B). In rad51Δ cells about half of the template switching events remained, suggesting a significant contribution of Rad51-independent template switching to the total number of tolerance events. This reflects the competition between template switching and TLS: when a template switching pathway is inhibited, TLS is favored. Upon introduction of the same lesion in the rad51-ED and rad51-KN mutants, we observed the same decrease in the use of template switching and the same increase in TLS (Fig. 9B).

Defects in binding to stalled forks and forming sister chromatid junctions
The recombinational processing of MMS-induced DNA lesions is associated with the formation of Rad51-dependent sister-chromatid junctions (SCJs). The previous genetic results suggested that there is a defect in rad51-ED and rad51-KN cells to process MMS-stalled replication forks. The accumulation of these recombination intermediates can be detected as X-shaped structures by 2D electrophoresis in sgs1Δ cells, which are defective in their dissolution73. To address the functionality of Rad51-ED and Rad51-KN in the recombinational response to MMS, we measured the accumulation of SCJs in cells synchronized in G1 and released into S phase in the presence of MMS (Fig. 10A). Rad51-ED and Rad51-KN are affected in their ability to promote SCJs as inferred from the behavior of the mutants. sgs1Δ rad51-KN cells displayed a 1.7-fold reduction in the amount of SCJs along the analyzed times that was mostly due to a delay in the accumulation of SCJs as compared to the wild type. Remarkably, sgs1Δ rad51-ED was severely affected in the accumulation of SCJs and reduced to ~20% of the wild type levels. We conclude that the rad51-ED and, to a lesser degree, rad51-KN mutants are defective in HR at MMS-stalled replication forks leading to increased usage of alternate post-replication repair pathways. Alternatively, it is possible that in these mutants such structures are less stable or more likely to be degraded.

The pronounced MMS sensitivity (Fig. 1, Supplementary Fig. 1) and a severe defect in SCJs formation (Fig. 10A) of rad51-ED cells contrasts with its near wild type behavior in direct genetic assays of HR (Fig. 2). A mechanistic difference between the roles of Rad51 in HR at DSBs and MMS-induced ssDNA is that the role of Rad51 at ssDNA lesions is coupled to replication35, whereas the role of Rad51 at DSBs is replication-independent74,75. Thus, Rad51-ED might be specifically affected in the binding to MMS-induced lesions during S-phase. To address this, we constructed chimeras of Rad51 with the MNaseI (Rad51-MN) and followed their binding to DNA lesions by Chromatin Endogenous Cleavage (ChEC) analysis. In this approach, the induction of the nuclease activity of the chimera by treating cells with Ca2+ will only generate a detectable cut in the DNA if the repair protein is targeted to a lesion that is not a DSB. Thus, we can infer Rad51 binding to DNA if DNA is digested upon Ca2+ treatment35. In the absence of MMS, most DNA remained as a single, high molecular DNA band on top of the gel; in the presence of MMS both Rad51-MN and Rad51-KN-MN generated a smear below the top band, whereas Rad51-ED-MN hardly digested the top band (Fig. 10B). These results were fully reproducible in two additional independent experiments (Supplementary Fig. 11A). We conclude that Rad51-ED is severely impaired in binding to MMS-stalled replication forks or that its DNA binding substrate is not formed or degraded. Control experiments showed that the Rad51-ED-MN fusion did not aggravate the DNA damage sensitivity of the rad51-ED mutant, suggesting that the observed defect is not a consequence of the MN fusion (Supplementary Fig. 11B). The Rad51-KN-MN fusion sensitized the rad51-KN mutant (Supplementary Fig. 11B), although the ChEC assays showed no defect for this mutant.

No effect on interaction with Rad55-Rad57
Rad51 paralog complex Rad55-Rad57 functions in the nucleation of Rad51 filaments on RPA-coated ssDNA76. It also stabilizes the Rad51-ssDNA filament against dissociation by the Srs2 helicase that has the ability to strip Rad51 from ssDNA, therefore acting as an anti-recombinase56–58. Detailed genetic analysis showed a strong similarity between the phenotypes of cells lacking Rad55 and the rad51-ED or rad51-KN mutants described here in their dependence on the TLS polymerase Rev377. This opened the possible interpretation that the rad51-ED and rad51-KN mutants affected the interaction of Rad51 with the Rad55-Rad57 complex. Using purified wild-type Rad51 as well as Rad51-ED and RAD51-KN proteins in pull-down experiments with the purified Rad55-Rad57 complex, we could demonstrate that both mutants interacted normally with the Rad51 paralog complex (Supplementary Fig. 12). We conclude that the phenotypes of the rad51-ED and rad51-KN mutants is not caused by a defect in interacting with Rad55-Rad57.

Methods
Yeast strains and growth conditions
Standard techniques for yeast growth and genetic manipulation were used96. All yeast strains used for genetic experiments are in the W303-1A background and WT RAD5 (Supplementary Table 1), except for some strains used in Supplementary Fig. 1C which are rad5-G535R as noted. See Supplementary Table 2 for all plasmids used. See Supplementary Table 3 for all oligonucleotides and their sequences.

Rad51 mutant isolation and genomic integration
To screen for rad51 separation-of-function mutants, a random pool was created by mutagenic PCR using the GeneMorph II EZClone Domain Mutagenesis Kit that employs two error prone polymerases to achieve balanced mutations in all four bases. The RAD51 open reading frame is 1.2 kb in length; the ideal pool of mutants would have one to a few mutations per gene. Mutagenic PCR conditions were empirically optimized to obtain a mutation rate of ~1 mutation per kb, by varying the amount of template DNA and reducing the number of PCR cycles using olWDH632 and olWDH633 which are 400 nt upstream and downstream of the RAD51 open reading frame, respectively, and plasmid pWDH957, which contains the wild type RAD51 gene with 1000 bp upstream and downstream of the RAD51 open reading frame cloned as a 3.2 kb HindIII-SacI fragment into Yep351 (pWDH958) using olWDH830 and olWDH1355. The pool used averaged two mutations per gene. The screen was conducted in a rad51Δ rad54ts strain (WDHY2546) to take advantage of the conditional phenotype. The rad54ts allele was determined to be caused by a single amino acid change (C692Y), which lead to loss of protein at the restrictive temperature. The mutagenized RAD51 pool was co-transformed with pWDH957 digested with StuI and NruI to cut out almost the entire RAD51 ORF from −22 to 47 bp upstream the stop codon for in vivo recombination to form functional circular YEp351 plasmids containing a mutagenized RAD51 gene. Approximately 2000 transformants were screened. Of these, 40 candidates exhibited improved viability when challenged with MMS compared to the rad51Δ rad54ts strain at 37°C. The plasmid dependence of the phenotype was confirmed. Plasmids were recovered in E. coli, the RAD51 sequence was established, and the phenotype re-confirmed by transformation of a host strain that has never been exposed to MMS.

The mutations rad51-ED and rad51-KN were transferred to the native RAD51 locus of WDHY2217 that contained a complete deletion of the URA3 gene to preclude reversion events using the PCR-based allele replacement method of Erdeniz et al.97 with the RAD51 primers olWDH1351 and 1353 as well as the K. lactis URA3 primers olWDH1357 and 1358. Correct integration and the integrity of the integrated sequences were confirmed by amplifying the RAD51 locus and sequencing the entire gene on both strands.

Serial dilution assays
The serial dilution spot assays were performed on solid YPD or YPD with the addition of MMS at the dilutions specified. To assay for sensitivity to ionizing radiation (IR), YPD plates were spotted with cells and allowed to dry before being exposed to IR (Cesium-137 source). Cells were spotted at 5-fold dilutions starting at approximately 2×107 cells/ml (OD600=1). The plates were incubated at 30°C and photographed daily for up to 4 days.

Immunoblots
Lysates were prepared by trichloroacetic acid (TCA) precipitations from cells grown to early log phase. Cells were re-suspended in 20% TCA before cell disruption utilizing a FastPrep machine (FP120, Bio101, Savant) on setting “4.5”, four times for 45 seconds. Beads were washed with 5% TCA. Precipitated protein was collected at 21,100×g in an Eppendorf centrifuge for 10 min, resuspended in 2x Laemmli sample buffer98, and neutralized with Tris-base before analysis by gel electrophoresis. Immunoblots were conducted as described92. Lysate from 0.5 OD600 units of cells were loaded and separated on a 10% SDS-PAGE gel before transfer to PVDF membrane. The membrane was blocked in 5% skim milk dissolved in TBST (10% Tris pH 7.5, 150 mM NaCl, 0.1% Tween-20) and incubated with either anti-Rad51 (1:1000, sc-133089, Santa Cruz Biotechnology) or anti-GAPDH (1:5000, GA1R, Thermo Scientific) antibodies. Membranes were developed by chemiluminescence (Clarity, Bio-Rad) and visualized using a GE Amersham Imager 600. All bands were quantified through densitometry using ImageLab (Bio-Rad).

Immunofluorescence
Rad51 foci were detected by immunofluorescence as described99. In short, log-phase cells (5 ODU) were crosslinked in 2% formaldehyde (37% stock) for 30 min at 30°C shaking. Cells were harvested at 800xg for 5 min in a 15 ml conical tube and rinsed once in 1.0 ml 1M potassium phosphate pH 6.5. The supernatant was removed, and cells were resuspended in 1.0 ml of solution P (0.1 M potassium phosphate pH 6.5, 1.2 M sorbitol), transferred to a microfuge tube and incubated with 15 µl zymolyase (10 mg/ml) in solution P and 5 µl β-mercaptoethanol for 30 min at 30°C shaking. While the cells were incubating, slides were prepared by overlaying 200 µl 0.3% poly-lysine (in ddH2O) for 15–30 min followed by a rinse with ddH2O. Spheroblasts were harvested at 800xg in a microfuge for 3 min, resuspended in 1–2.0 ml of solution P at ~0.5–1.0 ODU/ml and a minimum of 250 µl cells were incubated on each slide for 15–30 min. Cells were fixed by adding 100% ice cold MeOH dropwise onto the cell suspension. Slides were placed into a Coplin jar containing ice cold MeOH and incubate at −20°C for a minimum of 20 min. Slides were air dried and rehydrated in 1x PBS (phosphate-buffered saline: 137 mM NaCl, 2.7 mM KCl, 8 mM Na2HPO4, and 2 mM KH2PO4.) and then overlaid with 200 µL anti-Rad51 antiserum (1:100)92 in 3% BSA in PBS and incubated for 1 h at 37°C. The slides were washed twice in PBS/0.1% Tween 20 for 3 min each then 1x in PBS alone for 3 min while shaking. The slides were overlain with 200 µL goat anti-rabbit antibody conjugated to Alexa 488 (1:1,000; AbCam ab150077) in 3% BSA/PBS and incubated for 30 min at 37°C followed by the washing scheme outlined above. The slides were then stained for 1 min in DAPI (1 ug/mL))/1x PBS rinsed in ddH2O and air-dried. The slides were mounted in DABCO/20 mM Tris and imaged on a Zeiss Axioskop 2 at 100x magnification.

Direct-repeats sister-chromatid recombination assay
Single colonies were grown in synthetic complete (SC) media lacking uracil with raffinose as a carbon source for 24 h at 30°C. To determine spontaneous recombination rates, appropriate dilutions were plated on YPD media to determine viability, and on SC-LEU, and SC-URA-LEU to measure recombinants. 2% galactose was added to the cultures to induce expression of the HO endonuclease, which was expressed for 2 h before the cells were collected and plated to determine DSB-induced recombination frequencies. After 3 days of growth at 30°C, colonies were counted from at least 11 independent cultures per genotype by the method of the median100.

D-loop capture assay (DLC) assay
For DLC experiments, all strains were in the W303 RAD5 background. They contain a copy of the GAL1/10-driven HO endonuclease gene at the TRP1 locus on chromosome IV (Chr. IV). A point mutation inactivates the HO cut site at the mating-type locus (MAT) on Chr. III (MATa-inc). The DSB-inducible construct contains the 117-bp HO cut site, a 2086-bp-long homology A sequence (+4 to +2090 of the LYS2 gene), and a 327-bp fragment of the PhiX174 genome flanked by multiple restriction sites. The DLC assay was conducted following established protocols with slight adjustments101–103. Specifically, zymolyase lysed cells were immediately subjected to restriction digestion, ligation, and DNA purification steps following hybridization with oligonucleotides104. The control experiments monitoring DSB formation, ligation efficiency, and a normalization locus are shown in Supplementary Fig. 4.

Determination of BIR frequency
BIR frequency determination followed the method outlined in ref. 105. In brief, cells were cultivated to exponential phase in YEP-lactate medium and then plated on YPD plates. After 3 days, colonies were counted and subsequently replicated onto synthetic complete medium lacking lysine (LYS2 drop-out) plates or YPD plates containing geneticin (G418). Cell viability post-HO induction was calculated by dividing the number of colonies on YP galactose plates by those on YP glucose plates. The percentage of cells repairing via BIR was determined by dividing the number of cells on LYS2 drop-out plates by the number on YP galactose plates, normalized to the number on YPD. BIR frequencies were determined three times for each strain.

Detection of physical BIR products
Cells were cultured in YEP-lactate medium, followed by the addition of 2% galactose to induce HO endonuclease expression. Genomic DNA (25 ng) was then subjected to PCR amplification using Phusion High-Fidelity DNA Polymerase with the following cycling conditions: initial denaturation at 98°C for 30 s, followed by 25 cycles of denaturation at 98°C for 10 s, and annealing/extension at 72°C for 150 s in a 25 µl reaction volume. BIR product detection utilized P1 and P2 primers, while HO cut detection employed D1 and D2 primers. Normalization of P1-P2 and D1-D2 products was achieved using C1 and C2 primers, as described previously105,106.

Analysis of sister chromatid junctions (SCJs)
Yeast cells were grown at 30°C in a supplemented minimal medium (SMM). For G1 synchronization, cells were grown to mid-log phase and α-factor was added twice at 60 min intervals at 2 µg/ml (BAR1 strains). Then, cells were washed three times and released into fresh medium with 50 µg/ml pronase in the absence or presence of MMS at the indicated concentrations. SCJs (X-shaped molecules) were analyzed by 2D-gel electrophoresis from cells arrested with sodium azide (0.1% final concentration) and cooled down on ice as reported107. Briefly, total DNA was isolated with the G2/CTAB protocol, digested with EcoRV and HindIII, resolved by neutral/neutral two-dimensional gel electrophoresis, blotted to HybondTM-XL membranes, and analyzed by hybridization with the 32P-labeled A probe. Signals were quantified in a Fuji FLA5100 with the ImageGauge analysis program.

In vivo ChEC analysis
Chromatin endogenous cleavage (ChEC) of RAD51-MN, rad51-KN-MN and rad51-ED-MN cells was performed as reported35 from cultures grown at 30°C in SMM with or without 0.05% MMS for 2 h and arrested with sodium azide (0.1% final concentration). For cleavage induction, digitonin-permeabilized cells were incubated with 2 mM CaCl2 at 30°C under gentle agitation. Total DNA was isolated and resolved into 0.8% TAE 1× agarose gels. Gels were scanned in a Fuji FLA5100, and the signal profile quantified using ImageGauge. The area of the DNA digestion profile was equalized to eliminate DNA loading differences.

Direct post-replication repair analysis in cells
All strains used for the iDamage assay are derivatives of the EMY74.7 strain108. In order to study tolerance events, all strains are deficient in repair mechanisms: nucleotide excision repair (rad14) and mismatch repair (msh2). Gene disruptions were achieved using PCR-mediated seamless gene deletion109 or URA blaster110 techniques. Rad51 mutations were introduced using CRISPR/Cas9111. Integration of plasmids carrying the (6–4) TT and N2dG-AAF lesions (or control plasmids without lesion) was performed as previously described50. Lesions are inserted either on the lagging or leading strand. The lesion is located in the lacZ gene, allowing to score TS events as blue colonies. The non-damaged strand contains a +2 frameshift inactivating the lacZ gene, serving as a genetic marker for strand discrimination. Lesion tolerance rates were calculated as the relative integration efficiencies of damaged vs. non-damaged vectors normalized by the transformation efficiency of a control plasmid in the same experiment. DA events are calculated by subtracting TLS events from the total lesion tolerance events. All experiments were performed at least in triplicate. For each experiment, between 2000 and 4000 colonies were counted. Since we have not observed any difference in lesion tolerance pathways between lagging and leading strands, we plotted in the graphs average value of pooled data for leading and lagging strands. Graphs and statistical analysis were done using GraphPad Prism applying unpaired t-test. Bars represent the mean value ± sd.

Protein purifications
Rad51 mutant and wild type proteins were purified to homogeneity as described112,113. The host Saccharomyces cerevisiae strain (WDHY1611; Supplementary Table 1) had a chromosomal deletion of the RAD51 gene to avoid contaminating mutant protein preparation by wild type Rad51 protein. In brief, the mutant and wild type proteins (native, untagged) were purified using a four-column purification strategy including: Q-Sepharose, Cibacron Blue, Hydroxyapatite, and MonoQ. Rad54114 and RPA115 were purified as described. GST-Rad55-His6-Rad57 were purified as described (Liu et al. 2011b) and dialyzed into storage buffer containing 20 mM Tris-HCl pH 7.5, 0.2 M NaCl, 0.1 mM EDTA, 1 mM DTT and 50% glycerol. Sgs1 was expressed in Spodoptera frugiperda (Sf9) cells using the pFB-MBP-Sgs1 vector and purified by amylose and NiNTA affinity purification as previously described116. The N-terminal MBP tag was removed after the amylose purification step using PreScission Protease. Yeast Exo1 was purified from Sf9 cells with pFB-Exo1-FLAG by FLAG affinity and HiTrap SP HP (Cytiva) ion exchange chromatography as described previously117. Dna2 was expressed in the S. cerevisiae strain WDH668 from the pGAL18 Flag-HA-Dnawt-His: vector and purified by NiNTA and FLAG affinity chromatography as described118,119. Yeast RPA was expressed using the p11d–tRPA vector (a kind gift from M. Wold, University of Iowa) in BL21 (DE3)pLysS cells and purified on a HiTrap Blue column (Cytiva) followed by desalting with a HiTrap Desalting column (Cytiva) and finally with a HiTrap Q column (Cytiva)120.

Biochemical assays
ATPase assay: The ATP analysis was carried out using thin layer chromatography (TLC), as described121. 3 µM Rad51 was pre-incubated with 9 µM (nt or bp) DNA (phiX174 ss- or dsDNA), at 30°C, in 20 mM Tris HCl (pH 7.5), 10 mM MgCl2, 0.1 mg/ml BSA, 1 mM DTT, and 200 mM NaCl, then 0.05 mM ATP (1:100 nt radioactive) was added to initiate the reaction and start the time course. One µl aliquots were then spotted onto cellulose polyethyleneimine TLC plates. The plates were air dried and developed in 1 M formic acid and 0.5 M LiCl. The amount of ATP hydrolyzed was analyzed by PhosphorImager and quantified using ImageQuant software (Molecular Dynamics, Inc., Sunnyvale CA, USA).

DNA binding assays: DNA binding assays were performed as described122 and conducted as salt titration to assay the capability of Rad51 to bind DNA (ss- or dsDNA) in the presence of increasing concentrations of salt. DNA (10 µM nt/bp, ΦX174), salt (0–600 mM NaCl), and Rad51 (3.33 µM), were incubated for 15 min at 30°C in the following buffer: 20 mM TEA, 1 mM DTT, 25 µg/ml BSA, 4 mM magnesium acetate, and 2.5 mM ATP. The reactions were stopped by the addition of glutaraldehyde (0.25%) and incubated for an additional 15 min at 30°C, before being analyzed on 0.8% agarose gels in TAE buffer (40 mM Tris base, 20 mM acetic acid, 1 mM EDTA) 3.75 V/cm for 100 min. The dried gels were analyzed by PhosphoImager and quantified using ImageQuant software (Molecular Dynamics, Inc., Sunnyvale CA, USA). The data were plotted using Prism GraphPad software which also determined the salt-titration midpoint. Additional DNA binding assays were conducted as protein titrations. DNA (3 µM nt/bp, olWDH2086 ssDNA or olWDH2086 annealed to olWDH2184 dsDNA) and Rad51 (250, 500, 1000, or 2000 nM) were incubated for 30 min at 30°C in the following buffer: 35 mM Tris-acetate (pH 7.5), 50 or 100 mM NaCl, 7 mM magnesium acetate, 2 mM ATP, 1 mM DTT, and 0.25 mg/mL BSA. The reactions were analyzed on 1% 1x Tris-acetate agarose gels (Lonza) supplemented with 1 mM ATP and 2 mM magnesium acetate in TA buffer (40 mM Tris base, 20 mM acetic acid) at 3.33 V/cm for 60 min. Gels were imaged using a GE Amersham Imager 600. All bands were quantified through densitometry using ImageLab (Bio-Rad). Graphpad Prism was used for further analysis.

D-loop assay: Rad51-catalyzed D-loop assays were performed with slight modifications to the previously described methods123. A 5’ Cy5-labeled ssDNA oligonucleotide (25 nucleotides, olWDH2182 Supplementary Table 3) was annealed with another ssDNA oligonucleotide (100 nucleotides, olWDH2183 Supplementary Table 3) to serve as a tailed substrate. 0.25 µM wild-type Rad51, Rad51-ED, or Rad51-KN were incubated with 10 nM tailed substrate for 10 min at 30°C in a buffer containing 35 mM Tris-acetate (pH 7.5), 50 or 100 mM NaCl, 7 mM magnesium acetate, 2 mM ATP, 1 mM DTT, 0.25 mg/mL BSA, 20 mM phosphocreatine, and 100 µg/mL phosphocreatine kinase. Next, 42 nM RPA was added to the reaction mixture for another 10-minute incubation. Finally, Rad54 (120 nM) and supercoiled plasmid dsDNA (10 nM) were added, with samples taken at 0, 5, 10, 20, and 30 min as indicated. For the experiments testing the Rad54 dependence (Supplementary Fig. 9D), 20 nM of 95-mer (olWDH566, Supplementary Table 3) and Rad51 (0.67 µM) were incubated for 10 min at 30°C, in buffer (30 mM TrisOAc pH 7.5, 1 mM DTT, 50 mg/ml BSA, 5 mM MgOAc, 20 mM Phosphocreatine, 4 mM ATP, and 0.1 mg/ml creatine kinase). RPA (0.1 µM) was added and incubated for another 10 min at 30°C, followed by addition of Rad54 (0.112 µM) and dsDNA plasmid (20 nM molecules pUC19) to initiate the reaction and start the time course at 30°C. Samples were then deproteinized, separated on 1% agarose gels, and documented using a GE Amersham Imager 600. All bands were quantified through densitometry using Image Lab (Bio-Rad).

Preparation of oligonucleotide-based substrates: The oligonucleotide-based DNA substrate used for the helicase and nuclease assays was generated by labeling the BIO100C oligonucleotide (see Supplementary Table 1 for oligonucleotides sequence) at the 3’ end using terminal transferase (New England Biolabs) and [α-32P] dCTP (Hartmann Analytic) according to manufacturer’s instructions and purified on G25 columns (Cytiva). The purified oligonucleotide was mixed with a two-fold excess of BIO100 oligonucleotide (see Supplementary Table 1 for oligonucleotides sequence) in annealing buffer (10 mM Tris-HCl pH 8, 50 mM NaCl, 10 mM MgCl2), heated to 95°C for 3 min and cooled down to room temperature overnight.

Helicase and nuclease assays: Helicase and nuclease assays were performed with a 100 bp substrate (1 nM, in molecules, oligonucleotides BIO100C and BIO100) in reaction buffer containing 25 mM Tris-acetate pH 7.5, 1 mM dithiothreitol, 2 mM magnesium acetate, 1 mM ATP, 80 U/ml pyruvate kinase (Sigma), 1 mM phosphoenolpyruvate and 0.15 mg/ml bovine serum albumin (New England Biolabs), in a final volume of 15 µl. The reactions were performed in the presence of 45 nM (final) Saccharomyces cerevisiae RPA. The indicated Rad51 variant was added to the reaction followed by the indicated resection factors, Sgs1 (100 nM) for the helicase assays, and Exo1 (25 nM) or Dna2-Sgs1 (1 nM each) for the nuclease assays. The reaction was incubated for 30 min at 30°C and stopped by the addition of 5 µl of 2% STOP buffer (150 mM ethylenediaminetetraacetic acid [EDTA], 2% sodium dodecyl sulfate, 30% glycerol, bromophenol blue) supplemented with a 20-fold excess of unlabeled oligonucleotide with the same sequence as the labeled one (to prevent reannealing) and 1 µl of Proteinase K (18 mg/ml) (Roche). After deproteination for 30 min at 37°C, the reactions were separated on a 10% TBE gel. The gel was dried on 17 Chr paper (Whatman), imaged on a Typhoon Imager (Cytiva) and quantitated using the ImageJ software.

Protein pulldown assay: 100 nM GST–Rad55–His6–Rad57 or GST (Pierce) were incubated with 100 nM Rad51 in buffer P containing 25 mM Tris-acetate (pH 7.5), 10 mM magnesium acetate, 50 mM NaCl, 1 mM DTT, 10% glycerol and 0.01% NP-40 in a 25 µl volume for 1 h at room temperature. Equilibrated and BSA-treated glutathione agarose beads (Pierce) were added to the mixture and incubated for 1 h. The beads and supernatant were separated by centrifugation and the beads were washed twice with binding buffer P. The pulled-down protein complexes were eluted by heating at 65°C for 10 min in 12 µl 2x SDS–PAGE loading buffer and separated through a 10% SDS–PAGE gel. Anti-Rad51 (1:1000, sc-133089, Santa Cruz Biotechnology,) and anti-GST (1:5000, 27457701, Cytiva) antibodies were used for immunoblotting. Protein bands were visualized using Clarity Western ECL substrate (Bio-Rad) on immunoblots and quantified using ImageLab (Bio-Rad). 5% of the input and 100% of the elution for each reaction were loaded for comparison.

Electron microscopic imaging of Rad51 filaments
Rad51 filaments were assembled by incubating 2.5 µM Rad51, Rad51-ED, or Rad51-KN with 7.5 µM (bp or nt) of 1000 bp dsDNA or 700 nt ssDNA derived from phiX174 (SspI-Ahd1 fragment) at 30°C for 10 min in a buffer containing 35 mM Tris-acetate (pH 7.5), 50 mM NaCl, 4 mM MgOAC, 2 mM ATP, and 1 mM DTT. The reaction mixture was then freshly diluted tenfold in 10 mM Tris-HCl (pH 7.5), 50 mM NaCl, and 5 mM MgCl2. Samples were applied to 400-mesh carbon-coated copper grids (Ted Pella), negatively stained with 2% uranyl acetate, blotted dry, and then air-dried. Grids were imaged using a Talos Transmission Electron Microscope (Thermo Scientific), with images randomly captured from different grid regions. For each condition, 2–4 grids were used, and images were taken at 11,000x or 36,000x magnification. A total of 500 filaments per condition were manually measured using ImageJ.

Statistical analysis
The 95% or better confidence intervals for the median were calculated using the information provided in Supplementary Table 4. This utilizes the formula: (n+1)/2 ± 1.96(√n/2) to provide the upper and lower limit of the 95% or better confidence interval from the rank order of values. As such, this method will generate an uneven±confidence interval around the median, making it easier to compare multiple bar graphs on a single figure. The p-values were always generated between two datasets using pairwise unpaired t-test with Welch’s correction that does not assume equal standard deviations between datasets.

Data availability
All data supporting the findings of this study are available within the paper and its Supplementary Information. Source data are provided with this paper.

Peer review rounds
[{"round": 1, "reviewer_comments": {"reviewer_1": "The authors have done a very good job addressing comments raised. Specifically, they have\nprovided new data (Figs. 4, 5, and S7) and addressed questions raised regarding the nature of\nthe rad51-II3A and S181P mutants in the Discussion.\nThis is a nice contribution and I do not have have any more question.", "reviewer_3": "The manuscript by Meyer et al. originally described the identification and characterization of\nrad51 mutants that are efficient in homologous recombination (HR) but defective in post-\nreplicative repair, namely rad51-E135D (rad51-ED) and rad51-K305N (rad51-KN). In response\nto reviewers’ criticisms, the authors re-examined the impact of these mutations on HR and found\nthe defects to be more extensive than initially reported. Specifically, both rad51-ED and rad51-\nKN exhibited severe HR defects when homologous sequences were located on different\nchromosomes. The authors accordingly stepped back from their original claim that HR can be\nseparated from the activity required for post-replicative repair.\nOverall, I have major concerns with aspects of the data interpretation:\n(1) Fig. 4A\nThe authors claim that Rad51-ED binds ssDNA (100 nt) as efficiently as wild-type Rad51, and\nthat Rad51-KN does so even more efficiently. In contrast, the data suggest the opposite: both\nmutant proteins show a marked reduction in ssDNA binding. This is evident when examining the\nshifted bands and intermediates represented by the smear. If only the unshifted band is\nconsidered, one could plot the graphs and obtain the reported Kd values. However, this\napproach does not accurately reflect the binding activity. The more appropriate conclusion from\nFig. 4 is that both Rad51-ED and Rad51-KN are defective in ssDNA and dsDNA binding.\n(2) Fig. S6\nConsistent with Fig. 4, the results in Fig. S6 also indicate that Rad51-ED and Rad51-KN are\ndefective in both ssDNA and dsDNA binding. Although the authors interpret the ssDNA defect as\nreflecting local duplex formation associated with a longer ssDNA substrate, the straightforward\ninterpretation is that both mutant proteins are defective in ssDNA binding.\n(3) Fig. 2B\nThe authors claim that spontaneous recombination, as assayed using a direct-repeat\nrecombination system, is unaffected by rad51-ED and rad51-KN. However, the data do not\nconvincingly support this conclusion. Spontaneous recombination appears to be mildly reduced\nin rad51-KN and more so in rad51-ED. The apparent lack of statistical significance may reflect\nan insufficient sample size, if a statistical test was performed at all.\nThe authors have already shown that ATP hydrolysis stimulated by ssDNA is reduced in Rad51-\nED and Rad51-KN (Fig. 5A), more severely in Rad51-ED. Rad51 focus formation in response to\nMMS is also defective in both mutants (Fig. S8). These results are consistent with their defects\nin ssDNA binding.\nIn addition, Fig. 10A demonstrates that sister-chromatid joint formation is reduced in both rad51-\nED and rad51-KN. Even in the sgs1 background, the data suggest that inter-sister association is\ncompromised in these mutants, in agreement with my interpretation of Fig. 2B.\nTaken together, the evidence strongly supports the view that these two mutants are\nhypomorphic alleles with defects across a broader spectrum of HR functions than proposed by\nthe authors, with ED more severely affected than KN. Given that Rad51 has long been known to\nfunction in post-replicative repair, it is not surprising that partial defects in Rad51 activity\nproduce corresponding partial phenotypes.\nFurthermore, multiple studies have already reported that dsDNA binding of Rad51 is critical for\nprotecting stalled replication forks from nucleases (refs. 30, 31). This further diminishes the\nnovelty of the current work.\nMinor points\n(1) Line 219: there is no S2A (just S2).\n(2) Fig. S2: The figure appears to show raw CFU counts rather than viability, despite being titled\nas viability. Additionally, the elevated levels of rad51-KN under both induced and spontaneous\nconditions are confusing and should be more clearly explained in the main text.\n(3) Line 292: It would be useful to include ssDNA as well in the assay.\n(4) Line 424: Should this refer to Fig. S3B rather than S3A?\n(5) Line 425: The nature of the discrepancy mentioned is unclear and should be clarified.\n(6) Line 493: I was puzzled to find that the FigS11 does not show SCJ formation in the rad51\nnull mutant. Regarding the importance of including the rad51 null mutant in Fig10A. No matter\nhow many times it has been done previously, including a negative control is essential because it\nsets the base line for the data acquisition and interpretation.\n(7) Fig. 10B: The phenotypes of strains expressing Rad51-KN-MN and Rad51-ED-MN should\nbe compared directly with their untagged versions. The MN tag may alter the properties of these\nalleles.", "reviewer_4": "I have reviewed the responses to each critique made by all three reviewers. I think that by changing the terminology from separation of function mutants was appropriate, they addressed the biochemical concerns adequately by performing their binding assays with new substrates and their D-loop assays at higher salt concentrations, and by removing their alpha fold model structural analysis the authors have more than adequately addressed all reviewers concerns-particularly Reviewer 2."}, "author_response": "We greatly appreciate the helpful and constructive comments by all reviewers. Below we\ndiscuss point-by-point our responses and how we modified the manuscript to implement the\ncritiques. The text has been substantially revised and significant new data (Figures 3, 4, 5C-E,\nFigure S3C, S4, S5B,C, S7, S8, S9A-C, S11) were added, improving the manuscript\nsignificantly and offering additional insights into the mutants.\nIn short, while the mutants are fully proficient in inter-sister recombination, the analysis revealed\na defect in inter-homolog recombination (new Figure 3, S4), which in contrast to sister chromatid\nrecombination necessitates a genome-wide homology search. Moreover, additional biochemical\nanalysis revealed that the mutants form shorter filaments (new Figures S7, S8), which is likely\nthe reason for the inter-homolog HR defect.\nThe expanded biochemical analysis reinforced that the major defect of the mutants is binding to\ndsDNA, with good correlation between the in vitro and in vivo phenotypes showing a greater\nimpact of the E135D mutation (new Figure 4, 5C-E, S7, S9A-C).\nIn conclusion, we step back from the claim of separation-of-function mutants. The mutants\nreveal differences between sister-chromatid and interhomolog recombination, that align well\nwith recent results from the Piazza and Symington labs that genome-wide homology search\ndiffers from sister chromatid recombination. Importantly, our major conclusion stands reinforced\nthat the mutants primarily affect dsDNA binding and that the mutant phenotypes in vitro and in\nvivo correlate in severity with defects in fork protection leading to a shift in pathway usage at\nstalled forks and on over-reliance on TLS and fork regression in the mutants. We have revised\nthe discussion accordingly.\nWe added line numbers to the version with tracked changes (in red) to aid the reviewers.\nReviewer #1 (Remarks to the Author):\nIn this manuscript, the authors report the identification of novel mutations in Saccharomyces\ncerevisiae Rad51 that maintain its homologous recombination (HR) function but impair fork\nprotection. First, they found two Rad51 mutations (rad51-E135D (ED) and rad51-K305N (KN)\nthat suppress the MMS-sensitivity of a rad51∆/rad54∆ (or rad51∆/rad54ts) double mutant. The\nrad51-ED and -KN mutations induce an enhanced mutation rate and accentuate the MMS\nsensitivity of cells defective in post-replication repair, suggesting a shift in DNA repair pathway\nchoice from HR to alternate pathways. Biochemical analyses with rad51-ED and rad51-KN\nproteins show that they have diminished activities in ATP hydrolysis, DNA binding, and DNA\nprotection against nucleases, even though both appear to be proficient in D-loop formation.\nAdditionally, the authors present predicted 3D-structure of the Rad51-ssDNA filament\nreconstituted with the mutants, and provides insight into the structural basis of the alteration of\nRad51 interaction with duplex DNA.\nThe study is of general interests. However, I believe some claims could be more convincing if\nthe authors would examine other rad51 mutants in parallel. Additionally, it is unclear whether the\npresented phenotypes (e.g., MMS sensitivity), interpreted as changes in pathway choice (HR,\ntemplate switch, and translesion DNA synthesis), are solely due to Rad51's dsDNA binding or if\nthey are outcome of some other effects of the mutations.\nThank you for recognizing the general interest of the results. We would not venture to say that\nthe phenotype is ‘solely’ the consequence of the dsDNA binding defect, but that this defect best\ncorrelates with the phenotypic differences of the mutants. We made this point clearer in the text\n(lines 538-41, 554, 637).\n1. The rad51-ED and rad51-KN mutants were initially isolated from a screen using a double\nrad51/rad54 mutant (Fig. 1B), but subsequent studies with genomic integration of the mutations\ndo not show similar suppression (Fig 1E). The authors mention this difference on Page 7 but the\ndifference should be clarified further.\nThe reviewer is correct in pointing out that the rad51 mutants show significantly less\nsuppression on plates in single copy (Figure S1A) than when overexpressed (Figure 1B), as\nwas the case during the screen and the validation. Using a colony formation assay (Figure 1E)\nthe genomic integration showed no suppression. We explain this difference between the results\nfrom Figure 1B and Figure S1A to be a consequence of overexpressing the Rad51 mutants\nduring the screen and validation versus the genomic integration. The difference between plate\nassays which use serial dilutions (Figure 1B, S1B) and the colony formation assay in Figure 1E\nis explained by the former measuring a mixture of growth and survival, whereas the latter\nmeasures colony formation. We have added text to discuss these differences (lines 183-187).\n2. The mutator phenotype observed in rad51-ED and rad51-KN, despite the ir proficiency in HR,\nis an interesting finding. One wonder if this is specific to these two mutants or a general\nphenotype of rad51 mutants with decreased DNA binding affinity. The authors should test\nadditional hypomorphic DNA binding mutations that are equivalent to, e.g. T131P or I131A in\nhuman RAD51. This would add significantly to the study regardless of the outcome of the\nsuggested experiments.\nIt is unclear whether this is a general phenotype of hypomorphic rad51 mutants. Human\nRAD51-T131P is a semi-dominant mutation, whose viability depends on the presence of wild\ntype human RAD51 protein (PMID: 26253028). Our mutations were analyzed in a haploid\ncontext, which is not comparable. We are not familiar with an I131A mutation in human RAD51,\nand PubMed and Google searches were negative. The reviewer might be referring to the yeast\nrad51-II3A reported in Cloud et al. 2012 Science. This mutation is proficient in meiotic\nrecombination providing a short filament seed for the Dmc1 filament but is deficient in the mitotic\nHR function leading to IR sensitivity like the rad51 deletion mutant (Cloud et al. 2012). This\nphenotype clearly differs from our mutants which are not IR sensitive (rad51-KN) or significantly\nless than the deletion (rad51-ED). We also discuss a comparison with a recently published\nseparation of function mutant in human RAD51, RAD51S181P. We added a brief discussion of\nthese differences to the discussion on lines 609-622.\n3. Fig 4 & Fig S4: NaCl titration was used to examine DNA binding activity on ssDNA and\ndsDNA. However, protein titration that can be interpreted in terms of Kd would provide more\ninformative insights than NaCl titration. Additionally, the use of long ssDNA may underestimate\ndifferences in ssDNA binding due to secondary structures forming in the substrate. I suggest\ntesting shorter DNA substrates with a more quantitative method, e.g. MST.\nThe reviewer has three concerns. First, the EMSA conducted as salt midpoint titration to\nmeasure relative differences in DNA binding affinity. EMSA and salt midpoint titrations are a\nwell-established and accepted approach to compare DNA binding affinities, going back to the\npioneering work of Peter von Hippel and being frequently used for recombination proteins by\nSteve Kowalczykowski and others. Second, the reviewer prefers a protein titration to allow\ndetermining apparent Kd. Third, the reviewer is worried that the ssDNA substrate used in our\nEMSA experiments may form secondary structures, which may lead to an underestimation of\nthe ssDNA binding defect of the mutants. We accept the last two points and have conducted\nprotein titrations with 100 nt and 100 bp substrates to derive K values at 50 mM and 100 mM\nd\nNaCl (new Figure 4), the same concentrations at which we conducted the new D-loop\nexperiments (see point #4, new Figure 5C-E). As implied by the reviewer, the ssDNA binding\ndefect using the longer substrate was likely related to its secondary structure, as both mutant\nproteins showed no binding defect to the 100 nt substrate. These results reinforce the\nconclusions that the mutant proteins affect dsDNA binding, and that Rad51-ED displays the\nlarger effect than Rad51-KN.\nWe examined filament length by EM to further examine the dsDNA binding defect and found\nthat both mutant proteins formed significantly shorter filaments (new Figure S7). We also\nexamined filament formation in vivo by immunofluorescence (IF) and found that both mutants\ncaused a strong defect in Rad51 filament formation in response to DNA damage (new Figure\nS8). This result suggests that also filament formation on ssDNA is affected and may provide an\nexplanation for the defect in interhomolog recombination. In this respect our mutants are similar\nto the rad51-II3A mutant described by Dr. Bishop (see point #3), which also forms shorter\nfilament that are undetectable by IF. This point was added to the discussion (lines 609-610).\nThese results were unexpected but provide significant insights into the mutants. We thank the\nreviewer for the helpful comments that made us reexamine this aspect.\n4. Fig 7F: The D-loop assay was performed under low-salt condition, where the mutant does not\nshow much difference in DNA binding affinity, as indicated in Fig 7C. It is important to\nstandardize conditions for DNA binding and D-loop reactions to render conclusions robust.\nWe agree with the reviewer’s point and added new D-loop experiments that were conducted at\n50 mM and 100 mM NaCl using the same 100mer used for the new EMSA experiments (see\nresponse to point #3). These new data are presented in the new Figure 5.\n5. Results from the D-loop assay (Fig. 7F) indicate that rad51-ED exhibits 9same can be said\nabout rad51-KN, albeit to a lesser degree) more robust activity compared to wild type and the\nKN mutant proteins. The authors need to clarify whether this difference is due to intrinsic,\nenhanced recombinase activity of rad51-ED or resistance of D-loop coated by rad51-ED to\nRad54-mediated D-loop dissociation.\nThe reviewer points out that in D-loop experiments using short invading ssDNA two phases\nexist, an early D-loop formation phase and a later D-loop dissociation phase. The D-loop\ndissociation is a function of Rad54, as we have shown in Wright and Heyer (2014) PMID\n24486020. The reviewer makes a good point that the Rad51 mutant could affect either phase of\nthis process. We conducted new D-loop experiments under the same salt conditions as the\nEMSA experiments using a tailed substrate (new Figure 5). The kinetics shows no difference in\nthe formation phase, but faster dissociation of D-loops in the reaction with the mutant Rad51\nproteins. This is consistent with a functional defect in binding dsDNA, as after D-formation\nRad51 is bound to dsDNA, from where it needs to be displaced for D-reversal by Rad54 (Wright\nand Heyer (2014).\n6. Fig 8: The dsDNA binding properties and DNA protection observed are consistent with\nprevious reports on human proteins, and the results align with expectation based on their NA\nbinding activity. It will be good to test additional other rad51 mutants (see point #4).\nIt is unclear what can be gained by analyzing additional mutants, in particular the ones\nmentioned by the reviewer, see response to point #2. (The reviewer refers to #4, which appears\nto be incorrect.)\nReviewer #2 (Remarks to the Author):\nMeyer and Ceballos et al. investigate a role of central HR recombinase Rad51 in post-\nreplication repair using separation-of-function mutants that maintain recombination activities but\nare specifically defective in binding to dsDNA with consequent changes to in vivo repair results.\nThe multitude of genetic assays and in vitro characterization agree with the authors’ model\nimplicating a functional role of yeast Rad51 binding to dsDNA at stalled replication forks\nmodulating pathway choice. The authors infer potential structural mechanisms for the two\nmutants characterized by comparing mutant AlphaFold predictions with an experimentally\nderived structure, but it is not clear the evidence provided is sufficient to justify the authors\nassertions. Overall, the manuscript provides an interesting dissection of a non-standard role of\nRad51 through convincing and logical experiments, but the structural evidence is currently too\nspeculative and would need to be supported by additional experimental structures or other\ntechniques.\nWe thank the reviewer for the supportive comments and agree that the structural modeling does\nnot provide a satisfactory explanation for the biochemical and in vivo behavior of the proteins.\nSince we just had completed the high-resolution cryoEM structure of the yeast Rad51-ssDNA\nfilament with ADP-AlF , which is now published in Nucleic Acids Research (Liu et al. 2025), we\n3\nfelt compelled to try to model the mutants. However, we realize now this analysis does not add\nto the manuscript and we eliminated this part. We only retained as supplemental Figure S5A-C\nthe information on the sequence conservation around both residues and their locations with\nrespect to bound DNA and nucleotide co-factor.\nMajor comments:3\n1. The mutants were isolated in a gain-of-function genetic screen for alleles that allow MMS\nresistance in a temperature sensitive rad54 strain. It appears that this phenotype requires these\nalleles be on a high copy plasmid and no MMS resistance over wild-type protein was seen in\nnative locus replacements, and additionally the MMS resistance with high copy plasmid is much\nweaker in a rad54 delete than rad54ts at non-permissive temperature. Presumably the high\ncopy plasmid requirement means over-expression is necessary, but this is not specifically\nestablished. It is not clear how this screen would lead to the mutants and eventual phenotypes\nfound. It would be helpful for the authors to discuss this and perhaps speculate on what is\nhappening on a high copy plasmid with these mutants vs wild-type protein.\nSee above.\n2. The authors did not write the method section of Alphafold-prediction. Were ssDNA, ADP-\nAIF3, and magnesium included in the prediction to match the same cryoEM condition?\nThe authors compared individual side chain movements determined by cryoEM to the mutants\nthat were predicted by Alphafold. To validate prediction of side chain positions in Alphafold, a\nvery important thing to do first is the same Alphafold prediction of WT-Rad51 and compared to\ncryoEM structure of WT-Rad51. Would side chain positions match each other? Would\ncomparison of  helices positions look different from the Alphafold predictions of WT and the\nmutants like Fig 9C? if the authors can show WT-Rad51 Alphafold prediction and is indeed look\ndifferent from the mutants in overall filament shape + side chains, then I am convinced.\nHowever, if the WT-Rad51 Alphafold prediction looks more similar to other mutants Alphafold\nprediction, these differences the authors show in the paper may come from different\nmethodology (cryoEM vs Alphafold predictions).\nSee above.\n3. The authors claim on page 16 to be “detecting the movement of amino acid side chains in the\nRad51-ssDNA structure”, but this seems to be more of an inference based on different protomer\npositions in the filament structure. This language seems to imply a more direct technique was\nused. Additionally, the expected change is on dsDNA while the experimental structure is on\nssDNA with modeled dsDNA to infer mechanism. How do the authors know there aren’t\ndifferences in the relevant side chain position/dynamics on ssDNA vs dsDNA? This is\nparticularly relevant when very small changes are implied to cause an effect (see next\ncomment). Assertions throughout, including the statement on page 17 in the first paragraph of\nthe discussion “showing that the mutants render Rad51 less dynamic and more rigid” is not\nconvincing from the modeling.\nSee above.\n4. Pg16. “The Rad51-KN mutant has strengthened the interaction with M301 and A309,\nshortening the H-bond length from 3.1 Å to 3.0 Å, and 3.1 Å to 2.9 Å, respectively (Fig. 9D)”\nI don’t think this is valid argument...H-bond 3.1 to 3.0 to 2.9…\nSee above.\n5. In this paper, the authors compared a cryo-EM structure of yeast Rad51-ssDNA filament with\nADP-AIF3 (previously determined) with AlphaFold predicted structures of mutants (Rad51-ED\nand Rad51-KN). In the abstract and introduction parts, these are not described clearly in a way\nthat readers may expect to see new CryoEM structures at 2.4A resolution in the result section\nafter reading the abstract.\nSee above.\n6. Overall, the paper is too long with repetitive descriptions, may make readers lose interest\nwhile reading. Few descriptions/ data need to be shortened.\nWe shortened the descriptions and eliminated repetitions. Thank you for the comment.\nMinor comments:\n1. The last full sentence on page 3 is not clear/potential typo.\n2. Typo on page 15 “Wild type Rad51 inhibited DNA degradation by Rad51”\nThank you for spotting these issues, which are now corrected. Lines 86-88, line 340.\nReviewer #3 (Remarks to the Author):\nThe MS by Meyer et al. describes the identification and characterization of rad51 mutants that\nare efficient in HR but defective in HR-mediated template switching. In contrast to the null\nmutant, the rad51-E135D (rad51-ED) and -K305N (rad51-KN) mutants are genetically efficient\nin HR, based on HO-triggered sister chromatid assays. The purified Rad51-ED and -KN are at\nleast as efficient in D-loop formation as wild-type Rad51. In contrast, rad51-KN is partially\nsensitive to MMS and rad51-ED is as sensitive to MMS as the rad51 null mutant. These mutants\nshow strong synergy with post-replicative repair mutants in MMS sensitivity, suggesting their\ndefects in the generation of HR-mediated intermediates at replication forks, which is supported\nby their 2D gel analysis. They also showed that the post-replicative repair pathway is shifted to\ntranslesion synthesis in these rad51 mutants. Biochemically, these two mutant proteins,\nespecially the rad51-ED, exhibit a reduction in ATPase activity and DNA binding to both single-\nstranded (ss) and double-stranded (ds) DNA, with dsDNA binding more severely affected.\nConsistently, these mutant proteins fail to protect dsDNA from degradation by exonucleases\n(Exo1 and Dna2-Sgs1) and the Sgs1 helicase. Taken together, it is proposed that Rad51\nbinding to duplex DNA is critical to control the use of the pathway in post-replicative repair.\nThis MS would provide valuable information if these two rad51 mutations are indeed clear\nseparation-of-function alleles. On the other hand, it is always a bit tricky to interpret the data\nobtained with hypomorphic mutants. Regarding their phenotypes in post-replicative repair,\nrad51-ED is similar to the rad51 null, while rad51-KN is partial. Regarding biochemistry, their\nfailure to protect dsDNA from exonucleases and the Sgs1 helicase can be explained by their\ndefect/reduction in dsDNA binding. The question is how good are they at HR, which is claimed\nto be as good as wild-type based primarily on HO-triggered sister chromatid assays and D-loop\nassays. I feel I need to be further convinced of their HR proficiency.\nWe thank the reviewer for the helpful comments that led to significant new experimentation\nwhich greatly improved our understanding of the functional defects of the Rad51 mutants. This\nanalysis revealed a defect in inter-homolog recombination (new Figure 3), which in contrast to\nsister chromatid recombination necessitates a genome-wide homology search. Moreover,\nadditional biochemical analysis revealed that the mutant form shorter filaments (new Figures\nS7, S8), which is likely the reason for the inter-homolog HR defect.\nThe expanded biochemical analysis reinforced that the major defect of the mutants is in binding\nto dsDNA, with good correlation between the in vitro and in vivo phenotypes showing a greater\nimpact of the E135D mutation (new Figure 4).\nIn conclusion, we step back from the claim of separation-of-function mutants. The mutants\nreveal differences between sister-chromatid and interhomolog recombination, that align well\nwith recent results from the Piazza and Symington labs that genome-wide homology search\ndiffers from sister chromatid recombination. Our major conclusion stands reinforced that the\nmutants primarily affect dsDNA binding and that the mutant phenotypes in vitro and in vivo\ncorrelate in severity with defects in fork protection leading to a shift in pathway usage at\nstalled forks and on over-reliance on TLS and fork regression in the mutants. We made numerous text\nchanges accordingly.\nPlease provide line numbers for ease of review.\nWe added line numbers to indicate the changes in the version of the revised manuscript with\ntracked changes.\n(1) As explained above, rad51-ED is similar to rad51-null, while rad51-KN has partial MMS\nsensitivity, also in synergy with post-replicative repair mutants. In contrast, HR is fully competent\nin the sister chromatid assay. The proficiency of HR needs to be more concretely validated by\nfollowing the mating type switching process initiated by the GAL-induced HO endonuclease by\nSouthern blotting. In this way, not only the final efficiency of HR but also the kinetics can be\nclearly monitored.\nWe agree with the reviewer and added two key experiments: First, we use our newly developed\nD-loop capture assay to directly track D-loop levels. Second, we used a BIR assay that allows to\nphysically and genetically track the endpoints. These data are shown in Figure 3. Surprisingly\nand unexpectedly, both mutants showed a strong defect in interhomolog recombination that\nrequired a genome-wide homology search. Interestingly, this phenotype correlates with shorter\nRAD51 filaments, which may point to a difference between inter-sister and inter-homolog\nrecombination in the requirement for long Rad51 filaments (Figures S7, S8).\n(2) Regarding the D-loop assay shown in Fig. 7F. The data certainly show that the D-loop\nformation efficiency of rad51-ED and KN is at least as high as the wild-type level (or higher). On\nthe other hand, Figure 7A-E clearly show that these proteins are impaired in ATP hydrolysis and\nDNA binding. Although ATP hydrolysis may not be very relevant to D-loop formation, ssDNA\nbinding, as I understand, is a prerequisite for D-loop formation. Given that the physiological ionic\nstrength is ~120 mM NaCl (page 14, 6th line from the bottom of the 1st paragraph), D-loop\nassay results within this range (i.e., 0~120 mM) should also be presented and their HR\nperformance should be evaluated with these results included.\nWe agree with the reviewer and added new D-loop experiments under the same salt conditions\nas the new EMSA experiments at 50 mM and 100 mM NaCl. In fact, using a 100mer that is\ndevoid of dsDNA potential revealed that the mutant proteins had no defect binding ssDNA in the\nEMSA assay. See also our responses to points #4, 5 of reviewer 1 on this topic.\n(3) The aim and rationale of the suppressor screen needs to be described in more detail. I found\nthe relevance of using rad54 particularly confusing. I understand that Rad54 is not required for\nfork protection in mammals. The question is, is Rad54 required for fork protection in S.\ncerevisiae? MMS is introduced as a fork stalling agent, but according to Figure 1C,\nrad54 is as sensitive to MMS as rad51. Overall, I felt that the screening strategy was poorly explained, so it\nwas hard to understand what the goal and principle of this genetic screen was.\nThe reviewer is correct in noting that the MMS sensitivity of rad51 and rad54 cells is very similar\nbut that does not necessarily imply identical mechanisms. We have improved Figure 1 and\nadded clarification to the legend. The goal was to isolate novel rad51 mutants that separated\nRad51 function from Rad54 function. Reviewer 1 brought up a similar point (#1).\n(4) Related to (3), please explain how the identified properties of these mutants account for their\nsuppression of rad54 MMS sensitivity.\nReviewer 1 brought up the same point (#1), and we addressed this there.\n(5) Fig. 6B, rad51-ED ChEC result is not convincing. The main band of the MMS-treated rad51-\nED sample (15 min) shows a significant reduction in intensity. How could the band lose intensity\nwithout forming a smear?\nEven though we try to load similar amounts of total DNA, there are some differences as with the\nindicated case where the lower band intensity reflects less total DNA. To eliminate DNA loading\ndifferences, the area of the DNA digestion profiles was equalized as indicated in the methods\nsection. We added some explanation to the text (line 781-782) and added a figure of the repeat\nexperiment as Figure S11 as well as a comment that the results are reproducible.\nMinor points\n(6) Fig. 6A. Please include rad51-del (sgs1-del) in the 2D gel analysis, which is necessary to\nestablish the baseline for assessing the phenotypes of rad51-KN and -ED.\nRad51 is required for the formation of SCJs as determined by 2D gel analysis (less of 5% of the\nwild type levels are left in the rad51 null mutant). This observation was first reported by Foiani\nand colleagues (Liberi et al. (2005) Genes Dev. 19:339; PMID 15687257). This has been\nreproduced several times; lastly by Branzei and colleagues (Joseph et al. 2022; Nature Comm.\n13:2480; PMID 35513396 Figure 6B), using the same protocol and experimental conditions that\nwe have used in the current study (replication and recombination intermediates from ARS305 in\ncells released from G1 in 0.033% MMS). As can be concluded from our analysis in Figure 6A,\nthe rad51-ED mutant displayed a defect almost as severe as rad51∆ (10-15% of the wild type),\nwhereas the effect of rad51-KN was much milder. We have added this information to the results\nsection (lines 491-493).\n(7) Fig. 4. Please be consistent with the MMS sensitivity assay. rev3 and pol30 are spot assays\nwhile mms2 is a colony formation assay, making them difficult to compare. Please also include\nrad5-G535R results here. The results shown in Fig. S1AB are very confusing as the rad5-\nG535R results are included without any explanation until page 11. Also, please include the rad5-\nG535R single mutant in the spot assay, which is necessary to interpret the result.\nThe experiments in Figure 4A, B, F, G (now Figure 7) used serial dilution assays and C, D, E\nquantitative survival assays, because the mms2 analysis in C-E involved far fewer strains. This\nmore quantitative analysis with mms2 allowed more detailed insights into the relationship\nbetween the rad51 mutants and PCNA polyubiquitylation. The results clearly show the\nsynergy/additivity between the rad51 mutants and all PRR mutants in A-G. We are not sure,\nwhat the effort to conduct quantitative survival assays for another 48 strains would add.\nFigure S3 (formerly Figure S1) serves two purposes, first to show the IR sensitivity of the\nmutants, and second, the show the effect of the hypomorphic rad5-G535R mutation. We have\nreorganized the figure to reflect this sequence and guide the reader through the figure to focus\nfirst on the IR data in RAD5 wild type strains and then later on the rad5-G535R data.\nWe agree with the reviewer that the single rad5-G535R strain is missing from the analysis in\nFigure S1 (now Figure S3), as it was described and published to have no effect on growth and\nsurvival at his low MMS concentration. We added the direct comparison of rad5-G535R, wild\ntype, and rad51 to Figure S3 as part C. We can no longer conduct the IR experiment in Figure\nS1B, as our cesium source has been decommissioned. rad5-G535R was described as IR-\nresistant, and we added this information to the text (line 430-432).\n(8) Fig. 3 title \"rad51-ED and rad51-KN display a strong mutator phenotype similar to a\ncomplete HR defect\". It seems that the mutator phenotype of rad51-ED when combined with\nrad54 is stronger than rad51-del (24x vs 10x), so the title is misleading. It would also be helpful\nto explain how this could happen.\nWe agree with the reviewer that the title does not reflect that particular result. We rephrased the\ntitle (now Figure 8) and added some speculation, as to how this additive effect could be\nexplained on line 438, 450-451.\n(9) Fig. 5A. Overall, the system is poorly explained; everything is so small, and some critical\ninformation for understanding the system is missing. For example, what is lox71? Does it mean\nanything without explaining what lox66 is? Which strand is in frame and which is not? etc.\nPlease also clarify if rad51-ED and -KN are different from null. Also include how many\ncolonies were examined per session.\nThe experimental system is now better explained in the text (lines 459-461). Specifically, the\ndescription of the use of lox71 and lox66 sites has been added to the figure legend and in the\nresults section.\nThe figure has been improved according to the reviewer’s suggestions. The text in the figure\nhas been enlarged.\nIn response to the reviewer’s questions about the strands: The strand in frame is the strand\nwhere the lesion has been inserted. Our system allows insertion either on the lagging or lagging\nstrand. Since we did not observe any difference, the data shown is an average of leading and\nlagging strand integration. This is now better explained in the method section (lines 790-793,\n797-800). The number of colonies is also now mentioned in the method section.\nAs requested, we added the statistical tests comparing rad51-ED and rad51-KN to the rad51\nnull mutant. Some of the differences are not significant, and some have a high p-value\nindicating low significance. Given the small difference and high p-value, the differences are\nunlikely to be biological relevance.\nThank you for your comments to improve the presentation.\n(10) Page 20, last line of the 2nd paragraph \"Alternatively, it is possible that ...\"\nIt was not clear to me what exactly this line means.\nWe have eliminated that discussion.\n(11) Fig. 1B. It was not clear which experiment (or both) was done at 37C (although I can\ncertainly guess that ts experiments were done at 37C while the null was done at 30C).\nWe apologize for the oversight, both experiments were conducted at 37C, and this is now noted\nin the figure legend in addition to notation in the figure.\nResponse to reviews\nWe thank reviewers 1 and 4 for accepting our revision and reviewer 3 for thoughtful comments\nthat we are addressing in this second revision. These comments helped us to more accurately\ndescribe the results, and we appreciate the careful editorial comments.\nReviewer #1 (Remarks to the Author):\nThe authors have done a very good job addressing comments raised. Specifically, they have\nprovided new data (Figs. 4, 5, and S7) and addressed questions raised regarding the nature of\nthe rad51-II3A and S181P mutants in the Discussion.\nThis is a nice contribution and I do not have have any more question.\nResponse:\nThank you for your supportive comments.\nReviewer #3 (Remarks to the Author):\nThe manuscript by Meyer et al. originally described the identification and characterization of\nrad51 mutants that are efficient in homologous recombination (HR) but defective in post-\nreplicative repair, namely rad51-E135D (rad51-ED) and rad51-K305N (rad51-KN). In response\nto reviewers’ criticisms, the authors re-examined the impact of these mutations on HR and found\nthe defects to be more extensive than initially reported. Specifically, both rad51-ED and rad51-\nKN exhibited severe HR defects when homologous sequences were located on different\nchromosomes. The authors accordingly stepped back from their original claim that HR can be\nseparated from the activity required for post-replicative repair.\nOverall, I have major concerns with aspects of the data interpretation:\nResponse\nWe agree with major points 1 and 2 that there is a minor ssDNA binding defect that is not\ncaptured in the Kd and STM data but is indicated by the increased smear in the EMSA with\nssDNA. This may be related to a subtle stability defect in ssDNA binding, which we now\nacknowledge in the discussion. This should not distract from the major difference between the\nstrong dsDNA binding defect and the subtle ssDNA binding shown by three independent\nexperiments (Figure 4, S6, S7).\n(1) Fig. 4A\nThe authors claim that Rad51-ED binds ssDNA (100 nt) as efficiently as wild-type Rad51, and\nthat Rad51-KN does so even more efficiently. In contrast, the data suggest the opposite: both\nmutant proteins show a marked reduction in ssDNA binding. This is evident when examining the\nshifted bands and intermediates represented by the smear. If only the unshifted band is\nconsidered, one could plot the graphs and obtain the reported Kd values. However, this\napproach does not accurately reflect the binding activity. The more appropriate conclusion from\nFig. 4 is that both Rad51-ED and Rad51-KN are defective in ssDNA and dsDNA binding.\nResponse:\nThe reviewer is correct in pointing out that the data are plotted as bound DNA, which is the\ninverse of the unshifted substrate band. Our interpretation focused on the massive difference\nbetween ssDNA and dsDNA, which is reflected in the Kd data. The EMSA reveals additional\ninformation on the electrophoretic profile of the protein:DNA complexes. The reviewer points out\nthat there is more smear of not fully shifted substrate in the reactions with mutants. We now\nacknowledge this in the discussion of the data on page 9 and reworded the section heading on\npage 8 accordingly. This is likely related to subtly altered stability in ssDNA binding.\n(2) Fig. S6\nConsistent with Fig. 4, the results in Fig. S6 also indicate that Rad51-ED and Rad51-KN are\ndefective in both ssDNA and dsDNA binding. Although the authors interpret the ssDNA defect as\nreflecting local duplex formation associated with a longer ssDNA substrate, the straightforward\ninterpretation is that both mutant proteins are defective in ssDNA binding.\nResponse:\nAs the reviewer pointed out in the previous round of review, the long DNA substrates used in\nFigure S6 form secondary structures with dsDNA hairpin stems. For this reason, we added the\nexperiment in Figure 4 using ss oligonucleotide substrates devoid of potential for dsDNA\nformation to address this point. The EMSA data were quantified as in Figure 4 showing bound\nDNA, the inverse of free substrate. The salt titration midpoint (STM) is significantly more\naffected with dsDNA than with ssDNA. The reviewer is correct that the STM reduction for ssDNA\nreflects that a portion of the substrate was dsDNA, as was pointed out in the original review, and\npossibly altered affinity/stability in binding, as pointed out now. We acknowledge this latter\npossibility now specifically on page 9. Also, in Figure S6, the data clearly show a much stronger\ndefect in dsDNA binding, for example at 0 mM NaCl there is no defect for ssDNA but a 20% or\nmore reduction in binding dsDNA. Likewise, the STM values are much more reduced for dsDNA\n(58-75%), than for ssDNA (42-47%). Also the EM analysis of filaments (Figure S7) shows a\nstronger defect in binding dsDNA than ssDNA. These data independently validate the\nconclusion that dsDNA binding is much more affected than ssDNA binding.\n(3) Fig. 2B\nThe authors claim that spontaneous recombination, as assayed using a direct-repeat\nrecombination system, is unaffected by rad51-ED and rad51-KN. However, the data do not\nconvincingly support this conclusion. Spontaneous recombination appears to be mildly reduced\nin rad51-KN and more so in rad51-ED. The apparent lack of statistical significance may reflect\nan insufficient sample size, if a statistical test was performed at all.\nResponse:\nWe conducted statistical analysis for Figure 2B using a t-test for equal means revealing a p-\nvalue of 0.017 between wildtype and rad51-ED showing the means are significantly different.\nHowever, the fold difference of 3.86 when comparing means and 3.0 when comparing median\nvalues is minor when juxtaposed with rad52, rad51 and rad54 mutants, which have about a 100-\nfold effect. A p-value of 0.084 between wildtype and rad51-KN showed no significant difference\nalthough there is a fold difference of 2.15 when comparing means and 1.9 when comparing\nmedian values. Finally, there is a significant difference between rad51-ED and rad51-KN with a\np-value of 0.027. No other p-values showed any significance for the ED and KN mutants. We\nadded this information to the text on page 7.\nThe authors have already shown that ATP hydrolysis stimulated by ssDNA is reduced in Rad51-\nED and Rad51-KN (Fig. 5A), more severely in Rad51-ED. Rad51 focus formation in response to\nMMS is also defective in both mutants (Fig. S8). These results are consistent with their defects\nin ssDNA binding.\nResponse:\nWe agree with the reviewer, but the key point is that the defect with dsDNA is considerably\nstronger than with ssDNA. The defect in ATPase activity may or may not reflect an underlying\ndefect in DNA binding. It is unclear whether Rad51 focus formation in response to MMS reports\nthe ssDNA binding, dsDNA binding, or both.\nIn addition, Fig. 10A demonstrates that sister-chromatid joint formation is reduced in both rad51-\nED and rad51-KN. Even in the sgs1 background, the data suggest that inter-sister association is\ncompromised in these mutants, in agreement with my interpretation of Fig. 2B.\nResponse:\nWe agree with the reviewer and conclude in the manuscript that sister chromatid joint formation\nis reduced in both mutants. However, it is unclear whether ssDNA and/or dsDNA binding is\nrequired for this. Hence, the result does not directly relate to the interpretation of Figure 2\ndata.\nTaken together, the evidence strongly supports the view that these two mutants are\nhypomorphic alleles with defects across a broader spectrum of HR functions than proposed by\nthe authors, with ED more severely affected than KN. Given that Rad51 has long been known to\nfunction in post-replicative repair, it is not surprising that partial defects in Rad51 activity\nproduce corresponding partial phenotypes.\nFurthermore, multiple studies have already reported that dsDNA binding of Rad51 is critical for\nprotecting stalled replication forks from nucleases (refs. 30, 31). This further diminishes the\nnovelty of the current work.\nResponse:\nWe agree with the reviewers that these rad51 mutants are hypomorphs, and we have explicitly\ntaken back the claim of separation-of-function. The reviewer correctly points, and we discuss\nthis in the manuscript, that refs. 30, 31 showed that binding to dsDNA is critical for fork\nprotection by RAD51. The key novelty in our manuscript is that this affects pathway usage at\nstalled forks.\nMinor points\n(1) Line 219: there is no S2A (just S2).\nResponse:\nCorrected, thanks for noting.\n(2) Fig. S2: The figure appears to show raw CFU counts rather than viability, despite being titled\nas viability. Additionally, the elevated levels of rad51-KN under both induced and spontaneous\nconditions are confusing and should be more clearly explained in the main text.\nResponse:\nThe reviewer is correct. The KN data were erroneously swapped with the wt data. Thank you for\npointing this out. We apologize for our oversight and corrected the figure as well as numerical\ndata.\n(3) Line 292: It would be useful to include ssDNA as well in the assay.\nResponse:\nWe have conducted the experiment suggested by the reviewer and added the data as Figure\nS7E-H. The results confirm that the defect in binding dsDNA is more pronounced than the\ndefect on ssDNA.\n(4) Line 424: Should this refer to Fig. S3B rather than S3A?\nResponse:\nCorrected, thanks for noting.\n(5) Line 425: The nature of the discrepancy mentioned is unclear and should be clarified.\nResponse:\nWe added additional clarification.\n(6) Line 493: I was puzzled to find that the FigS11 does not show SCJ formation in the rad51\nnull mutant. Regarding the importance of including the rad51 null mutant in Fig10A. No matter\nhow many times it has been done previously, including a negative control is essential because it\nsets the base line for the data acquisition and interpretation.\nResponse:\nWe eliminated the comparison to the rad51 null mutant.\n(7) Fig. 10B: The phenotypes of strains expressing Rad51-KN-MN and Rad51-ED-MN should\nbe compared directly with their untagged versions. The MN tag may alter the properties of these\nalleles.\nResponse:\nWe conducted this comparison and added the data to Figure S11A.\nReviewer #4 (Remarks to the Author):\nI have reviewed the responses to each critique made by all three reviewers. I think that by\nchanging the terminology from separation of function mutants was appropriate, they addressed\nthe biochemical concerns adequately by performing their binding assays with new substrates\nand their D-loop assays at higher salt concentrations, and by removing their alpha fold model\nstructural analysis the authors have more than adequately addressed all reviewers concerns-\nparticularly Reviewer 2.\nResponse:\nThank you for your supportive comments."}]

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