The Dmc1 recombinase physically interacts with and promotes the meiotic crossover functions of the Mlh1–Mlh3 endonuclease

Abstract

The accurate segregation of homologous chromosomes during the Meiosis I reductional division in most sexually reproducing eukaryotes requires crossing over between homologs. In baker's yeast approximately 80% of meiotic crossovers result from Mlh1–Mlh3 and Exo1 acting to resolve double-Holliday junction intermediates in a biased manner. Little is known about how Mlh1–Mlh3 is recruited to recombination intermediates to perform its role in crossover resolution. We performed a gene dosage screen in baker's yeast to identify novel genetic interactors with Mlh1–Mlh3. Specifically, we looked for genes whose lowered dosage reduced meiotic crossing over using sensitized mlh3 alleles that disrupt the stability of the Mlh1–Mlh3 complex and confer defects in mismatch repair but do not disrupt meiotic crossing over. To our surprise we identified genetic interactions between MLH3 and DMC1, the recombinase responsible for recombination between homologous chromosomes during meiosis. We then showed that Mlh3 physically interacts with Dmc1 in vitro and in vivo. Partial complementation of Mlh3 crossover functions was observed when MLH3 was expressed under the control of the CLB1 promoter (NDT80 regulon), suggesting that Mlh3 function can be provided late in meiotic prophase at some functional cost. A model for how Dmc1 could facilitate Mlh1–Mlh3's role in crossover resolution is presented.

The authors identified genetic and biochemical interactions in baker’s yeast between Mlh1-Mlh3, which acts in meiotic crossover resolution, and Dmc1, the recombinase responsible for recombination between homologous chromosomes during meiosis. Partial complementation of Mlh3 crossover functions was observed when MLH3 expression was restricted to roughly the time of crossover resolution, suggesting that Mlh3 function can be provided late in meiotic prophase at some functional cost. A model for how Dmc1 could facilitate Mlh1-Mlh3’s role in crossover resolution is presented.

Introduction

Accurate segregation of homologous chromosomes during the reductional division (Meiosis I) in most sexually reproducing eukaryotes is accomplished by the physical tethering of homologs through crossing over and distal chromatid cohesion, enabling stable orientation of homologs on the meiotic spindle (Maguire 1974; Hunter 2015; Zickler and Kleckner 2015). Defects in crossing over result in gametes that are aneuploid and often nonviable. In humans such abnormalities have been linked to miscarriages, birth defects, and developmental disabilities (Hassold and Hunt 2001; Nagaoka et al. 2010; Hunter 2015).

Meiotic recombination events in yeast and mammals are initiated in meiotic prophase through Spo11-induced double-strand breaks (DSBs) that occur genome-wide (Keeney et al. 1997; Pan et al. 2011). These DSBs are resected in a 5′ to 3′ direction by the activities of factors that include Mre11-Rad50-Xrs2 (MRX; MRE11-RAD50-NBS1 in humans) and Exo1 (Cao et al. 1990; Fiorentini et al. 1997; Nicolette et al. 2010; Zakharyevich et al. 2010). The RecA homologs Rad51 and Dmc1 are loaded onto the resulting resected 3′ tails in a defined orientation, with Dmc1 located nearer to the 3′ end. Dmc1 then catalyzes a homology search for allelic loci, with Rad51 acting as an essential accessory factor in a noncatalytic role. Dmc1-coated 3′ tails invade a homologous donor locus, forming a displacement loop (D-loop) that can be extended by DNA polymerase activity (Bishop 1994; Cloud et al. 2012; Brown et al. 2015; Hunter 2015; Hinch et al. 2020).

In baker's yeast, approximately half of the 150–200 Spo11-induced meiotic DSBs are repaired as noncrossovers through the helicase-driven unwinding of extended D-loops (Synthesis-Dependent Strand Annealing, reviewed in Pyatnitskaya et al. 2019). The remainder are stabilized by the functionally diverse ZMM family of pro-crossover factors consisting of Zip1-4, Spo16, Msh4-Msh5, and Mer3 to form single-end invasion intermediates (SEI) that commit recombination to the Class I crossover pathway. These events are biased toward the use of a homologous chromosome as a donor for repair (Fig. 1a; Schwacha and Kleckner 1997; reviewed in Brown and Bishop 2014 and Hunter 2015). The SEI forms at roughly the same time as the synaptonemal complex, a structure that forms between homologs and is thought to aid in the removal of chromosomal interlocks during the homology search (Hunter and Kleckner 2001; Börner et al. 2004; Fung et al. 2004; Snowden et al. 2004; Lynn et al. 2007; Storlazzi et al. 2010; De Muyt et al. 2018; Pyatnitskaya et al. 2019). Following DNA synthesis steps and capture of the second end of the broken homolog, SEIs mature into symmetric double-Holliday junctions (dHJs) that are resolved asymmetrically to form crossovers through the actions of the mismatch repair (MMR) family member Mlh1–Mlh3 and the exo/endonuclease Exo1 to yield crossover products (Allers and Lichten 2001; Zakharyevich et al. 2012; reviewed in Pyatnitskaya et al. 2019). Crossovers dependent on Mlh1–Mlh3 bear the hallmarks of the Class I pathway; they are widely and evenly spaced (crossover interference) and display crossover assurance, a mechanism which ensures that each homolog pair receives at least one crossover (Szostak et al. 1983; Sym et al. 1993; Schwacha and Kleckner 1994; Sym and Roeder 1994; Schwacha and Kleckner 1995; Novak et al. 2001; Shinohara et al. 2003; Börner et al. 2004; Hillers 2004; Jones and Franklin 2006; Martini et al. 2006; Mancera et al. 2008; Shinohara et al. 2008; Zakharyevich et al. 2012; Chakraborty et al. 2017).

Fig. 1.

Model for a canonical resolvase role for Mlh1–Mlh3 in meiotic recombination and rationale for using mlh3-42 and mlh3-54 as baits (MMR⁻ CO⁺) in this study. a) Model for the Type I CO pathway in yeast meiotic prophase. Phase I, homology search: Following Spo11-mediated DSB formation and 5′ to 3′ resection of DSBs, a recombinase filament (Dmc1 filament initiates at the end of the Rad51 filament) forms at the 3′ resected ends. This filament initiates a search for the allelic locus on the homologous donor chromosome. Phase 2, CO designation: ZMM-facilitated stabilization of SEIs, D-loop migration, and second-end capture commit recombination intermediates to the Type I crossover pathway. Phase 3, dHJ resolution: Mlh1–Mlh3 and Exo1 participate in biased resolution of dHJs to produce crossover products. b) Crystal structure of the Mlh1–Mlh3 C-terminus (PDB 6RMN; Dai et al. 2021 ) with mutated residues highlighted by space filling (R552A, D553A, K555A, D556A for mlh3-42 and R682A, E684A for mlh3-54). c) Three regions within the Mlh3 C-terminus predicted to mediate the formation of an Mlh1–Mlh3 polymer (Dai et al. 2021) with amino acids changed in mlh3-42 highlighted as sticks for resolution purposes.

How might Mlh1–Mlh3 act in crossover resolution? Mlh3 contains a highly conserved (DQHA(X)2E(X)4E) endonuclease motif within its C-terminus (Kadyrov et al. 2006; Nishant et al. 2008). Disruption of this motif (mlh3-D523N) abolishes the endonuclease activity of Mlh1–Mlh3 in vitro and confers mlh3 null-like defects in crossing over (Nishant et al. 2008; Ranjha et al. 2014; Rogacheva et al. 2014). Curiously, Mlh1–Mlh3 does not display the biochemical activities characteristic of an intrinsic HJ resolvase. It is capable of binding model HJ substrates, but such binding inhibits its endonuclease activity. Rather, Mlh1–Mlh3 endonuclease activity is activated in vitro by polymer formation on large DNA substrates and mlh3 mutations predicted to disrupt polymer formation confer crossover defects (Hall et al. 2001; Manhart et al. 2017; Dai et al. 2021). Mlh1–Mlh3 and Exo1 form a constitutive complex throughout meiosis (Sanchez et al. 2020) and recent in vitro studies showed that Mlh1–Mlh3 endonuclease activity is stimulated by Msh4–Msh5, Exo1, and the DNA polymerase processivity factor PCNA (Cannavo et al. 2020; Kulkarni et al. 2020). While we do not have a clear understanding of how Mlh1–Mlh3 endonuclease activity is directed to form crossovers, several dHJ resolution models have been presented that incorporate the above Mlh1–Mlh3 activities and a role for Exo1 in protecting DNA nicks (Manhart et al. 2017; Marsolier-Kergoat et al. 2018; Kulkarni et al. 2020; Gioia et al. 2023).

Mlh1–Mlh3 is thought to act late in prophase I during dHJ resolution. However, several studies have suggested possible early roles for Mlh1–Mlh3 in meiosis: (1) Sanchez et al. (2020) reported in baker's yeast that Mlh3 localizes to DSB hotspots early in meiotic prophase, showing kinetics similar to the ZMM protein Zip3 (Serrentino et al. 2013). (2) Al-Sweel et al. (2017) observed in baker's yeast that the median gene conversion tract length associated with crossovers was longer in mlh3Δ compared to wild-type. (3) Marsolier-Kergoat et al. inferred through an analysis of genome-wide heteroduplex DNA patterns in baker's yeast that Mlh1–Mlh3 acts to limit inward branch migration (Marsolier-Kergoat et al. 2018; also see Martini et al. 2011; Peterson et al. 2020; Ahuja et al. 2021). (4) In mice, MLH3 forms foci in early pachynema prior to MLH1–MLH3 foci formation (Kolas et al. 2005). (5) Premkumar et al. (2023) obtained data in mice consistent with MLH3 playing an early structural role in differentiating SEIs further toward Class I resolution. (6) Work in the filamentous fungus Sordaria uncovered a role for Mlh1 in the resolution of chromosome entanglements during zygotene, prior to crossover formation in pachytene (Storlazzi et al. 2010). While the above studies are suggestive of early roles for Mlh1–Mlh3, specific mechanisms to support such an idea have not been presented.

To better understand the role of Mlh1–Mlh3 in meiotic prophase, we performed a gene dosage screen aimed at identifying factors that genetically interact with MLH3 in its meiotic procrossover role. Nine genes showed the strongest gene dosage phenotypes; we then narrowed the list further based on novelty of the interaction. Through this approach we obtained genetic and biochemical evidence that Mlh3 interacts with the recombinase Dmc1. Partial complementation of Mlh3 crossover functions was observed when MLH3 was expressed under the control of the CLB1 promoter. These observations led us to a model in which the Dmc1 recombinase filament provides a platform for Mlh1–Mlh3 polymer formation and subsequent endonuclease activation during crossover resolution.

Materials and methods

Media and chemicals

Saccharomyces cerevisiae yeast strains used in this study were grown at 30°C in yeast extract–peptone–2% dextrose (YPD) or yeast extract–peptone–2% lactate (YPL) media (Rose et al. 1990). When required, geneticin (Invitrogen, San Diego), nourseothricin (Werner BioAgents, Germany), or Hygromycin B (Invitrogen, San Diego) were added to media at recommended concentrations (Goldstein and McCusker 1999). Sporulation media was prepared as described (Argueso et al. 2004).

Plasmids

Plasmids used in this study are shown in Supplementary File 1 and oligonucleotides used to construct plasmids are listed in Supplementary File 2. All oligonucleotides used in this study were purchased from Integrated DNA Technologies (Coralville, IA). The yeast genes present on these plasmids are from the SK1 strain background (Kane and Roth 1974).

pEAI522 (7.8 kb; P_CLB1-MLH3) was created through a six fragment HiFi DNA Assembly (New England Biolabs) with BamHI digested pUC18 as the backbone and the following five fragments: (1) 560 bp homology 5′ of MLH3 ORF amplified from the SKI S. cerevisiae genome with AO4954/AO4955, (2) 540 bp CLB1 promoter amplified from SKI genome with AO4956/AO4957, (3) full MLH3 ORF consisting of 2,288 bp with 100 bp downstream of the stop triplet amplified from SKI genome with AO4958/AO4959, (4) KANMX amplified from genomic prep of KANMX containing SKI with AO4960/AO4961, (5) 560 bp homology downstream of MLH3 ORF amplified from SKI genome with AO4962/AO4963. Digestion of pEAI522 with XbaI and EcoRI yielded the DNA fragment used for integration.

pEAI523 (7.7 kb; pGAL1-MLH3) was created in the same manner as pEAI522 except for fragment (2) which was the GAL1 promoter amplified from SKI genome with AO4965/AO4966. To yield compatible fragment overhangs, 560 bp homology 5′ of MLH3 ORF was amplified from SKI genome with AO4954/AO4964 (fragment 1); likewise, the full MLH3 ORF consisting of 2,288 bp with 100 bp downstream of the stop triplet amplified from SKI genome with AO4967/AO4959 (fragment 3). All other fragments are identical to pEAI522.

pEAI524 (8.4 kb; pGAL1-MLH3-13XMyc) was created using a two fragment HiFi assembly. pEAI523 was amplified with AO5465/AO5466 to serve as the backbone. A 4.7 kb insert was amplified from pFA6a (Bahler et al. 1998; Longtine et al. 1998) with AO5467/AO5468 to fuse the 13XMyc tag to the C-terminal residues of MLH3.

pEAM370 (8.9 kb; pADH1-MLH1-2µ-URA3) was designed by a two-step HiFi assembly procedure. First, the MLH1 ORF along with 500 bp flanking sequence 5′ relative to the MLH1 start codon was amplified from the SK1 genome with AO5249/AO5250 and inserted into pRS426 at EcoRI using HiFi assembly. In the second step, an ADH1 promoter, amplified from pGADT7 (Supplementary File 1) with AO5375/AO5376, was inserted into the pRS426 2µ plasmid (Christianson et al. 1992) containing MLH1 which itself was amplified with AO5373/AO5374 to delete the native MLH1 promoter. The resulting plasmid contained an ADH1 promoter driving expression of MLH1 in a 2µ plasmid.

pEAM371 (8.9 kb; pADH1-MLH3-2µ-URA3) was designed in the same manner as pEAM370 apart from the MLH1 ORF being replaced by the MLH3 ORF. The MLH3 ORF along with 500 bp flanking sequence 5′ relative to the MLH3 start codon was amplified from the SK1 genome with AO5251/AO5252 and inserted into pRS426 at EcoRI using HiFi assembly. The ADH1 promoter was amplified from pGADT7 with AO5379/AO5380 inserted into the pRS426 2µ plasmid containing MLH3 amplified with AO5377/AO5378 to delete the native MLH3 promoter.

pEAM372 (12 kb; pADH1-MLH1-MLH3-2µ-URA3) was designed with HiFi assembly using fragments from pEAM370 and pEAM371. pEAM370 was amplified with AO5410/AO5411 whereas the pADH1-MLH3 fragment from pEAM371 was amplified with AO5412/AO5413 to insert pADH1-MLH3 downstream of pADH1-MLH1. The resulting plasmid contained both MLH1 and MLH3 ORFs in tandem, both driven by ADH1 promoters in a 2µ plasmid.

pEAM373 (pADH1-DMC1-2µ-URA3) was designed with HiFi assembly using pEAM370 as the backbone, amplified with AO5414/AO5415. The DMC1 ORF was then amplified from the SKI genome with AO5416/AO5417. The resulting plasmid contained the DMC1 ORF driven by the ADH1 promoter in a 2µ plasmid.

pEAA734 (6.6 kb; DMC1, CEN6-ARSH4, URA3) was constructed using two-fragment HiFi assembly (New England Biolabs). The DMC1 open reading frame with 300 bp upstream and 296 bp downstream sequences was amplified from the SK1 genome with AO4920/AO4921 and cloned by HiFi assembly into pRS416 (Christianson et al. 1992) digested with BamHI to generate pEAA734. dmc1 mutations were introduced into pEAA734 using Q5 site directed mutagenesis (New England Biolabs).

Yeast two-hybrid vectors pEAM350-369 (Supplementary File 1) were assembled using HiFi assembly primers (Supplementary File 2) designed for N-terminal GAL4 activation domain fusions (pGADT7), C-terminal GAL4 activation domain fusions (pGADCg), N-terminal GAL4 binding domain fusions (pGBKT7), and C-terminal GAL4 binding domain fusions (pGBKCg) to the complete open reading frames of MLH1, MLH3, DMC1, RAD51, and PMS1. MLH-N-terminal domain fusions (NTT) to these vectors were constructed that contained amino acids 1 to 504 of MLH1 and 1 to 476 of MLH3. MLH C-terminal domains fusions (CTT) to these vectors were constructed that contained amino acids 345 to 769 of Mlh1 and 375 to 715 of Mlh3.

The above plasmid sequences were confirmed with Sanger sequencing or Oxford Nanopore long read sequencing using Plasmidsaurus (https://www.plasmidsaurus.com).

Strains

Isogenic yeast strains used in this study were from the SK1 background (Kane and Roth 1974) except for strains used for RTK1 tetrad analysis which are SK1 congenic (Argueso et al. 2004; Supplementary File 3). Gene knockouts in yeast were generated as described below apart from EAY4651-EAY4653 (MATa, mlh2Δ::KANMX), which were generated by integrating a XbaI and EcoRI restriction digest fragment of pEAI480 (Supplementary File 1) using standard methods (Rose et al. 1990; Gietz et al. 1995; Goldstein and McCusker 1999). Genomic integration events were confirmed by PCR amplification of genomic extracts as described previously (Hoffman and Winston 1987).

Mutant dmc1 alleles were generated through Q5 site-directed mutagenesis (New England Biolabs) of pEAA734 (ARS::CEN, URA3) and transformed into EAY4911-EAY4913/EAY4914-EAY4916 diploids. Promoter swaps were achieved through integration of the appropriate plasmids (Supplementary File 1) designed as described above. Plasmid selection was maintained though omission of uracil in all growth and sporulation media.

EAY5071-EAY5075 strains contain the MLH3-13xMYC allele (13xMyc tag fused to the C-terminal residue of Mlh3). These strains were created by transforming the parental strains (SKY3576, SKY3575; Supplementary File 3) with a DNA fragment generated by PCR amplifying pFA6a (Bahler et al. 1998; Longtine et al. 1998) with AO5333 and AO5334 (Supplementary File 2). Transformants were selected on YPD media containing G418 and the presence of the MLH3-13xMYC allele was confirmed by DNA sequencing.

Spore-autonomous fluorescence assay

Crossing over in the CEN8-THR1 interval on Chromosome VIII was assessed in SKY3576/SKY3575 derived diploids using spore-autonomous fluorescence reporter constructs (Thacker et al. 2011). Briefly, haploids of opposite mating type were struck to singles on YPL plates and incubated at 30°C for 2–3 days. Single colonies for each mating type were then mixed in 25 µL of ddH₂O, spotted on YPD plates and incubated at 30°C for 5 h. Diploids were selected for by streaking the spot to single colonies on minimal media lacking tryptophan and leucine and incubated at 30°C for 48 h. Diploids were then patched on YPD and allowed to form a lawn before being transferred to sporulation media plates. After 2–4 days, spores were treated with 0.5% NP40 and briefly sonicated before analysis using a Zeiss AxioImager.M2. At least 500 tetrads for each genotype were counted to determine the % tetratype (Supplementary Files 4 and 5). At least two independent transformants were studied per genotype and the vast majority of genotypes were analyzed on separate days. When appropriate, ARS-CEN plasmid maintenance was achieved through growth and sporulation media lacking the necessary nutrient to ensure plasmid retention.

Tetrad analysis

Diploids derived from SK1 congenic strains EAY1108/EAY1112 were sporulated using the zero-growth mating protocol (Argueso et al. 2003). Haploid parental strains were mixed in 25 µL ddH₂O, mated on YPD for 5 h at 30°C, and then struck to singles on minimal plates lacking leucine and tryptophan and incubated at 30°C for 48 h. Diploids were then patched on YPD and allowed to form a lawn before being transferred to sporulation media plates and grown for 2–4 days at 30°C. Tetrads were dissected on minimal complete plates and then incubated at 30°C for 3–4 days. Spore viability was then assessed (Supplementary File 6). Spore clones were replica-plated onto relevant selective plates and assessed for growth after an overnight incubation. Genetic map distances were determined by the formula of Perkins (1949; Supplementary File 7). Statistical analysis was done using the Stahl Laboratory Online Tools (https://elizabethhousworth.com/StahlLabOnlineTools/).

Gene dosage screen

Gene knockout transformation cassettes were generated consisting of an MX antibiotic resistance marker flanked by 300 bp of upstream and downstream homology with respect to the open reading frame (ORF) of each gene of interest. These cassettes were amplified by PCR from genomic preps of the appropriate strains from the Saccharomyces genome deletion project (Giaever and Nislow 2014 ). In this collection, each ORF was replaced with KANMX4. Other MX knockout cassettes (e.g. NATMX, hphMX6) were generated through marker swapping of appropriate deletion strains.

EAY3486 (Supplementary File 3), a mlh3Δ::NATMX strain carrying a gene encoding a cyan fluorescent protein on chromosome VIII, was transformed with the PCR amplified knockout cassette. Cells were then plated on YPD-G418 plates and grown at 30°C for 3 days. At least two independent transformants were verified by confirming resistance to G418 and nourseothricin and PCR amplification using genomic preps of G418^r/NAT^r transformants. For PCR verification, primers annealing 350 bp upstream and downstream of the ORF of the gene of interest were utilized to ensure integration at the proper locus. Haploids were then crossed to four strains each carrying a gene encoding a red fluorescent protein on chromosome VIII. These four strains are as follows: EAY3252 (MLH3), EAY3255 (mlh3Δ), EAY3572 (mlh3-42), and EAY3596 (mlh3-54). Diploids were isolated by selecting on media lacking tryptophan and leucine and analyzed in the spore-autonomous fluorescence assay described below.

Our criteria for allele-specific interactions were those in which there was little to no change in percent tetratype in either an MLH3 and mlh3Δ background, but there was a significant drop of percent tetratype in either mlh3-42 or mlh3-54 backgrounds. Significance was assessed within backgrounds by chi-squared test. To minimize ⍺ inflation due to multiple comparisons, we applied a Benjamini–Hochberg correction at a 5% false discovery rate (Benjamini and Hochberg 1995). At least two independent transformants were analyzed for each genotype.

Two-hybrid analysis

Yeast two-hybrid assays were performed as previously described (Bentolila et al. 2021). Yeast two-hybrid expression strains, PJ69-4a and PJ69-4α (Supplementary File 3), were individually transformed with GAL4 activation domain fusion vectors or GAL4 binding domain fusion vectors, respectively. Vectors lacking any protein fusions served as negative controls. Single haploid transformants were selected for by plating on selective media. Crosses were performed as previously described and diploids were selected for by streaking to singles on minimal media plates lacking leucine and tryptophan. Interaction assays were performed by growing test diploids overnight in 3 mL minimal media lacking leucine and tryptophan. The following day, cultures were diluted in ddH₂O to OD₆₀₀ 0.5, 0.05, and 0.005 and spotted on -leucine, -tryptophan plates to assess plating efficiency and -leucine, -tryptophan, -histidine, -adenine plates to assess growth after 3 days of incubation at 30°C.

In vitro co-immunoprecipitation

yMlh1-FLAG-His₁₀-Mlh3-HA was purified using methods described in Rogacheva et al. (2014). yMlh1-FLAG-Pms1 was purified as described in Plys et al. (2012). 6XHis-yDmc1 and 6XHis-yRad51 (6xHis tag in Rad51 removed by a SUMO protease) were purified using previously published procedures (Steinfeld et al. 2019).

6xHis-hDMC1 was purified using the same procedure used to purify 6xHis-yDMC1 and is as follows. Rosetta 2 Bl21 (DE3) cells bearing pET11C-6xHis-hDMC1 expression plasmid (a kind gift from Michael Sehorn) was grown to an OD₆₀₀ between 0.6 and 0.8 at 37°C. The temperature was lowered to 16°C and cells were induced with 0.5 mM IPTG overnight. Cells were harvested by centrifugation and stored at −80°C. Cells were resuspended in buffer A (30 mM Na-Hepes pH 7.5, 1000 mM KCl, 5 mM MgCl₂, 0.01% NP-40, 2 mM ATP, 10% glycerol, 5 mM 2-mercaptoethanol, 0.2 mM PMSF, and Protease Inhibitor cocktail) and lysed by freeze–thaw. Cell lysate was sonicated on ice with 10 cycles with 65% duty cycle 15 s ON and 45 s OFF. Lysate was centrifuged at 10,000 × g for 45 min at 4°C. Clarified extract was mixed with preequilibrated Talon resin and incubated in batch for 1 h at 4°C. After 1 h the resin was centrifuged and washed 2× with buffer A, followed by two times washes with buffer B (30 mM Na-Hepes pH 7.5, 200 mM KCl, 5 mM MgCl₂, 0.01% NP-40, 2 mM ATP, 10% glycerol, 5 mM 2-mercaptoethanol, 0.2 mM PMSF, and Protease Inhibitor cocktail). The resin was then added to a disposable column. The protein was eluted in buffer C (30 mM Na-Hepes pH 7.5, 1,000 mM KCl, 5 mM MgCl_2, 0.01% NP-40, 2 mM ATP, 200 mM imidazole, 10% glycerol, 5 mM 2-mercaptoethanol, 0.2 mM PMSF, and Protease Inhibitor cocktail). Fractions were analyzed by SDS–PAGE and the peak fractions were pooled. The protein was further purified using a HiTrap Heparin HP column. The column was resolved using a 30 CV 20–100% gradient of buffer D (30 mM Na-Hepes pH 7.5, 0 mM KCl, 5 mM MgCl_2, 0.01% NP-40, 2 mM ATP, 10% glycerol, 5 mM 2-mercaptoethanol, and 1 mM EDTA) and buffer E (30 mM Na-Hepes pH 7.5, 1,000 mM KCl, 5 mM MgCl_2, 0.01% NP-40, 2 mM ATP, 10% glycerol, 5 mM 2-mercaptoethanol, and 1 mM EDTA). Fractions were analyzed by SDS–PAGE. Peak fractions were pooled and diluted to ∼200 mM KCl and loaded on a HiTrap Q HP column. The column was resolved using a 30 CV 20–100% gradient of Buffer F (30 mM Na-Hepes pH 7.5, 0 mM KCl, 5 mM MgCl_2, 0.01% NP-40, 2 mM ATP, 10% glycerol, 5 mM 2-mercaptoethanol, and 1 mM EDTA) and Buffer G (30 mM Na-Hepes pH 7.5, 1000 mM KCl, 5 mM MgCl_2, 0.01% NP-40, 2 mM ATP, 10% glycerol, 5 mM 2-mercaptoethanol, and EDTA). The fractions were analyzed by SDS–PAGE and peak fractions were pooled and concentrated by centrifugation at ∼1,100 × g at 4°C using a 10 kDa MWCO Vivaspin 6 column. The concentrated protein was stored in single use aliquots at −80°C.

Invitrogen Dynabeads Protein G Immunoprecipitation Kit (Catalog No. 10007D) was used for immunoprecipitations involving purified yMlh1-Mlh3, yMlh1-Pms1, yDmc1, yRad51, and hDmc1. For all anti-FLAG immunoprecipitations, 2 µg of purified recombinase (yDmc1, yRad51, and hDmc1) were mixed with 1 µg MLH complex (yMlh1-Mlh3 or yMlh1-Pms1) in 100–400 µL total volume Binding/washing buffer at 4°C for 30 min. For DNase treated samples, two units of DNase I (NEB) was added along with MgCl₂ to a final concentration of 2.5 mM. DNase I-treated samples were incubated at 37°C for 5 min before incubating protein mixtures at 4°C for 30 min. If 1 µg lambda DNA (NEB) was added to check for DNA degradation, 5 µL of DNase-treated samples was removed after 37°C incubation, treated with Proteinase K, and incubated at room temperature for 15 min before running on an 1% agarose gel. While protein mixtures were incubating at 4°C, 50 μL (1.5 mg) Dynabeads magnetic beads were prepared by removing the supernatant and incubating the beads at room temperature for 20 min with mild agitation in a total volume of 200 µL Binding buffer (pH 7.4: 137 mM NaCl, 2.7 mM KCl, 10 mM Na₂HPO₄, 1.8 mM KH₂PO₄, 0.02% Tween-20) with 5 µg ANTI-FLAG M2 antibody (Sigma, catalog number F1804). Beads were then washed once with Binding buffer before addition of the premixed proteins. The protein-bead-Ab mixture was incubated at room temperature for 30 min with mild agitation. The protein-bead-Ab mixture was then placed on a magnetic rack to remove the flow through fraction containing unbound proteins. Three times sample buffer supplemented with 50 mM DTT was added to the unbound fraction and proteins were loaded on a 10% SDS–PAGE gel. The protein-bead-Ab mixture was then washed 3× with 200 μL washing buffer (1× PBS, pH 7.4: 137 mM NaCl, 2.7 mM KCl, 10 mM Na₂HPO₄, 1.8 mM KH₂PO₄) and re-suspended in 100 μL washing buffer. The bead suspension was then transferred to a clean tube to prevent co-elution of protein bound to the tube wall. After removal of the supernatant, the beads were resuspended in 30 µL elution buffer and 15 µL 3× sample buffer with 50 mM DTT was added. The beads eluate was heated at 99°C for 6 min and loaded on a 10% SDS–PAGE gel. Bands were imaged and quantified by ImageJ (Schneider et al. 2012) after Coomassie Blue staining.

Meiotic time courses

SK1 yeast diploids were struck to singles on YPL plates and incubated at 30°C for 3 days. A single colony was used to inoculate a 3 mL YPD culture which was grown overnight to saturation. Seven hundred microliter of the saturated YPD culture was subcultured into 400 mL YPA (1% potassium acetate; 2% peptone; 1% yeast extract) and grown for 16–17 h at 30°C. For phenotypic analyses (e.g. spore autonomous assay of crossing over at CEN8::THR1), 50 mL of presporulated cells were washed with water and resuspended in 25 mL CSHSPO (1% potassium acetate; 0.1% yeast extract; 0.05% glucose) in a 250 mL flask to ensure adequate aeration. For protein expression or co-immunoprecipitation experiments, 400 mL of presporulated cells was washed with water and resuspended in ∼240 mL CSHSPO in a 2-L flask at an OD₆₀₀ of ∼1.5–2.5. All liquid sporulations were incubated at 30°C. T = 0 for all time-courses denotes the time when sporulation cultures were transferred to a 30°C shaker. When appropriate, β-estradiol (Millipore-Sigma) was added to 10 nM (complementation) or 1 µM (overexpression). To monitor meiotic progression, ∼500 µL aliquots from each sporulation culture were collected at appropriate timepoints. Cells were then fixed in 900 µL 40% EtOH, 0.1 M Sorbitol and either incubated at room temperature for 30–60 min or stored at 4°C. After fixation, cells were washed twice with 1× PBS. Two µL 0.5% NP40 and 1 µL DAPI (0.1 mg/mL stock) were added to cells prior to imaging and counting with a Zeiss AxioImager.M2. The number of cells with 1, 2, 3/4 nuclei were recorded. At least 600 cells were counted per genotype per timepoint except for the 0-h timepoint where at least 300 cells were counted. Two to four independent transformants were analyzed per genotype and experiments were performed on separate days to avoid batch effects.

Meiotic whole cell protein analysis

At appropriate timepoints, 10 OD₆₀₀ units of cells were collected from sporulation cultures (∼30–50 mg cell mass). Cells were collected by centrifugation, washed once with 1 mL of 1× TE (10 mM Tris 8.0, 1 mM EDTA) + 1 mM PMSF, and vortexed for 15 min at 4°C with 100 µL acid-washed glass beads (500 µm, Sigma G8772) and 200 µL lysis buffer (10% glycerol, 50 mM Tris–HCl pH 7.5, 0.2% NP-40, 150 mM NaCl, 5 mM EDTA, 1 mM PMSF, 1× Halt protease inhibitor cocktail (Thermo)). Lysate was cleared by centrifugation for 5 min at 4°C and 125 µL of cleared lysate was added to 50 µL 3× SDS protein sample buffer. Protein concentrations were determined with Bradford reagent (Thermo; Bradford 1976) and 20 µg total protein was loaded onto a 10% SDS–PAGE gel.

Meiotic co-immunoprecipitation

1.2 × 10⁹ cells were harvested at the desired time-point in meiosis and washed with 1× TE + 1 mM PMSF. Cells were lysed in ∼2 mL lysis buffer (10% glycerol, 50 mM Tris–HCl pH 7.5, 0.2% NP-40, 150 mM NaCl, 5 mM EDTA, 1 mM PMSF, 1× Halt protease inhibitor cocktail (Thermo)); 0 or 125 Units/mL Benzonase (Sigma) with ∼600 µL glass beads in a bead beater four times, 5 min each, with at least 5 min rest on ice between bead beating rounds. Lysate was cleared by centrifugation in a microfuge at 13,000 × g for 5 min at 4°C. 25 µL of supernatant was collected for input and mixed with 12.5 µL 3× sample buffer containing 50 mM DTT. Ten microliter (10 μg) of anti-Myc (clone 4A6, Sigma) was added to lysate and incubated at 4°C for 1 h with rotation. One hundred microliter of Dynabeads Protein G were washed twice 2:1 with lysis buffer. Lysate/antibody mixture was then added to washed magnetic beads and incubated at 4°C for 3 h with rotation. Beads were washed four times with 500 µL of lysis buffer. On the final wash step, the bead suspension was transferred to a clean tube. Washed beads were resuspended in 35 µL 3× SDS protein sample buffer containing 50 mM DTT and heated at 95°C for 10 min. The bead eluate was loaded onto a 10% SDS–PAGE gel.

Western blot analysis

For both whole cell protein extract and immunoprecipitation analyses proteins electrophoresed in SDS–PAGE were transferred to a 0.2 µm PVDF membrane (Thermo) at either 100 V for 2 h, 70 V for 4 h, or 30 V overnight. Proteins tagged with the Myc epitope were detected with mouse anti-Myc (1:1,000; clone 4A6, Sigma) and peroxidase conjugated antimouse secondary antibody (1:20,000, Sigma). Dmc1 was detected with goat anti-Dmc1 (1:2,000; kindly provided by Doug Bishop) and peroxidase conjugated antigoat (1:5,000; Thermo). Glucose-6-phosphate dehydrogenase was detected with rabbit anti-G6PDH (1:10,000; Sigma) and peroxidase conjugated anti-rabbit (1:20,000, Invitrogen). Signal was detected with Clarity Western ECL Substrate (Bio-Rad) with either film or ChemiDoc MP (Bio-Rad) and analyzed with Image Lab software (Bio-Rad).

Results

A dosage suppression screen for MLH3 interactors reveals known and novel interactions

We performed a screen in mlh3–42 (mlh3-R552A, D553A, K555A, D556A)/mlh3Δ and mlh3-54 (mlh3-R682A, E684A)/mlh3Δ strains in which 34 candidate genes were reduced in dosage from two copies to one. This is considered a double-haploinsufficiency screen because the mlh3 alleles in the test conditions are also present in one copy. The mlh3 alleles were chosen because they conferred separation of function phenotypes that we hypothesized would reveal synthetic phenotypes for crossing over with candidate factors that were also compromised for function. Strains bearing these alleles were proficient for meiotic crossing over but were defective in MLH3-dependent MMR and were disrupted for Mlh1–Mlh3 interactions as measured by yeast two-hybrid and protein purification analyses (Al-Sweel et al. 2017).

Both mlh3-42 and mlh3-54 map to the structured C-terminal domain of Mlh3 which binds to Mlh1 (Fig. 1b; PDB 6RMN; Dai et al. 2021). Residues 552–556 mutated in mlh3-42 map distal to the dimerization interface between Mlh1 and Mlh3 while residues 682–684 mutated in mlh3-54 map proximal to the interface. Dai et al. (2021) identified three regions within the Mlh3 C-terminus predicted to mediate the formation of an Mlh1–Mlh3 polymer; mutations in these regions disrupt meiotic crossing over. Interestingly, Mlh3 residues 552–556 map within a loop opposite to the second region (Fig. 1c). Based on the above structural information, we hypothesize that mlh3-54 destabilizes the canonical Mlh1–Mlh3 binding interface while mlh3-42 indirectly impairs Mlh1–Mlh3 functions by altering the conformation of the Mlh3 C-terminus to disrupt Mlh1–Mlh3 polymer formation.

The phenotypes exhibited by mlh3-42 and mlh3-54 mutants suggested to us that Mlh1–Mlh3 could be structurally stabilized by meiotic factors which promote its pro-crossover functions. An alternative hypothesis is that the pro-crossover function of Mlh1–Mlh3 is less dependent on complex stability than its MMR function. If our first hypothesis is correct, then lowering the protein abundance of factors that interact with Mlh3 would elicit allele-specific crossover defects. Because Mlh1–Mlh3 and Exo1 form a constitutive complex in meiotic prophase (Sanchez et al. 2020), we tested as proof of principle the effect of lowering gene dosage of EXO1 on crossing over in strains containing the mlh3-42 and mlh3-54 mutations.

We assessed effects of haploinsufficiency by measuring single crossover frequencies (tetratype class, abbreviated as TT) using spore autonomous fluorescent markers inserted in the 20-cM CEN8-THR1 interval on chromosome VIII (Fig. 2a; Thacker et al. 2011; Al-Sweel et al. 2017). Only TT events were scored because the marker configurations in the analyzed strains cannot distinguish nonparental ditype (NPD) from Meiosis I nondisjunction events. As shown in Fig. 2b, reduced EXO1 dosage did not affect crossing over in MLH3 and mlh3Δ strains, but significantly decreased crossing over in mlh3-42 (39% TT with two EXO1 copies compared to 29% TT with one) and mlh3-54 strains (37% TT with two copies compared to 29% with one). Reduced EXO1 dosage however, did not significantly impact the timing and efficiency of completing MI/MII divisions in the mlh3-42 and mlh3-54 strains (Fig. 2e, taking into account the standard error in biological replicates).

Fig. 2.

Gene dosage suppression analysis using mlh3-42 and mlh3-54 as baits. a) The spore-autonomous fluorescence assay was used to measure tetratypes (single crossover events) in the CEN8-THR1 interval on chromosome VIII (Thacker et al. 2011). In this strain construct NPD tetrads (all recombinant) cannot be distinguished from Meiosis I nondisjunction events and were not scored. b) Crossover analysis in the CEN8-THR1 interval on chromosome VIII expressed as percent tetratype for indicated EXO1 gene dosage in wild-type/mlh3Δ, mlh3Δ/mlh3Δ, mlh3-42/mlh3Δ, and mlh3-54/mlh3Δ strains. Dashed lines indicate wild-type, and mlh3Δ levels of crossing over. c) Crossover analysis in the CEN8-THR1 interval for indicated gene dosages of MLH3, EXO1, and exo1-MIP. Dashed lines indicate wild-type/wild-type, mlh3Δ/mlh3Δ, and exo1Δ/exo1Δ levels of crossing over. d) Crossover analysis in the CEN8-THR1 interval on chromosome VIII expressed as percent tetratype for indicated DMC1 gene dosage in wild-type/mlh3Δ, mlh3Δ/mlh3Δ, mlh3-42/mlh3Δ, mlh3-54/mlh3Δ, mlh3-42/mlh3-42, mlh3-54/mlh3-54, and mlh3-42/mlh3-54 strains. Dashed lines indicate wild-type/wild-type and mlh3Δ/mlh3Δ levels of crossing over. Significance in panels b to d was assessed according to χ²  P-values where P > 0.05 = n.s. (not significant), P < 0.05 = *, P < 0.01 = **, and P < 0.001 = ***. Blue asterisks indicate comparisons to wild-type/mlh3Δ and orange asterisks indicate comparisons to mlh3Δ/mlh3Δ. To account for multiple comparisons, we applied a Benjamini–Hochberg correction at a 5% false discovery rate (Benjamini and Hochberg 1995). See Materials and methods for details. e) Meiotic progression, as measured by completion of Meiosis I (MI), of sporulated cultures of the indicated genotypes. Error bars represent standard error for two to four biological replicates. Underlying data for panels b to e can be found in Supplementary File 4 and Supplementary File 5.

To complement the above analysis we examined MLH3–EXO1 interactions using the exo1-MIP allele (F447A, F448A) which disrupts interactions between Exo1 and Mlh1 and confers a meiotic crossover defect (Amin et al. 2001; Gellon et al. 2002; Tran et al. 2007; Zakharyevich et al. 2010). We detected an intermediate crossover defect (33% TT) in strains containing two copies of both exo1-MIP and MLH3; however, strains containing only single copies of exo1-MIP and MLH3 displayed crossover frequencies (23% TT) indistinguishable from mlh3Δ (Fig. 2c). We concluded that the reduction in crossover rates due to lowering the gene dosage of partner proteins in mlh3-42, mlh3-54 and exo1-MIP backgrounds was likely due to compromising the integrity of the Exo1–Mlh1–Mlh3 complex.

The above observations encouraged us to test the effect of gene dosage for 34 meiotic genes in the mlh3-42/mlh3Δ and mlh3-54/mlh3Δ backgrounds (Table 1; Supplementary File 3). Genes were chosen to represent various stages of meiotic recombination, including helicases, ZMMs, structure selective nucleases, synaptonemal complex factors, and those involved in the recombinase machinery and chromosome segregation. We generated haploid knockouts in the mlh3Δ strain EAY3486 and crossed the resulting strains to wild-type (EAY3252), mlh3Δ (EAY3255), mlh3-42 (EAY3572), and mlh3-54 (EAY3596) to generate diploids harboring single copies of the 34 genes. These strains also contained markers to detect crossing over in the CEN8-THR1 interval (Supplementary File 3). For all 34 genes we analyzed the statistical significance of reductions in crossing over in each of the four backgrounds, comparing two copies of the gene candidate to one copy, and categorized the strength of haploinsufficiency according to χ²  P-values, following Benjamini–Hochberg (Benjamini and Hochberg 1995) correction for multiple comparisons, where P > 0.05 = not haploinsufficient, P < 0.05 = mildly haploinsufficient, P < 0.01 = moderately haploinsufficient, and P < 0.001 = strongly haploinsufficient (Supplementary File 4). Lowered gene dosage rarely led to a significant drop in tetratype frequencies in the mlh3Δ backgrounds. The one exception was the Rad51 nucleoprotein filament disassembly gene SRS2 which conferred 14% TT frequency in the mlh3Δ background when present in one copy (P < 0.01; Krejci et al. 2003; Crickard et al. 2018).

Table 1.

Dosage suppression analysis in MLH3, mlh3Δ, mlh3-42 and mlh3-54 strains.

	P-value comparing two vs one gene copy
Gene tested	wild-type	mlh3Δ	mlh3-42	mlh3-54
Established Mlh1 and/or Mlh3 interactors
MLH1	n.s.	n.s.	n.s.	++
MLH2	n.s.	n.s.	n.s.	++
PMS1	n.s.	n.s.	++	+++
MSH4	n.s.	n.s.	+	++
MSH5	+	n.s.	+++	+++
EXO1	n.s.	n.s.	+++	+++
RTK1	n.s.	n.s.	+	++
CHD1	++	n.s.	+++	+++
CAF120	n.s.	n.s.	+	+
Crossover resolution
MUS81	n.s.	n.s.	+++	++
YEN1	n.s.	n.s.	n.s.	n.s.
SLX4	n.s.	n.s.	n.s.	n.s.
Homologous recombination
DMC1	n.s.	n.s.	+++	+++
MEI5	n.s.	n.s.	n.s.	+
RAD51	n.s.	n.s.	n.s.	n.s.
RAD54	n.s.	n.s.	++	n.s.
RDH54	++	n.s.	+++	+++
SGS1	n.s.	n.s.	++	n.s.
RMI1	+	n.s.	+++	+
MER3	+++	n.s.	+++	++
SRS2	n.s.	++	n.s.	+
Meiotic chromosome structure
HOP1	n.s.	n.s.	n.s.	n.s.
GMC2	n.s.	n.s.	n.s.	+
SPO16	n.s.	n.s.	n.s.	n.s.
ZIP1	n.s.	n.s.	n.s.	n.s.
ZIP2	n.s.	n.s.	n.s.	+
ZIP3	++	n.s.	+++	+++
ZIP4	n.s.	n.s.	+++	+
EMC11	n.s.	n.s.	n.s.	n.s.
RED1	n.s.	n.s.	n.s.	n.s.
REC8	n.s.	n.s.	+++	n.s.
PCH2	n.s.	n.s.	n.s.	n.s.
Homolog pairing and telomere clustering
NDJ1	n.s.	n.s.	n.s.	n.s.
CSM4	n.s.	n.s.	++	+++

Crossing over in the CEN8-THR1 interval on chromosome VIII was determined as % tetratype for the indicated gene present in two or one copies in wild-type/mlh3Δ, mlh3Δ/mlh3Δ, mlh3-42/mlh3Δ, and mlh3-54/mlh3Δ strains (see Supplementary File 4 for the complete data set). Differences in % tetratype in MLH3/mlh3 strains containing two vs one gene copy were assessed according to χ²  P-values where P > 0.05 = n.s. (not significant), P < 0.05 = +, P < 0.01 = ++, and P < 0.001 = +++. To account for multiple comparisons, we applied a Benjamini–Hochberg correction at a 5% false discovery rate (Benjamini and Hochberg 1995). Genes in bold indicate that there was no significant difference in % tetratype between two vs one gene copy in wild-type/mlh3Δ or mlh3Δ/mlh3Δ, but there was a significant difference in mlh3-42/mlh3Δ and mlh3-54/mlh3Δ.

Gene dosage effects were observed for 12 genes in the mlh3-42/mlh3Δ background (but not in MLH3/mlh3Δ or mlh3Δ/mlh3Δ) and 14 genes in the mlh3-54/mlh3Δ background (but not in MLH3/mlh3Δ or mlh3Δ/mlh3Δ; Table 1; Supplementary File 4; Supplementary Fig. 1). Nine of these genes (PMS1, MSH4, EXO1, RTK1, CAF120, MUS81, DMC1, ZIP4, CSM4) conferred dosage phenotypes in both mlh3 allele backgrounds. All nine genes in the Mlh1/Mlh3 interaction group showed a gene dosage effect with at least one mlh3 allele, including newly identified interactors CAF120, a component of the Ccr4-Not transcriptional regulatory complex, and the putative kinase RTK1 (Wild et al. 2019). Additionally, CSM4, which regulates telomere bouquet formation and chromosome pairing in meiosis, showed a gene dosage effect (Kosaka et al. 2008; Wanat et al. 2008) as did four factors that act in or stimulate strand invasion steps in meiotic recombination, DMC1, MEI5, RAD54, and SGS1. Lastly, we observed gene dosage effects for MSH5 (component of the Msh4–Msh5 complex; Pochart et al. 1997), CHD1 (Chd1 interacts with Mlh1–Mlh3; Wild et al. 2019), RDH54 (Rdh54 interacts with Dmc1; Dresser et al. 1997), RMI1 (component of the Sgs1–Top3–Rmi1 complex; Mullen et al. 2005), MER3 (Mer3 interacts with Mlh1–Mlh2; Duroc et al. 2017) and ZIP3 (part of a ZMM assembly complex that includes Zip4; Shinohara et al. 2008) in wild-type, mlh3-42, and mlh3-54 strains but not in mlh3Δ strains, suggesting that gene dosage interactions can be detected in some cases when MLH3 is fully functional.

The allele-specific gene dosage effect conferred by DMC1 was remarkably similar to that seen for EXO1 (compare Fig. 2d to Fig. 2b). Like the EXO1 dosage analysis, the efficiency of completing the MI/MII divisions and the timing of meiotic divisions was not dramatically impacted by lowering DMC1 dosage (Fig. 2e). In addition, we found that increasing the dosage of the mlh3-54 allele in strains containing one copy of DMC1 modestly increased crossing over, and when both mlh3-42 and mlh3-54 were expressed, crossing over was restored to wild-type (38%; Fig. 2d). We hypothesize that because mlh3-42 and mlh3-54 disrupt heterodimerization through distinct mechanisms (Fig. 1, b and c), a mix of the two alleles could offset their individual defects.

The above studies suggested to us that Mlh1–Mlh3 and Dmc1 physically interact (see below). Given the temporal separation between the functions of Dmc1 and Mlh3 in most models for meiotic recombination (Fig. 1a), we regarded the MLH3–DMC1 genetic interaction as unexpected and pursued it further. We performed an additional genetic analysis of RTK1 but concluded that Rtk1 is not essential for either Class I or Class II crossovers (Supplementary Fig. 2).

Yeast Mlh1–Mlh3 physically interacts with yeast Dmc1 and yeast Rad51 but not with human Dmc1

We purified yeast (y) Mlh1–Mlh3, yMlh1-Pms1, yRad51, yDmc1 and human (h) DMC1 and tested them for interactions by co-immunoprecipitation. yMlh1–Mlh3 robustly immunoprecipitated yDmc1 and yRad51 but not hDMC1 (Fig. 3, a and b). The yMlh1–Mlh3–yDmc1 interaction was retained in the presence of DNase I and in the absence of ATP, suggesting that the interaction was unlikely to be mediated through DNA (Supplementary Fig. 3; Busygina et al. 2013). Like yMlh1-Mlh3, yMlh1-Pms1 also immunoprecipitated yDmc1, suggesting that yMlh1 was likely important for interactions with yDmc1 (Fig. 3c).

Fig. 3.

In vitro co-immunoprecipitation reactions performed with yMlh1-FLAG-Mlh3, yMlh1-FLAG-Pms1, yDmc1, yRad51, and hDmc1. a) Unbound protein flow throughs showing that yDmc1, yRad51, and human Dmc1 do not bind nonspecifically to Dynabeads Protein G conjugated to anti-FLAG M2 antibody. b, c) Immunoprecipitation reactions containing the indicated purified proteins and Dynabeads Protein G conjugated to anti-FLAG M2 antibody. In all panels proteins were electrophoresed in 10% SDS–PAGE. The input lanes each show 10% of the mixture of the indicated proteins prior to immunoprecipitation. The entire amounts of each immunoprecipitation were loaded onto each lane. In panel (c) we calculated using ImageJ (Schneider et al. 2012) that 4.2% of yDmc1 was retained in the immunoprecipitate when Mlh1-Pms1 was included in the mixture (14% of the Mlh1-Pms1 input was in the immunoprecipitate), and 4.6% of yDmc1 was retained in the immunoprecipitate when Mlh1–Mlh3 was included (16% of the Mlh1–Mlh3 input was in the immunoprecipitate). See Materials and methods for details.

Our genetic analysis and in vitro pulldown experiments encouraged us to test if Mlh3 physically interacts with Dmc1 during meiosis through co-immunoprecipitation experiments. We used a Dmc1 polyclonal antibody to detect soluble Dmc1 present in cell extracts lysed in physiological (150 mM) NaCl concentrations (Fig. 4a; Materials and methods). We then used an anti-Myc antibody to immunoprecipitate Mlh3-13xMyc from cell extracts prepared from cultures harvested 0–8 h after transfer to sporulation media. Mlh3-13xMyc contains 13 copies of the Myc tag at its C-terminus. Strains bearing this allele were partially functional for meiotic crossing over (30% TT; Supplementary File 4). We only detected Mlh3 in Myc-tagged strains; untagged immunoprecipitates contained no cross-hybridizing bands. Interestingly, we could not detect a significant band corresponding to Dmc1 in Mlh3 pulldowns from extracts prepared 4 h after inducing sporulation, but a Dmc1-specific band was detected at 6 h (Fig. 4b). The Mlh3–Dmc1 interaction was retained when cell extracts were treated with Benzonase, confirming that DNA is not required for co-immunoprecipitation. We concluded that Mlh3 interacts physically with Dmc1 in meiotic prophase.

Fig. 4.

Mlh3 interacts with Dmc1 in vivo. a) Western blot showing specificity of yDmc1 antibody. Twenty microgram of whole cell extracts prepared in the indicated NaCl concentrations were loaded per lane. Five nanogram of purified yDmc1 was loaded in the yDmc1 lane. b) Time course pulldown of C-terminally tagged Mlh3-13xMyc at 0, 4, 6 h following transfer to sporulation media. Controls include dmc1Δ input (lane 1, left most lane) and inputs/pulldowns for untagged, wild-type strains (lanes 2–7). Inputs/pulldowns for Mlh3-13xMyc are in lanes 8–13. Lanes 14–15 show Mlh3-13xMyc pulldowns at 4 and 6 h when Benzonase was added to lysis buffer (125 Units/mL). c) Time course pulldown showing inputs/pulldowns for Mlh3-13xMyc at 0, 4, 6, 8 h. d) Meiotic progression, as measured by completion of Meiosis I, is presented for the Mlh3-13× Myc tagged strain compared to an untagged strain. The amount of sample loaded in panels (b) and (c) were normalized based on cell mass. In panels (b) and (c) “Input” lanes include 1.7% of the lysed cell supernatants from the indicated meiotic time points and “IP anti-Myc” lanes include the entire immunoprecipitate from the corresponding supernatants.

We tracked meiotic progression in sporulating cultures by DAPI staining of nuclei at 0, 4, 6, and 8 h postmeiotic induction. We detected the Dmc1–Mlh3 interaction at 6 and 8 h (Fig. 4c). Meiotic progression was delayed by ∼2 h relative to time courses performed with smaller culture volumes (compare Fig. 4d to Fig. 2e). This delay was also observed in the untagged wild-type strain. We inferred that a difference in methods used to sporulate small vs large meiotic cultures accounts for the 2 h delay rather than a phenotype associated with the tagged strain utilized for pulldown experiments.

Delayed expression of MLH3 results in partial complementation of MLH3 meiotic crossover functions

We performed an alanine scan mutagenesis of DMC1 with the goal of identifying dmc1 alleles which phenocopy the DMC1 haploinsufficiency phenotype observed in mlh3-42 and mlh3-54 backgrounds. This was performed by: (i) Aligning yDmc1 to human DMC1 (Supplementary Fig. 4a). (ii) Locating primarily charged residues that were not conserved between the yeast and human proteins. (iii) Mapping residues to the yDmc1 postsynaptic complex (PDB: 7EJ7) to confirm they were surface exposed and not part of Dmc1's self-interacting, or DNA binding interfaces (Steinfeld et al. 2019; Xu et al. 2021; Supplementary Fig. 4b, c). Twenty-one dmc1 alleles were analyzed for crossing over in the CEN8-THR1 interval. While one allele, dmc1-K101A, mildly reduced crossing over and deleting the first 21 amino acids conferred a null phenotype, no other mutations conferred significant defects in crossing over (Supplementary Fig. 4d).

Because the DMC1 mutagenesis focusing on charge surface residues did not identify specific residues that were likely to be important for Mlh1–Mlh3–Dmc1 interactions, and yeast two-hybrid analysis was unsuccessful in detecting such interactions (Supplementary Fig. 5), we performed experiments aimed at disrupting a Mlh3–Dmc1 interaction by modulating the timing of MLH3 expression. We hypothesized that delaying MLH3 expression to later timepoints in meiosis might impair Mlh3's ability to form a functional interaction with Dmc1.

To initially determine when MLH3 expression is critical in meiosis we placed MLH3 under the control of the GAL1 promoter in strains carrying a Gal4-Estrogen Receptor fusion (Benjamin et al. 2003). We induced MLH3 (untagged) expression with 10 nM β-estradiol at 0–6 h after inducing sporulation and monitored the effect of varying MLH3 induction on crossing over (Supplementary Fig. 6b). Inducing MLH3 upon transfer to sporulation media (defined as T = 0 h) resulted in full complementation of crossing over at CEN8-THR1 (39% TT), and a lack of β-estradiol induction (DMSO treatment) resulted in mlh3Δ levels (23% TT). MLH3 induction at 3 h posttransfer to sporulation medium was sufficient to impair crossing over (34% TT), and inductions at 4, 5, and 6 h after initiating sporulation led to crossover rates approaching no induction (25%, 24%, and 23% TT, respectively). Using strains carrying a Gal4-Estrogen Receptor fusion and MLH3-13XMyc under the control of the GAL1 promoter, we estimated that functional Mlh3-13XMyc levels were achieved 60–90 min post-β-estradiol induction (Supplementary Fig. 6), indicating a window from roughly 4 to 6 h in meiosis when Mlh3 protein expression was critical for its function. In these time courses at T = 7 h approximately 50% of cells had completed at least the MI division.

The above studies encouraged us to express MLH3 at a late stage in meiotic prophase to determine if such timing was sufficient for MLH3 function. We expressed MLH3 under the control of the CLB1 cyclin promoter, which is activated by the NDT80 transcription factor at a time when dHJs are resolved into crossovers (Chu and Herskowitz 1998; Brar et al. 2012) and monitored crossing over in the CEN8-THR1 interval. As shown in Fig. 5a, crossing over was somewhat impaired in P_CLB1-MLH3 (31.8% TT) but spore viability was indistinguishable from wild-type (Supplementary File 6), suggesting that late MLH3 expression can at least partially complement MLH3 functions. Lastly, compared to wild-type (MLH3/mlh3Δ), we observed a modest delay in completing MI/MII in mlh3Δ/mlh3Δ and P_CLB1-MLH3/mlh3Δ strains that was also seen in the absence of estradiol induction (DMSO treatment) time course (Fig. 5b, Supplementary Fig. 6c). An MI/MII delay in mlh3Δ strains was observed by Al-Sweel et al. (2017). A similar MI/MII delay was seen in exo1Δ by Zakharyevich et al. (2010), who showed that exo1Δ mutants displayed meiotic DSB kinetics similar to wild-type but a delay in the appearance of physical crossovers.

Fig. 5.

Modulating expression of MLH3 in meiotic prophase. a) MLH3 expression was placed under the control of the cyclin CLB1 promoter to drive late meiotic prophase expression. Crossing over in the CEN8-THR1 interval is presented for MLH3/mlh3Δ, mlh3Δ/mlh3Δ, and P_CLB1-MLH3/mlh3Δ strains. Asterisks indicate significance (P > 0.05 = n.s., P < 0.05 = *, P < 0.01 = **, and P < 0.001 = ***). b) Meiotic progression, as measured by completion of Meiosis I, is presented for the genotypes analyzed in Panel A. Error bars represent standard error for three to four biological replicates. Underlying data for panels (a) and (b) can be found in Supplementary File 4 and Supplementary File 5. c) A model proposing that Dmc1 in meiotic recombination intermediates nucleates the formation of stable Mlh1–Mlh3 polymers required for dHJ resolution.

Discussion

In this study we performed a screen for gene dosage effects in mlh3 backgrounds in which the stability of Mlh1–Mlh3 was predicted to be impaired (mlh3-42 and mlh3-54 strains). Our goal in this work was to identify new Mlh3 interactors. We screened 34 meiotic genes and identified established Mlh1 and Mlh3 interactors such as EXO1, MSH4, PMS1, and RTK1, a putative kinase of unknown function. Unexpectedly, one of the strongest genetic interactions involved the meiotic recombinase gene DMC1. We then showed that Mlh1–Mlh3 physically interacts with Dmc1 in vitro and in vivo.

The detection of an interaction between factors thought to act early (Dmc1) and late (Mlh1–Mlh3) in meiotic recombination is intriguing because cytological evidence in mice suggested that RAD51/DMC1 filaments are replaced with RPA prior to the recruitment of MLH1–MLH3 (Moens et al. 2002), and previous work in baker's yeast showed that Mlh1–Mlh3's acts late in meiotic prophase to resolve dHJs into crossovers in the Class I pathway (Zakharyevich et al. 2012; Arter et al. 2018). How can we reconcile the detection of Dmc1–Mlh1–Mlh3 interactions with the above observations? We hypothesize that in budding yeast Dmc1 and Mlh1–Mlh3 are present concurrently at least transiently on recombination intermediates. In such a model the formation of a Dmc1 filament acts to efficiently nucleate an Mlh1–Mlh3 polymer for its role in resolving dHJs into crossovers (Fig. 5c). It is important to note that the ZMM genes MSH4 and ZIP4 were identified in our dosage screen. How they and other ZMM genes, which are critical for the formation of Class I crossovers, could act in such a model remains unclear.

If Mlh3 interacts with Dmc1 on recombination intermediates, such a connection would likely be established shortly after strand invasion when Dmc1 is highly expressed and the local concentration of the two factors is likely to be high. Dmc1 levels reach a peak of ∼90,000 molecules/diploid cell in meiosis (Chan et al. 2019) while Mlh3 is constitutively expressed in meiosis at levels similar to that seen in mitosis (∼500 copies per haploid cell; Ho et al. 2018; Wild et al. 2019; Fig. 4). In support of this model, we observed: (1) Gene dosage interactions between DMC1 and mlh3 alleles that impacted meiotic crossing over. (2) Dmc1–Mlh3 physical interactions were detected at times in meiosis (6 and 8 h) when Dmc1 levels were high, but not at an earlier time point (4 h) when Dmc1 was expressed at somewhat lower levels (previous copy number studies suggest they are likely to be at much higher levels than for Mlh3; Fig. 4b, c; Chan et al. 2019; Ho et al. 2018). (3) P_CLB1-MLH3 only partially complemented the mlh3Δ phenotype, suggesting that the expression of Dmc1 and Mlh1–Mlh3 may need to be coordinated for crossover resolution. More precise time courses that include the measurement of physical recombination intermediates will be necessary to provide support for our model, and ChIP and immunofluorescence experiments would likely be useful to determine if Mlh3 recruitment to recombination intermediates is dependent on Dmc1. It will also be interesting to explore why an interaction between Dmc1 and Mlh1–Mlh3 was not observed at the 4-h time point; we speculate that the interaction requires additional factors or posttranslational modifications of Mlh1–Mlh3 and/or Dmc1 that are not present at this time point. Finally, because we have not obtained a mutation which disrupts the interaction between Mlh3 and Dmc1, our experiments do not rule out the possibility that other factors such as Mlh1 may be responsible for Mlh3 recruitment to recombination intermediates. Utilizing computational approaches such as AlphaFold (Varadi et al. 2022) may help us identify an Mlh1–Mlh3–Dmc1 protein–protein interaction that can be explored by mutagenesis.

Models proposing that Mlh3 plays primarily a late role in crossover resolution are supported by a study which showed that mlh3Δ mutants are functional for meiotic gene conversion (Abdullah et al. 2004). Further support was obtained by Arter et al. (2018) who showed that late meiotic expression of unrestrained alleles of the YEN1 resolvase (Yen1 does not act in the Class I crossover pathway) can restore crossing over in mlh1Δ and mlh3Δ strains. If Mlh1–Mlh3 play an exclusively late role in crossover resolution, how do we explain the partial P_CLB1-MLH3 complementation of the mlh3Δ crossover defect? We were encouraged to perform the CLB1 promoter studies after reading the work by Wild et al. (2019) who proposed that the chromatin remodeler Chd1 acts in meiotic prophase to activate the Mlh1-Mlh3 endonuclease to promote crossing over. They proposed such a model based on their observations that: (1) Exo1 associated with Chd1 at the prophase to metaphase I transition in meiosis. (2) chd1Δ, mlh1Δ, mlh3Δ, exo1Δ, and mlh1Δ chd1Δ double mutants displayed similar reductions in meiotic crossing over. (3) Mlh1, Mlh3, and Exo1 could still be recruited to pachytene chromosomes in chd1Δ. (4) Expression of CHD1 through the CLB1 promoter fully complement the chd1Δ crossover defect. One explanation for the partial mlh3Δ complementation by P_CLB1-MLH3 is that Mlh3 is that expression of MLH3 through the CLB1 promoter could result in a delay in achieving functional levels of Mlh3 (Mlh3 is normally constitutively expressed in meiosis) that would result in a competition with other resolvases such as Mms4-Mus81 that are activated when the Cdc5 kinase reaches functional levels (Matos et al. 2013). Such a scenario could result in some Class I dHJs being resolved by other resolvases (e.g. Mms4-Mus81) in an unbiased fashion that could result in the modest reductions in crossing over that were observed.

Haploinsufficiency as a tool to identify factors that interact in meiotic prophase

Haploinsufficiency screens have been performed to study factors that act in chromosome segregation in mitosis and meiosis (Baetz et al. 2004) and for expression of sporulation specific genes (Bungard et al. 2004). These approaches identified new interactions that were confirmed in biochemical and genetic assays. Our limited haploinsufficiency screen identified both previously known interactors of Mlh3 (see Table 1) and a new factor Dmc1, suggesting that a genome-wide screen, while labor intensive, would likely identify other components that interact with Mlh3 or other factors that act in crossing over.

Data availability

Strains and plasmids are available upon request. The authors affirm that all data necessary for confirming the conclusions of the article are present within the article, figures, tables, supplemental figures, and supplemental files. Supplementary Fig. 1 contains examples of haploinsufficiency analysis; Supplementary Fig. 2 displays the RTK1 analysis; Supplementary Fig. 3 shows co-immunoprecipitation analysis of yMlh1-Mlh3 and yDmc1 in the presence of DNase I; Supplementary Fig. 4 displays the effect of dmc1 mutations on meiotic crossing over in the CEN8-THR1 interval; Supplementary Fig. 5 provides a summary of yeast two-hybrid analysis; Supplementary Fig. 6 shows the effect of inducing GAL1-MLH3 with estradiol. Supplementary Files 1, 2, and 3, present plasmids, oligonucleotides, and strains used in this study, respectively; Supplementary File 4 displays the spore autonomous fluorescence assay data; Supplementary File 5 shows the raw data for the time-course phenotypic data; Supplementary File 6 shows the spore viability profiles; Supplementary File 7 displays the raw data for the Chromosome XV genetic map distance analysis of wild-type and rtk1Δ strains.

Supplemental material available at GENETICS online.

Funding

G.N.P., J.J.C., V.M., M.C.G., and E.A. were supported by the National Institute of General Medical Sciences of the National Institutes of Health (https://www.nih.gov/): R35GM134872. L.P. was funded by a fellowship from the Alfred P. Sloan Foundation and by National Institutes of Health grant F31GM145163. J.B.C. was supported by National Institute of General Medical Sciences of the National Institutes of Health (https://www.nih.gov/): R35GM142457. The content of this work is solely the responsibility of the authors and does not necessarily represent the official views of the National Institutes of Health. The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.