Casein kinase 1 (hhp) null mutants are meiotic recombination-deficient

To test for a possible role of casein kinase 1 homologs Hhp1 and Hhp2 in meiotic recombination of the fission yeast Schizosaccharomyces pombe, we measured recombination in hhp1Δ hhp2Δ double deletion mutants. Recombination was reduced by factors ranging from about 5 to 170, depending on the interval measured (Table 1). Both intergenic recombination (crossing over) and intragenic recombination (gene conversion) were reduced in the double mutant but only slightly in each single mutant, indicating that Hhp1 and Hhp2 have redundant roles in recombination, as previously reported for chromosome segregation [12]. Similar differential reductions, depending on the interval measured, are observed in cohesin- and LinE-deficient mutants [19,24,25], leading us to suspect that the Hhp substrate required for recombination is a cohesin subunit or LinE protein. Our results below bear out this suspicion.

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Table 1. Meiotic recombination depends on redundant Hhp1 and Hhp2 kinases.

https://doi.org/10.1371/journal.pgen.1005225.t001

An ATP analog-sensitive hhp mutant is DSB- and recombination-deficient even without analog

Because mitotic growth and viable spore yield are severely impaired in hhp1Δ hhp2Δ mutants [26,27], we used an Hhp1 mutant (hhp1-as encoding the Met⁸⁴ → Gly alteration) sensitive to 1-NM-PP1, an analog of the purine moiety of ATP, by alteration of its ATP-binding site [28,29]. In conjunction with hhp2Δ we could thereby allow Hhp function (in the absence of analog) or block Hhp function (in its presence). As expected from the results above, recombination was strongly reduced in the presence of the analog: we observed ~15- and 100-fold reductions in the two intervals tested (Table 2), comparable to the reductions seen in hhp1Δ hhp2Δ. Recombination was also reduced to essentially the same extent in the absence of the analog, a condition that allowed much higher viable spore yield and nearly wild-type chromosome segregation (Table 2 and S1 Table). This fortuitous result, presumably a reflection of hhp1-as allowing adequate phosphorylation of some substrates but not others, allowed us to conduct experiments under conditions allowing nearly wild-type mitotic growth and high viable spore yield. We discuss later the putative separation of functions of hhp1-as.

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Table 2. Even without ATP-analog, hhp1-as hhp2∆ mutant has strongly reduced meiotic recombination but only weakly reduced viable spore yield.

https://doi.org/10.1371/journal.pgen.1005225.t002

When we coupled the hhp1-as hhp2Δ mutations with rec12Δ, we obtained results indicating that, in the absence of analog, recombination but not Rec8 cleavage was defective in the hhp1-as hhp2Δ mutant. Because S. pombe has only three chromosomes and has a mechanism that enhances proper segregation of non-recombinant homologs [30], rec12Δ mutants form ~25% as many viable spores as wild type (Table 2) [31]. This yield was not further reduced by the hhp1-as hhp2Δ mutations in the absence of analog, but it was greatly reduced in its presence, as expected from failure of Rec8 to be cleaved under this condition [12,32]. In the absence of analog, the failure of rec12Δ to further reduce the viable spore yield of hhp1-as hhp2Δ indicates that without analog hhp1-as hhp2Δ blocks recombination but not chromosome segregation. We infer that one or more recombination-promoting protein(s) is not properly phosphorylated by Hhp1-as in the absence of the analog and that in the presence of the analog Rec8 is also hypo-phosphorylated.

Meiotic recombination requires both formation and repair of DSBs. hhp1Δ hhp2Δ mutants have DNA repair defects in vegetative cells [26]. To determine if meiotic DSB repair is blocked in the hhp1-as hhp2Δ mutant in the absence of the analog, we artificially introduced DSBs with the I-SceI homing endonuclease, controlled by the meiosis-specific rec12 promoter [33]. DSBs were introduced in the ade6 gene at the site of the ade6-3061 mutation, which can recombine with another mutation ade6-52 to generate Ade⁺ recombinants. In the absence of analog the frequency of recombinants was indistinguishable in wild type and in hhp1-as hhp2Δ, indicating that under this condition Hhp is not required for DSB repair (S2 Table). rec8Δ, however, reduced the recombinant frequency by a factor of ~3, suggesting that Rec8 is required for DSB repair of I-SceI DSBs, as it is at some chromosomal sites in budding yeast [6]. We infer that during meiosis Hhp is required for DSB formation, but we found no evidence for its having a role in DSB repair under this condition. This conclusion is consistent with the hhp mutant having high viable spore yield but low recombination-proficiency in the absence of analog (Table 2).

To directly test for a role of Hhp in DSB formation, we assayed DSBs by Southern blot hybridizations of DNA extracted from hhp1-as hhp2Δ mutants induced for meiosis in the absence of analog; DNA from wild type and rec8Δ was analyzed for comparison (Fig 1 and S1 Fig). In wild type there were six prominent, meiosis-dependent DSB hotspots on the 501 kb NotI restriction fragment J, as seen before [34,35]. DSBs were barely detectable at these sites in hhp1-as hhp2Δ, as is the case in rec8Δ and rec11Δ [14,18]. DSBs were also barely detectable in hhp1-as hhp2Δ at the strong DSB hotspot ade6-3049, as is the case in rec8Δ and rec11Δ [14,18]. In contrast, DSBs were detectable, though at reduced levels, at some hotspots on the 1500 kb NotI restriction fragment C in hhp1-as hhp2Δ and in rec8Δ and rec11Δ (Fig 1B) [14,18]. These results show that Hhp is required for most meiotic DSB formation, but residual DSBs with a spatial pattern similar to that in rec8Δ and rec11Δ, which are indistinguishable [14,18], remain in the absence of Hhp function.

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Fig 1. Meiotic DNA breakage in hhp, rec11, and rec8 mutants.

Strains with the indicated mutations were induced for meiosis in the absence of an ATP analog. At the indicated times, DNA was extracted and analyzed by Southern blot hybridization. All time points for each mutant were run on the same gel, one gel for the two NotI fragments and another for the ade6-3049 fragment (S1 Fig). Data below each lane with meiotically induced DNA are the percent of total DNA in the bands labeled a—f after subtraction of the intensity in the corresponding 0 hr lane. These data reflect DNA breakage at DSB hotspots. DSBs at the ade6-3049 hotspot are spread over about 1 kb, indicated by the bar (f) on the right. See also S1 Fig. (A) The 501 kb NotI fragment J on chromosome 1 was analyzed with a probe at its left end [60]. (B) The 1.5 Mb NotI fragment C on chromosome 2 was analyzed with a probe near its left end [18]. (C) The 6.6 kb AflII fragment containing ade6 on chromosome 3 was analyzed with a probe at its right end [61].

https://doi.org/10.1371/journal.pgen.1005225.g001

Rec8 phosphorylation-deficient mutants are not recombination-deficient

Because the residual patterns of DSB formation and recombination in hhp1-as hhp2Δ resemble those in rec8Δ (Fig 1) [14,18] and because Rec8 is an Hhp substrate [12,23], we tested the hypothesis that the Hhp substrate required for DSB formation is Rec8. One of the Hhp-dependent phosphorylation sites on Rec8 (S412) is critical for cleavage of Rec8 to allow chromosome segregation [12]. The non-phosphorylatable mutant rec8-S412A was, however, as recombination-proficient as wild type (S3 Table) and had DSB patterns on NotI fragments J and D similar to those of wild type. Similar recombination results were obtained with six additional rec8 mutants lacking seven, 12, 13, 17, or 18 phosphorylation sites (S3 Table). In S. cerevisiae, Rec8 phosphorylation is also not essential for meiotic recombination, although crossing-over is about one-half as frequent and delayed relative to wild type [36]. These results suggest that Rec8 is not, under the conditions used here, the major substrate of Hhp required for DSB formation and recombination, although minor effects cannot be ruled out.

Rec11 is phosphorylated in an Hhp-dependent manner

To search for additional Hhp substrates during meiosis, we immunoprecipitated TAP-tagged versions of Hhp1 and Hhp2 and analyzed the precipitated proteins by mass spectrometry (S2 Fig, S4 Table and S5 Table). In meiotically induced cells, but not in mitotically growing cells, we found that Hhp1 co-immunoprecipitated with Hhp2-TAP and, conversely, Hhp2 with Hhp1-TAP. We confirmed this interaction in meiotic extracts by standard co-immunoprecipitation and Western blotting (S3 Fig). Physical interaction between Hhp1 and Hhp2 was previously observed in checkpoint-activated mitotic cells [37]. Importantly, we found the cohesin subunits Rec8, Psm1, and Psm3 (orthologs of Smc1 and Smc3 in other species) in precipitates of Hhp1 and Hhp2 from meiotic cells (S4 Table). To test the possibility that other cohesin subunits interact with Hhp, we analyzed Rec11-TAP precipitates, in which we found known cohesin subunits and Hhp2, suggesting that cohesin and Hhp physically interact. Interestingly, we did not detect interaction between Hhp and cohesin in extracts from mitotically growing cells, suggesting that this interaction is stronger during meiosis or is meiosis-specific (S4 Table).

To test directly for phosphorylation of Rec11, we determined the mobility of Rec11-TAP by gel electrophoresis (Fig 2). The mobility of Rec11 from wild-type cells was considerably increased by phosphatase treatment (Fig 2A, lanes 1 and 2 vs. lanes 3 and 4), indicating that Rec11 indeed was phosphorylated during meiosis. In the presence of analog the mobility of Rec11 from the hhp1-as hhp2Δ mutant induced into meiosis was greater than that from wild-type cells (lane 5 vs. lane 1); its mobility was further increased by phosphatase treatment (lanes 7 and 8 vs. lanes 5 and 6). As expected, the mobility of Rec11 from wild type (hhp⁺) was the same with or without analog (lane 1 vs. lane 2), demonstrating that the increased mobility of Rec11 from the mutant was due to inhibition of Hhp and not to an off-target effect. In the absence of analog the mobility of Rec11 from the hhp1 mutant was also greater than that from wild type (Fig 2B, lanes 1 and 4 vs. lane 2) and was further increased by adding analog (Fig 2B, lane 2 vs. lane 3), as also evident in Fig 2A (lane 5 vs. lane 6).

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Fig 2. Rec11 is phosphorylated during meiosis, at least partially dependent on Hhp.

(A and B) pat1-114 rec11-TAP cells (hhp1⁺ hhp2⁺ or hhp1-as hhp2Δ, as indicated) were induced for meiosis in medium containing (+) or lacking (–) 30 μM 1-NM-PP1, an ATP analog. Extracted proteins were treated with alkaline phosphatase (AP) (+) or not (–) and separated by gel electrophoresis. Rec11-TAP protein was detected by western blotting using peroxidase anti-peroxidase antibody. (C) Positions of phosphorylation on Rec11-TAP isolated from meiotically induced pat1-114 cells were determined by mass spectrometry; S, serine; T, threonine. Their codons were mutated to create rec11-10A (encoding alanine at each position) and rec11-10D (encoding aspartate at each position).

https://doi.org/10.1371/journal.pgen.1005225.g002

These results indicate that Rec11 phosphorylation in the hhp1-as hhp2Δ mutant is reduced even in the absence of the inhibitor, consistent with the hhp1-as hhp2Δ mutant having a recombination-deficient phenotype in the absence of inhibitor (Table 2). They also indicate that Rec11 phosphorylation is reduced even more, but not completely, in the presence of analog. This outcome is consistent with viable spore yield being somewhat reduced in the absence of analog but reduced much more in the presence of analog (Table 2). Residual Rec11 phosphorylation in the presence of analog (Fig 2A, lane 5 vs. lane 7) may depend on residual Hhp1-as function or on another protein kinase.

Rec11 phosphorylation-site mutants are recombination- and DSB-deficient but segregation-proficient

To determine the nature of Rec11 phosphorylation during meiosis, we analyzed the immunoprecipitates of Rec11-TAP for phosphopeptides via mass spectrometry in two independent experiments. We found that Rec11 is phosphorylated on eight serine (S10, S22, S34, S43, S150, S439, S496 and S880) and two threonine (T60 and T70) residues during meiosis (Fig 2C).

To test the potential functional significance of Rec11 phosphorylation, we generated rec11 mutants encoding alanine at these ten phosphorylation sites (allele rec11-10A) or the phosphomimetic aspartate at those positions (rec11-10D) (Fig 2C). The recombination-proficiency of the rec11-10A mutant was lower than that of wild type by factors of ~5–10, whereas that of the rec11-10D mutant was near that of wild type (Tables 3). These data indicate that phosphorylation of Rec11 at one or more of these sites is important for recombination. [Since in recombination assays the phosphomimetic aspartate is less deleterious than alanine (Table 3), we presume that the deficiency in the rec11-10A mutant results from reduced phosphorylation, although other structural deficiencies of Rec11 cannot be excluded. The abundance of both Rec11 mutant proteins during meiosis was similar to that of wild-type Rec11 (S4 Fig), indicating that the mutations do not significantly alter the stability of Rec11.] Because the recombination-deficiency of the rec11-10A mutants was not as great as that of the rec11Δ null mutant or of the hhp1-as hhp2Δ mutant in the absence of analog, Rec11 presumably has additional phosphorylation sites important for its promotion of recombination; these may be the residual sites phosphorylated in hhp1-as hhp2Δ (Fig 2). Alternatively, Rec11 may have phosphorylation-independent functions important for recombination, still present in the rec11-10A mutants but lacking in rec11Δ.

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Table 3. Recombination defects in hhp1-as hhp2∆ and rec11 phosphorylation-site mutants resemble those in rec8∆ and rec11∆ mutants, and phosphomimetic rec11 mutation partially suppresses hhp1-as hhp2∆.

https://doi.org/10.1371/journal.pgen.1005225.t003

If phosphorylation of Rec11 depends on Hhp and is important for recombination, the rec11 phosphomimetic mutation, rec11-10D, might suppress the loss of Hhp function in the hhp1-as hhp2Δ mutant. Indeed, rec11-10D partially suppressed hhp1-as hhp2Δ for both gene conversion (ade6 intragenic recombination) and crossing-over (ade6—arg1 intergenic recombination). The rec11-10D mutation slightly raised the Ade⁺ recombinant frequency in the hhp1-as hhp2Δ mutant from 28 to 51 (per million viable spores; p = 0.05 by one-tailed t-test) and the ade6—arg1 crossover distance from 2.0 to 5.3 (cM; p < 0.001 by Fisher’s exact test) (Table 3). Suppression may be only partial owing to sites of phosphorylation other than those mutated in the rec11-10D mutant or to the phosphomimetic mutations being only partially effective, as is frequently observed [38,39].

As expected from the greater recombination-proficiency of rec11-10D than of rec11-10A, DSBs were more abundant in rec11-10D than in rec11-10A (Fig 1, middle two pairs of lanes in each panel). Only at one DSB hotspot, denoted “c” on NotI fragment C (Fig 1B, middle panel), were DSBs close to wild-type levels in rec11-10A; DSBs at this hotspot were also most abundant in hhp1-as hhp2Δ, rec8Δ, and rec11Δ (Fig 1) [14,18], showing that DSB formation at this hotspot is largely independent of these factors. At other hotspots, DSBs were slightly reduced in rec11-10D relative to wild type, but they were reduced more in rec11-10A though in most cases not to the level in hhp1-as hhp2Δ, rec8Δ, or rec11Δ (as noted above, DSB levels are indistinguishable in rec8Δ and rec11Δ [14,18]). These data are in accord with the recombination data in Table 3 —the rec11-10A mutation reduces both DSB formation and recombination more than the rec11-10D mutation but not as much as the hhp1-as hhp2Δ, rec8Δ, and rec11Δ mutations.

Because Hhp is required for cohesin removal and proper meiotic divisions [12,40], we tested whether Rec11 phosphorylation is required for these steps of meiosis. Although Rec8 phosphorylation is essential for its cleavage and removal and thus for segregation of chromosomes during meiosis I [12,40], meiotic divisions occurred normally in Rec11 phosphorylation-site mutants 10A and 10D, and no defect in Rec8 removal at the onset of anaphase I was observed (S5 Fig). Mutating one of the two separase cleavage sites on Rec8 (Rec8-RD1) leads to only a minor defect in chromosome segregation during meiosis [41]. We did not observe any further impairment of meiotic chromosome segregation when we analyzed cells expressing both Rec11-10A and Rec8-RD1 (S6 Fig). These results suggest that the Rec11 phosphorylation sites identified in our study are required for meiotic recombination but not for segregation of chromosomes during meiotic divisions.

Hhp promotes chromosomal loading of LinE proteins Rec10 and Rec27

Rec8 and Rec11 appear to act before the LinE proteins, because the formation of LinEs or nuclear LinE protein foci largely depends on Rec8 and Rec11, but Rec8 focus-formation does not depend on LinE proteins [22]; Rec11 focus-formation has not, to our knowledge, been similarly tested. We therefore tested the dependence on Hhp of focus-formation by LinE proteins, using fluorescence microscopy of Rec10-GFP and Rec27-GFP, both of which are functional [14,19,20]. The number of foci formed by each protein and the fraction of cells with visible foci were significantly reduced in hhp1-as hhp2Δ in the absence of analog (p < 0.01 by Fisher’s exact test) as well as in rec11-10A (p < 0.01) (Fig 3 and S7 Fig). Comparison of either time point for wt (3 and 3.5 hr) with either time point for hhp1-as hhp2∆ (4 and 4.5 hr) or for rec11-10A (3 and 3.5 hr) shows that the mutants have fewer foci and cells with foci. [We used later time points for the hhp1-as hhp2∆ mutant because meiosis is delayed in the mutant (S8 Fig).] The residual levels were similar to those reported in rec8Δ and rec11Δ mutants, which behave similarly in other recombination-related assays [19,20]. In contrast, Rec11-GFP appeared to localize to the nucleus to nearly the same extent in wild type and in hhp1-as hhp2Δ (S9 Fig) but not in rec8∆ (S10 Fig) as noted previously [42]. Rec11-GFP did not consistently form distinct foci, making quantification difficult. Nevertheless, the frequency of nuclei with Rec11-GFP fluorescence appeared to be similar in wild type and in hhp1-as hhp2Δ. These results show that the loading of LinE proteins depends on Hhp and likely on the phosphorylation of Rec11, which itself depends on Hhp (Fig 2).

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Fig 3. hhp and rec11-10A mutations reduce the number and intensity of foci of linear element proteins.

Strains with the indicated mutations were induced for meiosis in the absence of an ATP analog. At the indicated times, cells were fixed, stained with DAPI, and examined by fluorescence microscopy for foci of the indicated GFP fusion protein (green) and DNA (blue). Representative cells with Rec27-GFP are shown, and quantification is given in the table below. Data for the hhp1-as hhp2Δ mutant were taken 1 hr later than for hhp⁺ because replication is delayed by about 1 hr in the hhp1-as hhp2Δ mutant (S8 Fig). See also S6 Fig.

https://doi.org/10.1371/journal.pgen.1005225.g003

Separable functions of Hhp during meiosis

We were surprised that the hhp1-as (M84G) ATP-analog-sensitive mutant had a dramatic, differential phenotype even in the absence of added analog (Tables 2 and 3; Figs 1, 2, and 3)—it strongly reduced recombination but had much less effect on viable spore yield or chromosome segregation (Tables 2 and 3). We infer that this mutation differentially alters the substrate specificity or activity of Hhp1 such that in the absence of analog the mutant Hhp1 adequately phosphorylates Rec8 but not Rec11 (at least not completely) (Tables 2 and 3; Figs 1, 2, and 3). This fortuitous result greatly aided our experiments because hhp1Δ hhp2Δ mutants grow poorly and have very low viable spore yields [26–28], whereas the hhp1-as mutants grow like wild type and have quite high viable spore yield in the absence of analog (Table 2). There are precedents for such differential inactivation of protein kinases by ATP-binding site mutations. For example, mutation of the “gatekeeper” residue in the ATP-binding site can reduce kinase activity even without analog present [29]. In addition, differential regulation of cellular events can arise from differential threshold levels for kinase activity [44,45].

During meiosis, Hhp plays major roles both in the timely removal of cohesin from chromosome arms at the onset of anaphase I, via phosphorylation of Rec8, and in DSB formation and recombination, via phosphorylation of Rec11. Since both of these processes are meiosis-specific, it is not surprising that we found physical interaction between Hhp and cohesin only in meiotic cells (S4 Table and S3 Fig); this association may aid the coordination of Rec8 cleavage and recombination. The abundance of hhp1 and hhp2 transcripts is greatly increased during meiosis [46], a feature consistent with Hhp playing especially important roles during meiosis. Hhp directly phosphorylates Rec8 [12,23], and it may directly phosphorylate Rec11 or activate another protein kinase that does; our data do not distinguish these possibilities, but it was recently reported that Hhp1 and Hhp2 can directly phosphorylate Rec11 [43]. Hhp clearly regulates separately two events essential for the formation of viable gametes and species propagation.

A role for Hhp in the regional specificity of meiotic recombination

We found that the Hhp mutants (hhp2Δ coupled with hhp1-as or hhp1Δ) reduced recombination more in some intervals than in others (Tables 1, 2, and 3, and S3 Table), much like the region-dependent reductions by rec8 and rec11 mutations, including deletions [18,25]. Furthermore, hhp1-as abolished meiotic DSBs at some hotspots but not at others (Fig 1 and S1 Fig). The residual DSB patterns are reminiscent of those of cohesin and certain LinE mutants [14,18]. These observations lead us to propose that Rec11 phosphorylation is required for the loading of the putative Rec25-Rec27-Mug20 complex and Rec10 at DSB hotspots [14]. This proposal is consistent with the reduction, but not elimination, of Rec27-GFP foci in the hhp1-as hhp2Δ mutant (Fig 3 and S7 Fig). Rec11 phosphorylation seems not to be required for Rec11 to localize to the nucleus and possibly to load onto chromosomes, since Rec11-GFP nuclear localization was similar in wild type and in the hhp1-as hhp2Δ mutant (S9 Fig).

Our results, coupled with previous data [14], suggest the following scheme for formation of meiotic DSBs and crossovers (Fig 4). Rec8 is loaded onto chromosomes during S phase. Rec11 is concurrently or subsequently loaded in a Rec8-dependent manner (S10 Fig) [42] and then phosphorylated by Hhp, which allows the preferential loading of Rec25-Rec27-Mug20 at DSB hotspots. Rec10 is loaded at low levels independently of any of these proteins and, in their absence, activates the Rec12 complex to form DSBs at low level across the genome and at a few DSB hotspots. The Rec25-Rec27-Mug20 complex, whose loading at high levels at DSB hotspots depends on Rec8 and phosphorylated Rec11, along with Rec10 strongly activates the Rec12 complex to form DSBs at the hundreds of DSB hotspots dependent upon these proteins [14]. In this view, the main role of Hhp is to promote high level DSB formation at hotspots, which collectively account for about 70% of all DSBs and about half of all crossovers across the genome [47].

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Fig 4. Scheme for dual action of casein kinase 1 (Hhp) on cohesin subunits to regulate meiotic chromosome segregation via Rec8 phosphorylation and to promote linear element formation, meiotic DSB formation, and recombination via Rec11 phosphorylation.

Cohesin subunits Rec8 and Rec11 are loaded onto chromosomes prior to premeiotic replication, during which sister chromatid cohesion is established. Before, during, or after loading they are phosphorylated by casein kinase homologs Hhp1 and Hhp2. Right: Phosphorylation (P’n) of Rec11 leads to loading of the linear element (LinE) proteins Rec10, Rec25, Rec27, and Mug20, which in turn activate Rec12 (Spo11 homolog) and its partner proteins for DSB formation; DSBs lead to crossovers, which allow segregation of homologous centromeres, not sister centromeres, at the first meiotic division (MI). Left: Phosphorylation of Rec8 (generating P-Rec8) allows its cleavage, first in the arms, which allows segregation of homologous centromeres at MI; later, cleavage of Rec8 in the centromeres allows segregation of sister centromeres at MII. Rec8 is also needed for loading of Rec11 (S10 Fig) [42] and is thus indirectly required for DSB formation and recombination [14,18]. The precise timing of Rec8 and Rec11 phosphorylation is unknown. Rec8 and Rec11 (STAG3 functional homolog) and their phosphorylation may play similar roles in mammalian gametogenesis (see Discussion).

https://doi.org/10.1371/journal.pgen.1005225.g004

Conservation of meiosis-specific cohesin subunits and their roles

The proteins discussed here are widely conserved across all eukaryotic phyla, including humans. To our knowledge, casein kinase homologs are present, often as multiple paralogs, in all eukaryotes examined, and all except apparently the protist Tetrahymena thermophila have meiosis-specific Rec8 cohesin subunits [7]. In only rare cases, such as the budding yeast Saccharomyces cerevisiae and T. thermophila, is there no identified meiosis-specific homolog of the Rec11 cohesin subunit [6,7,48,49]. Vertebrates harbor three Rec11-like STAG (stromal antigen) proteins, STAG1, STAG2, and STAG3. STAG3 is meiosis-specific, is required for meiotic sister chromatid cohesion and chromosome axis formation, and is closely related to the Rec11 protein studied here [50–52]. Murine STAG3 is phosphorylated during meiosis, and this modification appears to be required for meiotic progression [53]. STAG3-deficient mice, both male and female, are sterile and display severe meiosis I defects [54,55]. A STAG3 frameshift mutation is apparently the cause of premature ovarian failure in humans [56]. Thus, our observations on Rec11 phosphorylation and its role in meiotic chromosome behavior are likely to pertain to meiosis and fertility in most species, including humans.

S. pombe strains, mutant constructions, and genetic methods

Strains were constructed by standard meiotic crosses [57]; genotypes of strains and the figures and tables in which each was used are in S6 Table. Mutations were introduced into cloned genes using the QuikChangeII kit (Agilent Technologies), which were inserted into the genome by transformation to antibiotic-resistance [28]. Transformants were confirmed by PCR-based analysis and, in some cases, by nucleotide sequencing.

Meiotic crosses and analysis of random spore colonies were conducted as described [57]. The ATP analog 1-NM-PP1 (Toronto Research Chemicals) was added to the sporulation agar (SPA) at 50 μM; required amino acid, purine, and pyrimidine supplements were added at 100 μg/ml. Ade⁺ recombinant frequencies were determined by differential plating on yeast extract agar (YEA) with and without guanine (200 μg/ml). To determine intergenic recombinant frequencies, spore suspensions were plated on YEA; colonies were transferred with toothpicks to YEA supplemented with adenine (100 μg/ml), incubated overnight, and replicated to appropriate media to determine phenotypes. Recombinant frequencies were converted to genetic distance (cM) using Haldane’s equation [x = -½ ln(1-2R), where x is the distance in Morgans (M) and R is the recombinant frequency].

To prepare large meiotic cultures, cells were grown to log phase at 25° C in supplemented liquid minimal medium (EMM2), washed in H₂O, resuspended in supplemented EMM2 without a nitrogen source, and incubated for 18–19 hr, at which time NH₄Cl was added to 5 mg/ml and the temperature raised to 34° C to inactivate the Pat1-114 temperature-sensitive protein kinase [58]. Cells were harvested at appropriate times after induction and analyzed for DNA content by flow cytometry (to determine the time of replication) and for other features as described below.

DNA and protein analysis

To determine DSBs, meiotic cells were washed, concentrated by centrifugation, and embedded in agarose plugs, which were sequentially treated with lytic enzymes, proteinase K, RNase, and appropriate restriction enzymes [58]. The digested DNA was separated by gel electrophoresis and analyzed by Southern blot hybridization.

To analyze phosphorylation and electrophoretic mobility of Rec11, cells expressing Rec11-TAP were arrested in G1 by nitrogen starvation, and meiosis was induced by shifting the culture to 34°C. Four hr later, cells from 20 mL of culture were concentrated by centrifugation, suspended in IPP150 buffer (50 mM Tris-HCl pH 8.0, 150 mM NaCl, 10% glycerol, 0.1% NP-40), and homogenized using glass beads (0.4–0.6 mm diameter). Extracted proteins were immunoprecipitated with IgG Sepharose 6 Fast Flow beads (GE Healthcare), treated with alkaline phosphatase (Thermo Scientific) as indicated and separated by electrophoresis through 5% polyacrylamide gels containing SDS (0.1%). Proteins were transferred to a PVDF membrane (Millipore), and Rec11-TAP and Hhp2-TAP were detected using rabbit antiperoxidase antibody linked to peroxidase (Dako; 1:30,000 dilution) in 0.1% PBS-T (8 gm NaCl, 0.2 gm KCl, 1.44 gm Na₂HPO₄, 0.24 gm KH₂PO₄, 1 mL Tween-20 per L). Hhp1-PK9 was detected using mouse-anti-PK (V5) antibody (Serotec; 1:2000 dilution) and goat anti-mouse IgG-HRP secondary antibody (Santa Cruz Biotechnology; 1:5000 dilution) in 0.1% PBS-T.

Affinity purification of proteins

Cells from six-liter (mitotic) or fifteen-liter (meiotic) cultures of strains expressing TAP-tagged proteins were collected by centrifugation; meiotic cultures were harvested at 2.5–3.5 hr after induction of meiosis. Yeast cell powder was made from frozen pellets using a SamplePrep 6870 Freezer Mill (SPEX, Inc.). Proteins were extracted using IPP150 buffer containing complete protease and phosphatase inhibitors (Roche) and 1 mM PMSF (Sigma). All washing steps were performed in Poly-Prep columns (Bio-Rad) by gravity flow. IgG Sepharose™ 6 Fast Flow beads (500 μl; GE Healthcare) were washed with IPP150 buffer, mixed with protein extract, and rotated for 2 hr at 4°C. Beads were washed with IPP150 buffer and then with TEV cleavage buffer (TCB: 10 mM Tris-HCl pH 8.0, 150 mM NaCl, 10% glycerol, 0.1% NP-40, 0.5 mM EDTA, 1 mM DTT). Protein cleavage was performed in 2 ml of TCB buffer supplemented with 400 units of AcTEV protease (Life Technologies) for 2 hr at 16°C. The eluate (2 ml) was supplemented with 6 μl of 1 M CaCl₂ and mixed with 6 ml of calmodulin binding buffer 1 (CBB1: 10 mM Tris-HCl pH 8.0, 150 mM NaCl, 10% glycerol, 0.1% NP-40, 1 mM imidazole, 1 mM Mg(OAc)₂, 2 mM CaCl₂, 10 mM β-mercaptoethanol). Calmodulin Sepharose 4B beads (150 μl; GE Healthcare) were washed with CBB1 buffer, added to a mixture of eluate and CBB1 buffer, and incubated for 2 hr at 4°C. The beads were washed with CBB1 and calmodulin binding buffer 2 (CBB2: 10 mM Tris-HCl pH 8.0, 150 mM NaCl, 1 mM Mg(OAc)₂, 2 mM CaCl₂, 1 mM β-mercaptoethanol). The proteins were step-eluted using one bed volume of elution buffer (EB: 10 mM Tris-HCl pH 8.0, 150 mM NaCl, 1 mM Mg(OAc)₂, 2 mM EGTA, 1 mM β-mercaptoethanol). Eluted proteins were separated by SDS-PAGE and silver stained. Eluates from peak fractions were analyzed by LC-MS/MS as described in S1 Text and by Cipak et al. [59].

Fluorescence microscopy

Haploid pat1-114 cells were arrested by nitrogen starvation for 16 hr and released into meiosis at 34°C by inactivation of Pat1 and addition of nitrogen. Cells were harvested at the indicated time points after meiotic induction, fixed with 70% ethanol (Fig 3 and S7 Fig) or with 99.8% methanol (S9 Fig and S10 Fig), and stained with DAPI; nuclei were counted in 100 cells at each time point. The Rec10-GFP protein was visualized in unfixed cells (S7 Fig) as follows. Cells from 100 μl of culture were collected by centrifugation, washed once with water, and spread on a cover slip coated with poly-L-lysine. Slides with a drop of mounting medium containing DAPI were covered with the cover slips, and the cells were analyzed the next day. All images were from a single focal plane; out-of-focus cells were not scored. Images for Fig 3 and S7 Fig were obtained with a Zeiss Axio Imager.Z2 microscope equipped with a Plan Apochromat 63x/1.4 oil-immersion lens and an AxioCamMRm camera; images were analysed with ZEN 2011 software. Images for S9 Fig and S10 Fig were obtained with a Zeiss Apotome microscope equipped with a 100X /1.4 oil-immersion lens and were analyzed using Axiovision software. All inductions and localization analyses were performed at least twice. Haploid strains were analysed because diploid hhp1-as hhp2∆ cells grow poorly even without analog and produce grossly abnormal asci.