MutSγ crossovers rely on residues within Zip1’s extreme N terminus

The primary amino acid sequence of Zip1 suggests that the region encompassing residues ~175–748 of the 875 residue protein has the capacity to assemble an extended coiled-coil structure, while the flanking N- and C-terminal regions are likely unstructured. Mirror-image Zip1 units assemble in a head-to-head fashion to span the ~100 nm width of the budding yeast SC central region [12, 13]; Zip1’s C termini orient toward aligned chromosome axes (lateral elements) while its N termini orient toward the central element substructure (comprised of–at least–the SUMO, Ecm11, and Gmc2 proteins) at the midline of the budding yeast SC. We previously reported that the non-null zip1[Δ21–163] mutant phenocopies SC central element-deficient ecm11 and gmc2 null mutants. In the zip1[Δ21–163], ecm11 (null) or gmc2 (null) mutant, tripartite SC assembly fails but MutSγ-mediated crossover recombination events occur in excess [24]. In order to identify residues within the Zip1 and Zip1[Δ21–163] proteins that are critical for Zip1’s crossover activity, we created and analyzed additional non-null zip1 alleles (Fig 1A). We found that alleles encoding disruptions in Zip1’s first twenty residues severely diminish Zip1’s capacity to promote MutSγ crossovers (Fig 1B, 1C and 1D).

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Fig 1. Zip1’s N terminal twenty residues are critical for MutSγ crossing over.

A) Line illustrations represent Zip1’s N terminal region (approximately residues 1–200); red brackets highlight residues that are deleted in the altered versions of Zip1 analyzed in this study (precise deletion information is indicated at the left of each line). B) Cartoons illustrate the seven genetic intervals that were utilized to assess meiotic crossover recombination in this study; strains carried two distinguishable alleles (genetic markers) corresponding to five loci whose position span the length of chromosome III, and four loci whose positions encompass more than half of chromosome VIII. Crossover recombination is measured by examining the frequency of chromatids for which adjacent genetic markers have become unlinked from one another during meiosis; a higher frequency of crossover recombination results in a larger map distance between genetic markers that define an interval. (C, D) Bar graphs plot the sum of map distances across all seven intervals in each strain, normalized to the control value. Note that the control for normalization in (C) is the pch2 value, while wild type is the control in (D). Random spore analysis was used to calculate the map distances shown in the hatched bars graphed in (C), while tetrad analysis was used to calculate the map distances illustrated by the solid bars in (C, D); see Materials and methods for map distance calculation procedures. Exact values corresponding to % of control map distances are indicated in gray color above each bar in (C, D). Map distances for individual intervals are reported in Table 1 (pch2 strains) and Table 2 (PCH2 strains). pch2 strains used in (C) are: AM3724 (ZIP1 MSH4), AM4025 (msh4), AM4023 (zip1), AM4026 (zip1 msh4), AM3725 (zip1[Δ2–163]). Strains used in (D) are: K842 (wt), K852 (msh4), AM3684 (zip1[Δ2–20]), K1000 (zip1[Δ2–20] msh4), MP43 (zip1[Δ2–9]), MP46 (zip1[Δ2–9] msh4), SYC107 (zip1[Δ10–14]), SYC149 zip1[Δ10–14] msh4, AF8 (zip1[Δ15–20]), K914 (zip1[Δ15–20] msh4), AF6 (zip1[Δ21–163]), and SYC151 (zip1[Δ21–163] msh4). Data for wild-type, msh4, zip1[Δ21–163], and zip1[Δ21–163] msh4 strains were reported previously [24]. Zip1’s first twenty amino acid residues are illustrated below the graph in (D), with colors corresponding to the residues removed in each of three zip1 mutant alleles: red = zip1[Δ2–9], blue = zip1[Δ10–14], green = zip1[Δ15–20].

https://doi.org/10.1371/journal.pgen.1008201.g001

Unlike zip1[Δ21–163] mutants but similar to a zip1 null (corresponding to a complete deletion of the ZIP1 ORF) in our BR1919 strain background, zip1[Δ2–163] meiotic cells exhibit a severely diminished capacity to form spores (1% in zip1[Δ2–163] vs 5% in zip1 vs 57% in the wild-type strain, n > 1000; S1 Table). The meiotic checkpoint that prevents zip1 null meiotic cells from producing spores relies on the AAA+ ATPase protein, Pch2 [40, 41], thus we removed PCH2 activity from zip1[Δ2–163] strains in order to evaluate the spore viability and meiotic crossover recombination phenotypes associated with this allele. As expected, the diminished spore formation phenotype of zip1[Δ2–163] is partially bypassed by removal of PCH2 (28%; S1 Table). We found that meiotic products from pch2 zip1[Δ2–163] strains were only slightly more viable than those from pch2 zip1 strains (58% in zip1[Δ2–163] vs. 39% in zip1; S1 Table), raising the possibility that the pro-crossover activity of Zip1 (and of Zip1[Δ21–163]) relies on Zip1 residues 2–20. Indeed, consistent with previously published data implicating Zip1 in the formation of MutSγ crossovers, tetrad or random spore analysis to deduce meiotic crossover recombination frequencies within seven distinct intervals (spanning most of chromosome III and a substantial length of chromosome VIII; Fig 1B) revealed that pch2 msh4, pch2 zip1, and pch2 zip1[Δ2–163] meiotic cells exhibit similarly diminished interhomolog crossing over relative to the pch2 single mutant (Fig 1C, Table 1). These data in conjunction with our prior phenotypic analysis of zip1[Δ21–163] and zip1[Δ21–163] msh4 [24] indicates a critical role for the first twenty residues of Zip1 in promoting MutSγ crossovers.

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Table 1. Genetic map distances in mutant strains; pch2 background.

https://doi.org/10.1371/journal.pgen.1008201.t001

We next assessed the sporulation and interhomolog crossover recombination phenotypes of zip1 alleles that encode proteins with smaller internal deletions of Zip1’s N terminus (Fig 1A). We found that, in contrast to zip1[Δ2–163], strains expressing zip1[Δ2–9], zip1[Δ10–14], and zip1[Δ15–20] sporulate as efficiently as zip1[Δ21–163] and wild type BR strains (57%, 54%, and 51%, respectively; S1 Table). zip1[Δ2–20] gave an intermediate sporulation efficiency at 20%. These data indicate that Zip1’s residues 2–20 and 21–163 contribute redundantly to the mechanism(s) that normally prevent a PCH2-mediated checkpoint block to meiotic progression.

Despite their near normal sporulation efficiency and high spore viability (S1 Table), zip1[Δ2–20], zip1[Δ2–9], zip1[Δ10–14], and zip1[Δ15–20] mutants exhibit deficiencies in MutSγ-mediated crossover recombination. Removal of the MutSγ complex protein, Msh4, reduces detectable crossovers in chromosome III and VIII genetic intervals to about 50% of wild-type levels (Fig 1D, Table 2). By contrast to zip1[Δ21–163] single mutants, which exhibit elevated crossovers [24], zip1[Δ2–20], zip1[Δ2–9], zip1[Δ10–14], and zip1[Δ15–20] mutants exhibit 69%, 57%, 57% and 76% of the corresponding wild-type crossover level, respectively; Fig 1D, Table 2). Importantly, removal of MSH4 from zip1[Δ2–20], zip1[Δ2–9], and zip1[Δ10–14] strains resulted in little change in the crossover phenotype relative to their zip1 single mutant allele counterparts, (zip1[Δ2–20] msh4, zip1[Δ2–9] msh4, and zip1[Δ10–14] msh4 double mutants exhibit 63%, 52%, and 54% of wild-type crossovers, respectively). Because removal of Msh4 does not cause a substantial further reduction in crossovers, we conclude that few, if any, MutSγ-mediated crossovers form in zip1[Δ2–20], zip1[Δ2–9], and zip1[Δ10–14] mutant strains. By contrast, removal of MSH4 did reduce the crossover level of the zip1[Δ15–20] mutant (76% in zip1[Δ15–20] versus 55% in zip1[Δ15–20] msh4; Fig 1C and Table 2), suggesting that the Zip1[Δ15–20] protein may support an intermediate level of MutSγ-mediated crossing over. Taken together, the interhomolog crossover recombination phenotypes of zip1[Δ2–9], zip1[Δ10–14], and zip1[Δ15–20] mutants indicate that Zip1’s first twenty residues are critical for its meiotic crossover promoting activity.

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Table 2. Genetic map distances in mutant strains; PCH2 background.

https://doi.org/10.1371/journal.pgen.1008201.t002

We note that each msh4 strain bearing one of these four non-null zip1 alleles exhibits a slightly higher crossover frequency than the msh4 single mutant. This phenomenon is particularly dramatic for the zip1[2–20] msh4 strain, which exhibits 63% of wild type crossover frequency whereas the msh4 single mutant exhibits 46% of wild-type crossovers (Fig 1D, Table 2). The curious result that crossovers are elevated in msh4 mutants when Zip1’s N terminus is altered (but not when Zip1 is absent altogether; Fig 1C), raises the possibility that Zip1’s N terminal residues might normally constrain the processing of some interhomolog recombination intermediates in a manner that ensures they are MutSγ-dependent.

Removal of residues 2–9 or 10–14 results in a novel separation-of-function Zip1 protein that is crossover-deficient but synapsis-proficient

Residues 21–163 within Zip1’s N terminal unstructured region are dispensable for Zip1’s function in crossover recombination but essential for tripartite SC assembly [24]. We investigated whether residues 2–20 are critical for the formation of mature SC by asking whether coincident linear structures of the SC transverse filament protein (Zip1) and the SC central element protein Ecm11 assemble on surface-spread meiotic prophase nuclei from strains carrying wild-type or a mutant zip1 allele and missing the Ndt80 transcription factor. Ndt80 is required for progression beyond a mid-meiotic prophase stage when full-length SCs are normally assembled [42], thus the ndt80 null background allows us to maintain cells in sporulation medium for prolonged periods in order to assess the overall capacity of a strain to assemble SC. We utilized a polyclonal antibody targeted against Zip1’s C terminal 264 residues [43] together with an antibody against the MYC epitope tag that is fused to the C terminus of one copy of the ECM11 gene in these strains.

As expected based on the SC-deficient phenotype of the zip1[Δ21–163] mutant, meiotic prophase nuclei from the zip1[Δ2–163] mutant strain fail to exhibit extensive Zip1 or Ecm11-MYC coincident linear structures on meiotic chromosomes. At 24 hours after placement into sporulation medium, when ~85% of ZIP1 ndt80 strains in our BR genetic background exhibit nearly full synapsis [44], zip1[Δ2–163] mutants instead display Zip1 or Ecm11-MYC foci of varying sizes, sometimes accompanied by a large “polycomplex” aggregate of these SC central region proteins (S1 Fig). Interestingly, ndt80 meiotic cells expressing zip1[Δ2–20] also fail to exhibit any detectable SC formation, even after 24 hours in sporulation medium (S1 Fig). These data in conjunction with the deficient SC assembly phenotype of zip1[Δ21–163] [24] indicate that residues within both the 2–20 and 21–163 regions of Zip1 are required for Zip1’s capacity to assemble SC.

However, we found that not all residues within Zip1’s 2–20 region are critical for SC assembly. In our initial examination of surface-spread meiotic prophase nuclei in zip1[Δ2–9] and zip1[Δ10–14] strains at the 24 hour time point, we observed many nuclei with extensive SC, as reflected by long linear assemblies of coincident anti-Zip1 and anti-Ecm11-MYC label (Fig 2). In addition, we observed extensive SCs in meiotic nuclei from a strain expressing zip1[10–14→A], where each of Zip1’s residues 10–14 is replaced by alanine (Fig 2).

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Fig 2. SC assembly requires Zip1 residues 15–20 but not residues 2–14.

Panels show representative surface-spread mid-meiotic prophase nuclei from wild-type (top row), and various zip1 internal deletion or residue-substitution alleles (genotypes indicated at left; strains are K320 (ZIP1), AM4064 (zip1[Δ2–9]), AM4194 (zip1[Δ10–14]), AM4069 (zip1[10–14→A]), SYC79 (zip1[Δ15–20]), and K981 (zip1[Δ15–20→A]). Note that strains carrying a short internal deletion of Zip1 residues 10–14 or 15–20 exhibit a similar phenotype, respectively, as strains in which Zip1 residues 10–14 or 15–20 are replaced with alanine. All strains carry an ndt80 null allele, which allows meiotic cultures to accumulate at mid-late prophase stages when full-length SCs are normally present. Mid-meiotic prophase chromosomes are stained with DAPI to label DNA (white), anti-Zip1 (green), and anti-MYC to label Ecm11 (magenta). The merge between Zip1 and Ecm11 channels is shown in the final column. Quantitation of the number and cumulative length of SC linear assemblies in wild type as well as each internal deletion zip1 mutant strain is given in Fig 4. Arrows point to polycomplex structures. Arrowhead indicates a large focus or pair of foci that measures at 0.7 μm and thus may have been included in the assessment of “linear” SC structures (see Fig 4). Scale bar, 1 μm.

https://doi.org/10.1371/journal.pgen.1008201.g002

The tripartite SC in budding yeast is assembled by Zip1 transverse filament proteins whose C termini orient toward homologous axes and whose N termini orient nearby to the central element substructure positioned at the midline of the SC. We used structured illumination microscopy (SIM) to ask whether the SC structures in zip1[Δ2–9] meiotic nuclei have a canonical, tripartite organization. This tripartite organization can be detected using SIM on surface-spread meiotic chromosomes dually labeled with antibodies that target the C terminal region of Zip1 and SC central element components [13]. With the increased resolution that SIM affords, our anti-Zip1 antibody localizes as a wide ribbon on linear SC structures; one can often observe parallel tracts of Zip1 C termini flanking the central element protein(s) within subsections of such a Zip1 linear element. We observed no detectable difference in the organization of Zip1 and the central element proteins within SCs assembled by wild type Zip1 versus Zip1[Δ2–9] protein: Antibodies targeting the C terminus of Zip1 were observed to flank the SC central element substructure within SCs assembled by Zip1[Δ2–9] (Fig 3).

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Fig 3. Zip1[Δ2–9] assembles with proper orientation within SCs.

Structured illumination was used to probe the organization Zip1 and Ecm11 within linear assemblies in wild type (K1268; top panels) and zip1[Δ2–9] (AM4064; bottom panels) strains. Antibodies against the C terminal 264 residues of the Zip1 protein (green) were applied in conjunction with antibodies against a mixture of Ecm11 and Gmc2 partial proteins (magenta; see Methods for antibody information). In surface-spread meiotic nuclei containing wild-type SC (top panels), C terminal Zip1 antibody label often displays a wide ribbon in which parallel tracts are sometimes visible with the resolution power of structured illumination (~120 nm) [13]. Such parallel tracts of Zip1 (see zoomed insets) flank a central narrow band of Ecm11-Gmc2 protein, and reflect Zip1’s orientation within the mature SC, where Zip1’s C termini interface with lengthwise-aligned homologous chromosome axes and Zip1’s N termini orient closer to the SC central element (comprised of Ecm11 and Gmc2). DNA is labeled by DAPI in the experiment but not shown in the figure. Bottom panels show a representative spread where wild-type Zip1 organization is apparent within the SCs built by the Zip1[Δ2–9] protein. Scale bar, 1 μm.

https://doi.org/10.1371/journal.pgen.1008201.g003

These observations indicate that, in reciprocal fashion to residues 21–163, residues 2–15 are critically required for Zip1’s MutSγ crossover-promoting activity but dispensable for its capacity to assemble tripartite SC.

In contrast to the robust synapsis observed in zip1[Δ2–9] and zip1[Δ10–14] strains at the 24 hour time point, extensive coincident linear assemblies of Zip1 and Ecm11 were not detectable in meiotic prophase nuclei from zip1[Δ15–20] or zip1[15–20→A] mutant strains (Fig 2). Instead, the vast majority of meiotic prophase nuclei from these strains exhibit foci of Zip1 and Ecm11 of varying sizes, which are sometimes, but not always, coincident.

Our phenotypic assessment of three novel non-null zip1 alleles thus indicates that Zip1’s N terminal twenty residues correspond to adjacent regions that function somewhat independently of one another: residues 2–14 are essential for normal MutSγ-dependent crossovers but dispensable for SC assembly, whereas residues 21–163; [24], and possibly residues 15–20 are less critical for MutSγ crossovers but crucial for SC assembly.

SC assembles early but may be more labile in MutSγ crossover-deficient but SC-proficient zip1 and zip3 mutants

Our initial 24 hour time point data hinted at a possible deficiency in the number of meiotic nuclei with extensive SC in zip1[Δ2–9], and zip1[Δ10–14], relative to wild-type strains. To explore the possibility that SC assembly occurs with altered timing in these zip1 mutants, we examined SC abundance in at least 50 meiotic surface-spread nuclei from zip1[Δ2–9], zip1[Δ10–14] and zip1[[Δ15–20] strains at 15, 18, 21 and 24 hours after placement into sporulation medium. Nuclei were selected solely based on DAPI-stained morphology of the DNA; this method can be used to select nuclei in mid or late meiotic prophase. In addition to a wild-type control strain, our analysis included the zip3 mutant, which resembles zip1[Δ 2–9] and zip1[Δ10–14] mutants in its failure to form MutSγ crossovers but proficiency for SC assembly (albeit diminished). We quantified the extent of SC assembly in the selected meiotic nuclei at each time point by measuring the number of linear assemblies of Zip1, Ecm11-MYC, or coincident Zip1-Ecm11-MYC (Fig 4, left column) and the cumulative length of Zip1, Ecm11-MYC, and coincident Zip1-Ecm11-MYC linear structures (Fig 4, center column).

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Fig 4. SC assembly requires Zip1 residues 15–20 but not residues 2–14.

Each circle in the scatterplots in the far left column represents the number of linear assemblies of Zip1 (blue), Ecm11 (orange) or coincident Ecm11 and Zip1 (purple) detected in surface spread meiotic nuclei of wild-type, zip1 or zip3 mutant strains (genotype indicated at the far right; strain names listed in Fig 2 legend), at 15, 18, 21 or 24 hours after placement into sporulation medium (time points indicated on the x axis). 50 nuclei were examined for each strain at every individual time point, except for wild-type strains and zip3 mutant strains at the 15 hour time point, where 125 and 100 nuclei were examined, respectively. Assemblies of SC proteins were considered to be linear if they measured 0.7 μm or greater in length, although some large or adjacent foci potentially were included in these calculations (see arrowhead in Fig 2 and Fig 2 legend). Individual circles in the middle column of scatterplots indicate the cumulative length of the linear assemblies of Zip1 (blue), Ecm11 (orange) or coincident Ecm11 and Zip1 (purple, “SC”) detected in the surface spread nuclei of indicated strains. Dark and light grey bars indicate the mean, and standard error of the mean, respectively. Bar graphs at far right indicate the percentage of nuclei in these datasets that exhibited a Zip1 polycomplex aggregate (see arrow in Fig 2 for an example). The vast majority of polycomplex structures detected contained both Zip1 and Ecm11 proteins. An asterisk indicates zero polycomplex structures detectable in any nuclei at the indicated time point. Raw data for Fig 4 plots is provided in S5 Table.

https://doi.org/10.1371/journal.pgen.1008201.g004

Our analysis revealed that SC cumulative length per meiotic nucleus (as measured by coincident Zip1 and Ecm11-MYC; purple circles in Fig 4, center scatterplots) was only slightly reduced in zip1[Δ2–9] and zip1[Δ10–14] meiocytes relative to wild type: zip1[Δ2–9] and zip1[Δ10–14] populations of meiotic nuclei exhibited a maximum of 32 and 38 microns, and on average 17 and 20 microns of cumulative SC length per nucleus, respectively, at the time point displaying the most abundant SC. ZIP1 control meiotic nuclei exhibited a maximum of 41 and average of 23 microns at the time point with the most abundant SC. zip3 mutants exhibited a more dramatic reduction in SC cumulative length relative to wild type, with a maximum of 25 and a mean of 8 microns at the time point exhibiting most abundant SC (Fig 4, center column).

Interestingly, SC cumulative length was highest for wild-type populations at the 24 hour time point, while SC cumulative length in zip1[Δ2–9], zip1[Δ10–14], and zip3 strains peaked at the 21 hour time point in our time course. Moreover, at the earliest time point (15 hour) both the number of SC linear assemblies and SC cumulative length per meiotic nucleus was substantially higher in zip1[Δ2–9], zip1[Δ10–14] and zip3 populations (exhibiting a maximum of 25, 31, 17 microns and mean of 4, 4, 2 microns, respectively, at this time point) relative to the ZIP1 ZIP3 control population, which exhibited a mean of 0.04 micron of cumulative SC length per nucleus. The difference in SC accumulation at the earliest time point in zip1[Δ2–9], zip1[Δ10–14] and zip3 relative to wild type populations of meiotic nuclei is significant (using an unpaired t-test or the non-parametric Mann Whitney test, the two-tailed P value is <0.0001 for each mutant strain) and suggests that SC initiates earlier and/or assembles faster in zip1[Δ2–9], zip1[Δ10–14], and zip3 meiocytes, relative to wild type. We repeated a similar time course analysis of SC assembly on these strains and observed consistent results (S2 Fig). Early synapsis may have been missed by two earlier analyses of SC assembly in zip3 because of technicalities: In one study, wild-type nuclei already exhibited substantial SC at the earliest time point examined [45]. An earlier time course spanned appropriate pre-synapsis time points, but only those nuclei with extensive SC were tallied, thus the less extensive SC structures potentially present in the zip3 mutant at early time points were likely excluded [28].

Using a lacO array inserted near the centromere of chromosome IV and GFP-LacI expressed in trans, we observed that SC assembles between aligned homologous chromosomes in most meiotic nuclei that display moderate to extensive synapsis in zip1[Δ2–9], zip1[Δ10–14] and zip3 strains (n > 50; Fig 5). Thus, while SC assembly initiates early and potentially in a manner that is uncoupled from normal regulatory cues in zip1[Δ2–9], zip1[Δ10–14] and zip3 strains, SC assembly in these mutants remains an event that is triggered downstream of the homologous pairing process.

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Fig 5. SC in meiotic nuclei from zip1[Δ2–9], zip1[Δ10–14], and zip3 strains assembles predominantly between homologs.

A) Representative surface-spread meiotic nuclei from wild-type (AM4458 or AM4462), zip1[Δ10–14] (AM4446), or zip3 (AM4453), strains homozygous for an ndt80 null allele, a centromere IV-associated lacO array, and GFP-LacI. Surface-spread nuclei are labeled with DAPI-labeled DNA (light blue in top images), anti-Gmc2 (magenta), and anti-GFP (green). Meiotic nuclei were examined 23 hours after placement into sporulation medium. Only those nuclei exhibiting approximately 30% minimum SC assembly were assessed for distance between GFP foci. GFP foci were considered paired if they were positioned within 0.5 micron of one another. While most meiotic nuclei examined from all strains exhibit paired GFP signals, occasionally unpaired GFP signals were observed in zip1[Δ2–9] (AM4440 or AM4441), zip1[Δ10–14] (AM4446), or zip3 (AM4453) mutants; examples are shown in the two lower right panels (zip3 strains). Scale bar, 1 μm. B) The table lists the number of paired versus unpaired GFP-LacI foci observed in at least fifty meiotic nuclei from ZIP1 ZIP3 (AM4458 or AM4462), zip1[Δ2–9] (AM4440 or AM4441), zip1[Δ10–14] (AM4446), or zip3 (AM4453) strains.

https://doi.org/10.1371/journal.pgen.1008201.g005

SC assembly appeared less extensive in zip1[Δ2–9], and zip1[Δ10–14] mutants at the 24 hour relative to the 21 hour time point (Fig 4). Although the decrease in cumulative SC at 24 hours is not dramatic, both the range and average cumulative SC length per nucleus shifted lower at the 24 hour time point compared to the 21 hour time point in zip1[Δ2–9], and zip1[Δ10–14] strains (Mann Whitney two-tailed P = 0.015 for zip1[Δ2–9] and 0.071 for zip1[Δ10–14]), while cumulative SC increased in wild-type meiotic nuclei at 24 versus 21 hour timepoints (P = 0.0014). Furthermore, Zip1 polycomplex aggregates, while undetectable in 100 zip1[Δ2–9] and zip1[Δ10–14] meiotic nuclei from 15 and 18 hour time points, were observed at the 21 hour time point and were observed even more frequently (30–40% of nuclei, n = 50) at the 24 hour time point (Fig 4, right column). As polycomplex structures tend to assemble when SC assembly is disrupted, the sub-peak level of assembled SC at the 24 and 26 hour time points and the increased occurrence of polycomplex structures together suggest the possibility that SCs assembled in zip1[Δ2–9], and perhaps zip1[Δ10–14] strains are less stable than SCs assembled in wild-type strains. The early appearance of polycomplex structures and overall lower extent of assembled SC observed in the zip3 mutant (ranging from 1.7 microns at 15 hours to 8.4 microns at the time point with peak SC assembly, 21 hours) furthermore suggests that SCs assembled in the absence of Zip3 may be unstable. A replicate time course analysis comparing 21 to 26 hour timepoints gave consistent results (S2 Fig): While the cumulative length of SC in populations of wild-type meiotic nuclei increased between 21 and 26 hour time points (Mann Whitney two-tailed P = 0.010), SC decreased between 21 and 26 hours in zip1[Δ2–9], zip1[Δ10–14] and zip3 populations of meiotic nuclei (Mann Whitney two-tailed P = 0.0015, <0.0001, and 0.0289 respectively).

Interestingly, a small number of meiotic nuclei with relatively extensive synapsis from zip1[Δ2–9], zip1[Δ10–14] and zip3 mutant strains displayed unpaired GFP-LacI-lacO signals, one or more of which associated with a linear assembly of SC protein (Fig 5). This observation is consistent with the possibility that SC structures prematurely fall apart in these mutants, and either re-assemble on or remain attached to chromosome axes after an original SC breaks down.

Our time course experiment furthermore confirmed that zip1[Δ15–20] meiotic nuclei fail to assemble extensive SC at any point during meiotic prophase (prior to the ndt80 late meiotic prophase arrest). Out of the 200 zip1[Δ15–20] meiotic nuclei analyzed over the time course, zero exhibited robust long linear structures containing coincident Zip1 and Ecm11-MYC. However, large or adjacent foci of coincident Zip1 and Ecm11 were sometimes included in our SC measurements (which recorded any Zip1 or Ecm11 continuous structures with a dimension of 0.7 micron or more), and occasionally a meiotic nucleus displayed linear elements of Ecm11 and Zip1 with a frayed and diffuse appearance (Fig 6). The average cumulative length of SC per nucleus detected in zip1[Δ15–20] populations was 1.7 microns at the peak time point (24 hours; Fig 4). Zip1 polycomplex structures were observed only at the 24 hour time point in zip1[Δ15–20] strains (Fig 4, right column).

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Fig 6. Zip1[Δ15–20] protein occasionally assembles fragile-appearing structures built of coincident Zip1 and Ecm11.

While the vast majority of meiotic nuclei from zip1[Δ15–20] strains (SYC97) exhibit only abundant and varying-sized foci of Zip1 and Ecm11 (Figs 2 and 4), frail-looking linear assemblies of Zip1 (green) were occasionally observed accompanying similar types of Ecm11 linear assemblies (magenta) on surface-spread meiotic chromosomes (labeled with DAPI, white). The abnormal-looking linear assemblies appear wavy, and often taper at their ends. These assemblies (as well as instances of adjacent large focal deposits of Zip1 and Ecm11) were included in the linear assembly measurements reported in Fig 4. The top row presents the singular rare nucleus, out of the more than 100 nuclei examined, in which these frail linear assemblies were most abundant. We note that unlike the robust linear assemblies of coincident Ecm11 and Zip1 observed in zip1[Δ2–9] or zip1[Δ10–14] strains, these diffuse linear assemblies do not appear to join lengthwise-aligned chromosomes. Scale bar, 1 μm.

https://doi.org/10.1371/journal.pgen.1008201.g006

Crossover-deficient, synapsis-proficient zip1 mutants initiate SC assembly from both centromeric and interstitial chromosomal sites

While the earliest SC assembly events that occur during meiosis in budding yeast have been found to preferentially initiate at centromeres [29], SC assembly events are also associated with non-centromeric sites (presumably recombination sites) in wild-type meiotic nuclei at intermediate stages of synapsis. In the zip3 mutant, by contrast, new SC assembly events associate predominantly with centromeres at both early and later meiotic prophase stages [29]. As the SC assembly and meiotic crossover phenotypes in zip1[Δ2–9] and zip1[Δ10–14] strains resemble the phenotypes found in zip3 mutant meiotic cells, we asked whether nascent SC assembly events in zip1[Δ2–9] and zip1[Δ10–14] meiotic nuclei associate with centromeres more often than wild-type nuclei at intermediate stages of synapsis. Surface-spread meiotic chromosomes from 15, and 18 hour time points were co-labeled with antibodies that target Zip1, and that target the MYC epitope that is fused to the Ctf19 centromere protein in these strains. In order to enrich for new, singular SC assembly events in our analysis, we identified the total number of Zip1 linear stretches measuring between 0.7–1.0 micron in length, and measured the number that are directly adjacent to (overlapping) a Ctf19-MYC focus. We found that 83% (29 out of 35) of such short SC structures were associated with a Ctf19-MYC focus in zip3 meiotic nuclei, consistent with previously-published data [29]. By contrast, 48%, 52% and 64% (35/73, 27/52, and 23/36) of short Zip1 linear structures were associated with Ctf19-MYC in zip1[Δ2–9], zip1[Δ10–14], or ZIP1 ZIP3 meiocytes, respectively (Fig 7). Thus, in contrast to zip3 mutants, zip1[Δ2–9], and zip1[Δ10–14] mutants display a wild-type capacity to initiate SC assembly from non-centromeric sites on meiotic chromosomes.

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Fig 7. SC comprised of Zip1[Δ2–9] or Zip1[Δ10–14] protein initiates at both centromeric and non-centromeric chromosomal sites.

A) Representative surface-spread meiotic nuclei from wild-type (AM4203; top row), zip3 (AM4204; second row), zip1[Δ2–9] (AM4231; third row) or zip1[Δ10–14] (AM4175; bottom row) strains producing the centromere-associated Ctf19-MYC protein. Surface-spread nuclei are labeled with DAPI-labeled DNA (white), anti-Zip1 (magenta), and anti-MYC (green). While all strains at this early stage of synapsis exhibit a large number of short Zip1 assemblies associated with a centromere, fewer examples of ≤1 μm Zip1 assemblies without an associated centromere were observed in zip3 strains, relative to wild-type, zip1[Δ2–9] or zip1[Δ10–14] strains. Scale bar, 1 μm. Quantitation of the number of ≤1 μm Zip1 assemblies with or without an associated Ctf19-MYC signal is shown in the bar graphs in (B), where light shading represents the percentage of total ≤1 μm Zip1 assemblies in each strain that are unassociated with a centromere signal. In wild-type strains, 23 out of 36 ≤1 μm Zip1 assemblies (in 10 surface-spread nuclei) were associated with a centromere; In zip3 strains, 29 out of 35 ≤1 μm Zip1 assemblies (in 11 nuclei) were associated with a centromere; In zip1[Δ2–9] strains, 35 out of 73 ≤1 μm Zip1 assemblies (in 21 nuclei) were associated with a centromere; In zip1[Δ10–14] strains, 27 out of 52 ≤1 μm Zip1 assemblies (in 11 nuclei) were associated with a centromere. A Fishers Exact Test found no significant difference between the proportion of centromere-associated Zip1 stretches in wild-type versus zip3 in this data set (two-tailed P value = 0.107), but did find a significant difference between zip1[Δ2–9], and zip3 (two-tailed P value = 0.0007), and between zip1[Δ10–14] and zip3 (two-tailed P value = 0.0033).

https://doi.org/10.1371/journal.pgen.1008201.g007

Adjacent regions within Zip1’s first twenty residues have opposing effects on the SUMOylation of an SC central element protein

Humphryes et. al (2013) demonstrated that SUMOylated Ecm11 is required for SC assembly, and that the central element component Gmc2, transverse filament Zip1, along with SIC proteins Zip2, Spo16 and Zip4 (but not Zip3) are required for robust Ecm11 SUMOylation during meiosis. This report also revealed that Ecm11 is hyper-SUMOylated in mutants missing the putative SUMO E3 ligase and SIC protein, Zip3. To ask whether the N terminal twenty residues of Zip1 are required for its capacity to regulate Ecm11 SUMOylation, we evaluated the abundance of Ecm11 forms in meiotic extracts from strains homozygous for ZIP1, zip1[Δ2–9], zip1[Δ10–14], zip1[Δ15–20], a zip1 null, or a zip3 null allele. These strains also are homozygous for the ndt80 mutation, in order to allow our asynchronous meiotic cultures to accumulate (by 24 hours after placement into sporulation medium) at a mid-late meiotic prophase stage, when the Ecm11 SUMOylation that accompanies synapsis is at a maximum [42].

A Western blot can readily detect three forms of Ecm11-MYC in protein extracts from meiotic cells homozygous for MYC-tagged Ecm11 [13, 35, 38]. UnSUMOylated Ecm11-MYC migrates near the 75 kD marker on a protein gel, whereas monoSUMOylated and polySUMOylated Ecm11-MYC is positioned near the 100 kD and 150 kD positions, respectively. HyperSUMOylated Ecm11-MYC, which is abundant in zip3 meiotic extracts, migrates at various positions between the 150 kD and 250 kD markers (Fig 8A).

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Fig 8. Zip1 residues 15–20 promote, while residues 2–14 limit, SUMOylation of the SC central element protein Ecm11.

A) A representative Western blot using an anti-MYC antibody reveals unSUMOylated, mono-SUMOylated, poly-SUMOylated and hyper-SUMOylated forms of Ecm11-MYC in meiotic extracts prepared from ZIP1 ZIP3, zip3, or various zip1 mutant strains (protein alterations caused by each zip1 allele is indicated on the x axis). All strains carry an ndt80 null allele, which causes a meiotic arrest that ensures maximal enrichment of mid-meiotic prophase stage cells at 24 hours after placement into sporulation medium [44]. Meiotic extracts were prepared at 24 hours after placement into sporulation media, as previously described [13, 38]. Strains included in this analysis are: AM2712 (ZIP1), MP39 (zip1[Δ2–9]), SYC96 (zip1[Δ10–14]), SYC97 (zip1[Δ15–20]), AM3719 (zip3), AM3662 (zip1[F4A,F5A]), AM3628 (zip1[N3A,R6A,D7A]), AM3656 (zip1[P14A,P16A]), AM2784 (zip1Δ), K986 (zip1[I18A,F19A]). B) The stacked bar graph plots the percentage of mono-SUMOylated (dark shaded bar), poly-SUMOylated (light bar), or hyper-SUMOylated (gray shaded bar) forms of Ecm11-MYC detected in each strain at 24 hours after placement into sporulation media. The absence of a light or gray bar in some strains indicates that this form of Ecm11 was detected in less than 1% of the total population. Error bars represent the range of values from three independent meiotic cultures, except in the case of the zip1[F4A, F5A] strain, where only one experiment was performed. Data plotted is listed in S6 Table. * Note zip1[I18A, F19A] is not present in the blot shown in (A).

https://doi.org/10.1371/journal.pgen.1008201.g008

We found the proportion of SUMOylated Ecm11-MYC in ZIP1 ZIP3 ndt80 meiotic extracts at the 24 hour time point to be, on average, 14%, wherein 11% of total Ecm11-MYC was of the monoSUMOylated form and 3% was of the polySUMOylated form (three replicates; Fig 8B). Consistent with prior results [13, 38], zip1 null strains exhibited a relatively low level of SUMOylated Ecm11: An average of 4% of total Ecm11-MYC was of the monoSUMOylated form, while polySUMOylated Ecm11-MYC was below levels of detection (less than 1%; Fig 8B). Again consistent with prior findings [38], zip3 meiotic extracts exhibited not only an elevated level of polySUMOylated Ecm11-MYC (15% of total Ecm11-MYC, on average, over three replicates), but also an abundance of hyperSUMOylated Ecm11 (19% of total Ecm11-MYC, on average, over three replicates; Fig 8B).

We found that residues 15–20 are important for Zip1’s capacity to promote Ecm11 SUMOylation. In zip1[Δ15–20] meiotic extracts at the 24 hour time point, on average only 8% of total Ecm11 was of a SUMOylated form (Fig 8B). Given the SC assembly defect of zip1[Δ15–20] strains, this result bolsters a direct correlation between Ecm11 SUMOylation and SC assembly.

In striking contrast, cells expressing the zip1[Δ2–9] or zip1[Δ10–14] alleles exhibit robust levels of Ecm11 SUMOylation as well as the hyperSUMOylated forms of Ecm11 that are characteristic of the zip3 mutant. An average of 40% of Ecm11-MYC was SUMOylated in zip1[Δ2–9] meiotic extracts at the 24 hour time point, with 9% in the hyperSUMOlated form (Fig 8B). Likewise, an average of 54% of Ecm11-MYC was SUMOylated in zip1[Δ10–14] meiotic extracts, with 13% in the hyperSUMOylated form (Fig 8B).

It has been proposed that the hyperSUMOylated forms of Ecm11 that occur in zip3 mutant meiotic cells correspond to Ecm11 linked to poly-SUMO branched chain structures of various sizes and shapes [38, 46]. The accumulation of such extensively SUMOylated Ecm11 protein in zip1[Δ2–9] and zip1[Δ10–14] mutants indicates that residues within the 2–14 region of Zip1 are dispensable for Ecm11 SUMOylation per se, but regulate the extent and/or manner of Ecm11 SUMOylation. Zip1’s residues 2–14 appear to control similar aspects of Ecm11 SUMOylation as the putative SUMO E3 ligase, Zip3, raising the possibility that Zip1’s N terminal residues regulate Ecm11 SUMOylation in part through an interaction with Zip3.

The phenotype of two- or three- residue alterations within Zip1’s N terminus resemble corresponding small internal deletion alleles

We found that zip1 alleles encoding proteins with alanine substitutions in place of dual or triple residues within Zip1’s N terminus exhibit the distinguishing Ecm11 SUMOylation and synapsis phenotypes of corresponding internal deletion zip1 alleles. zip1[N3A, R6A, D7A], zip1[F4A, F5A], and zip1[P14A, P16A] exhibited elevated levels of SUMOylated Ecm11 during meiosis, reminiscent of the zip1[Δ2–9] and zip1[Δ10–14] mutants, although we note that zip1[N3A, R6A, D7A] and zip1[P14A, P16A] strains exhibit a particular abundance of monoSUMOylated relative to polySUMOylated and hyperSUMOylated Ecm11, which differs slightly from the distribution of SUMOylated Ecm11 forms in zip1[F4A, F5A], zip1[Δ2–9], zip1[Δ 10–14] or zip3 mutants (Fig 8B). By contrast, meiotic extracts from zip1[I18A, F19A] strains exhibit a dramatic reduction in SUMOylated Ecm11, reminiscent of meiotic extracts from zip1[Δ15–20] and zip1 null strains (Fig 8B).

We also found that zip1[N3A, R6A, D7A] and zip1[I18A, F19A] exhibit SC assembly phenotypes that are generally reminiscent of corresponding internal deletion zip1 alleles. Linear stretches of coincident Zip1 and Ecm11-MYC were often detectable on surface-spread meiotic chromosomes from zip1[N3A, R6A, D7A] ndt80 and zip1[F4A, F5A] ndt80 strains at multiple time points in a meiotic time course, while such extensive SC structures were absent from meiotic chromosomes in zip1[I18A, F19A] ndt80 strains at all time points (Fig 9).

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Fig 9. zip1[N3A, R6A, D7A] and zip1[F4A, F5A] and zip1[I18A, F19A] mutants resemble corresponding small deletion alleles with respect to synapsis.

Panels show representative surface-spread mid-meiotic prophase nuclei from zip1[N3A, R6A, D7A] (K969; top panel including 3 rows), zip1[F4A, F5A] (AM4067; middle panel including two rows) and zip1[I18A, F19A] (K985; bottom panel including three rows) mutants, with genotypes indicated at left. All strains carry an ndt80 null allele, which allows meiotic cultures to accumulate at mid-late prophase stages when full-length SCs are normally present. Mid-meiotic prophase chromosomes are stained with DAPI to label DNA (white), anti-Zip1 (green), and anti-MYC to label the epitope-tagged Ecm11 expressed in these strains (magenta). The merge between Zip1 and Ecm11-MYC channels is shown in the final column.

https://doi.org/10.1371/journal.pgen.1008201.g009

We furthermore found that meiotic crossovers are reduced in zip1[N3A, R6A, D7A], zip1[F4A, F5A], and zip1[I18A, F19A] point mutants in a manner that resembles the corresponding deletion strain (S2 Table). Specifically, zip1[F4A, F5A] exhibited the most dramatic deficit in meiotic crossovers, and crossovers in this mutant are not further reduced by removal of MSH4, indicating that zip1[F4A, F5A] meiotic cells lack MutSγ-mediated crossovers (S2 Table). Crossovers did further diminish from their intermediate level when MSH4 was removed from zip1[I18A, F19A] mutants (S2 Table).

The phenotypes of these novel zip1 dual- and triple-residue substitution alleles strengthen the idea that Zip1’s first twenty residues encompass both crossover recombination and SC assembly functionalities and that adjacent sites within this region maintain different and independent roles in regulating synapsis.

Residues within the 2–14 region influence Zip1’s capacity to interface with Zip3 at polycomplex structures

The shared phenotypes of zip1[Δ2–9], zip1[Δ10–14] and zip3 mutants prompted us to wonder whether the N terminus of Zip1 directly or indirectly interacts with the Zip3 protein. Prior evidence for an interaction between Zip1 and Zip3 includes the observation that Zip3 is detected throughout Zip1 polycomplex structures that assemble in contexts where SC assembly fails [28, 34]. To explore the possibility that Zip1’s N terminus mediates an interaction with the Zip3 protein, we examined the distribution of Zip3 at Zip1 polycomplex structures assembled in spo11 meiotic cells, which fail to initiate recombination and thus also SC assembly [45, 47].

Of the polycomplex structures assembled by wild-type Zip1 and Zip1[Δ15–20] protein, 100% (20/20) exhibited Zip3-MYC distributed uniformly across the entire structure (Fig 10A). Frequently, additional “capping” structures of coincident Zip3-MYC and Zip4-HA protein flank the Zip1 polycomplex, as has been reported previously [34]. Intriguingly, however, among more than 20 meiotic nuclei examined from spo11 zip1[Δ2–9] and spo11 zip1[Δ10–14] strains, Zip3-MYC was completely absent from the bulk of the Zip1 polycomplex structure (Fig 10A). Instead, Zip3-MYC co-localized with Zip4-HA in the capping configuration at opposite ends of the polycomplex. Often these “capping” structures of coincident Zip3-MYC and Zip4-HA were observed at a substantial distance away from the polycomplex aggregate of Zip1 protein.

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Fig 10. Zip1 residues 2–14 promote the localization of Zip3 to Zip1 polycomplex structures.

A) Four groups of panels each show three representative images of Zip1 polycomplex structures in spo11 meiotic prophase nuclei expressing either wild-type ZIP1 (AM4174; far left group), zip1[Δ2–9] (AM4253; second group), zip1[Δ10–14] (AM4173; third group), or zip1[Δ15–20] (AM4256; far right group). Zip1 polycomplex is shown in green (first row) on surface-spread meiotic prophase nuclei. The localization of Zip3-MYC (second row, magenta) and Zip4-HA (fourth row, magenta) is also assessed. Merged images between either Zip3-MYC and Zip1 or Zip4-HA and Zip1 are shown in the third and fifth rows, respectively. The number of polycomplexes (n = 20) for which Zip3 is nearly fully coincident with Zip1 is displayed at the bottom of the corresponding strain’s images (n = 20). (B) Three groups of panels each show three different surface spread spo11 meiotic nuclei with a Zip1 polycomplex structure (labeled by anti-Zip1; green, top row). The localization of Zip3-MYC (second row, magenta) and Zip4-HA (fourth row, magenta) is also assessed on these surface-spread meiotic prophase nuclei. Merged images between either Zip3-MYC and Zip1 or Zip4-HA and Zip1 are shown in the third and fifth rows, respectively. Nuclei at the far left correspond to strains homozygous for zip1[N3A, R6A, D7A] (AM4350); the middle panel corresponds to strains homozygous for zip1[F4A, F5A] AM4340; and nuclei in the far right panel correspond to strains homozygous for zip1[Δ21–163] (AM4343). Scale bar, 1 μm.

https://doi.org/10.1371/journal.pgen.1008201.g010

We furthermore found that Zip3 is diminished at polycomplexes assembled by Zip1[N3A, R6A, D7A] or Zip1[F4A, F5A] protein (Fig 10B), although the absence of Zip3 from the bulk of these Zip1 polycomplex structures is less dramatic than what is observed at Zip1[Δ2–9] or Zip1[Δ10–14] polycomplex. Finally, as expected based on the robust localization of Zip3 to Zip1[Δ15–20] polycomplex, we found that Zip3 also localizes uniformly throughout polycomplexes built of Zip1[Δ21–163] protein (Fig 10B).

These data indicate that, at least in the context of polycomplex structure, Zip1’s residues 2–14 mediate a direct or indirect interaction with the pro-crossover and putative E3 SUMO ligase protein, Zip3.

Residues within Zip1’s 2–14 region are critical for Zip3 recruitment to recombination initiation sites

Zip3 and other SIC proteins (such as Zip2, Zip4 and Spo16) form foci that co-localize with MutSγ along aligned homologous chromosomes at mid meiotic prophase [28, 34]. Consistent with the notion that such Zip3 foci mark recombination intermediates, Zip3 has been detected at DSB hotspots using chromatin immunoprecipitation (ChIP) [37, 48]. Zip1 was found to be required for the recruitment of Zip3 to the DSB sites examined, thus we asked whether the capacity of Zip1 to recruit Zip3 to DSB sites relies on the N terminal residues of Zip1 that facilitate Zip3’s localization to Zip1 polycomplex.

We performed ChIP in conjunction with quantitative PCR (ChIP-qPCR) on meiotic cell extracts from ZIP1, zip1[Δ2–9], zip1[Δ10–14], and zip1 null strains expressing a Zip3 protein with three copies of the FLAG epitope fused to its C terminus. Strains for this experiment were built in the SK1 genetic background, to ensure maximal synchrony over a meiotic time course (SK1 strains enter and/or progress through meiosis more synchronously than the BR strain background used for all other experiments in this study). ChIP-qPCR was performed at multiple time points during sporulation, and the time course experiment was performed in duplicate for each strain except for the zip1 null negative control, where the single experiment performed gave results that are consistent with prior published data [48].

We examined Zip3-6xHIS-3xFLAG association with chromosomal sites corresponding to three known DSB hotspots, a centromere, or the chromosome axis [37, 48]. Sequences enriched for Rec8 that are embedded in the proteinaceous chromosome axis are generally anti-correlated with DSB sites, but are thought to associate with DSB repair intermediates, according to a “loop-tether” model for DSB formation in budding yeast [49]. A sequence internal to the large NFT1 open reading frame was previously found to be devoid of Zip3 binding [50, 51] and thus served as a negative control for Zip3 enrichment.

In strains carrying wild-type ZIP1 within two hours after placement of into sporulation medium, centromeric DNA was more abundant in Zip3 immunoprecipitates relative to axis or DSB site DNA, consistent with previously published results [37, 48]. Between two and four hours after placement into sporulation medium, DNA sequences corresponding to chromosome axis sites and three DSB hotspots (GAT1, BUD23, and ERG1; Fig 11) significantly increased their abundance within Zip3 immunoprecipitates, reflecting Zip3 recruitment to these chromosomal sites. Zip3 localization to all sites peaked at the 4 hour time point in ZIP1 meiotic cells, which corresponds to maximal DSB activity at the BUD23 locus in this SK1 strain background [48]. At this four hour time point, Zip3 enrichment was found to be two to three fold greater at GAT1 and BUD23 compared to DNA sequences at the chromosome axis (Fig 11). Consistent with DSB repair timing in this genetic background, Zip3 enrichment at all sites dramatically diminished between four and six hours, and was at pre-meiotic levels by eight hours after placement in sporulation medium. Consistent with prior findings, Zip3 was virtually undetectable at DSB, axis and centromere sites in the zip1 null strain (in which the ZIP1 ORF is deleted; Fig 11; [48]).

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Fig 11. Residues 2–14 are required for Zip1’s capacity to recruit Zip3 to recombination initiation sites.

Chromatin immunoprecipitation (ChIP) followed by quantitative PCR (qPCR) to monitor the association of Zip3-FLAG with three DSB sites (GAT1, BUD23, ERG1, dark blue, green and light blue lines, respectively) as well as to centromere and axis sites (magenta and red lines, respectively) in SK1 strains carrying ZIP1 (ORD9670), zip1[Δ2–9] (AM3946/VBD1872), zip1[Δ10–14] (AM3951/VBD1873), or zip1Δ (ORD9689) alleles. x axes indicate number of hours after placement in sporulation medium. The relative abundance of indicated chromosomal sites detected by qPCR in Zip3-FLAG immunoprecipitates is expressed as a percentage of the abundance of each site detected in the input, prior to immunoprecipitation. Values are the average ± standard deviation from two independent experiments. A single experiment was performed in the case of the zip1 null strain. Data plotted is listed in S6 Table.

https://doi.org/10.1371/journal.pgen.1008201.g011

Similar to a zip1 null strain, little Zip3 was detectable at any of the three DSB sites examined, nor at the axis or centromere site, in zip1[Δ2–9] or zip1[Δ10–14] strains (Fig 11). The phenotype of zip1[Δ2–9] in this experiment appeared indistinguishable from the zip1 null, whereas slight Zip3 enrichment was detected at the BUD23 DSB site in the zip1[Δ10–14] meiotic time course. These data indicate that the capacity of Zip1 to recruit Zip3 to DSB sites during meiosis relies on Zip1’s N terminal residues 2–14.

As expected based on ultrastructural images of recombination nodules along the length of synapsed chromosomes [20, 22, 52], we found that meiotic recombination proteins embed within the SC central region in budding yeast. Structured illumination microscopy (SIM) in conjunction with antibodies targeting the meiotic axis protein Red1 and the central element protein(s) Ecm11 or Gmc2 reveal MutSγ and Zip3 foci at the midline of the SC, embedded within the SC central element. Singular foci of epitope-tagged MutSγ protein Msh4-MYC and Zip3-MYC localize directly between aligned Red1-labeled axes, where the SC central element substructure is positioned (Figs 12 and 13). When antibodies targeting SC central element proteins are used to label the SC central element directly, Msh4 foci are observed embedded directly in the linear Ecm11-Gmc2 structures at the midline of the SC (Fig 13).

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Fig 12. Zip3-MYC foci localize to the SC central element sub-structure in wild-type meiotic nuclei and are diminished in number and intensity in zip1[Δ2–9] and zip1[Δ10–14] strains.

Images show representative surface-spread meiotic nuclei labeled with antibodies against the chromosomal axis protein Red1 (magenta) and antibodies against the MYC epitope to label Zip3-MYC (green). (A) In wild-type strains (AM4171), structured illumination microscopy reveals Zip3-MYC foci directly in between lengthwise-aligned homologous chromosome axes in mid-meiotic prophase nuclei, reflecting the embedded distribution of Zip3 complexes within the central region of SCs. (B) The position of Zip3-MYC foci in between lengthwise-aligned axes is unchanged but the number of robust Zip3-MYC foci is severely diminished in zip1[Δ2–9] strains (AM4277). Zip3-MYC’s location within the central element substructure of the SC in this strain is also indicated in this image, where Zip3-MYC (green) is co-labeled with antibodies that target Ecm11-Gmc2 (magenta). Scale bar, 1 μm.

https://doi.org/10.1371/journal.pgen.1008201.g012

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Fig 13. Msh4-MYC localizes to the SC central element sub-structure.

Images show representative surface-spread meiotic nuclei from a wild-type strain (K1268) carrying a MYC-tagged MSH4 gene. The surface-spread nuclei are labeled with antibodies against Ecm11-Gmc2 central element proteins (magenta, top row), or the chromosomal axis protein Red1 (magenta, bottom row), in conjunction with antibodies against the MYC epitope (green). The increased resolving power of structured illumination microscopy reveals that Msh4-MYC foci directly embed within the central element substructure of the SC (see zoomed inset in top row). Scale bar, 1 μm.

https://doi.org/10.1371/journal.pgen.1008201.g013

In SCs assembled by Zip1[Δ2–9] protein, we observe Zip3-MYC foci embedded within the SC central element, but such Zip3-MYC foci appear strongly diminished in both number and intensity relative to Zip3-MYC foci on meiotic chromosomes from ZIP1 strains (Fig 12). Similarly, using conventional wide-field fluorescence microscopy we observe a diminished number of bright Msh4-MYC foci on aligned mid-meiotic prophase chromosomes in strains expressing zip1[Δ2–9], zip1[Δ10–14] and zip1[Δ15–20] relative to ZIP1 strains, mimicking the Msh4-MYC pattern seen in a zip1 null strain (Fig 14A and 14B). Furthermore, the Msh4-MYC foci observed on synapsed meiotic prophase chromosomes in zip1[Δ2–9] strains do not robustly co-localize with other SIC proteins, such as Zip4-HA, relative to the Msh4-MYC foci assembled on synapsed chromosomes in wild-type meiotic nuclei (S3 Fig).

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Fig 14. Msh4-MYC foci are reduced in number and diminished in size in zip1[Δ2–9] and zip1[Δ10–14] strains.

Images show representative surface-spread meiotic nuclei labeled with antibodies that target Ecm11-Gmc2 (magenta, top rows) or antibodies against the chromosomal axis protein Red1 (magenta, bottom rows) in conjunction with antibodies that target the MYC epitope to label Msh4-MYC (green). (A) In wild-type strains (AM4278; top row), conventional fluorescence microscopy reveals Msh4-MYC foci directly embedded within the SC central element substructure, consistent with the position of Zip3 complexes (which colocalize with Msh4 [28]) within the central region of SCs as shown in Fig 12. The position of Msh4-MYC foci within the central region of the SC is unchanged but the number of robust Msh4-MYC foci is severely diminished in zip1[Δ2–9] strains (AM4274; middle row) and in zip1[Δ15–20] strains (AM4265; bottom panel including two rows). (B) The distribution of Msh4-MYC foci with respect to Red1-labeled chromosome axes is shown for wild type (K1268; top row), zip1[Δ10–14] strains (AM4270; middle row), as well as zip1 null strains (AM4263; bottom row). Scale bar, 1 μm.

https://doi.org/10.1371/journal.pgen.1008201.g014

Our ChIP and cytological studies together indicate that the N terminal twenty residues of the budding yeast transverse filament protein, Zip1, are essential for the proper enrichment of pro-crossover protein Zip3 to meiotic centromeres, to DSBs, and to chromosomal axis sites, and for the accumulation of robust Zip3 and MutSγ foci within the central region of the SC.

Adjacent functionally distinct regions within Zip1’s N terminal twenty residues: Interaction domains?

Our analysis of the novel zip1 alleles reported here leads us to propose that Zip1’s first twenty residues correspond to adjacent interaction domains that directly engage with pro-crossover and pro-synapsis machinery and/or mechanisms, as illustrated in Fig 15B. The absence of Zip1’s residues 2–9 or 10–14 confers a phenotype that strongly resembles the unique phenotype of cells missing the pro-crossover SIC protein, Zip3: zip1[Δ2–9], zip1[Δ10–14] and zip3 strains display some SC assembly despite severely diminished MutSγ crossovers, and hyperSUMOylated forms of the SC central element protein, Ecm11. This constellation of phenotypes is a striking contrast to the synapsis-deficiency and diminished Ecm11 SUMOylation phenotype displayed by mutants that are missing Zip1 altogether or missing other SIC proteins that colocalize with Zip3 at presumed recombination intermediates embedded in the SC, such as Zip2, Zip4, Spo16 or the MutSγ complex [32–34, 37, 38, 55]. The unique, zip3-like phenotype of zip1[Δ2–9] and zip1[Δ10–14] mutants suggests that these mutant meiocytes are missing a specialized capacity to promote Zip3 function.

Zip3 localizes, along with other SIC proteins, to MutSγ foci on mid-meiotic prophase chromosomes in budding yeast and is (along with the other SIC proteins) required for MutSγ -mediated crossover formation [28, 35]. Unlike Zip2, Zip4 and Spo16, however, Zip3 has been implicated in preventing unwarranted (recombination-independent) SC formation [45] and is not required per se for SC assembly [28]. Zip4 directly interacts with a Zip2-Spo16 complex that is capable of binding branched DNA structures, and can interact with the meiotic axis component Red1, the MutSγ factor Msh5, as well as Zip3 [37]. One possibility is that while Zip2, Zip4 and Spo16 are absolutely critical at recombination sites to establish a suitable foundation on which to initiate SC assembly, Zip3 may be specifically involved in the licensing of SC assembly at certain recombination sites.

Interestingly, we found evidence in support of a specific regulatory role for Zip3 in meiotic recombination as well. Our data indicates that crossover levels in zip3, zip3 msh4, or zip1[2–20] msh4 mutants are elevated over the msh4 single mutant (Table 2, S2 Table) while crossovers in zip1 meiotic cells resemble the level observed in msh4 and zip1 msh4 strains (Table 1). This observation suggests that Zip3 acts not only to promote MutSγ crossing over, but also to ensure that Zip1-associated recombination intermediates are processed in an MutSγ-dependent manner. Under this model, recombination intermediates associated with both Zip3 and Zip1 fail to resolve into interhomolog crossover events when Msh4 is absent, but when Zip3 or both Msh4 and Zip3 are absent, at least some Zip1-associated recombination intermediates can resolve into interhomolog crossovers (through a MutSγ-independent pathway). This model is supported by sequence signatures observed at interhomolog recombination events in msh4 versus zip3 mutants [31], which suggest that unbiased resolution of joint molecule recombination intermediates—a type of resolution associated with MutSγ-independent pathways [5, 56]—occurs to a greater extent in zip3 relative to msh4 mutants.

Zip3 is a RING domain protein that has been shown to have SUMO ligase activity in vitro [46], and to be responsible for the SUMOylation of a fraction of Red1, a meiosis-specific axis associated protein that is required for normal recombination and SC assembly [57–59]. On the other hand, Zip3 normally prevents hyperSUMOylation of the SC central element protein, Ecm11. Thus Zip3 likely impacts meiotic recombination and SC assembly mechanisms through both negatively and positively regulating the SUMOylation of several distinct target proteins.

Consistent with a particular role for Zip3 in preventing unwarranted SC assembly via an interaction with Zip1’s N terminal residues, and also with the idea that this Zip1-Zip3 interaction might be required to “trigger” synapsis initiation at MutSγ crossover sites, zip1[Δ2–9] and zip1[Δ10–14] mutants display an absence of MutSγ crossovers, a failure of stable Zip3 recruitment to several DSB sites, and premature SC assembly (albeit still between homologs). These phenotypes are consistent with the idea that, through an interaction with Zip1’s N terminus, Zip3 not only constrains Zip1-associated recombination intermediates to be MutSγ-dependent (possibly through the SUMOylation of targets such as the Red1 axis-associated protein), but also negatively regulates SC assembly until the successful completion of a specific intermediate event associated with the recombination process. Perhaps negative regulation of Ecm11 SUMOylation is linked to Zip3’s switch-like function in ensuring that SC assembly occurs “in the right place at the right time” (i.e. at a specific type of recombination intermediate, such as one designated to become a MutSγ crossover event). We note, however, that SC assembly is more robust and non-centromeric synapsis initiation events are more abundant in zip1[Δ2–9] and zip1[Δ10–14] relative to zip3 mutant strains; this difference could reflect additional pro-SC assembly activities of Zip3, carried out in a manner that is independent of Zip3’s engagement with Zip1’s N terminal tip.

While multiple attempts at two-hybrid and pull-down experiments have failed to reveal evidence of a strong physical interaction between Zip3 and Zip1’s N terminus, cytological support for this interaction comes from the localization of Zip3 at polycomplex structures, aggregates of Zip1 and other SC-associated proteins that form when SC assembly is compromised. While many if not all SIC proteins have been found to localize to Zip1 polycomplex structures, Zip3’s localization shows a greater degree of coincidence with Zip1 throughout the bulk of the polycomplex, for example relative to Zip4 ([34]; Fig 10A) or the MutSγ component, Msh4 [60]. Importantly, Zip3’s localization throughout the bulk of Zip1 polycomplex is not abolished when Zip4 (which has been found to interact with Zip3 [37]) is absent [34], however Zip3’s localization to the bulk of Zip1 polycomplex is completely abolished by the loss of Zip1’s residues 2–9 or 10–14 (Fig 10A). Finally, Zip1[Δ2–9] and Zip1[Δ10–14] have lost Zip1’s capacity to recruit Zip3 to sites of recombination initiation (Fig 11). Given our inability to observe a stable interaction between Zip1 and Zip3 via pull downs or two hybrid methods, it seems likely that Zip3 interfaces with Zip1 in a manner that is dependent on other proteins, on molecular structures nearby, such as the DNA joint molecule itself, or on a relatively unstable Zip1 structural configuration.

Based on the absence of SC assembly in meiotic cells expressing zip1[Δ15–20], zip1[15–20→A] and zip1[I18A, I19A], we furthermore conclude that Zip1’s first twenty residues correspond to (perhaps in an overlapping manner) at least one interaction domain for an SC assembly factor or complex of factors. Unlike the limited nature of the region within Zip1’s N terminal 163 amino acids that is required for MutSγ crossovers (twenty residues, based on the fact that zip1[Δ21–163] is fully capable of MutSγ crossing over [24]), groups of residues that are critical for allowing Zip1 to assemble SC may be distributed throughout the entire N terminal region encompassed by residues 15–163, as both zip1[Δ15–20] and zip1[Δ21–163] fail to assemble SC. Nevertheless, we propose that the region overlapping residues 15–20 interfaces with components serving an SC assembly function, based on the fact that alteration of two adjacent residues (I18 and I19) at Zip1’s extreme N terminus (a change that is unlikely to alter the overall length or structure of the rod-like protein) completely abolishes Zip1’s capacity to build the SC.

We previously demonstrated that zip1[Δ21–163] phenocopies the ecm11 and gmc2 null mutant phenotype (a failure in SC assembly but proficiency in crossing over), suggesting that this N terminal region of Zip1 functionally interacts with the central element in order to assemble SC [24]. Moreover, Leung et. al (2015) demonstrated that Zip1’s N terminal 346 residues is sufficient to promote Ecm11 SUMOylation in vegetative (non-meiotic) cells, provided that Gmc2 is also expressed. These data suggest that the N terminal region of Zip1 is able to engage with the Ecm11-Gmc2 proteins, perhaps in a direct manner or perhaps indirectly through a protein expressed in both meiotic as well as mitotic cells [61]. Similar to the uncertainty about whether Zip3 interacts with Zip1 in a direct manner, apart from the genetic interactions found for Zip1 and Ecm11 and the coincidence of Ecm11 and Gmc2 at the midline of SC (where Zip1 N termini also reside [13]), strong evidence of a direct physical interaction between Zip1 and Ecm11 or Gmc2 does not yet exist.

The adjacency between Zip1’s pro-crossover and a pro-synapsis regions may serve as a liaison to coordinate SC assembly with intermediate steps in recombination

Finally, we note the tantalizing possibility that the adjacency between the putative pro-crossover and pro-synapsis regions of Zip1’s N terminus is functionally important for ensuring that SC assembly occurs in coordination with intermediate steps in the MutSγ crossover recombination pathway. Specifically, we speculate that Zip1 may physically connect crossover recombination events to SC assembly through a mechanism that is based, at least in part, on its capacity to stabilize Zip3 at its N terminus. Here, Zip3 would be expected to be in close proximity to putative pro-synapsis factors stabilized (perhaps in conjunction with other SIC proteins) by the adjacent region in Zip1, and thus could potentially be oriented appropriately to regulate the extent of SUMOylation of SC central element protein Ecm11.

SCs assembled in the absence of MutSγ crossing over in budding yeast may be less stable

During the course of analyzing SC assembly over a time course of meiotic progression in our mutants we found that SCs assembled in zip3, zip1[D2-9] and zip1[10–14] strains (corresponding to SCs assembled in the absence of MutSγ crossovers) assemble earlier than wild-type SC structures, and appear to be less capable of persisting during an ndt80-mediated, meiotic prophase arrest (Fig 4, S2 Fig). Pattabiraman (2017) found that a MutSγ-associated process affects the dynamic properties of C. elegans SC [39]; our set of preliminary observations raises the intriguing possibility that the MutSγ crossover pathway influences the structure and/or dynamics of budding yeast SCs in a similar fashion.

Strains

Yeast strains used in this study are isogenic to BR1919-8B [62] and were created using standard genetic crosses and manipulation procedures. CRISPR-Cas9 methodology was utilized to create unmarked alleles (described below). Strains for crossover analysis carry an hphMX4 cassette inserted near the chromosome III centromere, ADE2 inserted upstream of the RAD18 locus, a natMX4 cassette inserted near the HMR locus, TRP1MX4 inserted 62 bp downstream of the SPO11 locus (Kee and Keeney, 2002), URA3 replacing SPO13, and LYS2 inserted on chromosome VIII at coordinate 210,400 bp. Strains SYC107, SYC149, SYC151 and K914 do not carry THR1 nor LYS2 on chromosome VIII. Zip3 and Zip4 epitope tags (MYC and HA, respectively) are positioned internal to the gene ORFs, as described in [34]. ChIP and qPCR experiments were performed in strains of the SK1 genetic background (S4 Table).

Creation of unmarked zip1 alleles using CRISPR-Cas9

Custom zip1 alleles were created in two sequential transformation steps: The first step is the creation of a “base strain”, which entails replacing the DNA sequences to be altered (in this case, within the ZIP1 locus) with the kanMX4 dominant drug resistance cassette. Next, ~500 ng of a CRISPR-Cas plasmid pRS425-Cas9-kanMX, created by Gang Zhao in Bruce Futcher’s laboratory (Stony Brook University), is transformed into the zip1-kanMX4 base strain along with custom DNA sequences (“healing fragments”) that carry the desired alteration in DNA sequence as well as homology to sequences flanking the kanMX4 insert. pRS425-Cas9-kanMX is a yeast two micron plasmid carrying sequences that encode the LEU2 gene, the CAS9 gene (driven by the TEF1 promoter) and a unique CRISPR guide RNA (driven by the RNA polymerase III promoter, SNR52) that targets the kanMX gene. For our strains, “healing fragments” typically correspond to two overlapping ZIP1 DNA sequences sharing at least twenty bases of overlap that encode the DNA changes desired in the new allele. These DNA fragments were amplified by PCR using a template with sequences containing the wild type ZIP1 gene. The 5’ healing fragment is created using a reverse primer containing the desired DNA changes and a forward primer that has homology to ZIP1 sequences 5’ of the kanMX4 insertion, whereas the 3’ healing fragment is created using a forward primer containing the desired DNA changes (usually a reverse complement DNA fragment to the reverse primer used to create the 5’ fragment) and a reverse primer corresponding to ZIP1 sequences 3’ to the kanMX4 insertion. Primers are positioned such that the PCR products contain have at least 50bp of DNA homologous to the regions flanking kanMX4 in the base strain. After transformation of the zip1-kanMX4 strain with pRS425-Cas9-kanMX plasmid DNA along with both “healing” DNA fragments, cells are plated onto synthetic medium lacking leucine and incubated for up to 10 days at thirty degrees Celsius. Leu+ colonies survive due to the presence of the pRS425-Cas9-kanMX plasmid and because of a DNA repair event that replaced kanMX4 with “healing fragment” sequences. Leu+ colonies are screened for G418 sensitivity; G418-sensitive transformants are struck out on YPD media to isolate single colonies and then genotyped by PCR. For potential mutant allele transformants (based on the PCR genotyping), the entire gene ORF is sequence verified.

Cytological analysis and imaging

Meiotic nuclei were surface spread on glass slides and imaged as described in [24]. The following primary antibodies were used: affinity purified rabbit anti-Zip1 (1:100, raised at YenZym Antibodies, LLC, against a C terminal fragment of Zip1, as described in [43], mouse anti-cMYC (1:200, clone 9E10, Abcam). Mouse anti-Gmc2 antibodies were raised against purified Gmc2 protein, and guinea pig anti-Gmc2_Ecm11 antibodies were raised against a co-purified protein complex (ProSci Inc.). These antibodies were used at 1:800. Chicken anti-HA (1:100, Abcam), and rabbit anti-Red1 (1:100, a kind gift from G.S. Roeder, [59]) were also used. Secondary antibodies conjugated with Alexa Fluor dyes were purchased from Jackson ImmunoResearch and used at 1:200 dilution. Microscopy and image processing were performed using a Deltavision RT imaging system (General Electric) adapted to an Olympus (IX71) microscope. Measurements of Zip1, Ecm11, and SC linear structures in Fig 4 and S2 Fig (raw data in S5 Table) were measured manually by K.V.M. (“ImageK”), using the measurement tool in the SoftWorx program associated with the Deltavision RT system. Structured illumination microscopy was carried out using Applied Precision’s OMX Blaze Structured Illumination Microscope system at The Rockefeller University’s Bio-Imaging Resource Center.

Calculations and statistical analysis

Genetic crossover data was compiled and processed using an Excel Linkage Macro program, created by Jonathan Greene (Rhona Borts, pers. comm.) and donated by Eva Hoffmann (University of Copenhagen, Denmark). Crossover values (and their standard errors) were obtained using the Stahl lab online tools (https://elizabethhousworth.com/StahlLabOnlineTools/), with the method of Perkins [63]. Non-mendelian segregation is reported in S3 Table. Recombinant spore values were calculated according to the following: 100(r/t), where r = the number of colonies carrying a chromosome which is recombinant in the interval and t; the total number of colonies assessed. Standard error (S.E.) values for random spore analysis were calculated according to the formula: 100(√(r/t)(1-r/t)/t) [64]. All other statistical analyses were carried out using Graphpad Prism or Graphpad InStat (www.graphpad.com).

Western blot

Western blotting was performed as described previously [13] with the following modifications: Amersham Protran 0.2μm NC was used as the transfer membrane following the manufacturer’s recommendation; after secondary antibody incubation the membrane was processed with a final wash in 100mM Tris-Cl pH 9.5, 100mM NaCl, 5mM MgCl₂ to boost the HRP- mediated chemiluminescence using Amersham ECL Prime Western Blotting Detection Reagent. Signals were detected on a Syngene G:Box and measured using Syngene GeneTools software; the areas being assessed for measurements were manually refined in order to ensure all and only appropriate data from each lane is collected.

Chromatin immunoprecipitation

Meiotic cells were processed as described [65], with the following modifications: Lysis was performed in Lysis buffer plus 1 mM PMSF, 50 μg/mL Aprotinin and 1X Complete Mini EDTA-Free (Roche), using 0.5 mm zirconium/silica beads (Biospec Products, Bartlesville, OK). 2 μg of the mouse monoclonal anti-FLAG antibody M2 (Sigma) and 30 μL Protein G magnetic beads (New England Biolabs) were used. Quantitative PCR was performed from the immunoprecipitated DNA or the whole-cell extract using a 7900HT Fast Real-Time PCR System (Applied Biosystems, Thermo Scientific) and SYBR Green PCR master mix (Applied Biosystems) as described [65]. Results were expressed as % of DNA in the total input present in the immunoprecipitated sample. Primers for GAT1, BUD23, ERG1, Axis and NFT1 loci have been described [50, 51, 66].