Meiosis-Specific Functions of Kinesin Motors in Cohesin Removal and Maintenance of Chromosome Integrity in Budding Yeast

Kinesin motors provide the molecular forces at the kinetochore-microtubule interface and along the spindle to control chromosome segregation. During meiosis with two rounds of microtubule assembly-disassembly, the roles of motor proteins remain unexplored. We observed that in contrast to mitosis, Cin8 and Kip3 together are indispensable for meiosis. While examining meiosis in cin8Δ kip3Δ cells, we detected chromosome breakage in the meiosis II cells.

ABSTRACT

Kinesin motors provide the molecular forces at the kinetochore-microtubule interface and along the spindle to control chromosome segregation. During meiosis with two rounds of microtubule assembly-disassembly, the roles of motor proteins remain unexplored. We observed that in contrast to mitosis, Cin8 and Kip3 together are indispensable for meiosis. While examining meiosis in cin8Δ kip3Δ cells, we detected chromosome breakage in the meiosis II cells. The double mutant exhibits a delay in cohesin removal during anaphase I. Consequently, some cells fail to undergo meiosis II and form dyads, while some, as they progress through meiosis II, cause a defect in chromosome integrity. We believe that in the latter cells, an imbalance of spindle-mediated force and the simultaneous persistence of cohesin on chromosomes cause their breakage. We provide evidence that tension generated by Cin8 and Kip3 through microtubule cross-linking is essential for signaling efficient cohesin removal and the maintenance of chromosome integrity during meiosis.

INTRODUCTION

Meiotic chromosome segregation comprises certain unique events distinct from mitosis. In budding yeast, these events include late assembly of the mature kinetochore competent to connect the microtubules, pairing of homologs, mono-orientation of the sister kinetochores with respect to the spindle pole in metaphase I, a stepwise dissolution of cohesin from chromatin, two rounds of chromosome segregation with spindle assembly and disassembly, and partial dephosphorylation of the cyclin-dependent kinase (CDK) substrates adequate for spindle disassembly at meiosis I but not for DNA replication. Furthermore, while dephosphorylation of CDK substrates by Cdc14 phosphatase released by the mitotic exit network (MEN) pathway from its inhibited state is essential for cell cycle exit in both mitosis and meiosis, Cdc14 released by the Cdc14 fourteen early anaphase release (FEAR) pathway appears to be dispensable for the same in mitosis but not in meiosis, as it is required to exit from meiosis I (1–3). Nevertheless, FEAR-dependent Cdc14 release has significant roles in both mitosis and meiosis for coherent segregation of all chromosomal loci and for stability and proper orientation of the microtubule spindle (2, 4–7).

Irrespective of the type of cell cycle, the formation of a microtubule-based spindle and movement of chromosomes along the spindle being attached to the microtubule play a pivotal role during chromosome segregation. The occurrence of these events relies on the polymerization-depolymerization property of the microtubules, which is facilitated by the functions of several microtubule-associated proteins (MAPs) and microtubule-based motors (8). In Saccharomyces cerevisiae, four nuclear motors of the kinesin superfamily (kinesin-related proteins [KRPs]), namely, Cin8, Kip1, Kip3, and Kar3, have essential roles in chromosome segregation (9–13). However, due to functional redundancy among these motors, they are nonessential for growth (14, 15).

Cin8 and Kip1 belong to the BimC or kinesin-5 family of proteins, where the motor domain is at the amino-terminal end of the protein and motor movement is directed toward the plus end of the microtubule (16, 17). Cin8 and Kip1 form homotetramers, and their plus-end-directed functions are to extend the spindle by pushing the poles apart and to maintain the kinetochores in a clustered form through cross-linking of antiparallel and parallel microtubules, respectively (18–22). Later, through in vitro assays, minus-end-directed movement of both single motors has been identified when they work singly on an individual microtubule (23–26). Recently, it has been shown under in vivo conditions that Cin8 clusters at the minus end and spindle pole bodies (SPBs) during the early stage of mitosis for capturing the microtubules emanating from opposite SPBs, which facilitates bipolar spindle formation (27). However, the implication of Kip1 minus-end-directed movement has not been explored. In addition to the cross-linking function, Cin8 and, to a lesser extent, Kip1 can also depolymerize kMT (kinetochore-microtubule) in a length-dependent manner, which is believed to be essential for congression of the chromosomes (28). The regulation of Cin8 and Kip1 functions depends on the phosphorylation status of these proteins, where their phosphorylation by Cdk1 during early mitosis mediates SPB separation (29). In metaphase, Cin8 and Kip1 are localized at the centromeres and along the length of the microtubule (13). Since the phosphorylation of Cin8 inhibits its association with the microtubules (30), following the metaphase-to-anaphase transition, dephosphorylation of Cin8 by protein phosphatase 2A regulatory subunit Cdc55 (PP2A^Cdc55) and Cdc14 phosphatase results in its accumulation near the spindle poles and at the spindle midzone, which is crucial for spindle elongation (31, 32). However, it is not known if a similar dephosphorylation also occurs in Kip1. During early anaphase, anaphase-promoting complex-bound activator protein Cdc20 (APC^Cdc20) degrades Kip1 (33), whereas Cin8 is degraded during late anaphase by anaphase-promoting complex-bound activator protein Cdh1 (APC^Cdh1) (34). On the other hand, the primary function of the Kip3 motor, belonging to the kinesin-8 family of proteins, is the depolymerization of microtubule plus ends by a mechanism similar to that of kinesin-13 motors (12, 35), which has a role in the movement of chromosomes during anaphase (13, 36). However, Kip3 also slides and clusters the microtubules by cross-linking antiparallel and parallel microtubules, respectively, through its tail domain (37). However, the cross-linking function of Kip3 is trivial compared to kinesin-5 proteins owing to its intrinsic structural ability to form homodimers but not the homotetramers observed in kinesin-5 motors (18–22, 37). Kip3 activity appears to be regulated spatially and temporally based on the length of the spindle and the exact localization of the motor. On a short spindle, it helps in clustering and alignment of the kinetochores by cross-linking of the parallel microtubules and depolymerase activity at the plus ends. During an increase in the spindle length, Kip3 cross-links and slides the antiparallel interpolar microtubules. Finally, when the spindle reaches its maximum length, Kip3 localizes at the plus ends and causes spindle disassembly by its depolymerization activity (22, 38). Kar3 (a minus-end-directed kinesin-14 family protein) is another microtubule depolymerizer present in the cell and is functionally antagonistic to Cin8/Kip1 spindle elongation activity. Kar3 pulls two spindle poles together; therefore, the spindle collapse observed in the absence of both Cin8 and Kip1 can be suppressed by reducing the activity of Kar3 (39). Additionally, Kar3 appears to promote kinetochore-microtubule attachment, as in mitosis, it is found to occupy a subset of kinetochores on which microtubule attachments are slow to form (13).

As described above, several groups have elucidated the functions of nuclear kinesin motors in chromosome segregation in mitosis. Given the mechanistic uniqueness in chromosome segregation in meiosis, as outlined above, it is intriguing to investigate their functions during this cell cycle. However, a KAR3 mutant was found to be arrested at prophase I (40, 41), which makes it difficult to analyze the meiotic events in the absence of Kar3. Therefore, in this study, we focused on elucidating the functions of three motors, Cin8, Kip1, and Kip3, in meiosis. Using knockout mutants, we observed that these motors are required for homolog pairing. Strikingly, we noticed that cells with a loss of both Cin8 and Kip3 harbor chromosome breakage. Further investigation argues for a defect in Rec8-cohesin removal from chromatin in these cells. We propose that the conditions in the absence of Cin8 and Kip3 perhaps create an imbalance between the microtubule-mediated force generated by other motors and the resisting force by persistent cohesin, which may lead to chromosome breakage. From our findings, we suggest that the tension generated by the cross-linking activity of Cin8 and Kip3 is crucial to signal cells for cohesin cleavage. Thus, our study reveals significant roles of kinesin motors in meiosis and hints at the essentiality of these proteins in suppressing aneuploidy during gametogenesis.

RESULTS

The motors are required for faithful meiosis.

In the first set of experiments, we compared spore viabilities, a readout for faithful meiosis, between the wild type and the individual motor mutants. Given that there are functional redundancies among the motors, we observed a marginal decrease in spore viability in kip1Δ and kip3Δ mutants (approximately 89 and 92%, respectively). However, the cin8Δ mutant showed an ∼65% reduction in spore viability, suggesting that this protein is more significant in meiosis (Fig. 1A). It is expected that the pace of meiotic progression can slow down if there is any perturbation in meiosis. To test this, the wild-type and mutant strains were released into synchronized meiosis. Consistent with the spore viability data, we observed that the cin8Δ mutant showed a delay at metaphase I compared to the wild type and the kip1Δ and kip3Δ mutants (Fig. 1Bii), suggesting that some defect is occurring during early meiotic events in the absence of Cin8 and that perhaps, due to functional redundancy, the defect is not apparent in kip1Δ and kip3Δ mutants. To investigate if the defect causes chromosomes to missegregate, we marked both CenV homologs with the TetO/TetR-green fluorescent protein (GFP) system (see Materials and Methods) and observed their distribution at the end of meiosis. Following faithful meiosis, a tetranucleated cell would show one GFP dot in each nucleus (type I) (Fig. 1D). However, four GFP dots in three and two nuclei (types II and III) or in one nucleus account for chromosome missegregation. Meiotic induction, unless otherwise mentioned, was carried out at 33°C, as the phenotype of the loss of Cin8 becomes aggravated at a higher temperature (13, 42, 43). We observed around 50, 22, and 17% (type II and type III) chromosome missegregation in cin8Δ, kip1Δ, and kip3Δ cells, respectively (Fig. 1D), suggesting that the spore viability defect is probably due to the generation of aneuploidy. As in the cin8Δ mutant, the delay in the cell cycle occurs in metaphase I, we presumed that at least some defects might be occurring during the preceding events of chromosome segregation, which include chromatid cohesion, homolog pairing, and sister chromatid mono-orientation. To investigate cohesion between the sisters and the orientation of their spindle attachment, both sisters of one homolog were marked with the TetO/TetR-GFP system. In metaphase I-arrested cells, a defect in sister chromatid mono-orientation would appear as two GFP dots. On the other hand, noncohesed sisters in the cycling cells would produce binucleates with one GFP dot in each nucleus. However, we failed to detect any defect in either sister chromatid mono-orientation or their cohesion (Fig. 2A and B). Although not for Cin8, Kip1, or Kip3, the role of Kar3 in sister chromatid cohesion in mitosis has been reported previously (44).

FIG 1.

Meiosis in the motor mutants. (A) Spore viabilities of wild-type (SGY5001) (n = 40), cin8Δ (SGY315) (n = 60), kip1Δ (SGY317) (n = 107), and kip3Δ (SGY314) (n = 91) cells were analyzed after induction of meiosis at 30°C. “n” represents the number of tetrads dissected. (B) The indicated strains were induced for synchronized meiosis and analyzed for meiotic progression. At the indicated time points, the fraction of cells at different stages of meiosis was determined by anti-α-tubulin staining. At least 100 cells were counted for each time point. (C) Representative images of cin8Δ kip3Δ cells showing the maximum population at the anaphase I stage at 5 h of meiotic induction as determined by anti-α-tubulin and DAPI staining. Arrows indicate anaphase I cells. Bar, 5 μm. (D) The indicated strains harboring homozygous CenV-GFP (see Materials and Methods) were analyzed for meiotic chromosome segregation at 33°C in tetranucleated cells (n = 90 to 150). Error bars represent the standard deviations from the mean values obtained from three independent experiments. Bar, 2 μm.

FIG 2.

Motor proteins are required for homolog pairing and their disjunction but do not have a role in sister chromatid mono-orientation or cohesion. (A and B) Heterozygously marked CenV-GFP dots were analyzed for meiotic chromosome segregation at 33°C. (A) Metaphase I-arrested wild-type (SGY5143) (n = 90), cin8Δ (SGY5078) (n = 124), kip1Δ (SGY5018) (n = 91), and kip3Δ (SGY5034) (n = 71) cells were analyzed for the percentage of mononucleated cells with one or separated sister centromeres (n = 70 to 120). (B) Wild-type (SGY5006) (n = 95), cin8Δ (SGY5090) (n = 122), kip1Δ (SGY5077) (n = 97), and kip3Δ (SGY309) (n = 96) cells were analyzed for the percentage of binucleates with one or two GFP dots in one nucleus (types I and II, respectively) or one dot each in two nuclei (type III). (C) Wild-type (SGY263), cin8Δ (SGY5197), kip1Δ (SGY5347), and kip3Δ (SGY5190) cells arrested at prophase I by Ndt80 depletion and harboring homozygous CenV-GFP were analyzed by chromosome spreads for the number of GFP dots. One, two, or more than two 2 GFP dots were scored as paired, unpaired, or unpaired with noncohesed sister chromatids, respectively. (D) Cells analyzed in panel C were also observed for the Zip staining. n = 150 to 220 for panels C and D. (E) Strains in Fig. 1A were analyzed at the binucleated stage for the disjunction of CenV homologs (n = 50 to 110). Types I and II indicate disjunction and nondisjunction of the homologs, respectively. Error bars represent the standard deviations from the mean values obtained from three independent experiments, and “n” represents the number of cells scored. Bars, 2 μm.

However, we observed an increased defect in homolog pairing in the cin8Δ mutant when both CenV homologs were marked with GFP (type II and type III) (Fig. 2C). Consistent with this, we observed a higher percentage of mislocalization (polycomplex formation) of Zip1, a component of the synaptonemal complex (SC) that reinforces pairing (Fig. 2D). A similar result was also obtained previously, where homologs failed to synapse in the absence of Kar3 (40). Although both the cin8Δ and kip1Δ mutants showed defects in the localization of Zip1, only cin8Δ cells showed a homolog pairing defect. Since Cin8 has a more significant role than Kip1 in mitosis (16) and in meiosis (as shown by spore viability assays) (Fig. 1A), more roles of Cin8 than of Kip1 are expected. Following disassembly of the SC, Zip1 is maintained at the centromeres until the proper bipolar attachment of the homologs is achieved (45). As the Zip1 localization was compromised in the cin8Δ or kip1Δ mutant, we examined the homolog biorientation of motor mutants where both CenV homologs were marked with GFP. About 37% of the binucleated cells of the cin8Δ mutant showed homolog nondisjunction, compared to only 7% in the wild type (type II) (Fig. 2E), while for the kip3Δ and kip1Δ mutants, the populations exhibiting such a defect were relatively smaller (14% and 11%, respectively) (Fig. 2E). The above-described results suggest that the absence of motor proteins, especially Cin8, can affect homolog pairing in meiosis.

Meiosis is profoundly compromised in the cin8Δ kip3Δ double mutant.

Since Cin8, Kip1, and Kip3 share overlapping functions in microtubule cross-linking and depolymerization (10, 12, 13, 16, 28, 38, 46), we argued that their functions cannot be properly revealed by studying only the single mutants. Therefore, we generated the two only possible viable double mutants, kip1Δ kip3Δ and cin8Δ kip3Δ, as the cin8Δ kip1Δ mutant has been reported to be inviable (9, 16, 19, 47). Although the sporulation efficiencies of the cin8Δ, kip3Δ, and cin8Δ kip3Δ strains were similar (63%, 85%, and 70%, respectively) (Table 1) after 12 h of sporulation induction, strikingly, we observed a precipitous drop (approximately 16%) in the spore viability of the double mutant compared to the cin8Δ and kip3Δ single mutants (Fig. 1A and Fig. 3A). To further investigate the probable roles of Cin8 and Kip3 together in meiosis, we monitored meiotic progression in the wild type and the motor mutants and noticed that in comparison to the wild-type, cin8Δ, or kip3Δ strain, the cin8Δ kip3Δ double mutant proceeded through meiosis slowly, and the majority of the cells were arrested transiently at anaphase I, with one spindle and an improper disjunction of nuclei (Fig. 1Biii and Fig. 1C).

TABLE 1.

Sporulation efficiencies

Strain	% sporulation efficiency
Wild type	81
cin8Δ strain	62.9
kip1Δ strain	74.5
kip3Δ strain	85.2
cin8Δ kip3Δ strain	69.4

FIG 3.

Meiosis in cin8Δ kip3Δ cells results in the frequent formation of dyads with aneuploid spores. (A and B) kip1Δ kip3Δ (SGY5104) (n = 54) and cin8Δ kip3Δ (SGY5089) (n = 117) cells were analyzed for spore viability (A) and the formation of dyads and tetrads (B) following 12 h of meiotic induction (n = 195 to 351). The maximum population of cin8Δ kip3Δ sporulated cells forms dyads, with a small population of tetrads. “n” represents the total numbers of tetrads dissected (A) and sporulated cells (B). (C) Percentages of dyads with one GFP dot each in two spores (1:1) and one or two GFP dots in one spore (1:0 or 2:0, respectively) in cin8Δ kip3Δ (SGY5154) cells (n = 136) harboring heterozygous CenV-GFP. Bar, 2 μm. (D) Spore viability of the dyads formed in cin8Δ kip3Δ (SGY5089) cells following meiosis at 30°C. Sixty dyads were dissected for viability estimation. Error bars represent the standard deviations from the mean values obtained from three independent experiments.

Given that Cin8 and Kip3 can cross-link and slide the antiparallel microtubules causing spindle elongation, our results indicate that cells lacking both Cin8 and Kip3 cause slow spindle elongation and defects in chromosome disjunction during meiosis I that might be responsible for the delay in spindle disassembly and completion of meiosis I. Due to this delay, around 50% of the cin8Δ kip3Δ cells proceeded to meiosis II without completing meiosis I, as assessed by the separation of sister chromatids on the meiosis I spindle and produced dyads (Fig. 3B and C). The inability to complete meiosis I due to a defect in spindle elongation but the ability to proceed to meiosis II and generate dyads with two diploid spores are hallmarks of the FEAR mutants (1, 2, 48, 49). Additionally, similar to the FEAR mutants (2), in cin8Δ kip3Δ cells, we also observed reductional segregation of chromosomes in the dyads, as both the heterozygously tagged CenV-GFP dots (sister chromatids) were found in one spore in 76% of the dyads (Fig. 3C). However, cosegregation of the sister chromatids per se does not imply prevention of meiosis II in cin8Δ kip3Δ cells, since in many dyads, we observed stained nuclei that were not included in the spores, suggesting that massive missegregation had occurred during meiosis II as well. Consequently, the viability of the dyad spores obtained from cin8Δ kip3Δ cells was extremely poor (10%) (Fig. 3D).

We believe that the phenotypes of cin8Δ kip3Δ cells are similar to those of the FEAR mutants, as the Cdc14 phosphatase released by the FEAR network promotes spindle elongation through dephosphorylation of Cin8, which facilitates its binding to the spindles and sliding of the antiparallel microtubules (7, 31). However, the removal of Cin8 alone did not exhibit as severe a phenotype as that of the FEAR mutants due to functional redundancy in spindle elongation between Cin8 and Kip1/Kip3 and due to additional functions of the FEAR network (50). It is also expected that the FEAR mutant-like phenotypes observed in meiotic cin8Δ kip3Δ cells will also be observed in mitosis. Since the FEAR mutants exhibit a delay in mitotic exit (51), the wild type and the motor mutants were released synchronously using α-factor into fresh yeast extract-peptone-dextrose (YPD) medium to compare the paces of mitosis. As observed for the esp1-1 FEAR mutant (51, 52), we observed a delay in cell cycle progression in cin8Δ and cin8Δ kip3Δ cells. Wild-type, kip1Δ, and kip3Δ cells completed one cycle of mitosis in approximately 55 min, while in the cin8Δ and cin8Δ kip3Δ mutants, it was delayed (around 75 min) (as shown by the dashed and dotted lines, respectively, in Fig. S1A at http://www.bio.iitb.ac.in/~santanu/wp-content/uploads/2020/05/Supplementary_file-final.pdf). In the time window between 90 and 100 min (see Fig. S1B at the URL mentioned above), the wild-type, kip1Δ, and kip3Δ strains exhibit a second peak for metaphase cells, while the cin8Δ strain demonstrates only one peak, whereas in the cin8Δ kip3Δ strain, the same peak is broadened and further extended until 105 min, suggesting that the metaphase-to-anaphase transition is maximally delayed in the double mutant. Additionally, we also observed a phenotype in the cin8Δ and cin8Δ kip3Δ mutants at equal frequencies, where we found a persistent population of cells with elongated nuclei spanning the mother cell and daughter bud but with a bipolar spindle of a metaphase-specific length (type II) (see Fig. S1C at the URL mentioned above). Such a phenotype could be due to an inability to extend the spindle but with the to-and-fro movement of the short spindle resulting in nuclear elongation, which has been observed previously with metaphase-arrested short spindles (53).

From the above-described results, it is apparent that the absence of both Cin8 and Kip3 causes defects in spindle elongation and the metaphase-to-anaphase transition in both mitosis and meiosis. However, it is apparent that while in meiosis, cin8Δ kip3Δ cells show more defects in spindle elongation and in the metaphase-to-anaphase transition than the single mutants, such a difference is absent in mitosis. Consequently, in contrast to the poor spore viability following meiosis observed for the cin8Δ kip3Δ mutant (Fig. 3A), we failed to observe any difference in viabilities among the wild type, the single mutants, and the cin8Δ kip3Δ double mutant following mitosis (see Fig. S1D at the URL mentioned above). This is further supported by the fact that while the pace of meiosis was affected to a greater extent in the cin8Δ kip3Δ mutant than in the wild type or the single mutants (Fig. 1B), the mitotic growth rates were not affected to that extent (see Fig. S1E at the URL mentioned above). These results suggest that the loss of both Cin8 and Kip3 perhaps causes some meiosis-specific defects, as revealed below.

Meiotic chromosome segregation is largely perturbed in the cin8Δ kip3Δ mutant.

To examine if there are any meiosis-specific defects in the cin8Δ kip3Δ mutant, we sought to investigate meiotic chromosome segregation under these conditions. We used wild-type, cin8Δ kip3Δ and kip1Δ kip3Δ double mutant, and the corresponding single mutant cells where both homologs of chromosome V were marked with GFP. Since we observed that the cin8Δ kip3Δ mutant did not sporulate at a temperature of 33°C, meiosis induction was carried out at 30°C. We analyzed tetranucleated cells to ensure that both meiosis I and meiosis II had occurred. Wild-type, kip3Δ, kip1Δ, and kip1Δ kip3Δ cells mostly showed (100%, 92%, 84%, and 88%, respectively) four nuclei with one GFP dot in each nucleus (type I) (Fig. 4A), which was reduced slightly in cin8Δ cells and largely in cin8Δ kip3Δ cells (79% and 44%, respectively). The type II and III categories, having GFP dots in three or two nuclei, respectively, and which account for missegregation of the chromosomes, were found correspondingly more often in the mutants. Unexpectedly, a significant population of tetranucleates (approximately 30%) harboring >4 (termed “supernumerary”) CenV-GFP dots was found in cin8Δ kip3Δ cells (type IV category) (Fig. 4A), while a minute population of this category was observed in kip1Δ kip3Δ cells (6%). The difference observed between the double mutants can be expected since Cin8 is known to have more significant cell cycle functions than Kip1 from a previous mitotic study (16). The supernumerary GFP dot phenotype is not specific for chromosome V since the same phenotype was also observed (approximately 35%) (type IV) in the cin8Δ kip3Δ mutant when chromosome III (ChrIII) was marked using the LacO/LacI-GFP system at the pericentromeric region (22 kb away from CenIII) (Fig. 4B). To determine the stage of the cell cycle at which these supernumerary foci start appearing, Tub1 was N terminally tagged with cyan fluorescent protein (CFP) in the cin8Δ kip3Δ strain harboring homozygous CenV-GFP. Chromosome abnormality was found only in the cells with two spindles, suggesting that >4 foci were generated in cells that had passed through meiosis II (Fig. 4C). This numerical abnormality is specific for meiosis and did not occur due to aneuploidy generated as a legacy of an error during previous mitosis since we failed to obtain >2 GFP dots before entry into meiosis (see Fig. S2A at http://www.bio.iitb.ac.in/~santanu/wp-content/uploads/2020/05/Supplementary_file-final.pdf) or during any stage of mitosis (see Fig. S2B at the URL mentioned above) in cin8Δ kip3Δ cells.

FIG 4.

The cin8Δ kip3Δ mutant generates aneuploid tetranucleated cells showing supernumerary GFP foci of the marked chromosomes. (A) Tetranucleated cells of the wild-type (SGY5001), cin8Δ (SGY315), kip1Δ (SGY317), kip3Δ (SGY314), kip1Δ kip3Δ (SGY5104), and cin8Δ kip3Δ (SGY5089) strains harboring homozygous CenV-GFP were analyzed for meiotic chromosome segregation at 30°C (n = 100 to 309). “n” represents the number of tetranucleated cells scored for chromosome segregation. (B) Tetrads from the wild-type (SGY5407) (n = 129) and cin8Δ kip3Δ (SGY5329) (n = 119) strains harboring homozygous ChrIII-GFP marked at the LEU2 locus 22 kb away from the centromere were analyzed as described above for panel A. Type I represents one GFP focus in each of the four spores, whereas types II and II represent four GFP foci in three or two spores, respectively. Type IV is the tetrad with >4 GFP foci. (C) Supernumerary CenV-GFP foci are observed only in those cin8Δ kip3Δ cells (SGY5385) that are in meiosis II, as judged by the presence of two spindles (marked by arrows). Error bars represent the standard deviations from the mean values obtained from three independent experiments. Bars, 2 μm.

Chromosome breakage occurs in cin8Δ kip3Δ cells during meiosis II.

We next sought to address the reason for the generation of supernumerary GFP foci in cin8Δ kip3Δ cells. At least two possibilities can be envisaged for this. First, leaky chromosome replication between meioses I and II may amplify the operator arrays and cause >4 GFP foci. However, this possibility seems unlikely because if there is leaky replication of the operator array, due to close proximity (within 1.4 kb), CenV would also have been replicated, and in that case, >4 kinetochore foci would have been observed. However, we found the normal four Ndc10 (kinetochore) foci in 100% of cells with supernumerary CenV foci (see Fig. S2C at the URL mentioned above). Second, due to an imbalance of spindle force acting on the centromeres, the chromosomes may break, and since the operator arrays in our assays remain closed to the centromere, the arrays can also break to give >4 GFP foci. Given the functions of the motors in moderating spindle-chromosome interactions through force generation, the latter possibility is more likely. To investigate if there is indeed any chromosome breakage, a single-cell gel electrophoresis assay, known as the comet assay, was performed (54). As it is difficult to lyse the tetrad because of the robust spore wall, cells were analyzed for chromosome breakage at the tetranucleated stage before the formation of the spore wall. For the comet assay, the cells were released synchronously in meiosis. At the time of harvesting of the cells, there were >50% tetranucleated cells (observed by nucleus staining) in all the strains utilized for the assay, and ≥17% of the cells were at the binucleated stage (Fig. 5A). This observation suggests the completion of S phase in most of the cells and negates the possibility of the presence of any replication intermediates that might give a tail-like appearance in the comet assay. H₂O₂ (10 mM)-treated cells were used as a positive control for breakage (55). Interestingly, we obtained a notable population of DNA masses that formed tails or a comet phenotype in cin8Δ kip3Δ (approximately 20%) cells compared to wild-type (2.5%) or cin8Δ (1%) cells, while in the H₂O₂-treated sample, almost 46% of the cells exhibited the comet phenotype (Fig. 5B). To quantify the defect, we compared the percentages of DNA in the head and tail parts of the comet among the test samples (see Materials and Methods) (56). A reduction in the percentage of DNA in the head region is accompanied by an increase in the percentage of DNA in the tails of H₂O₂-treated and cin8Δ kip3Δ spheroplasts (Fig. 5C). The lack of tail structures in wild-type and cin8Δ cells is not due to the presence of sporulated cells that are resistant to Zymolyase treatment, since in the population of tetranucleated cells used for the comet assay, approximately 89%, 59%, and 66% of wild-type, cin8Δ, and cin8Δ kip3Δ cells were nonsporulated, respectively (Fig. 5D). Furthermore, the spores contained the spore wall, which was not visible in the spheroplasted nuclei of the above-described test samples under a bright-field microscope (Fig. 5E). Note that for comparison among the test samples, only spheroplasted cells were analyzed for the detection of the comets. Based on these criteria, the above-described results suggest that chromosome breakage occurs in cin8Δ kip3Δ cells.

FIG 5.

cin8Δ kip3Δ cells cause DNA breakage as they proceed through meiosis. (A) Nuclear stages of meiotic cells harvested for the comet assay (see Materials and Methods) from the sporulation medium (SPM) culture before the formation of the tetrads, which is at 8 h for cin8Δ kip3Δ (SGY5089) cells, 7 h for cin8Δ (SGY315) cells, and 5 h for wild-type (SGY40) cells. (B) Cells analyzed as described above for panel A were used for the comet assay. As a positive control, wild-type cells were treated with 10 mM H₂O₂ for 30 min at 4°C. The histograms correspond to the percentages of the cells that formed comets. Representative gel images (with “+” and “−” polarities) show ethidium bromide-stained DNA from each strain. “+” and “−” indicate the anode and the cathode, respectively. The arrow indicates an unspheroplasted cell. A total of 80 to 156 nuclei were observed for comet formation. Bars, 5 μm. (C) Percentages of DNA in the tail and head regions were calculated as described in Materials and Methods and plotted. (D) Cells with four nuclei observed in panel A were analyzed for the presence or absence of spore wall formation. (E) Representative images showing that, compared to spheroplasted cells, the cell wall is visible in cells that are not spheroplasted. Arrows indicate sporulated cells. (F to H) Chromosome spreads showing Rad52-EGFP foci stained using anti-GFP antibody in the wild-type (SGY5414), cin8Δ (SGY5422), and cin8Δ kip3Δ (SGY5415) strains. At least 40 spreads were analyzed for each type. Spindle poles marked by anti-α-tubulin antibody were used to judge the cell cycle stage of the spreads. Foci were counted after merging of the z-stacks, with the maximum intensities for tubulin and Rad52 keeping the threshold the same for all the fields. One tubulin dot (1 SPB) within one DAPI mass represents prophase I, while 2 dots (2 SPBs, 1 dot each in two DAPI masses) represent anaphase I, and 4 dots (4 SPBs, 1 dot each in four DAPI masses) represent the meiosis II stage. Bars, 2 μm.

To further reconfirm the chromosome breakage, we looked at the localization pattern of Rad52, which is required for the repair of DNA double-stranded breaks (DSBs) generated due to intrinsic or extrinsic factors (57–60). In meiosis, at prophase I, programmed DSBs occur for recombination, and consequently, Rad52 foci are visible at this stage in the wild type. However, once the DSBs are repaired, the average number of Rad52 foci reduces but persists in later stages of meiosis (61, 62). We counted and compared the Rad52-enhanced GFP (EGFP) foci in wild-type, cin8Δ, and cin8Δ kip3Δ chromosome spreads harboring 1 SPB within a single nucleus (prophase I stage) (Fig. 5F); 2 SPBs, 1 in each of the 2 nuclei (anaphase I stage) (Fig. 5G); and 4 SPBs, 1 in each of the 4 nuclei (anaphase II/post-meiosis II stage) (Fig. 5H). At prophase I, we observed an average of 29 foci in wild-type spreads, which was reduced to 9 ± 3 foci in cin8Δ spreads (Fig. 5F), which is consistent with the defective homolog pairing observed in cin8Δ cells (Fig. 2C). However, in cin8Δ kip3Δ cells, the average count was nearly 21, which suggests that the loss of Kip3 by an unknown mechanism rescues the defect of the cin8Δ mutant. While analyzing the spreads at anaphase I, we noticed no significant difference in Rad52-EGFP staining between wild-type and cin8Δ cells (wild-type spreads, 11 ± 7 foci; cin8Δ spreads, 10 ± 4 foci) (Fig. 5G) but observed a slight increase in staining for the double mutant (cin8Δ kip3Δ spreads, 15 ± 7 foci) (Fig. 5G). However, a drastic accretion in Rad52-EGFP staining was observed in cin8Δ kip3Δ spreads at anaphase II/post-meiosis II (tetranucleated stage) over the wild-type or cin8Δ spreads (wild-type spreads, 13 ± 6 foci; cin8Δ spreads, 11 ± 6 foci; cin8Δ kip3Δ spreads, 24 ± 6 foci) (Fig. 5H). These results indicate that as cin8Δ kip3Δ cells pass through meiosis II, they accumulate DNA damage in the form of DSBs (Fig. 4C).

Previously, we noticed supernumerary SPB formation in kinetochore mutants as the cells entered meiosis II (63). As both Cin8 and Kip3 also have some functional roles at the centromeres, we reasoned that in cin8Δ kip3Δ cells, following interphase II (stage between meioses I and II), maybe >4 SPBs or spindle poles are generated, and the resulting extra pole(s) may cause an imbalance of force and, hence, chromosome breakage. However, analysis of tetranucleated cin8Δ kip3Δ cells harboring >4 CenV-GFP foci showed only 4 SPBs (see Fig. S3 at http://www.bio.iitb.ac.in/~santanu/wp-content/uploads/2020/05/Supplementary_file-final.pdf), indicating that multipolarity is not the cause of chromosome breakage.

The cin8Δ kip3Δ strain hinders cohesin removal from chromatin in meiosis.

In budding yeast, cohesin is removed from the chromosome arms during anaphase I, while the removal of centromeric cohesin occurs during meiosis II. However, in FEAR mutants, the loss of cohesin from the arms is delayed during anaphase I (2). Since we noticed that cin8Δ kip3Δ cells exhibit phenotypes similar to those of FEAR mutants during meiosis (Fig. 1Biii and Fig. 3B), we therefore investigated if the double mutant is compromised in cohesin removal. We monitored Rec8-EGFP staining at different stages of meiosis in wild-type and cin8Δ kip3Δ cells. Meiotic stages were determined on the basis of the number of and distance between the Spc42 foci. Centromeric Rec8 was judged by its staining to be present only in the vicinity of the SPBs due to the proximity of the centromeres to the SPBs, whereas nuclear Rec8 comprised of arm plus centromeric Rec8, was identified by its presence spanning a broader region between the two SPBs (Fig. 6A and B). We observed that for the wild type, 64% of anaphase I cells displayed centromeric Rec8, which was reduced to 35% in cin8Δ kip3Δ cells (Fig. 6C). Rather, we noticed more cells with nuclear Rec8 in the double mutant (65%) than in the wild type (36%), suggesting a defect in cohesin removal during the metaphase I-to-anaphase I transition. Given that cohesin removal is completed during meiosis II, strikingly, nuclear Rec8 was observed even during the meiosis II stage in a staggering population (45%) of cin8Δ kip3Δ cells, whereas for the wild type, this population was insignificant (3%) (Fig. 6A, B, and D). It is possible that the expression of Rec8 is upregulated in cin8Δ kip3Δ cells, causing its accumulation and, hence, a defect in its efficient removal. To test this, we compared the levels of Rec8 expression between the wild-type and cin8Δ kip3Δ strains at the mRNA level. Since Rec8 expression peaks during the early stage of meiosis (64), we examined RNA levels at this stage. Based on the meiotic progression assay, we observed that following meiosis induction, at the 5-h time point, both strains harbored similar populations of metaphase I cells (Fig. 6G), and hence, RNA was isolated at this time point and quantified by reverse transcription followed by quantitative PCR-quantitative PCR (RT-qPCR). However, we failed to detect any significant difference in Rec8 levels between the wild type and the double mutant (Fig. 6H). This suggests that the greater Rec8 staining observed for the cin8Δ kip3Δ strain during anaphase I and subsequent stages is not due to an increase in its expression but is due to a defect in its removal.

FIG 6.

Removal of Rec8 cohesin is defective in the cin8Δ kip3Δ mutant. (A to D) Wild-type (SGY5557) and cin8Δ kip3Δ (SGY5523) cells harboring Rec8-EGFP and Spc42-CFP were analyzed for Rec8 localization at different stages of meiosis. (A) Representative images where Rec8 staining appearing as two tight-knit dots at the vicinity of each of the two SPBs or as a single such dot each between the two SPBs of two pairs was scored as centromeric Rec8 at anaphase I or meiosis II (metaphase II and anaphase II), respectively. Meiosis II cells with no Rec8 staining were also scored. (B) Representative images where Rec8 staining appearing in a broader region close to the SPBs or between the SPBs was scored as nuclear Rec8 (arm plus centromeric) at anaphase I or at meiosis II, respectively. (C and D) Percentages of anaphase I (C) and meiosis II (D) cells with the types of Rec8 staining in wild-type and cin8Δ kip3Δ strains. For panels C and D, 148 and 267 cells were analyzed for the wild-type and cin8Δ kip3Δ strains, respectively. Bars, 5 μm. (E and F) Chromosome spreads from wild-type (SGY5497) (n = 83), cin8Δ (SGY5501) (n = 114), and cin8Δ kip3Δ (SGY5500) (n = 183) cells harboring Rec8-EGFP were monitored at different stages of meiosis by EGFP immunostaining. (E) Quantitative analysis of Rec8 localization as only two foci (centromeric Rec8) or distributed throughout the chromatin (nuclear Rec8) in binucleated chromosome spreads. Representative images of each type are shown on the right. (F) Quantitative analysis of Rec8 localization with a tiny (single dot) or large (nuclear) appearance on the chromosome spreads from the tetranucleates. The tetranucleated stage of the spreads was determined by tubulin immunostaining. cin8Δ kip3Δ cells show significant Rec8 throughout the chromatin (nuclear), even in the tetranucleated stage. Representative images of each type are shown on the right. (G) Wild-type (SGY5533) and cin8Δ kip3Δ (SGY5534) strains tagged with Rec8-6HA were analyzed for different stages of meiosis at the indicated time points by tubulin immunostaining and DAPI staining. (H) Normalized mRNA expression level of Rec8 in the strains used for panel G. The Rec8 mRNA expression level does not change between the wild-type and cin8Δ kip3Δ strains. The values from the three independent biological replicates were averaged and plotted. Error bars represent the standard deviations from the mean values. A P value of ≥0.05 is considered nonsignificant (ns). Bars, 2 μm.

We obtained similar results in chromosome spreads immunostained for Rec8-EGFP, where both binucleated and tetranucleated spreads had high levels of nuclear Rec8 (82% and 54%, respectively) in the cin8Δ kip3Δ strain with respect to the wild type (19% and 11%, respectively) (Fig. 6E and F). Notably, we observed that in cin8Δ cells, nuclear Rec8 also persisted in a larger population of binucleated spreads (70%) (Fig. 6E). However, in the majority of the spreads at the tetranucleated stage, Rec8 appeared as a single dot, indicating the presence of a negligible amount on the chromatin, in contrast to the dispersed Rec8 signal present all over the chromatin in the double mutant in a higher percentage of the spreads (Fig. 6F). Altogether, these results suggest that a prolonged cohesin-chromatin association occurs in cin8Δ kip3Δ cells and, to a lesser extent, in cin8Δ cells. Due to this defect and the associated delay in spindle elongation and disassembly, cin8Δ and cin8Δ kip3Δ cells show a delay in the meiosis I-to-meiosis II transition (Fig. 1Biii). It is tempting to speculate that due to a higher level of cohesion retention in cin8Δ kip3Δ cells, the chromosomes cannot disjoin properly when subjected to pulling force exerted by the other motors during anaphase I and anaphase II spindle elongations, and they eventually break, causing very low spore viability.

As we observed a delay in spindle elongation and cell cycle progression in cin8Δ and cin8Δ kip3Δ cells, it is critical to address if the prolonged retention of Rec8 on chromatin is due to a delay in the degradation of securin (Pds1), a condition that releases separase to cleave Rec8. Under unperturbed conditions, Pds1 is degraded during the metaphase I-to-anaphase I transition following a reappearance in metaphase II and degradation in anaphase II. We monitored the levels of Rec8 and Pds1 through synchronized meiosis in wild-type, cin8Δ, and cin8Δ kip3Δ cells by immunoblotting (Fig. 7A and B). As described above (Fig. 1B), the pace of meiosis was delayed in cin8Δ kip3Δ cells compared to wild-type and cin8Δ cells (Fig. 7C); Pds1 degradation in the same strains also followed the same regime (Fig. 7A and B). Notably, with the disappearance of Pds1, all Rec8 was removed in wild-type cells, while removal was deferred in cin8Δ cells and, to a greater extent, in cin8Δ kip3Δ cells (Fig. 7A and B). Consistent with our cell biological data (Fig. 6), we noticed that in the double mutant, a significant level of Rec8 was persistent even at 15 h in meiosis, when around 90% of the cells had either entered into anaphase II or sporulated, whereas in such cells of either the wild-type or the cin8Δ strain, Rec8 was absent (Fig. 7A to C). Live-cell imaging of Pds1-EGFP also revealed that there is no difference in Pds1 stability on the anaphase I spindle between the wild type and the double mutant (Fig. 7D). These results suggest that protracted Rec8 retention on chromatin in cin8Δ kip3Δ cells is not due to a biochemical delay imposed by persistent Pds1.

FIG 7.

The cin8Δ kip3Δ mutant perturbs the coordination between Pds1 and Rec8 degradation in meiosis but does not affect the dynamics of Mcd1 removal in mitosis. (A) Western blot analysis of wild-type (SGY5534), cin8Δ (SGY5532), and cin8Δ kip3Δ (SGY5533) cells for the levels of Rec8-6HA and Pds1-13Myc at the indicated time points during meiotic progression. Tubulin was used as a loading control. (B) Densitometric analysis of Rec8-6HA and Pds1-13Myc signals obtained in panel A after normalization to the tubulin signal using ImageJ software. As evident from the graph, the drop in the Rec8 level, unlike Pds1, is much slower in cin8Δ kip3Δ than in wild-type or cin8Δ cells. (C) Percentages of prophase I, metaphase I, anaphase I, metaphase II, and anaphase II/sporulated cells determined by tubulin immunostaining at the time points utilized for Rec8-6HA and Pds1-13Myc detection in panels A and B (n ≥ 90 cells for each time point). (D) Analysis of Pds1-EGFP localization in wild-type (SGY5570) (n = 44) and cin8Δ kip3Δ (SGY5567) (n = 75) cells during the anaphase I stage determined by the distance between the two SPBs marked with Spc42-mCherry as shown in the representative images at the bottom. Pds1 staining under wild-type conditions, i.e., at the metaphase I stage, is also represented. (E) Localization of Mcd1-EGFP in wild-type (SGY5629) and cin8Δ kip3Δ (SGY5630) cells corresponding to the three-dimensional (3D) distances between the two spindle poles marked by Spc42-mCherry. Fields of view of metaphase (≤1.2-μm distance) and anaphase (≥2.5-μm distance) cells show the presence and absence of Mcd1-EGFP, respectively. Error bars represent the standard deviations from the mean values obtained from three independent experiments. Bar, 2 μm.

The failure of proper cohesin removal in post-anaphase I cin8Δ kip3Δ cells instigated us to examine if a similar defect prevails in mitosis. For the estimation of Mcd1 (mitotic cohesin) localization with respect to the distinct mitotic stages, cells were released synchronously from G₁ arrest and examined for the presence or absence of an Mcd1-GFP nuclear signal. With reference to the distances between the SPBs, we found no significant difference in Mcd1-EGFP staining between wild-type and cin8Δ kip3Δ cells. The cells with interpolar distances in the range of 1.2 to 2.2 μm (metaphase/preanaphase) were positive for Mcd1-GFP, while beyond that (postanaphase), no Mcd1 staining was visible (Fig. 7E). These results indicate that mitotic cohesin removal is not perturbed in cin8Δ kip3Δ cells.

The cin8Δ kip3Δ strain causes homolog nondisjunction and aberrant meiosis II.

As the defect in cohesin removal hinders homolog separation during meiosis I (65, 66), we analyzed homolog segregation in wild-type and cin8Δ kip3Δ binucleated cells harboring homozygous CenV-GFP (Fig. 8A). Such cells with proper homolog disjunction will exhibit an equal number of CenV-GFP foci in each nucleus (2:2; type I), whereas nondisjunction will result in an unequal distribution of GFP foci (1:0, 1:3, and 4:0; type II). We detected the type II phenotype in approximately 26% of cin8Δ kip3Δ cells, compared to 12% of wild-type cells. Notably, a unique third category (around 17%) (type III) was observed only in cin8Δ kip3Δ cells, where CenV-GFP dots were present in the middle of a stretched DAPI (4′,6-diamidino-2-phenylindole) signal, where due to incomplete segregation, instead of two equal masses, DAPI is stained as one single elongated mass. We believe that the type III phenotype was generated as the sustained cohesin perturbs chiasma resolution and impedes the disjunction of the homolog, since we observed a significant reduction in the distance between the two homologs in binucleated meiosis I cells (Fig. 8B). In support of this, the removal of chiasmata by the spo11Δ strain resulted in a reduction of the type III phenotype of cin8Δ kip3Δ spo11Δ cells (Fig. 8A). However, the spo11Δ strain caused an increased type II frequency, as the loss of homolog pairing is known to perturb homolog biorientation and disjunction (65, 67).

FIG 8.

Homologous chromosomes and nuclear separation are impeded in cin8Δ kip3Δ cells. Wild-type (SGY9002) and cin8Δ kip3Δ (SGY5338) cells harboring homozygously marked CenV-GFP and Spc42-mCherry were released into meiosis at 30°C and analyzed for the homolog (CenV) and nuclear segregation. (A) Segregation of the CenV-GFP homologs at the binucleated stage for wild-type (n = 185), cin8Δ kip3Δ (n = 167), and cin8Δ kip3Δ spo11Δ (SGY5444) (n = 97) cells. Representative images for the types of CenV homolog separation are shown. (B) 3D distances between the two CenV-GFP homologs plotted with respect to the interpolar (SPB-SPB) distances. The distances were measured using Imaris software (see Materials and Methods). (C and D) Nuclear separation in meiotic cells harboring 2 SPBs (C) and 4 SPBs (D) (for panel C, n = 70 wild-type cells and n = 95 cin8Δ kip3Δ cells; for panel D, n = 166 wild-type cells and n = 166 cin8Δ kip3Δ cells). “n” represents the total number of cells scored for the analysis. (E) Analysis of tubulin morphology in wild-type (SGY5001) (n = 103) and cin8Δ kip3Δ (SGY5089) (n = 91) meiosis II cells by tubulin immunostaining. Error bars represent the standard deviations from the mean values obtained from three independent experiments. A P value of ≥0.05 is considered nonsignificant (ns). Bars, 2 μm.

A blockage in Rec8 cleavage in the esp1-1 separase mutant hinders nuclear separation; however, following prolonged arrest, the cells embark on abrupt meiosis II (65). Since homolog nondisjunction was found to be impaired in cin8Δ kip3Δ binucleated cells, which include both anaphase I as well as metaphase II cells (Fig. 8A), we examined meiosis I and II nuclear segregations in cells harboring two SPBs (Fig. 8C) and four SPBs (Fig. 8D), respectively. Given that cohesin retention in the cin8Δ kip3Δ strain is not due to Pds1 stability, i.e., cell cycle arrest (Fig. 7A to D), we argued that these cells would progress through meiosis I in spite of having a physical barrier in nuclear separation. As expected, we observed a significant population of anaphase I cells of the cin8Δ kip3Δ strain with incomplete nuclear division, as evident from the “stretched” nuclear morphology (approximately 40%) (Fig. 8C).

We also observed a meager population (approximately 10%) of anaphase I cells with three connecting nuclear lobes (“crossed” morphology) only for the cin8Δ kip3Δ strain. This category of DAPI segregation resembles the one obtained for FEAR mutants resulting from the initiation of meiosis II on the meiosis I spindle (2). The population of cells in the stretched and crossed categories either evade meiosis II, forming dyads (Fig. 3B), or abruptly enter meiosis II, where they showed mostly asymmetric (26%) and no nuclear (mononucleates) (41%) separation with 4 SPBs (Fig. 8D). Similar phenotypes were observed in mam1Δ cells due to delayed nuclear division (68). Furthermore, due to the prolonged anaphase I and the subsequent abrupt initiation of meiosis II, there was a significant difference in the lengths of the two spindles in meiosis II in around 34% of cin8Δ kip3Δ cells (Fig. 8E). This phenotype is similar to the ones observed in meiosis II cells of the mam1Δ strain in budding yeast (68) and a recombination-defective rec8 mutant of Schizosaccharomyces pombe (69), where the common responsible factor is delayed nuclear separation.

Kip1 degradation is delayed in cin8Δ kip3Δ cells.

In cin8Δ kip3Δ cells, although delayed, spindle elongation occurs, and we believe that Kip1 executes this function in a protracted way. In mitosis, Kip1 is degraded during the onset of anaphase by Cdc20 (34). To investigate if Kip1 becomes more stable in the absence of Cin8 and Kip3, we compared the Kip1 levels between wild-type and cin8Δ kip3Δ cells during different stages of meiosis by immunoblotting (Fig. 9A and B). Given the difference in the paces of the cell cycle, the 10-h stage of wild type cells was considered equivalent to the 12-h stage of cin8Δ kip3Δ cells, as the percentages of tetranucleated cells observed were almost similar (approximately 82% in wild-type and 74% in cin8Δ kip3Δ cells) (Fig. 9C). As expected, Kip1 was found to be stable for a longer duration in cin8Δ kip3Δ than in wild-type cells (Fig. 9A and B). To further examine this, we monitored the localization of Kip1 in wild-type and cin8Δ kip3Δ cells undergoing meiosis with live-cell imaging. Stages were judged on the basis of the number of SPBs and the distance between two SPBs. In wild-type anaphase I cells, Kip1 was localized either along the spindle (42%) or near the poles (12%), while in 46% of cells, Kip1 was absent, suggesting that it is degraded toward the end of meiosis I. In contrast, Kip1 was absent in only 4% of anaphase I cin8Δ kip3Δ cells (Fig. 9D). In metaphase II, while almost 100% of wild-type cells showed a polar localization of Kip1, almost 45% of cin8Δ kip3Δ cells exhibited a single spindle-like localization spanning the 4 SPBs, suggesting that Kip1 degradation is deferred in the latter cells (Fig. 9E). However, overall, the Kip1 level was not altered, as determined by comparing Kip1-EGFP intensities between wild-type and cin8Δ kip3Δ cells at anaphase I and metaphase II (Fig. 9F and G). For intensity measurements within anaphase I cells, only spindle-localized Kip1-EGP intensities were compared between the wild type and the mutant. These results indicate that in the absence of Cin8 and Kip3, spindle elongation can still be possible, perhaps through positive regulation of Kip1 function. From the above-described localization and immunoblot studies, we propose that in the absence of a spindle localization of Cin8 and Kip3, Kip1 exhibits a greater distribution along the spindle, which may result in delayed degradation of this protein.

FIG 9.

Increased stability of Kip1 in cin8Δ kip3Δ cells during meiosis. (A) Wild-type (SGY5539) and cin8Δ kip3Δ (SGY5540) cells harboring Kip1-6HA were induced for synchronized meiosis and analyzed for the levels of Kip1-6HA at the indicated time points during meiotic progression. Tubulin was used as a loading control. (B) Densitometric analysis of Kip1-6HA bands obtained in panel A after normalization with the respective tubulin bands using ImageJ software. (C) Percentages of prophase I, metaphase I, anaphase I, metaphase II, and anaphase II/sporulated cells determined by tubulin immunostaining of wild-type and cin8Δ kip3Δ cells at the time points utilized for Kip1-6HA detection in panels A and B (n ≥ 90 cells for each time point). The arrow indicates a reference time point for comparison between wild-type and cin8Δ kip3Δ cells. (D and E) Localization of Kip1-EGFP in wild-type (SGY5051) and cin8Δ kip3Δ (SGY5561) cells during anaphase I (wild-type cells, n = 56; cin8Δ kip3Δ cells, n = 97) (D) and metaphase II (wild-type cells, n = 47; cin8Δ kip3Δ cells, n = 165) (E). The stage of a meiotic cell was determined by the number of and distance between the SPBs. “n” represents the number of cells analyzed for the assay. (F and G) Dot plots of Kip1-EGFP intensities in wild-type and cin8Δ kip3Δ cells during anaphase I (F) and metaphase II (G). Each signal intensity value was normalized to the background and Spc42-mCherry intensity values. Error bars represent the standard deviations from the mean values obtained from three independent experiments. A P value of ≥0.05 was considered nonsignificant (ns). Bars, 2 μm.

The tension generated by microtubule-mediated force drives efficient Rec8 removal.

From the above-described results, it is evident that in the absence of both Cin8 and Kip3, Rec8 is not efficiently removed from chromatin, and this condition perhaps leads to chromosome breakage during meiosis II. What could be the reason for Rec8 retention when Cin8 and Kip3 are not present? We argue that in cin8Δ kip3Δ cells, due to the absence of microtubule cross-linking and depolymerization activities, there is inadequate microtubule-based pulling force acting on the cohesin between the sisters during meiosis I and meiosis II. Given this, we hypothesize that the generation of tension on cohesin is perhaps a novel determinant for efficient Rec8 removal. If this is true, then the generation of microtubule force in cin8Δ kip3Δ cells can rescue Rec8 cleavage and, therefore, chromosome integrity and spore viability. To test this, we expressed in these cells a phosphodeficient allele of CIN8 (Cin8-3A) that is retained on the spindle and can generate force for an extended period or a phosphomimic allele of CIN8 (Cin8-3D) that fails to bind to the microtubule and create force and thus exhibits a diffuse nuclear localization (31). The spindle localization of Cin8-3A but the diffuse localization of Cin8-3D in anaphase I cells confirmed their modes of action (Fig. 10A). Remarkably, in the chromosome segregation assay with homozygous CenV-GFP, we observed a drop in the percentage of tetranucleates harboring >4 GFP dots, a readout of chromosome breakage, in cin8Δ kip3Δ Cin8-3A cells (15%) compared to cin8Δ kip3Δ cells (29%) (type IV) (Fig. 10B). There is no significant difference in the patterns of chromosome segregation observed among the cin8Δ, cin8Δ cin8-3A, and cin8Δ cin8-3D mutants, suggesting that CIN8 phosphomutants by themselves do not exhibit any additional defects (Fig. 10B). In accordance with this, the spore viability obtained for cin8Δ kip3Δ cells (approximately 16%) was ameliorated to a great extent upon the expression of Cin8-3A (approximately 48%) (Fig. 10C). These results suggest that the microtubule binding ability of Cin8-3A can partially mitigate the defects found in cin8Δ kip3Δ cells. The observed rescue effect is specific to the ability of Cin8-3A to bind to the microtubule, as cin8Δ kip3Δ cells expressing Cin8-3D showed phenotypes similar to those of cin8Δ kip3Δ cells alone (Fig. 10B and C). These results indicate that the tension generated by Cin8 and Kip3 collectively via the microtubule perhaps creates a signal for efficient cleavage and subsequent removal of Rec8.

FIG 10.

The absence of microtubule cross-linking activity of Cin8 is partly responsible for the defect observed in the cin8Δ kip3Δ mutant. (A) cin8Δ kip3Δ cin8-3A–13MYC (SGY5546) and cin8Δ kip3Δ cin8-3D–13MYC (SGY5547) cells expressing phosphodeficient or phosphomimic alleles of CIN8, respectively, were examined during meiosis for the localization of the corresponding mutant proteins with respect to the spindle. While Cin8-3A–13Myc was detected along the long spindles (white arrows) and at the poles (arrowheads), Cin8-3D–13Myc was observed as diffuse nuclear signals (red arrows). Bars, 5 μm. (B) Homozygous CenV-GFP segregation in tetranucleated cells of the wild-type (SGY5001), cin8Δ (SGY315), cin8Δ cin8-3A–13MYC (SGY5715), cin8Δ cin8-3D–13MYC (SGY5716), and cin8Δ kip3Δ (SGY5089) strains and the two strains used in panel A. More than 80 tetranucleates were counted for each strain. The frequency of tetranucleates with supernumerary GFP dots (type IV) was reduced in cells expressing Cin8-3A, which can bind and cross-link the microtubules. Bar, 2 μm. (C) Percentages of spore viability in the above-mentioned strains. More than 70 tetrads were dissected for each strain. Error bars represent the standard deviations from the mean values obtained from three independent experiments.

To further test whether tension is an additional factor required for cohesin removal in meiosis, we monitored Rec8 localization after mimicking the conditions of loss of tension by two distinct approaches. During meiosis I, the tension between the homologs and on the cohesin is generated as the bipolar pulling force by the microtubule is opposed by chiasmata formed between the homologs and the cohesion formed between the sister chromatids. We inhibited chiasma formation by deleting SPO11, examined Rec8 localization, and compared it with the above-described results (Fig. 6A to D and Fig. 11A). Nuclear Rec8 localization was observed in around 92% of spo11Δ anaphase I cells, which was far greater than that observed in cin8Δ kip3Δ (65%) or in wild-type (36%) cells (Fig. 6A to C and Fig. 11A), suggesting that loss of tension indeed resists efficient cohesin removal. However, as spo11Δ cells proceeded to meiosis II, the Rec8 staining pattern in metaphase II became similar to that of wild-type cells. This was expected since the spo11Δ strain can alleviate tension only during meiosis I. In another approach to investigate the role of tension in Rec8 removal, we depolymerized microtubules using benomyl (see Materials and Methods) in cells depleted for the spindle assembly checkpoint protein Mad2 using the CLB2 promoter so that the cells could proceed through meiosis (70). We treated the cells with benomyl after 5.5 h of meiotic release when most of the cells had passed the prophase I stage (Fig. 11B). In the absence of Mad2, benomyl-treated cells were able to go through meiosis I and meiosis II although not as efficiently as mock-treated cells (Fig. 11C). Due to the absence of microtubules, as SPB separation was improper, we were unable to distinguish between metaphase I and anaphase I cells, and therefore, only the cells with 4 SPBs were analyzed. We observed that a notable population (69%) of cells harbored robust nuclear Rec8 staining in the benomyl-treated culture but no or minimal centromeric Rec8 staining in the mock-treated culture (Fig. 11D). This suggests that the removal of microtubules by benomyl reduces tension and that this in turn perturbs Rec8 cleavage. Consequently, it is expected that benomyl-treated cells harboring homozygous CenV-GFP would cause chromosome breakage during meiosis II and show supernumerary GFP foci. Although DAPI segregation in the presence of a sublethal concentration of benomyl was not as efficient as that of the mock-treated culture, we observed ∼32% tetranucleated cells with supernumerary GFP foci in the presence of the drug, which was a meager 5% under unperturbed conditions (Fig. 11E). The two above-described investigations indicate that a reduction of tension can cause inefficient cohesin removal, and we suggest that this condition eventually leads to chromosome breakage, as observed in cin8Δ kip3Δ cells.

FIG 11.

Tension is indispensable for the timely removal of Rec8, and deletion of REC8 suppresses the formation of supernumerary centromeric foci in cin8Δ kip3Δ cells during meiosis. (A) Localization of Rec8-EGFP in spo11Δ (SGY5610) cells (n = 220) harboring Spc42-CFP in the anaphase I and metaphase II stages with respect to wild-type and cin8Δ kip3Δ cells, as shown in Fig. 6C and D. Representative images of spo11Δ cells are shown on the left, whereas those of wild-type and cin8Δ kip3Δ cells are shown in Fig. 6A and B, respectively. Based on the distribution of Rec8 staining, cells were categorized as having centromeric or nuclear Rec8, as shown by arrowheads or arrows, respectively, in the representative images. (B and C) Mad2 depletion relieves cell cycle arrest caused by microtubule disruption. Wild-type (SGY5557) and P_CLB2-MAD2 (SGY5628) cells harboring Rec8-EGFP and Spc42-CFP were released into synchronized meiosis. The progression of the cells through meiosis before or after the addition of benomyl was analyzed by tubulin immunostaining and DAPI staining. (B) Percentages of cells that had progressed beyond prophase I following 5.5 h of meiotic release into drug-free medium. (C) Each meiotic culture was either mock treated (dimethyl sulfoxide [DMSO]) or treated with benomyl, and percentages of tetranucleated/sporulated cells at the indicated time points were determined. (D) Localization of Rec8-EGFP in Mad2-depleted cells harboring P_CLB2-MAD2 (SGY5628) in the presence or absence of benomyl. Cells showing 4 SPBs marked by Spc42-CFP were scored for centromeric, nuclear, or no signals of Rec8. (E) P_CLB2-MAD2 cells harboring homozygous CenV-GFP (SGY3248) were analyzed for the segregation of CenV-GFP at the tetranucleated stage following microtubule depolymerization by benomyl. For panels D and E, approximately 120 to 150 cells were analyzed from benomyl-treated or untreated culture, and the drug or DMSO was added following 5.5 h of meiotic induction. (F) Analysis of CenV-GFP-marked homolog segregation in the tetranucleates of rec8Δ (SGY5667) (n = 142) and cin8Δ kip3Δ rec8Δ (SGY5670) (n = 74) cells along with the cin8Δ kip3Δ data shown in Fig. 4A. “n” represents the total number of tetranucleates scored for chromosome segregation. Error bars represent the standard deviations from the mean values obtained from three independent experiments. Bar, 2 μm.

If the retention of Rec8 is responsible for chromosome breakage in cin8Δ kip3Δ cells, then the removal of Rec8 in these cells should alleviate the defect. To examine this, we deleted REC8 in the wild-type and cin8Δ kip3Δ strains harboring homozygous CenV-GFP. Since meiosis is severely compromised in the absence of Rec8 (71), we observed a much smaller population of tetranucleates in rec8Δ or in cin8Δ kip3Δ rec8Δ cells. Due to the high rate of chromosome nondisjunction in the absence of cohesin, the percentage of tetranucleates with GFP dots in all four nuclei was negligible (approximately 1%) (type I) (Fig. 11F); instead, we observed a predominant population of tetranucleates with GFP dots in 2 nuclei in rec8Δ and cin8Δ kip3Δ rec8Δ cells (47% and 58%, respectively) (type III), while the remaining population contained GFP dots either in three of the four nuclei (31% and 17% of rec8Δ and cin8Δ kip3Δ rec8Δ cells, respectively) (type II) or in only one of the four nuclei (21% and 25% of rec8Δ and cin8Δ kip3Δ rec8Δ cells, respectively) (type IV). This gross chromosome missegregation was also evident from the asymmetric DAPI staining observed in the tetranucleated cells. However, as we expected, none of the triple mutant cells exhibited >4 CenV-GFP dots, indicating that defective cohesin removal in cin8Δ kip3Δ cells is indeed responsible for chromosome breakage, which is also depicted in our model (Fig. 12).

FIG 12.

A possible mechanism responsible for chromosome breakage in cin8Δ kip3Δ cells. The model shows meiotic chromosome segregation in wild-type (A) and cin8Δ kip3Δ (B) cells. (I) Due to the absence of Cin8 and Kip3, the transduction of pulling force on the chromosomes by only Kip1 through sliding of the antiparallel microtubules is less in the double mutant than in the wild type. This causes a lack of tension on the cohesin, perturbing their removal. (III) The persistence of cohesin resists chiasma resolution, which, along with weak kinetochore-microtubule attachment in the cin8Δ kip3Δ mutant, results in homolog nondisjunction. These cells, after a transient delay at anaphase I, either form dyads or enter into meiosis II, showing mononucleates with 4 SPBs, perhaps due to structural blockage in disjoining chromatids, which can subsequently produce tetranucleates with broken chromosomes. (IV and V) However, in some cells, homolog disjunction occurs but with persistent cohesin (IV), which results in chromosome breakage when the cells enter meiosis II (V).

DISCUSSION

Faithful chromosome segregation relies on the coordinated interaction between the microtubule and chromosomes. The molecular motors profoundly influence this interaction not only by ensuring the proper attachment of the kinetochores to microtubules but also by temporally and spatially regulating the microtubule spindle. Roles of motor proteins in the context of mitotic chromosome segregation have been described in several studies (9, 14, 43, 46). However, given the differences in the patterns of chromosome movement between mitosis and meiosis, including two-time chromosome segregation with concomitant assembly, extension, and disassembly of the spindle in meiosis, it is intriguing to investigate the functions of these motors in meiosis. In this work, we analyzed the functions of three microtubule plus-end-directed kinesin motors in meiosis.

Loss of Cin8 perturbs homolog pairing and homolog disjunction.

Analysis of the single-motor mutants revealed that the loss of Cin8 affects meiosis more than the loss of either Kip1 or Kip3 (Fig. 1). At early meiosis, Cin8 appears to promote homolog pairing and, consequently, homolog disjunction (Fig. 2C to E). In homolog pairing, it is required that each homolog locate each other, which requires the functions of the cytoskeleton and the motors that are believed to facilitate pairing by enhancing the search rate. The function of the dynein motors with the help of the nuclear envelope-spanning SUN/KASH proteins in homolog pairing has been demonstrated in Caenorhabditis elegans (72, 73). In S. pombe, the dynein motors drive the “horsetail nuclear movement” that facilitates homolog pairing (74). In S. cerevisiae, the rapid prophase movements (RPMs) of chromosomes in meiosis I are believed to occur via actin and nuclear envelope motor proteins, including Mps3-Ndj1-Csm4, through interactions with telomeres (75–77). While RPMs and telomere-led movements of the chromosomes promote homolog pairing, it is plausible that the nuclear kinesins may facilitate RPMs, or they may function in pairing independently. However, the former possibility is unlikely since no interaction among the nuclear envelope proteins and the kinesin motors has been demonstrated. With better microtubule cross-linking activity than the kinesin-8 motor (Kip3), the kinesin-5 motors (Cin8/Kip1) may have a greater role in the movement of one homolog than the other during the search for the pairing partner. The fact that we observed a more significant effect of Cin8 than of Kip1 on homolog pairing and, for that matter, on meiosis (Fig. 2C), despite both proteins belonging to the kinesin-5 family, is not surprising since it has been demonstrated that in mitosis, Cin8 plays a larger role than Kip1 in chromosome segregation (78). This is perhaps due to structural differences between these two proteins (23, 79).

Cin8 and Kip3 together are essential for timely exit from meiosis I and completion of meiosis II.

Our analysis revealed that cin8Δ kip3Δ cells share the phenotypes of the FEAR mutants, which include a delay in spindle elongation and disassembly and the generation of dyads (Fig. 1Bii and iii and Fig. 3B). We believe that this happens in FEAR mutants because Cin8 dephosphorylation by FEAR pathway-released Cdc14 is essential for maintaining Cin8 at the spindle (7). Due to the lack of Cdc14 in the FEAR mutants, the phosphorylated form of Cin8 mediated by Cdk1 is enriched, which dissociates Cin8 from the spindle (31). Therefore, the spindle phenotypes observed for the FEAR mutants of meiotic cells resemble those of cells devoid of both Cin8 and Kip3. Notably, the cin8Δ mutant alone does not show a FEAR-like phenotype, indicating that the Kip3 function is parallel to that of Cin8, at least at the spindle, and its function might be similarly modulated by the absence of Cdc14. Although Kip3 function has not been reported to be regulated by Cdc14, in a screen using a yeast proteomic library, Kip3 was identified as one of the Cdk1 substrates (80). Given that Cdc14 is known to undo most of the Cdk1-mediated phosphorylations and that in S. pombe, one of the kinesin-8 members, Klp-6, is a substrate of the Cdc14 homolog Clp1 (81), it is possible that Cdc14 regulates Kip3 function in S. cerevisiae. In addition, similar to the FEAR mutants, cin8Δ kip3Δ cells mostly showed reductional segregation in two spores of the dyads. However, some cells completed meiosis II and produced tetranucleates but with dire consequences, as discussed below.

Improper cohesin removal in cin8Δ kip3Δ cells causes chromosome breakage in meiosis.

The finding of >4 CenV-GFP foci in homozygously GFP-marked cin8Δ kip3Δ cells specifically in meiosis but not in mitosis was surprising (Fig. 3E and F; see also Fig. S2B at http://www.bio.iitb.ac.in/~santanu/wp-content/uploads/2020/05/Supplementary_file-final.pdf). Further analysis revealed that this happens due to chromosome breakage in cells that attempt to complete meiosis II (Fig. 4C). Unexpectedly, our investigations suggest that this breakage is due to the improper removal of cohesin from chromatin during both the metaphase I-to-anaphase I and metaphase II-to-anaphase II transitions (Fig. 6 and Fig. 7A to D). We believe that the anaphase I delay in cin8Δ kip3Δ cells, besides the lack of sliding of the antiparallel microtubules, is also due to the inefficient removal of cohesin from the arm regions.

Since in cin8Δ kip3Δ cells, we observed the uncoupling of Pds1 degradation from Rec8 removal (Fig. 7A to D), it is reasonable to propose that in meiosis, efficient Rec8 cleavage perhaps requires an additional factor besides the release of separase.

Microtubule based tension: a novel determinant to cleave Rec8- but not Mcd1-cohesin?

It is important to address why Rec8 removal is compromised in the absence of Cin8 and Kip3 together. Both Cin8 and Kip3 localize at the kinetochore, where Kip3 is a part of the core kinetochore and is involved in kinetochore-microtubule attachment (13). On the other hand, the lack of Cin8 and Kip3 together, but not individually, causes a reduced transient separation of the sister kinetochores compared to the wild type in preanaphase mitotic cells (13), and we noticed that the metaphase-to-anaphase transition is delayed (see Fig. S1B at the URL mentioned above). These results suggest that Cin8 and Kip3 together are involved in force generation on the chromosomes toward the opposite spindle poles, which is consistent with the fact that these motors have microtubule cross-linking (19, 20, 38, 82) and depolymerase (28, 37, 38) activities. Therefore, in cin8Δ kip3Δ meiotic cells, the homologs and, thus, the sisters are not under tension in meiosis I and meiosis II, respectively. We propose a model (Fig. 12) where the efficient cleavage of Rec8 in both meioses I and II requires the homologs and the sisters, respectively, to be under tension. In support of this, in a phosphodeficient Cin8 mutant that remains bound to the spindle for a longer duration and can generate force, Rec8 removal is supposedly better, and hence, we observed less chromosome breakage and improved spore viability, the opposite of which was found in the case of a phosphomimic mutant that fails to bind to the spindle and generate force (Fig. 10B and C). To reconfirm our tension model of Rec8 cleavage, we created tensionless conditions by removing either chiasmata (SPO11) or microtubules in a Mad2-depleted strain and observed defective Rec8 removal under both conditions (Fig. 11A and D). Importantly, we failed to observe any perturbation of Mcd1 removal in cin8Δ kip3Δ cells (Fig. 7E), and we believe that this is why the cells perform better in mitosis (see Fig. S1D and E at the URL mentioned above). To address how tension might drive Rec8 but not Mcd1 cleavage, we reason that tension may by some means promote the phosphorylation of Rec8, and previous studies have shown that phosphorylation of Rec8, but not Mcd1, is indispensable for cohesin cleavage (4, 83, 84). Tension can influence the maintenance of certain proteins on chromatin responsible for Rec8 phosphorylation, or as a direct effect on the cohesin, it may expose the Rec8 sites for phosphorylation. Alternatively, tension may facilitate phosphorylated Rec8 amenable to cleavage by separase. It is possible that in cin8Δ kip3Δ cells, the spindle assembly checkpoint becomes activated because of a loss of tension and faulty kinetochore-microtubule attachment, which can keep APC inactivated and resist cohesin cleavage. However, we believe that this is unlikely as we observed that Pds1 degradation occurred in the double mutant at a normal pace of the cell cycle (Fig. 7B and D). To explain why a tension-based mechanism has evolved to sensitize cohesin removal in meiosis, it can be argued that in this cell cycle, unlike in mitosis, chiasmata are formed, and the removal of arm cohesin is required for their resolution. During resolution of chiasmata, the “terminalization” of the crossover point that occurs due to pulling of the homologs might subject arm cohesin to tension, which perhaps signals their removal. However, how the prolonged retention of cohesin with reduced tension acting on the chromosomes (due to the absence of Cin8 and Kip3) can eventually lead to chromosome breakage is not clear from our study. We observed that in cin8Δ kip3Δ cells, spindle disassembly is delayed (stretched category) (Fig. 8C), and Kip1 activity is protracted (Fig. 9). Additionally, the loss of these proteins can potentially cause abnormally extended kinetochores-microtubules since they also possess microtubule depolymerase activity (28, 37, 38). It is plausible that when these conditions together prevail over an extended period of time spanning two rounds of spindle assembly/disassembly and chromosome movement in meiosis, an imbalance of force is generated on the chromosomes, causing them to break. Although tension is known to cause a reorientation of the unipolar-attached chromosome and resumption of the cell cycle from arrest (85), here, we report for the first time the requirement of tension for the efficient removal of cohesin in meiosis and the importance of kinesin-5 and kinesin-8 motors in promoting this event and, thus, maintaining chromosome integrity. Given that meiosis and the functions of kinesins are conserved across eukaryotes, it would be tempting to investigate if an attenuation of motor functions could be one of the reasons for the generation of aneuploid gametes that occurs at an alarming rate during human gametogenesis.

MATERIALS AND METHODS

Yeast strains and media.

All the strains used in this study were of the SK1 background. A list of strains and plasmids with their genotypes can be found in Table S1 at http://www.bio.iitb.ac.in/~santanu/wp-content/uploads/2020/05/Supplementary_file-final.pdf. The plasmids utilized for C-terminal protein tagging and deletion of a gene were obtained from Euroscarf and were PCR based (86). Transformation of the cells with the PCR cassettes was performed as mentioned previously (87). In the case of selection of the cells on dropout medium along with the antibiotic G418, the medium was used as described previously (88), where instead of ammonium sulfate, monosodium glutamate was used to restore the sensitivity to G418. For metaphase I and prophase I arrest, P_CLB2 and P_GAL1 constructs were used to shuffle the endogenous promoters of CDC20 and NDT80, respectively, as described previously (4, 89). For chromosome segregation assays, chromosome V and chromosome III were marked with GFP by integrating repeats of tet operators at 1.4 kb and lac operators at 22 kb away from the centromeres, respectively, in cells expressing TetR-GFP and LacI-GFP, respectively (90, 91).

Fluorescence microscopy.

For live-cell imaging, 1 ml of the culture at an optical density at 600 nm (OD₆₀₀) of 1 was fixed with formaldehyde (final concentration, 5%) for 5 to 10 min. The pellet was washed two times with 0.1 M phosphate buffer (pH 7.5). For DAPI staining, after fixation of the sample with formaldehyde, the pellet was washed once with 50% ethanol and then resuspended in DAPI at a final concentration of 1 μg/ml. The image was acquired with z-stacking (spacing of 0.5 μm) using a Zeiss AxioObserver Z1 inverted microscope (63×, 1.4-numerical-aperture [NA] objective). Processing and merging of images were done using AxioVs40 V 4.8.2.0 software. The exposure time was set according to the fluorescence signal and was kept constant among the samples used for comparison (mainly, it was 1.5 s for EGFP, CFP, and mCherry fluorophore excitation). In order to avoid bleed-through of the intense Spc42-CFP signal in the GFP channel, a Zeiss confocal laser scanning microscope (LSM 780) equipped with a 32-array GaAsP detector was used. Images were acquired using Zeiss Zen 2012 software.

Image analysis.

Images were generated by merging the planes projecting maximum intensity and further analyzed. The quantification of the fluorescence intensity of the images, acquired using a Zeiss AxioObserver Z1 microscope, was performed using ImageJ software (92). A region of interest covering the fluorescence signal was defined, and the integrated intensity of that region was estimated, following background reduction, by averaging the integrated intensities of three random nonfluorescent areas multiplied by the area of the fluorescent signal region. Estimation of the number of Rad52 or Rec8 foci per chromosome spread was performed using an automatic spot detection algorithm (Imaris3D reconstitution software), keeping the threshold limit constant for all the images.

Growth conditions and meiotic induction.

Before meiotic induction, the cells were patched onto YPG (1% yeast extract, 2% peptone, 2% glycerol) to restrain the growth of petite colonies and then transferred to presporulation medium overnight. This was followed by meiotic induction in sporulation medium (0.02% raffinose, 1% potassium acetate) as described previously (93, 94).

In order to prevent the loss of the centromeric plasmid containing a mutated open reading frame (ORF) of CIN8, in the presporulation medium (PSP2) used for the meiosis synchronization, instead of yeast extract-potassium acetate (YPA) medium, selective medium (synthetic complete media without uracil) supplemented with 0.1% yeast extract was used (95, 96).

For mitotic synchronization, cells were arrested at G₁ using α-factor at a concentration of 5 μg/ml in cells at an OD₆₀₀ of 0.3 (97, 98). After 3 h, when >90% of cells exhibited shmoo formation, the cells were washed and released into fresh YPD medium.

For enhancing the expression of LacI-GFP that is under the control of the HIS3 promoter, 3-aminotriazole was added at a final concentration of 20 mM to sporulation medium.

Unlike mitosis, disruption of microtubules before or during meiotic S phase causes cells to arrest at G₁ or G₂ phase, respectively (99). Therefore, for microtubule depolymerization in meiosis, the cells were treated with benomyl at a concentration of 60 μg/ml after 5.5 h of meiotic induction, when most of the cells passed through S phase. Stages of the cell cycle and microtubule morphology before and after drug treatment were determined by tubulin immunofluorescence. In order to avoid spindle checkpoint-mediated arrest in the absence of microtubules, Mad2 was depleted in meiosis using the CLB2 promoter (70).

Comet assay.

A comet assay was performed as described previously (100). After meiotic induction in SPM for 8 h, 1 ml of the sporulating culture at a concentration of 10⁷ cells per ml was harvested. Cells were then resuspended in buffer (1 M sorbitol, 25 mM KH₂PO₄, 50 mM β-mercaptoethanol [β-ME]) containing Zymolyase 20T (20 mg/ml; MP Biomedicals). The pH of the buffer was adjusted to 6.5 using NaOH. The cells were then incubated for 1/2 h for making spheroplasts. Spheroplasted cells were then mixed with 1.5% low-melting-point (LMP) agarose and spread immediately onto a glass slide precoated with 0.5% normal-melting-point (NMP) agarose. Slides were placed on ice for the agarose to solidify, in which the embedded cells formed cavities in the gel. Subsequently, the slides were submerged in lysing buffer (30 mM NaOH, 1 M NaCl, 0.05% sodium dodecyl sulfate, 50 mM EDTA, 10 mM Tris-HCl [pH 10]) for 20 min at 4°C. Following lysis of the spheroplasts, the cavities formed by the spheroplasted cells contained only high-molecular-weight DNA, while the other biomolecules diffused out. The slides were then placed into electrophoresis buffer (30 mM NaOH, 10 mM EDTA, 10 mM Tris-HCl [pH 10]) at 4°C for 20 min for the unwinding of the DNA, which was followed by electrophoresis for 20 min at 0.7 V/cm. Upon application of an electric current of 300 mA at 24 V, the fragmented DNA, named the “tail,” moved toward the anode (positive electrode), while the compact mass of DNA remained in the cavity, giving a “comet”-like appearance on the gel. For each comet, the head is the mass of supercoiled DNA. Only if there is a DNA break is supercoiling released. The loose ends extend toward the anode (+) during electrophoresis, giving a tail-like appearance against the immobile compact mass of DNA. Following this, the slides were incubated in neutralization buffer (10 mM Tris-HCl [pH 7.4]) at room temperature for 10 min. The slides were incubated in 76% and 96% ethanol for 10 min each at room temperature. The slides were then incubated with a solution containing ethidium bromide (10 μg/ml) for 5 min and observed using an epifluorescence microscope (excitation filter, 546 nm; emission filter, 575 nm). Wild-type cells treated with 10 mM H₂O₂ were used as positive controls. We used a method to calculate the percentage of DNA in the tail as described previously by Braafladt et al. (56), using the formula % DNA in tail = 100 × I_t/(I_h + I_t), where I_t is the total tail intensity and I_h is the total head intensity.

Immunostaining.

Immunostaining was performed as described previously (94). Cells from the meiotic culture were harvested and fixed with 5% formaldehyde. Spheroplasts were made using Zymolyase and placed onto a polylysine-coated slide. The spheroplasts were permeabilized by Triton X-100 or methanol-acetone and then incubated with primary followed by secondary antibodies. DAPI (4′,6-diamidino-2-phenylindole) at a concentration of 1 μg/ml in 0.1 M phosphate buffer was used to stain the DNA. Primary antibodies, including rat antitubulin (catalog number MCA78G; Serotec) and mouse anti-Myc (catalog number 11667149001; Roche), were used at dilutions of 1:5,000 and 1:200, respectively. The secondary antibodies used, obtained from Jackson, were tetramethyl rhodamine isothiocyanate (TRITC)-labeled goat anti-rat (catalog number 115-485-166), Alexa Fluor 488-labeled goat anti-rat (catalog number 112-545-167), and TRITC-labeled goat anti-mouse (catalog number 115-025-166) antibodies at a dilution of 1:200.

Chromosome spread.

The protocol for chromosome spread formation was performed as described previously (94, 101). Two milliliters of a meiotic culture was spheroplasted using Zymolyase 20T (10 mg/ml) for 1 h with 1.42 M β-ME. The reaction was stopped by the addition of 200 μl of stop solution (0.1 M morpholineethanesulfonic acid [MES], 1 mM EDTA, 0.5 mM MgCl₂, 1 M sorbitol [pH 6.4]). Spheroplasted cells were fixed on acid-washed slides with a paraformaldehyde solution (4% paraformaldehyde and 3.4% sucrose with 2 drops of NaOH to dissolve the paraformaldehyde), followed by the addition of 1% Lipsol to burst the cells. The slides were kept drying overnight at room temperature after homogenously smearing the spheroplasts on the slide. The next day, the slides were washed with 2 ml of 0.4% Photoflow-200 (Kodak), followed by washing in phosphate-buffered saline (PBS) for 10 min. Before the addition of primary antibodies, 100 μl of blocking solution (5% skim milk) was added to the slide for 30 min. Primary antibodies were diluted in PBS supplemented with 0.1% BSA (bovine serum albumin). Primary antibodies used were rabbit anti-Zip1 (catalog number SC 33733; Santa Cruz Biotechnology) (1:100), mouse antihemagglutinin (anti-HA) (catalog number MMS-101P; Covance) (1:200), and mouse anti-GFP (catalog number 11814460001; Roche) (1:200). The slides were coated with 100 μl of primary antibody for 1 h, followed by washing with PBS three times with a 5-min incubation each time. A similar treatment with secondary antibody was performed. Secondary antibodies from Jackson, TRITC-labeled goat anti-rat and Alexa Fluor 488-labeled goat anti-mouse (catalog number 115-485-166) antibodies, were used at a dilution of 1:200. Chromatin was stained using DAPI.

Immunoblotting and its quantification.

Whole-cell proteins were extracted by NaOH treatment as described previously (102), with some modifications. Cells from 10 ml of the culture at an OD₆₀₀ of 1 were pelleted down and treated with 0.1 N NaOH for 30 min. After alkaline treatment, pelleted cells were resuspended in electrophoresis sample buffer (2% SDS, 10% glycerol, 80 mM Tris [pH 6.8], 2% bromophenol blue, 100 mM dithiothreitol) (104) and boiled for 5 min at 100°C. The supernatant obtained after centrifugation was used for immunoblotting. Primary rabbit anti-Myc antibody (catalog number ab9106; Abcam) was used at a dilution of 1:5,000 in a 1:20 mixture of Tris-buffered saline plus Tween (TBST)–5% skim milk. Horseradish peroxidase (HRP)-conjugated secondary antibodies (Jackson) used for detection were goat anti-mouse (catalog number 115-035-166) (1:5,000), goat anti-rabbit (catalog number 111-035-003) (1:10,000), and goat anti-rat (catalog number 112-035-167) (1:10,000) antibodies. Blots were developed using ECL reagents (catalog number 170-5060; Bio-Rad Laboratories). The intensities of the bands at different time points were quantified using ImageJ software. The ratio of the protein bands to the loading control band was used for comparison between the wild-type and mutant strains.

RNA extraction and cDNA synthesis.

Total RNA was extracted from the meiotic cell culture at an OD₆₀₀ of 1.5 to 2 (∼1.5 × 10⁷ cells) using TRIzol reagent and a PureLink RNA minikit (Ambion Life Technologies). The cell pellet was resuspended in TRIzol, and cell lysis was done using diethyl pyrocarbonate (DEPC)-treated sterile 0.5-mm glass beads. The rest of the protocol was performed according to the instructions provided by the supplier (PureLink RNA minikit). RNA was eluted in DEPC (Sigma-Aldrich Chemicals Pvt. Ltd.)-treated sterile water. DNase treatment was done at 37°C for 10 min using RNase-free DNase I (Thermo Fisher Scientific) in the buffer supplied with the enzyme. The DNase was inactivated by heating at 65°C for 10 min. To ensure the absence of DNA impurity, endpoint PCR was performed with the same primers as the ones used for RT-qPCR.

cDNA synthesis was done using an iScript cDNA synthesis kit (Bio-Rad Laboratories Inc., USA) as instructed by the supplier.

RT-qPCR and data analysis.

All the quantitative PCR (qPCR) experiments were performed using the CFX96 Touch real-time PCR detection system (Bio-Rad Laboratories Inc., USA). We used two reference genes, TDH3 and CDC19, as internal controls to normalize the expression level of the gene of interest, REC8. The fold change in mRNA expression was calculated using the “ΔΔC_T” method (103). The change in threshold cycle (ΔC_T) value was determined by normalizing the Rec8 value to the values for both reference genes for the individual strains. For ΔΔC_T calculation, ΔC_T values of the test samples were further normalized to the ΔC_T value of the wild type. Finally, the fold change was obtained by using the ΔΔC_T value determined by using the equation $2^{-\Delta \Delta C_T}$. The primers and conditions utilized for setting up RT-qPCR are shown in Tables S2 and S3, respectively, at http://www.bio.iitb.ac.in/~santanu/wp-content/uploads/2020/05/Supplementary_file-final.pdf.