Evidence of 14-3-3 proteins contributing to kinetochore integrity and chromosome congression during mitosis Free

The 14-3-3 proteins are a family of highly conserved low-molecular-mass acidic proteins (Aitken et al., 1992; Martens et al., 1992) known for their myriad functions in eukaryotic cells (Kumar, 2017). They act by recognizing cellular proteins via a conserved phosphorylated motif (Muslin et al., 1996) and/or an amphipathic groove (Masters et al., 1999) to alter the activity, stability, subcellular localization and ability to physically interact with other proteins of the target protein (Kumar, 2017; Obsilova and Obsil, 2022). Among eukaryotes, the number of structurally conserved 14-3-3 isoforms varies with species; the ectopic expression of such isoforms can provide inter-species functional complementation (Van Heusden et al., 1995; Kumar, 2017). Despite such functional complementation, there are non-shared (unique) functions for individual isoforms (Wilkert et al., 2005; Herod et al., 2022).

In budding yeast, there are two 14-3-3 isoforms, namely Bmh1 and Bmh2 (collectively Bmh), characterized as having high sequence similarity with the mammalian isoform 14-3-3ε (Gelperin et al., 1995; Van Heusden et al., 1995). Bmh1 and Bmh2 exist as homo- and hetero-dimers, of which heterodimers are more prevalent within the cell (Chaudhri et al., 2003). Deletion mutants of either isoform show no impact on cell growth, but the double deletion is lethal in most genetic backgrounds of budding yeast (Van Heusden et al., 1995); however, in certain genetic backgrounds such as SK1 and ∑1278, the double mutants are viable with severe growth defects (Roberts et al., 1997; Slubowski et al., 2014). The viable genetic background strains might have evolved alternative pathways to alleviate the double deletion defects. For instance, overexpression of Ras/cAMP-dependent protein kinase (Tpk1) partially rescued viability loss caused by bmh double deletions (Gelperin et al., 1995) which might explain the viability of the double deletion in ∑1278 strain background where Ras/cAMP signaling is hyperactive (Stanhill et al., 1999). Nevertheless, comparing the functions of Bmh proteins in different genetic background strains might help in understanding their non-shared individual and cumulative contributions in budding yeast; therefore, we assessed the cumulative functions of Bmh proteins in the SK1 background strains, where a bmh double mutant is viable, and non-shared functions in the W303 background strains, where a bmh double mutant is lethal.

Consistent with their diverse functions across the cell cycle, the Bmh proteins are localized to almost all the sub-cellular locations throughout the mitotic and meiotic cell cycle stages (Slubowski et al., 2014). Their functions during S phase in DNA replication and damage repair have been demonstrated (Lottersberger et al., 2003). The M phase functions argue for their roles in chromosome segregation. For instance, Bmh proteins are involved in spindle assembly- and spindle position checkpoint-mediated cell cycle arrest, and hence the mutants are sensitive to spindle damage and spindle misalignment, respectively (Grandin and Charbonneau, 2008; Caydasi et al., 2014). Previously, we observed that there is a change in the overall cellular protein concentration ratio of Bmh1 and Bmh2 between mitotic and meiotic cells (Kumar et al., 2014; Kumar, 2018), suggesting their involvement in distinct chromosome segregation events. Notably, there are several lines of evidence depicting that Bmh isoforms have non-shared functions in chromosome segregation. For instance, a role for Bmh1, but not Bmh2, has been shown in silencing of the spindle assembly checkpoint (SAC), where it facilitates centromeric localization of an intermediate filament protein, Fin1 (Bokros et al., 2016; 2021). Bmh1 acts as a component of spindle position checkpoint (SPOC), which monitors proper orientation of the spindle along the mother–bud axis, ensuring successful disjunction of duplicated spindle pole bodies (SPBs) (Caydasi et al., 2014). Recently, Bmh1 has been shown to contribute to chromosome compaction through centromeric recruitment of the histone deacetylase Hst2 (Jain et al., 2021).

Given the role of Bmh proteins in chromosome segregation through microtubule spindle and chromosome morphogenesis, we hypothesize that these proteins might also influence kinetochore function due to following observations. First, the anti-microtubule drug sensitivity of the bmh mutants (Grandin and Charbonneau, 2008) is also shared by the kinetochore protein mutants (Sanyal et al., 1998; Poddar et al., 1999; Ghosh et al., 2001). Second, centromere-localized Aurora B kinase (Ipl1) phosphorylates histone H3 at serine 10, which is recognized by Bmh1, which in turn recruits Hst2 at the centromeres to remove H4K16 acetylation required to initiate chromosome compaction from the centromeres in cis (Jain et al., 2021). Third, the kinetochore protein Fin1 (Akiyoshi et al., 2009b) has been shown to have physical interaction with Bmh proteins in vitro (Bokros et al., 2016). Forth, computational analyses have revealed that several kinetochore proteins harbor Bmh interaction motifs and finally, in a high-throughput study the kinetochore protein Iml3 found to co-purify with Bmh1 (Kakiuchi et al., 2007).

To examine the hypothesis the Bmh proteins have a role in kinetochore function, we investigated the functions of the Bmh proteins and observed that they promote high-fidelity chromosome segregation perhaps by contributing to the maintenance of a kinetochore ensemble facilitating stable kinetochore–microtubule (KT–MT) dynamics. In addition, we provide evidence for a role of Bmh proteins in synchronous chromosome congression during metaphase. Thus, this work reports additional functions of Bmh proteins related to faithful chromosome segregation.

Budding yeast cells with mutations in proteins involved in KT–MT-related functions of chromosome segregation show hypersensitivity to microtubule-destabilizing drugs (Guarente, 1993). To elucidate the contribution of Bmh proteins in similar functions, we compared benomyl sensitivity of the bmh single and double mutants to wild-type and the non-essential kinetochore mutants (ctf19Δ and iml3Δ) (Fig. 1A). With increasing concentrations of benomyl, bmh2Δ and bmh1Δ bmh2Δ mutants showed growth defects that were similar to those of kinetochore mutants, as reported previously (Hyland et al., 1999; Ghosh et al., 2001), whereas the bmh1Δ mutant showed hardly any defect (Fig. 1A). We also tested the benomyl sensitivity in conditional bmh double mutants where BMH2 was deleted, and Bmh1 was degraded using the AID degron system (Nishimura et al., 2009) in W303 strains, where double deletion is lethal (Van Heusden et al., 1995). We compared the bmh single and conditional double mutants with the wild-type and with the kinetochore mutant (ctf19Δ). As expected, ctf19Δ, bmh2Δ and conditional double mutants displayed benomyl hypersensitivity (Fig. S1A), consistent with observations in SK1 strains (Fig. 1A), suggesting that the bmh mutant phenotypes are likely to be similar across different yeast genetic backgrounds. Noticeably, the severity of benomyl sensitivity varied among different genetic backgrounds of budding yeast (compare Fig. 1A with Fig. S1A). The hypersensitivity of bmh single mutants to microtubule-destabilizing drugs has also been observed elsewhere (Grandin and Charbonneau, 2008). However, the bmh1Δ bmh2Δ mutant showed a synergistic growth defect compared to the bmh single mutants (Fig. 1A) indicating possible non-shared functions of Bmh1 and Bmh2 with respect to chromosome segregation. In coherence with our observation, non-redundant functions of 14-3-3 isoforms were reported earlier in higher eukaryotes (Wilkert et al., 2005).

Given the benomyl sensitivity (Fig. 1A) and the slow growth of the bmh mutants, we wished to know at what stage of the cell cycle the defect is occurring. The budding index revealed that a high percentage of large-budded cells accumulated in the bmh1Δ bmh2Δ mutant (Fig. 1B) indicating cells are being delayed at G2/M and/or at anaphase. Similar accumulations of large-budded cells were observed in W303 background strains (Fig. S1B). To pinpoint the cell cycle stage of the delay, we analysed the segregation pattern of the DNA mass (DAPI) and the SPBs (with Spc42–EGFP) in the large-budded cells (Fig. 1C). We found that whereas the wild-type and the single mutants showed normal segregation (equally segregated DAPI and SPB signal in mother and daughter buds), the bmh1Δ bmh2Δ mutant largely (∼60%) showed defective segregation (an undivided single DAPI mass with both SPBs in the mother bud). Notably, in the cells showing defective segregation, the presence of DAPI and the spindle well away from the bud neck (double-headed arrow), despite there being a large bud, indicates that the movement of the nucleus to the bud neck and subsequent separation of the SPBs are delayed in the bmh1Δ bmh2Δ mutant. In order to visualize the dynamics of kinetochore and SPB separation, we fluorescently labelled kinetochores (Ndc80–CFP) and SPBs (Spc42–GFP) in the wild-type and bmh1Δ bmh2Δ mutant, and observed the cell cycle by time-lapse imaging of the live cells (Fig. 1D). Considering that the cells are in similar cell cycle stage at the start point (t=0 mins) (see Materials and Methods), a delay in the successful duplication followed by the bi-lobed metaphase configuration of kinetochores and SPBs (marked by arrowhead) was observed in bmh1Δ bmh2Δ mutant (t=60 mins) compared to the wild-type (t=20 mins). The delay in forming the bi-lobed configuration in bmh1Δ bmh2Δ mutant might be because of previously reported G1/S transition delay (Lottersberger et al., 2006). After forming the bi-lobed configuration, the wild-type cells successfully segregated their kinetochores and SPBs between mother and daughter buds within the range of normal doubling time (t=80–100 mins); in contrast, the bmh1Δ bmh2Δ mutant displayed a substantial delay in further segregation where the SPBs remained within the mother bud even after 180 min, indicating cells were either delayed at the metaphase to anaphase transition or, even after anaphase was triggered, that SPBs were not able to separate due to physical constraints. Interestingly, compared to the wild-type, reduced kinetochore (Ndc80–CFP) signal intensity was consistently observed in the bmh1Δ bmh2Δ mutant after it achieved a bi-lobed configuration (t=60 mins), suggesting potential kinetochore perturbation. We conclude that loss of both the Bmh proteins causes a high percentage of chromosome missegregation, perhaps due to defects in KT–MT-related functions. However, negligible growth defects (Fig. 1B,C) in bmh single mutants imply functional complementation between the Bmh proteins despite their independent contribution to chromosome segregation.

Given that the bmh1Δ bmh2Δ mutant showed benomyl sensitivity (Fig. 1A), reduced kinetochore (Ndc80–CFP) signal intensity after bi-lobed configuration (Fig. 1D), and a substantial delay in anaphase separation of the chromosomes (Fig. 1D), we hypothesized that the KT–MT attachment dynamics could be perturbed in these cells. Previous studies have reported that non-essential kinetochore proteins contribute to kinetochore organization and thereby promote high-fidelity chromosome segregation (reviewed in Mehta et al., 2022). Therefore, we wished to examine kinetochore organization in the absence of Bmh proteins using a chromatin spread, expecting altered localization of kinetochore proteins at centromeres. As we observed an accumulation of bmh1Δ bmh2Δ cells with undivided DAPI signals (Fig. 1C), we compared the localization of kinetochore proteins in the cells with an undivided DAPI mass. On the chromatin spreads made from asynchronously grown wild-type and bmh1Δ bmh2Δ cells, we chose to monitor the localization of the epitope-tagged outer kinetochore protein Ndc80–HA, which has been used previously (Janke et al., 2001). Based on the Ndc80–HA signal within undivided DAPI mass, we categorized the spread patterns as follows: ‘clustered kinetochores’, ‘declustered kinetochores’, and ‘weak-signal’ representing single or bi-lobed tight-knit foci, multiple foci, and multiple weak-intensity foci, respectively (Fig. 2A). In comparison with wild-type, bmh1Δ bmh2Δ mutant displayed a significant increase in the percentages of spreads harbouring declustered kinetochores or weak signal (∼57%) with nearly a cumulative equivalent percentage reduced in the clustered kinetochores category (Fig. 2B). The observed increase in the percentage of spreads with weakened signals and declustered kinetochores implies that Bmh proteins are crucial for the stability and clustering of kinetochores. We further biochemically assessed Ndc80–HA localization at the centromeres by performing a chromatin immunoprecipitation (ChIP) assay using anti-HA antibodies in asynchronously grown wild-type and bmh1Δ bmh2Δ cells. In accordance with the chromatin spread results, a reduced binding of Ndc80–HA was observed at the centromeres in bmh1Δ bmh2Δ mutant compared to the wild-type (Fig. 2C), indicating that Bmh proteins contribute to kinetochore stability. To explore their role in kinetochore clustering, we fluorescently labelled kinetochores (Ndc80–CFP) and SPBs (Spc42–EGFP) and performed live-cell imaging to compare the kinetochore distribution between wild-type and bmh1Δ bmh2Δ mutant during metaphase (cells with an SPB–SPB distance of 2.0–4.0 µm). Based on the Ndc80–CFP signal, we categorized the metaphase cells into ‘clustered kinetochores’, showing tight-knit bi-lobed foci between two SPBs, and ‘declustered kinetochores’, showing multiple foci linearly distributed between two SPBs (Fig. 2D, left). We observed a substantial increase in the percentage of cells with declustered kinetochores in bmh1Δ bmh2Δ mutant (>70%) as compared to the wild-type (∼20%) (Fig. 2D, right). For better visualization of kinetochore (Ndc80–CFP) distribution, line scans were undertaken between SPBs, which clearly depicted the deterioration of bimodal distribution of the kinetochore clusters in the double mutant (Fig. 2E, bottom) compared to the wild-type (Fig. 2E, top). The observed defect in kinetochore protein localization and in its clustering is not specific to Ndc80, as a similar pattern was observed for another kinetochore protein, Mtw1 in bmh1Δ bmh2Δ mutant (Fig. S2A–E).

As a G1/S transition delay has been reported in temperature-sensitive bmh mutants, where cells remained unbudded for up to 3 h compared to wild-type (Lottersberger et al., 2006), there is a possibility that the observed Ndc80–HA reduction at the centromeres (Fig. 2C) could be due to accumulation of unbudded cells undergoing kinetochore reassembly. To examine this, we synchronized the wild-type and bmh1Δ bmh2Δ mutant cells at G1 stage (using α factor) and released them into fresh medium, observing only a 20–30 min delay in bud emergence in the mutant. Despite this delay, the mutant progressed to produce a peak of large-budded cells at 90 mins and unlike wild-type, continued with that level of large-bud population till at least 150 min (Fig. S3A). Therefore, in an asynchronous population, G1/S (unbudded or small-budded) cells are expected to be rather fewer in number in the mutant than in the wild-type culture. This coincides with the budding index measurements from the asynchronous cultures (Fig. 1B). Hence, the reduction in Ndc80–HA in the mutant is likely not due to a difference in the pace of the cell cycle compared with the wild-type. Further, to confirm that the kinetochore protein reduction in bmh1Δ bmh2Δ mutants is not due to cell cycle differences, we have quantified and compared the fluorescence intensity of two kinetochore proteins (Ndc80–CFP and Mtw1–EGFP) specifically from metaphase cells of wild-type and bmh1Δ bmh2Δ mutants. As expected, a significant reduction in the signal intensity of both the kinetochore proteins was observed in bmh1Δ bmh2Δ mutants (Fig. S3B,C).

Alternatively, the kinetochore declustering might happen due to differences in the overall expression level of kinetochore proteins and/or defective spindle formation in the absence of Bmh proteins. To investigate expression differences, we estimated and compared protein levels of Ndc80 and Mtw1 in the wild-type and bmh1Δ bmh2Δ mutants by western blotting and found no significant difference (Fig. S4A,B). To investigate spindle defects, we examined spindle morphology and kinetochore (Mtw1) localization in both wild-type and bmh1Δ bmh2Δ mutants. Irrespective of kinetochore conformation (clustered or declustered), spindle morphology showed no discernible difference (Fig. S4C,D). In support of our observation, declustered kinetochores with unperturbed spindle morphology has been previously reported in kinesin motor mutants (Tytell and Sorger, 2006). In conclusion, our findings suggest that in the absence of both the Bmh proteins, kinetochore integrity is compromised, which perhaps alters the dynamics of KT–MT attachment causing kinetochores to be irregularly distributed (declustered) between the two SPBs.

The formation of bi-lobed clusters of sister kinetochores requires synchronized poleward movement of the chromosomes, termed chromosome congression, following their attachments to the kinetochore microtubules (kMTs) (Xiangwei et al., 2000; Pearson et al., 2004). Defective chromosome congression can be identified through the following phenotypes. First, there is hyperstretching of the pericentromeric chromatin due to deregulated pulling and pushing forces resulting from alteration of the plus- and/or minus-end growth–shrinkage dynamics of the kMTs during the metaphase–anaphase transition (Thrower and Bloom, 2001; Tytell and Sorger, 2006; Wargacki et al., 2010), and, second, there is an accumulation of lagging chromosomes resulting in a linearly declustered distribution of the kinetochores between two SPBs (Tytell and Sorger, 2006). To examine chromatin hyperstretching, we fluorescently labelled the pericentric chromatin 1.4 kb away from CEN5 (CEN5–GFP) and SPBs (Spc42–mCherry) in the wild-type and bmh1Δ bmh2Δ cells. We selectively scored cells either in the metaphase or in early anaphase stage judged by the SPB–SPB distance (2.0–4.0 µm) and categorized the behaviour of the sister chromatids based on the type of CEN5–GFP fluorescent signal as follows (Fig. 3A). Type-I and type-II have two sister CEN5–GFP signals that are coalesced into one focus or divided into two foci, respectively; type-III, aer cells with either one (top row) or two (bottom row) hyperstretched CEN5–GFP signals. Here, type-I represents the cells in early metaphase stage (SPB–SPB distance <2.5 µm), whereas type-II and -III represent the cells in late metaphase to early anaphase cells (SPB–SPB distance=2.5–4.0 µm). When comparing with wild-type, we observed a significant increase in the percentage of type-III population (∼27%) in bmh1Δ bmh2Δ mutants, with nearly an equivalent percentage reduction in the type-II population (∼24%); no significant change in type-I population (∼3%) was observed (Fig. 3B). These data indicate that the cells with hyperstretched chromatids are getting accumulated in bmh1Δ bmh2Δ mutants specifically during the metaphase–anaphase transition, and the hyperstretching is not prevalent during the early metaphase stages. Note that pericentric hyperstretching denotes a strong poleward force exerted over centromeres by the kMTs, which is possible only after the successful establishment of KT–MT attachment. Given that nearly 50% of the bmh1Δ bmh2Δ cells show hyperstretched chromatin, this indicates that KT–MT attachment is not perturbed in the majority of the cells.

Previously observed linear declustering of the kinetochores (Fig. 2D) clearly demonstrates the presence of lagging chromosomes between two SPBs during metaphase. However, similar measurements in anaphase cells (SPB–SPB distance>4 µm) showed that there was no significant increase in the declustering phenotype in the mutant over the wild-type suggesting an amelioration of the kinetochore clustering with time (Fig. S5A,B). In accordance with this, kinesin-8 motor (kip3Δ) mutants display lagging chromosomes with prolonged chromatid hyperstretching near centromeres due to altered microtubule dynamics during the metaphase–anaphase transition (Tytell and Sorger, 2006; Wargacki et al., 2010). Overall, we conclude that the kinetochore declustering (Fig. 2A,D) and chromatin hyperstretching (Fig. 3A,B) phenotypes observed in bmh1Δ bmh2Δ double mutants are perhaps due to defective chromosome congression, which is a consequence of the altered kMT dynamics, as shown by the schematics in Fig. 3C.

It has been shown previously that mutants with perturbations in KT–MT attachment-related functions activate the SAC and are synthetically lethal or sick with the SAC mutants (Wang and Burke, 1995; Pangilinan and Spencer, 1996; Hyland et al., 1999; Cheeseman et al., 2001; Daniel et al., 2006). As bmh1Δ bmh2Δ cells displayed both kinetochore perturbations and defective kMT dynamics, we believed that the observed G2/M delay (Fig. 1B,D) is SAC dependent and therefore the bmh1Δ bmh2Δ mutant would be expected to interact genetically with the SAC mutants. To examine this, bmh1Δ bmh2Δ mutant was crossed with an isogenic strain deleted for MAD2, the major SAC component (Li and Murray, 1991). The resulting diploids were sporulated and tetrads were dissected (genotypes of the obtained spores are listed in Table S3). We observed that the mad2Δ bmh1Δ bmh2Δ mutant spores were viable (Fig. 4A, solid squares) but their growth was marginally weaker than the bmh1Δ bmh2Δ mutant spores (Fig. 4A, dotted squares), indicating a synthetic negative interaction between mad2Δ and bmh1Δ bmh2Δ. Consequently, in the presence of benomyl mad2Δ bmh1Δ bmh2Δ cells performed very poorly compared to the corresponding double or single mutants (Fig. 4B). Notably, the mad2Δ bmh1Δ mutant grew less well than mad2Δ bmh2Δ. This is not unexpected as Bmh1 has been shown previously to be involved in SAC silencing and SPOC function (Grandin and Charbonneau, 2008; Caydasi et al., 2014). Interestingly, the mad2Δ bmh1Δ bmh2Δ cells grew less well than the mad2Δ bmh1Δ cells, indicating that Bmh2 contributes towards kinetochore-related functions in a manner that is not shared with Bmh1.

Abrogation of SAC-mediated G2/M arrest in presence of any perturbations in KT–MT interaction leads to progression into the cell cycle that generates multi-bud phenotype in budding yeast (Grandin and Charbonneau, 2008; Caydasi et al., 2014). Given that we observed that the bmh mutants genetically interact with mad2Δ (Fig. 4A,B) indicating that Bmh proteins might also have a role in SAC activation, we hypothesize that a fraction of bmh mutants might abrogate SAC-mediated G2/M arrest and generate multi-budded cells. While assessing phenotypes of the budded cells, we broadly categorized them as ‘cycling’ [small-budded cells with undivided DAPI with two or one SPB(s) in the mother compartment plus the large-budded cell with one each of DAPI and SPB in both the compartments], ‘G2/M’ (large-budded cells harbouring undivided DAPI with two SPBs within mother compartment), and ‘multi-budded’ (cells with multiple buds containing fragmented DAPI mass and super-numerary SPBs). We indeed observed a significant fraction of cells (∼23%) showed the multi-bud phenotype in the bmh1Δ bmh2Δ mutant, indicating that although there is a sizable fraction (∼47%) of cells at G2/M stage, accounting for SAC-mediated arrest, the double mutant is partially compromised in that arrest (Fig. 4D). Further removal of Mad2 in these cells caused, as expected, a nearly 2-fold increase (∼44%) in the multi-bud phenotype (Fig. 4D, compare black bars of bmh1Δ bmh2Δ and mad2Δ bmh1Δ bmh2Δ mutants) with a nearly equivalent reduction in the percentage of cells with the G2/M arrest phenotype again indicating that the latter phenotype in bmh1Δ bmh2Δ is largely dependent on SAC. However, compared with the wild-type (∼4%), no statistically significant increase in the percentage of multi-budded phenotype was observed in mad2Δ bmh1Δ (∼8%) or mad2Δ bmh2Δ (∼7%) cells, perhaps due to functional complementation between the Bmh proteins. Altogether, we conclude that the G2/M delay observed in the bmh1Δ bmh2Δ mutants (Fig. 1B,D) is perhaps due to Mad2-dependent transient activation of the SAC in response to the altered KT–MT attachment dynamics. Notably, in an previous study, bmh1Δ mutant cells where shown to display normal SAC activation and retained the securin Pds1 upon microtubule depolymerization (Caydasi et al., 2014). In another study, upon inducing tension defects, bmh1Δ mutants where shown to display normal SAC activation but failed to maintain the SAC arrest at later time points (Bokros et al., 2016). Therefore, a transient SAC activation and/or its persistence in bmh1Δ bmh2Δ mutants might reveal a marginal difference in Pds1 degradation kinetics between the mutant and the wild type. However, the bmh1Δ bmh2Δ mutants grow slower (∼2-fold) than the wild-type (Fig. 1A), making it challenging to detect subtle time differences in Pds1 degradation timings.

Given that bmh1Δ bmh2Δ mutant cells showed G2/M delay (Fig. 1) and KT–MT-related defects (Fig. 2), we hypothesize that the kinetochore function could be weak in these cells. Here, we utilized the W303 background strain (in which bmh double mutant is lethal) to explore the distinct functions of individual Bmh proteins and to eliminate the possibility of salvage pathways (see Fig. 7). To assess this, we tested the genetic interaction between bmh and kinetochore (ctf19Δ) mutants. We could not find any synthetic growth defect in ctf19Δ bmh1Δ or ctf19Δ bmh2Δ mutants under normal conditions (Fig. 5A, DMSO). However, in the presence of benomyl, ctf19Δ showed a considerable growth defect as reported previously (Hyland et al., 1999); the ctf19Δ bmh2Δ mutant grew worse than ctf19Δ (Fig. 5A). Interestingly, the growth of ctf19Δ became marginally better upon removal of Bmh1 (Fig. 5A). These results suggest that the loss of Bmh proteins, and Bmh2 in particular, might influence the kinetochore function. We also tested genetic interaction between bmh and kinetochore mutants in the SK1 background using another non-essential kinetochore protein mutant, iml3Δ, which is similar to ctf19Δ (Ghosh et al., 2001; Pot et al., 2003). We presumed that as the bmh double mutant in the SK1 background is viable, the removal of both the Bmh proteins might be required to detect genetic interactions with iml3Δ. As expected, we observed minimal growth discrepancies between iml3Δ, iml3Δ bmh1Δ, and iml3Δ bmh2Δ mutants when exposed to benomyl (Fig. S6). Although the iml3Δ bmh1Δ and iml3Δ bmh2Δ mutants exhibited slightly poorer growth than the iml3Δ mutant, the subtle differences between the double mutants were not discernible in the spotting assay. However, the bmh1Δ bmh2Δ iml3Δ mutant displayed significantly poorer growth compared to iml3Δ or bmh1Δ bmh2Δ mutants, indicating the collective impact of Bmh proteins on kinetochore function (Fig. S6).

The requirement of any non-essential protein for kinetochore function in budding yeast can be assessed by measuring the mitotic stability of minichromosomes (Maine et al., 1984; Sanyal et al., 1998; Poddar et al., 1999; Ghosh et al., 2001). In accordance with the where shown to display observations (Fig. 5A), an enhanced loss of the minichromosome in ctf19Δ bmh2Δ mutant compared to ctf19Δ or bmh2Δ single mutant denotes that Bmh2 might be required for proper kinetochore function (Fig. 5B). Notably, although minichromosome instability can also arise due to defects in DNA replication from ARS, we believe that kinetochore dysfunction plays here the major role as further evidence of kinetochore alteration has been provided below. In the transcriptional read-through assay, transcription across a centromere and a downstream reporter is perturbed in wild-type cells due to kinetochore complex formation at the centromere (Doheny et al., 1993). However, in cells with compromised kinetochore complex, the transcription can proceed causing the reporter expression (Fig. 5C). The β-galactosidase activity, as a read-out of LacZ reporter expression, from the bmh mutants and kinetochore mutant ctf19Δ was compared with that in wild-type cells. Similar to what was seen with the ctf19Δ mutant, the relative β-galactosidase activity was found to be high in bmh2Δ but not in bmh1Δ cells (Fig. 5D), indicating the requirement of Bmh2 but not Bmh1 for kinetochore structural integrity. Dicentric plasmids within the wild-type cells end up with breakage due to the microtubule-mediated opposite pulling force acting on two kinetochores formed on two centromeres. Hence, a dicentric plasmid will be unstable and will be converted into a monocentric plasmid of different sizes in different cell lineages upon rearrangements and ligation (Fig. 5E). Consequently, the colonies that appeared after transformation of the cells with dicentric plasmid will be of heterogeneous sizes (Koshland et al., 1987; Sanyal et al., 1998). Conversely, a kinetochore mutant cell avoids the breakage due to the weak KT–MT interaction, causing the dicentric plasmid to be stabilized, producing homogenous transformant colonies. As expected from the above results and previous reports, ctf19Δ and bmh2Δ mutants produced homogenous colonies indicating that they have compromised kinetochore integrity, whereas the bmh1Δ mutant showed heterogenous colonies like in the wild-type (Fig. 5F).

In summary, the above observations together suggest that Bmh proteins are important for proper functioning of budding yeast kinetochore. Among the Bmh proteins, Bmh2 promotes the functions of the kinetochore, perhaps by influencing its integrity. By contrast, a significant increase in mitotic stability of a minichromosome in ctf19Δ bmh1Δ over ctf19Δ alone (Fig. 5B), a β-galactosidase activity lower than the wild-type in transcriptional read-through assay (Fig. 5D) and dicentric plasmid instability (Fig. 5F) in bmh1Δ cells hint towards a positive impact of loss of Bmh1 on kinetochore integrity. As a corollary, overall stress tolerance and enhancement in the life span of budding yeast cells with bmh1Δ mutation have been reported previously (Wang et al., 2009).

Bmh-interacting motifs have been identified in several kinetochore and SPB proteins (Kumar, 2018) and in a previous high-throughput screening of direct physical interactors of Bmh proteins, the kinetochore protein Iml3 was co-purified (Kakiuchi et al., 2007). These observations and our findings strongly suggest that Bmh proteins contribute to the kinetochore function. In this context, we hypothesize that Bmh proteins are physically present at the kinetochores to execute their roles. To investigate this, we constructed strains where both Ndc80 and either Bmh1 or Bmh2 were tagged with 6×HA and 13×Myc at their C-terminals, respectively. The fused proteins were functional as evident from previous studies (Janke et al., 2001; Slubowski et al., 2014). Their localizations were assessed simultaneously on chromatin spreads made from the cells harvested from different stages of the cell cycle that were judged from the spreads as follows: G1/S or metaphase stage as a single DAPI mass harbouring a single Ndc80 focus (Fig. 6A) or bi-lobed Ndc80 foci (Fig. 6B), respectively; and anaphase stage as two (separated or connected) DAPI masses each harbouring one Ndc80 focus (Fig. 6C). Given that Bmh proteins are known for their binding to the cruciform DNA and they localize on the chromatin at the autonomously replicating sequence (ARS) sites (Yahyaoui et al., 2007; Yahyaoui and Zannis-Hadjopoulos, 2009), we expected multiple foci of Bmh proteins of which a subset will be kinetochore specific on the spreads. We indeed observed chromatin localization of Bmh proteins as multiple foci on the spreads from different cell cycle stages (Fig. 6A–C). From each cell cycle stage, the extent of colocalization between Ndc80 and Bmh proteins was measured using a Pearson's correlation coefficient (PCC), and the obtained values were compared with no-tag control for statistical significance (Fig. 6D). After thorough analysis (see Materials and Methods), we found that a significant percentage (>60%) of Bmh proteins colocalized either partially or completely with Ndc80 throughout the cell cycle (Fig. 6E). The colocalization patterns ‘No’ (PCC<0.1), ‘partial’ (PCC=0.1–0.5), and ‘complete’ (PCC>0.5) were categorized based on PCC values as described elsewhere (Zinchuk and Grossenbacher-Zinchuk, 2014; Prajapati et al., 2017). These results indicate that at least a proportion of Bmh proteins resides at or close to the centromeres.

To further investigate whether the Bmh proteins are targeted to the centromeres, we performed a ChIP assay. As Bmh proteins colocalize with kinetochore in all cell cycle stages (Fig. 6E), asynchronously grown cultures were analysed for centromere localization estimating centromere (CEN) fragment enrichment. Surprisingly, no significant localization of Bmh proteins on the centromere was observed (Fig. S7A,B). Failure to detect centromeric localization by ChIP is perhaps due to the transient nature of the physical interaction between Bmh proteins and the kinetochore protein(s). In summary, we conclude that the Bmh proteins either partially or completely localize at the kinetochores across the cell cycle stages to functionally contribute to the kinetochore functions.

The 14-3-3 family of proteins are conserved across eukaryotes and serve diverse regulatory functions in cells. Our investigation in budding yeast reveals that 14-3-3 homologs (Bmh proteins) play a crucial role in KT–MT-related functions. Specifically, Bmh proteins promote kinetochore organization, stabilize KT–MT dynamics and facilitate proper chromosome congression during mitosis.

In budding yeast, Bmh1 physically interacts with an intermediate filament protein Fin1, to resist its kinetochore localization, which has implications in SAC silencing (Akiyoshi et al., 2009b; Bokros et al., 2016; 2021). Nevertheless, whether these proteins have a direct impact on kinetochore function was never investigated. Here, we demonstrate a functional correlation between Bmh proteins and kinetochore function using transcriptional read-through, dicentric stabilization, genetic interaction and subcellular colocalization assays.

Given that Iml3, which harbours a Bmh-interacting motif, physically interacts with Bmh1-interacting protein Fin1 (Kakiuchi et al., 2007; Akiyoshi et al., 2009b), we speculate that Bmh proteins might be targeted at or be close to the centromeres through trimeric interaction of Iml3–Fin1–Bmh. Alternatively, but not mutually exclusively, Bmh proteins might directly recognize centromere cruciform structures (Callejo et al., 2002); however, direct or indirect CEN DNA binding of Bmh proteins is unlikely, as we failed to detect these proteins at the centromeres by ChIP. Bmh proteins can be targeted to centromeres through an association with phosphorylated H3S10 mark written by the yeast Aurora B kinase Ipl1 (Jain et al., 2021). Given that Bmh proteins capable of forming homo- and hetero-dimers can cross-link two phosphoproteins, it is not surprising that Bmh proteins might have roles in correct kinetochore assembly by bridging kinetochore proteins, as several of them are phosphorylated. For instance, in humans, a few 14-3-3 isoforms have been reported to interact with phosphorylated CENP-A and CENP-C to form a connecting bridge and thereby stabilize the kinetochore ensemble (Goutte-Gattat et al., 2013). Our observations of kinetochore declustering (Fig. 2D) and mislocalization of the kinetochore proteins (Fig. 2C) in absence of both the Bmh proteins further support this notion. As Bmh isoforms are known to interact with phosphorylated Fin1 (Mayordomo and Sanz, 2002; Akiyoshi et al., 2009b), the observed Bmh2-specific interaction with kinetochores (Fig. 5A–F) might also be Fin1 dependent. Given that early Fin1 localization and consequent SAC silencing at kinetochores were specific to bmh1Δ mutants (Bokros et al., 2016), it can be argued that Bmh2 perhaps holds non-shared functions at kinetochores. In summary, we report here both cumulative and non-shared contributions of Bmh isoforms related to kinetochore functions. Notably, although the strains like ∑1278 and SK1 can survive in absence of both the Bmh proteins by upregulation of Ras-cAMP signalling through overexpression of Ras-cAMP-dependent protein kinase (Tpk1) (Stanhill et al., 1999; Gelperin et al., 1995), the same is not likely to rescue the observed kinetochore function defect in bmh1Δ bmh2Δ as Tpk1 overexpression alone has been found to be detrimental to kinetochore function (Lina et al., 2012; Shah et al., 2019).

In the context of non-shared functions, it is interesting to observe that, unlike bmh2Δ, the bmh1Δ mutant alleviated the impact of non-essential kinetochore mutants (Fig. 5A–F). In general, 14-3-3 isoforms have non-shared functions reported in higher eukaryotes due to their differences in transcription levels, subcellular localization, structure and their preference to form homo- or hetero-dimers (Chaudhri et al., 2003; Obsil and Obsilova, 2011; Slubowski et al., 2014; Abdrabou et al., 2020). Given that the dimerization preferences of 14-3-3 isoforms are linked to unique cellular functions (Aitken, 2002), the non-shared functions of Bmh isoforms in budding yeast might also arise from their specific dimerization preferences. However, the majority of Bmh isoforms endogenously exist as heterodimers (Chaudhri et al., 2003), suggesting although they remain together, one has a predominant role in a context-dependent manner. For example, whereas only Bmh1 has a role in Fin1 localization at the kinetochore for SAC removal (Bokros et al., 2016; 2021), strong physical affinity is reported between Fin1 and both the Bmh isomers (Akiyoshi et al., 2009b). Therefore, in the majority of cases, the loss of one isoform can support cell survival as another isoform can function optimally as homodimers. However, the non-shared functions are unmasked under specific conditions when perhaps the full functional activity of Bmh proteins requires certain condition-specific ratios of both homo- and hetero-dimers. Another potential explanation could be that the specific kinases that phosphorylate the targets of the Bmh proteins function in a manner that translates to non-shared functions of the Bmh proteins.

We noticed several phenotypes of bmh double mutants that account for the involvement of Bmh proteins in multiple events of chromosome segregation. The observed delay in SPB–kinetochore duplication, their disjunction (Fig. 1D), and the accumulation of multi-budded cells (Fig. 4D) in bmh mutants are perhaps due to the contributions of Bmh proteins in DNA replication, kinetochore functions and DNA damage checkpoint (DDC) activation, respectively (Lottersberger et al., 2003; Usui and Petrini, 2007; Grandin and Charbonneau, 2008; Bokros et al., 2016).

The increase in multi-budded cells in bmh double mutants upon removal of Mad2 (Fig. 4D) also suggests that lack of Bmh proteins causes kinetochore perturbation resulting in altered KT–MT dynamics and SAC activation, which might also contribute to the delay in SPB–kinetochore disjunction (Fig. 1D). In general, the multi-budded phenotype is attributed to various cellular malfunctions, such as impaired SAC, SPOC, mitotic exit and cytokinesis. Given that Mad2 functions through a distinct pathway from SPOC protein Bub2 (Fraschini et al., 1999), and both bmh1Δ and bmh2Δ single mutants by themselves alone do not affect SPOC function (Caydasi et al., 2014), we can rule out perturbation of SPOC as being responsible for the generation of multi-budded phenotype in the bmh1Δ bmh2Δ mutants. However, as Bmh proteins are involved in multiple cellular pathways, we cannot rule out that defective mitotic exit or cytokinesis underlie the multi-budded phenotype. In Drosophila oocytes, 14-3-3 isoforms are reported to perform SAC functions by regulating the localization of the chromosomal passenger complex (CPC) component Borealin (Repton et al., 2022). Notably, it is possible that DNA damage generated due to improper DNA replication in bmh1Δ bmh2Δ mutant might also activate the SAC, given that in budding yeast it has been reported that DNA damage-induced epigenetic alteration at the centromeres can alter kinetochore assembly to activate the SAC (Dotiwala et al., 2010). Overall, the phenotypes of the cell cycle delay as well as the presence of a subpopulation of multi-budded cells in bmh1Δ bmh2Δ mutants, suggest that the mutants not only activate a DDC and/or the SAC but are also compromised in sustaining the checkpoint-mediated cell cycle arrest. In support of this, it has been reported that bmh mutants are sensitive to DNA-damaging and microtubule-depolymerizing drugs (Grandin and Charbonneau, 2008), and the bmh1Δ mutant, in particular, has been found to cause premature SAC silencing (Bokros et al., 2016).

The presence of declustered kinetochores (Fig. 2D) and chromatid hyperstretching phenotype (Fig. 3A) in bmh mutants could possibly be due to malfunctioning of microtubule-associated motors as Bmh proteins were co-purified with kinesin-related motors (Cin8 and Kar3), a Kar3-associated protein (Cik1) and a protein involved in nuclear orientation and congression (Bim1) during mitosis (Kakiuchi et al., 2007). cin8Δ and kar3Δ mutants display a kinetochore declustering phenotype similar to that of the bmh mutants (Tytell and Sorger, 2006; Jin et al., 2012), and cik1Δ cells, like bmh mutants, show activation of the SAC and an increase in the syntelic attachment rate with DNA replication errors (Liu et al., 2011; Jin et al., 2012). Moreover, Bim1 and Kar3 are known to have a substantial role in nuclear orientation and migration during mating and mitosis (Beach et al., 2000; Molk et al., 2006). Noticeably, in a substantial percentage of bmh1Δ bmh2Δ mutant cells, the DAPI mass (nucleus) remains away from the bud neck perhaps due to underlying motor protein-mediated nuclear migration defects (Fig. 1C). Direct functional correlations between 14-3-3 isoforms and microtubule-associated proteins including motor proteins have been observed in higher eukaryotes (Dorner et al., 1999; Lu and Prehoda, 2013); however, whether this holds true in budding yeast requires further investigation.

Given that proper phosphorylation of kinetochore proteins, such as Cse4, Ndc80 and Dam1, by Ipl1 kinase is essential for correct sister kinetochore biorientation and the formation of the canonical bilobed structure (Akiyoshi et al., 2009a; Boeckmann et al., 2013; Jin et al., 2017; Mittal et al., 2019), we believe that altered phosphorylation of the kinase substrates at the kinetochores might also cause the declustering phenotype in bmh mutants. To corroborate, it has been reported that PP1 phosphatase can dephosphorylate Ipl1 substrates at the kinetochore, and Bmh1 restrains PP1 activity by sequestering it in a trimeric Bmh1–Fin1–PP1 complex (Bokros et al., 2016; 2021). Therefore, it is plausible that a pre-mature dephosphorylation of the kinetochore substrates of Ipl1 can happen in bmh double mutants, resulting in the kinetochore declustering phenotype. In support of our argument, upon depletion of Ipl1, kinetochore declustering phenotypes have been observed in Candida albicans (Varshney and Sanyal, 2019). It would be interesting to investigate the functional correlation between Bmh proteins and Ipl1 substrates on kinetochores. In summary, we contribute to the growing list of cell cycle functions of Bmh proteins by providing evidence that these proteins influence the kinetochore ensemble, stable KT–MT dynamics, synchronous chromosome congression and sustained activation of the checkpoints. Based on our observations and previous literature, we generated a model explaining the phenotypes of bmh1Δ bmh2Δ double mutants in comparison with wild-type (Fig. 7).

The 14-3-3 proteins recognize their binding partners via phosphorylated motifs. Recently, these motifs have been found within the intrinsically disordered regions (IDRs) of 14-3-3-binding partners (Bustos, 2012; Sluchanko and Bustos, 2019), and numerous 14-3-3 partners are predicted to have propensity to phase separate as membrane-less entities (Huang et al., 2022). In this context, the literature suggests that 14-3-3 isoforms often temporally sequester their binding partners from their native cellular localization sites. For instance, 14-3-3 proteins sequester Cdc25 from the nucleus (Lopez-Girona et al., 1999; Zeng and Piwnica-Worms, 1999; Meng et al., 2013), Fin1 from kinetochore (Bokros et al., 2016), Bfa1 from the SPB (Caydasi et al., 2014), Borealin and kinesin-14/Ncd from microtubules (Repton et al., 2022), and BAD proteins from the apoptotic pathway (Tan et al., 2000). We presume that the binding partners, while under such sequestration, might exist as a phase-separated entity. Hence, the function of the Bmh proteins at the kinetochore might also be manifested by sequestration of the kinetochore proteins into phase-separated entities so that the latter become available at the centromeres spatiotemporally, regulating their hierarchical assembly at the kinetochore. Investigating further on the mechanism of Bmh protein-mediated regulation of kinetochore integrity is an interesting prospective.

All Saccharomyces cerevisiae strains are of either SK1 or W303 genetic background. The detailed descriptions of strains and plasmids used in this study are listed in Table S1. Unless specified, as a standard condition, the strains were grown in rich medium [YEPD; 1% yeast extract (HiMedia, #RM027), 2% peptone (HiMedia, #RM001) and 2% dextrose (Merck, #MF4M741178)] at 30°C till mid-log phase [optical density at 600 nm (OD₆₀₀)=∼1.0] before harvesting for the experiments. Gene modifications like C-terminal fluorescence or affinity tagging, gene deletions, and linearized plasmid integration into specific genome loci were achieved through lithium acetate transformation (Gietz and Woods, 2002) using homologous recombination. The linear cassettes for gene modifications were PCR amplified using template plasmids acquired from Euroscarf (Wach et al., 1997), and the modifications were verified by diagnostic PCR. For conditional mutants of essential genes, auxin-based degron systems (Nishimura et al., 2009) were utilized, as mentioned elsewhere (Mehta et al., 2014). For chromatin hyperstretching experiments, we inserted [TetO]₂₂₄ arrays at 1.4kb away from CEN5 in the cells constitutively expressing TetR–GFP, as performed elsewhere (Michaelis et al., 1997; Tanaka et al., 2000). For G1 synchronization of cells, α factor was used as described elsewhere (Breeden, 1997; Kumar et al., 2021). For genetic interaction with Mad2, the spores with required genotypes were obtained through meiotic induction of appropriate diploids, as described previously (Mehta et al., 2014) followed by tetrad dissection using a Zeiss Axio (Scope A1) micromanipulator (20x objective) in YEPD plates and incubation at 30°C for 2–3 days for the spore growth.

Cells grown in standard conditions till mid-log phase were sonicated briefly following which they were counted using a haemocytometer (Rohem, India, #10269/BS 748), 10-fold serially diluted and spotted on control (DMSO) and benomyl-containing YEPD plates. The appropriate concentrations of benomyl (Sigma, #45339) dissolved in DMSO were added while preparing the plates. After spotting, the plates were incubated at 30°C for 2–5 days depending on the growth rate of mutants and genetic background of the strains, before they were photographed.

For whole-cell protein extraction, nearly 10 ml of mid-log phase (OD₆₀₀=∼1.0) cells were harvested and treated with 0.1 NaOH solution, as described before (Kushnirov, 2000). Subsequently, the processing of extracted protein samples, western blottin, and relative band intensities quantifications were performed, as mentioned elsewhere (Mittal et al., 2020). The antibodies and their dilutions (in 1× Tris-buffered saline with 0.1% Tween 20; TBST) were as follows. Primary antibodies: rat anti-HA (Roche, #12158167001) at 1:5000, mouse anti-GFP (Roche, #11814460001) at 1:3000, and rat anti-tubulin (Serotec, MCA78G) at 1:5000. Secondary antibodies: horseradish peroxidase (HRP)-conjugated goat anti-rat-IgG (Jackson, # 112-035-167) and goat anti-mouse-IgG (Jackson, # 115-035-166) at 1:5000.

To access minichromosome (CEN plasmid) stability, we followed the method as described earlier (Prajapati et al., 2017; Kumar et al., 2021). Typically, cells harbouring CEN plasmids (YCplac3; Gietz and Sugino, 1988) were grown overnight in selective medium (SC-Ura; HiMedia, #G155), and re-inoculated at OD₆₀₀=0.005 into non-selective medium (YEPD) and incubated at 30°C for 30–40 h to allow cells to reach ‘N’ generations. The plasmid-containing cell fractions at the initial (f0) and final (fN) time points in YEPD were estimated by plating equal volumes of cultures on SC-Ura and YEPD plates. The percentage minichromosome loss rate was calculated using the following equation, loss rate (%)=(1/N) [ln (f0/fN)]×100, with N=number of generations; f0=fraction of plasmid-bearing cells at ‘0’ generation; fN=fraction of plasmid-bearing cells at ‘N’ generations.

The dicentric plasmid stability assay and transcriptional read-through assay were performed as described elsewhere (Doheny et al., 1993). To construct a dicentric plasmid pRT1 (pGAL1-CEN4-LacZ), a 110 bp long CEN4 fragment was PCR amplified (using CEN4 primers, Table S2) and cloned into SalI site placed upstream to LacZ ORF of a centromeric plasmid pAKD06 (Rizvi et al., 2017) harbouring a pGAL1-LacZ cassette. As the readout for mitotic instability of dicentric plasmid was reported earlier as colony size heterogeneity on the transformant plate (Mann and Davis, 1983; Koshland et al., 1987; Doheny et al., 1993), mid-log phase (OD₆₀₀=∼1.0) grown cells were transformed with pRT1 plasmid, and the size heterogeneity of transformant colonies were relatively compared between wild-type and mutants on the selection plates.

We performed the transcriptional read-through assay as described elsewhere (Mann and Davis, 1983; Koshland et al., 1987; Doheny et al., 1993). We utilized the pRT1 plasmid (this study) harbouring pGAL1-CEN4-LacZ to assess the GAL promoter-driven LacZ expression with the transcriptional block imposed by kinetochores assembled over CEN4. To do so, the pGAL1-CEN4-LacZ cassette was excised from the pRT1 plasmid and cloned into pRS406 (a yeast integrative plasmid) using HindIII and SpeI restriction enzymes to construct the pRT2 plasmid. pRT2 was linearized using StuI and was integrated into URA3 locus of wild-type and mutant cells and the transformants were maintained under galactose to keep the integrated CEN4 (pGAL1-CEN4-LacZ) inactive. In the wild-type, a kinetochore formed on the integrated CEN4 offers resistance to LacZ expression, which was assayed by β-galactosidase activity. In contrast, in the mutants, due to compromised kinetochore integrity, the resistance is attenuated causing increased LacZ expression. The β-galactosidase activity was determined by an o-nitrophenyl β-D-galactopyranoside (ONPG) liquid assay (quantitative) as mentioned elsewhere (Clontech Laboratories, 2008). β-galactosidase activity from each sample was calculated from at least three biological replicates and the concentration of cells (OD₆₀₀) was normalized to compare the β-galactosidase activity between wild-type and mutants.

Fluorescence imaging was undertaken as described elsewhere (Mittal et al., 2020). Briefly, cells grown in standard conditions until mid-log phase (OD₆₀₀=∼1.0) were fixed by adding 5% formaldehyde solution directly to the culture medium and incubating at room temperature (RT) for 5 mins. The fixed cells were harvested and washed twice with 0.1 M phosphate buffer (pH 7.5) before imaging. For chromatin visualization, the cells harvested after formaldehyde fixation were briefly washed once with 50% ethanol, followed by two 0.1 M phosphate buffer (pH 7.5) washes. Finally, the cells were re-suspended in freshly prepared DAPI (Invitrogen, #D1306) (1 μg/ml) solution before imaging. Optionally, a brief sonication step was introduced to de-clump cells before imaging. The images were acquired through a Zeiss Axio Observer Z1 fluorescence microscope (63× or 100× objective) in z-stack mode (0.2–0.5 μm spacing). For intensity comparison and 3D distance measurements, a Zeiss confocal laser scanning microscope (LSM 780) with a 32-array GaAsP detector was used. Based on the fluorescence signal intensity, the exposure time was determined and kept constant across the samples for comparison.

The z-stack images acquired through Zeiss Axio (Observer Z1) fluorescence microscope or confocal laser scanning (LSM 780) microscope were merged and processed for further analyses using Zeiss Zen 3.1 (blue edition) software. In represented merged images, the cell boundaries (dotted lines) were delineated referencing their brightfield images. For 3D distance measurements, the distance between any two fluorescent foci was determined by marking their centroids across z-stacks using the Imaris 8.0.2 (Bitplane/Slice tool) software, as described elsewhere (Mittal et al., 2020). The obtained SPB–SPB and CEN–SPB 3D distances (in nm) were utilized to determine the cell cycle stage and CEN–SPB proximity, respectively, as described elsewhere (Tytell and Sorger, 2006; Sau et al., 2014).

The line scans were performed over live-cell images after merging their z-stacks projecting maximum fluorescence intensities using Zeiss Zen 3.1 (blue edition) software. The kinetochore signal distribution in the merged images was obtained using the ‘Profile’ tool in Zeiss Zen 3.1 (blue edition) software.

The Pearson's correlation coefficient (PCC) values to assess the colocalization of any two fluorescent signals were calculated from merged z-stack images using Imaris 8.0.2 (Coloc tool) software, as described elsewhere (Prajapati et al., 2017). The obtained PCC values were categorized as mentioned previously (Zinchuk and Grossenbacher-Zinchuk, 2014) to determine the colocalization significance.

For live-cell time-lapse imaging, cells were grown overnight at 30°C in YEPD medium supplemented with adenine (5 mg/ml; SRL, #50300) following which they were re-inoculated into SC medium and grown until they reached and OD₆₀₀=0.5. Subsequently, 0.2–0.5 ml of culture was added over confocal dishes (Alkali Scientific) pre-coated with concanavalin A (0.25 mg/ml) solution and air-dried for 30 mins in sterile conditions for cell adherence. Unbound cells were washed away using sterile SC broth. The confocal discs were mounted over the microscope stage (temperature maintained at 30°C) to acquire automated time-lapse images at specified time intervals (at each interval, multiple fields were acquired with z-stacks) for 12–15 h. Images were acquired using a Nipkow spinning disc (Yokogawa CSU-X1 automated model) microscope equipped with an EMCCD camera. Post-acquisition processing was performed using Zeiss Zen 3.1 (blue edition) software.

Immunofluorescence was performed as mentioned previously (Mittal et al., 2020; Shah et al., 2023). Briefly, 5 ml of mid-log phase (OD₆₀₀=∼1.0) cells were fixed by adding 5% formaldehyde solution directly to the culture medium and incubated at room temperature (RT) for 15 mins. The fixed cells were harvested, washed once with PBS and resuspended in spheroplasting solution (1.2 M sorbitol, 0.1 M phosphate buffer pH 7.5), and treated with Zymolyase 20 T (10 mg/ml, MP Biomedicals, #32092) for 1 h at 30°C for spheroplasting before mounting on poly-lysine-coated wells on a slide. After washing the unbound cells with PBS, the cells adhered to poly-lysine-coated wells were permeabilized and flattened by immersing the slide in −20°C methanol for 5 mins and in acetone for 30 s. After allowing slides to air-dry for 1–2 mins, blocking was undertaken using 5% skimmed milk solution prepared in dilution buffer (10 mg/ml BSA in PBS). Subsequently, the cells were incubated with appropriate primary and secondary antibodies, and DAPI (Invitrogen, #D1306) (1 μg/ml) solutions. The intermittent washes were performed in PBS. Finally, after spreading the mounting solution (1 mg/ml phenylenediamine in 90% glycerol), the glass slides were sealed with coverslips for imaging. The antibodies and their dilutions (in dilution buffer) were as follows. Primary antibodies: rat anti-HA (Roche, #12158167001) at 1:200, mouse anti-GFP (Roche, #11814460001) at 1:100, mouse anti-Myc (Roche, #11667149001) at 1:200 and rat anti-tubulin (Serotec, MCA78G) at 1:5000. Secondary antibodies: (TRITC)-labelled goat anti-rat-IgG (Jackson, #115485166), Alexa Fluor 488-labeled goat anti-rat-IgG (Jackson, #112545167) and TRITC-labelled goat anti-mouse-IgG (Jackson, #115025166) at 1:200.

Chromatin spreads were made as mentioned previously (Prajapati et al., 2018; Mittal et al., 2020). Briefly, 2 ml of mid-log phase (OD₆₀₀=∼1.0) cells were harvested, washed once with PBS and spheroplasted using Zymolyase 20T, as mentioned above in the indirect immunofluorescence protocol section. After spheroplasting, the reaction was stopped using an ice-cold stop solution (0.1 M Morpholineethanesulfonic acid, 1mM EDTA, 0.5mM MgCl₂, 1 M sorbitol pH 6.4); and the spheroplasted cells were harvested and washed gently in PBS before mounting on a clean glass slide. The cells were treated with freshly made paraformaldehyde solution (4% paraformaldehyde, 3.4% sucrose and a few drops of NaOH solution to dissolve paraformaldehyde), followed by 1% Lipsol (LIP Equipment and Services) solution for cell lysis. The lysed cells were homogenously spread over the slide and air-dried at RT. After overnight drying, the slides were treated with 0.4% Kodak Photoflow-200 to avoid photobleaching, followed by 5% skimmed milk as blocking solution. Subsequently, the slides were treated with appropriate primary and secondary antibodies and DAPI (1 μg/ml) solutions. The antibodies and their dilutions are identical to those used for indirect immunofluorescence. The intermittent washes were performed in PBS solution. Finally, after spreading the mounting solution (1 mg/ml phenylenediamine in 90% glycerol), the glass slides were sealed with coverslips for imaging.

The ChIP assay was performed as described previously (Mehta et al., 2014; Shah et al., 2023). Briefly, 50 ml of mid-log phase (OD₆₀₀=∼1.0) cells were fixed with 1% formaldehyde for 1–2 h (the duration depends on chromatin proximity of the targeting protein) at 25°C. The fixed cells were harvested, washed twice with 1× TBS, and resuspended in lysis buffer [50 mM HEPES-KOH, pH 7.5, 140 mM KCl, 1 mM EDTA, 1% Triton X-100, 0.1% sodium deoxycholate, and 1× PIC (protease inhibitor cocktail, Roche)]. The cells were lysed using 0.5 mm glass beads using a mini-bead beater (BIOSPEC), followed by chromatin fragmentation using water bath sonicator (Diagenode SA, Picoruptor, BC100) to obtain 200–600 bps fragments. After clarifying the lysate fractions, 3–5 μg of appropriate antibodies were added and incubated at 4°C overnight with gentle rotation. Subsequently, Protein-A conjugated Sepharose beads (GE Healthcare, 17-0780-01) were added and incubated at 4°C for 2 h. Finally, the chromatin fragments were eluted by sequential washing, de-crosslinking, Proteinase K treatment and phenol:chloroform:isoamyl (PCI) purification. The enrichment of obtained chromatin fragments was estimated using qPCR (Bio-Rad CFX96 Real-Time machine) using specific primers targeting specific and non-specific (negative control) chromatin loci, as listed in Table S2. As mentioned previously (Mehta et al., 2014; Mittal et al., 2020), the following equation was used to estimate the percentage chromatin enrichment. ChIP efficiency=Enrichment/Input X 100; Enrichment/Input=E∧-ΔCT; ΔCT=CT(ChIP)−[CT(Input)−LogEX(D)]; E=primer efficiency value, CT=Threshold values obtained from qPCR, D=Input dilution factor. E was estimated as {[10^(–1/slope)]– 1} from standardization graphs of CT values against dilutions of the input DNA.

Error bars in the individual bar graphs represent the s.e.m. derived from the standard deviation (s.d.) of at least three independent biological replicates. The ‘N’ values denote the total number of cells analysed, obtained from ‘n’ number of biological replicates for individual assays. For all the figures, the statistical significance (P) was calculated using a two-tailed Student's t-test (paired or unpaired, depending on the data type). P≤ 0.05 is categorized as significant differences and ‘ns’ represents statistically not significant. The s.d., s.e.m. and statistical significance (P) values were calculated using automated modules of Microsoft Excel/GraphPad Prism 9.0 (Version 9.4.1) software.

We thank the central instrumentation facility at the BSBE department of IIT Bombay for their help.

The peer review history is available online at https://journals.biologists.com/jcs/lookup/doi/10.1242/jcs.261928.reviewer-comments.pdf