Abstract
The accurate segregation of chromosomes during mitosis relies on the attachment of sister chromatids to microtubules from opposite poles, called biorientation. Sister chromatid cohesion resists microtubule forces, generating tension which provides the signal that biorientation has occurred. How tension silences the surveillance pathways that prevent cell cycle progression and correct erroneous kinetochore-microtubule remains unclear. Here we identify SUMOylation as a mechanism that promotes anaphase onset upon biorientation. SUMO ligases modify the tension-sensing pericentromere-localized chromatin protein, shugoshin, to stabilize bioriented sister kinetochore-microtubule attachments. In the absence of SUMOylation, Aurora B kinase removal from kinetochores is delayed. Shugoshin SUMOylation prevents its binding to protein phosphatase 2A (PP2A) and release of this interaction is important for stabilizing sister kinetochore biorientation. We propose that SUMOylation modulates the kinase-phosphatase balance within pericentromeres to inactivate the error correction machinery, thereby allowing anaphase entry in response to biorientation.
Introduction
Mitosis divides the nucleus to produce two genetically identical daughter cells. Prior to mitosis, DNA replication produces sister chromatids, linked together by the cohesin complex. Sister chromatids are aligned at metaphase, thus allowing microtubule spindles to be captured by kinetochores assembled on centromeres. The correct form of attachment is termed ‘biorientation’, meaning that the kinetochores on the two sister chromatids are attached to microtubules emanating from opposite spindle poles. Biorientation creates tension, because cohesin holding sister chromatids together resists the pulling force of microtubules [1]. The fulfilment of biorientation allows securin degradation and, consequently, the activation of the protease separase, which cleaves cohesin, triggering sister chromatid separation (reviewed in [2]).
The conserved shugoshin protein plays key roles in promoting biorientation in mitosis and preventing cell cycle progression where biorientation fails [3, 4]. Budding yeast possesses a single shugoshin gene, SGO1. Sgo1 localizes to both the core ∼125bp centromere, where the kinetochore resides, and the surrounding ∼20kb cohesin-rich chromosomal region called the pericentromere [5]. The kinetochore-localized Bub1 kinase promotes Sgo1 enrichment at the pericentromere through phosphorylation of S121 on histone H2A [6–8]. Sgo1, in turn, recruits condensin and protein phosphatase 2A, PP2A-Rts1, to the pericentromere and maintains the chromosome passenger complex (CPC) containing Aurora B kinase at centromeres during mitosis [9, 10]. Condensin at pericentromeres is thought to bias the conformation of the sister chromatids to favour biorientation. The CPC recognizes erroneous microtubule-kinetochore attachments and destabilizes them, thereby maintaining the activity of the spindle assembly checkpoint (SAC) to prevent anaphase entry (reviewed in [11]). In vertebrate cells, PP2A-B56 protects cohesin in pericentromeres from removal by the so-called prophase pathway, which removes cohesin through a non-proteolytic mechanism that is independent of separase [12, 13]. In budding yeast, PP2A-Rts1 is recruited by shugoshin despite the absence of the prophase pathway [9,10,14]. Instead, PP2A-Rts1 has been implicated in ensuring the equal segregation of sister chromatids during mitosis, since mutants failing to recruit PP2A-Rts1 to the centromere are unable to respond to a lack of inter-sister kinetochore tension and mis-segregate chromosomes upon recovery [9, 14].
Sgo1 both directs and responds to cell cycle cues as chromosomes establish and achieve biorientation, upon which anaphase entry is triggered. A key signal that biorientation has occurred is the tension-dependent removal of Sgo1 from the pericentromere during metaphase, resulting in the delocalization of its effectors, PP2A-Rts1, condensin and the CPC [8, 14]. Upon anaphase I onset, Sgo1 is ubiquitinated and degraded by APC/C-Cdc20 [14, 15]. Similarly, human shugoshin is degraded in anaphase as a result of APC/C-Cdc20 activity [16]. Shugoshin can be stabilized by mutation of its APC-Cdc20-dependent destruction sequence in both yeast and human cells, however, this does not impair the metaphase-anaphase transition [13,14,17]. Nevertheless, there is evidence that Sgo1 inactivation is important, since SGO1 overexpression results in a pronounced metaphase delay and a block to cohesin cleavage [18]. Since the Sgo1-induced metaphase delay is abrogated by deletion of BUB1, it is likely that Sgo1 must be localized at the pericentromere to prevent anaphase onset [18]. Accordingly, we showed that Sgo1 and its associated proteins are released from centromeres upon sister chromatid biorientation and tension [8]. Potentially, the temporal separation between Sgo1 removal from each pericentromere and its later degradation could ensure the reversibility of the mechanism that monitors biorientation right up until the moment that the commitment to anaphase is made. However, this model poses a conundrum: with the initiation of cohesin cleavage at anaphase onset, tension between sister kinetochores is lost, which could lead to re-activation of the error correction and biorientation pathways. What is more, it was unclear whether Sgo1-associated PP2A-Rts1 and CPC depart from centromeres simply by interacting with Sgo1, or by a more sophisticated mechanism involving finely orchestrated interplay between kinase and phosphatase activities.
Here, we identify Sgo1 SUMOylation as a mechanism that inactivates the pericentromeric signalling hub and thereby ensures timely anaphase onset. Small ubiquitin-like modifier (SUMO) is a 12 kDa protein that is covalently added to lysine residues of SUMO substrates. SUMOylation is carried out by the sequential activities of E1 activation enzyme, E2 conjugation enzyme and E3 ligase [19]. We isolated the E3 SUMO ligase, Siz2, as a negative regulator of Sgo1 through a genetic screen. We show that Sgo1 is SUMOylated in a manner dependent on Siz2 together with its paralog, Siz1, and that a failure to SUMOylate Sgo1 causes a delay to anaphase onset. We provide evidence that Sgo1 SUMOylation modulates its interaction with PP2A-Rts1 and facilitates the removal of CPC to stabilize sister chromatid biorientation, thereby enabling efficient entry into anaphase.
Results
SUMO ligases reverse the effects of SGO1 overexpression
To identify negative regulators of Sgo1, we screened for high copy suppressors of the poor growth caused by SGO1 overexpression [18]. We recovered a number of plasmids that improved the growth of cells carrying multiple copies of SGO1 under galactose-inducible control (pGAL-SGO1) (Figure S1A; Table S1), including one carrying a ∼5kb fragment containing truncated SLP1 and PUP1 together with full length ISN1 and SIZ2 (Figure 1A). SIZ2, encoding one of three budding yeast SUMO E3 ligases and sharing functional redundancy with its paralog, Siz1, is an attractive candidate for an Sgo1 antagonist, since Siz1/Siz2 have functions in chromosome segregation and cell division (Johnson and Gupta, 2001; Makhnevych et al., 2009; Montpetit et al., 2006; Takahashi et al., 2006). We used live cell imaging to confirm that overexpression of SIZ2 (by placement under control of the copper-inducible promoter, pCUP1-SIZ2) counteracts the delay in metaphase caused by SGO1 overexpression (Figure 1B). Metaphase duration, estimated as the time between spindle pole body separation (emergence of two Spc42-tdTomato foci) and anaphase onset (dispersal of Cdc14-GFP from the nucleolus), was markedly reduced in pGAL-SGO1 pCUP1-SIZ2 cells compared to pGAL-SGO1 cells (Figure 1C). These findings suggest that Siz1/Siz2 promote anaphase onset by counteracting Sgo1. Indeed, siz1Δ siz2Δ cells show a prolonged metaphase, measured as the time between formation of a short metaphase spindle (Tub1-YFP) and Cdc14-GFP dispersal (Figure 1D and E; [20]). Consistently, short metaphase spindles and securin (Pds1) persist in siz1Δ siz2Δ cells after synchronous release from a G1 arrest (Figure 1F and 1G). Deletion of SGO1 in siz1Δ siz2Δ cells partially rescued the metaphase delay, as judged by both decreased accumulation of short spindles and reduced Pds1 stabilization (Figure 1F and 1G), as did deletion of CDC55 (Figure S1B), previously shown to rescue the metaphase delay of pGAL-SGO1[18], further indicating that ectopic Sgo1 activity is at least partially responsible for the metaphase delay of cells lacking SIZ1 and SIZ2. We note, however, that the metaphase delay was more pronounced in sgo1Δ siz1Δ siz2Δ cells than in sgo1Δ cells, indicating that additional factors prevent timely anaphase onset in the absence of Siz1-Siz2. We conclude that Siz1/Siz2 promote anaphase onset, in part by antagonising Sgo1.
A chromatin-associated pool of Sgo1 is SUMOylated by Siz1 and/or Siz2
To determine whether Siz1/Siz2 might counteract Sgo1 by direct SUMOylation, we assayed Sgo1-SUMO conjugates in vivo. Cells carrying SGO1-6HA and a plasmid producing His-tagged yeast SUMO (7His-Smt3) were lysed under denaturing conditions and SUMOylated proteins were isolated using nickel affinity chromatography (Figure 2A). Anti-HA western blotting identified two slow-migrating bands corresponding to SUMOylated Sgo1 in wild type but not siz1Δ siz2Δ cells (Figure 2B). We further demonstrated direct SUMOylation of purified Sgo1 (Figure S2) in vitro by either Siz1 or Siz2 (Figure 2C).
Sgo1 is recruited to the pericentromeric chromatin during S phase, released into the nucleoplasm upon sister kinetochore biorientation at metaphase and degraded in anaphase [8]. Sgo1 SUMOylation was maximal prior to anaphase onset, reflecting Sgo1 abundance (Figure 2D). We assessed whether Sgo1 recruitment to pericentromeric chromatin is important for its SUMOylation. Loss of Bub1 or inactivation of its kinase activity which is required for phosphorylation of histone H2A-S121 and Sgo1 recruitment to pericentromeres [6, 21] greatly diminished Sgo1 SUMOylation (Figure 2E). Similarly, the sgo1-100 and sgo1-700 alleles, which carry point mutations that delocalize Sgo1 from pericentromeres [10], also markedly reduced Sgo1 SUMOylation (Figure 2E). To test whether Sgo1 removal from pericentromeres in response to tension coincides with loss of SUMOylation, we analysed cells arrested in metaphase (by depletion of CDC20) either in the presence (-tension) or absence (+tension) of microtubule depolymerising drugs. Indeed, spindle tension largely abrogated Sgo1 SUMOylation, though total Sgo1 levels were comparable to the no tension condition (Figure 2F). We conclude that Sgo1 SUMOylation is promoted by its association with pericentromeres.
Sgo1 SUMOylation requires its coiled-coil
If Sgo1 SUMOylation mediates the effect of Siz1/Siz2 in the metaphase-anaphase transition, a mutant that specifically abrogates Sgo1 SUMOylation, either by losing the SUMOylation sites, or by reducing Sgo1’s interaction with the E3 ligases, is expected to recapitulate siz1Δ siz2Δ’s metaphase delay. With the aim to map the SUMOylation sites, we performed mass spectrometry analysis of elutes from nickel affinity chromatography identified Lys124 as an in vivo SUMOylation site (Table S2). However, the sgo1-K124R mutant retained high levels of SUMOylation, suggesting the presence of additional SUMOylation sites (Figure 3A and B). As an alternative approach to mass spectrometry, we analysed a series of Sgo1 truncations (Figure S3A). SUMOylation was abolished in Sgo1-Δ2-208 and reduced in Sgo1-Δ2-108, suggesting that the N-terminal 108 amino acids are important for SUMOylation together with K124R (Figure S3B). Further analysis revealed robust SUMOylation of Sgo1-Δ2-40 and Sgo1-Δ2-40-K124R but not Sgo1-Δ2-108 K124R (Figure S3B), suggesting that SUMOylation requires amino acids 40-108. Interestingly, this region encompasses the coiled-coil domain of budding yeast Sgo1, which is known to directly bind PP2A-Rts1 and is also important for maintaining CPC at the centromeres (Verzijlbergen et al., 2014; Xu et al., 2009). We mutated the Lys residues in this region in several combinations (Figure 3A). SUMOylation was progressively reduced in the Sgo1-K56R K85R (Sgo1-2R), Sgo1-K56R K64R K70R K85R (Sgo1-4R) and Sgo1-K56R K64R K70R K85R K124R (Sgo1-5R) mutants (Figure 3B). The reduced SUMOylation of Sgo1-4R was recapitulated in the in vitro SUMOylation assay (Figure 3C). Therefore, lysines in Sgo1 coiled-coil enable its SUMOylation and may be direct targets of Siz1/Siz2. Moreover, mutation of these lysines specifically reduced SUMOylation in Sgo1 both in vivo and in vitro, providing a tool to study the effects of reduced Sgo1 SUMOylation in vivo.
Time course analysis and live cell imaging revealed a negligible or mild metaphase delay in the sgo1-K124R and sgo1-2R mutants, respectively (Figure S3C and D) and the sgo1-2R mutation did not exacerbate the metaphase delay of siz1Δ siz2Δ cells (Figure S3E). The sgo1-4R mutant showed a more pronounced metaphase delay (Figure 3D), but additional mutation of K124R (sgo1-5R) largely abrogated this delay (Figure S3F), suggesting adverse effects on Sgo1 protein function (see also Figure 5B below). Therefore, we focused on sgo1-4R for further analysis of the effects of reduced Sgo1 SUMOylation.
Sgo1 is not a direct target of Slx5-Slx8-mediated ubiquitination
How might SUMOylation of Sgo1 reverse its inhibitory effects on anaphase onset? Sgo1 was stabilized in cells lacking E3 ligases, Siz1 and Siz2 (Figure S1B), or where E2 Ubc9 function was impaired (Figure S4A). Cells lacking the kinetochore-associated Slx5/Slx8 SUMO-targeted ubiquitin ligase (STUbL) complex show increased centromeric PP2A-Rts1 [22] and slx5Δ cells exhibit a metaphase delay in which Sgo1 is stabilized (Figure 4A). This led us to test whether Sgo1-SUMO conjugates are targeted for proteosomal degradation by Slx5/8 [23]. We measured the half-life of Sgo1 protein levels after transient expression of ectopic pGAL-SGO1 in G1-arrested cells followed by quenching with glucose/cycloheximide (Figure S4B). This revealed a partial stabilization of Sgo1 in both siz1Δ siz2Δ and slx5Δ, compared to wild type cells (Figure S4C), indicating that Siz1/Siz2 and Slx5/Slx8 promote Sgo1 degradation.
To examine whether Sgo1 could be a direct target of Slx5/Slx8 we analysed ubiquitin conjugates in siz1Δ siz2Δ, slx5Δ and sgo1-4R cells. Since Sgo1 degradation also requires APC/C-dependent ubiquitination within a C-terminal degradation box on Sgo1 [14], we also analyzed mutants lacking this region (Δdb). Strikingly, Sgo1 ubiquitination was abolished in cells lacking the APC/C-dependent degradation box, but not detectably reduced in siz1Δ siz2Δ, slx5Δ or sgo1-4R cells, indicating that Sgo1 is unlikely to be a direct target of STUbL enzymes (Figure 4B). Consistently, Sgo1-SUMO conjugates do not accumulate in slx5Δ cells (Figure 4C) and the half-life of Sgo1-2R was not increased over wild type Sgo1 (Figure S4D). Therefore, Slx5/Slx8 regulates anaphase onset, but not by directly ubiquitinating Sgo1, and although Siz1/Siz2 and Slx5 affect Sgo1 stability, they do so indirectly, for example, by activating proteosome function [24].
Moreover, Sgo1 degradation appears dispensable for anaphase onset since sgo1-Δdb cells show no metaphase delay [14] and abolishing the APC/C recognition sites in Sgo1 did not exacerbate the metaphase delay of the sgo1-2R mutant (sgo1-2R Δdb, Figure S3D and S4E). Importantly, Sgo1-Δdb was SUMOylated to a similar extent to wild type Sgo1 (Figure 4C). Therefore, Sgo1 SUMOylation is able to promote anaphase onset even in the absence of APC/C-mediated degradation.
Sumoylation does not promote Sgo1 removal from chromatin under tension
Sgo1 is released from pericentromeres under tension [8] but whether this is critical for the metaphase-anaphase transition remained unclear. To address this, we asked if artificial tethering of Sgo1 to the kinetochore can prevent tension-dependent removal: GFP-binding protein (GBP) -tagged Sgo1 was produced from a galactose-inducible promoter (replacing endogenous Sgo1) in cells where the kinetochore protein Mtw1 was tagged with GFP (Mtw1-GFP) (Figure 4D). On its own, pGAL-SGO1-GBP expression caused a modest metaphase delay compared to the Mtw1-GFP control, presumably due to mild overexpression (Figure 4D). However, Sgo1-GBP expression in a strain producing Mtw1-GFP resulted in a severe delay in metaphase (Figure 4D). This delay required Sgo1 association with PP2A-Rts1 and/or CPC, because kinetochore tethering of the Sgo1-3A mutant protein, which has lost these interactions, resulted in a more modest metaphase delay (Figure 4D). Therefore, kinetochore-associated Sgo1 prevents anaphase onset in a manner depending on its ability to bind PP2A-Rts1 and/or CPC, showing that tension-dependent removal of Sgo1 is critical for anaphase entry.
Based on these findings, we considered that Siz1/Siz2 may promote anaphase entry by triggering the release of Sgo1 from chromosomes upon sister kinetochore biorientation. In wild type metaphase-arrested cells, chromatin immunoprecipitation followed by qPCR (ChIP-qPCR) showed that Sgo1 associates with a centromeric site in the absence, but not presence, of spindle tension and this pattern was largely unchanged in siz1Δ siz2Δ, sgo1-2R, sgo1-4R or sgo1-5R cells with reduced SUMOylation (Figure 4E and F; Figure S4F and G). We confirmed that SUMOylation is dispensable for the tension-dependent release of Sgo1 during anaphase by live cell imaging. In wild type cells, Sgo1-GFP first appeared as a bright focus which dissociated upon splitting of Mtw1-tdTomato foci at metaphase and this occurred with similar timing in siz1Δ siz2Δ cells (Figure 4G, Figure S4H). Instead, the observed delay in anaphase entry occurred after bulk Sgo1 removal (Figure 4G). Hence, although tension-dependent release of Sgo1 is critical for anaphase entry, this occurs independently of Sgo1 SUMOylation.
Sgo1 SUMOylation is required for stabilizing biorientation
Our findings indicate that SUMOylation neither targets Sgo1 for STUbL-mediated destruction nor does it facilitate Sgo1 removal under tension. To probe the mechanism connecting Sgo1 SUMOylation to timely anaphase onset, we visualized the efficiency of sister kinetochore biorientation in the Sgo1 SUMO mutants. We analysed the initial establishment of sister kinetochore biorientation by monitoring the separation of sister CEN4-GFP foci as spindles reformed after nocodazole wash-out, while maintaining a metaphase arrest (by depletion of Cdc20, Figure 5A). In contrast to sgo1-3A cells which have impaired biorientation, likely due to impaired CPC binding[10], all mutants with reduced Sgo1 SUMOylation except sgo1-5R (siz1Δ siz2Δ, sgo1-2R and sgo1-4R) showed proficient sister kinetochore biorientation (Figure 5B). Therefore, error correction and sister kinetochore biorientation pathways are functional in the absence of Sgo1 SUMOylation.
Next, we assessed the stability of biorientation in the SUMO mutants. Cells were released from nocodazole washout, and the separation of CEN4-GFP was monitored as cells progressed into anaphase. A single CEN4-GFP focus was observed initially, and two CEN4-GFP foci appeared upon attachment of sister kinetochores to microtubules from opposite poles. Stable attachment led to further separation of the two CEN4-GFP foci, which eventually segregated to opposite poles in anaphase (Figure 5C). In contrast, if attachments are unstable, the two CEN4-GFP foci reassociate prior to their splitting and segregation. In sgo1Δ and sgo1-3A mutants, which are defective in sensing and correcting attachment errors, the visualisation of two CEN4-GFP foci was delayed and the number of missegregation events was increased, but the levels of reassociation of two CEN4-GFP foci was similar to wild type (Figure 5D and E, Figure S5A and B). siz1Δ siz2Δ and sgo1-4R mutants, in stark contrast, were proficient in the initial establishment of biorientation and did not show increased missegregation (Figure 5D and E, Figure S5A and B). Instead, both mutants showed ∼15% increase in the number of cells in which the two CEN4-GFP foci reassociated, indicative of unstable biorientation. We conclude that Sgo1 SUMOylation is important to maintain the bioriented state.
The metaphase delay in siz1Δ siz2Δ is rescued by inactivating mutations in CPC/SAC
Unstable biorientation in SUMO-deficient mutants is expected to generate unattached kinetochores and engage the SAC, potentially explaining the metaphase delay of these cells. Consistent with this idea, deletion of MAD2 partially rescued the metaphase delay of siz1Δ siz2Δ cells (Figure S6A). The CPC-dependent error correction pathway is likely responsible for the instability of kinetochore-microtubule interactons in the SUMO mutants because inhibition of the CPC component, Ipl1 (ipl1-as1) also reduced the metaphase delay of siz1Δ siz2Δ cells (Figure 6A). Consistently, Sgo1 interaction with PP2A-Rts1 and/or CPC, is important for the metaphase delay in the absence of SUMOylation because both time course analysis (Figure S6B) and live cell imaging (Figure 6B) revealed that sgo1-3A siz1Δ siz2Δ cells spent less time in metaphase than siz1Δ siz2Δ cells. Interestingly, however, unlike Ipl1 (Figure 6A), the PP2A regulatory subunit, Rts1, was largely dispensible for the metaphase delay of siz1Δ siz2Δ cells (Figure S6C). Therefore, the CPC-dependent error correction pathway is responsible for the metaphase delay observed in the absence of Sgo1 SUMOylation.
Sgo1 SUMOylation promotes Ipl1 relocalization
The kinase activity of Ipl1 (Aurora B kinase) is required for error correction and Ipl1 is known to re-localize from centromeres to the spindle mid-zone upon the establishment of biorientation [25, 26]. Unstable biorientation in Sgo1 SUMOylation mutants suggested that this removal may be incomplete. We monitored Ipl1-GFP and its co-localization with Mtw1-tdTomato by imaging cells released from G1. In both siz1Δ siz2Δ and sgo1-4R, Ipl1 recruitment to the kinetochore-proximal regions occurred normally (Figure 6C). However, as kinetochores separated, Ipl1-GFP persisted close to kinetochores in the Sgo1 SUMO mutants (Figure 6C). Increased centromeric Ipl1 was also measured by ChIP in SUMO-deficient cells arrested in metaphase with kinetochores under tension (Figure S7A and B). Hence, Sgo1 SUMOylation facilitates the re-localization of Ipl1from centromeres to prevent persistent error correction and SAC activation.
Sgo1 SUMOylation is incompatible with PP2A-Rts1 binding
Interestingly, the coiled-coil domain of Sgo1 is both required for its SUMOylation (Figure 3) and for PP2A-Rts1 binding [10, 27]. This raised the question of whether Sgo1 SUMOylation also impacts PP2A-Rts1 binding. Sgo1 SUMOylation was increased in the Sgo1-3A mutant (Figure 7A), suggesting that PP2A-Rts1 binding normally dampens Sgo1 SUMOylation. Structural modelling of budding yeast Sgo1-PP2A-Rts1 interaction revealed that PP2A-Rts1 binding to Sgo1 would be incompatible with SUMOylation on these residues (Figure S7C). We used an in vitro binding assay to test the effects of Sgo1 SUMOylation on Rts1 binding. Purified Sgo1 was SUMOylated on beads in vitro, beads were stringently washed to remove components of the SUMO reaction and subsequently incubated with cell-free extract from sgo1Δ Rts1-9Myc cells. This revealed that, as expected, Rts1-9Myc bound robustly to unSUMOylated Sgo1, however Rts1 binding was greatly reduced by Sgo1 SUMOylation, consistent with the prediction that SUMOylation and PP2A-Rts1 binding are mutually exclusive (Figure 7B). Similarly, immunoprecipitated Rts1-9Myc bound in vitro SUMOylated Sgo1 less well than unSUMOylated Sgo1 (Figure 7C). Analysis of the SUMO-deficient Sgo1-4R protein showed that, unexpectedly, binding of Rts1 was reduced to a similar extent to Sgo1-3A, even in the absence of SUMOylation (Figure 7B). Despite the fact that both mutants fail to bind PP2A-Rts1, they result in very different outcomes in vivo: while sgo1-3A shows defective initial biorientation of sister kinetochores and chromosome mis-segregation after nocodazole washout, sgo1-4R does not (Figure 5B and E). Conversely, unstable sister kinetochore biorientation and a metaphase delay is observed in sgo1-4R but not sgo1-3A cells (Figure 3D, Figure 5D, Figure 6B, Figure S6B). This indicates that a failure to bind PP2A-Rts1 cannot be the primary cause of these defects. Instead, sister kinetochore biorientation and chromosome segregation defects in sgo1-3A cells are attributed to defective CPC maintenance at kinetochores [10] while, conversely, our data indicate that CPC persists at kinetochores in sgo1-4R cells causing unstable biorientation and a metaphase delay (Figure 6C). Collectively, these data indicate that the ability of Sgo1 to bind and release CPC underlies the establishment and stabilization of biorientation, respectively (Figure S7D).
Dissociation of shugoshin and PP2A-Rts1 stabilizes sister kinetochore biorientation
To understand the importance of the Sgo1-PP2A-Rts1 interaction in stabilizing sister kinetochore biorientation, we asked whether tethering of Rts1 to wild type Sgo1 or Sgo1-4R could rescue the instability of attachments. Although forcing Rts1 interaction with Sgo1-4R throughout the cell cycle did not affect initial biorientation (Figure 7D), this state was unstable, as judged by the increased number of cells switching between two and one CEN4-GFP dots (Figure 7E), though chromosome segregation was ultimately successful in the majority of cells (Figure 7F). This further confirms that loss of Rts1 binding is not the cause of unstable sister kinetochore biorientation in sgo1-4R cells. However, interestingly, forced interaction between Rts1 and wild type Sgo1 also resulted in frequent switches (Figure 7E). Therefore, release of the Sgo1-Rts1 interaction is important to stabilize bioriented sister kinetochore-microtubule attachments. Together with our finding that PP2A-Rts1 binding is incompatible with Sgo1 SUMOylation, this suggests that PP2A-Rts1 dissociation as a result of Sgo1 SUMOylation, is also important to stabilize biorientation.
Discussion
Identification of shugoshin regulators
Starting with an unbiased genetic screen we have identified SUMO ligases as negative regulators of the pericentromeric hub that responds to a lack of tension between kinetochores. Sgo1, the central pericentromeric adaptor protein is one key target of the Siz1/Siz2 SUMO ligases. Kinetochore-microtubule interactions are unstable in Sgo1-SUMO deficient cells (siz1Δ siz2Δ and sgo1-4R). Persistent cycles of kinetochore detachment and re-attachment engage the SAC, explaining why a failure to SUMOylate Sgo1 results in a metaphase delay. Consistently, we find that inactivation of components of the error correction pathway (Ipl1, Mad2) advanced anaphase timing in siz1Δ siz2Δ cells.
Sgo1 inactivation and removal from kinetochores is essential for timely anaphase entry ([18], Figure 4D) and here we have identified one mechanism that contributes to this inactivation. However, Sgo1 also prevents anaphase onset by inhibiting separase independently of securin [18]. PP2A-Cdc55 dependent dephosphorylation of separase and, potentially also cohesin itself is likely to underlie this inhibition [28, 29]. Notably, ZDS2, a negative regulator of PP2A-Cdc55 [30], was also isolated in our screen along with HOS3, the cohesin deacetylase [31–33] indicating that further mechanisms await discovery.
SUMOylation of Sgo1 ensures efficient entry into anaphase
How does Sgo1 SUMOylation regulate anaphase entry? We found that Sgo1 SUMOylation is dispensable for its tension-dependent release from pericentromeres and that, although SUMOylation promotes Sgo1 degradation indirectly, this is not required for efficient anaphase entry. Instead, our work suggested that Sgo1 SUMOylation likely promotes anaphase entry by silencing the error correction process, as biorientation was highly unstable in Sgo1 SUMO-deficient mutants (Figure 5D). Remarkably, we found that Ipl1 removal from kinetochores was incomplete in the Sgo1 SUMO mutants (Figure 6C), suggesting a key role of this modification in modulating the subcellular localization of Ipl1.
Meanwhile, we showed that Rts1 binds preferentially to unSUMOylated Sgo1, and that tethering Sgo1 to Rts1 destabilized biorientation in a similar way as the Sgo1 SUMO mutants. These findings suggest that Sgo1 SUMOylation-mediated Rts1 dissociation has an important role in stabilizing microtubule-kinetochore attachment. Interestingly, PP2A-B56 dampens the effects of Aurora B to allow initial attachments in human cells [34–36], suggesting a potential explanation for our observations. Overall, our findings indicate that SUMOylation modulates the kinase-phosphatase balance at the kinetochore to dampen CPC activity and allow initial kinetochore-microtubule attachments (Figure 7G). Different mutants affect this balance in distinct ways, leading to the observed outcomes on the establishment and stabilization of biorientation (Figure S7D).
SUMOylation – a generalized mechanism of centromere regulation with implications for disease?
Accumulating evidence indicates that SUMOylation might play a specific role at centromeres to fine-tune chromosome segregation. The SUMO isopeptidase Ulp2/Smt4 is important for maintenance of cohesion specifically at centromeres, in part through regulating Topoisomerase II [37]. The Pds5 subunit of cohesin is also known to prevent polySUMOylation of cohesin [38] and centromeric cohesin may be particularly susceptible since it lacks Pds5 [39]. PIAS SUMO ligases are known to be localized at centromeres in vertebrate mitotic cells and oocytes [40–42]. Moreover, the SUMO pathway is required to prevent cohesion loss during meiosis II, which centrally requires Sgo2-PP2A, and it is conceivable that modulation of the PP2A-Sgo1 interaction as we find in yeast underlies these observations in mouse oocytes [43]. Indeed, global studies found that shugoshins are SUMOylated in fission yeast and human cells, though the function has yet to be tested [44, 45]. Furthermore, there is ample evidence that CPC function may be subject to regulation by SUMOylation. Aurora B SUMOylation in Xenopus and human cells was shown to promote its enrichment at centromeres and it was proposed that this modification may serve as a reversible mechanism to dampen Aurora B kinase activity [46], while in C. elegans meiosis the localization of both Aurora B and Bub1 kinase are influenced by SUMOylation [47–49]. This suggests that SUMO may have a general role in CPC/error correction pathways and we speculate that multi-lateral SUMO-SIM interactions [50] enable the coordinated re-localization of surveillance factors. Notably, mis-regulation of the SUMO pathway is wide-spread in different cancers [51, 52]. Potentially, reductions in chromosome segregation fidelity caused by SUMO malfunction at centromeres, as we show here, could be a contributing factor in driving these malignant states.
Author contributions
Conceptualization, A.L.M., X.B.S., C.S., L.W., O.N., D.C., A.W. and Z.S.; Methodology, X.B.S, M.W., C.S., O.N., T.T. and R.H. Investigation: X.B.S., M.W., C.S., D.C., O.N., A.W., T.T. and Y.W. Software: D.A.K. Writing – original draft, X.B.S.. and A.L.M.; Writing – Review and Editing, all authors; Visualization, X.B.S., A.A.J. and A.L.M.; Supervision, A.L.M., R.H. and Z.S.; Funding acquisition, A.L.M.
Declaration of interests
The authors declare no competing interests.
Methods
Yeast strains and plasmids
Yeast strains are derivatives of W303 and are listed in Supplementary Table S3. Plasmids and primers are listed in Supplementary Tables S4 and S5, respectively. StuI-digested AMp1239 was transformed into a CDC14-GFP strain to make the YFP-TUB1 CDC14-GFP parent strain. Genes were deleted or tagged using PCR-based transformation. K-R mutant plasmids were generated using Quikchange II XL site-directed mutagenesis kit (Agilent), with primers listed in Table S5. K-R mutants were PCR amplified from the resulting plasmid using primers AM16 and AM3177 and were integrated into an sgo1Δ strain (AMy827). 7HIS-SMT3 and HIS-UBI plasmids were kind gifts from Dr. H. Ulrich.
Yeast growth and synchronization
Unless otherwise stated, yeast strains were grown in YEP supplemented with 2% glucose and 0.3 mM adenine (YPDA). For the benomyl sensitivity assays, plates were made by adding 10 μg/mL benomyl (Sigma) or the equivalent volume of DMSO (solvent control) to boiling media.
To synchronize cells in G1, overnight cultures were inoculated to OD600 = 0.2 and grown for 1 h at room temperature, before diluting back to OD600 = 0.2. α-factor was added to 5 μg/mL for 90 min and then re-added to 2.5 μg/mL for every 90 min, until > 95% cells exhibited shmooing morphology. To release cells from G1, α-factor was washed out using 10 × volume relevant media. For pMET-CDC20-containing strains, cells were arrested in G1 in methionine dropout medium. After α-factor wash-out, cells were released into YPDA (+ DMSO or + 30 μg/mL benomyl and 15 μg/mL nocodazole) + 8 mM methionine for 1 h. 4 mM methionine and DMSO/15 μg/mL nocodazole were re-added for 1 h.
Metaphase duration measurements by live cell imaging and mitotic time course
Synthetic complete/dropout media were used for growing and washing cells for live cell imaging. Cells released from G1-arrest were loaded onto μ-slide dishes (Ibidi) coated with concanavalin-A (Sigma). Images were taken at indicated time intervals using a Zeiss inverted microscope, in a temperature-controlled chamber (25°C for glucose-based media and 30°C for raffinose-based media). Movies were analyzed using the ImageJ software.
Mitotic time course experiments were performed at 25°C. To inhibit Ipl1-as5, NA-PP1 (Toronto Research Chemicals) was added to a final concentration of 50 μM upon the emergence of small-budded cells. For every time point, samples were either treated with 5% trichloroacetic acid (TCA) for protein extraction, or fixed in 3.7% formaldehyde for immunofluorescence. TCA-treated pellets were snap frozen in liquid nitrogen, washed in acetone and air-dried. Protein samples were prepared by bead beating and boiling in SDS sample buffer. For immunofluorescence, Fixed cells were spheroplasted using zymolyase (AMS Biotechnology) and glusulase (Perkin Elmer), fixed in methanol for 3 min, followed by 10 s incubation in acetone. Rat anti-α-tubulin (Abd Serotec) antibody was used at 1:50 and anti-Rat FITC (Jackson ImmunoResearch) was used at 1:16.7. Spindle morphology of 200 cells was analyzed for each time point.
Western blotting
Proteins were separated in 8% bis-tris acrylamide gels and were transferred to nitrocellulose membranes (except anti-ubiquitination blots, for which PVDF membranes were used). Membranes were blocked in 3-5% milk in phosphate-buffered saline + 0.1% Tween-20 (PBST) or tris-buffered saline + 0.1% Tween-20 (TBST). All antibodies were diluted in 2% milk PBST (except anti-ubiquitination blot, for which 2% milk TBST was used). Anti-c-Myc (9E10, Biolegend), anti-V5 (AbD Serotec), anti-Flag (M2, Sigma) and anti-HA (12CA5, Roche) antibodies were used at 1:1000 dilution. Anti-Pgk1 (lab stock) and anti-Kar2 (lab stock) loading controls were used at 1:10000 and 1:100,000 dilution, respectively. The non-quantitative blots were detected by ECL (Thermofisher) and autoradiograms. 20% Femto-ECL (Thermofisher) diluted in Pico-ECL was used to detect SUMOylated-Sgo1 signals. Quantitative blots were detected using a BioRad Chemidoc (Figure S4C) or a LiCOR Odyssey Clx machine (Figure S4D). Protein expression was quantified using ImageJ (Figure S4C) or ImageStudio (Figure S4D).
Analysis of in vivo SUMOylation
Cultures were inoculated to OD600 = 0.2 in 200 mL synthetic dropout media and grown for 4 h at room temperature. Equal OD of cells were collected for samples in the same experiment. Cell pellets were resuspended in 20 mL cold H2O and incubated with 3.2 mL solution containing 1.85 M sodium hydroxide and 7.5% β-mercaptoethanol. After 20 min incubation on ice, 1.65 mL 100% trichloroacetic acid was added and cells were incubated on ice for a further 20 min. Cell pellets were drop-frozen in liquid nitrogen, and subsequently lysed by bead-beating in 300 µL buffer A (6 M guanidine hydrochloride, 100 mM sodium phosphate buffer pH 8.0, 10 mM Tris-HCl pH 8.0). Lysate was diluted three-fold in buffer A and 10 µL was saved as input controls. Lysate was applied to a column packed with 600 μL 50% slurry Ni-NTA agarose beads (Qiagen), washed twice with buffer A + 0.05% Tween-20, twice with buffer B (8 M urea, 100 mM sodium phosphate buffer pH 6.8, 10 mM Tris-HCl pH 6.8) + 0.05% Tween-20 and once with buffer B + 0.05% Tween-20 + 20 mM imidazole. SUMOylated proteins were eluted by buffer B + 0.05% Tween-20 + 200 mM imidazole. Input and elute samples were concentrated by centrifugation using Vivaspin columns (Sartorius).
Mass spectrometry
A pGAL-SGO1 strain with 7HIS-SMT3 was inoculated to 0.2 OD in 2% raffinose media. After 3 h growth at room temperature, cells were diluted back to 0.2 OD and 2% galactose was added to induce SGO1 overexpression. Cells were harvested after 3 h and SUMOylated proteins were purified as described above. Proteins eluted from the Ni-NTA column were separated on a 4-12% NuPage Bis-Tris gel (Invitrogen) and the gel slice encompassing SUMOylated Sgo1 (between ∼100 kDa and 135 kDa, based on immunoblotting of a parallel gel) was excised for mass spectrometry analysis.
Cloning, expression and purification of recombinant Sgo1
Full-length wild type SGO1 was amplified from plasmid AMp899, to replace SMT3 in plasmid AMp773 by Gibson assembly using primers AM8849 to AM8852. V5 tag was inserted by Gibson assembly using plasmid AMp970 and primers AM8866 to AM8869 to generate N-terminal tobacco etch virus (TEV) protease cleavable Hisx7-V5 tag-tagged SGO1 (AMp1738) under the control of a pCUP1 promoter. sgo1-4R was PCR amplified from plasmid AMp1340 and ligated into AMp1738 by Gibson assembly using primers AM8850, AM8851, AM9124 and AM 9125 to generate N-terminal TEV protease cleavable Hisx7-V5 tag-tagged sgo1-4R (AMp1783).
A protease-deficient yeast strain (AMy8184) was transformed with the resulting plasmids (AMp1738 or AMp1783) and the transformants were inoculated into 8 L uracil dropout media. When OD600 reached 0.5 - 0.7, 0.5 mM CuSO4 was added to induce Hisx7-V5 -SGO1 (or sgo1-4R) expression. Cells were harvested 6 h after induction. Cell pellets were snap frozen in liquid nitrogen and ground to powder in a ball breaker machine (Retsch). All purification steps were performed at 4°C or on ice. Cell powder was resuspended in lysis buffer (25 mM Tris-HCl pH 7.5, 150 mM NaCl, 1 mM MgCl2, 10 % glycerol, 0.1% NP-40, 0.05 mM EDTA, 0.05 mM EGTA, 1 mM DTT, 1xCLAAPE, 1 mM Pefabloc, 0.4 mM Na Orthovanadate, 0.1 mM Microcystin, 1 mM NEM, 2 mM B-glycerophosphate, 1 mM Na pyrophosphate, 5 mM NaF, complete EDTA-free protease inhibitor (Roche)). Cell lysates were treated with 40 u/mL benzonase for 1.5 h. The crude lysate was diluted with 25 mM sodium phosphate buffer pH 7.5, 500 mM NaCl, 10 % glycerol, 0.1% NP-40, 10 mM imidazole, 1 mM DTT, 0.25 mM PMSF, cleared by ultracentrifugation (50,000 ×g at 4°C for 30 min), filtered through a 0.22 µm filter and loaded onto a HiTrap IMAC FF 1 ml column (GE Healthcare) charged with Ni2+ and attached to an AKTA system. The column was washed with 25 mM sodium phosphate buffer pH 7.5, 500 mM NaCl, 10 % glycerol, 0.1% NP-40, 25 mM imidazole, 1 mM DTT, 0.25 mM PMSF. Recombinant Sgo1 protein was eluted with an increasing imidazole gradient (25 - 500 mM) over 40 column volumes and then loaded onto a gel filtration Superose 6 10/300 column in 50 mM Tris-HCl pH 7.5, 500 mM NaCl, 10% glycerol, 0.1 mM EDTA, 0.1 mM EGTA, 1 mM DTT, 0.25 mM PMSF. Fractions containing the eluted protein were run on an SDS-PAGE gel and the purity of recombinant Sgo1 was assessed by InstantBlue (Expedeon) staining.
Cloning, expression and purification of of SUMOylation enzymes
SUMO E1 enzymes Uba2 and Aos1 (from vectors pET11a-UBA2 and pET28a-AOS1, [53]) were co-expressed in BL21 CodonPlus (DE3) RIL (Agilent Technologies) in the presence of 0.05 mM IPTG at 18 °C for 20 h. Expression of SUMO E2 enzyme Ubc9 and SUMO Smt3 (pET21b-UBC9 and pET21a-SMT3 [53]) was induced by 0.1 mM IPTG at 18 °C for 20 h in BL21 CodonPlus (DE3) RIL. E1, E2 and SUMO were purified using Ni-NTA agarose resin (Qiagen) with 20 mM sodium phosphate pH7.5, 500 mM NaCl, 10 to 200 mM imidazole, followed by Superdex 200 16/600 column (hand poured using prep grade resin from GE Healthcare) in 50 mM Tris-HCl pH 7.5, 150mM NaCl, 1mM DTT, 10% glycerol.
siz1-167-508 and siz2-130-490 were amplified from plasmids pGEX-4T-1-SIZ1-HF [53]) and AMp1528 (YEPlac112-SIZ2, this work), respectively. The respective PCR product was cloned into pMAL-c2X to generate N-terminal MBP-tagged siz1-167-508 (AMp1679) and N-terminal MBP-tagged siz2-130-490 (AMp1680). Siz1 (167-508) or Siz2 (130-490) were expressed in LB containing 10 μM ZnCl2, 0.1 mM IPTG and appropriate antibiotics at 16 °C for 20 h using BL21 CodonPlus (DE3) RIL (Agilent Technologies). Siz1 (167-508) or Siz2 (130-490) was purified from MBPTrap HP 1 mL column (GE Healthcare) with 50 mM Tris-HCl pH 7.5, 150 mM NaCl, 10 μM ZnCl2, 0 to 10 mM maltose gradient, followed by gel filtration with a Superdex 200 16/600 column (hand poured using prep grade resin from GE Healthcare) in 50 mM Tris-HCl pH 7.5, 150 mM NaCl, 10 μM ZnCl2, 10% glycerol.
In vitro SUMOylation assays and SUMOylated Sgo1 pull down assays
Purified Sgo1 (1 μM) was mixed with 5 mM ATP, 15 μM SUMO, 0.5 μM E1 (unless otherwise indicated), 0.5 μM E2 (unless otherwise indicated), 0.1 μM (unless otherwise indicated) Siz1 (167-508) or Siz2 in a reaction buffer consisting of 25 mM HEPES pH 7.5, 150 mM KCl, 10 mM MgCl2, 15 % glycerol, 0.1 % NP-40, 0.1 mM DTT, 0.25 mM PMSF. The reaction was incubated at 30 °C for 2 h (unless otherwise indicated). For detection of SUMOylated Sgo1, the reaction was boiled in SDS sample buffer before analyzed by anti-V5 western blotting.
For the binding assay, product of in vitro SUMOylation was incubated with 50 μL of anti-V5 coupled protein G magnetic dynabeads (Thermofisher) in Binding buffer A (25 mM HEPES pH 7.5, 150 mM KCl, 2 mM MgCl2, 15 % glycerol, 0.1% NP-40, 0.1 mM EDTA, 0.5 mM EGTA, 0.25 mM PMSF) for 2.5 h at 4 °C. After washing three times with Binding buffer A, beads were incubated with yeast lysate containing tagged Rts1 or Bir1 for 2 h at 4 °C. Beads were washed five times with binding buffer A + 10 mM NEM and were heated at 65 °C for 15 min to elute bound proteins.
Analysis of in vivo ubiquitination
Cultures were inoculated to OD600 = 0.2 in 100 mL 2% raffinose-containing dropout media and were arrested in G1 as described above. 2% galactose was added for 30 min to induce pGAL-SGO1-9MYC expression. Cells were treated, lysed in buffer A and diluted as described in in vivo SUMOylation assay. Lysates were incubated with Ni-NTA beads, 0.05% Tween-20 and 15 mM imidazole overnight at 4°C, with gentle rotation. Beads were washed twice with buffer A + 0.05% Tween-20, and four times with buffer C (8M urea, 100 mM sodium phosphate buffer pH 6.3, 10 mM Tris-HCl, pH 6.3) + 0.05% Tween-20. For elution, beads were heated at 60°C for 10 min in HU buffer (8M urea, 200 mM Tris-HCl pH 6.8, 1 mM EDTA, 5% (w/v) SDS, 0.1% (w/v) bromophenol blue, 1.5% (w/v) DTT).
Sgo1 half-life measurement
Cultures were inoculated to OD600 = 0.2 in YEP + 2% raffinose + 0.3 mM adenine. 5 μg/mL alpha-factor was added for 1.5 h and re-added to 2.5 μg/mL every hour until the end of the experiment. SGO1 overexpression was induced by the addition of 2% galactose for 30 min. De novo synthesis of Sgo1 protein was quenched by the addition of 2% glucose and 1 mg/mL cycloheximide (Acros Organics).
Co-immunoprecipitation
Cultures were inoculated to OD600 = 0.2 in 2 L YPDA and grown at 30 °C until OD600 reached 1-1.5. Cell pellets were harvested by centrifugation and drop-frozen in liquid nitrogen. Cells were ground, lysed and benzonase-treated as described in ‘Purification of recombinant protein’. After centrifugation at 3,600 rpm for 10 min, protein concentrations were measured by Bradford assay and approximately 85 mg of protein was incubated with 4 mg of epoxy beads (Thermofisher) coupled to IgG for 1.5 h at 4°C. Beads were washed four times with the lysis buffer and were heated at 70°C for 10 min in 50 μL 1 × LDS sample buffer + 2.5% β-mercaptoethanol. 5 μL of input and 20 μL of IP samples were separated on an 8% acrylamide gel and were transferred to PVDF membranes. Western blot was performed using TBST-based buffer.
Chromatin immunoprecipitation (ChIP)
Metaphase-arrested cells were fixed in 1.1% formaldehyde for at least 30 min. Cells were washed twice with TBS (20 mM Tris-HCl pH 7.5, 150 mM NaCl), once with FA lysis buffer (50 mM HEPES-KOH pH 7.5, 150 mM NaCl, 1 mM EDTA, 1% Triton-X, 0.1% sodium deoxycholate) + 0.1% SDS, and drop frozen in liquid nitrogen. Cells were lysed by bead beating in FA lysis buffer + 0.5% SDS + 1 mM PMSF + EDTA-free protease inhibitor cocktail (Roche). The resulting pellets were sonicated in a Bioruptor machine. Sheared chromatin was incubated with the relevant antibody and protein G dynabeads (Thermofisher) at 4°C overnight. Anti-HA (12CA5, Roche Diagnostics), anti-Flag M2 (Sigma), or anti-V5 (Abd Serotec) antibodies were used as relevant. Beads were washed once with FA lysis buffer + 0.1% SDS+ 275 mM NaCl, once with FA lysis buffer + 0.1% SDS + 500 mM NaCl, once with wash buffer (10 mM Tris-HCl pH 8, 0.25 M LiCl, 1 mM EDTA, 0.5% NP-40, 0.5% sodium deoxycholate) and once with TE (10 mM Tris-HCl, pH 8, 1 mM EDTA). Chromatin was eluted by boiling in the presence of 10% Chelex beads, treated with proteinase K (Life technologies) for 30 min and boiling for a further 10 min. Quantitative PCR was done using Express SybrGreenER (Thermofisher) (CEN4 and ARM4) or Luna (NEB) (all other chromosomal sites) on a Lightcycler machine (Roche). Primers used were listed in Table S5. ChIP enrichment was determined using the formula E-ΔCt, where ΔCt = (Ct(ChIP) – [Ct(input) – logE(Input dilution factor)] and E represents primer efficiency.
Biorientation assay in fixed and live cells
Cells carrying pMET-CDC20, SPC42-tdTOMATO and CEN4-GFP were arrested in metaphase in the presence of nocodazole and benomyl as described in ‘yeast growth and synchronization’.
For analyzing biorientation in metaphase-arrested cells (Figure 5C and 5D), drugs were washed out by filtering with 10 times the volume of YEP and cells were released into YPDA + 8 mM methionine. Samples were taken at the indicated time points, fixed in 3.7% formaldehyde for 9 min, washed in 80% ethanol and resuspended in 1 μg/mL DAPI in PBS. 200 cells were analyzed for each time point using fluorescent microscope.
For analyzing biorientation in cells going into anaphase (Figure 5E and 7F), nocodazole-arrested cells were loaded onto the ONIX Microfluidic Perfusion System (CellAsic) and visualized with a Zeiss inverted microscope coupled to an EMCCD camera at 25°C. Imaging started as soon as cells were released into methionine dropout media without drugs.
Supplementary information
Table S1. Complete list of high copy suppressors of GAL-SGO1 sickness identified in the screen shown in Figure S1A.
Table S2. (A) List of SUMOylated peptides identified in mass spectrometry (B) List of peptides identified in mass spectrometry
Table S3. Yeast strains used in this study
Table S4. Plasmids used in this study
Table S5. Oligonucleotides used in this study
Acknowledgments
We are grateful to Yoshiko Kikuchi and Helle Ulrich for yeast strains and plasmids, respectively, to Weronika Borek for help with R programming. We thank Lorraine Pillus, Emily Petty, Federico Pelisch and Andrew Goryachev for helpful discussions and comments on the manuscript. This work was funded by Wellcome through Senior Research Fellowships to AM [[107827]] and AAJ [202811], and core funding for the Wellcome Centre for Cell Biology [203149].
Footnotes
↵† Lead contact
References
- [1].↵
- [2].↵
- [3].↵
- [4].↵
- [5].↵
- [6].↵
- [7].
- [8].↵
- [9].↵
- [10].↵
- [11].↵
- [12].↵
- [13].↵
- [14].↵
- [15].↵
- [16].↵
- [17].↵
- [18].↵
- [19].↵
- [20].↵
- [21].↵
- [22].↵
- [23].↵
- [24].↵
- [25].↵
- [26].↵
- [27].↵
- [28].↵
- [29].↵
- [30].↵
- [31].↵
- [32].
- [33].↵
- [34].↵
- [35].
- [36].↵
- [37].↵
- [38].↵
- [39].↵
- [40].↵
- [41].
- [42].↵
- [43].↵
- [44].↵
- [45].↵
- [46].↵
- [47].↵
- [48].
- [49].↵
- [50].↵
- [51].↵
- [52].↵
- [53].↵
- [54].↵