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. Author manuscript; available in PMC: 2016 May 31.
Published in final edited form as: Cell Rep. 2016 Apr 28;15(6):1254–1265. doi: 10.1016/j.celrep.2016.04.017

Slx5/Slx8 promotes replication stress tolerance by facilitating mitotic progression

Yee Mon Thu 1, Susan Kaye Van Riper 2, LeeAnn Higgins 1, Tianji Zhang 1, Jordan Robert Becker 1, Todd William Markowski 1, Hai Dang Nguyen 1,#, Timothy Jon Griffin 1, Anja-Katrin Bielinsky 1,*
PMCID: PMC4864160  NIHMSID: NIHMS776746  PMID: 27134171

SUMMARY

Loss of minichromosome maintenance protein 10 (Mcm10) causes replication stress. We uncovered that S. cerevisiae mcm10-1 mutants rely on the E3 SUMO ligase Mms21 and the SUMO-targeted ubiquitin ligase complex Slx5/8 for survival. Using quantitative mass spectrometry, we identified changes in the SUMO proteome of mcm10-1 mutants and revealed candidates regulated by Slx5/8. Such candidates included subunits of the chromosome passenger complex (CPC), Bir1 and Sli15, known to facilitate spindle assembly checkpoint (SAC) activation. We show here that Slx5 counteracts SAC activation in mcm10-1 mutants under conditions of moderate replication stress. This coincides with the proteasomal degradation of sumoylated Bir1. Importantly, Slx5-dependent mitotic relief was not only triggered by Mcm10 deficiency but also by treatment with low doses of the alkylating drug methyl methanesulfonate. Based on these findings, we propose a model in which Slx5/8 allows for passage through mitosis when replication stress is tolerable.

INTRODUCTION

Replication stress is a condition in which lesions in the template strand or intrinsic defects in the replication machinery stall the progression of replication forks. Stalled forks unable to be reactivated will eventually collapse and cause chromosome breakage (Zeman and Cimprich, 2014). Minichromosome maintenance protein 10 (Mcm10), an evolutionally conserved replication factor, is a suppressor of chromosome breakage (Chattopadhyay and Bielinsky, 2007; Lukas et al., 2011). Mcm10 facilitates the activation of the replicative helicase and is required for origin unwinding. During the subsequent elongation step, Mcm10 is anchored to the Mcm2-7 complex, which comprises the core of the replicative Cdc45:Mcm2-7:GINS (CMG) helicase. It transiently interacts with DNA polymerase-α/primase and proliferating cell nuclear antigen (PCNA) both of which cycle on and off DNA during lagging strand synthesis (Thu and Bielinsky, 2014). Accordingly, depletion of Mcm10 compromises fork elongation, most notably at fragile sites of the human genome (Miotto et al., 2014).

A recent synthetic genetic array analysis (SGA) of mcm10-1, a temperature-sensitive mutant strain of Saccharomyces cerevisiae, revealed a strong negative genetic interaction between genes encoding the small ubiquitin-like modifier (SUMO)-targeted ubiquitin ligase (STUbL) complex Slx5/8 (Slx5/8; synthetically lethal with sgs1) and mcm10-1 (Thu and Bielinsky, 2013, 2014). The Slx5/8 heterodimer functions as an E3 ubiquitin ligase responsible for the turnover of poly-sumoylated proteins, and has been implicated in the maintenance of genome integrity (Galanty et al., 2012; Vyas et al., 2013; Xie et al., 2007; Yin et al., 2012). In yeast, the complex mediates nuclear-pore associated repair of broken forks (Nagai et al., 2008). In mammalian cells, RNF4 is responsible for the collapse of stalled replication forks and subsequent double-strand break (DSB) formation in order to allow for appropriate repair when checkpoint signaling is defective (Ragland et al., 2013). STUbL-dependent regulation of replication stress and DNA damage repair suggests that diverse sets of chromatin-bound proteins must be sumoylated under such conditions (Cremona et al., 2012; Psakhye and Jentsch, 2012; Wu et al., 2014).

S. cerevisiae expresses three SUMO E3 ligases, Siz1, Siz2 (SAP and mIZ-finger domain) and Mms21 (methyl methanesulfonate sensitivity 21), and two SUMO isopeptidases, Ulp1 and Ulp2 (UbL-specific protease) (Jentsch and Psakhye, 2013). Mms21-dependent sumoylation serves in enhancing fork stability when the replication machinery encounters obstacles (Branzei et al., 2006) and Mms21 targets are largely distinct from Siz1- and Siz2 substrates (Albuquerque et al., 2013; Reindle et al., 2006). Moreover, proteins that belong to the same molecular complex are often collectively sumoylated (Gibbs-Seymour et al., 2015; Hendriks et al., 2015; Psakhye and Jentsch, 2012; Sarangi and Zhao, 2015). One possible fate of SUMO conjugates is STUbL-mediated ubiquitination and degradation. To date, only a few in vivo substrates of Slx5/8 and RNF4 have been experimentally determined. Those include mediator of DNA damage checkpoint 1 (MDC1), centromere protein-I (CENP-I), Fanconi anemia complementation group D2 and I (FACND2/FACNI) and Jumonji/ARID1 B (JARID1B) in mammalian cells (Galanty et al., 2012; Gibbs-Seymour et al., 2015; Hendriks et al., 2015; Luo et al., 2012; Mukhopadhyay et al., 2010; Yin et al., 2012), and modifier of transcription 1 (Mot1), MATα2 repressor, topoisomerase 1 (Top1), Siz1 and mitotic chromosome determinant 1 (Mcd1) in yeast (D’Ambrosio and Lavoie, 2014; Steinacher et al., 2013; Wang and Prelich, 2009; Westerbeck et al., 2014; Xie et al., 2010).

In this study, we developed a quantitative mass spectrometry (MS) method named DRIPPER (Directed RIPPER) to explore the SUMO proteome in the context of Mcm10 deficiency-induced replication stress. RIPPER is an algorithm for intensity-based label-free peptide quantification (Van Riper et al., 2016). DRIPPER employs directed MS rather than the workhorse of mass spectrometry-based proteomic identification and quantification workflows, data dependent acquisition (DDA). Whereas DDA is a logical choice for many studies, DDA analyses are biased toward highly abundant proteins, because DDA automatically selects peptides with the highest intensities. This effectively limits the dynamic range of possible identifications and, consequently, low abundance proteins are often missed. To increase the dynamic range for the discovery of differentially abundant proteins, we turned to directed MS (Letarte et al., 2008; Schiess et al., 2009; Schmidt et al., 2008). DRIPPER departs from directed methods in the manner in which it decouples quantification and identification into two stages. This two-stage method selects only differentially abundant peptides from MS1 analyses on three technical replicates and generates a list of several thousand peptides for identification via directed MS/MS (more details in Supplemental Experimental Procedures). DRIPPER allowed us to perform unbiased quantification and identification of SUMO conjugates that were regulated differently under normal conditions or induced replication stress.

We chose mcm10 mutants as a genetic model for replication stress, since these cells are synthetically sick with deletions of SLX5 and SLX8 (Thu and Bielinsky, 2013, 2014). We analyzed the SUMO proteome and identified proteins potentially regulated by Slx5/8. Our data suggested that two subunits of the chromosomal passenger complex (CPC), Sli15 (synthetically lethal with ipl1) and Bir1 (baculoviral IAP repeat-containing protein 1) are affected by the Slx5/8 pathway in a manner consistent with promoting replication stress tolerance.

RESULTS

Mcm10 deficient cells require the STUbL complex Slx5/8 for optimal growth

The temperature-sensitive mcm10-1 allele encodes a P269L substitution, which renders the protein functional at 25°C, but unstable at 37°C (Homesley et al., 2000). Heat-induced depletion of Mcm10 causes replication stress displaying the typical hallmarks of Rad53 phosphorylation and PCNA ubiquitination (Becker et al., 2014). SGA analysis identified SLX5 and SLX8 as top hits that exhibited synthetic sickness with the mcm10-1 allele at 30°C (Thu and Bielinsky, 2013, 2014). We validated the SGA results in a different genetic background and found that an approximately 100-fold growth defect in mcm10-1 slx5Δ mutants was apparent at 33°C compared to either single mutant (Figure 1A). Tetrad dissection of diploid mcm10-1 slx5Δ and mcm10-1 slx8Δ strains confirmed the synthetic interaction (Figure S1A). The enhanced temperature sensitivity of mcm10-1 slx5Δ cells was fully reversed at 30°C by expressing wild-type SLX5 under the control of its endogenous promoter (Figure 1A). The functional RING domain of Slx5 was necessary to reverse the growth defect of mcm10-1 slx5Δ cells (Figure 1A), arguing that the catalytic function of Slx5/8 was required to confer resistance to replication stress (Xie et al., 2007). However, with increasing temperatures, SLX5 expression gradually lost the ability to rescue the viability of mcm10-1 slx5Δ mutants (Figure 1A), likely because the cells accumulated too much DNA damage (Becker et al., 2014). Re-expressing MCM10 under the control of its endogenous promoter fully rescued the growth defect of the double mutants at all temperatures (Figure 1B). Taken together, these results demonstrate that viability of mcm10-1 cells strongly depends on the activity of the Slx5/8 complex.

Figure 1. mcm10-1 mutants are synthetically sick with slx5Δ.

Figure 1

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(A, B) Successive 10-fold serial dilutions of indicated strains carrying an empty vector (EV), SLX5, CC561/564SS (RING mutant of SLX5) or MCM10 were grown on synthetic complete medium lacking uracil.

See also Figures S1.

mcm10-1 negatively interacts with mutations in SUMO pathways

Since Mcm10 deficient cells activate the Slx5/8 complex, we reasoned that poly-sumoylation must be crucial for mutant survival. To substantiate this idea, we mutated all lysine residues in SUMO to prevent the formation of poly-SUMO chains in the mcm10-1 strain. The double mutants exhibited a greater growth defect than either single mutant, underscoring the importance of poly-SUMO conjugates in mcm10-1 cells (Figure S1B). We also disrupted the functions of the E3 SUMO ligases Siz1, Siz2 and Mms21 in mcm10-1 mutants. As Siz1 and Siz2 are known to exhibit functional redundancy (Albuquerque et al., 2013; Reindle et al., 2006), we generated mcm10-1 siz1Δ siz2Δ triple mutants (Figures S1C and D). Mcm10 deficient cells were synthetically sick with mms21-CH, a mutant allele of MMS21, but not with siz1Δ or siz2Δ, either alone or in combination (Figures S1C–E).

Poly-sumoylation was crucial for mcm10-1 survival, however, we wondered whether aberrant accumulation of long SUMO chains negatively impacted cell growth. Interestingly, mcm10-1 mutants were sensitive to genetic ablation of the isopeptidase Ulp2 (Figure S1F). A similar genetic interaction has been observed for Ulp1 (Makhnevych et al., 2009). Based on our results, we concluded that the balance between poly-sumoylation and deconjugation of SUMO chains is necessary for mcm10-1 mutant survival.

Identification of SUMO conjugates in mcm10-1 cells by DRIPPER

To better understand the mechanistic role of SUMO in the context of replication stress, we set out to identify SUMO conjugates that were differently regulated in wild-type (WT) and Mcm10 deficient cells. We speculated that SUMO conjugates that exhibited a significant decrease in mcm10-1 cells might be potentially regulated by Slx5/8. To purify sumoylated proteins, we integrated a synthetic histidine-tagged SUMO gene (His8SUMO) at the endogenous SMT3 locus (Figure 2A). This minimizes the identification of false positive targets that may result from SUMO overexpression (Wohlschlegel et al., 2004). Control strains carried the untagged SUMO gene and a selectable marker at the SMT3 locus. Addition of the His8-tag did not affect SUMO expression, substrate conjugation, or temperature sensitivity of the mcm10-1 strain (Figures 2A and B). However, the sumoylation pattern in mcm10-1 mutants appeared different from that in WT cells when the strains were shifted to the restrictive temperature for 3 h (red bar in Figure 2B). To identify poly-SUMO conjugates, we purified His8-tagged proteins on cobalt resins under denaturing conditions. Cobalt was more efficient than nickel in enriching for poly-sumoylated proteins, the preferred substrates of the Slx5/8 heterodimer (Figure 2C) (Mullen and Brill, 2008). We observed established SUMO targets, such as Rfa1, Rfa2 (subunits of replication protein A, RPA) and PCNA using this approach (Figures 2D–F) (Cremona et al., 2012; Parker et al., 2008; Psakhye and Jentsch, 2012). In addition, we achieved a high degree of SUMO enrichment in His8-tagged samples (Figure S2A).

Figure 2. Cobalt affinity purification of SUMO conjugates.

Figure 2

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(A) Top: The cartoon illustrates the His8-tagged SUMO gene (SMT3) and the 3′ UTR integrated at the endogenous SMT3 locus. TRP1 (TRP) is a selection marker. Bottom: Indicated strains were grown on YPD plates in successive 10-fold serial dilutions. (B) SUMO patterns of whole cell extracts (WCEs) are shown for indicated strains. The red line indicates differences between WT and mcm10-1 mutants at 37°C. Tubulin was a loading control. (C) Sumoylated PCNA isolated by nickel or cobalt affinity purification from mcm10-1 mutants expressing SMT3 or HIS8SMT3 at 37°C. (D–F) Indicated strains were grown at 37°C. Eluates from cobalt affinity purification were analyzed with an anti-PCNA (D), -Rfa1 (E) or -Rfa2 (F) antibody. Free SUMO was used as a loading control.

See also Figure S2.

Purified SUMO conjugates from untagged and His8-tagged WT or mcm10-1 samples were separated on SDS-polyacrylamide gels and stained with Coomassie blue. The fraction above 75 kD showed signals highly specific to His8-tagged samples (bracket in Figure S2B). It was excised and analyzed via mass spectrometry using the DRIPPER methodology (Figure 3A). During the first stage of DRIPPER (quantification), we used MS1-only analysis on three technical replicates for each sample. We quantified peptide intensities by first extracting ion chromatograms (XICs) from MS1 spectra and then computing the XICs’ area under the curve using RIPPER (Van Riper et al., 2016). The result was an inclusion list of ~3000 candidate peptides fulfilling the following criteria: 1) peptides from His8SUMO samples that were not present in the untagged controls (intensity=0 in untagged WT and mcm10-1 samples); 2) peptides, which were differentially abundant in WT and mcm10-1 strains (either enriched or depleted in mcm10-1 samples) with a Student’s t-test p-value <0.001 (Figure 3A). During the second stage of DRIPPER (identification), we used the inclusion list to direct the selection of peptides for MS2. Finally, the results from DRIPPER’s quantification and identification were matched based on m/z value (±Δ 0.005 units) and retention time (±Δ 2 min). Because some peptide sequence matches overlapped in the m/z and retention time window, the final list was manually curated using m/z tolerances, retention drift observations, missing data observations, and statistical significance criteria to remove duplicate matches (details in Supplemental Experimental Procedures). Strong correlation (R2=0.99851) between retention times of MS1 and MS2 spectra indicated accurate peptide match (Figure S3A). In addition, 93% of the proteins were identified by multiple peptides, which behaved uniformly showing either an increase or decrease in mcm10-1 samples (Figure S3B). To compare the performance of DRIPPER to conventional DDA, we subjected the same biological samples to DDA. DRIPPER revealed several fold more unique peptides and almost twice as many proteins as DDA (Figures S3C and D). Moreover, DRIPPER identified more peptides per protein and more peptides in the low intensity range (Figures S3E and F).

Figure 3. Directed RIPPER (DRIPPER) reveals SUMO conjugates enriched or depleted in mcm10-1 cells.

Figure 3

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(A) DRIPPER separates peptide quantification (MS1) from identification (MS2). (B) The scatter plot illustrates the relative abundance of SUMO conjugates from the inclusion list. Relative intensities were determined by the ratios of peptide intensities from mcm10-1 to WT samples. Blue or red shaded areas represent arbitrary units (>8 = peptide intensities of 0 in WT, <−8 = peptide intensities of 0 in mcm10-1). Proteins from the same macromolecular complex or of similar function were depicted with matching symbols. Not all proteins are marked.

See also Figure S3 and Table S1.

Proteins in DNA damage response pathways and genome stability networks are differentially sumoylated in mcm10-1 mutants

Using DRIPPER, we identified a total of 96 sumoylated proteins that displayed a difference in abundance in mcm10-1 and WT cells (Figure 3B and Table S1). Among those, 77 had been reported to be sumoylated (Figure S4A and Table S2) (Albuquerque et al., 2013; Denison et al., 2005; Hannich et al., 2005; Panse et al., 2004; Wohlschlegel et al., 2004; Wykoff and O’Shea, 2005; Zhou et al., 2004). SUMO was present in both samples but slightly enriched in mcm10-1 cells (Figure 3B). We performed DRIPPER analysis of an independent biological replicate and observed that all, but four, proteins identified in both runs exhibited similar behaviors, i.e., they were either depleted or enriched in the mutant (Figure 4A and Table S1). Gene ontology (GO) analysis of all SUMO conjugates uncovered roles in transcription, DNA damage repair, DNA replication, cell cycle regulation, and chromosome segregation (Figure 4B and Table S3).

Figure 4. SUMO conjugates down-regulated in mcm10-1 mutants are potential substrates of the Slx5/8 complex.

Figure 4

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(A) The bar graph displays SUMO conjugates commonly identified by two independent DRIPPER analyses. (B) Gene ontology (GO) analysis was performed on SUMO targets identified by experiment 2. Top ten enriched GO terms are shown. (C) The Venn diagram presents overlap between SUMO targets identified in experiment 2 and potential Slx5/8 targets revealed in a study by Albuquerque et al. (Albuquerque et al., 2013). Listed proteins represent SUMO conjugates that were depleted (black) or enriched (red) in mcm10-1 cells.

See also Figure S4, Tables S2 and S3.

Sumoylated proteins highly enriched in mcm10-1 mutants included factors involved in homologous recombination (HR), such as Rad52, Rad59, Sgs1, Rfa1 and Rfa2 (Figure 3B). Accumulation of these SUMO conjugates had been reported in response to DNA damage (Branzei et al., 2006; Cremona et al., 2012; Psakhye and Jentsch, 2012). Enrichment of Rfa1, Rfa2 and PCNA in mcm10-1 samples was expected based on our initial western blots (Figures 2D–F). However, our proteomic screen also revealed chromatin-associated SUMO conjugates, such as Fob1, Top1, Tof2 and Sir4 that were underrepresented in mcm10-1 mutants compared to WT controls (Figures 3B and S4B). Other SUMO conjugates depleted in mcm10-1 mutants included CPC subunits, Bir1 and Sli15, and components of three different chromatin remodeling complexes, the Swr1 - (Vps72, Swc3 and Bdf1), Rsc - (Rsc2 and Rsc8) and SWI/SNF complex (Swi3) (Figure 3B). Identification of more than one subunit of a macromolecular complex (CPC, Swr1 - or Rsc complex) or multiple members of the same biological pathway (HR) is consistent with the notion that proteins that cooperate functionally are co-regulated by SUMO (Figure 3B) (Gibbs-Seymour et al., 2015; Hendriks et al., 2015; Psakhye and Jentsch, 2012).

Slx5/8 affects the abundance of sumoylated CPC subunits, Bir1 and Sli15, in mcm10-1 cells

We reasoned that the SUMO proteome of mcm10-1 cells likely contained factors regulated by the Slx5/8 pathway, and that their identification would provide clues as to how this STUbL promotes cell survival. To determine possible Slx5/8 substrates, we first overlapped our list of candidates with the list of potential Slx5 targets identified under normal growth conditions (Albuquerque et al., 2013). We determined that 20 proteins were present in both studies (Figure 4C). We performed a similar analysis with the list of potential Mms21 targets from the same study and identified 24 overlapping candidates (Figure S4C). These comparisons revealed SUMO conjugates that are potentially regulated by both Slx5 and Mms21 in mcm10-1 mutants (Figure S4D).

Candidates for Slx5/8 regulation included the CPC subunits, Bir1 and Sli15, and the Swr1 complex subunit, Vps72 (Figure 4C). We epitope-tagged (3HA-HIS8) the corresponding genes and induced Mcm10 depletion by temperature shift. Nickel affinity purification of tagged Bir1 under denaturing conditions revealed that the sumoylated form was less abundant when Mcm10 was degraded (Figure 5A), consistent with the proteomics data. Moreover, we found that the sumoylated form of the protein was enriched in mcm10-1 slx5Δ mutants compared to mcm10-1 cells (Figure 5A). Similarly, sumoylated Sli15 was more highly abundant in mcm10-1 slx5Δ than mcm10-1 mutants (Figure 5B). The levels of sumoylated Bir1 and Sli15 in mcm10-1 slx5Δ mutants were not quite as high as in WT cells (Figures 5A and B). This suggests that additional STUbLs or SUMO isopeptidases compensate for the loss of Slx5/8 activity in order to restrict the accumulation of poly-SUMO conjugates, which negatively affects mcm10-1 survival (Figures 1A and S1F). To assess whether Slx5/8 affects the steady-state levels of total Bir1 and Sli15, we examined whole cell extracts. Deletion of SLX5 caused only a slight enrichment in total protein (Figures S5A and B).

Figure 5. Slx5 destabilizes sumoylated Bir1 and Sli15 in mcm10-1 cells.

Figure 5

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Bir1 (A) and Sli15 (B) were purified from indicated strains grown at 35°C for 3 h. Eluates were immunoblotted with a SUMO- or HA-specific antibody. Equal amounts of total protein were loaded. The asterisk indicates a non-specific band. (C–E) Successive 10-fold serial dilutions of indicated strains were spotted on YPD plates and grown at different temperatures.

See also Figure S5.

In addition to CPC subunits, sumoylated Vps72 was diminished in mcm10-1 cells, independently confirming the results of our proteomic screen (Figure S5C). However, sumoylated Vps72 was not significantly increased in mcm10-1 slx5Δ cells, implying that Slx5/8 does not play a role in its regulation (Figure S5C).

Deletion of SLI15 and mitotic checkpoint genes supports survival of mcm10-1 mutants

One possible scenario of how Slx5/8 supports mcm10-1 survival is by interfering with CPC function. To test this idea genetically, we knocked out the non-essential CPC gene SLI15 in WT and mcm10-1 cells. Ablation of SLI15 partially rescued the growth defect resulting from Mcm10 depletion (Figure 5C). The CPC complex regulates chromosome bi-orientation during mitosis and participates in spindle assembly checkpoint (SAC) activation (Carmena et al., 2012). To understand whether CPC depletion affected mcm10-1 viability through its regulation of SAC, we knocked out MAD1 and MAD2, two genes involved in this pathway. Indeed, deletion of MAD1 or MAD2 rescued the growth defect of mcm10-1 cells (Figures 5D, E and S5D). More importantly, deletion of MAD2 partially rescued the temperature sensitivity of the mcm10-1 slx5Δ double mutants (Figure 5E). Together, our data suggested that one function of Slx5/8 is to reduce SAC activation, possibly by targeting sumoylated Sli15 and Bir1.

The Slx5/8 complex relieves mitotic arrest following moderate replication stress

To explore whether Slx5/8 relieves mitotic arrest, we examined cell cycle progression of mcm10-1 and mcm10-1 slx5Δ cells at elevated temperatures (Figures 6A and B). At 33°C, both mutants were initially delayed at G2/M phase. However, mcm10-1 cells with functional Slx5/8 eventually completed mitosis and entered the subsequent G1 phase of the cell cycle (Figure 6A). In contrast, mcm10-1 slx5Δ mutants exhibited a prolonged G2/M arrest (Figure 6A). To conclusively demonstrate that this arrest was due to SAC activation, we analyzed the status of Pds1. Pds1 is the homolog of securin in budding yeast and prevents sister chromatid separation. When the SAC is activated, Pds1 is stabilized until the checkpoint is turned off (Cohen-Fix et al., 1996). Consistent with the mitotic block, Pds1 was stabilized in mcm10-1 slx5Δ double mutants over the entire course of the experiment (8.5 h) (Figure S6A). In contrast, when the double mutants were released from a nocodazole block at 25°C, Pds1 was degraded with kinetics similar to WT cells (Figure S6B). These results argued that Slx5 suppresses SAC activation when cells encounter moderate replication stress.

Figure 6. Slx5 allows for mitotic progression when replication stress is moderate.

Figure 6

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(A, B) Asynchronous cultures of mcm10-1 and mcm10-1 slx5Δ mutants were grown at 33°C or 35°C. Samples were collected at indicated times and DNA content was analyzed by flow cytometry analysis (FACS). (C, D) Rad53 activation was monitored in samples shown in A and B. Tubulin was a loading control. Numbers represent the ratios of hyper-phosphorylated to unmodified Rad53. Red circles indicate unmodified Rad53. (E) Bir1 was purified from indicated strains grown at 33°C for 3 h treated with DMSO or MG132. Eluates were immunoblotted with a SUMO- or HA-specific antibody. Equal amounts of total Bir1 protein were loaded. (F) The experiment was performed as described in (E) and WCEs were immunoblotted with a HA-specific antibody. Tubulin was a loading control.

See also Figures S6 and S7.

At higher temperatures, however, when cells accumulated more severe DNA damage (Becker et al., 2014), the majority of mcm10-1 and mcm10-1 slx5Δ mutants remained arrested in G2/M (Figure 6B), consistent with a more robust Rad53 activation observed at 35°C compared to 33°C (Figures 6C and D). Hyper-phosphorylation of Rad53 observed at 35°C in mcm10-1 mutants was slightly diminished when SLX5 was knocked out, implying that Slx5 contributes to checkpoint signaling under extreme DNA damage conditions (Figure 6D).

Sumoylated Bir1 is subject to replication stress-induced proteasomal degradation

Our results suggested that Slx5 activity, which primes sumoylated proteins for proteasomal degradation, shows the strongest effect on mcm10-1 survival at the semi-permissive temperature of 33°C (Figures 1A, 6A and B). Thus, we examined sumoylated Bir1 at 33°C in the presence or absence of the proteasome inhibitor MG132. Whereas the treatment had insignificant effects in WT cells, it resulted in a substantial accumulation of sumoylated Bir1 in mcm10-1 cells (Figure 6E). When we examined total Bir1 levels in whole cell extracts, the protein was stabilized in both WT and mcm10-1 cells upon treatment with MG132 (Figure 6F). We interpreted this to mean that the proteasomal degradation of sumoylated Bir1 was linked to replication stress, as it only occurred in mcm10-1 mutants (Figure 6E). We also attempted to detect ubiquitinated Bir1 in the presence of MG132. Unfortunately, these experiments were inconclusive (data not shown).

Replication stress contributes to depletion of sumoylated Bir1 in mcm10-1 cells

Because replication stress induces accumulation of mcm10-1 cells in G2/M and this coincides with depletion of sumoylated CPC subunits (Figures 5A, B, and 6A, B), we tested whether sumoylated Bir1 was intrinsically unstable in G2/M under unperturbed conditions. WT cells were synchronized in G1, released and collected in G2/M (Figure S7A). We found that the amount of sumoylated Bir1 in asynchronous and G2/M cells was similar (Figure S7B). Thus, the drastic decrease of sumoylated Bir1 in mcm10-1 mutants under semi-permissive conditions was linked to replication stress and not an artifact of cell cycle arrest (Figure 5A).

Slx5/8-mediated mitotic escape is not unique to Mcm10 deficiency

To determine whether Slx5/8 allows for mitotic escape under a different form of replication stress, we treated WT and slx5Δ cells with different concentrations of methyl methanesulfonate (MMS) (Figures 7A and S7C, D). Rad53 activation in response to 0.003% MMS was moderate, similar to mcm10-1 cells at 33°C (Figures 6C and 7B). Under these conditions, slx5Δ mutants were delayed in traversing through G2/M phase compared to WT (Figure 7A), consistent with the prolonged retention of Pds1 (Figure S7E). We observed a similar effect with the lower dose of MMS (0.0015%), however at a higher concentration (0.01%), significant differences between WT and slx5Δ mutants were no longer detectable (Figures S7C and D). Therefore, Slx5/8 regulation of mitotic escape in the presence of tolerable replication stress was not restricted to Mcm10 deficiency.

Figure 7. Slx5/8 promotes replication stress tolerance.

Figure 7

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(A, B) Asynchronous cultures of WT and slx5Δ mutants were treated with 0.003% MMS. Samples were collected at indicated times for FACS analysis (A) or Rad53-specific immunoblots (B). Tubulin was a loading control. Numbers represent the ratios of hyper-phosphorylated to unmodified Rad53. Red circles indicate unmodified Rad53. (C) Model for a role of Slx5/8 in mitotic progression under conditions of high or moderate replication stress. Replication stress causes exposure of RPA-coated ssDNA. Moderate stress triggers low-level Rad53 activation, allowing cells to progress through S phase. Checkpoint activation may also promote the activity of Mms21 (not shown), resulting in sumoylation of chromatin-associated proteins. Under these circumstances, Slx5/8 regulation of CPC, composed of Bir1, Sli15, Ipl1 (increase in ploidy 1) kinase, Nbl1 (N-terminal-Borealin like protein 1), promotes escape from mitotic arrest. When stress levels are high, robust hyperactivation of Rad53 inhibits S phase progression. In this case, Slx5/8 dependent regulation of CPC has no effect.

DISCUSSION

We demonstrate here that Slx5/8 and the SUMO network play crucial roles in promoting replication stress tolerance. To profile changes in sumoylation upon debilitating normal replication fork progression, we developed a quantitative MS method called DRIPPER. This technique accurately determined the relative abundance of differentially sumoylated proteins in mcm10-1 mutants and WT cells, and provided a list of candidate targets of the Slx5/8 pathway.

DRIPPER improves some of the shortcomings of DDA-based methods

Our findings suggest that the label-free quantitative method DRIPPER can be a cost-effective alternative to widely used DDA-based methods. Here, we highlight a few advantages. First, DRIPPER is inherently unbiased since quantification of peptides precedes the identification step. Second, unlike DDA, DRIPPER is an efficient tool to identify differentially abundant proteins and bypasses the need for labeling amino acids in culture, which can be costly. Third, DRIPPER is not restrained by the number of available isotopes or chemical groups for peptide labeling. Most importantly, DRIPPER does not automatically couple chromatographic MS and peptide identification. Thus, it is not restricted to identifying the most abundant peptides but provides a wider dynamic range. In support of this notion, we detected a higher number of total peptides and peptides of low abundance by DRIPPER than by DDA (Figures S3C and F).

SUMO regulates the genome stability network of Mcm10 deficient cells

Our study extends previous reports, which determined changes in sumoylation induced by DNA damaging agents (Cremona et al., 2012; Psakhye and Jentsch, 2012). Consistent with the above studies, we found that sumoylation of HR factors (Rad52 and Rad59) and Rpa subunits (Rfa1 and Rfa2) was upregulated upon Mcm10 depletion (Figure 3B). However, since Siz2, the SUMO ligase that targets these factors (Psakhye and Jentsch, 2012), has a negligible role in mcm10-1 mutants (Figure S1C), their sumoylation might merely be a by-product of replication stress.

Our proteomics results overlap considerably with an analysis by the Zhou laboratory, which identified potential targets of Mms21 and Slx5/8 under unperturbed conditions (Figures 4C and S4C, D). This observation agrees with our findings that both Mms21 and Slx5/8 contribute to the survival of mcm10-1 mutants (Figures 1A and S1A, E). Interestingly, Mcm10 depleted cells depend on Mms21 but did not rely on the other two SUMO E3 ligases, Siz1 and Siz2 (Figures S1C–E). Mms21 has been reported to target proteins localized to centromeres and kinetochores, including Bir1 (Yong-Gonzales et al., 2012), but needs to be further corroborated. Thus, Mms21 may support mcm10-1 survival directly through sumoylation of Bir1. Interestingly, phosphorylation of Mms21 by the Mec1 checkpoint kinase stimulates its SUMO ligase activity (Carlborg et al., 2015). Since Rad53, a downstream target of Mec1, is activated in mcm10-1 cells, checkpoint signaling may promote Mms21-dependent sumoylation of chromatin-associated proteins and relay the stress signal from replication forks to other chromosome organizing regions, such as centromeres (Figure 7C).

Sumoylated CPC subunits are subject to Slx5/8 regulation and proteasomal degradation in Mcm10 deficient cells

Changes in the SUMO proteome prompted us to explore potential Slx5/8 targets and their roles in mcm10-1 mutants. We found that sumoylated Bir1 and - Sli15 were enriched in the absence of Slx5 (Figures 5A and B). Although this effect may be indirect, this observation raises the possibility that targeting sumoylated CPC subunits is one mechanism by which Slx5/8 promotes growth of Mcm10 deficient cells. How might disruption of CPC function assist mcm10-1 mutant cell survival? In early mitosis, the CPC is targeted to the centromere-kinetochore interface, and this localization is mediated by Bir1 (Cho and Harrison, 2012; Kawashima et al., 2010; Yoon and Carbon, 1999). Despite its interaction with kinetochore proteins, CPC is not an anchor for the kinetochore (Buvelot et al., 2003; Tanaka et al., 2002), but required to sense the tension generated between sister chromatids once they are attached to opposite spindle poles (Sandall et al., 2006). A lack of tension will activate the SAC (Biggins and Murray, 2001; Carmena et al., 2012; Shimogawa et al., 2009). These signaling events are responsible for delaying mitosis until chromosome bi-orientation is achieved. In addition to misoriented chromosomes, lack of replication can ablate the tension between sister chromatids (Stern and Murray, 2001). Moreover, aberrant replication intermediates at telomeres or the rDNA locus arrest cells at prometaphase in a SAC-dependent manner (Nakano et al., 2014), arguing that incomplete replication at diverse genomic regions can trigger a mitotic delay.

Our data shows that Slx5/8 regulates sumoylated Bir1 (Figure 5A), which is subject to proteasomal degradation when cells undergo replication stress (Figure 6E). Under the same conditions, Slx5 counteracts SAC activation (Figure S6A). Together, these findings are consistent with the model that targeted degradation of sumoylated Bir1 by Slx5/8 disrupts CPC function and allows for mitotic entry of mcm10-1 mutants (Figure 7C). This is also in agreement with the observation that inactivation of the SAC by MAD2 deletion partially rescued the loss of SLX5 (Figure 5E). Our model predicts that sumoylated Bir1 functions at the kinetochore, which is proximal to the site of Slx5 localization (Montpetit et al., 2006; Mukhopadhyay et al., 2010; Sun et al., 2007; van de Pasch et al., 2013). It would then be plausible that degradation of sumoylated Bir1 counteracts SAC activation (Figure 7C).

Slx5/8 promotes mitotic progression in the face of moderate replication stress

The ability of Slx5/8 to rescue mcm10-1 cells is clearly temperature dependent (Figure 1A). This suggested that targeted degradation of SUMO substrates by this STUbL is effective when replication stress is moderate, but ineffective when stress conditions become severe. We utilized the level of Rad53 phosphorylation to gauge the degree of inflicted stress. Our data is consistent with a model in which persistent, but low-level Rad53 activation allows cells to progress through the cell cycle until they temporarily accumulate at G2/M. In the presence of Slx5/8 cells pass through mitosis (Figure 6A), but in its absence, this process is greatly delayed (Figures 6B). In contrast, under more severe replication stress, Rad53 is hyper-activated and triggers robust G2/M arrest (Figures 6D and 7C). In this case, the function of Slx5/8 is inconsequential. Importantly, Slx5/8 dependent mitotic relief is not unique to mcm10-1 mutants, as we observed a similar effect when we treated cells with a low dose of MMS (Figure 7A). The idea that cells enter mitosis with a genome that triggers detectable Rad53 activation and, therefore, must have RPA-coated single-stranded regions or DNA breaks is supported by previous reports. Budding yeast sic1 mutants enter mitosis while they are still replicating their genome and mammalian cells that harbor replication intermediates or unresolved DNA structures – including those resulting from Mcm10 deficiency – progress into G1 phase with 53BP1 foci (Chan et al., 2009; Harrigan et al., 2011; Lengronne and Schwob, 2002; Lukas et al., 2011). Collectively, these data imply that cellular mechanisms have evolved to ensure escape from mitotic arrest when the integrity of the genome can be restored in the next generation. Our data makes it conceivable that Slx5/8 actively participates in this process.

EXPERIMENTAL PROCEDURES

Construction of yeast strains

Yeast strains used in this study are isogenic derivatives of W303-1a and the relevant genotypes are shown in Table S4. Details on construction of knockout and epitope-tagged strains are described in Supplemental Experimental Procedures.

Purification of His8SUMO conjugates for mass spectrometry analysis

Half a liter of asynchronous wild-type and mcm10-1 strains expressing SMT3 or His8SMT3 from the endogenous promoter were grown in YPD to log-phase at 25°C and shifted to 37°C for 3 h in pre-warmed YPD. Proteins from these samples were extracted using NaOH/β-mercaptoethanol and then precipitated by 50% TCA. Protein pellets were resuspended in urea buffer and 50 mg of total protein were incubated with Talon Metal Affinity Resin (Clontech) for 2 h at RT in a gravity flow column (Bio-Rad). Detailed wash and elution conditions are described in Supplemental Experimental Procedures.

Label-free quantification of peptides from mass spectrometry analysis

After in-gel trypsin digestion, we analyzed peptide mixtures by capillary liquid chromatography-mass spectrometry (LC-MS) on an Eksigent 1D plus LC with a MicroAS autosampler (Dublin) online with an Orbitrap Velos MS system (Thermo Fisher Scientific, Inc.). We analyzed four sample types by 1D LC-MS for each quantification measurement consecutively on the same analytical column. The four sample types were: sample 1) WT expressing untagged SUMO; sample 2) WT expressing His8SUMO; sample 3) mcm10-1 mutants expressing untagged SUMO; sample 4) mcm10-1 mutants expressing His8SUMO. The quantitative analysis strategy included the following sample sets: A) triplicate injections of samples 1 to 4 analyzed in random order in MS1 (survey scan) only; B) directed MS/MS with inclusion lists for analytes differentially quantified between samples 2 and 4 that were undetected in control samples 1 and 3. Details of the trypsin digestion, LC-MS/MS, generetaion of an inclusion list, database searching, matching quantification and identification runs, optimization of directed MS runs and DDA analysis are in Supplemental Experimental Procedures. The proteomics data have been deposited to the ProteomeXchange Consortium (Vizcaino et al., 2014) via the PRIDE partner repository (PRIDE: PXD002607).

GO analysis and prediction of potential Slx5 or Mms21 targets

GO analysis was performed using Saccharomyces Genome Database Gene Ontology Slim Mapper. Potential Slx5 or Mms21 targets were predicted using the study by Albuquerque et al. (Albuquerque et al., 2013). Details are described in Supplemental Experimental Procedures.

Additional experimental procedures

Methods on construction of yeast strains, temperature shift experiments, western blotting, purification of SUMO targets, cell cycle analysis, assessment of Pds1 degradation, serial dilution assays and tetrad dissection are described in Supplemental Experimental Procedures.

Supplementary Material

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Acknowledgments

We thank members of the Bielinsky laboratory for helpful discussions and Eric Hendrickson for critical reading of the manuscript. We also thank B. Stillman, X. Zhao, J.F.X. Diffley, G. Brush, M. Hochstrasser, and D. Moazed for generously sharing their reagents. We wish to acknowledge the University of Minnesota Flow Cytometry Resource and the Minnesota Supercomputing Institute. This work was supported by NIH grant GM074917 (AKB) and partly through a scholarship by the Leukemia & Lymphoma Society LLS1023-09 (AKB). The authors recognize the Center for Mass Spectrometry and Proteomics at the University of Minnesota.

Footnotes

AUTHOR CONTRIBUTIONS

Conceptualization, A.K.B. and Y.T.; Methodology, Y.T., L.H., S.K.V.R. and A.K.B.; Software, S.K.V.R.; Validation, Y.T. and J.R.B.; Formal Analysis, Y.T., S.K.V.R., L.H. and A.K.B.; Investigation, Y.T., S.K.V.R., L.H., T.W.M., J.R.B., T.J. and H.D.N.; Data Curation, S.K.V.R. and L.H.; Writing – Original Draft, Y.T. and A.K.B.; Writing – Reviewing and Editing, Y.T., S.K.V.R., L.H., T.W.M., T.J.G., A.K.B.; Visualization, Y.T., S.K.V.R. and J.R.B.; Supervision, A.K.B. and T.J.G.; Project Administration, A.K.B. and Y.T.; Funding Acquisition, A.K.B.

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NIHMS776746-supplement-1.xlsx (275.8KB, xlsx)
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