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The Late S-Phase Transcription Factor Hcm1 Is Regulated through Phosphorylation by the Cell Wall Integrity Checkpoint Takahiro Negishi,a Jiri Veis,b David Hollenstein,b Mizuho Sekiya,a Gustav Ammerer,b,c Yoshikazu Ohyaa Department of Integrated Biosciences, Graduate School of Frontier Sciences, University of Tokyo, Chiba, Japana; Department of Biochemistry, Max F. Perutz Laboratories, University of Vienna, Vienna, Austriab; Christian Doppler Laboratory for Proteome Analysis, University of Vienna, Vienna, Austriac

The cell wall integrity (CWI) checkpoint in the budding yeast Saccharomyces cerevisiae coordinates cell wall construction and cell cycle progression. In this study, we showed that the regulation of Hcm1, a late-S-phase transcription factor, arrests the cell cycle via the cell wall integrity checkpoint. Although the HCM1 mRNA level remained unaffected when the cell wall integrity checkpoint was induced, the protein level decreased. The overproduction of Hcm1 resulted in the failure of the cell wall integrity checkpoint. We identified 39 Hcm1 phosphorylation sites, including 26 novel sites, by tandem mass spectrometry analysis. A mutational analysis revealed that phosphorylation of Hcm1 at S61, S65, and S66 is required for the proper onset of the cell wall integrity checkpoint by regulating the timely decrease in its protein level. Hyperactivation of the CWI mitogen-activated protein kinase (MAPK) signaling pathway significantly reduced the Hcm1 protein level, and the deletion of CWI MAPK Slt2 resulted in a failure to decrease Hcm1 protein levels in response to stress, suggesting that phosphorylation is regulated by CWI MAPK. In conclusion, we suggest that Hcm1 is regulated negatively by the cell wall integrity checkpoint through timely phosphorylation and degradation under stress to properly control budding yeast proliferation.

C

ritical events during cell cycle progression, such as DNA replication and mitosis, are regulated by cell cycle checkpoints to ensure the proper completion of cell division. The cell cycle in the budding yeast Saccharomyces cerevisiae is coordinated with bud growth to secure the morphological space for cell division (1). One of the cell cycle checkpoints required to monitor budding morphology is the morphogenesis checkpoint (2), and the other is the cell wall integrity (CWI) checkpoint (3). They function independently to coordinate bud growth and cell wall construction, respectively, with cell cycle progression (4). Without a supply of cell wall materials for bud growth, the budding yeast cell cycle is arrested before entry into M phase and cannot proceed to mitosis (3). Compared to the morphogenesis checkpoint that regulates Swe1 phosphorylation and its degradation to activate cyclin-dependent kinase (CDK) (5), the primary function of the cell wall integrity checkpoint is inhibiting cell cycle progression to mitosis, which is achieved essentially by downregulating M-phase cyclin CLB2 (3, 4). During functions of the active cell wall integrity checkpoint, cells accumulate at the small budded state. Budding starts as cells proceed to S phase; thus, the checkpoint must be activated during S phase to prevent later cell cycle events. However, the manner by which the regulatory signaling cascade induced by the cell wall integrity checkpoint feeds into the complex transcriptional network of cell cycle progression has not been fully investigated. Genome-wide transcriptional studies have revealed the transcription cascade of cell cycle-related transcription factors (6–9). Studies have shown major transcription factors regulating each other at specific cell cycle stages, such as Ace2/Swi5 (M- to G1-phase transition), Swi4/Swi6 and Swi6/Mbp1 (late G1- to Sphase transition), Hcm1 (late S phase), and Ndd1/Fkh2 (M phase). The only transcription activator that functions in the network during S phase is Hcm1, suggesting that the cell wall integrity checkpoint regulates Hcm1 to subsequently inhibit M-phase cyclin CLB2 through the transcription network. HCM1 is one of the four budding yeast genes homologous to

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the Forkhead family of eukaryotic transcription factors (10). Hcm1 expression is regulated by Swi4/Swi6 (Swi4/Swi6 cell cycle box binding factor; SBF complex) and Swi6/Mbp1 (MluI cell cycle box binding factor; MBF complex) (11, 12). It takes the role of a transcription activator for a number of downstream genes with important functions, such as spindle pole body (SPB) assembly, spindle dynamics, chromosome segregation, and budding (12– 14). In addition, Hcm1 induces the expression of the M-phasespecific transcription factors Fkh2 and Ndd1 (14), which are part of the mitotic cyclin CLB2 transcription complex (15), supporting the notion that it is a part of the upstream effectors controlling mitotic cyclin at the cell wall integrity checkpoint. Landry et al. (16) reported predicted phosphorylation sites for positive and negative functions of Hcm1 through a mutagenesis analysis. Interestingly, the loss of potential N- and C-terminal phosphorylation sites revealed negative (degradation) and positive (transcriptional activation) Hcm1 regulation, respectively, via phosphorylation, suggesting that Hcm1 activity is coordinated by phosphorylation through the cell cycle. However, the actual phosphorylation sites and possible in vivo kinases that may regulate Hcm1 have not been fully investigated. In this study, we investigated the mechanism by which the cell wall integrity checkpoint controls Hcm1. We identified 39 in vivo

Received 18 October 2015 Returned for modification 11 November 2015 Accepted 24 December 2015 Accepted manuscript posted online 4 January 2016 Citation Negishi T, Veis J, Hollenstein D, Sekiya M, Ammerer G, Ohya Y. 2016. The late S-phase transcription factor Hcm1 is regulated through phosphorylation by the cell wall integrity checkpoint. Mol Cell Biol 36:941–953. doi:10.1128/MCB.00952-15. Address correspondence to Yoshikazu Ohya, [email protected]. Supplemental material for this article may be found at http://dx.doi.org/10.1128 /MCB.00952-15. Copyright © 2016, American Society for Microbiology. All Rights Reserved.

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phosphorylation sites, including 26 novel sites, using tandem mass spectrometry (MS/MS) analysis of affinity-purified Hcm1. A mutagenesis analysis of in vivo Hcm1 phosphorylation sites revealed the role of the protein during the cell wall integrity checkpoint. We show that three phosphorylation sites, S61, S65, and S66, are required for proper functioning of the cell wall integrity checkpoint. Finally, the phosphorylation-induced timely degradation of the protein when the checkpoint was activated and the involvement of mitogen-activated protein kinase (MAPK) activity during degradation is described. MATERIALS AND METHODS Yeast strains, plasmids, oligonucleotides, and culture media. The strains, plasmids, and oligonucleotides used in this study are listed in Tables S3 to S5 in the supplemental material, respectively (3, 17–22). Cells were grown in standard YPD (1% Bacto yeast extract [BD Biosciences, San Diego, CA], 2% polypeptone [Wako Chemicals, Osaka, Japan], 2% glucose [Wako Chemicals]), YPGS (1% Bacto yeast extract [BD Biosciences], 2% Bacto peptone [BD Biosciences], 0.1% sucrose [Wako Chemicals], and 2% galactose [Wako Chemicals]), and SD medium (0.67% yeast nitrogen base without amino acids [BD Biosciences] and 2% glucose), appropriately supplemented to maintain plasmids. Solid medium was prepared by adding 2% agar powder (Shoei, Tokyo, Japan). All strains were derivatives of YPH499 and YPH500 (23), with the exception of YOC4915, which was a w303 derivative. Gene disruption, transformation, and plasmid construction. HCM1 deletion strains were generated by the PCR-mediated gene disruption method using the Candida glabrata URA3 (Cg-URA3) gene (20). CgURA3 was amplified with the appropriate pair of primers (no. 3122 and 3123) with flanking sequences derived from the upstream and downstream regions of the corresponding HCM1 gene using pYO2241 as the template, and it was transformed into YOC1001 or YOC1087 to generate YOC4876 and YOC4877, respectively. YOC4879 and YOC5058 were transformed with pYO326 and pYO2382 to construct YOC5086, YOC5087, YOC5088, and YOC5089. pYO3057 (pWR168) was used with a primer pair (no. 3290 and 3291) to construct YOC4911 and YOC4915, as described in reference 17. pYO3030 was generated by removing the 2␮ region in pYES/NT (Invitrogen, Carlsbad, CA) (3), and the HCM1 genome was cloned into the pYO3030 BamHI site to generate pYO3046. YOC4462 then was constructed by transforming StuI-linearized pYO3046 to integrate Gal1pHCM1 into the URA3 locus. The HCM1 genome was tagged with 3⫻ hemagglutinin (HA) using one-step HA tagging to generate pYO3105, as described previously (19), with the pYO1150 plasmid and a primer pair (no. 3169 and 3170). The tagged HCM1-HA was amplified using a primer pair (no. 3216 and 3217) that amplified ⫾500 bp of the HCM1-HA open reading frame attaching the HindIII sites for cloning into the vector system. The amplified fragment was cloned into pYO2899 (pUC19) at the HindIII site, yielding pYO3105 for subsequent site-directed mutagenesis. YOC5096 and 5097 were constructed according to Longtine et al. (24) using the appropriate primer pair (no. 3122 and 3123). Site-directed mutagenesis. High-fidelity PCR using KOD⫹ polymerase and the DpnI digest was performed according to the manufacturer’s protocol (Toyobo, Tokyo, Japan) using pYO3105 as the template plasmid and primer pairs 3276-3277, 3278-3279, 3280-3281, 3282-3283, and 3284-3285 to yield pYO3106, pYO3194, pYO3107, pYO3195, and pYO3108, respectively. The 3286-3287 primer pair was used to generate the S65A_S66A mutation to construct pYO3196, and the 3288-3289 primer pair was used to generate S61A in addition to S65A_S66A. Each clone sequence was verified by sequencing analysis. After the alleles were constructed as pUC19-based plasmids, about 5 ␮g of each plasmid was digested with HindIII and transformed into YOC4876 or YOC4877, and the Cg-URA3 replacement with the wild-type or mutant HCM1-HA allele was selected using plates supplemented with 5-fluoroorotic acid (Toronto

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Research Chemicals, Toronto, ON, Canada). The use of pYO3105, pYO3106, pYO3194, pYO3107, pYO3108, and pYO3196 yielded YOC4878, YOC4879, YOC4839, YOC5084, YOC4896, YOC5085, YOC4841, YOC5057, and YOC5058, respectively. Transformants later were checked by PCR and sequencing. Cell synchronization. Cell synchronization by centrifugal elutriation was described by Suzuki et al. (3). Briefly, a 1-liter culture of cells was prepared at 1 ⫻ 107 cells/ml and 25°C. Cells were collected by centrifugation, and the cell pellet was resuspended in 50 ml of sterile water and sonicated briefly. The cells then were applied to an elutriation device equipped with an elutriation rotor (R5E; Hitachi, CA), and the pump speed was adjusted so that the sterile water flow rate was ⬃18 to 25 ml/ min. About 300-ml volumes of G1 cells were collected in tubes placed on ice. Typically, about 1 ⫻ 109 cells (about 10% of input) or fewer were acquired. Cells then were collected by centrifugation and released into fresh medium at the specified temperature or condition. For acute heat treatment at 37°C, a half volume of prewarmed YPD at 50°C was added to YPD at room temperature. Evaluation of bud index and observation of spindle morphology by immunofluorescence microscopy. Cell samples were collected and fixed with 3.6 to 3.8% (vol/vol) formaldehyde (Wako Chemicals) by gentle shaking for 30 min. Cells then were collected by centrifugation, resuspended in 1 to 2 ml of 10% phosphate-buffered saline (PBS; TaKaRa Bio Inc.), and stored for no more than 3 days. The bud index of cell samples under the indicated conditions or at the indicated time points was evaluated by counting and categorizing cells (n ⬎ 200) into “no,” “small,” and “medium-to-large” buds, where a small bud was defined as cells possessing a bud size of less than about one-third the size of the mother and medium-to-large buds as cells with a bud size of more than about one-third the size of the mother. Spindle morphology was visualized by indirect immunofluorescence. For tubulin staining, anti-tubulin 1/34 (antitubulin rat IgG2a monoclonal antibody, 1:100 dilution) and Alexa Fluor 564-labeled goat anti-rat IgG and IgM (1:100 dilution) antibodies were used. Simultaneous 4=,6-diamidino-2-phenylindole (DAPI) staining was used to visualize DNA. Cells showing bipolar spindles were counted, and the percentage in the population was determined (n ⬎ 200) under the indicated conditions or at the indicated time points. Cells were observed using an Axio Imager M1 with a 100⫻ Plan-Apochromat with an oil-immersion objective lens (Carl Zeiss, Germany) equipped with a CoolSNAP HQ cooled charge-coupled device camera (Roper Scientific, AZ) and AxioVision software (Carl Zeiss). RNA extraction and Northern blotting. RNA extraction and Northern blotting were performed as described previously (3). The primers used to construct the probes for visualizing each mRNA transcript are listed in Table S5 in the supplemental material (YHP1, no. 3185-3186; CIN8, no. 3210-3211; DSN1, no. 3206-3207; SPC34, no. 3208-3209; WHI5, no. 3204-3205; FKH1, no. 3187-3188; NDD1, no. 3176-3178; FKH2, no. 3193-3194; HCM1, no. 3120-3121; and ACT1, no. 3274-3275). Liquid chromatography-tandem mass spectrometry (LC-MS/MS) analysis for the identification of Hcm1 phosphorylation sites. Protein purification and mass spectrometric analysis were based on methods described in reference 25, with the modifications described below. Purified proteins were digested using trypsin (recombinant, proteomics grade; 5%, wt/wt, of the estimated protein amount; Roche). The digest was performed overnight at 37°C. MS analysis was performed using a reversed-phase nano-high-performance liquid chromatograph (HPLC) (U3000 or U3000 nanoRSLC; Thermo Fisher Scientific) directly coupled to an LTQ OrbitrapVelos or Q Exactive Plus mass spectrometer (Thermo Fisher Scientific) via a nanoelectrospray ion source (Thermo Fisher Scientific). A total of 17 HPLC-MS/MS runs were analyzed. MS-GF⫹ (26) (v. 9979), Comet (27) (v. 2014010), and X! Tandem (28) (2013.09.01) were used for peptide spectrum matching. Searches were performed against the SGD database (6,717 entries; database was last updated for the search performed on 3 February 2011) plus contaminants.

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FIG 1 Overexpression of Hcm1 induces a defective cell wall integrity checkpoint. (A) Hcm1 overexpression induces a defective cell wall integrity checkpoint. Cells of the indicated strains (YOC1087 and YOC4462) were synchronized at G1 phase by elutriation and released in YPD or YPGS medium at 37°C. The inhibition of bud growth (left) and bipolar spindle formation (right) was quantified at the indicated time points (n ⬎ 200 for each time point). (B) Representative images of data from panel A 240 min after release. Arrows indicate the bipolar spindle.

A decoy search was performed against the reversed database and used for false discovery rate (FDR) calculations. A precursor ion mass tolerance of 10 ppm was applied for all search engines. If possible, a fragment ion mass tolerance of 0.8 Da for CID data sets or 0.05 Da for HCD data sets was allowed. Up to four of the following variable modifications per peptide were allowed: phosphorylation of serine, threonine, and tyrosine; deamidation of asparagine and glutamine; oxidation of methionine; and protein N-terminal methionine removal and acetylation. In addition, the carbamidomethylation of cysteine was used as a fixed modification. An in-house Python tool (utilizing the pyteomics library v2.4.3 [29]) was used for further data processing. Peptide spectrum matches of individual search engines were combined and ambiguous identifications removed, resulting in an overall FDR below 1%. Phosphosite localization probabilities were calculated using the Java standalone version of phosphoRS (30). A phosphosite probability of ⱖ75% was considered confidently localized. A more detailed description of individual search settings and data processing is available upon request. Cycloheximide chase assay. Cells were grown to mid-log phase and were harvested and released in fresh YPD medium at a concentration of 5 ⫻ 106 cells/ml. Cells were allowed to grow for 2 h prior to cycloheximide (Wako) treatment. Cycloheximide stock solution was prepared at 10 mg/ml in water and was applied at a final concentration of 100 ␮g/ml in YPD. Cells were cultured and collected at the indicated time points and were frozen until their downstream application. Protein extraction, phosphatase treatment, electrophoresis, Western blotting, and quantification. Protein extracts were prepared for gel electrophoresis. Phos-tag gel electrophoresis was conducted as described previously (31). Briefly, ⬃2 ⫻ 108 to 4 ⫻ 108 cells for each sample or time point were used for extraction. Cells were collected by centrifugation and immediately frozen in liquid nitrogen prior to protein extraction. The concentration and amount applied were adjusted prior to electrophoresis using the bicinchoninic acid method (BCA protein assay kit; Pierce, IL) so that each lane contained ⬃50 ␮g of protein. For phosphatase treatment, calf intestinal phosphatase (10 U/␮l; TaKaRa, Japan) was used, and 0.2 U was added to about 50 ␮g of the protein extract and left at 4°C overnight. Electrophoresis gels were prepared using the standard method with an acrylamide concentration of 10%. Transfer was conducted using a standard wet method at 400 mA for 2 h on ice. For HA and Myc detection, the membrane was blocked with 5% milk, followed by the application of the primary 11MO anti-HA (Covance) or 9E10 anti-Myc (Calbiochem, Darmstadt, Germany) antibody, respectively, and incubation overnight at 4°C in 5% milk at a concentration of 1:1,000 in Tris-buffered saline with

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Tween 20 (TBS-T; pH 7.75). For Cdc28 detection, the membrane was blocked with 5% bovine serum albumin (BSA), followed by incubation with the primary PSTAIRE antibody (Santa Cruz Biotechnology, TX) overnight at 4°C in 5% milk at a concentration of 1:5,000 in TBS-T (pH 7.75). The Phos-tag gel was prepared as described in the manufacturer’s protocol (Wako) by adjusting the Phos-tag concentration to 27 ␮M in a 7.5% acrylamide separation gel. Before transfer to the polyvinylidene difluoride (PVDF) membrane, it was critical to incubate the gels in 1 mmol/ liter EDTA in the transfer buffer for 10 min, wash them with 20% ethanol for 5 min, and then wash them with the transfer buffer without EDTA for 10 min. Transfer was conducted using a standard wet method at 400 mA for 2 h on ice, and the membrane was blocked for 1 h at room temperature (RT) in 3% milk, followed by incubation with the primary antibody 11MO (Covance) overnight at 4°C in 3% milk at a concentration of 1:1,000 in TBS-T (pH 7.75). For secondary antibodies, anti-mouse antibody– horseradish peroxidase (HRP) and anti-rabbit antibody–HRP (Invitrogen) were used at a concentration of 1:2,500 in 5% BSA and TBS-T at RT for 1 h for HA and Cdc28, respectively. The ECL prime system (GE Healthcare) combined with LAS1000 (Fujifilm, Japan) was used for detection. Quantification of total, phospho-, and dephospho-Hcm1-HA was carried out as follows. First, the Hcm1-HA level was normalized to that of Cdc28 (control) by quantifying the blots using Image Gauge software (Fuji Film). The untagged background was set as 0, and the highest level of Hcm1 was set to 1.0. The amounts of phospho- and dephosphoHcm1-HA then were quantified from the results of the Phos-tag gel, and the proportion of the previously determined Hcm1-HA level was calculated.

RESULTS

The cell wall integrity checkpoint regulates Hcm1 protein level to inhibit cell cycle progression. The temperature-sensitive FKS1 mutant (catalytic subunit of 1,3-␤-glucan synthase [32]) fks11154 loses its synthase activity at the restrictive temperature of 37°C due to an impaired supply of cell wall material to growing buds (33). As a consequence, fks1-1154 cells undergo budding cycle arrest with no or small-budded cells and, more interestingly, with duplicated and adjacent SPBs and replicated DNA, which is described as the cell wall integrity cell cycle checkpoint (3). Similar cell cycle arrest phenotypes have been reported in other cell wall mutants, such as glycosylphosphatidylinositol-anchored membrane protein mutants (Dcw1 and Dfg5 [34]) and a mannosyl

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FIG 2 Late-S-phase transcription factor Hcm1 is downregulated by the cell wall integrity checkpoint. (A) Hcm1 protein level was examined during the activation of the cell wall integrity checkpoint. HA-tagged Hcm1 from FKS1 (YOC4878) and fks1-1154 (YOC4879) cells were synchronized to G1 phase by elutriation, grown at 37°C, and subjected to Western blotting in a time course manner together with the asynchronized untagged controls (YOC1001 and YOC1087). (B) Quantification of Hcm1 level normalized to that of the Cdc28 loading control from panel A. AU, arbitrary units. (C) Hcm1 protein level was regulated similarly in FKS1 (YOC4878) and fks1-1154 (YOC4879) strains at the permissive temperature of 25°C. HA-tagged Hcm1 from FKS1 (YOC4878) and fks1-1154 (YOC4879) cells were prepared as described for panel A and were cultured at 25°C, followed by Western blotting in a time course manner together with untagged controls (YOC1001 and YOC1087). (D) Quantification of Hcm1 level normalized to that of the Cdc28 loading control from panel C. (E) HCM1 transcription was examined during functions of the active cell wall integrity checkpoint. Cells of the indicated strains (YOC1001 and YOC1087) were synchronized to G1 phase by elutriation and grown at 37°C, followed by Northern blotting in a time course manner. The ACT1 transcript level is shown as the loading control. (F) Hcm1-regulated gene expression, including the key transcription factors NDD1 and FKH2, was delayed or downregulated by activation of the cell wall integrity checkpoint. Cells were prepared as described for panel E. The expression of genes reported to be regulated by Hcm1 in the genome-wide transcription study of Pramila et al. (14) was examined while the cell wall integrity checkpoint was active.

transferase (MNN10) deletion mutant (3). We first tested whether the overproduction of Hcm1 affected the cell cycle arrest phenotype of fks1-1154 to investigate the role of Hcm1 in the checkpoint. For this purpose, Hcm1 was produced under the control of a galactose-inducible GAL1 promoter. Budding and spindle morphology were examined by microscopy following synchronization of the cells at G1 phase by centrifugal elutriation and release in galactose (Gal)- or glucose (Dex)-containing medium. Bud growth did not occur after release regardless of the medium, because glucan synthase activity was perturbed (Fig. 1A, left, and B). The bipolar spindle did not form when Hcm1 was not overexpressed (i.e., in Dex medium or when Hcm1 was not expressed under the control of the GAL1 promoter) (Fig. 1A, right, and B), indicating a functional checkpoint. In contrast, when Hcm1 was overexpressed under the GAL1 promoter in galactose medium,

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bipolar spindles formed and accumulated starting 120 min after release (Fig. 1A, right, and B), indicating a defective checkpoint. Similar to findings of a previous report on the G2/M-phase Forkhead transcription factor Fkh2 (3), overexpression of the late-Sphase transcription factor Hcm1 resulted in a defective checkpoint. We next determined whether the Hcm1 protein level and HCM1 transcription level are regulated when the cell wall integrity checkpoint is activated. We found that the protein level was downregulated under the checkpoint-activated condition (fks1-1154; at 37°C) to ⱕ50% of that of the wild type at 60 and 120 min after release from G1 phase (Fig. 2A and B). The Hcm1 protein level was not downregulated under the checkpoint-uninduced condition (Fig. 2C and D). However, the HCM1 transcript level was not affected by the cell wall integrity checkpoint, peaking around the

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151 ASWQNSIRHN LSLNDAFIKT EKSCDGKGHF WEVRPGAETK FFKGENRGYE 201 FVKDSLQDIG KYFEIDSTLD ELEQVESGEG NDDLPDEEER EEAGKFPSIE 251 IQLNSSPILR VSQLHHIPQL KTDNSVLNPH ENLESMRNMI ENDVNNIDSL

Hcm1p-HA

301 EPPYVMKKYH TSLGLPSLVN AKDHFQAGVK NNNITQANRF NTLPITSAKS 351 PQNFRKYFTS FNSNFEDLSP LRSNVGAGSL LDPLPYSPLK LYDQKNLALM

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401 SKPQSQQSYS NSQLPPPPSS HGSDLLKTPK MRHSDGLEKT PSRLISTPKD 451 GNSILRKWQT PSHLFEDLYC SPLFRAIETP IRYITTPGGT LETQISPRKS

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501 SAPDVLTSAT NSKFASSGLF GVDVYSVWKR ATEKISDGNN TTDSNQKHHP 551 YHNHPSNDSG NEKN*

This Study (39)

ISPB Phosphopep DB (6)

D

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Albuquerque et al. 2008 (5)

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In vivo P+ site (T/S) [26] In vivo P+ Proline-directed site (T/S-P) [12] In vivo P+ Tyrosine site (Y) [5]

CDK preferred site (T/S-P-x-K/R) [5] MAPK preferred site (P-x-T/S-P) [2] DNA binding domain

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3 Soufi et al. 2009 (8)

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FIG 3 Hcm1 is a phosphoprotein. (A) Cells harboring HA-tagged (YOC4878) or untagged Hcm1 (YOC1001) were subjected to protein extraction following phosphatase treatment with or without inhibitors. The protein extracts were subjected to SDS-PAGE supplemented with the Phos-tag or without supplement. Western blotting was performed using anti-HA (11MO) antibody for Hcm1-HA and anti-Cdc2/p32 (PSTAIRE) antibody for Cdc28/Pho85 as loading controls. (B) The 39 in vivo serine, threonine, and tyrosine phosphorylation sites (in red) identified in this study in reference to the Hcm1 primary amino acid sequence (see Table S2 in the supplemental material). Red boldface S and T (serine and threonine) show proline-directed amino acid residues. Underlining shows the DNA-binding domain (from SGD or Superfamily 1.73, HMM server). (C) The phosphorylation sites identified in this study were compared to those reported previously from systematic studies and the ISPB PhosphoPep database (see Table S2). Numbers in parentheses indicate the number of sites identified in each study. (D) A schematic model is shown for Hcm1 with the DNA-binding domain and the phosphorylation sites investigated in this study. In addition to the sites identified in panel B, the in vivo phosphorylation sites identified in the ISPB phosphopep database and in Soufi et al. are listed, and the color keys indicate the characteristics of each site (41). Some of the phosphorylation sites are preferred by CDK and MAPK (44).

same time under the checkpoint-activated condition (60 to 120 min after release from G1) (Fig. 2E). Taken together, these results indicate that Hcm1 is downregulated only at the protein level. Hcm1 is a transcription factor of NDD1 and FKH2 (14), comprising the G2/M-phase transcription complex (Mcm1-Fkh2Ndd1 complex) (15). To determine whether the G2/M-phase transcription complex is inhibited upon checkpoint activation, we analyzed NDD1 and FKH2 transcript levels in the fks1-1154 mutant. Northern blot analysis showed that NDD1 and FKH2 transcription were reduced when the checkpoint was activated (Fig. 2F). Similar to FKH2 and NDD1, the transcription of a number of other HCM-cluster genes, such as YHP1, CIN8, DSN1, SPC34, WHI5, and FKH1, was downregulated or delayed under the checkpoint-activated condition (Fig. 2F), further supporting the notion that Hcm1 is functionally inhibited when the checkpoint is active. Affinity-purified tandem mass spectrometry analysis of Hcm1 reveals 29 phosphorylation sites in vivo. Cell cycle-related transcription factors often are regulated by phosphorylation (15, 35–38), and Hcm1 is a suggested CDK substrate (39). We first demonstrated that Hcm1 is phosphorylated in vivo using Phos-tag gel electrophoresis. Cells harboring HA-tagged Hcm1 were grown to the mid-log phase and subjected to protein extraction with or without phosphatase and a phosphatase inhibitor, followed by gel electrophoresis and Western blot immunodetection (Fig. 3A).

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When Phos-tag was incorporated into the gel during electrophoresis, Hcm1 retarded its mobility in the gel, which collapsed completely when treated with phosphatase (Fig. 3A). As the slowermigrating bands were restored in the presence of a phosphatase inhibitor, it was evident that phosphorylated Hcm1 was successfully detected. In vivo Hcm1 phosphorylation has been reported in systematic studies (40–42); however, studies involving affinity purification and phosphopeptide enrichment are expected to reveal a more complete set of protein phosphorylation sites. We exhaustively identified in vivo Hcm1 phosphorylation sites by mass spectrometry analysis with 15 runs of exponentially growing cell cultures and two runs of ␣-factor-arrested cell cultures in this study. We employed phosphopeptide-enriched tandem mass spectrometry analysis with a strain carrying endogenously expressing HTB (6⫻His-2⫻Tev-biotin)-tagged Hcm1. HTB tagging has been used successfully for tandem affinity purification of transcription factors in budding yeast (17, 43). Detailed MS methods are provided in Materials and Methods, and the data are provided in Table S1 in the supplemental material. As a result of the MS phosphorylation site mapping, Hcm1 was phosphorylated at 39 serine, threonine, or tyrosine amino acid residues in vivo (Fig. 3B; STY denoted in red), which covered most of the reported sites from systematic studies but also revealed 26 novel sites (Fig. 3C; also see

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FIG 4 Hcm1 S61, S65, and S66 are required for the cell wall integrity checkpoint. (A) Hcm1 phosphorylation site amino acid replacement mutants were constructed to produce nonphosphorylatable alleles. (B) Cells of the indicated strains (YOC4879, YOC4839, YOC5084, YOC4896, YOC5085, and YOC4841) harboring alleles depicted in panel A were tested for their function in the cell wall integrity checkpoint. Cells were synchronized to the G1 phase by centrifugal elutriation and budding morphology (top; med-large bud indicates a bud that is approximately one-third larger than the mother) was investigated at each time point after release of the cells at the permissive growth temperature of 25°C, and bipolar spindle morphology (bottom) was investigated by visualizing the spindle using indirect immunofluorescence microscopy (n ⬎ 200 for each time point). The percentage of med-large budded cells or cells with a bipolar spindle in the population after release is shown for each time point. Strains are as described in the graph. (C) Cells depicted in panel B also were investigated at the restrictive temperature of 37°C in terms of budding (top) and bipolar spindle morphology (bottom). (D) Representative images of the cells described in panel C at the indicated time points are shown. Arrows indicate bipolar spindles.

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Table S2), implying the novelty and the confidence of this study. Schematic mapping of the phosphorylation sites showed six phosphosites in the N-terminal region and the remainder in the C-terminal region. Twelve of the phosphorylation sites possessed sequences that were proline directed and preferred by CDK (T/SP-X-K/R; five sites) or MAPK (P-X-T/S-P; two sites) (Fig. 3D) (44). Phosphorylation of Hcm1 at S61, S65, and S66 is required for the cell wall integrity checkpoint. Landry et al. showed that potential Hcm1 phosphorylation sites in the N-terminal region, namely, T39, S61, and S66, are required for degradation (16). Our MS analysis revealed other phosphorylation sites (S57A and S65A) in proximity to these sites. Therefore, we constructed mutagenized Hcm1 alleles at five phosphorylation sites replaced with nonphosphorylatable alanine (T39A, S57A, S61A, S65A, and S66A) (Fig. 4A) to elucidate the possible role of Hcm1 phosphorylation in the cell wall integrity checkpoint. These mutants were evaluated for budding progression and spindle formation at the permissive temperature (checkpoint-uninduced condition) and showed cell cycle progression similar to that of the wild type (Fig. 4B). We then tested whether the alanine replacements at these sites affected the cell wall integrity checkpoint (Fig. 4C). While cell budding was inhibited in all mutants tested (Fig. 4C, top), the T39A and S57A Hcm1 mutants showed a cell cycle arrest phenotype and an inability to form bipolar spindles, suggesting that the loss of phosphorylation has no significant effect on checkpoint function (Fig. 4C, bottom). In contrast, the S61A Hcm1 mutants showed a strong bipolar spindle formation phenotype, as almost 50% of the cells developed a bipolar spindle 180 to 240 min after release from the G1 phase (Fig. 4C, bottom, and D). About 30% of S65A and S66A Hcm1 mutant cells formed a bipolar spindle 180 to 240 min after release from the G1 phase (Fig. 4C, bottom, and D). We also tested whether each single mutation affected the protein level of Hcm1 when the cell wall integrity checkpoint was active, and we found that the protein level was markedly higher in the defective check-

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point mutants (S61A, S65A, and S66A) (Fig. 5A and B). These findings strongly suggest that the loss of Hcm1 downregulation causes defects in the cell wall integrity checkpoint. We constructed an S61A, S65A, and S66A collective mutant and evaluated its function at the checkpoint to assess the additive effect of S61, S65, and S66 phosphorylation. We then evaluated whether the collective mutant showed delayed degradation that was similar to what was described previously (16). As shown in Fig. 6A and B, in response to the addition of cycloheximide, an inhibitor of protein synthesis, the Hcm1 protein level became markedly higher in the mutant after 5, 15, and 30 min, supporting the model that the protein degradation was delayed in the collective mutant. Using the cell wall synthase-defective mutant fks11154, we next showed that the collective mutant overrode the cell cycle arrest phenotype (Fig. 6C and D), indicating a defective cell wall integrity checkpoint. In addition, the manner in which the bipolar spindle was formed was similar to that of S61A in Fig. 4, suggesting no additive effect among the S61A, S65A, and S66A mutations. We also evaluated whether the loss of phosphorylation sites affected the Hcm1 protein level. We found that the collective mutant showed an increased protein level, almost 2-fold that of the wild-type control, at 60 to 120 min after release from G1 phase (Fig. 6E and F). Phos-tag electrophoresis revealed that the levels of nonphosphorylated Hcm1 species were significantly higher in the collective mutant (Fig. 6E, top, and F), suggesting phosphorylation at those sites. Since the transcription profiles of the collective mutant were similar to those of the control strain (Fig. 6G, HCM1), the increase in protein level in the collective mutant was not due to transcriptional upregulation. Taken together, these results suggest that S61, S65, and S66 phosphorylation or N-terminal phosphorylation of Hcm1 contributes to its degradation when the checkpoint is activated. In accordance with the increase in Hcm1 level, the expression of FKH2, a G2/M-phase transcription factor, was upregulated in the collective mutant (Fig. 6G, FKH2), likely due to the failure to

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degrade Hcm1. This implies that nondegraded Hcm1 contributes to cell cycle progression, but we cannot exclude the possibility that N-terminal phosphorylation affects the transcriptional activation of this protein. As a consequence, cell cycle arrest due to the cell wall integrity checkpoint likely was overridden in the collective mutant. To this end, we examined the expression of M-phase cyclin Clb2 by Western blotting and showed that, at 180 min after the release from G1 phase, Clb2 expression in the collective mutant is almost 2-fold that in the wild type (Fig. 6H and I). Both the wild type and mutant showed similar kinetics of cell cycle-dependent Clb2 expression under the checkpoint-uninduced condition (Fig. 6J and K). Taken together, these data suggest that the S61, S65, and S66 phosphorylation sites are required for the timely degradation of Hcm1 upon activation of the cell wall integrity checkpoint, which inhibits transcription of the G2/M-phase transcription factor FKH2, resulting in cell cycle arrest with duplicated and adjacent SPBs. CWI MAPK activity regulates Hcm1 protein level in a manner similar to that of the cell wall integrity checkpoint. In response to cell wall stress, the CWI MAPK signaling pathway is activated to control a number of downstream effectors (45). Therefore, we tested the possibility that the CWI MAPK pathway is an upstream kinase that triggers phosphorylation when the cell wall integrity checkpoint is activated. First, the Hcm1 protein level of the collective mutant under heat stress was assessed by Western blotting. The mutant protein level was markedly higher than the wild-type Hcm1 level in the fks1-1154 mutant at 10, 30, and 60 min after the temperature shift (Fig. 7A and B). A similar result was obtained in the wild-type FKS1 background (data not shown). These results suggest that heat stress affects Hcm1 protein level by N-terminal phosphorylation. We next determined whether Slt2, a MAPK in the pathway, is required for Hcm1 downregulation under heat stress. The Hcm1 protein level was about 1.4-fold higher in the slt2⌬ mutant, even in the absence of stress (Fig. 7C and D, time zero). The difference became larger after heat stress, reaching almost 2.4-fold at 30 min after heat stress was introduced (Fig. 7C and D, 30 min). It should be noted that the levels of nonphospho-Hcm1 species increased in the slt2⌬ mutant at 30 and 60 min after the temperature shift (Fig. 7C, top, D, black bars). This implies that Slt2 contributes to Hcm1 protein level by phosphorylation. Taking these findings together, the loss of Hcm1 phosphorylation sites and the loss of a kinase in the CWI MAPK pathway impose similar effects on Hcm1 protein levels under heat stress.

We examined the transcription of HCM1 under similar conditions by Northern blotting. Although the transcription level decreased markedly after the temperature shift, HCM1 transcription in the slt2⌬ mutant and the wild type changed in a similar manner (Fig. 7E). This result suggests that the SLT2 deletion mutation does not affect the HCM1 transcript level. In addition, Hcm1 in the slt2⌬ mutant showed a sustained protein level compared to that of the wild type upon cycloheximide addition at an elevated temperature, further suggesting that MAPK contributes to the protein level through degradation (Fig. 7F and G). Finally, we introduced Bck1-20, a constitutively active form of MAPK kinase kinase (MAPKKK), into the CWI MAPK pathway (46) to determine whether elevated CWI MAPK pathway activity affects the Hcm1 level. We found that the level of Hcm1 protein was decreased in the BCK1-20 mutant, while no such reduction occurred in the collective mutant (Fig. 7H and I). Thus, the activation of the CWI MAPK pathway likely triggers Hcm1 degradation, which is dependent on the Slt2 MAPK and S61, S65, and S66 phosphorylation sites. DISCUSSION

In this study, we have shown the involvement of Hcm1 posttranslational modification in the cell wall integrity checkpoint. Hcm1 protein level is downregulated during the active cell wall integrity checkpoint to prevent the transcription of G2/M-phase genes, including the FKH2 and NDD1 transcription factors, which leads to the downregulation of the M-phase cyclin CLB2. We found that the S61, S65, and S66 phosphorylation sites are critical for the cell wall integrity checkpoint by identifying in vivo Hcm1 phosphorylation sites followed by an evaluation of nonphosphorylated mutants. Analyses of the collective mutant suggested that phosphorylation promotes the timely degradation of Hcm1 to inhibit the transcription of G2/M-phase genes and bipolar spindle formation when the checkpoint is activated. Furthermore, phosphorylation at these sites likely is promoted by the CWI MAPK signaling pathway to regulate Hcm1 protein level. We showed that Hcm1 is degraded after cell wall perturbation, which is promoted by the phosphorylation of S61, S65, and S66 N-terminal regions. These results are in agreement with a report that phosphorylation of the N-terminal region of Hcm1 promotes its degradation (16). Our MS analysis revealed many in vivo Hcm1 phosphorylation sites at different positions outside the N-terminal region, which includes some of the C-terminal phosphorylation sites suggested to regulate Hcm1 function (16, 47). The

FIG 6 S61, S65, and S66 Hcm1 phosphorylation sites are required to regulate the protein level of the cell wall integrity checkpoint. (A) The loss of Hcm1 phosphorylation sites inhibits Hcm1 protein degradation. Cells of the indicated strains (YOC4878 and YOC5057) were subjected to cycloheximide chase assay, and protein extracts were collected at different time points, followed by Western blotting together with the untagged control (YOC1001). (B) Quantification of images shown in panel A. (C) Cells harboring hcm1 S61A_S65A_S66A have a defective cell wall integrity checkpoint. Cells of the indicated strains (YOC4879 and YOC5058) were synchronized at the G1 phase by elutriation and released into YPD media at 37°C. The inhibition of bud growth (left) and bipolar spindle formation (right) were quantified at the indicated time points (n ⬎ 200 for each time point). (D) Representative images of data shown in panel C 0 and 180 min after release. Arrows indicate bipolar spindles. (E) Downregulation of Hcm1 during the functioning of the cell wall integrity checkpoint is lost in cells harboring hcm1 S61A_S65A_S66A. Cells were prepared as described for panel C and subjected to SDS-PAGE, followed by anti-HA Western blotting after Phos-tag gel electrophoresis, along with control electrophoresis and loading controls (PSTAIRE). (F) Quantification of Hcm1 level to that of the Cdc28 loading control from panel E. Proportions of phospho- and dephospho-Hcm1 are indicated by gray and black bars, respectively, at each level of Hcm1 detected. (G) Cells prepared in a manner similar to that for panel C were examined for HCM1, FKH2, and ACT1 transcription during functions of the active cell wall integrity checkpoint by Northern blotting in a time course manner. (H) Inhibition of Clb2 expression by the cell wall integrity checkpoint is partially suppressed by the loss of Hcm1 phosphorylation sites. Cells of the indicated strains (YOC5096 and YOC5097) were prepared as described for panel E, and their Myc-tagged Clb2 protein levels were determined by Western blotting. (I) Quantification of Clb2 level normalized to that of the Cdc28 loading control from panel F. (J) Cells of the indicated strains (YOC5096 and YOC5097) were prepared as described for panel H, except they were cultured at the permissive temperature of 25°C. (K) Quantification of Clb2 level normalized to that of the Cdc28 loading control from panel H.

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positive regulatory phosphorylation of those sites is thought to be promoted during the cell cycle in a CDK-dependent manner (16, 39). Arsenault et al. reported that environmental stresses dephosphorylate C-terminal phosphorylation sites through the Cnb1 calcium-activated phosphatase, thereby negatively regulating Hcm1 function, and suggested Hcm1 as a rheostat that controls cell proliferation (47). Our analyses indicated that N-terminal phosphorylation sites influence the cell wall stress response by negatively regulating Hcm1 protein level at the cell wall integrity checkpoint,

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suggesting a role for Hcm1 as a brake. Since phosphospecies of Hcm1 were present in the collective mutants, the positive regulatory C-terminal phosphorylation persists during the checkpoint or independent of the degradation signal. Therefore, it is clear that Hcm1 is a transcription factor with a dual role, one of which is to positively promote the cell cycle and the other to negatively inhibit proliferation via the cell wall integrity checkpoint. It is suggested that the positive function of Hcm1 regulated by C-terminal phosphorylation, at least in part, persists during the checkpoint, since, in the collective mutant used in this study, Hcm1 function was not

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abolished. Although, as shown previously, stress-inductive Cnb1 phosphatase activity contributes to the Hcm1 function in the cellular response to environmental stresses (47), there may be temporally or functionally different roles in the regulatory mechanisms, which should be revealed by further analysis of combinations of phosphosite mutations. Our results demonstrate that the activation of the CWI MAPK pathway promotes the degradation of Hcm1, suggesting that the CWI MAPK pathway is activated to induce degradative phosphorylation of Hcm1 when the checkpoint is activated. However, it is unknown how the signaling pathway is triggered during the checkpoint and how it affects Hcm1 phosphorylation. At present, we do not have substantial evidence showing either that CWI MAPK directly phosphorylates Hcm1 or that it is the only kinase with this function. Interestingly, we identified that Hog1, the high-osmolarity glycerol (HOG) MAPK of the signaling pathway, and its upstream SHO1-branch factors are indispensable for the cell wall integrity checkpoint (unpublished data), as opposed to the conventional upstream sensors of the CWI MAPK pathway. Previous studies have shown that zymolyase, specifically the 1,3␤-glucanase component of zymolyase, activates the CWI MAPK pathway in a Hog1-dependent manner through SHO1-branch sensors (48, 49). There are considerable similarities between the cell wall integrity checkpoint and the adaptive response to zymolyase, such as the gene products required for proper responses and the nature of stress targeting 1,3-␤-glucan in the cell wall, suggesting that MAPK is activated in a similar manner. We propose that the key regulatory mechanism of the cell wall integrity checkpoint is the regulation of Hcm1 via MAPK activation, leading to the inhibition of downstream cell cycle regulators such as the Fkh2 complex and Clb2. Although a number of genome-wide studies have contributed to our understanding of the transcriptionally activated gene clusters specific to each cell cycle phase (7, 9, 16, 50), relatively less is known about the cell cycle checkpoint controlling the transcriptional network. One example is the DNA replication and DNA damage checkpoint that controls the Dun1 kinase pathway and Nrm1 transcription repressor via Rad53, which affects transcription of the G1/S cluster (9, 51–53). Negative regulation of Hcm1 by the cell wall integrity checkpoint is an example of a cell cycle checkpoint signal feeding into a key component of the transcription network during late S phase. Another interesting aspect of the checkpoint is the involvement of the MAPK signaling pathway in the regulatory mechanism of a cell cycle-related transcription factor. A previous study showed that although it is independent of cell wall stress, Pkc1, an upstream kinase of the CWI MAPK pathway, downregulates the G2/Mphase Forkhead transcription factor complex by phosphorylating Ndd1 (54). Therefore, other mechanisms may exist in which MAPK signaling controls cell cycle transcription factors. Considering the complexity of the transcription network of the cell cycle as well as the fact that a considerable number of genes are regulated by MAPK signaling pathways, other regulatory mechanisms, in addition to the inhibition of Hcm1, may contribute to the function of the cell wall integrity checkpoint. Similar to the cell wall integrity checkpoint, environmental stressors that control the cell cycle have been investigated in other organisms. StyI MAPK, the Hog1 homologue in Schizosaccharomyces pombe, delays the cell cycle during the G2/M phase in response to osmostress by regulating Cdc25 localization (55). Interestingly, StyI regulation of Cdc25 is independent of Wee1

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function, similar to the cell wall integrity checkpoint being independent of Swe1 (3, 56). Evidence has accumulated for the existence of a CWI MAPK pathway in plants that is similar to that of budding yeast (reviewed in reference 57). One example of reduced cell wall material controlling cell growth is when pectin, a component of the Arabidopsis thaliana cell wall, is reduced through mutagenesis or use of an inhibitor, a negative feedback mechanism in the brassinosteroid (BR) signaling pathway that reduces BR production while inducing the expression of cell wall remodeling genes to ensure proper growth (58). Thus, mechanisms that regulate proliferation in response to cell wall stresses in other organisms have been reported, suggesting conservation of the checkpoint mechanism to coordinate the cell cycle under conditions in which the cell wall is under stress. ACKNOWLEDGMENTS We thank S. Nogami for assistance in the initiation of this study and K. Suzuki and members of the Signal Transduction laboratory for valuable discussion. This work was supported by grants from the Ministry of Education, Science and Sports and Culture of Japan (Y.O.) and a grant from the Austrian Science Fund (FWF) F3411-B19 (G.A.).

FUNDING INFORMATION Ministry of Education, Culture, Sports, Science, and Technology (MEXT) provided funding to Yoshikazu Ohya under grant numbers 19057002 and 15H04402. Austrian Science Fund (FWF) provided funding to Gustav Ammerer under grant number F3411-B19.

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