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Yeast Yeast 2014; 31: 145–158. Published online in Wiley Online Library (wileyonlinelibrary.com) DOI: 10.1002/yea.3004

Research Article

Involvement of Sac1 phosphoinositide phosphatase in the metabolism of phosphatidylserine in the yeast Saccharomyces cerevisiae Motohiro Tani* and Osamu Kuge Department of Chemistry, Faculty of Sciences, Kyushu University, Fukuoka, Japan

*Correspondence to: M. Tani, Department of Chemistry, Faculty of Sciences, Kyushu University, 6-10-1 Hakozaki, Higashi-ku, Fukuoka 812-8581, Japan. E-mail: [email protected]

Received: 5 October 2013 Accepted: 20 February 2014

Abstract Sac1 is a phosphoinositide phosphatase that preferentially dephosphorylates phosphatidylinositol 4-phosphate. Mutation of SAC1 causes not only the accumulation of phosphoinositides but also reduction of the phosphatidylserine (PS) level in the yeast Saccharomyces cerevisiae. In this study, we characterized the mechanism underlying the PS reduction in SAC1-deleted cells. Incorporation of 32P into PS was significantly delayed in sac1Δ cells. Such a delay was also observed in SAC1- and PS decarboxylase gene-deleted cells, suggesting that the reduction in the PS level is caused by a reduction in the rate of biosynthesis of PS. A reduction in the PS level was also observed with repression of STT4 encoding phosphatidylinositol 4-kinase or deletion of VPS34 encoding phophatidylinositol 3-kinase. However, the combination of mutations of SAC1 and STT4 or VPS34 did not restore the reduced PS level, suggesting that both the synthesis and degradation of phosphoinositides are important for maintenance of the PS level. Finally, we observed an abnormal PS distribution in sac1Δ cells when a specific probe for PS was expressed. Collectively, these results suggested that Sac1 is involved in the maintenance of a normal rate of biosynthesis and distribution of PS. Copyright © 2014 John Wiley & Sons, Ltd. Keywords: phospholipids; phospholipid metabolism; phosphoinositide; Saccharomyces cerevisiae

Introduction Phosphoinositides (PIPs) are phosphorylated derivatives of phosphatidylinositol (PI), which regulates many biological processes, such as vesicular trafficking, cytoskeletal organization and cell proliferation (Di Paolo and De Camilli, 2006; Odorizzi et al., 2000). PI can be phosphorylated at the D-3, D-4 or D-5 position of the inositol head group, and PI 4-phosphate [PI(4)P], PI 4,5-bisphosphate [PI(4,5)P2], PI 3-phosphate [PI(3)P] and PI 3,5bisphosphate [PI(3,5)P2] are produced in the yeast Saccharomyces cerevisiae (Odorizzi et al., 2000). In yeast, PI(4)P is synthesized from PI by three PI 4-kinases, Stt4, Pik1 and Lsb6, and PI(4,5)P2 from PI(4)P by a PI(4)P 5-kinase, Mss4 (see supporting Copyright © 2014 John Wiley & Sons, Ltd.

phosphatidylserine;

Sac1;

information, Figure S1). Stt4 and Pik1, which act as major PI 4-kinases in yeast, are essential for cell growth. Stt4 and Pik1 produce distinct pools of PI (4)P and PI(4,5)P2 and play different roles in cellular functions. For instance, Stt4 is required for normal vacuole morphology, organization of the actin cytoskelton and cell wall integrity, whereas Pik1 is required for endocytosis, vacuolar protein sorting and protein secretion (Audhya et al., 2000). PI(3) P is synthesized from PI by Vps34. PI(3)P is further phosphorylated by Fab1, which results in the production of PI(3,5)P2 (Odorizzi et al., 2000) (see supporting information, Figure S1). Vps34 forms two different multi-subunit complexes, which function in distinct biological processes, the autophagy/ cytosol-to-vacuole transport and vacuolar protein

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sorting pathways, respectively (Backer, 2008; Kihara et al., 2001). PIPs are dephosphorylated by the lipid phosphatases Sac1, Inp51, Inp52, Inp53, Inp54, Fig4 and Ymr1 (Odorizzi et al., 2000; Parrish et al., 2004). These phosphatases regulate the cellular levels of PIPs, together with PI kinases, and the temporal and spatial distribution of PIP signalling. Sac1 hydrolyses PI(4)P, PI(3)P and PI(3,5)P2 to PI, but PI(4)P produced by Stt4, but not Pik1, is a preferred substrate (Foti et al., 2001; Tahirovic et al., 2005). Mutation of SAC1 prevents the defect of the early secretory pathway from the Golgi to plasma membranes caused by mutation of SEC14, encoding the PI/phosphatidylcholine (PC) transfer protein (Bankaitis et al., 1989; Whitters et al., 1993). Moreover, the SAC1 mutation causes multiple aberrant phenotypes, including dramatic accumulation of PI(4)P, a defect in actin cytoskeleton organization, inositol auxotrophy, cold sensitivity, high sensitivity to multiple drugs, cell wall defects and delayed endocytosis and vacuolar protein sorting (Cleves et al., 1989; Hughes et al., 1999; Mayinger et al., 1995; Schorr et al., 2001; Tahirovic et al., 2005; Whitters et al., 1993). Furthermore, sac1 mutant cells exhibit aberrant lipid metabolism, i.e. alteration of the metabolism of sphingolipids (Breslow et al., 2010; Brice et al., 2009), accelerated PC biosynthesis from choline and a reduction in the cellular phosphatidylserine (PS) level (Rivas et al., 1999; Tani and Kuge, 2010), indicating that Sac1 is involved in the regulation of several lipid metabolism pathways other than the metabolism of PIPs. Previously, we found that the double mutation of SAC1 and a non-essential sphingolipid-metabolizing enzyme gene (CSG1, CSG2, IPT1 or SCS7) causes a synthetic growth defect phenotype in yeast (Tani and Kuge, 2010). Restoration of the PS level by overexpression of PS synthase partly suppressed the synthetic growth defect phenotype of csg1Δ, ipt1Δ and scs7Δ cells under SAC1-repressive conditions, indicating that the reduction in the PS level caused by the SAC1 defect causes a synthetic growth defect with the deletion of each of the sphingolipidmetabolizing enzyme genes. However, it remains unclear how the PS level is decreased in sac1Δ cells. In this study, we characterized the mechanism underlying the PS reduction in sac1Δ cells. The rate of biosynthesis of PS in sac1Δ cells was significantly lower than that in wild-type cells; however, the expression level of PS synthase was the same. Copyright © 2014 John Wiley & Sons, Ltd.

M. Tani and O. Kuge

Moreover, an aberrant distribution of PS was observed in sac1Δ cells when a specific probe for PS was expressed in the cells. These results suggested that Sac1 is important for the maintenance of normal biosynthesis and a normal subcellular distribution of PS.

Materials and methods Yeast strains and media The S. cerevisiae strains used are listed in Table 1. Disruption of SAC1, PSD1, PSD2, EPT1, CPT1 and VPS34 was performed by replacing their open reading frames with the URA3 marker from the pRS406 vector (Sikorski and Hieter, 1989), the kanMX4 marker from a genome from a yeast knockout library or the pFA6a–kanMX4 vector (Wach et al., 1994), or the natMX4 marker from the p4339 vector (pCRII–TOPO::natMX4) (Tong and Boone, 2006). Occasionally, kanMX4 and natMX4 were replaced with the hygromycin B-resistant gene (from the pFA6a–hphNTI vector; Janke et al., 2004) to create hphMX4. To generate a yeast in which the expression of STT4 is regulated by doxycycline (Dox), the STT4 upstream region was replaced with a tetracycline operator cassette containing a repressor binding site (tetO2), the gene encoding the TetR–VP16 tTA transactivator, and a kanMX4 marker, as described previously (Belli et al., 1998). For tagging of the C-terminus of Cds1 with six copies of the HA epitope (6 × HA), a 6 × HA fusion cassette with the hphNT1 marker from the pYM16 vector was introduced immediately upstream of the stop codon of chromosomal CDS1, as described previously (Janke et al., 2004). For overexpression of CDS1, a TEF promoter cassette with the natNT2 marker from pYM-N19 was introduced immediately upstream of the initiator ATG of chromosomal CDS1, as described previously (Janke et al., 2004). To tag the N-terminus of Pss1 and Pis1 with three copies of the FLAG epitope (3 × FLAG), a 3 × FLAG tag was introduced immediately downstream of the initiator ATG of chromosomal PSS1 or PIS1 without changing the potential promoter region, as described below. A DNA fragment of the PSS1 or PIS1 ORF without the initiator ATG was amplified by PCR using a 5′ primer with a HindIII restriction site (5′-TAAAAGCTTGTTGAATCAG ATGAAG ATTTCGCA-3′ for PSS1; or 5′-TAAA Yeast 2014; 31: 145–158. DOI: 10.1002/yea

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Table 1. Strains used in this study Strain BY4741 MTY1013 MTY1016 MTY1027 MTY1376 MTY1377 MTY285 MTY1167 MTY1148 MTY1156 MTY1177 MTY1180 MTY1144 MTY1154 MTY1158 MTY1163 MTY1252 MTY1251 MTY1357 MTY1364 MTY1358 MTY1365 MTY1009 MTY1010 MTY90 MTY1112

Genotype

Source

MATa his3Δ1 leu2Δ0 met15Δ0 ura3Δ0 BY4741, sac1Δ::kanMX4 BY4741, psd1Δ::hphMX4 psd2Δ::natMX4 BY4741, psd1Δ::hphMX4 psd2Δ::natMX4 sac1Δ::kanMX4 BY4741, ept1Δ::hphMX4 cpt1Δ::kanMX4 BY4741, ept1Δ::hphMX4 cpt1Δ::kanMX4 sac1Δ::natMX4 BY4741, 3xFLAG-PSS1::hphNT1 BY4741, sac1Δ::natMX4 3xFLAG-PSS1::hphNT1 BY4741, CDS1-6xHA::hphNT1 BY4741, sac1Δ::kanMX4 CDS1-6xHA::hphNT1 BY4741, 3xFLAG-PIS1::hphNT1 BY4741, sac1Δ::natMX4 3xFLAG-PIS1::hphNT1 BY4741, TEFp-CDS1::natNT2 BY4741, sac1Δ::kanMX4 TEFp-CDS1::natNT2 BY4741, TEFp-CDS1-6xHA::natNT2, hphNT1 BY4741, TEFp-CDS1-6xHA::natNT2, hphNT1 sac1Δ::kanMX4 BY4741, ADHp-yeGFP-PSS1::natNT2 PHO88-eqFP611::kanMX4 BY4741, sac1Δ::URA3 ADHp-yeGFP-PSS1::natNT2 PHO88-eqFP611::kanMX4 BY4741, ADHp-yeGFP-PIS1::natNT2 PHO88-eqFP611::kanMX4 BY4741, sac1Δ::URA3 ADHp-yeGFP-PIS1::natNT2 PHO88-eqFP611::kanMX4 BY4741, ADHp-yeGFP-CDS1::natNT2 PHO88-eqFP611::kanMX4 BY4741, sac1Δ::URA3 ADHp-yeGFP-CDS1::natNT2 PHO88-eqFP611::kanMX4 BY4741, tetO2-STT4::kanMX4 BY4741, tetO2-STT4::kanMX4 sac1Δ::URA3 BY4741, tetO7-SAC1::kanMX4 BY4741, tetO7-SAC1::kanMX4 vps34Δ::natMX4

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AGCTTAG TTCGAATTCAACACCAGA-3′ for PIS1), a 3′ primer with a BamHI restriction site (5′-CAAGGATCCCTATGGCTTTGGAATTTTC AAGCTC-3′ for PSS1; or 5′-AAGGATCCTCA GTAAGTCTTGTT CTTCTCGTT-3′ for PIS1), and yeast genomic DNA as a template. The PCR product was inserted into the HindIII and BamHI sites of the p3 × FLAG-CMV™-7.1 vector (Sigma, St. Louis, MO, USA). A DNA fragment of 3 × FLAG-PSS1 or 3 × FLAG-PIS1 was amplified by PCR using a 5′ primer (5′- TCTATTTGATTC AATCAAAAAACAAAAATAAAACTATATATTAAAAAATGGACTACAAAGACCATGACGG3′ for PSS1; or 5′-GGAGCCTTCAAGTAATGTA AATAAGAGGGAAAGTGTGATAGTACAAGA TGGACTACAAAGACCATGACGG-3′ for PIS1), a 3′ primer (5′-AATGATCGTTCCACTTTTTACT ATGGCTTTGGAATTTTCAAGCTC-3′ for PSS1; or 5′-GTTCCACTTTTTATCAGTAAGTCTTGTT CTTCTCGTT-3′ for PIS1), and the p3 × FLAGCMV™-7.1 vector containing the PSS1 or PIS1 ORF as a template. A DNA fragment of the hphNTI marker was amplified by PCR using a 5′ primer (5′-TTGAAAATTCCAAAGCCATAGTAAAAAG Copyright © 2014 John Wiley & Sons, Ltd.

TGGAACGATCATTCAAGA-3′ for PSS1; or 5′AAGACTTACTGATAAAAAGTGGAACGATCATTCA-3′ for PIS1), a 3′ primer (5′- AGTTATA TGTACAAATTTTTTTTGACGCCAGGCATGAACAAAAACTACTAATCGATGAATTCGAGCTCG-3′ for PSS1; or 5′-AAGATTAGCGCAATTA AAAGGAGAAAAAAAGTAAGAAACTCATCC TATCAATCGATGAATTCGAGCTCG-3′ for PIS1), and pYM16 (Janke et al., 2004) as a template. These two DNA fragments were extended by PCR, and the resultant DNA fragment (3 × FLAGPSS1::hphNT1 or 3 × FLAG-PIS1::hphNT1) was used to transform the cells. For tagging of the Nterminus of Pss1, Pis1 and Cds1 with a yeastenhanced green fluorescent protein (yeGFP) tag, a yeGFP fusion cassette with the ADH promoter and natNT2 marker from the pYM-N9 vector was introduced immediately downstream of the initiator ATG of chromosomal PSS1, PIS1 or CDS1, as described previously (Janke et al., 2004). For tagging of the C-terminus of Pho88 with a red fluorescent protein (eqFP611; Wiedenmann et al., 2002) tag, a eqFP611 fusion cassette with the kanMX4 marker from the pYM51 vector was introduced Yeast 2014; 31: 145–158. DOI: 10.1002/yea

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immediately upstream of the stop codon of chromosomal PHO88, as described previously (Janke et al., 2004). The cells were grown in either YPD medium [1% yeast extract (BD Difco, Heidelberg, Germany), 2% peptone (BD Difco) and 2% glucose] or synthetic complete (SC) medium [0.67% yeast nitrogen base without amino acids (BD Difco) and 2% glucose] containing nutritional supplements.

Plasmids The GFP-tagged Lact-C2 expression plasmid, pAGX2–GFP–Lact-C2, was a kind gift from Dr Hiroyuki Arai and Dr Tomohiko Taguchi (Tokyo University) (Uchida et al., 2011). A single-copy plasmid (pRS416) containing SAC1–6 × HA or SAC1–C392S–6 × HA and its 5′- and 3′-untranslated regions (540 and 74 bp, respectively) was constructed as described below. For tagging of the C-terminus of Sac1 with 6 × HA, a 6 × HA fusion cassette with the hphNT1 marker from the pYM16 vector was introduced immediately upstream of the stop codon of chromosomal SAC1, as described previously (Janke et al., 2004). A DNA fragment was amplified by PCR using a 5′ primer (5′-TGTTCCATC CCTCGAAACACT-3′), a 3′ primer with a KpnI site (5′-TTGGGTACCGGGGACGAGGCAAGCT AAAC-3′) and genomic DNA of SAC1–6 × HA:: hphNT1 strain as a template. A DNA fragment of SAC1–C392S–6 × HA was obtained by fusing the upstream and downstream DNA fragments of SAC1-6 × HA. The upstream fragment was amplified with a 5′ primer (5′-TGTTCCATCCCTCGAA ACACT-3′), a 3′ primer (5′-CCAAACAATCCAT ACTGTTTGTTCTTAC-3′; underlining shows the location of the mutation) and genomic DNA of strain SAC1–6 × HA::hphNT1 as a template. The downstream fragment was amplified with a 5′ primer (5′-GTAAGAACAAACAGTATGGATTG TTTGG-3′; underlining shows the location of the mutation), a 3′ primer (5′-TTGGGTACCGGGGA CGAGGCAAGCTAAAC-3′) and genomic DNA of the SAC1–6 × HA::hphNT1 strain as a template. These fragments were extended by PCR. The fragment obtained was subcloned into the pGEM-T-easy vector (Promega, Madison, WI, USA). The vector was digested with KpnI and SacI and the fragment obtained was subcloned into pRS416. Copyright © 2014 John Wiley & Sons, Ltd.

M. Tani and O. Kuge

[32P] orthophosphate labelling For pulse labelling, cells were cultured overnight in YPD medium, diluted (0.1 A600 U/ml) in YPD medium and then incubated for 7 h. The cells were collected by centrifugation, resuspended in 500 μl fresh YPD medium to 2 A600 U/ml, and then labelled with [32P] orthophosphate (1 μCi/1 A600 units of cells; MP Biomedicals, Morgan Irvine, CA, USA) for the indicated times at 30 °C. For steady-state labelling, cells were cultured overnight in YPD medium, with or without 10 μg/ml Dox, diluted (0.05 A600 U/ml) in 1.5 ml fresh YPD medium, with or without Dox, and then labelled with 3 μCi [32P] orthophosphate for 15 h at 30 °C. The cells were resuspended in 1 ml fresh YPD medium, with or without Dox, to 0.2 A600 U/ml and labelled with 2 μCi [32P] orthophosphate for 6 h. The radiolabelled cells were chilled on ice, collected by centrifugation, washed twice with distilled water and then suspended in 250 μl ethanol:water:diethylether:pyridine:15 N ammonia (15:15:5:1:0.018 v/v). After 15 min of incubation at 65 °C, the residue was centrifuged at 10 000 × g for 1 min and extracted once more in the same manner. The resulting supernatants were dried and the lipids were suspended in 20 μl chloroform:methanol: water (5:4:1 v/v) and then spotted on Silica Gel 60 thin layer chromatography (TLC) plates (Merck, Whitehouse Station, NJ, USA). Two-dimensional (2D) TLC with chloroform:methanol:4.2 N ammonia (9:7:2 v/v; dimension 1) and chloroform:methanol:acetic acid:water (15:3.9:2:1 v/v; dimension 2) was performed to separate the phospholipids; a typical result of TLC is shown in Figure S2 (see supporting information). Quantification of each phospholipid was performed using a Bio Imaging analyser FLA-2000 (Fuji Photo Film, Kanagawa, Japan).

Vacuole labelling with FM4-64 Yeast cells grown in SC medium lacking uracil but containing 2 mM choline (SC – Ura + Cho medium) were collected by centrifugation and then resuspended in fresh SC – Ura + Cho medium to 1 A 600 U/ 100 μl. FM4-64 (Molecular Probes, Eugene, OR, USA) was added to the cells to a final concentration of 20 μM, followed by reaction at 30 °C for 15 min, washing three times with 1 ml SC – Ura + Cho medium and then chasing in 1 ml fresh SC – Ura + Yeast 2014; 31: 145–158. DOI: 10.1002/yea

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Cho medium without FM4-64 for 120 min. The cells were collected by centrifugation and viewed under a fluorescence microscope (Leica DMRB; Leica, Solms, Germany).

SDS–PAGE and western blotting Protein extraction, SDS–PAGE and western blotting were performed as described previously (Tani and Kuge, 2010). Anti-HA (Sigma), anti-FLAG (Stratagene, La Jolla, CA, USA) and anti-Pgk1p (Molecular Probes) were used as primary antibodies. Horseradish peroxidase-conjugated anti-mouse IgG (Biosource, Camarillo, CA, USA) were used as secondary antibody.

Results Reduction in rate of biosynthesis of PS in sac1Δ cells It was previously reported that sac1Δ and sac1-22 mutant cells show a reduction in the PS level (Rivas et al., 1999; Tani and Kuge, 2010); however, the mechanism underlying alteration of the phospholipid remains unclear. To determine the rate of biosynthesis of PS, wild-type and sac1Δ cells were pulse-labelled with [32P] orthophosphate for 15, 30 and 60 min. As shown in Figure 1B, the rate of biosynthesis of PS in sac1Δ cells was reduced by approximately 50% as compared with that in wild-type cells. The rate of biosynthesis of phosphatidylinositol (PI) was slightly reduced in sac1Δ cells. Instead, phosphoinositides (PIPs), including PI(4)P and PI(3)P, accumulated dramatically (Figure 1B). It should be noted that the PI level was found not to be reduced in sac1Δ cells when the cellular levels of phospholipids were examined by means of steady-state radiolabelling with [32P] orthophosphate (Figure 1A). PS can be converted to PE by PS decarboxylase, Psd1 and Psd2 (Birner et al., 2001; Henry et al., 2012) (see supporting information, Figure S1). To investigate whether or not the conversion of PS to PE is related to the reduction in the PS level in sac1Δ cells, the rate of biosynthesis of PS was examined in psd1Δ psd2Δ and psd1Δ psd2Δ sac1Δ cells. As shown in Figure 1D, the rate of biosynthesis of PS in psd1Δ psd2Δ sac1Δ cells was also reduced by approximately 50% as compared with that in psd1Δ psd2Δ cells. Furthermore, the steady-state PS level Copyright © 2014 John Wiley & Sons, Ltd.

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in psd1Δ psd2Δ sac1Δ cells was also reduced as compared with that in psd1Δ psd2Δ cells (Figure 1C). Thus, it was indicated that the reduction in the PS level in sac1Δ cells occurs in the absence of PS decarboxylation. We also investigated the effect of SAC1 deletion under Kennedy pathway-deficient conditions. To block the Kennedy pathway, genes encoding ethanolamine- and choline-phosphotransferases, EPT1 and CPT1 (Henry et al., 2012) (see supporting information, Figure S1) were deleted. As shown in Figure 1E, F, the steady-state PS level and the rate of biosynthesis of PS in ept1Δ cpt1Δ sac1Δ cells was also reduced as compared with that in ept1Δ cpt1Δ cells. It should be noted that slight enhancement of phosphatidylcholine (PC) biosynthesis was observed by the deletion of SAC1 (Figure 1A, B). The enhancement was also observed in PS decarboxylation- or Kennedy pathway-deficient conditions (Figure 1C–F). These results may suggest that PC biosynthesis from both PS and the Kennedy pathway is upregulated by the deletion of SAC1. It was previously reported that PC biosynthesis from choline is activated by the SAC1 defect (Rivas et al., 1999). Collectively, these results suggested that the biosynthesis of PS is reduced by the SAC1 deletion.

Protein expression levels of PS, PI and CDP–DAG synthase in sac1Δ cells In yeast, the biosynthesis of PS and PI share the common liponucleotide precursor CDP–diacylglycerol (CDP–DAG), i.e. PS is synthesized from CDP– DAG and serine, and PI from CDP–DAG and inositol (Henry et al., 2012) (see supporting information, Figure S1). Thus, it is thought that the cellular PS level is influenced not only by PS synthase but also by PI and CDP–DAG synthase (Kelley et al., 1988; Shen and Dowhan, 1997). To examine the influence of SAC1 deletion on the expression levels of PS synthase (Pss1), PI synthase (Pis1) and CDP–DAG synthase (Cds1), these proteins were detected by western blotting. The chromosomal PSS1 and PIS1 were tagged with 3 × FLAG at the N-terminus without changing the potential promoter region, and the proteins were detected with an anti-FLAG antibody (Figure 2A, B). The chromosomal CDS1 was tagged with 6 × HA at the C-terminus and the protein was detected with an anti-HA antibody (Figure 2C). Yeast 2014; 31: 145–158. DOI: 10.1002/yea

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Figure 1. Steady-state and pulse radiolabelling of SAC1-deleted cells with [32P] orthophosphate. (A) Steady-state radiolabelling of wild-type and sac1Δ cells with [32P] orthophosphate. The radiolabelled phospholipids were extracted and resolved by TLC. Glycerophospholipids {cardiolipin (CL), phosphatidic acid (PA), phosphatidylethanolamine (PE), phosphatidylserine (PS), phosphatidylinositol (PI), phosphatidylcholine (PC), phosphoinositides [PIPs: PI(4)P and PI(3)P] and others [unidentified bands (see supporting information, Figure S2)]} were quantified using a Bio Imaging analyser. (B) Wild-type and sac1Δ cells were pulse-radiolabelled for 15, 30 and 60 min with [32P] orthophosphate at 30 °C. The results are expressed as relative radioactivity; the total radioactivity incorporated into phospholipids (PE, PC, PS, PI and PIPs) upon incubation of wild-type cells for 60 min was taken as 1. (C) Steady-state radiolabelling of psd1Δ psd2Δ and psd1Δ psd2Δ sac1Δ cells with [32P] orthophosphate. (D) Pulse radiolabelling of psd1Δ psd2Δ and psd1Δ psd2Δ sac1Δ cells with [32P] orthophosphate. The total radioactivity incorporated into phospholipids (PE, PC, PS, PI and PIPs) on incubation of psd1Δ psd2Δ cells for 60 min was taken as 1. (E) Steady-state radiolabelling of ept1Δ cpt1Δ and ept1Δ cpt1Δ sac1Δ cells with [32P] orthophosphate. (F) Pulse radiolabelling of ept1Δ cpt1Δ and ept1Δ cpt1Δ sac1Δ cells with [32P] orthophosphate. The total radioactivity incorporated into phospholipids (PE, PC, PS, PI and PIPs) on incubation of ept1Δ cpt1Δ cells for 60 min was taken as 1. Data shown are representative of at least three independent experiments. Details are given in Materials and methods

3xFLAG-Pss1 was detected as two protein bands, which are the phosphorylated and dephosphorylated forms, respectively (Figure 2A) (Choi et al., 2010). The expression levels of both 3xFLAG-phosphoCopyright © 2014 John Wiley & Sons, Ltd.

Pss1 and 3xFLAG-Pss1 in sac1Δ cells were comparable to those in wild-type cells (Figure 2A, D), indicating that the expression and phosphorylation levels of Pss1 are not affected by the SAC1 deletion. Yeast 2014; 31: 145–158. DOI: 10.1002/yea

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Figure 2. Expression levels of Pss1, Pis1 and Cds1 in sac1Δ cells. (A–C) Western blotting analysis of 3xFLAG-Pss1 (A), 3xFLAG-Pis1 (B) and Cds1-6xHA (C). Wild-type and sac1Δ cells expressing each tagged enzyme were grown to the exponential phase of growth in YPD medium. Yeast cell extracts were immunoblotted using anti-FLAG, anti-HA or anti-Pgk1. (D) The relative amounts of 3xFLAG-Pss1 and 3xFLAG-phospho-Pss1 were determined using ImageJ software (NIH). The total amount of Pss1 (3xFLAG-Pss1 + 3xFLAG-phospho-Pss1)/Pgk1 in wild-type cells was taken as 1. (E, F) Determination of the relative amounts of 3xFLAG-Pis1 (E) and Cds1-6xHA (F). The amount of Cds1-6xHA or 3xFLAG-Pis1/Pgk1 in wild-type cells was taken as 1. Data shown are the averages of at least three independent experiments. (G) Western blotting analysis of overexpressed Cds1-6xHA. Wild-type and sac1Δ cells expressing Cds1-6xHA by the native promoter or TEF promoter were grown to the exponential phase of growth in YPD medium. Yeast cell extracts were immunoblotted using anti-HA or anti-Pgk1. (H) Steady-state radiolabelling of wild-type, sac1Δ, TEFp-CDS1 and TEFp-CDS1 sac1Δ cells with [32P] orthophosphate. Data shown are representative of at least three independent experiments. Details are given in Materials and methods

No significant difference in the expression level of 3 × FLAG-Pis1 was observed between wild-type and sac1Δ cells (Figure 2B, E). In addition, in the in vitro enzyme assay, no significant difference in the activities of PS synthase and PI synthase was Copyright © 2014 John Wiley & Sons, Ltd.

observed between wild-type and sac1Δ cells (see supporting information, Figure S3). The expression level of Cds1-6HA in sac1Δ cells was reduced by 30% compared to that in wild-type cells (Figure 2C, F). To investigate whether or not overexpression of Yeast 2014; 31: 145–158. DOI: 10.1002/yea

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Figure 3. Subcellular localization of Pss1, Pis1 and Cds1 in sac1Δ cells. Wild-type and sac1Δ cells expressing yeGFP-Pss1, yeGFP-Pis1 or yeGFP-Cds1 and Pho88-eqFP611 were cultured overnight in YPD medium, diluted (0.1 A600 U/ml) in fresh YPD medium, and then incubated for 7 h at 30 °C. GFP and RFP fluorescence was observed by fluorescent microscopy

CDS1 can prevent the reduction in the PS level in sac1Δ cells, Cds1 was overexpressed under the control of a TEF promoter (TEFp–CDS1). The overexpression of Cds1 was confirmed by tagging with 6 × HA at the C-terminus of Cds1 (Figure 2G). No significant difference in the PS level was observed between sac1Δ and TEFp–CDS1 sac1Δ cells (Figure 2H), suggesting that the reduction in expression level of Cds1 in sac1Δ cells (Figure 2C, F) is not related to the PS reduction. In contrast, the overexpression of CDS1 caused a slight increase in the PI level in both SAC1 and sac1Δ cells. We also confirmed the subcellular localization of Pss1, Pis1 and Cds1 by tagging of their N-termini with a yeastenhanced green fluorescent protein (yeGFP). In both wild-type and sac1Δ cells, yeGFP–Pss1, yeGFP– Pis1 and yeGFP–Cds1 were well co-localized with Pho88-eqFP611, a marker protein of the ER (Tavassoli et al., 2013), indicating that the SAC1 deletion does not cause abnormal localization of Pss1, Pis1 and Cds1 (Figure 3).

Defects of both dephosphorylation and biosynthesis of PIPs cause a reduction in the PS level To investigate whether or not the PIP phosphatase activity of Sac1 is necessary for maintenance of the PS level, a single-copy plasmid harbouring Copyright © 2014 John Wiley & Sons, Ltd.

6 × HA-tagged SAC1 or SAC1–C392S, a catalytic inactive mutant (Manford et al., 2010), or an empty vector was introduced into sac1Δ cells. As shown in Figure 4A, the expression levels of Sac1–6 × HA and Sac1–C392S–6 × HA were comparable when each protein was expressed in sac1Δ cells. The expression of Sac1–6 × HA, but not Sac1–C392S–6 × HA, suppressed the growth defect of sac1Δ cells at 15 °C, a typical phenotype of sac1Δ cells (Manford et al., 2010), indicating that the mutation of C392S causes loss of the PIP phosphatase activity of Sac1–6 × HA (Figure 4B). Figure 4C shows the phospholipid composition when cells were radiolabelled to steady state with [32P] orthophosphate. sac1Δ cells expressing Sac1–6 × HA exhibited an approximately two-fold increase in the PS level as compared with that in sac1Δ cells with the empty vector, whereas that in sac1Δ cells expressing Sac1–C392S–6 × HA did not increase. These results indicated that the PIP phosphatase activity of Sac1 is necessary for complementation of the reduced PS level in sac1Δ cells. We also investigated whether or not deletion of other PIP phosphatases (Inp51, Inp52, Inp53, Inp54, Fig4 and Ymr1) affect the PS level. As shown in Figure S4 (see supporting information), the deletion of one of these PIP Yeast 2014; 31: 145–158. DOI: 10.1002/yea

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Figure 4. Expression of a catalytic inactive mutant of Sac1 does not suppress the reduction in the PS level in sac1Δ cells. (A) Western blotting analysis of sac1Δ cells harbouring pRS416-SAC1-6xHA, pRS146-SAC1-C392S-6xHA or the empty vector. Cells were grown to the exponential phase of growth in SC medium lacking uracil but containing 2 mM choline (SC – Ura + Cho medium). Yeast cell extracts were immunoblotted using anti-HA or anti-Pgk1. (B) Cold sensitivity of sac1Δ cells. Wild-type and sac1Δ cells harbouring each plasmid were cultured overnight in SC – Ura + Cho medium. Cells were spotted onto SC – Ura + Cho plates in 10-fold serial dilutions, starting with a density of 0.7 A600 U/ml. The plates were incubated at 30 °C for 1 day or 15 °C for 6 days. (C) Steady-state radiolabelling of cells harbouring each plasmid with [32P] orthophosphate. Cells were cultured overnight in SC – Ura + Cho medium, diluted (0.1 A600 U/ml) in 1.5 ml fresh SC – Ura + Cho medium, and then labelled with 3 μCi [32P] orthophosphate for 15 h at 30 °C. The cells were resuspended in 1 ml fresh SC – Ura + Cho medium to 0.2 A600 U/ml and then labelled with 2 μCi [32P] orthophosphate for 6 h. Data shown are representative of at least three independent experiments. Details are given in Materials and methods

phosphatases did not have significant effects in the phospholipid composition (Figure S4). Mutation of SAC1 causes dramatic accumulation of PI(4)P, and the bulk of the accumulated PI(4)P is generated by PI 4-kinase Stt4 (Foti et al., 2001; Tahirovic et al., 2005). STT4 is essential for cell growth; however, the deletion of SAC1 can suppress the growth defect of a temperaturesensitive mutant of STT4, due to suppression of Copyright © 2014 John Wiley & Sons, Ltd.

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the degradation of PI(4)P generated by Stt4 (Tahirovic et al., 2005). Thus, we next investigated whether or not the reduction in PS level in sac1Δ cells is suppressed by repression of STT4. To repress the expression of Stt4, a mutant strain that carries the STT4 gene under the control of a tetO2 promoter (tetO2-STT4) was used. As shown in Figure S5 (see supporting information), the growth of tetO2–STT4 cells was inhibited by the addition of Dox, and this growth inhibition was suppressed by the SAC1 deletion. Figure 5A shows the steadystate phospholipid compositions in tetO2–STT4, Dox-treated tetO2–STT4, tetO2–STT4 sac1Δ and Dox-treated tetO2-STT4 sac1Δ cells. As expected, the accumulation of PIPs in Dox-treated tetO2– STT4 sac1Δ cells was significantly lower than that in tetO2–STT4 sac1Δ cells (Figure 5Ac), indicating that the repression of STT4 by a tetracyclineregulatable promoter can suppress the accumulation of PIPs caused by the SAC1 deletion. In Dox-treated tetO2-STT4 cells, the PS level was reduced compared to untreated cells. Moreover, the repression of STT4 enhanced the reduction in the PS level caused by the SAC1 deletion (Dox-treated tetO2STT4 sac1Δ cells) (Figure 5A). These results indicated that both repression of STT4 and deletion of SAC1 cause a reduction in the PS level. Sac1 can also dephosphorylate PI(3)P and PI(3,5)P2 because slight accumulation of PI(3)P and PI(3,5)P2 has been observed in sac1Δ cells (Guo et al., 1999; Tahirovic et al., 2005). VPS34 encoding PI 3-kinase is responsible for the production of PI(3)P in yeast, and its deletion causes depletion of PI(3)P and PI(3,5)P2. Thus, we next examined the effect of double mutation of VPS34 and SAC1 on the PS level. Since double deletion of VPS34 and SAC1 causes synthetic lethality in yeast cells (Tahirovic et al., 2005), expression of Sac1 was repressed under the control of a tetO7 promoter (tetO7–SAC1) (Tani and Kuge, 2010). As shown in Figure 5B, reduction in the PS level was observed with the single deletion of VPS34 as well as the repression of STT4. Treatment of tetO7-SAC1 cells with Dox caused a reduction in the PS level; however, the reduction was not prevented by the deletion of VPS34 (Figure 5B). Collectively, these results indicated that a defect of dephosphorylation of PIPs by Sac1 or synthesis of PIPs by Stt4 and Vps34, or the double defect of the dephosphorylation and the synthesis of PIPs, causes a reduction in the PS level. Yeast 2014; 31: 145–158. DOI: 10.1002/yea

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Figure 5. Effects of the combination of mutations of SAC1 and PI kinase genes at the PS level. (A) Steady-state radiolabelling of tetO2–STT4 and tetO2–STT4 sac1Δ cells with [32P] orthophosphate in the presence or absence of 10 μg/ml Dox; (b, c) radiolabelled PS and PIPs levels in each lot of cells, respectively, extracted from (a). (B) Steady-state radiolabelling of tetO7–SAC1 and tetO7–SAC1 vps34Δ cells with [32P] orthophosphate in the presence or absence of 10 μg/ml doxycycline (Dox). Data shown are representative of at least three independent experiments. Details are given in Materials and methods

Aberrant intracellular distribution of PS in sac1Δ cells The C2 domain of lactadherin (Lact-C2) is a specific probe for PS and is used for observation of its intracellular distribution (Yeung et al., 2008). To examine the effect of SAC1 deletion on the intracellular distribution of PS, GFP-tagged Lact-C2 was expressed in wild-type and sac1Δ cells. As reported previously, GFP–Lact-C2 was observed predominantly at the plasma membrane of wild-type cells, whereas the intracellular distribution of GFP–Lact-C2 was increased in sac1Δ cells (Figure 6A). Figure 6B shows the results of Copyright © 2014 John Wiley & Sons, Ltd.

semiquantification of the intracellular fluorescence intensity of GFP–Lact-C2 in wild-type and sac1Δ cells. A significant increase in intracellular fluorescence intensity was observed in sac1Δ cells (Figure 6B). A large part of the intracellular GFP– Lact-C2 in sac1Δ cells exhibited a large and round structure, which is likely a vacuole. To visualize vacuolar membranes, sac1Δ cells expressing GFP– Lact-C2 were labelled with the lipophilic dye FM464 for 15 min, and then chased for 2 h at 30 °C, conditions under which the dye is transported from plasma membranes to vacuolar membranes (Vida and Emr, 1995). As shown in Figure 6C, intracellular Yeast 2014; 31: 145–158. DOI: 10.1002/yea

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Figure 6. Distribution of GFP–Lact-C2 in sac1Δ cells. (A) Wild-type and sac1Δ cells harbouring pAGX2–GFP–Lact-C2 were cultured overnight in SC – Ura + Cho medium, diluted (0.1 A600 U/ml) in fresh SC – Ura + Cho medium and then incubated for 8 h at 30 °C. GFP fluorescence was observed by fluorescent microscopy. Arrowheads and arrows indicate plasma membrane localization and intracellular accumulation of GFP–Lact-C2, respectively. (B) Frequency distribution of intracellular fluorescence intensity of GFP–Lact-C2 in individual cells. The fluorescence intensity was quantified using ImageJ software (NIH). The percentage of intracellular fluorescence in each individual cell was calculated as follows: intracellular fluorescence in individual cells (%) = percentage of intracellular fluorescence relative to total cellular fluorescence. Data represent the value for 100 cells pooled from three independent experiments for each strain. (C) Co-localization of GFP–Lact-C2 and vacuolar membranes in sac1Δ cells. To visualize vacuolar membranes, cells were labelled with FM4-64 for 15 min and then chased for 120 min at 30 °C. Details are given in Materials and methods

GFP–Lact-C2 was co-localized with FM4-64 in sac1Δ cells. Thus, these results suggested that the deletion of SAC1 causes an aberrant distribution of PS, which mainly exhibits vacuolar membrane localization.

Discussion In the present study, the reduction in PS level caused by deletion of SAC1 was characterized. It was found that the apparent rate of biosynthesis Copyright © 2014 John Wiley & Sons, Ltd.

of PS in sac1Δ cells was significantly reduced as compared with that in wild-type cells in a pulselabelling experiment involving [32P] orthophosphate. In yeast, PS is an intermediate of PE and PC biosynthesis, i.e. PS is converted to PE by PS decarboxylases Psd1 and Psd2, and subsequently methylated (Henry et al., 2012) (see supporting information, Figure S1). The reduction in the PS biosynthesis rate in sac1Δ cells was also observed in psd1Δ psd2Δ sac1Δ cells (Figure 1D), suggesting that PS reduction in sac1Δ cells is caused by a delay of PS biosynthesis from CDP–DAG and serine, but Yeast 2014; 31: 145–158. DOI: 10.1002/yea

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not by an aberration of decarboxylation of PS. A slight reduction in the rate of biosynthesis of PI was observed in sac1Δ cells, while PIPs were dramatically accumulated (Figure 1B), suggesting that the actual rate of biosynthesis of PI from CDP–DAG and inositol is not affected by the deletion of SAC1. It should also be noted that a significant difference in the total amount of major phospholipids (PE, PC, PI and PS) was not observed between SAC1 and SAC1-deleted cells when total cellular lipids were separated by TLC and visualized with a copper sulphate and orthophosphoric acid reagent (data not shown). In yeast, PS and PI synthases (Pss1 and Pis1) have a common substrate, CDP–DAG, and partitioning of CDP–DAG between PS and PI synthesis is an important decisive factor for the final levels of PS and PI (Kelley et al., 1988; Shen and Dowhan, 1997). For instance, PSS1-deleted mutant cells exhibit not only loss of PS but also an increase in the PI level (Bailis et al., 1987), indicating competition between PI and PS biosynthesis. Therefore, we investigated the expression levels of Pss1 and Pis1 in sac1Δ cells; however, these expression levels were not affected by the deletion of SAC1 (Figure 2A, B). Thus, it is suggested that the delay of PS biosynthesis caused by SAC1 deletion is not due to alteration of the expression levels of PS synthase and PI synthase, but by other factors. It should be noted that yeast PS synthase activity was found to be regulated by several factors, including glycerophospholipids, sphingoid bases, inositol and CTP in an in vitro enzyme assay (Carman and Henry, 1999). Investigation of factors inducing the delay of PS biosynthesis caused by the SAC1 deletion is an important issue for further studies. It was found that expression of a catalytic inactive mutant of Sac1 could not complement the PS reduction in sac1Δ cells (Figure 4), indicating that the PIP phosphatase activity of Sac1 is necessary for maintenance of the PS level. Sac1 preferentially dephosphorylates PI(4)P generated by Stt4. The SAC1 deletion causes dramatic accumulation of PI(4)P, and the accumulation is suppressed by the repression of STT4 (Foti et al., 2001). Moreover, STT4 repression can suppress the delay of endocytosis caused by the SAC1 deletion (Tahirovic et al., 2005). On the other hand, the SAC1 deletion can suppress defects associated with the STT4 repression, such as growth defects Copyright © 2014 John Wiley & Sons, Ltd.

M. Tani and O. Kuge

(see supporting information, Figure S5) (Tahirovic et al., 2005) and aberrant organization of the actin cytoskelton (Foti et al., 2001), indicating functional connections between Sac1 and Stt4. However, it was found that repression of STT4 caused a reduction in the PS level, like the SAC1 deletion. Moreover, a combination of STT4 repression and SAC1 deletion did not suppress the PS reduction (Figure 5A). Similar to the case of STT4, loss of PI (3)P and PI(3,5)P2, endogenous substrates for Sac1, due to deletion of VPS34 caused PS reduction and did not suppress the PS reduction caused by the SAC1 deletion (Figure 5B). These results indicated that defects of both the degradation and synthesis of PIPs cause a reduction in the PS level, implying that normal metabolism of PIPs is important for the maintenance of the cellular PS level. In yeast, PS plays various roles in cellular functions, such as intracellular trafficking of v-SNARE Snc1 from post-Golgi endosomes to the late Golgi and siderophore–iron chelates transporter Arn1 from vacuoles to plasma membranes (Furuta et al., 2007; Guo et al., 2010; Tani and Kuge, 2012), regulation of cell polarity via small GTPase Cdc42 (Das et al., 2012; Fairn et al., 2011), transport of amino acids (Nakamura et al., 2000) and maintenance of vacuole morphology (Hamamatsu et al., 1994). Thus, elucidation of the regulation of PS metabolism has become an important area of study. At present the physiological significance of the maintenance of PS by Sac1 remains unclear. Very recently, it was reported that the phosphatase activity of Sac1 is activated by PS, and that deletion of PSS1 causes accumulation of PI(4)P, probably due to inactivation of Sac1, suggesting a functional relationship between PS and Sac1 in regulation of the metabolism of PIPs (Zhong et al., 2012). Further detailed investigation of the molecular mechanism underlying the regulation of PS by Sac1 and the functional relationship between PS and Sac1 will provide new insights into the physiological significance of PS and PIPs. Acknowledgements We wish to thank Dr T. Ogishima (Kyushu University) for valuable suggestions regarding this study. The study was funded by the Nagase Science and Technology Foundation, the Noda Institute for Scientific Research and the Cosmetology Research Foundation, Japan. Yeast 2014; 31: 145–158. DOI: 10.1002/yea

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Supporting Information Additional supporting information may be found in the online version of this article at the publisher’s web-site: Table S1. Strains used in the supplementary figures Figure S1. Phospholipid sysnthesis and degradation pathway in the yeast S. cerevisiae Figure S2. Two-dimensional TLC of radiolabelled phosholipids Figure S3. Enzyme activities of PS synthase and PI synthase in sac1Δ cells Figure S4. Effects of deletion of PIP phosphatase on phospholipid composition Figure S5. Effects of SAC1 deletion and STT4 repression on yeast cell growth

Yeast 2014; 31: 145–158. DOI: 10.1002/yea