Ypk1 and Ypk2 kinases maintain Rho1 at the plasma membrane by flippase-dependent lipid remodeling after membrane stresses

The plasma membrane (PM) frequently suffers from physical stress, including turgor pressure caused by imbalanced osmolality and membrane stretching/shrinking caused by muscle contraction (Lessey et al., 2012; Hohmann, 2002). Moreover, abnormal lipid composition causes PM stress even without environmental perturbations (Berchtold et al., 2012). To quickly manage such PM stress, eukaryotic cells have developed sophisticated signaling circuits that involve the target of rapamycin complex 2 (TORC2) kinase complex. TORC2 is activated by PM stress, such as hypotonic shock or inhibition of sphingolipid biosynthesis (Berchtold et al., 2012), and regulates various cellular functions, including actin organization, cell motility and morphogenesis (Loewith and Hall, 2011; Cybulski and Hall, 2009; Loewith et al., 2002; Zoncu et al., 2011).

In both yeast and mammals, the Rho-family of GTPases is a crucial target of TORC2 (Schmidt et al., 1997; Jacinto et al., 2004; Ho et al., 2008; Helliwell et al., 1998a). In budding yeast, the RhoA homolog Rho1 plays essential roles in actin organization and in stress responses (Levin, 2011). After activation by its guanine nucleotide exchange factors (GEFs), Rho1 binds to and recruits its effector Pkc1 (Andrews and Stark, 2000; Kamada et al., 1996), the only protein kinase C (PKC) in budding yeast, to the PM. Then, Rho1-GTP and the membrane lipid phosphatidylserine cooperatively activate Pkc1 (Kamada et al., 1996), which in turn activates the downstream MAP kinase cascade for transcriptional responses (Kamada et al., 1995) and remodels actin organization partly through downregulation of the formin Bni1 and the exocyst subunit Sec3 (Kono et al., 2012). Several models have been proposed to explain how TORC2 regulates Rho1 and Pkc1, including TORC2-dependent activation of Rho1 GEFs (Schmidt et al., 1997; Ho et al., 2008) and Pkc1 phosphorylation by TORC2 (Nomura and Inoue, 2015). However, the physiological significance of these mechanisms is not well understood.

The best-characterized TORC2 targets are the Akt/SGK families of protein kinases, which are directly phosphorylated and activated by TORC2 (Kamada et al., 2005). The yeast Akt (and/or SGK) homologs Ypk1/Ypk2 regulate sphingolipid synthesis via the homologs Orm1 and Orm2, and Lac1 and Lag1 (Roelants et al., 2011; Muir et al., 2014; Sun et al., 2012; Aronova et al., 2008), phospholipid flipping via the flippase kinases Fpk1/Fpk2 (Roelants et al., 2010), and production and efflux of glycerol via Gpd1 and Fps1, respectively (Lee et al., 2012; Muir et al., 2015). Among these effectors, Fpk1/Fpk2, whose kinase activities are inhibited by phosphorylation through Ypk1/Ypk2 (Roelants et al., 2010), are of particular importance because a recent quantitative phosphoproteomic approach revealed that regulation of actin polarization and endocytosis through the TORC2-Ypk1/Ypk2 pathway is largely mediated by the Fpk1/Fpk2 axis (Rispal et al., 2015). A recent study demonstrated that Fpk1/Fpk2 negatively regulate the Rho1-Pkc1 pathway, at least in part via the Rho1 GEF Rom2 (Niles and Powers, 2014). However, the Fpk1/Fpk2 substrates that regulate Rho1-Pkc1 remain unknown.

In this study, we define a key signaling mechanism that links the TORC2-Ypk1/Ypk2-Fpk1/Fpk2 kinase cascade to Rho1-Pkc1. Ypk1/Ypk2 promote cortical localization of Rho1 through inhibition of the Lem3-containing lipid flippase complex, an established target of Fpk1/Fpk2. The flippase complex determines subcellular distribution of phosphatidylserine that is required for Rho1 localization and cell viability under stress. We propose that Ypk1/Ypk2-dependent rearrangement of phosphatidylserine compensates for the reduction of anionic charge at the PM caused by stress-triggered degradation of phosphatidylinositol 4,5-bisphosphate [PI(4,5)P₂].

The TORC2-Ypk1/Ypk2 cascade regulates actin polarity (Rispal et al., 2015). To examine the relationship between YPK1/YPK2 and RHO1, we first tested their genetic interaction. Overexpression of RHO1 rescued the temperature-sensitive (ts) growth defect of the ypk1-1 ypk2Δ strain (hereafter referred to as ypk^ts) (Fig. 1A). Moreover, the actin organization defect of ypk^ts cells was largely rescued by overexpression of RHO1 (Fig. 1B). These results suggest that Ypk1/Ypk2 regulate actin organization via or in parallel with Rho1.

TORC2 activates Rho1 GEFs through an unknown mechanism (Schmidt et al., 1997; Ho et al., 2008), and Fpk1/Fpk2 regulate the localization of the Rho1 GEF Rom2 (Niles and Powers, 2014). If Ypk1/Ypk2 regulate Rho1 solely via GEFs, then active mutants of Rho1 would rescue ypk^ts. To test this possibility, we expressed two Rho1 active forms in ypk^ts cells: the RHO1-Q68H mutant, which is defective in GTP hydrolysis, and the RHO1-F35L mutant, which has high intrinsic nucleotide exchange activity. These RHO1 mutants are dominantly active and rescue the lethality of the cells that lack all three Rho1 GEFs, i.e. Rom1, Rom2 and Tus1 (Yoshida et al., 2009). Neither of these Rho1 active mutations suppressed the ypk^ts growth defect (Fig. 1C). A plausible interpretation of these results is that Ypk1/Ypk2 regulates Rho1 through GEF-independent mechanisms.

Next, we tested the possibility that Ypk1/Ypk2 regulate Rho1 localization. In wild type cells, Rho1 is concentrated at the bud cortex during polarized growth; but we found that localization of GFP-Rho1 at the bud cortex was slightly reduced in ypk^ts cells at the restrictive temperature, although Rho1 protein was expressed at normal levels (Fig. 1D,E). Ypk1/Ypk2 are activated upon PM stresses, such as hypotonic shock or treatment with myriocin (an inhibitor of sphingolipid synthesis), and are essential for the resistance to these stresses (Berchtold et al., 2012). Based on these notions, we examined GFP-Rho1 localization under these stresses. Under both stress conditions, ypk^ts cells showed a prominent impairment of cortical localization of Rho1 compared with that seen in unstressed cells (Fig. 1D,F). These results suggest that Ypk1/Ypk2 support PM localization of Rho1, especially when the PM is stressed.

Rho GTPases Rho1 and Cdc42 are delivered to the inner cortex of growing daughter cell tips through polarized secretion (Wedlich-Soldner et al., 2003; Abe et al., 2003). Once delivered, Rho1 and Cdc42 are anchored to the inner PM by two mechanisms: by a hydrophobic interaction with the PM via covalently attached geranylgeranyl moiety and by an electrostatic interaction with anionic PM lipids via the poly-basic sequence (Heo et al., 2006). Therefore, elaborate regulation of anionic lipids is essential for stable localization of Rho1. Although Ypk1/Ypk2 are well-established regulators of sphingolipids and neutral glycerophospholipids (Roelants et al., 2010, 2011; Muir et al., 2014; Sun et al., 2012; Aronova et al., 2008), it is unclear whether and how anionic lipids are regulated by Ypk1/Ypk2.

We have previously shown that Rho1 localizes to the PM in part by interaction of the C-terminal Rho1 poly-basic sequence with PI(4,5)P₂ (Yoshida et al., 2009). Therefore, we first tested the role of Ypk1/Ypk2 in the regulation of PI(4,5)P₂. GFP-2PH^PLC, a specific biosensor for PI(4,5)P₂ (Stefan et al., 2002), exclusively decorated the PM in both wild type and ypk^ts cells (Fig. 2A), suggesting that Ypk1/Ypk2 are not required to maintain PI(4,5)P₂ levels.

Next, we examined the role of Ypk1/Ypk2 in the subcellular distribution of phosphatidylserine, another important anionic lipid, by using the specific phosphatidylserine biosensor GFP-2PH^evt-2 (Uchida et al., 2011; Lee et al., 2015). Phosphatidylserine has been implicated in polarized growth through PM recruitment of the Rho GTPase Cdc42 in both yeast and humans (Fairn et al., 2011; Das et al., 2012; Bruurs et al., 2015). GFP-2PH^evt-2 showed peripheral localization in wild type cells, reflecting enrichment of phosphatidylserine at the inner leaflet of the PM (Fig. 2B). In sharp contrast, GFP-2PH^evt-2 was mainly lost from the PM and relocated to the early endosomes in ypk^ts cells (Fig. 2B,C). A similar relocation of GFP-2PH^evt-2 has been observed in ypk1(L424A) ypk2Δ, the analog-sensitive ypk mutant (Roelants et al., 2011) (Fig. 2D). Displacement of GFP-2PH^evt-2 from the PM suggests that either the total amount of phosphatidylserine at the inner PM is reduced or that the ability of phosphatidylserine to recruit GFP-2PH^evt-2 is impaired. The latter event can be caused by an altered lateral distribution of phosphatidylserine, because recent evidence suggested that nanoclustering of phosphatidylserine at the inner PM increases its ability to recruit proteins, such as human K-Ras (Zhou et al., 2015). In either case, our data suggests that Ypk1/Ypk2 regulate the spatial organization of phosphatidylserine.

Next, we sought the Ypk1/Ypk2 effector(s) of phosphatidylserine regulation. Ypk1/Ypk2 regulate phospholipid asymmetry by inhibiting Fpk1/Fpk2 kinases (Roelants et al., 2010). Fpk1/Fpk2 phosphorylate and activate the Dnf1/Dnf2-Lem3 lipid flippase complex, which flips the neutral lipids phosphatidylcholine and phosphatidylethanolamine from the outer leaflet to the inner leaflet of the PM (Nakano et al., 2008; Saito et al., 2004; Kato et al., 2002; Baldridge and Graham, 2012, 2013; Baldridge et al., 2013; Panatala et al., 2015; Iwamoto et al., 2004). In human cells, phosphatidylcholine flipping is proposed to dilute the local concentration of phosphatidylserine at the inner PM (Miyano et al., 2016). We therefore examined the involvement of the flippase complex in phosphatidylserine regulation. We found that deletion of LEM3 largely restores PM localization of the phosphatidylserine probe GFP-2PH^evt-2 in ypk^ts (Fig. 3A). This result suggests that Ypk1/Ypk2 regulate phosphatidylserine distribution via inhibition of the Lem3-containing flippase complex.

We also found that LEM3 deletion efficiently rescues delocalization of GFP-Rho1 upon hypotonic shock or myriocin treatment, as well as temperature-sensitive growth defects and myriocin hypersensitivity of ypk^ts (Fig. 3B,C,D), raising the possibility that redistribution of phosphatidylserine contributes to Rho1 regulation by Ypk1/Ypk2. To test the requirement of phosphatidylserine in Rho1 regulation, we depleted cellular phosphatidylserine by deleting CHO1, the only gene encoding phosphatidylserine synthase in budding yeast. GFP-Rho1 was excessively enriched in the bud cortex in unstressed lem3Δ cells, whereas CHO1 deletion cancelled this effect (Fig. 3E). Furthermore, in lem3Δ cells, GFP-Rho1 was maintained in the bud cortex even after myriocin treatment; this phenotype was also suppressed by CHO1 deletion (Fig. 3F). Thus, phosphatidylserine is required for the flippase inhibition-dependent cortical localization of Rho1.

We further analysed the mechanism of Rho1 regulation by Lem3. Phosphatidylserine supports cortical localization of yeast Cdc42 by preventing its extraction by Rho guanine nucleotide dissociation inhibitor (GDI) (Das et al., 2012). Rho GDI is potentially involved in flippase-mediated Rho1 regulation as well, because enhancement of PM localization of Rho1 by LEM3 deletion was not significant in rdi1Δ cells, which lack the only Rho GDI in budding yeast (Fig. 3G). A recent study revealed that Ypk1/Ypk2 maintain the Rho1 GEF Rom2 at the PM through inhibition of Fpk1/Fpk2 (Niles and Powers, 2014), suggesting that Rom2 contributes to Rho1 regulation by Ypk1/Ypk2. However, enhanced PM localization of Rho1 in by LEM3 deletion was still evident in the yeast strains that lack all three Rho1 GEFs Rom1, Rom2 and Tus1 (ΔGEF) (Fig. 3H), suggesting that Rho1 regulation by flippase is not solely mediated by GEFs.

Next, we examined whether Rho1 physically associates with phosphatidylserine. An in vitro binding assay using PIP strips indicated that the Rho1 poly-basic sequence has an affinity for phosphatidylserine, supporting the idea that phosphatidylserine contributes to the cortical localization of Rho1 through electrostatic interaction – although the interaction was weaker than that of other anionic lipids (Fig. 3I). In vivo, a high concentration of phosphatidylserine at the PM may compensate for the low affinity of phosphatidylserine for the Rho1 poly-basic sequence, as phosphatidylserine accounts for more than 10% of PM phospholipids, whereas PI(4,5)P₂ composes only 1% (Fairn and Grinstein, 2012; Leventis and Grinstein, 2010; van Meer et al., 2008; Martin, 2015). Nanoclustering of phosphatidylserine at the PM might also contribute to Rho1 binding, as is the case for human K-Ras (Zhou et al., 2015). We confirmed that the effect of lem3Δ on Rho1 PM localization in vivo is mediated by the poly-basic sequence, as the Rho1^5KA mutant, whose five lysine residues within its poly-basic sequence are mutated to alanine (Yoshida et al., 2009), did not accumulate in the PM in lem3Δ cells (Fig. 3J). Taken together, our data supports the model that flippase inhibition by Ypk1/Ypk2 promotes PM localization of Rho1 by enhancing the electrostatic interaction between the Rho1 poly-basic sequence and anionic lipids, including phosphatidylserine.

As phosphatidylserine is a highly abundant anionic lipid composing up to 10% of PM phospholipids (Fairn and Grinstein, 2012; Leventis and Grinstein, 2010; van Meer et al., 2008), its contribution to the net negative charge of the PM is significant. Interestingly, however, the cho1Δ mutant is viable, indicating that phosphatidylserine is dispensable for normal cell proliferation. Based on our finding that phosphatidylserine is under the control of Ypk1/Ypk2 (Fig. 2), we speculated that phosphatidylserine plays an important role under PM stress. To test this hypothesis, we examined GFP-Rho1 localization in the cho1Δ mutant under PM stress. In cho1Δ cells, GFP-Rho1 normally localized to the PM in the unstressed condition. In contrast, upon hypotonic shock, cortical localization of GFP-Rho1 was severely impaired (Fig. 4A). Because phosphatidylserine is a precursor of phosphatidylethanolamine, CHO1 deletion also significantly decreases the level of phosphatidylethanolamine (Fairn et al., 2011). To rule out the possibility that defective GFP-Rho1 localization in cho1Δ cells is an indirect consequence of phosphatidylethanolamine reduction, we examined Rho1 localization in the psd1Δ mutant, which lacks the main phosphatidylethanolamine synthase (Trotter et al., 1993). Regardless of stress, psd1Δ cells were only slightly defective in the cortical localization of GFP-Rho1, suggesting that phosphatidylethanolamine has a minor role in Rho1 localization (Fig. 4B). Importantly, hypotonic shock did not cause severe delocalization of GFP-Rho1 in psd1Δ cells. Thus, the loss of cortical localization of Rho1 in cho1Δ cells under PM stress is not due to the loss of phosphatidylethanolamine. Analogous to hypotonic shock, Rho1 delocalization induced by treatment with myriocin was exacerbated in cho1Δ cells (Fig. 4C). These results suggest that phosphatidylserine is crucial for the maintenance of cortical localization of Rho1, specifically under PM stress.

Rho1 recruits its effector protein Pkc1 to the bud cortex (Andrews and Stark, 2000). Analogous to GFP-Rho1, delocalization of Pkc1-GFP after treatment with myriocin was more prominent in cho1Δ cells and less obvious in lem3Δ cells (Fig. S1), suggesting that phosphatidylserine ensures not only localization but also function of Rho1 under PM stress.

Delocalization of Pkc1-GFP in cho1Δ cells raised the possibility that phosphorylation of Mpk1, the major downstream factor of Pkc1 signaling pathway, could be decreased in these cells. However, phosphorylation of Mpk1 in response to hypotonic shock was observed even in cho1Δ cells (data not shown). This suggests that localization of Pkc1 to the bud cortex is not necessarily essential for stress-triggered activation of the downstream MAP kinase cascade, consistent with the recent observation that ectopic activation of the MAP kinase cascade by Pkc1 occurs at endosomes when PI(4,5)P₂ is artificially eliminated (Fernández-Acero et al., 2015). It is possible that phosphatidylserine-dependent regulation of Rho1-Pkc1 axis is crucial for Pkc1 targets other than the MAP kinase cascade. As an alternative readout of Rho1-Pkc1 function, we examined organization of the actin cytoskeleton, a defect associated with Rho1-Pkc1 malfunction (Delley and Hall, 1999), in cho1Δ cells. Under normal growth conditions, the yeast actin cytoskeleton polarizes at the small- to medium-sized buds to sustain polarized bud growth (Pruyne and Bretscher, 2000; Moseley and Goode, 2006). Under conditions such as heat shock or damage of the cell wall, the actin cytoskeleton is rapidly depolarized to suspend polarized cell growth and to repair damage (Levin, 2011). Consistent with a recent report (Gualtieri et al., 2004), the cortical actin cytoskeleton was rapidly depolarized upon transient (20 min) hypotonic shock, and eventually repolarized within 3 h in wild type cells (Fig. 4D). The actin repolarization process required Pkc1 activity, as pkc1-2 temperature-sensitive mutant cells failed to repolarize actin 3 h after hypotonic shock (Fig. 4D). We found that cho1Δ cells show an even more severe actin depolarization phenotype than pkc1-2 cells. Furthermore, cho1Δ cells failed to repolarize even after 3 h (Fig. 4D), suggesting requirement of phosphatidylserine for rapid repolarization and/or the completion of repair processes.

Then, we monitored cell proliferation after hypotonic shock. After the shock, wild type cells exhibited only a transient growth arrest (less than 1 h) (Fig. 4E). In contrast, both pkc1-2 and cho1Δ cells failed to proliferate for at least 5 h after release from the shock (Fig. 4E). This was not simply due to cell death or lysis because these cells remained viable and eventually restarted growth in 24 h (Fig. S2). The psd1Δ strain restarted growth within 1 h , which was comparable to the recovery time for wild type cells (data not shown), excluding the possibility that the growth recovery defects in cho1Δ are due to the loss of phosphatidylethanolamine. These results suggest that phosphatidylserine and Pkc1 are required for recovery after hypotonic shock.

We found cho1Δ cells highly sensitive to PM stress induced by myriocin treatment but not psd1Δ cells (Fig. 4F). The myriocin sensitivity of cho1Δ cells is likely to be due to the loss of phosphatidylserine but not phosphatidylethanolamine, as the psd1Δ mutant is not hypersensitive to myriocin (Fig. 4F). In contrast to cho1Δ cells, lem3Δ cells are resistant to myriocin – as is the flippase mutant dnf1Δ dnf2Δ dnf3Δ (Roelants et al., 2010). Myriocin resistance in the lem3Δ strain was largely cancelled by CHO1 deletion but not by PSD1 deletion (Fig. 4F), confirming the contribution of phosphatidylserine, but not of phosphatidylethanolamine, to PM stress resistance.

Genetic evidence suggests that one of the crucial targets of phosphatidylserine in the PM stress response is the Rho1-Pkc1 pathway: the myriocin sensitivity of cho1Δ cells was partially rescued by overexpressing Rho1 in the presence of Pkc1-R398P, the constitutively active allele of Pkc1 (Nonaka et al., 1995) (Fig. 4G). Neither overexpression of Rho1 alone nor expression of Pkc1-R398P alone detectably rescued myriocin sensitivity, indicating that phosphatidylserine promotes both recruitment of Rho1 to the PM and activation of Pkc1 during PM stress.

Why is phosphatidylserine only required for Rho1 localization when the PM is stressed (Fig. 4A)? Under normal conditions, PI(4,5)P₂ plays a major role in cortical localization of Rho1 (Yoshida et al., 2009). However, upon various stimuli, such as mechanical stress to the PM, a large fraction (∼50%) of PI(4,5)P₂ is immediately degraded by phospholipase C (PLC) to produce the second messengers diacylglycerol (DAG) and inositol triphosphates (IP₃) (Storch et al., 2012; Perera et al., 2004). Based on these facts, we hypothesized that phosphatidylserine becomes essential only when PI(4,5)P₂ levels are reduced. Consistent with this idea, the cho1Δ mutant shows a synthetic growth defect in combination with the temperature-sensitive mutation of MSS4 (Sun and Drubin, 2012), which encodes the only PI4P 5-kinase in yeast (Homma et al., 1998; Desrivieres et al., 1998).

We confirmed the rapid loss of PI(4,5)P₂ upon PM stress (Perera et al., 2004) with a visual assay using GFP-2PH^PLC. We observed a rapid redistribution of GFP-2PH^PLC from the PM into the cytoplasm within 2–4 min after hypotonic shock (Fig. 5A).

To test our hypothesis that phosphatidylserine compensates for PI(4,5)P₂, we examined the effect of flippase inhibition on Rho1 localization when PI(4,5)P₂ was absent. In mss4-1 cells, Rho1 failed to localize to the PM at the restrictive temperature (Fig. 5B), confirming the requirement of PI(4,5)P₂ in Rho1 localization. However, deletion of LEM3 significantly restored Rho1 localization without detectably restoring PI(4,5)P₂ production in the mss4-1 mutant (Fig. 5B). Both the temperature-sensitive growth defect and myriocin hypersensitivity of mss4-1 cells were rescued by deletion of LEM3 (Fig. 5C). Collectively, our data suggest that inhibition of flippase, which is triggered by activation of TORC2-Ypk1/Ypk2, supports localization of Rho1 to the PM and cell survival when levels of PI(4,5)P₂ are limited. Because enhanced Rho1 localization and myriocin resistance of lem3Δ cells are dependent on CHO1 (Figs 3E,F and 4F), phosphatidylserine should have a significant role in phenotypic rescue of mss4-1 by LEM3 deletion.

Although previous genetic studies placed TORC2 upstream of the Rho1-Pkc1 pathway (Helliwell et al., 1998a,b), the molecular link between them is poorly understood. Here, we show that Ypk1/Ypk2, essential substrates of TORC2, regulate peripheral localization of Rho1 through flippase-mediated rearrangement of phospholipids. Our results pinpoint the fact that phosphatidylserine is the key lipid responsible for Rho1 localization and function during PM stress, and that the spatial organization of phosphatidylserine is controlled largely by the Ypk1/Ypk2-flippase pathway. Based on our observations that phosphatidylserine is required for Rho1 localization under stress and that flippase inhibition bypasses the requirement of PI(4,5)P₂, we propose that Ypk1/Ypk2 redistribute phosphatidylserine as a backup for PI(4,5)P₂, thereby allowing production of PI(4,5)P₂-derived second messengers without abandoning Rho1-Pkc1 signaling (Fig. 5D).

The precise mechanism by which the Dnf1/Dnf2-Lem3 flippase complex regulates phosphatidylserine distribution needs to be elucidated. Phosphatidylserine is not a good substrate for Dnf1, which rather prefers phosphatidylcholine and phosphatidylethanolamine (Baldridge and Graham, 2012); phosphatidylserine regulation by this flippase might, therefore, be a consequence of phosphatidylcholine/phosphatidylethanolamine flipping.

We notice that LEM3 deletion only partially rescues the actin polarization defect in ypk^ts cells (data not shown) that is similar to the defect in response to pharmacological inhibition of TORC2 (Rispal et al., 2015). Therefore, Rho1 targeting does not appear to be the only mechanism by which the TORC2-Ypk1/Ypk2 pathway regulates actin organization. Further studies are needed to fully understand the role for Ypk1/Ypk2 in this process.

In mammals, the PM of certain cell types, such as cardiomyocytes and smooth muscles, are regularly exposed to physical stress. Because crucial roles of TORC2 and Rho-PKC pathways in cardiac function and stress response have been demonstrated (Kajimoto et al., 2011; Volkers et al., 2013; Yano et al., 2014; Sciarretta et al., 2015; Zhao et al., 2014; Moschella et al., 2013), flippases and phosphatidylserine might also be key signaling mediators in these cell types.

The Saccharomyces cerevisiae strains used in this study are listed in Table S1. Strains were constructed using standard yeast genetics procedures (Longtine et al., 1998). Plasmids used in this study are listed in Table S2.

For experiments using plasmids, cells were grown in synthetic complete (SC) medium lacking appropriate nutrients for plasmid preservation. In experiments including the cho1Δ strain, 1 mM ethanolamine was added to SC medium for growth support (Hikiji et al., 1988). Yeast extract, peptone, dextrose (YPD) medium was used for experiments without plasmids unless specified. Unless specified, cells were grown to mid-log phase at 30°C.

In hypotonic shock experiments, cells were acclimatized to hypertonic conditions by growing them in the presence of 1 M sorbitol. The acclimatized cells in mid-log phase were collected by centrifugation and re-suspended 1:20 volume of the same medium and then diluted 1:20 with distilled water. For actin staining and growth curve assays, which were performed at 40°C in order to inactivate Pkc1 in the pkc1-2 mutant, cells were collected after hypotonic shock for 20 min and re-cultured in the original medium (with 1 M sorbitol).

Fluorescence images were acquired by using an Eclipse E600 fluorescence microscope (Nikon) equipped with a DC350F charge-coupled device camera (Andor) and an oil 60× objective (NA 1.4). The images were captured and analysed with NIS-Elements software (Nikon).

Cells were fixed for 40 min with 4% formaldehyde (final concentration). After collecting by centrifugation, cells were stained for 30 min with 0.66 μM Alexa Fluor 488 phalloidin (A12379, Molecular Probes) and washed with phosphate-buffered saline.

Yeast cells overexpressing GFP-tagged Rho1 C-terminal tail were collected in mid-log phase. Cell lysate was prepared by disrupting the cell pellet with glass beads in lysis buffer [50 mM Tris-HCl pH 8.0), 10 mM EDTA, 100 mM NaCl, 0.5% Triton X-100, 1 mM phenylmethylsulfonyl fluoride, and complete Mini Protease Inhibitor Cocktail (Roche)]. The PIP Strips membrane (Echelon) was blocked with 3% bovine serum albumin and incubated with cell lysate (0.5 mg/ml total protein) overnight at 4°C. The signal was detected with anti-GFP antibody (7.1 and 13.1, Roche) and anti-mouse horseradish-peroxidase-conjugated IgG (NA931, GE Healthcare) and developed with ECL Prime (GE Healthcare).

Lysates were prepared as previously described (Hatakeyama et al., 2010). Briefly, cells were treated with 7.2% w/v trichloroacetic acid (final concentration), pelleted, washed with 70% ethanol and then dissolved in 6 M urea buffer. After boiling in Laemmli SDS sample buffer, samples were subjected to regular SDS-PAGE and immunoblotting experiments.

We thank Tomohiko Taguchi (Univ. Tokyo, Tokyo, Japan) and Shoken Lee (Univ. Tokyo, Tokyo, Japan) for providing plasmids and for insightful discussions. We thank Scott Emr (Cornell, Ithaca, USA), Yoshikazu Ohya (Univ. Tokyo, Kashiwa, Japan), Daniel Lew (Duke, Durham, USA), Jeremy Thorner (UC Berkeley, Berkeley, USA) and David Pellman (Harvard, Boston, USA) for providing strains and plasmids. We thank members of the Yoshida Lab and Nan Pang for technical support, and helpful discussions.