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Down-regulation of TORC2-Ypk1 signaling promotes MAPK-independent survival under hyperosmotic stress

  1. Alexander Muir
  2. Françoise M Roelants
  3. Garrett Timmons
  4. Kristin L Leskoske
  5. Jeremy Thorner  Is a corresponding author
  1. University of California, Berkeley, United States
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Cite this article as: eLife 2015;4:e09336 doi: 10.7554/eLife.09336

Abstract

In eukaryotes, exposure to hypertonic conditions activates a MAPK (Hog1 in Saccharomyces cerevisiae and ortholog p38 in human cells). In yeast, intracellular glycerol accumulates to counterbalance the high external osmolarity. To prevent glycerol efflux, Hog1 action impedes the function of the aquaglyceroporin Fps1, in part, by displacing channel co-activators (Rgc1/2). However, Fps1 closes upon hyperosmotic shock even in hog1∆ cells, indicating another mechanism to prevent Fps1-mediated glycerol efflux. In our prior proteome-wide screen, Fps1 was identified as a target of TORC2-dependent protein kinase Ypk1 (Muir et al., 2014). We show here that Fps1 is an authentic Ypk1 substrate and that the open channel state of Fps1 requires phosphorylation by Ypk1. Moreover, hyperosmotic conditions block TORC2-dependent Ypk1-mediated Fps1 phosphorylation, causing channel closure, glycerol accumulation, and enhanced survival under hyperosmotic stress. These events are all Hog1-independent. Our findings define the underlying molecular basis of a new mechanism for responding to hypertonic conditions.

https://doi.org/10.7554/eLife.09336.001

Introduction

In Saccharomyces cerevisiae, target of rapamycin (TOR) Complex 2 (TORC2)-dependent signaling responds to multiple plasma membrane-perturbing stresses, including sphingolipid depletion (Roelants et al., 2010, 2011), heat shock (Sun et al., 2012), and both hypotonic (Berchtold et al., 2012) and hypertonic (Lee et al., 2012) conditions. The essential downstream effector of TORC2 is the protein kinase Ypk1 (and its paralog Ypk2) (Casamayor et al., 1999; Roelants et al., 2002, 2004).

To understand how TORC2-Ypk1 signaling elicits cellular responses, we performed a genome-wide screen to discern previously unidentified Ypk1 substrates and thereby discovered that ceramide synthase activity is stimulated by TORC2-Ypk1 (Muir et al., 2014). Among other potential targets, our screen pinpointed two proteins involved in glycerol metabolism, aquaglyceroporin Fps1 (Luyten et al., 1995) and Gpt2/Gat1, an enzyme that converts glycerol-3P to phosphatidic acid (Zheng and Zou, 2001), a precursor to other phospholipids and triacylglycerol (Henry et al., 2012). Both candidates warranted further investigation because, as we showed, Ypk1-mediated phosphorylation inhibits another enzyme in glycerol metabolism, Gpd1, which generates glycerol-3P (Lee et al., 2012), and GPD1 expression is highly induced by hyperosmotic stress (Albertyn et al., 1994b). Second, accumulation of intercellular glycerol is essential for yeast cell survival under hyperosmotic conditions (Westfall et al., 2008; Saito and Posas, 2012; Hohmann, 2015). Thus, we suspected that TORC2-Ypk1 signaling might play an as yet unrecognized role in the cellular response to hyperosmotic shock.

Hyperosmotic conditions evoke two known signaling modalities. Pathways coupled to alternative osmosensors (Sln1 and Sho1) activate MAPK Hog1, which drives both transcription-independent and -dependent responses that markedly increase both production and intracellular retention of glycerol (Westfall et al., 2008; Saito and Posas, 2012; Hohmann, 2015). Hyperosmotic shock also increases cytosolic [Ca2+] thereby activating calcineurin (CN) (Denis and Cyert, 2002), promoting processes that stimulate retrieval of excess plasma membrane (Guiney et al., 2015). Although both CN-deficient and hog1∆ cells are quite sensitive to the ionic imbalances caused by high salt (e.g., 1 M NaCl), hog1∆ cells are significantly more sensitive to hypertonic stress per se, such as a high concentration of an uncharged impermeant osmolyte (e.g., 1 M sorbitol).

Our understanding of the response to high osmolarity remains incomplete, however. Although it is well documented that preventing glycerol efflux through the aquaglyceroporin Fps1 is essential for yeast to survive hyperosmolarity (Luyten et al., 1995; Tamás et al., 1999; Duskova et al., 2015), and that activated Hog1 can negatively regulate this channel by displacing the Fps1-activating proteins Rgc1/2 (Lee et al., 2013), Fps1 still closes in response to hyperosmotic shock in hog1∆ cells (Tamás et al., 1999; Babazadeh et al., 2014). Therefore, we explored the possibility, as suggested by our screen, that Fps1 is an authentic target of TORC2-dependent Ypk1-mediated phosphorylation, that this modification is important for Fps1 function, and that it is under regulation by hyperosmotic conditions.

Results

Ypk1 phosphorylates Fps1 and hyperosmotic shock inhibits this phosphorylation

The 743-residue enzyme Gpt2 contains one Ypk1 phospho-acceptor motif (646RSRSSSI652). At such sites, Ser residues just penultimate to the canonical one (in red) can be phosphorylated in a Ypk1-dependent manner (Roelants et al., 2011). Therefore, we generated a Gpt2(S649A S650A S651A) mutant. One or more of these three Ser residues is phosphorylated in vivo because, compared to wild-type, Gpt23A exhibited a distinctly faster mobility upon SDS-PAGE, a hallmark of decreased phosphorylation (Figure 1A), just like wild-type Gpt2 treated with phosphatase (Figure 1—figure supplement 1). However, this phosphorylation did not appear to be dependent on Ypk1 because little change occurred in Gpt2 mobility when an analog-sensitive ypk1-as ypk2Δ strain was treated with the cognate inhibitor (3-MB-PP1) (Figure 1A).

Figure 1 with 5 supplements see all
Fps1 (but not Gpt2) is phosphorylated by Ypk1.

(A) Wild-type (BY4741) or ypk1-as ypk2Δ (yAM135-A) cells expressing plasmid borne Gpt2-3xFLAG (pAX238) or Gpt23A-3xFLAG (pAX244) were grown to mid-exponential phase and then treated with vehicle (−) or 10 μM 3-MB-PP1 (+) for 90 min. Cells were harvested, extracts prepared, resolved by SDS-PAGE, and blotted as in ‘Materials and methods’. (B) Wild-type cells expressing either Fps1-3xFLAG (yGT21) or Fps13A-3xFLAG (yGT22) from the FPS1 promoter at the normal chromosomal locus, or ypk1-as ypk2Δ cells expressing either Fps1-3xFLAG (yAM281) or Fps13A-3xFLAG (yAM284-A) from the FPS1 promoter at the normal chromosomal locus, were grown to mid-exponential phase and treated as in (A) with vehicle or 3-MB-PP1 for 60 min. Cells were harvested, extracts prepared, resolved by Phos-tag SDS-PAGE, and blotted as in ‘Materials and methods’. Unphosphorylated Fps1 (red asterisk). (C) A tor2-as strain (yKL5) expressing Fps1-3xFLAG (pAX274) or Fps13A-3xFLAG (pAX275) was grown to mid-exponential phase and then treated with vehicle (−) or 2 μM BEZ-235 (+) for 30 min. Cells were harvested, extracts prepared, resolved and analyzed as in (B). (D) Wild-type (BY4741) or tor2-29ts (JTY5468) cells expressing Ypk17A-myc (pFR252) were grown at 30°C (left panel) or 26°C (right panel) to mid-exponential phase, then diluted into fresh YPD in the absence (−) or presence of 1 M sorbitol (final concentration). After the indicated times (1–15 min), culture samples were collected, lysed and the resulting extracts resolved by Phos-tag SDS-PAGE and analyzed by immunoblotting with anti-myc mAb 9E10, as described in ‘Materials and methods’. (E) As in (D), except for the genotype (strain) expressing Ypk17A-myc (pFR252), which were, aside from the wild-type control, hog1Δ (YJP544), sho1Δ (JTY5540), ssk1Δ (JTY5541), ssk22Δ (JTY5539), ssk2Δ (JTY5538) or pbs2Δ (JTY5537), and the treatment with 1 M sorbitol was for 1 min. (F) Wild-type (BY4741) or otherwise isogenic cna1∆ cna2∆ (JTY5574) cells expressing Ypk17A-myc (pFR252) were grown to mid-exponential phase then diluted into fresh YPD in the absence (−) or presence (+) of 1 M sorbitol (final concentration). After 1 min, the cells were collected, lysed and the resulting extracts resolved by Phos-tag SDS-PAGE and analyzed by immunoblotting with anti-myc mAb 9E10, as described in ‘Materials and methods’. (G) Wild-type cells expressing either Fps1-3xFLAG (yGT21) or Fps13A-3xFLAG (yGT22) from the chromosomal FPS1 locus, were diluted into fresh YPD in the absence (−) or presence of 1 M sorbitol (final concentration) for the indicated times and then extracts of the cells prepared and analyzed as in (B).

https://doi.org/10.7554/eLife.09336.002

In marked contrast, three of four predicted Ypk1 sites in the 669-residue Fps1 channel (176RRRSRSR182, 180RSRATSN186, 565RARRTSD571) (Figure 1—figure supplement 2A) are phosphorylated in vivo, as indicated by the effect of site-directed mutations to Ala on electrophoretic mobility (Figure 1—figure supplement 2B), and their phosphorylation requires Ypk1 activity, because, in inhibitor-treated ypk1-as ypk2Δ cells, the mobility of wild-type Fps1 was indistinguishable from that of Fps1(S181A S185A S570A) (Figure 1B), just like wild-type Fps1 treated with phosphatase (Figure 1—figure supplement 2C). Moreover, a C-terminal fragment of Fps1 containing Ser570, one of the apparent Ypk1 phosphorylation sites delineated in vivo, is phosphorylated by purified Ypk1 in vitro and solely at the Ypk1 site (S570) (Figure 1—figure supplement 3). Furthermore, as for other Ypk1-dependent modifications (Muir et al., 2014), phosphorylation of these same sites in Fps1 in vivo was also TORC2-dependent, because treatment with a TORC2 inhibitor (NVP-BEZ235) (Kliegman et al., 2013) also reduced Fps1 phosphorylation (Figure 1C). Thus, Fps1 is a bona fide Ypk1 substrate.

We documented elsewhere using Phos-tag gel mobility shift that Ypk1 phosphorylation at T662, one of its well-characterized TORC2 sites, is eliminated when cells are subjected to hyperosmotic shock for 10 min (Lee et al., 2012), and the same effect is observed using a specific antibody (Niles et al., 2012) that monitors phosphorylation of Ypk1 at the same site (Figure 1—figure supplement 4A). Using Ypk17A, which also permits facile detection of mobility shifts arising from TORC2-specific phosphorylation (K Leskoske and FM Roelants, unpublished results) (Figure 1—figure supplement 4B), we followed the kinetics of this change. Loss of TORC2-mediated Ypk1 phosphorylation upon hyperosmotic shock occurs very rapidly (within 1 min) and persists for about 15 min (Figure 1D), but is transient. By 20 min after hyperosmotic shock, TORC2-mediated Ypk1 phosphorylation is again detectable and is nearly back to the pre-stress level by 75 min (Figure 1—figure supplement 5A). Rapid reduction in TORC2-mediated Ypk1 phosphorylation under hypertonic stress was still observed in mutants lacking the Sho1- or Sln1-dependent pathways that converge on Hog1 or Hog1 itself (Figure 1E) or CN (Figure 1F). Thus, loss of TORC2-mediated Ypk1 phosphorylation upon hyperosmotic shock occurs independently of other known response pathways.

Given that Ypk1 phosphorylates Fps1 and that hyperosmotic stress rapidly abrogates TORC2-dependent phosphorylation and activation of Ypk1, Ypk1 modification of Fps1 should be prevented under hyperosmotic stress. As expected, Ypk1 phosphorylation of Fps1 is rapidly lost upon hyperosmotic shock (Figure 1G), yielding a species with mobility indistinguishable from Fps13A, remains low for at least 20 min, but returns by 75 min (Figure 1—figure supplement 5B), mirroring the kinetics of loss and return of both TORC2-mediated Ypk1 phosphorylation (Figure 1D and Figure 1—figure supplement 5A) and Ypk1-dependent phosphorylation of Gpd1 that we observed before (Lee et al., 2012). Thus, hyperosmotic stress dramatically down-modulates Ypk1-mediated phosphorylation of Fps1.

Ypk1 phosphorylation of Fps1 promotes channel opening and glycerol efflux

In its open state, the Fps1 channel permits entry of toxic metalloid, arsenite, which inhibits growth (Thorsen et al., 2006), whereas lack of Fps1 (fps1∆) or the lack of channel activators (rgc1∆ rgc2∆) (Beese et al., 2009) or an Fps1 mutant that cannot open because it cannot bind the activators (Fps1∆PHD) (Lee et al., 2013) are arsenite resistant. We found that Fps13A was at least as arsenite resistant as any other mutant that abrogates Fps1 function (Figure 2A). Thus, Fps13A acts like a closed channel, suggesting that Ypk1-mediated phosphorylation promotes channel opening. Loss of individual phosphorylation sites led to intermediate levels of arsenite resistance (Figure 2B). Thus, modification at these sites contributes additively to channel opening.

Phosphorylation by Ypk1 opens the Fps1 channel.

(A) Cultures of Fps1-3xFLAG (yGT21), Fps13A-3xFLAG (yGT22), Fps1ΔPHD-3xFLAG (yAM307-A), rgc1Δ rgc2Δ (DL3188) and fps1Δ (yAM181-A) were adjusted to A600 nm = 1.0 and serial dilutions were then spotted onto YPD plates containing the indicated concentration of arsenite. Cells were allowed to grow for 4 days at 30°C prior to imaging. (B) As in (A), except Fps1-3xFLAG (yGT21), Fps1(T147A)-3xFLAG (yAM310-A), Fps1(S181A S185A)-3xFLAG (yAM301-A), Fps1(S570A)-3xFLAG (yGT24) or Fps13A-3xFLAG (yGT22) cultures were used and cells were grown for 2 days at 30°C prior to imaging. (C) Triplicate exponentially-growing cultures of wild-type (BY4742), fps1Δ (yAM181-A), Fps1-3xFLAG (yGT21) and Fps13A-3xFLAG (yGT22) strains were harvested, extracted, and the glycerol and protein concentration measured as described in ‘Materials and methods’. Values represent the ratio of glycerol-to-protein (error bar, standard error of the mean). (D) Extracts from the strains in (B) were resolved by standard SDS-PAGE using 8% acrylamide gels. (E) fps1Δ (yAM181-A) cells expressing Fps1-GFP (pAX290), Fps1(S181A S185A)-GFP, (pAX294), Fps1(S570A)-GFP (pAX293) or Fps13A-GFP (pAX295) were viewed by fluorescence microscopy as described in ‘Materials and methods’. Representative fields are shown.

https://doi.org/10.7554/eLife.09336.008

Others have shown that intracellular glycerol is elevated in fps1∆ cells in the absence of hyperosmotic stress (Tamás et al., 1999). If Fps13A favors the closed-channel state, then it should also cause constitutive elevation of intracellular glycerol concentration. Indeed, in the absence of any osmotic perturbation, Fps13A mutant cells accumulated ∼twofold as much glycerol as otherwise isogenic FPS1+ strains (Figure 2C). Consistent with this result, we observed before that loss of Ypk1 (and Ypk2) activity caused an increase in glycerol level compared to control cells (Lee et al., 2012).

Consistent with Ypk1-dependent phosphorylation affecting Fps1 channel function per se, immunoblotting (Figure 2D) and fluorescence microscopy (Figure 2E) showed that the steady-state level and localization of Fps1 are unaffected by the presence or absence of these modifications.

Hyperosmotic stress-evoked down-regulation of Ypk1 phosphorylation of Fps1 promotes cell survival independently of known Fps1 regulators

Fps1 can be negatively regulated by Hog1 via two mechanisms: Hog1 phosphorylation of Fps1 stimulates its internalization and degradation (Thorsen et al., 2006; Mollapour and Piper, 2007); Hog1 phosphorylation closes the channel by displacing bound Fps1 activators (Rgc1 and Rgc2) (Beese et al., 2009; Lee et al., 2013). We found, however, that Fps13A was still in the closed state, as judged by arsenite resistance, in the total absence of Hog1 (hog1Δ) (Figure 3A), or in an Fps1 mutant (Fps1IVAA) that cannot bind Hog1 or where the activator cannot be displaced from Fps1 by Hog1 phosphorylation (Rgc27A) (Lee et al., 2013) (Figure 3B). Thus, closure of the Fps1 channel by lack of Ypk1 phosphorylation occurs independently of any effects requiring Hog1. Consistent with this conclusion, presence or absence of Ypk1-mediated Fps1 phosphorylation had no effect on Fps1-Rgc2 interaction (Figure 3C).

TOR Complex 2 (TORC2)-dependent Ypk1-mediated regulation of Fps1 is independent of Hog1 and Rgc1 and Rgc2.

(A) Cultures of Fps1-3xFLAG (yGT21), Fps1570A-3xFLAG (yGT24), Fps13A-3xFLAG (yGT22), Fps1-3xFLAG hog1Δ (yAM275), Fps1570A-3xFLAG hog1Δ (yAM291-A) and Fps13A-3xFLAG hog1Δ (yAM278) strains were adjusted to A600 nm = 1.0 and serial dilutions were then spotted onto YPD plates containing the indicated concentration of arsenite. Cells were allowed to grow for 2 days at 30°C prior to imaging. (B) As in (A), except Fps1IVAA-3xFLAG (yAM308-A), Fps1(3A)IVAA-3xFLAG (yAM309-A), Rgc27A-HA (yAM315) and Fps13A-3xFLAG Rgc27A-HA (yAM318) strains were tested. The Fps1IVAA mutation prevents Hog1 binding to and regulation of Fps1, and Rgc27A cannot be displaced from Fps1 because it cannot be phosphorylated by Hog1; both mutations render the channel constitutively open and make cells arsenite sensitive (Lee et al., 2013). (C) Fps1-3xFLAG (yAM271-A) or Fps13A-3xFLAG (yAM272-A) strains were co-transformed with PMET25-Rgc2-HA (p3151) and PMET25-Fps1-3xFLAG (pAX302) or PMET25-Fps13A -3xFLAG (pAX303) plasmids. After Rgc2-HA and Fps1-3xFLAG expression, Fps1 was immuno-purified with anti-FLAG antibody-coated beads (see ‘Materials and methods’). The bound proteins were resolved by SDS-PAGE and the amount of Rgc2-HA present determined by immunoblotting with anti-HA antibody. (D) Wild-type (BY4741), hog1Δ (YJP544) or Fps13A-3xFLAG hog1Δ (yAM278) strains were grown and serial dilutions of these cultures plated onto synthetic complete medium lacking tryptophan with 2% dextrose and the indicated concentration of sorbitol. Cells were grown for 3 days prior to imaging.

https://doi.org/10.7554/eLife.09336.009

Collectively, our results show that, independently of Hog1, hypertonic conditions drastically diminish TORC2-dependent Ypk1 phosphorylation, in turn dramatically decreasing Ypk1-mediated Fps1 phosphorylation, thereby closing the channel and causing intracellular glycerol accumulation. Thus, absence of Ypk1 phosphorylation should allow a cell lacking Hog1 to better survive hyperosmotic conditions. Indeed, Fps13A hog1Δ cells are significantly more resistant to hyperosmotic stress than otherwise isogenic hog1Δ cells (Figure 3D). This epistasis confirms that, even when Hog1 is absent, loss of Ypk1-mediated Fps1 channel opening is sufficient for cells to accumulate an adequate amount of glycerol to physiologically cope with hyperosmotic stress.

Discussion

Aside from further validating the utility of our screen for identifying new Ypk1 substrates (Muir et al., 2014), our current findings demonstrate that TORC2-dependent Ypk1-catalyzed phosphorylation of Fps1 opens this channel and, conversely, that loss of Ypk1-dependent Fps1 phosphorylation upon hypertonic shock is sufficient to close the channel, prevent glycerol efflux, and promote cell survival. In agreement with our observations, in a detailed kinetic analysis of global changes in the S. cerevisiae phosphoproteome upon hyperosmotic stress (Kanshin et al., 2015), it was noted that two sites in Fps1 (S181 and T185), which we showed here are modified by Ypk1, become dephosphorylated.

We previously showed that Gpd1, the rate-limiting enzyme for glycerol production under hyperosmotic conditions (Remize et al., 2001), is negatively regulated by Ypk1 phosphorylation (Lee et al., 2012). Thus, inactivation of TORC2-Ypk1 signaling upon hyperosmotic shock has at least two coordinated consequences that work synergistically to cause glycerol accumulation and promote cell survival, a similar outcome but mechanistically distinct from the processes evoked by Hog1 activation (Figure 4). First, loss of TORC2-Ypk1 signaling alleviates inhibition of Gpd1, which, combined with transcriptional induction of GPD1 by hyperosmotic stress, greatly increases glycerol production. Second, loss of TORC2-Ypk1 signaling closes the Fps1 channel, thereby retaining the glycerol produced.

Saccharomyces cerevisiae has two independent sensing systems to rapidly increase intracellular glycerol upon hyperosmotic stress.

(A) Hog1 MAPK-mediated response to acute hyperosmotic stress (adapted from Hohmann, 2015). Unstressed condition (top), Hog1 is inactive and glycerol generated as a minor side product of glycolysis under fermentation conditions can escape to the medium through the Fps1 channel maintained in its open state by bound Rgc1 and Rgc2. Upon hyperosmotic shock (bottom), pathways coupled to the Sho1 and Sln1 osmosensors lead to Hog1 activation. Activated Hog1 increases glycolytic flux via phosphorylation of Pkf26 in the cytosol and, on a longer time scale, also enters the nucleus (not depicted) where it transcriptionally upregulates GPD1 (de Nadal et al., 2011; Saito and Posas, 2012), the enzyme rate-limiting for glycerol formation, thereby increasing glycerol production. Activated Hog1 also prevents glycerol efflux by phosphorylating and displacing the Fps1 activators Rgc1 and Rgc2 (Lee et al., 2013). These processes act synergistically to elevate the intracellular glycerol concentration providing an osmolyte to counterbalance the external high osmolarity. (B) Unstressed condition (top), active TORC2-Ypk1 keeps intracellular glycerol level low by inhibition of Gpd1 (Lee et al., 2012) and because Ypk1-mediated phosphorylation promotes the open state of the Fps1 channel. Upon hyperosmotic shock (bottom), TORC2-dependent phosphorylation of Ypk1 is rapidly down-regulated. In the absence of Ypk1-mediated phosphorylation, inhibition of Gpd1 is alleviated, thereby increasing glycerol production. Concomitantly, loss of Ypk1-mediated phosphorylation closes the Fps1 channel, even in the presence of Rgc1 and Rgc2, thereby promoting glycerol accumulation to counterbalance the external high osmolarity. Schematic depiction of TORC2 based on data from Wullschleger et al. (2005); Liao and Chen (2012); Gaubitz et al. (2015).

https://doi.org/10.7554/eLife.09336.010

Presence of two systems (TORC2-Ypk1 and Hog1) might allow cells to adjust optimally to stresses occurring with different intensity, duration, or frequency. Reportedly, Hog1 responds to stresses occurring no more frequently than every 200 s (Hersen et al., 2008; McClean et al., 2009), whereas we found TORC2-Ypk1 signaling responded to hypertonic stress in ≤60 s. Also, the Sln1 and Sho1 sensors that lead to Hog1 activation likely can respond to stimuli that do not affect the TORC2-Ypk1 axis, and vice-versa.

A remaining question is how hyperosmotic stress causes such a rapid and profound reduction in phosphorylation of Ypk1 at its TORC2 sites. This outcome could arise from activation of a phosphatase (other than CN), inhibition of TORC2 catalytic activity, or both. Despite a recent report that Tor2 (the catalytic component of TORC2) interacts physically with Sho1 (Lam et al., 2015), raising the possibility that a Hog1 pathway sensor directly modulates TORC2 activity, we found that hyperosmolarity inactivates TORC2 just as robustly in sho1Δ cells as in wild-type cells. Alternatively, given the role ascribed to the ancillary TORC2 subunits Slm1 and Slm2 (Gaubitz et al., 2015) in delivering Ypk1 to the TORC2 complex (Berchtold et al., 2012; Niles et al., 2012), response to hyperosmotic shock might be mediated by some influence on Slm1 and Slm2. Thus, although the mechanism that abrogates TORC2 phosphorylation of Ypk1 upon hypertonic stress remains to be delineated, this effect and its consequences represent a novel mechanism for sensing and responding to hyperosmolarity.

Materials and methods

Construction of yeast strains and growth conditions

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S. cerevisiae strains used in this study (Supplementary file 1) were constructed using standard yeast genetic manipulations (Amberg et al., 2005). For all strains constructed, integration of each DNA fragment of interest into the correct genomic locus was assessed using genomic DNA from isolated colonies of corresponding transformants as the template and PCR amplification with an oligonucleotide primer complementary to the integrated DNA and a reverse oligonucleotide primer complementary to chromosomal DNA at least 150 bp away from the integration site, thereby confirming that the DNA fragment was integrated at the correct locus. Finally, the nucleotide sequence of each resulting reaction product was determined to confirm that it had the correct sequence. Yeast cultures were grown in rich medium (YPD; 1% yeast extract, 2% peptone, 2% glucose) or in defined minimal medium (SCD; 0.67% yeast nitrogen base, 2% glucose) supplemented with the appropriate nutrients to permit growth of auxotrophs and/or to select for plasmids.

Plasmids and recombinant DNA methods

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All plasmids used in this study (Supplementary file 2) were constructed using standard laboratory methods (Green and Sambrook, 2012) or by Gibson assembly (Gibson et al., 2009) using the Gibson Assembly Master Mix Kit according to the manufacturer's specifications (New England Biolabs, Ipswich, Massachusetts, United States). All constructs generated in this study were confirmed by nucleotide sequence analysis covering all promoter and coding regions in the construct.

Preparation of cell extracts and immunoblotting

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Yeast cell extracts were prepared by an alkaline lysis and trichloroacetic acid (TCA) precipitation method, as described previously (Westfall et al., 2008). For samples analyzed by immunoblotting, the precipitated proteins were resolubilized and resolved by SDS-PAGE, as described below. For samples subjected to phosphatase treatment, the precipitated proteins were resolubilized in 100 µl solubilization buffer (2% SDS, 2% β-mercaptoethanol, 150 mM NaCl, 50 mM Tris-HCl [pH 8.0]), diluted with 900 µl calf intestinal phosphatase dilution buffer (11.1 mM MgCl2, 150 mM NaCl, 50 mM Tris-HCl [pH 8.0]), incubated with calf intestinal alkaline phosphatase (350 U; New England Biolabs) for 4 hr at 37°C, recollected by TCA precipitation, resolved by SDS-PAGE, and analyzed by immunobotting. To resolve Gpt2 and its phosphorylated isoforms, samples (15 μl) of solubilized protein were subjected to SDS-PAGE at 120 V in 8% acrylamide gels polymerized and crosslinked with a ratio of acrylamide:bisacrylamide::75:1. To resolve Fps1 and Ypk1 and their phosphorylated isoforms, samples (15 μl) of solubilized protein were subjected to Phos-tag SDS-PAGE (Kinoshita et al., 2009) (8% acrylamide, 35 µM Phos-tag [Wako Chemicals USA, Inc.], 35 µM MnCl2) at 160 V.

After SDS-PAGE, proteins were transferred to nitrocellulose and incubated with mouse or rabbit primary antibody in Odyssey buffer (Li-Cor Biosciences, Lincoln, Nebraska, United States), washed, and incubated with appropriate IRDye680LT-conjugated or IRDye800CW-conjugated anti-mouse or anti-rabbit IgG (Li-Cor Biosciences) in Odyssey buffer with 0.1% Tween-20 and 0.02% SDS. Blots were imaged using an Odyssey infrared scanner (Li-Cor Biosciences). Primary antibodies and dilutions used were: rabbit anti-HA, 1:1000 (Covance Inc., Dedham, Massachusetts, United States); mouse anti-HA, 1:1000 (Covance Inc.); mouse anti-FLAG, 1:5000 (Sigma–Aldrich, St. Louis, Missouri, United States); rabbit anti-FLAG, 1:5000 (Sigma–Aldrich); tissue culture medium containing mouse anti-c-myc mAb 9E10, 1:100 (Monoclonal Antibody Facility, Cancer Research Laboratory, University of California, Berkeley); rabbit anti-Ypk1(P-T662), 1:20,000 (generous gift from Ted Powers, University of California, Davis); and, rabbit anti-yeast Pgk1, 1:10,000 (this laboratory).

Protein purification and in vitro kinase assay

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Ypk1 and GST-Fps1(531-0669) proteins were purified as previously described (Muir et al., 2014). Following protein purification, Ypk1 in vitro kinase assays were performed as previously described (Muir et al., 2014).

Measurement of intracellular glycerol accumulation

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Measurement of intracellular glycerol was conducted as described (Albertyn et al., 1994a). Briefly, samples (∼40 ml) of exponentially-growing cultures were harvested by centrifugation, washed with 1 ml of medium, recollected and the resulting cell pellets frozen in liquid N2 and stored at −80°C prior to analysis. Each cell pellet was boiled for 10 min in 1 ml of 50 mM Tris-Cl (pH 7.0). This eluate was clarified by centrifugation for 15 min at 13,200 rpm (16,100×g) in a microfuge (Eppendorf 5415D). Glycerol concentration in the resulting supernatant fraction was measured using a commercial enzymic assay kit (Sigma Aldrich) and normalized to the protein concentration of the same initial extract as measured by the Bradford method (Bradford, 1976).

Fluorescence microscopy of Fps1-GFP

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An fps1Δ strain was transformed with plasmids expressing wild-type Fps1-GFP or the mutant Fps1-GFP derivatives and grown in selective medium to mid-exponential phase. Samples of the resulting cultures were viewed directly under an epifluorescence microscope (model BH-2; Olympus America, Inc.) using a 100× objective fitted with appropriate band-pass filters (Chroma Technology Corp.). Images were collected using a CoolSNAP MYO charge-coupled device camera (Photometrics, Tucson, Arizona, United States).

Co-immunoprecipitation of Fps1 and Rgc2

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Co-immunoprecipitation experiments were performed with minor modifications as previously described (Lee et al., 2013). Cells expressing Fps1-3xFLAG (yAM271-A), Fps13A-3xFLAG (yAM272-A) or untagged Fps1 (BY4742) were transformed with empty vector or the same vector expressing Fps1-3xFLAG (pAX302) or Fps13A-3xFLAG (pAX303) under control of the MET25 promoter. These transformants were then co-transformed with a plasmid expressing Rgc2-3xHA under control of the MET25 promoter (Lee et al., 2013). Cultures of each were grown to mid-exponential phase in SCD-Ura-Leu. Cultures were then diluted to A600 nm = 0.2 in 1 l of SCD-Ura-Leu-Met to induce expression of Rgc2-3xHA and Fps1-3xFLAG and grown at 30°C for 4 hr. Cells were harvested by centrifugation and resuspended in 5 ml of TNE+Triton+NP-40 (50 mM Tris-Cl [pH 7.5], 150 mM NaCl, 4 mM NaVO4, 50 mM NaF, 20 mM Na-PPi, 5 mM EDTA, 5 mM EGTA, 0.5% Triton-X100, 1.0% NP-40, 1× cOmplete protease inhibitor [Roche, Pleasanton, California, United States]). The cells were then lysed cryogenically using Mixer Mill MM301 (Retsch GmbH, Haan, Germany). The lysate was thawed on ice and then clarified by centrifugation for 20 min at 10,500 rpm (13,000×g) in the SS34 rotor of a refrigerated centrifuge (Sorvall RC-5B). Protein concentration of the clarified lysate was measured using BCA reagent (Thermo Fisher Scientific, Waltham, Massachusetts, United States) and then Fps1-3xFLAG was immunoprecipitated from a volume of extract containing a total of 10 mg protein using 50 μl of mouse anti-FLAG antibody coupled-agarose resin (Sigma Aldrich) equilibrated in TNE+Triton+NP-40. Binding was allowed to occur for 2 hr at 4°C. The resin was then washed extensively with TNE+Triton+NP-40 and the proteins remaining bound were then resolved by SDS-PAGE and analyzed by immunoblotting with appropriate antibodies to detect both Fps1-3xFLAG and Rgc2-3xHA.

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Decision letter

  1. Tony Hunter
    Reviewing Editor; Salk Institute, United States

eLife posts the editorial decision letter and author response on a selection of the published articles (subject to the approval of the authors). An edited version of the letter sent to the authors after peer review is shown, indicating the substantive concerns or comments; minor concerns are not usually shown. Reviewers have the opportunity to discuss the decision before the letter is sent (see review process). Similarly, the author response typically shows only responses to the major concerns raised by the reviewers.

Thank you for submitting your work entitled “Down-regulation of TORC2-Ypk1 signaling promotes MAPK-independent survival under hyperosmotic stress” for peer review at eLife. Your submission has been favorably evaluated by Tony Hunter (Senior editor) and two reviewers. One of the two reviewers, Susan Henry, has agreed to share her identity.

The reviewers have discussed the reviews with one another, and the Reviewing editor has drafted this decision to help you prepare a revised submission.

The reviewers were interested in your elucidation of a new TORC2-Ypk1-Fps1 pathway that serves as a Hog1-independent backup mechanism for cell survival in response to hyperosmotic stress acting by closing the Fps1 channel for glycerol efflux, resulting in glycerol accumulation and cell survival during hyperosmotic stress, and they are both in favor of publication in eLife with revision.

Please address the specific points raised by Reviewer 2, which all appear feasible to do within a short time frame.

Reviewer 1:

Prior work from this group had shown that phosphorylation of the Ypk1 protein kinase at its TORC2-dependent sites is eliminated rapidly during hyperosmotic shock but that this loss is attenuated within 10-15 min, but this attenuation was found to be independent of known stress response pathways, including the Hog1 MAPK. Their previous proteomic screen had also identified Fps1p as being a target of the Ypk1p protein kinase.

This current paper provides strong and clear evidence that the aquaglyceroporin Fps1p is a bona fide target of Ypk1, a TORC2-dependent protein kinase, upon exposure of yeast cells to hypertonic stress. This current work also demonstrates for the first time that the open channel state of Fps1 is dependent on phosphorylation by Ypk1p. In addition, and importantly, the authors show that in wild-type cells, under hyperosmotic stress conditions, TORC2-dependent phosphorylation of Fps1 by Ypk1 is blocked. They show that this results in closure of the Fps1 channel for glycerol efflux, resulting in glycerol accumulation during hyperosmotic stress, which in turn leads to increased cell survival under these conditions.

The evidence presented also indicates that these responses are not dependent on the Hog1 MAPK pathway, which is also activated during hypertonic stress. Thus, this work identifies a novel mechanism involved in survival under hypertonic stress that is Hog1 independent, but TORC2-dependent.

Overall, the data are of high quality and importance and reveal a novel mechanism.

Reviewer 2:

Muir et al. follow up a previous genome-wide screen in which they identified potential Ypk1 substrates. They now confirm that aquaglyceroporin (Fps1) is a target of Ypk1. The authors demonstrate that TORC2-Ypk1 signaling initiates Fps1 phosphorylation to promote Fps1 channel opening. Phospho-deficient Fps1 is in a closed state and confers resistance to hyperosmotic stress and arsenite. They also show that hyperosmotic stress inhibits TORC2-Ypk1 signaling.

1) It has been reported already that the Slm proteins activate TORC2-Ypk1 signaling in response to hypo-osmotic stress (Berchtold et al. Nat Cell Biol. 14:542, 2012). Do the Slm proteins inhibit TORC2-Ypk1 in response to hyperosmotic stress? Also, the Berchtold et al. studies should be discussed.

2) Figure 1A-C: The authors show that TORC2-Ypk1 signaling promotes phosphorylation of Fps1 on three serine residues (Ser181, Ser 185, Ser570) as determined by resolving “phospho-”Fps1 in phos-tag gels. The phos-tag gel assay requires a phosphatase-treated control to claim that that the slow-migrating band is indeed phospho-Fps1. Figure 1A (experiment for Gpt2 phosphorylation) also needs a phosphatase-treated control.

3) Figure 1C: The authors demonstrate that acute inactivation of TORC2, by BEZ235 treatment of an analog-sensitive tor2 (tor2-as) mutant, reduces Fps1 phosphorylation. Is there a positive control to show that TORC2 activity is indeed inhibited? The authors could possibly look at TORC1 activity (e.g. Sch9 phosphorylation), since TOR2 is also part of TORC1.

4) Figure 1D: The authors show that TORC2 activity is inhibited upon sorbitol treatment, as determined by mobility shift of Ypk1 in a phos-tag gel. For this experiment, they used a phospho-deficient mutant of Ypk1 in which bona fide TORC2 target sites (Ser644 and Thr662) are mutated. This suggests that the slow migrating form of Ypk1 (this also needs phosphatase treatment) may be phosphorylated on yet to be characterized sites and it is these sites that the authors are monitoring. A more reliable assay of TORC2 activity would be to blot with phospho-Ypk1 antibody (Thr662) (Berchtold et al. Nat Cell Biol. Op. cit.).

5) The authors state: “Thus, Fps1 is a bona fide Ypk1 substrate.” To make this claim, they should provide evidence that Ypk1 directly phosphorylates Fps1. This can be done by performing a Ypk1 kinase assay in vitro with recombinant Fps1-WT and Fps1-3A. The authors have previously performed in vitro Ypk1 assays.

6) Figure 2A: The authors show that phospho-deficient Fps1-3A cells are the most arsenite resistant cells among fps1 mutants tested. They concluded that the Fps13A mutant is a closed channel such that toxic arsenite cannot enter cells. Why are Fps13A mutant cells more resistant than fps1∆ cells or Fps1-∆PHD cells in which Fps1 channel is constantly absent or closed, respectively?

7) Figure 2A-B: Based on data showing that Fps13A mutant cells are resistant to arsenite, the authors conclude that Ypk1-mediated phosphorylation promotes Fps1 channel opening. This conclusion would be strengthened if the authors also demonstrated that a phospho-mimetic mutant of Fps1 (Fps13D or Fps13E) confers arsenite hyper-sensitivity.

https://doi.org/10.7554/eLife.09336.013

Author response

The reviewers were interested in your elucidation of a new TORC2-Ypk1-Fps1 pathway that serves as a Hog1-independent backup mechanism for cell survival in response to hyperosmotic stress acting by closing the Fps1 channel for glycerol efflux, resulting in glycerol accumulation and cell survival during hyperosmotic stress, and they are both in favor of publication in eLife with revision.

Please address the specific points raised by Reviewer 2, which all appear feasible to do within a short time frame.

We are very pleased that the editor and the two referees are in favor of publication of our Research Advance in eLife, pending appropriate revision in response to the remarks of Reviewer #2. In this regard, and as enumerated below, several of the issues raised by Reviewer #2 are already addressed in prior published work from ourselves and others, and certain other issues are addressed by results now provided in the revised manuscript (which were not originally included in the initially submitted manuscript because of the length limitations of the Research Advance format). Some of the newly added data were generated by graduate student Kristin L. Leskoske and, hence, she has now been added as a co-author of this study.

Reviewer 2:

Muir et al. follow up a previous genome-wide screen in which they identified potential Ypk1 substrates. They now confirm that aquaglyceroporin (Fps1) is a target of Ypk1. The authors demonstrate that TORC2-Ypk1 signaling initiates Fps1 phosphorylation to promote Fps1 channel opening. Phospho-deficient Fps1 is in a closed state and confers resistance to hyperosmotic stress and arsenite. They also show that hyperosmotic stress inhibits TORC2-Ypk1 signaling.

We thank this referee for the generally correct synopsis of the novel findings reported in our study. However, this referee felt that we needed to better address and/or experimentally document several of our findings. Our disposition of those concerns is described below.

1) It has been reported already that the Slm proteins activate TORC2-Ypk1 signaling in response to hypo-osmotic stress (Berchtold et al. Nat Cell Biol. 14:542, 2012). Do the Slm proteins inhibit TORC2-Ypk1 in response to hyperosmotic stress? Also, the Berchtold et al. studies should be discussed.

The main focus of this Research Advance was to demonstrate unequivocally that Fps1 is indeed an authentic in vivo substrate of the TORC2-Ypk1 signaling axis and to elucidate the physiologically role of these post -translational modifications. Moreover, given the space limitations of a Research Advance, we feel that it is well beyond the scope of this report to demand that we also elucidate the mechanism by which hyperosmotic shock ablates transiently the TORC2-dependent phosphorylation of Ypk1, and whether the Slm1 and Slm2 proteins may or may not be involved at this level of regulation.

Nonetheless, in this same regard, existing data in the literature indicate that it would be technically very difficult to directly address the question posed by Reviewer #2 about whether the Slm1 (and/or Slm2) proteins are required for the observed down-regulation of TORC2-Ypk1 signaling under hyperosmotic conditions, for the following reasons. There are only two known conditions under which slm1∆slm2∆ cells are viable and in which, theoretically, we could ask whether or not 1 M sorbitol is still able to cause a drop in TORC2-mediated phosphorylation of Ypk1 when the Slm1 and Slm2 proteins are absent.

First, a slm1∆slm2∆sac7∆ triple mutant is viable, as shown by work from Scott Emr's lab [Audhya A et al. (2004) EMBO J. 23: 3747 -3757]. SAC7 encodes a primary GAP for yeast Rho1 and, as shown in collaborative work by Yoshi Ohya and David Levin [Kamada Y et al. (1996) J. Biol. Chem. 271: 9193-9196], Rho1 is the activator of yeast Pkc1 [misnamed because it really is more related to the mammalian Rho- (and Rac-) activated PKR2/PKNγ; see Vincent S, Settleman J (1997) Mol. Cell. Biol. 17: 2247-2256; Mukai H (2003) J. Biochem. (Tokyo) 133: 17-27]. In any event, loss of Sac7 up-regulates Pkc1 signaling and, as we demonstrated before [Roelants FM et al. (2002) Mol. Biol. Cell 13: 3005 -3028], any perturbation that up-regulates Pkc1 function bypasses the inviability of cells deficient in Ypk1 and Ypk2 signaling, presumably because some aspects of the cell wall integrity pathway act in parallel or are semi-redundant with functions normally controlled by Ypk1 and Ypk2. The problem arises, however, from observations reported by Ted Powers lab [Niles BJ et al. (2012) PNAS 109: 1536–1541]. They examined the state of Ypk1 phosphorylation in slm1∆slm2∆sac7∆ triple mutant and found that Ypk1 phosphorylation at both its activation loop (T504) and at one of its critical TORC2 sites (T662), as judged for the latter using a phospho-site-specific antibody they themselves generated, were greatly reduced. Thus, because TORC2- mediated phosphorylation is already way down in this background, it would be very challenging and probably not meaningful to examine whether the level of T662 phosphorylation goes down further in response to challenge with 1 M sorbitol.

Second, a slm1∆slm2∆ double mutant expressing a constitutively-active allele, Ypk1(D242A), is also able to grow. However, this allele is able to bypass the need for functional TORC2, as judged by its ability to rescue the lethality of a tor2ts mutant or treatment of cells with a TORC2 inhibitor [Roelants FM et al. (2011) PNAS 108: 19222-19227] or avo3∆ cells, which are deficient in TORC2 activity [Niles, op. cit.]. However, the same problem arises here too because, as judged by using exactly the same antibody (generously provided by Ted Powers), we have found that T662 phosphorylation is virtually undetectable on Ypk1(D242A) in slm1∆slm2∆ cells (K. Leskoske, unpublished data). Thus, once again, because TORC2-mediated phosphorylation is already way down in this background, it would be very challenging and probably not meaningful to examine whether the level of T662 phosphorylation goes down further in response to challenge with 1 M sorbitol.

Finally, although the lethality of Slm1- and Slm2-deficient cells clearly arises from a lack of sufficient Ypk1 function, exactly why Slm1- and Slm2-deficient cells have low apparent TORC2-mediated phosphorylation of Ypk1 is not clear. Slm1 and Slm2 were identified originally as PP2B/calcineurin-binding proteins nearly contemporaneously in three different labs [Bultynck G et al. (2006) Mol. Cell. Biol. 26: 4729-4745; Tabuchi M et al. (2006) Mol. Cell. Biol. 26 : 5861 -5875; Daquinag A et al. (2007) Mol. Cell. Biol. 27: 633-650]. Hence, in the absence of Slm1 and Slm2, there will be a lack of proper sequestration of PP2B. Thus, the low level of Ypk1 phosphorylation observed when Slm1 and Slm2 are absent could arise, potentially, from an enhanced rate of dephosphorylation of Ypk1 itself or of some site in a component of TORC2 critical for its activity on Ypk1, or both. Alternatively, because Slm1 and Slm2 contain lipid-binding PH domains near their C-terminus [Yu JW et al. (2004) Mol. Cell 13 : 677-688; Gallego O et al. (2010) Mol. Syst. Biol. 6: 430.1-430.15] and both are considered subunits of TORC2 [Gaubitz C et al. (2015) Mol. Cell 58: 977- 988] it is possible that, in the absence of Slm1 and Slm2, inefficient or improper membrane tethering of TORC2 is responsible for the reduction in Ypk1 phosphorylation observed in slm1∆slm2∆ cells. Neither we nor any other yeast researcher has yet adequately resolved these issues. Nonetheless, as requested by Reviewer #2, we have added a sentence to the Discussion pointing out that it is possible that the response to hyperosmotic shock might be mediated by some influence on the Slm1 and Slm2 proteins, and we cite the Berchtold et al. reference in this context (although the Berchtold paper was already cited in our original manuscript).

2) Figure 1A-C: The authors show that TORC2-Ypk1 signaling promotes phosphorylation of Fps1 on three serine residues (Ser181, Ser 185, Ser570) as determined by resolving “phospho-”Fps1 in phos-tag gels. The phos-tag gel assay requires a phosphatase-treated control to claim that that the slow-migrating band is indeed phospho-Fps1. Figure 1A (experiment for Gpt2 phosphorylation) also needs a phosphatase-treated control.

We showed, using Phos-tag™ gels, that the migration of Fps13A in vivo exactly mirrors the collapse of isoforms of wild-type Fps1 observed in vivo upon inhibition of Ypk1 function. Thus, the inescapable conclusion is that Fps1 is indeed phosphorylated at these three sites in the cell and that Ypk1 is the enzyme responsible for these phosphorylations in vivo. Nonetheless, as requested by Reviewer #2, we now provide new data (new Figure 1–figure supplement 2C) that directly address this issue by showing that CIP treatment of WT Fps1 collapses the isoforms to a migration pattern identical to Fps13A. As also requested by Reviewer #2, the same analysis is now also provided for Gpt2 (new Figure 1–figure supplement 1), even though it is not the primary focus of the findings presented in this paper.

3) Figure 1C: The authors demonstrate that acute inactivation of TORC2, by BEZ235 treatment of an analog-sensitive tor2 (tor2-as) mutant, reduces Fps1 phosphorylation. Is there a positive control to show that TORC2 activity is indeed inhibited? The authors could possibly look at TORC1 activity (e.g. Sch9 phosphorylation), since TOR2 is also part of TORC1.

Data fully documenting the efficacy and specificity of the yeast tor2-as allele for inhibition by NVP-BEZ235 were already published by the lab of Kevan Shokat [Kliegman JI et al. (2013) Cell Rep. 5: 1725-173]. We see no necessity to recapitulate those experiments, especially because our results indicate that the inhibitor reduces phosphorylation of Fps1 at its Ypk1 sites, as expected because Ypk1 activity is TORC2-dependent, fully consistent with the conclusions of Kliegman [op. cit.].

4) Figure 1D: The authors show that TORC2 activity is inhibited upon sorbitol treatment, as determined by mobility shift of Ypk1 in a phos-tag gel. For this experiment, they used a phospho-deficient mutant of Ypk1 in which bona fide TORC2 target sites (Ser644 and Thr662) are mutated. This suggests that the slow migrating form of Ypk1 (this also needs phosphatase treatment) may be phosphorylated on yet to be characterized sites and it is these sites that the authors are monitoring. A more reliable assay of TORC2 activity would be to blot with phospho-Ypk1 antibody (Thr662) (Berchtold et al. Nat Cell Biol. Op. cit.).

As will be described in detail in another manuscript (Leskoske K, Roelants FM and Thorner J), TORC2 does indeed phosphorylate Ypk1 both in vivo and in vitro at three additional sites aside from its classical “turn” (S644) and “hydrophobic” (T662) motifs. So, the isoforms generated on Phos-tag™ gels by the Ypk17A mutant used are a reliable reporter of its TORC2-mediated modification. Nonetheless, as requested by Reviewer #2, we now provide new data (new Figure 1–figure supplement 4A) that directly address this issue by showing using the reagent recommended by the referee, namely anti-P-T662 site antibody (again, generously provided by Ted Powers), that phosphorylation at this site is rapidly and completely abrogated when cells are treated with 1 M sorbitol, in agreement with our independent analysis using Ypk17A. In addition, we now demonstrate by phosphatase treatment that the observed isoforms are indeed due to phosphorylation (new Figure 1–figure supplement 4B), as also requested by Reviewer #2.

5) The authors state: “Thus, Fps1 is a bona fide Ypk1 substrate.” To make this claim, they should provide evidence that Ypk1 directly phosphorylates Fps1. This can be done by performing a Ypk1 kinase assay in vitro with recombinant Fps1-WT and Fps1-3A. The authors have previously performed in vitro Ypk1 assays.

Like other demonstrated targets of Ypk1 (e.g. Orm1, Orm2, Lac1, Lag1), Fps1 is a polytopic integral membrane protein. Hence, as for these other substrates, we use large soluble fragments of the protein that contain its cytosolically disposed domains as the phospho-acceptor in in vitro kinase reactions. Thus, as requested by Reviewer #2, and as an illustrative example, we now provide new data (new Figure 1–figure supplement 3) that directly address this issue by documenting that a large (139-residue) fragment of Fps1 containing one of the three primary Ypk1 phosphorylation sites observed in vivo (S570) is indeed phosphorylated in Ypk1-specific manner in vitro and solely at this Ypk1 site (S570), as we previously described as “data not shown” [Muir A et al. (2014) eLife 3: e03779.1-e03779.34].

6) Figure 2A: The authors show that phospho-deficient Fps1-3A cells are the most arsenite resistant cells among fps1 mutants tested. They concluded that the Fps13A mutant is a closed channel such that toxic arsenite cannot enter cells. Why are Fps13A mutant cells more resistant than fps1∆ cells or Fps1-∆PHD cells in which Fps1 channel is constantly absent or closed, respectively?

The particular image provided to illustrate that the Fps13A mutant confers arsenite resistance was included because, as Reviewer #2 noted, it contained a full set of controls. Moreover, we have done the comparison multiple times and Fps13A-expressing cells do grow slightly better than cells totally lacking Fps1 (fps1∆ mutant) on both YPD alone and YPD + arsenite. The modest, but measurable, decrease in vegetative growth rate caused by the complete absence of Fps1 has been noted previously by others [Yoshikawa K et al. (2011) Yeast 28: 349-361; Lourenco AB et al. (2013) PLoS One 8: e55439.1-e55439.12; Marek A, Korona R (2013) Evolution 67 : 3077-3086]. Presumably, retention of this channel [∼1,000 copies per cell; Ghaemmaghami S et al. (2003) Nature 425: 737-741], even in closed form, preserves some aspect of cell function that becomes slightly compromised when this integral plasma membrane protein is missing.

7) Figure 2A-B: Based on data showing that Fps13A mutant cells are resistant to arsenite, the authors conclude that Ypk1-mediated phosphorylation promotes Fps1 channel opening. This conclusion would be strengthened if the authors also demonstrated that a phospho-mimetic mutant of Fps1 (Fps13D or Fps13E) confers arsenite hyper-sensitivity.

We tested, of course, whether an Fps13E mutant, which could possibly mimic the “persistently phosphorylated” and thus the “permanently open state” of the channel, might confer a degree of arsenite sensitivity equivalent to or even greater than that conferred by WT Fps1. In this instance, however, we found that an Fps13E mutant exhibited arsenite resistance, comparable to that displayed by Fps13A. Therefore, it seems that, in the case of this particular protein and these particular sites, Ser-to-Glu mutations are not an adequate mimic for authentic phosphate groups.

https://doi.org/10.7554/eLife.09336.014

Article and author information

Author details

  1. Alexander Muir

    1. Division of Biochemistry, Biophysics and Structural Biology, Department of Molecular and Cell Biology, University of California, Berkeley, Berkeley, United States
    2. Chemical Biology Graduate Program, University of California, Berkeley, Berkeley, United States
    Present address
    Vander Heiden Lab, Department of Biology and Koch Institute for Integrative Cancer Research, Massachusetts Institute of Technology, Cambridge, United States
    Contribution
    AM, Conception and design, Acquisition of data, Analysis and interpretation of data, Drafting or revising the article
    Competing interests
    The authors declare that no competing interests exist.
  2. Françoise M Roelants

    Division of Biochemistry, Biophysics and Structural Biology, Department of Molecular and Cell Biology, University of California, Berkeley, Berkeley, United States
    Contribution
    FMR, Conception and design, Acquisition of data, Analysis and interpretation of data, Drafting or revising the article
    Competing interests
    The authors declare that no competing interests exist.
  3. Garrett Timmons

    Department of Chemistry, University of California, Berkeley, Berkeley, United States
    Contribution
    GT, Conception and design, Acquisition of data, Drafting or revising the article
    Competing interests
    The authors declare that no competing interests exist.
  4. Kristin L Leskoske

    Division of Biochemistry, Biophysics and Structural Biology, Department of Molecular and Cell Biology, University of California, Berkeley, Berkeley, United States
    Contribution
    KLL, Acquisition of data, Drafting or revising the article
    Competing interests
    The authors declare that no competing interests exist.
  5. Jeremy Thorner

    Division of Biochemistry, Biophysics and Structural Biology, Department of Molecular and Cell Biology, University of California, Berkeley, Berkeley, United States
    Contribution
    JT, Conception and design, Analysis and interpretation of data, Drafting or revising the article
    For correspondence
    jthorner@berkeley.edu
    Competing interests
    The authors declare that no competing interests exist.

Funding

National Institute of General Medical Sciences (NIGMS) (T32 GM07232)

  • Alexander Muir
  • Kristin L Leskoske

University of California Berkeley (University of California, Berkeley) (Predoctoral Fellowship)

  • Alexander Muir

National Institute of General Medical Sciences (NIGMS) (R01 GM21841)

  • Jeremy Thorner

Foundation of the American College of Allergy, Asthma & Immunology (ACAAI Foundation) (Senior Investigator Award 11-0118)

  • Jeremy Thorner

The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.

Acknowledgements

This work was supported by NIH Predoctoral Training Grant GM07232 and a Predoctoral Fellowship from the UC Systemwide Cancer Research Coordinating Committee (to AM), by NIH Predoctoral Training Grant GM07232 (to KLL), by NIH R01 Research Grant GM21841 and Senior Investigator Award 11-0118 from the American Asthma Foundation (to JT). We thank Stefan Hohmann (Univ. of Göteborg, Sweden), David E Levin (Boston Univ., Boston, MA), and Ted Powers (Univ. of California, Davis) for generously providing strains, plasmids and reagents, Hugo Tapia (Koshland Lab, UC Berkeley) for helpful discussions and reagents for measuring intracellular glycerol, and Jesse Patterson and the other members of the Thorner Lab for various research materials and thoughtful suggestions.

Reviewing Editor

  1. Tony Hunter, Salk Institute, United States

Publication history

  1. Received: June 10, 2015
  2. Accepted: August 13, 2015
  3. Accepted Manuscript published: August 14, 2015 (version 1)
  4. Version of Record published: August 28, 2015 (version 2)

Copyright

© 2015, Muir et al.

This article is distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use and redistribution provided that the original author and source are credited.

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