The target of rapamycin (TOR) kinase, a central regulator of eukaryotic cell growth, exists in two essential, yet distinct, TOR kinase complexes in the budding yeast Saccharomyces cerevisiae: rapamycin-sensitive TORC1 and rapamycin-insensitive TORC2. Lst8, a component of both TOR complexes, is essential for cell viability. However, it is unclear whether the essential function of Lst8 is linked to TORC1, TORC2, or both. To that end, we carried out a genetic screen to isolate lst8 deletion suppressor mutants. Here we report that mutations in SAC7 and FAR11 suppress lethality of lst8Δ and TORC2-deficient (tor2-21) mutations but not TORC1 inactivation, suggesting that the essential function of Lst8 is linked only to TORC2. More importantly, characterization of lst8Δ bypass mutants reveals a role for protein phosphatase 2A (PP2A) in the regulation of TORC2 signaling. We show that Far11, a member of the Far3-7-8-9-10-11 complex involved in pheromone-induced cell cycle arrest, interacts with Tpd3 and Pph21, conserved components of PP2A, and deletions of components of the Far3-7-8-9-10-11 complex and PP2A rescue growth defects in lst8Δ and tor2-21 mutants. In addition, loss of the regulatory B′ subunit of PP2A Rts1 or Far11 restores phosphorylation to the TORC2 substrate Slm1 in a tor2-21 mutant. Mammalian Far11 orthologs FAM40A/B exist in a complex with PP2A known as STRIPAK, suggesting a conserved functional association of PP2A and Far11. Antagonism of TORC2 signaling by PP2A-Far11 represents a novel regulatory mechanism for controlling spatial cell growth of yeast.
THE target of rapamycin (TOR) kinase is a phosphatidylinositol kinase-related protein kinase that controls eukaryotic cell growth and proliferation in response to nutrient conditions (Inoki et al. 2005; Wullschleger et al. 2006; Zoncu et al. 2011). The TOR kinase is inhibited by the complex of rapamycin and Fpr1, a peptidyl-prolyl cis–trans isomerase. The TOR kinase is conserved in eukaryotes. Unlike fungal species, which may possess two TOR kinases, higher eukaryotes such as humans possess only one. The TOR kinase exists in multi-protein complexes, which have been purified from many different eukaryotic systems. There exist two distinct TOR kinase complexes. In yeast, rapamycin-sensitive TORC1 consists of Tor1 or Tor2, Lst8, Kog1, and Tco89, and rapamycin-insensitive TORC2 consists of Tor2, Lst8, Avo1, Avo2, Avo3, and Bit61 (Loewith et al. 2002; Wedaman et al. 2003; Reinke et al. 2004). Both complexes are partially conserved in mammals: mTORC1 contains the yeast Kog1 ortholog raptor, while mTORC2 contains mSin1 and rictor, orthologs of yeast Avo1 and Avo3, respectively; GβL, the ortholog of yeast Lst8, exists in both mTORC1 and mTORC2 (Zoncu et al. 2011).
TOR regulates cell growth by sensing and responding to changes in nutrient conditions (Schmelzle and Hall 2000). TORC1 has an essential function involving the regulation of cell growth that is carried out when either Tor1 or Tor2 is in the complex. Under favorable growth conditions, TORC1 promotes cell growth by maintaining robust ribosome biogenesis (Marion et al. 2004; Martin et al. 2004; Zurita-Martinez and Cardenas 2005). When TORC1 is inactive, there is a dramatic downregulation of general protein translation, an upregulation of autophagy, accumulation of the storage carbohydrate glycogen, increased sorting and turnover of amino acid permeases, and activation of stress-responsive transcription factors via nuclear translocation (Wullschleger et al. 2006). TOR inhibition via rapamycin treatment activates a subset of stress-responsive transcription factors (Beck and Hall 1999; Cardenas et al. 1999; Shamji et al. 2000; Cooper 2002; Chen and Kaiser 2003). Rapamycin treatment can also lead to reduced gene expression, including encoding ribosomal proteins (RP) (Wullschleger et al. 2006).
TORC2 has a separate essential function that is Tor2 specific, which involves cell cycle-dependent polarization of the actin cytoskeleton (Cybulski and Hall 2009). TORC2 mediates the organization of the actin cytoskeleton through the activation of a Rho1/2p GTPase switch, composed of the Rho GTPases Rho1 and Rho2, the Rho GDP-GTP exchange factor Rom2, and the Rho GTPase-activating protein Sac7. Activated GTP-bound Rho1 activates Pkc1, which activates the cell-wall integrity pathway MAP kinase cascade, Bck1-Mkk1/2-Mpk1. Activation of Rho1 and the cell-wall integrity pathway restores cell growth and actin polarization to tor2 mutant cells. sac7 mutations suppress TORC2 deficiency by increasing the levels of GTP-bound Rho1. How TORC2 mediates the organization of the actin cytoskeleton is unclear and might involve three TORC2 substrates: Slm1, Slm2, and Ypk2 (Audhya et al. 2004; Fadri et al. 2005; Kamada et al. 2005; Tabuchi et al. 2006; Aronova et al. 2008).
Lst8 is essential for cell viability in Saccharomyces cerevisiae (Roberg et al. 1997). It is unknown whether the essential function of Lst8 is linked to TORC1, TORC2, or both. In TORC2, Lst8 binds to the C-terminal kinase domain of Tor2, independently of Avo1/2/3, and Lst8 depletion destabilizes the interaction between Tor2 and Avo2 or Avo3 (Wullschleger et al. 2005). Lst8 is also required for full Tor2 kinase activity in vitro, and its depletion leads to a depolarized actin cytoskeleton similar to tor2 mutations (Loewith et al. 2002; Wullschleger et al. 2005). Overexpression of MSS4 encoding a phosphatidylinositol-4-phosphate 5-kinase, RHO1/2, ROM2, PKC1, MKK1, or BCK1 suppresses tor2 and avo1 mutations (Helliwell et al. 1998a,b; Loewith et al. 2002); however, these suppressors were reported to be unable to suppress an lst8 mutation (Loewith et al. 2002). The question remains whether the essential function of Lst8 is linked to TORC2, and the role of Lst8 in TORC1 is largely unknown.
Here, we provide evidence that the essential function of Lst8 is linked to TORC2, but not to TORC1. We show that components of the Far3-7-8-9-10-11 complex, which have been implicated in pheromone-induced cell cycle arrest, vacuolar protein sorting, and cell fitness, negatively regulate TORC2 signaling. We find that Far11 interacts with protein phosphatase 2A and that mutations in the PP2A-Rts1 subcomplex suppress TORC2 deficiency. We propose that the Far3-7-8-9-10-11 complex and PP2A-Rts1 antagonize TORC2 signaling by promoting dephosphorylation of TORC2 substrates.
Materials and Methods
Strains, plasmids, and growth media and growth conditions
Yeast strains and plasmids used in this study are listed in Table 1, Table 2, Table S2, and Table S3. Yeast cells were grown at 30° or 37° in SD (0.67% yeast nitrogen base plus 2% dextrose), YNBcasD (SD medium plus 1% casamino acids), Ura Leu dropout (SD plus Complete Supplement Mixture without uracil and leucine; Bio101), or YPD (1% yeast extract, 2% peptone, 2% dextrose) medium as indicated in the text and in the figure legends. For lst8Δ bypass assays, SD medium with or without 1 g/liter 5-fluroorotic acid was used to select for growth of cells that have lost URA3 plasmids. When necessary, amino acids, adenine, and/or uracil were added to the growth medium at standard concentrations to cover auxotrophic requirements (Amberg et al. 2005).
lst8Δ mutant bypass genetic screens were conducted using transposon mutagenesis as described (Liu et al. 2001). Briefly, ade2Δ lst8Δ cells carrying plasmid pRS412-LST8 were used for transposon mutagenesis; after mutagenesis, cells were plated on YPD medium to select for colonies that were red or sectoring, indicating loss of the pRS412-LST8 plasmid. Putative lst8Δ bypass mutants were confirmed to be recessive, single-gene mutations using standard yeast genetic techniques. Determination of transposon insertion sites were carried out as described (Burns et al. 1994). Briefly, a recovery plasmid encoding the ampicillin-resistance gene (AmpR) and URA3 was integrated into the transposon integration site of the leb1 and leb2 mutants by homologous recombination, and transformants were selected on uracil-dropout medium. Genomic DNA was then isolated, digested with EcoRI, and ligated with T4 DNA ligase. Ligation products were transformed into bacteria, and AmpR transformants were selected for on LB broth supplemented with ampicillin. Transposon integration sites were determined by sequencing of the recovered plasmids.
Total cellular RNAs were isolated using a hot phenol method as described (Giannattasio et al. 2005). Cells were grown in appropriate medium to ∼OD600 0.6 and collected for isolation of total cellular RNA. 32P-labeled probes against RPL3, RPS6A, and ACT1 were used to probe mRNA immobilized on nylon membranes. PhosphorImager was used to record signals of the RNA transcripts.
Cellular extract preparation, immunoblotting, and immunoprecipitation
Total cellular protein extracts were prepared by disrupting yeast cells in extraction buffer (1.85 N NaOH–7.5% β-mercaptoethanol) followed by precipitation with trichloroacetic acid as described (Yaffe and Schatz 1984). Phosphatase treatment of total cellular proteins was conducted as described (Liu et al. 2008). For co-immunoprecipitation experiments, cellular lysates were prepared in immunoprecipitation (IP) buffer (50 mM Tris–HCl, pH 7.6, 150 mM NaCl, 0.5% Triton X-100, and protease inhibitors). Cell extracts (∼3 mg proteins) were incubated at 4° for 1 hr with anti-myc antibody (9E10; Roche), after which 30 μl of a 50% slurry of protein G-Sepharose (Roche) was added to each sample, and the samples were further incubated at 4° for 2 hr. Washed immunoprecipitates bound to the sepharose beads were released by boiling in 1× SDS-PAGE loading buffer. The released immune complexes were analyzed by Western blotting. myc- and HA-tagged proteins were probed with anti-myc antibody and anti-HA antibody (3F10; Roche), respectively. Chemiluminescence images of Western blots were captured using the Bio-Rad Chemi-Doc photo documentation system.
Actin staining and GFP fluorescence microscopy
The actin cytoskeleton was visualized in rhodamine phalloidin-stained, formaldehyde-fixed cells, as described (Amberg et al. 2005). Overnight cultures were diluted to ∼OD600 0.1 and allowed to grow at 30° for 2 hr and then switched to 37° for 3 hr before formaldehyde was added to a final concentration of 3.7%. After fixing for 1 hr, 1 ml of fixed cells were collected, washed in PBS buffer, and stained with rhodamine phalloidin conjugate (Invitrogen) and visualized by fluorescence microscopy. GFP fluorescence of GFP-tagged proteins was analyzed in cells grown to log phase. Fluorescence images of rhodamine phalloidin-labeled actin structures and GFP-tagged proteins were acquired with a Photometrics Coolsnap fx CCD camera and Metamorph Imaging Software and processed using ImageJ (National Institutes of Health) and Adobe Photoshop.
Preparation of recombinant 6×His-tagged Slm1
PCR-amplified SLM1 coding sequence was cloned into the SacI and XhoI sites of pET24a vector (Novagen). The resultant plasmid was transformed into BL21(DE3)-competent cells (Novagen), and expression of 6×His-tagged Slm1 was induced by adding 1 mM IPTG to bacterial cultures grown at 20° for 16 hr. Recombinant C-terminal 6xHis-tagged Slm1 was purified under native conditions using Ni-NTA agarose beads (Qiagen) as described in the product instruction manual. Slm1-His6 (14 mg) was obtained from 1 liter of induced culture.
In vitro kinase assay of Slm1 phosphorylation by Tor2-HA
Yeast strains expressing 3×HA-tagged Tor2 were grown overnight to mid-log phase, and cellular lysates were prepared in lysis buffer (50 mM Tris–HCl, pH 7.6, 150 mM NaCl, 1% Triton X-100, and protease inhibitors) by vortexing with glass beads. Cell extracts (∼3 mg proteins) were incubated with 100 μl protein A-agarose beads (Roche) at 4° for 1 hr to remove nonspecific binding proteins. Precleared cell lysates were then incubated with 16 μg anti-HA antibody (12CA5, Roche) for 1 hr, after which 40 µl of a 50% slurry of protein A-agarose beads was added to each sample, and the samples were further incubated at 4° for 1 hr. Precipitates were washed twice with 1 ml lysis buffer, twice with 1 ml wash buffer (50 mM Tris–HCl, pH 7.6, 300 mM NaCl, and protease inhibitors), and once with 1 ml kinase buffer (20 mM Tris–HCl, pH 7.6, 50 mM NaCl, 5 mM MgCl2, 1 mM PMSF). After the final wash, beads were resuspended in 25 µl of kinase buffer plus 1 mM dithiothreitol (DTT). The reaction was initiated by adding 25 µl reaction mixture [kinase buffer with 1 mM DTT, 0.4 mM ATP, 5 µCi of [γ-32P]ATP (PerkinElmer), 5 µg recombinant Slm1-His6]. After incubation for 30 min at 30°, the reaction was terminated by adding 25 μl 3× SDS gel-loading buffer (150 mM Tris–HCl, pH 6.8, 6% SDS, 30% glycerol, 0.3% bromophenol blue) and 8.5 μl 1 M DTT and boiling for 4 min. Samples (20 µl) were fractionated by SDS–PAGE on 7.5% polyacrylamide gels in triplicate, with one dried for detecting 32P-incorporation in Slm1 by autoradiography (Molecular PhosphorImager FX, Bio-Rad), one stained by Coomassie Blue for detecting 6×His-tagged Slm1, and one processed for Western blotting for detecting HA-Tor2. A mock treatment experiment was conducted using anti-HA immunoprecipitates from cells expressing nontagged Tor2.
Mutations in SAC7 and FAR11 suppress lethality due to an lst8Δ mutation
To gain insights into the essential function of Lst8, we conducted a genetic screen to search for mutations that allow cells to survive without Lst8 by employing an ade2 colony sectoring assay. This assay takes advantage of a build-up of purine precursors in the vacuole, which results in colonies that appear red in ade2 mutant cells. We utilized a previously constructed Tn3::lacZ::LEU2 mutagenesis library (Burns et al. 1994) to introduce mutations in an lst8Δ ade2Δ mutant carrying plasmid pRS412-LST8 (CEN LST8 ADE2) and screened for red or sectoring lst8Δ bypass (leb) mutant cells on YPD medium, which had lost or were in the process of losing the pRS412-LST8 plasmid. Of ∼30,000 Leu+ transformants, 49 viable solid red or sectoring colonies were selected for further analysis. Of the 49, 11 colonies were pure red. Crossing to an ade1Δ strain resulted in noncomplementation of the red phenotype in 1 of the 11 pure red colonies, indicating that the red phenotype was due to a mutation in ADE1 and not to the loss of the pRS412-LST8 plasmid. Alternatively, crossing to an ade2Δ strain resulted in noncomplementation of the red phenotype in the remaining 10 pure red colonies, indicating that the red phenotype is due to a mutation in ADE2 or to loss of the pRS412-LST8 plasmid. However, wild-type LST8 was found by PCR to be present in all 11 pure red colonies, which were deemed false positives. The remaining 38 sectoring colonies were analyzed similarly, and the red sectoring phenotype of one was due to an ADE1 mutation while 35 were due to mutations in ADE2. PCR genotyping confirmed the absence of pRS412-LST8 in the remaining two mutants, which we termed leb1 and leb2. In these two leb mutants, the lst8Δ bypass phenotype was found to cosegregate with Leu+ after crossing to an lst8Δ leu2 ade2Δ pRS412-LST8 strain of the opposite mating type. Tetrads were dissected, indicating that the transposon insertion had produced the mutant phenotype. Figure 1A shows that, in contrast to wild-type cells, leb1 and leb2 mutants form both red and sectoring colonies, indicating loss of the pRS412-LST8 plasmid.
We identified the transposon insertion site in the leb1 and leb2 mutants by plasmid rescue and sequencing of the recovered plasmids as described by Burns et al. (1994). The transposon insertion sites in the leb1 and leb2 mutants were found to be in the open reading frames of SAC7 and FAR11, respectively. Consistently, Figure 1, B and C, shows that wild-type SAC7 and FAR11 on a centromeric plasmid can complement leb1 and leb2 mutations, respectively. Furthermore, a sac7Δ or far11Δ mutation in an lst8Δ ade2Δ pRS412-LST8 strain also led to lst8Δ bypass (loss of pRS412-LST8 is indicated by the red colony phenotype in Figure 2A).
sac7Δ and far11Δ mutants are sensitive to rapamycin
Whether the essential function of Lst8 is linked to TORC1, TORC2, or both has yet to be determined. We tested the ability of sac7Δ and far11Δ mutations to suppress TORC1 inactivation by rapamycin treatment and TORC2 deficiency due to a temperature-sensitive tor2-21 mutation. An fpr1Δ mutation enables cells to grow in the presence of rapamycin; however, far11Δ lst8Δ, far11Δ (far11Δ lst8Δ pRS412-LST8), sac7Δ lst8Δ, and sac7Δ (sac7Δ lst8Δ pRS412-LST8) cells were unable to grow on YPD medium supplemented with 200 nM rapamycin (Figure 2A), demonstrating that these two lst8Δ bypass mutations do not suppress a severe or complete loss of TORC1 activity. Consistently, sac7Δ and far11Δ mutations failed to restore cell growth to tor1Δ tor2-21 double mutants grown at 37°, which have defects in the function of both TORC1 and TORC2 (Supporting Information, Figure S1) (Schmidt et al. 1997). In contrast, sac7Δ and far11Δ mutations restored cell growth to a tor2-21 mutant grown at 37°, which causes a specific defect in only TORC2 (Figure 3C and Figure 4C), consistent with previous findings that sac7Δ suppresses a tor2-21 mutation (Schmidt et al. 1997). These data suggest that the essential function of Lst8 may be linked to TORC2, but not to TORC1.
It is conceivable that lst8Δ may result in a partial loss of TORC1 activity, which is not sufficient to support cell growth but is not severe enough to prevent sac7Δ and far11Δ mutations from restoring partial cell growth to lst8Δ mutant cells. To test this possibility, we tested the sensitivity of wild-type, fpr1Δ, sac7Δ, and far11Δ mutant cells to lower concentrations of rapamycin (Figure S2). It has been reported that sac7Δ and far11Δ mutant strains in the BY4741 background are hypersensitive to treatment with 10 nM rapamycin (Xie et al. 2005). In the presence of <10 nM rapamycin, a far11Δ mutant in the BY4741 background has been reported to grow better than wild-type cells (Huber et al. 2009). We analyzed cell growth in the presence of 2–20 nM rapamycin and found that sac7Δ resulted in hypersensitivity to rapamycin treatment. A far11Δ mutant, in contrast, grew slightly better than wild-type cells when treated with 3 and 5 nM rapamycin. In the presence of 7–20 nM rapamycin, however, far11Δ cells no longer grew better than wild-type cells. In our strain background, treatment of wild-type cells with 10 nM rapamycin likely reduces TORC1 activity to just below the threshold that supports cell growth. Our observations that sac7Δ and far11Δ bypass lst8Δ but not treatment with 10 nM rapamycin strongly suggest that the essential function of lst8 is not linked to TORC1.
To further corroborate our hypothesis that Lst8 is not essential for TORC1 activity, we examined the effect of lst8Δ on the expression of genes encoding RPs, which are positively regulated by TORC1. Utilizing sac7Δ to obtain a viable lst8Δ mutant, we compared the expression of two RP genes, RPL3 and RPS6A, encoding the L3 protein of the large (60S) ribosomal subunit and the S6 protein of the small (40S) ribosomal subunit, respectively, in LST8 sac7Δ vs. lst8Δ sac7Δ mutant cells by Northern blot analysis. As expected, TORC1 inactivation due to rapamycin treatment inhibited expression of both RPL3 and RPS6A (Figure 2B). In contrast, lst8Δ only mildly reduced the expression of RPL3 and RPS6A, suggesting that lst8Δ does not lead to severe loss of TORC1 activity.
lst8Δ causes mislocalization of Bit61 and Avo3, but not of Kog1
Recent research has demonstrated that the TORC1 components are located on intracellular membranes with a concentration on the vacuolar membrane while TORC2 components appear as punctate spots at the plasma membrane (Wedaman et al. 2003; Araki et al. 2005; Sturgill et al. 2008; Berchtold and Walther 2009; Binda et al. 2009). It has been proposed that plasma membrane localization of TORC2 is essential for cell viability (Berchtold and Walther 2009). Isolation of lst8Δ mutant suppressors allowed us to examine the role of Lst8 in the cellular localization of TOR complex components. GFP fluorescence was analyzed in sac7Δ far11Δ double mutants and sac7Δ far11Δ lst8Δ triple mutants expressing the GFP-tagged TORC1 component Kog1 and TORC2 components Bit61 and Avo3. As previously reported, Bit61 and Avo3 localized to the plasma membrane as punctate spots in wild-type LST8 cells (Figure 2C) (Berchtold and Walther 2009). An lst8Δ mutation, however, largely abolished punctate plasma membrane localization of Bit61 and Avo3. In contrast, lst8Δ did not affect localization of Kog1 to the vacuolar membrane (Figure 2C). These findings are consistent with our hypothesis that the essential function of Lst8 is linked to TORC2, but not to TORC1, and further suggest that Lst8 is required for proper cellular localization of TORC2.
Mutations of the Far3-7-8-9-10-11 complex bypass lst8Δ and tor2-21 mutations
Far11 has been shown to be involved in cell cycle arrest in response to a mating pheromone in a multi-protein complex with Far3, Far7, Far8, Far9, and Far10 (Horecka and Sprague 1996; Kemp and Sprague 2003). Interactions among these six Far proteins are based mostly on yeast two-hybrid assays (Kemp and Sprague 2003; Lai et al. 2011); however, Far3 has been reported to interact with Far11 by co-immunoprecipitation. We generated strains coexpressing HA-tagged Far11 and myc-tagged Far7, Far8, Far9, Far10, and Far11 under the control of their respective endogenous promoters to test whether Far11 interacts with Far7, Far8, Far9, Far10, or itself by co-immunoprecipitation. FAR11-HA was found to be functional by its ability to complement a far11Δ mutation using the colony-sectoring assay described for Figure 1C (Figure S3). myc-tagged Far proteins are functional as described previously (Kemp and Sprague 2003). Far11-HA in cell lysates prepared for co-immunoprecipitation exists as two bands on Western blots (Figure 3A). The faster mobility form of Far11-HA is likely to be a proteolytically truncated form of Far11 since Far11-HA exists as a single band on Western blots when the total cellular proteins are prepared by disrupting cells in the presence of 1.85 N NaOH–7.5% β-mercaptoethanol followed by precipitation with trichloroacetic acid (Figure S4). myc-tagged proteins were precipitated from cell lysates using anti-myc antibody. Figure 3A shows that Far11-HA co-immunoprecipitates with myc-tagged Far7, Far8, Far9, Far10, but not Far11, demonstrating that Far11 interacts with other Far proteins in vivo.
Since Far11 is part of a multi-protein complex, it is possible that the entire complex is involved in TORC2 signaling. We examined whether mutations of the Far complex bypass lst8Δ and tor2-21 as well. lst8Δ farΔ double mutants, each carrying a centromeric plasmid encoding LST8 and URA3 ([CEN URA LST8]), were grown on SD medium without or with 5-fluoroorotic acid (5-FOA), which selects for cells that have lost the URA3 plasmid. On the basis of their relative growth in the presence of 5-FOA, mutations in FAR3, FAR7, FAR8, FAR9, FAR10, and FAR11 bypass lst8Δ to varying degrees: far11 > far8/9 > far3/7 > far10 (Figure 3B). farΔ mutations were subsequently found to suppress a tor2-21 mutation at 37° (Figure 3C). The tor2-21 suppression phenotypes of far3, far7, far8, far9, far10, and far11 mutations largely mirror that of their respective lst8Δ bypass (Figure 3, B and C), indicating that the function of Lst8 is tightly linked to TORC2.
Far9 and Far10 are homologous proteins with 31% sequence identity and 47% sequence similarity, yet they have different lst8Δ and tor2-21 suppression phenotypes. We generated a tor2-21 far9Δ far10Δ triple mutant and found that its growth was only marginally better than that of the tor2-21 far9Δ double mutant (Figure S5), indicating that, of the two, Far9 plays the primary role in TORC2 signaling.
It is unclear whether an lst8Δ or tor2-21 mutation leads to complete loss of TORC2 activity. Deletion of TOR2, encoding the only TOR kinase in TORC2, abolishes TORC2 activity. To test whether far11Δ bypasses tor2Δ, spores of tetrads from FAR11/far11 TOR2/tor2Δ::kanMX4 diploid cells were assessed for viability. All tetrads produced at most two viable spores, none of which were geneticin resistant (kanXM4 confers geneticin resistance) (Figure S6). Since FAR11 and TOR2 are not located on the same chromosome, 25% of spores should have the genotype tor2Δ::kanMX4 far11Δ. Failure to obtain viable geneticin-resistant spores indicates that far11Δ is unable to bypass tor2Δ. Similarly, we found that far11Δ failed to bypass avo1Δ or avo3Δ (Figure S6) . Since far11Δ is able to suppress lst8Δ and tor2-21 mutations, it is likely that lst8Δ and tor2-21 mutations do not completely abolish TORC2 activity and that far11Δ can restore growth to cells with a severe loss, but not a total loss, of TORC2 activity.
sac7Δ and far11Δ additively suppress tor2-21
TORC2 is involved in the organization of the actin cytoskeleton. A tor2-21 mutant shows depolarization of the actin cytoskeleton, and several tor2-21 suppressors can restore actin polarization to tor2-21 mutant cells (Schmidt et al. 1997). We compared actin structures in wild-type, tor2-21, tor2-21 sac7Δ, and tor2-21 far11Δ mutant cells to establish whether actin polarization defects caused by a tor2-21 mutation could be restored by a far11Δ mutation. As expected, a sac7Δ mutation restored polarization of the actin cytoskeleton in tor2-21 mutant cells grown at 37° (Figure 4A). Similarly, a far11Δ mutation restored polarization of actin structures in tor2-21 mutant cells grown at 37° (Figure 4B). A recent genome-wide study of genetic interactions in yeast showed that far11Δ restored actin polarization to tsc11-1 (avo3-1) mutant cells (Baryshnikova et al. 2010). Together, these data establish that Far11 negatively regulates TORC2-mediated polarization of the actin cytoskeleton.
Similar phenotypes of far11Δ and sac7Δ mutations prompted us to determine whether they act through the same molecular mechanism. Accordingly, we compared the growth of a tor2-21 sac7Δ far11Δ triple mutant to wild-type, tor2-21, tor2-21 sac7Δ, and tor2-21 far11Δ mutants at 30° vs. 37°. Figure 4C shows that, while all strains grew equally well at 30°, the tor2-21 sac7Δ far11Δ mutant grew better than either the tor2-21 far11Δ or the tor2-21 sac7Δ mutant at 37°, indicating that far11Δ and sac7Δ have additive effects in suppressing tor2-21. To examine whether sac7Δ and far11Δ have an additive effect in suppressing an actin depolarization defect in tor2-21 mutant cells, we determined the percentage of cells with a polarized actin cytoskeleton in wild-type and isogenic tor2-21, tor2-21 sac7Δ, tor2-21 far11Δ, and tor2-21 sac7Δ far11Δ mutant cells. Table S1 shows that the effect of sac7Δ and far11Δ on the restoration of actin structure polarization is additive. Sac7 and Rom2 have opposing roles in mediating TORC2 function, and it has been proposed that Tor2 activates Rho1 via Rom2 (Schmidt et al. 1997). Therefore, to determine whether tor2-21 suppression by far11Δ is Rom2 dependent, we introduced a rom2Δ mutation into the tor2-21 far11Δ mutant. Figure 4D shows that a rom2Δ mutation greatly reduced the tor2-21 suppression phenotype of a far11Δ mutation at 37° but did not abolish it, suggesting that far11Δ suppression of TORC2 deficiency is not entirely dependent on the Rom2-mediated Rho1/2 GTPase switch.
Far11 interacts with Tpd3 and Pph21, components of PP2A
Our data so far raise the question: How does the Far3-7-8-9-10-11 complex mediate TORC2 signaling? The human ortholog of yeast Far11 exists in the human STRIPAK complex, which also contains components of PP2A (Goudreault et al. 2009). The Far11 ortholog in Drosophila has also been reported to interact with PP2A in the dSTRIPAK complex (Ribeiro et al. 2010). Therefore, we tested whether Far11 in yeast also exists in a complex with PP2A. In yeast, the heterotrimeric PP2A phosphatase consists of the regulatory A subunit Tpd3, the regulatory B subunit Cdc55 or B′ subunit Rts1, and one of the two homologous and functionally redundant catalytic C subunits, Pph21 or Pph22 (Duvel and Broach 2004). Among ∼75 proteins that genetically or biochemically interact with Far11 in various genome-wide gene/protein interaction studies, Tpd3 has been found to interact with Far11 by yeast two-hybrid analysis (Uetz et al. 2000). The significance of this interaction remains unknown.
To establish the interaction between Far11 and PP2A in yeast, lysates from cells coexpressing 3× HA-tagged Far11 and 3× myc-tagged Tpd3 or Pph21 were subjected to immunoprecipitation using anti-myc antibody. TPD3-myc and PPH21-myc constructs were found to be functional by their ability to rescue growth defects of tpd3Δ and pph21/22Δ mutants, respectively (Figure S7). No Far11-HA was detected in the IP pellet from cells expressing Far11-HA alone. In contrast, Far11-HA was recovered in the IP pellet from cells coexpressing Tpd3-myc and, to a lesser extent, from cells coexpressing Pph21-myc (Figure 5A) likely because C-terminal tagging of Pph21 perturbs methylation at its C terminus required for PP2A complex stability (Wei et al. 2001). These findings establish that Far11 interacts with PP2A phosphatase.
Defects in PP2A-Rts1 bypass lst8Δ and tor2-21 mutations
To investigate whether PP2A is involved in TORC2 signaling, we examined whether mutations in PP2A components bypass lst8Δ. We analyzed the growth of an lst8Δ tpd3Δ double mutant, an lst8Δ rts1Δ double mutant, an lst8Δ cdc55Δ double mutant, and an lst8Δ pph21Δ pph22Δ triple mutant, each carrying a centromeric plasmid encoding URA3 and LST8 on SD medium without or with 5-FOA. tpd3Δ, rts1Δ, and pph21/22Δ mutations, but not a cdc55Δ mutation, were able to bypass lst8Δ (Figure 5, B and C), indicated by their ability or inability to grow in the presence of 5-FOA, suggesting that reduced activity in the PP2A-Rts1 subcomplex results in lst8Δ bypass. We then tested whether rts1Δ and tpd3Δ mutations suppress tor2-21. Figure 5D shows that rts1Δ suppresses a tor2-21 mutation by restoring cell growth at 37°. A tpd3Δ mutation led to temperature-sensitive growth defects in the TOR2 wild-type strain used in our study; therefore, we could not assay tor2-21 suppression by tpd3Δ. These findings indicate that mutations in genes encoding components of the PP2A-Rts1 subcomplex suppress TORC2 deficiency.
far11Δ and rts1Δ restore phosphorylation of Slm1 to a tor2-21 mutant
One known function of TORC2 is its role in the organization of the actin cytoskeleton possibly by phosphorylating Slm1, Slm2, and Ypk2. Our data above show that far11Δ, sac7Δ, and rts1Δ mutations suppress a tor2-21 mutation and that Far11 interacts with PP2A. Therefore, we tested the possibility that Far11 may mediate dephosphorylation of Slm1, Slm2, and/or Ypk2 via PP2A by evaluating the phosphorylation states of Slm1, Slm2, and Ypk2 in wild-type, tor2-21, tor2-21 far11Δ, tor2-21 sac7Δ, and tor2-21 rts1Δ mutant cells, each expressing Slm1-HA, Slm2-HA, or Ypk2-HA from their respective endogenous promoters. The phosphorylation states of Ypk2 and Slm2 did not differ between the wild-type and the tor2-21 mutant grown at 37° (Figure S8). Therefore, Ypk2 and Slm2 were not studied further. Consistent with previous reports on the phosphorylation state of GFP-tagged Slm1 (Audhya et al. 2004), Figure 6A shows that Slm1-HA is phosphorylated: lambda protein phosphatase (λPPase) treatment resulted in increased levels of the faster mobility forms of Slm1-HA with concomitant reduced levels of the slower mobility forms of Slm1-HA; phosphatase inhibitors largely abolished the effect of λPPase treatment. As reported previously, Figure 6B shows that Slm1 is dephosphorylated in tor2-21 mutant cells grown at 37° (Audhya et al. 2004). Remarkably, far11Δ, but not sac7Δ, restored Slm1-HA phosphorylation to tor2-21 mutant cells grown at 37° (Figure 6B), implicating Far11 in Slm1 dephosphorylation and suggesting that suppression of TORC2 deficiency by a sac7Δ mutation takes place downstream of Slm1. Furthermore, Slm1-HA was phosphorylated in tor2-21 rts1Δ mutant cells grown at 37° (Figure 6C). These data suggest that Far11-PP2A-Rts1 may antagonize TORC2 activity and decrease the levels of the phosphorylated form of the TORC2 substrate Slm1.
Effects of sac7Δ and far11Δ mutations on Tor2 kinase activity
Sac7 has been proposed to function downstream of TORC2. Our genetic data suggest that Far11-PP2A-Rts1 antagonizes TORC2 signaling. It is possible that Far11-PP2A-Rts1 may function downstream of TORC2 and promote dephosphorylation of TORC2 substrates. It is also likely that Far11-PP2A-Rts1 may function upstream of TORC2 and negatively impact TORC2 activity. To differentiate between these two possibilities, the activity of immunopurified Tor2 with a N-terminal 3×HA tag from wild-type and isogenic far11Δ mutant cells was determined in an in vitro kinase assay using Slm1 as a substrate. Slm1 has been reported to be a TORC2 substrate in in vitro kinase assays (Audhya et al. 2004; Fadri et al. 2005). For our assays, recombinant 6×His-tagged Slm1 was added to kinase reactions with immunopurified HA-Tor2 and [γ-32P]ATP. Figure 7 shows that far11Δ has no significant effect on kinase activity of immunopurified HA-Tor2, suggesting that Far11 functions at a site downstream of TORC2.
Similarly, we performed an in vitro kinase assay using HA-Tor2 from sac7Δ mutant cells. Surprisingly, sac7Δ slightly increases kinase activity of immunopurified HA-Tor2 (Figure 7). Mutations in SAC7 have been proposed to activate Rho1, which in turn activates Pkc1 and the cell-wall integrity MAP kinase cascade. Increased activity of Tor2 in sac7Δ mutant cells suggests positive feedback regulation in TORC2 signaling. Differential effects of sac7Δ and far11Δ mutations on Tor2 kinase activity further support the notion that Sac7 and Far11 mediate TORC2 signaling through different mechanisms.
Lst8 is an essential protein that exists in both TOR kinase complexes. We found that mutations in genes encoding the PP2A-Rts1 subcomplex and the Far3-7-8-9-10-11 complex bypass lst8Δ and TORC2 deficiency. Analysis of these mutants led us to propose that the essential function of Lst8 is linked to TORC2. The Far3-7-8-9-10-11 complex components are partially conserved in Drosophila and mammals and have been reported to interact with PP2A phosphatase in the STRIPAK complex. We showed that the Far3-7-8-9-10-11 complex and PP2A negatively regulate TORC2 signaling possibly by mediating dephosphorylation of the TORC2 substrate Slm1, depicted by our proposed model in Figure 8. Our results not only demonstrate that the essential function of Lst8 is linked only to TORC2, but also, more importantly, reveal a novel link between the two major signaling protein complexes PP2A and TORC2.
Essential function of Lst8 is linked to TORC2, but not to TORC1
Yeast Lst8 has been reported to be important for TORC2 complex integrity in vivo and for Tor2 kinase activity in vitro (Wullschleger et al. 2005). Underlying its importance in TORC2 activity, the presence of Lst8 in TORC2 has been reported in multiple organisms including yeast, slime mold, worms, flies, and mammals (Cybulski and Hall 2009). Consistently, our data demonstrate that the essential function of Lst8 is linked to TORC2. Delocalization of Bit61 and Avo3 from punctate structures at the plasma membrane in lst8Δ mutant cells likely results from compromised TORC2 integrity in the absence of Lst8, indicating that Lst8 is also required for proper localization of the TORC2 complex.
Lst8 interacts with the kinase domain of Tor2 in yeast TORC2 (Wullschleger et al. 2005). Since far11Δ bypasses lst8Δ and tor2-21, but not tor2Δ, avo1Δ, and avo3Δ, our data suggest that neither lst8Δ nor the temperature-sensitive tor2-21 mutation leads to total loss of TORC2 activity. This possibility helps explain our observation that far11Δ and rts1Δ can restore Slm1 phosphorylation to tor2-21 mutant cells at the restrictive temperature: If tor2-21 led to the total loss of TORC2 activity and consequent loss of Slm1 phosphorylation, PP2A inactivation could not restore Slm1 phosphorylation unless other protein kinases also phosphorylate Slm1. In this scenario, TORC2 would share a redundant function with the other putative kinase in phosphorylating Slm1 and may have another essential function separate from its kinase activity, for example, by maintaining interactions with other proteins to conduct downstream signaling.
Three TORC2 substrates in yeast are Slm1, Slm2, and Ypk2. A constitutively active Ypk2 mutant can restore growth to tor2Δ mutant cells, leading to the proposal that the essential function of TORC2 is linked mainly to Ypk2 phosphorylation (Kamada et al. 2005; Cybulski and Hall 2009). There is strong evidence that functionally redundant Slm1 and Slm2 are essential substrates of TORC2. First, Slm1 has been reported to interact with TORC2 and to be phosphorylated by TORC2 (Audhya et al. 2004; Fadri et al. 2005). Second, TORC2-dependent phosphorylation of Slm1 seems to correlate with its plasma membrane association (Audhya et al. 2004). Slm1 contains a PH domain that binds to multiply phosphorylated phosphoinositides and is required for Slm1’s plasma membrane localization (Fadri et al. 2005), and putative loss of plasma membrane association of Slm1/2 leads to cell death. Third, a sac7Δ mutation suppresses both a tor2-21 mutation and a slm1Δ slm2Δ double mutation, and the actin cytoskeleton is depolarized in both slm1/2 and tor2-21 mutant cells. Although we could not test whether phosphorylation of Slm2 and Ypk2 in a tor2-21 mutant is restored by far11 and rts1 mutations, it is likely that, due to mutations in Far11-PP2A, increased phosphorylation of Slm1, Slm2, and/or Ypk2 leads to cell viability in TORC2-deficient cells.
The presence of Lst8 in TORC1 is conserved from yeast to mammals. In yeast, Lst8 localizes not only to the TORC1 compartment at the vacuolar membrane, but also to the TORC2 compartment as punctate structures at the plasma membrane (Berchtold and Walther 2009). Similar to its interaction with Tor2 in TORC2, Lst8 interacts with the Tor1 kinase domain in the TORC1 complex in yeast and humans (Kim et al. 2003; Adami et al. 2007). Therefore, our observation that Lst8 is not required for TORC1-dependent expression of genes encoding ribosomal proteins and Kog1 localization at the vacuolar membrane is surprising and raises the question: Is Lst8 required for TORC1 activity at all? Missense mutations in LST8 increase the expression of a subset of TORC1-target genes (Chen and Kaiser 2003; Giannattasio et al. 2005). Therefore, Lst8 is likely to be required for optimal TORC1 activity, but an lst8Δ mutation may not reduce TORC1 activity severely enough to lead to cell death. In mammals, the role of Lst8 in mTORC1 is unclear. mLst8 knockdown in human immortalized cell lines suggested that mLst8 is important for mTORC1 activity (Kim et al. 2003). Later in mice, mLst8 was found to be important for mTORC2, but not for mTORC1 function during mouse development (Guertin et al. 2006). The discrepancy could be attributed to the differences between a developing mouse embryo and an immortalized human cell line. However, the conclusions concerning the role of Lst8 in both yeast and mice are similar: the essential function of Lst8 is linked to TORC2 but not to TORC1.
Far3-7-8-9-10-11-PP2A as a negative regulator of TORC2 signaling
One of the important findings presented here is a negative regulatory role of the Far3-7-8-9-10-11 complex and PP2A in TORC2 signaling. Considering the extensive studies on PP2A and TORC2, it is surprising that, to our knowledge, our study may represent the first to present a direct genetic interaction between TORC2 and PP2A. More importantly, our data mirror a recent study in Drosophila demonstrating that the Drosophila Far complex works in concert with PP2A in the regulation of a different kinase pathway, the Hippo signaling pathway (Ribeiro et al. 2010). Thus, PP2A regulation by the Far protein complex appears to be evolutionarily conserved. In a proteomics study, the yeast Far11 orthologs Fam40A and Fam40B (STRIP1/2) were isolated in the STRIPAK complex, which also contains components of PP2A. Interestingly, far11Δ leads to the strongest suppression of TORC2 deficiency, and among the six Far proteins, Far11 is the most conserved. Far9 and Far10 are homologous proteins, and their Drosophila and mammalian orthologs show limited sequence homology, mostly in an FHA domain, which is known to interact with phosphothreonine epitopes on target proteins (Durocher and Jackson 2002). Far8 shows very limited sequence homology to striatin (Goudreault et al. 2009). The question remains: How does the Far3-7-8-9-10-11 complex affect PP2A activity? Interaction between Far11 and Tpd3, the scaffolding subunit of PP2A, suggests that the Far complex may directly regulate PP2A by either targeting the TORC2 substrate Slm1 and/or mediating PP2A activity. Unlike slow cell growth phenotypes due to a tpd3Δ single mutation or a pph21/22Δ double mutation, mutations in FAR3, FAR7, FAR8, FAR9, FAR10, and FAR11 have little or no growth defects, suggesting that the Far complex is not integral to PP2A activity. In both the Drosophila study and ours, mutations in PP2A and/or Far components of the STRIPAK complex lead to increased phosphorylation of target proteins. Thus, it is possible that the Far complex might target certain substrates to PP2A.
Our study demonstrates that Far11-PP2A-Rts1 modulates Slm1 phosphorylation by counteracting the kinase activity of TORC2, providing a molecular mechanism to explain how mutations in FAR11 and genes encoding PP2A-Rts1 components might suppress TORC2 deficiency. The calcineurin phosphatase also mediates Slm1 dephosphorylation and counteracts TORC2 signaling (Bultynck et al. 2006; Mulet et al. 2006; Daquinag et al. 2007). It was further shown that mutations in cnb1, encoding the regulatory subunit of calcineurin, suppress an avo3 temperature-sensitive mutation (Aronova et al. 2008). It remains to be determined whether a cnb1 mutation restores phosphorylation of Slm1 to TORC2-deficient cells. Ypk2 is another essential effector of TORC2. Mutations in calcineurin could potentially restore phosphorylation of Ypk2 in avo3 mutant cells, thereby suppressing the avo3 temperature-sensitive growth phenotype. Furthermore, in a genome-wide study on genetic interactions in yeast, mutations in PPG1, encoding a PP2A-like phosphatase, also suppress TORC2 deficiency (Baryshnikova et al. 2010). Therefore, it is likely that these phosphatases may work together to mediate TORC2 signaling.
Various genetic screens in fungi have isolated mutations in the Far complex. In Neurospora crassa, a mutation in ham-2, the ortholog of yeast FAR11, leads to defects in hyphal fusion (Xiang et al. 2002). In Sordaria macrospora, mutations in PRO22, encoding the yeast Far11 ortholog, generate a novel type of sterile mutant with a defect in ascogonial septum formation (Bloemendal et al. 2010). In yeast, mutations in FAR9/VPS64 and FAR11/YNL127w result in vacuolar sorting defects (Bonangelino et al. 2002), and mutations in FAR3, FAR7, FAR8, FAR10, and FAR11 create long-lived mutants (Fabrizio et al. 2010). In all of these studies, the underlying mechanisms are unknown. In light of our findings, it is possible that these disparate phenotypes are due to the perturbation of PP2A activity and/or TORC2 signaling in these mutants.
We thank Michael Hall, Michael Snyder, George Sprague, Jr., Tobias Walther, and Terrance Cooper for providing us with strains and plasmids; Melissa Overstreet and Sylvester Tumusiime for technical support; Mary Clancy for in vitro kinase assays; the W. M. Keck Foundation for the Keck Facility; and Robin Rowe for sequencing. This work was supported by grant LEQSF(2008-11)-RD-A-31 from the State of Louisiana Board of Regents.
Supporting information is available online at http://www.genetics.org/content/suppl/2012/01/31/genetics.111.138305.DC1.
Communicating editor: A. P. Mitchell
- Received December 29, 2011.
- Accepted January 18, 2012.
- Copyright © 2012 by the Genetics Society of America