Phosphorylation at distinct subcellular locations underlies specificity in mTORC2-mediated activation of SGK1 and Akt

The mechanistic target of rapamycin (mTOR) is a key regulatory node in the signaling pathways that control numerous essential cellular and physiological functions including proliferation, growth, metabolism and ion balance. mTOR is the catalytic core of two functionally distinct multi-protein complexes, mTORC1 and mTORC2. Each complex interacts with a distinct set of targets enabling mTOR to respond uniquely to growth factors, nutrient availability and other stimuli. mTORC1 consists of mLST8, DEPTOR, PRAS40 (also known as AKT1S1) and RAPTOR (also known as RPTOR). mTORC2, also contains mLST8 and DEPTOR, but is defined by the presence of RICTOR, SIN1 (also known as MAPKAP1) and PROTOR (also known as PRR5) (Kennedy and Lamming, 2016). Only mTORC1 is acutely inhibited by the macrolide rapamycin owing to the presence of RAPTOR and absence of RICTOR. Consequently, mTORC1 signaling and physiological function are relatively well defined. Much less is known about mTORC2 signaling and function (Gaubitz et al., 2016).

The best-characterized mTORC2 substrates belong to the AGC family of serine/threonine kinases, which include Akt proteins (hereafter referred to generically as Akt unless a specific isoform is meant), serum/glucocorticoid-induced kinases (SGKs) and protein kinase C (PKC). mTORC2 directly phosphorylates two conserved motifs within the C-terminal tails of these kinases, termed the turn motif and hydrophobic motif (HM). In addition, they also require phosphorylation at a site within their activation loop (T-loop) in the kinase catalytic domain mediated by the 3-phosphoinositide-dependent protein kinase 1 (PDK1, also known as PDPK1). Phosphorylation of the turn motif site is generally thought to occur constitutively, during or soon after translation, and functions to increase protein stability and stabilize the active conformation of the kinase (Alessi et al., 2009). In contrast, HM phosphorylation is induced in response to PI3K activation. Phosphorylation at both the T-loop and HM sites is required for maximal kinase activity (Pearce et al., 2010).

Within the AGC family, SGK and Akt share the greatest degree of homology, as they possess a similar domain structure and ∼55% sequence identity through their catalytic domain (Park et al., 1999). As a result, Akt and SGK recognize a similar substrate consensus motif (R-x-R-x-x-S/T) and phosphorylate an overlapping, but not identical, set of proteins (Brunet et al., 2001; Collins et al., 2003; Pearce et al., 2010). Despite their considerable similarities, Akt and SGK family members exhibit distinct physiological functions in vivo. Akt2 plays a major role in regulation of metabolism (Toker and Marmiroli, 2014), whereas SGK1 is primarily implicated in the control of ion transport (Lang et al., 2006). Accordingly, they are activated under distinct physiological circumstances, and disruption of their function results in distinct pathophysiological consequences. For example, Akt2 dysregulation is implicated in the development of insulin resistance and diabetes (Toker and Marmiroli, 2014), while impaired SGK1 function contributes to defects in blood pressure control and electrolyte balance (Lang et al., 2006). How these related kinases respond to distinct physiological cues, and achieve functional specificity in their signaling outputs is not clear. One mechanism for controlling selective activation of SGK1 and Akt could occur through distinct regulation of their catalytic activities by mTORC2. How mTORC2 selectively regulates substrate activation is not currently known. This issue has important clinical implications in that hypertension is frequently a major complication associated with obesity and diabetes and an obligate parameter for the diagnosis of metabolic syndrome (Carey, 2011). The coincidence of these disease states likely reflects crosstalk in the signaling pathways that regulate Akt and SGK proteins, and emphasizes the need for greater understanding of the regulatory mechanisms that control mTORC2 activity and confer mTORC2 substrate selectivity.

Recently, the mTORC2 subunit SIN1 has been implicated as a critical signaling integrator for mTORC2 activity. SIN1, along with RICTOR, is required for mTORC2 complex assembly (Gaubitz et al., 2016). In addition, SIN1 recruits several mTORC2 substrates to the complex through direct interaction (Cameron et al., 2011; Lu et al., 2011). We previously showed that mutation of a single amino acid (Q68) in SIN1 disrupts the interaction of SIN1 with SGK1 but not with Akt or PKC (Lu et al., 2011). This same mutation also disrupts mTORC2-dependent activation of SGK1, but not of Akt or PKC, demonstrating that mTORC2 can be induced to selectively activate one of its substrates, but not others, through mutation of one of its defining subunits, SIN1. More recently, several groups have demonstrated that growth factor signaling can influence mTORC2 activity through phosphorylation of different subunits. Growth factor-induced phosphorylation of RICTOR at T1135 by p70S6K (also known as RPS6KB1, hereafter S6K) triggers 14-3-3 binding and inhibits mTORC2 substrate activation (Dibble et al., 2009; Treins et al., 2010). Growth factors also modulate mTOR activity through SIN1 phosphorylation at T86 and T398 (Humphrey et al., 2013; Yang et al., 2015; Liu et al., 2013). However, the phosphorylation sites in SIN1 and RICTOR identified so far appear to modify mTORC2 function globally, as opposed to influencing the activity of mTORC2 towards a particular substrate. Indeed, whether the modification state of mTORC2 subunits plays a role in dictating substrate specificity is unknown. In particular, the effect of mTORC2 subunit phosphorylation on SGK HM (S422) phosphorylation has not been addressed.

In this study, we set out to determine whether there are conditions under which mTORC2 selectively activates SGK1 but not Akt. We chose angiotensin II (AngII) as a candidate hormone for selective regulation of SGK1. AngII is a potent regulator of Na⁺ balance and blood pressure homeostasis. In volume depletion, AngII stimulates the Na⁺-retaining hormone aldosterone, and also directly affects salt reabsorption through regulation of ion transporters located in various segments of the kidney tubules (Herrera and Coffman, 2012). Recent evidence supports a role for SGK1 as a mediator of AngII signaling: AngII stimulates induction of SGK1 transcription (Baskin and Sayeski, 2012; Stevens et al., 2008) and knockdown of SGK1 in human proximal tubule cells impairs AngII-induced Na⁺ uptake through the sodium proton exchanger 3 (NHE3) (Stevens et al., 2008). Importantly, AngII stimulates epithelial Na⁺ transport in kidney tubules without affecting Akt-dependent processes such as inhibition of gluconeogenesis. Here, we find that AngII selectively triggers SGK1 S422 phosphorylation without inducing phosphorylation of Akt at either critical activating site, T308 or S473. AngII stimulates conventional PKC-dependent phosphorylation of SIN1 at S128, S315 and S356. Inhibition of conventional (c)PKC activity re-localizes SIN1 and SGK1 from the nucleus and perinuclear compartment to the plasma membrane and selectively blocks SGK1 S422 but not Akt S473 phosphorylation. Thus, we show that mTORC2 achieves specificity in signaling to SGK1 and Akt through phosphorylation of their hydrophobic motif sites at distinct cellular membranes.

To determine whether AngII stimulates mTORC2-dependent SGK1 hydrophobic motif phosphorylation, we treated human embryonic kidney (HEK)-293T cells, expressing the angiotensin type I receptor (AT₁R, also known as AGTR1) and FLAG-tagged SGK1, with AngII either in the presence of vehicle (DMSO), an AT₁R receptor blocker (losartan) or an mTOR catalytic site inhibitor (PP242). HEK-293 cells have the machinery for AngII signaling when AT₁R is expressed (Hein et al., 1997). AngII stimulated SGK1 S422 phosphorylation and this was blocked by inhibition of AT₁R or inactivation of both mTOR complexes (PP242) (Fig. 1A). Phosphorylation of extracellular-regulated kinase 1 and 2 (ERK1/2, also known as MAPK3 and MAPK1, respectively) is a well-known consequence of AT₁R activation (Shah et al., 2004); therefore, we used ERK1/2 phosphorylation as a positive control for activation of AT₁R signaling. By comparing inhibition of mTORC1 activity (rapamycin) with total mTOR inhibition (PP242) (Fig. 1B), we found that AngII-induced SGK1 S422 phosphorylation was not blocked with rapamycin but completely prevented by PP242 suggesting that SGK1 S422 is a substrate for mTORC2 downstream of AT₁R activation. As expected, rapamycin and PP242 prevented phosphorylation of S6K, an mTORC1 target, at its hydrophobic motif, T389. In addition, we found that SGK1 S422 phosphorylation requires PI3K activity as PI3K inhibition (PIK90) prevented S422 phosphorylation (Fig. 1B). Similar to what was seen in HEK-293 cells, we found that AngII also stimulated SGK1 S422 phosphorylation predominantly through mTORC2 in the opossum proximal tubule cell line OKP (Fig. S1A) (Moe et al., 1991). Interestingly, in both HEK-293 and OKP cell lines inhibition of mTORC1 with rapamycin led to a small decrease in SGK1 S422 phosphorylation (Fig. 1B; Fig. S1A).

To further characterize the role of mTORC2 and its subunit SIN1 in regulation of AngII-induced SGK1 S422 phosphorylation, we generated HEK-293T SIN1-knockout (SIN1^−/−) cells by using CRISPR technology (Fig. 1C; Fig. S2). Consistent with our mTOR inhibitor studies, inactivation of mTORC2 by SIN1 deletion abolished both basal and AngII-stimulated SGK1 S422 phosphorylation. Reconstitution of mTORC2 activity by expression of V5-tagged wild-type SIN1 restored SGK1 S422 phosphorylation in response to AT₁R activation (Fig. 1C).

Although AngII strongly stimulated SGK1 S422 phosphorylation, surprisingly, we did not observe an increase in Akt S473 phosphorylation (Fig. 1A,C) indicating that, in response to AngII, mTORC2 targeted the HM site in SGK1 but not Akt. Note that the antibodies used to detect Akt phosphorylation cross-react with all Akt isoforms (at equivalent sites to Akt1 T308 or S473). To confirm that AngII preferentially triggers SGK1 HM phosphorylation, we directly compared AngII and insulin stimulation in SIN1^−/− cells expressing wild-type V5-tagged SIN1 (Fig. 1D). Both AngII and insulin induced SGK1 S422 phosphorylation; however, only insulin induced Akt S473 phosphorylation. We also observed AngII-dependent SGK1 S422 phosphorylation without Akt S473 phosphorylation in OKP cells (Fig. S1A,B).

As a start to identifying a mechanism for distinct regulation of mTORC2-dependent SGK1 activation, we first investigated whether AngII stimulation influenced the interaction between SGK1 and SIN1. By co-immunoprecipitation experiments, we found that the amount of SGK1 associated with SIN1 did not change in response to AngII (Fig. 2A) or in response to inhibition of PI3K or mTOR activity. Instead, we observed a decrease in the mobility of SIN1 in response to AngII by performing western blotting (Fig. 2B), suggesting that SIN1 might become phosphorylated in response to this ligand. To determine whether the AngII-induced SIN1 mobility shift was due to phosphorylation, we immunoprecipitated SIN1–V5 from HEK-293-AT₁R cell extracts followed by incubation with the protein phosphatase from phage λgt10 (Fig. 2B). HEK-293-AT₁R cells stably express HA- and Flag-tagged AT₁R (Hunyady et al., 2002). Phosphatase treatment reversed the AngII-stimulated decrease in SIN1 mobility establishing that AngII had induced phosphorylation of SIN1. In addition to PI3K and mTOR, AT₁R signaling leads to activation of various downstream mediators including PKC, protein kinase D (PKD) and ERK1/2 (Hunyady and Catt, 2006; Rozengurt, 2007). Neither PI3K nor mTOR inhibition prevented the SIN1 mobility shift (Fig. 2A), suggesting these kinases were not the AngII-induced SIN1 kinase. Next, we tried inhibition of ERK1/2 (via MEK inhibition with U0126) or all PKC isoforms (Gö6983) and looked for the effect of these inhibitors on the AngII-induced decreased SIN1 mobility (Fig. 2C). We found that only PKC inhibition completely prevented the AngII-induced mobility shift of SIN1. Furthermore, co-expression of SIN1 and wild-type (WT) PKCα triggered a robust mobility shift in SIN1 in response to PMA that was not observed with a kinase-dead PKCα mutant (PKCαD463N) (Fig. 2D). These results implicated PKC, most likely a conventional isoform, as a mediator of SIN1 phosphorylation in response to AngII signaling.

To determine whether there was a functional consequence for PKC-dependent SIN1 phosphorylation, we examined the effect of cPKC inhibition on SGK1 and Akt HM phosphorylation. Expression of the kinase-dead PKCα mutant (PKCαD463N) completely blocked SGK1 HM phosphorylation in response to AngII but had little effect on basal Akt S473 phosphorylation (Fig. 3A–C). We also assessed SIN1 T86 phosphorylation, a site that previously was shown to correlate with increased mTORC2 activity (Yang et al., 2015). AngII led to an increase in SIN1 T86 phosphorylation that was inhibited under conditions of PKCα inactivation. Consistent with these results, acute pharmacological inhibition of cPKC with LY333531 also blunted AngII-induced SGK1 S422 and SIN1 T86 phosphorylation without inhibiting Akt S473 phosphorylation (Fig. 3D–F). In addition, pretreatment with CID655763, which at 25 µM inhibits both PKD and PKC, impaired AngII-stimulated SGK1 phosphorylation (Fig. 3G). In OKP cells, PKCαD463N overexpression or acute pharmacological inhibition of cPKC also blocked AngII-induced SGK1 S422 phosphorylation (Fig. 3H,I). To determine whether the effect of PKC on mTORC2-dependent SGK1 activation was selective for AT₁R signaling or a more general requirement for SGK1 activation, we examined the effect of PKC inactivation on insulin-stimulated SGK1 and Akt HM phosphorylation. Similar to AT₁R signaling, when PKCα kinase activity was disrupted either by expression of PKCαD463N or pre-treatment with LY333531, SGK1 S422 phosphorylation was markedly reduced in response to insulin compared to control (Fig. 4A,B). In striking contrast, insulin-stimulated Akt S473 phosphorylation was not significantly altered with PKCα kinase inactivation (Fig. 4A,B). Similar results were obtained in mouse embryonic fibroblasts (MEFs) and OKP cells transfected with PKCαD463N and Flag–SGK1 (Fig. 4C,D). Combined, these results suggest that cPKC activity is a general requirement for SGK1 S422 phosphorylation and, importantly, that Akt S473 phosphorylation occurs through a distinct mechanism.

The results described above are consistent with a role for PKC-mediated phosphorylation of SIN1 in regulation of SGK1 S422 phosphorylation. To identify the site(s) of PKC-mediated phosphorylation on SIN1, we performed mass spectrometry on SIN1 immunoprecipitated from HEK-293T cell extracts overexpressing Flag-tagged SIN1 and either WT PKCα or PKCαD463N and stimulated with phorbol 12-myristate 13-acetate (PMA). We observed three serine residues, S128, S315 and S356 (mouse sequence), whose phospho-occupancy increased in the presence of WT PKCα but not with the catalytically inactive PKCαD463N (Fig. 5A; Figs S3, S4A, S5 and Tables S1 and S2). We compared these results to those obtained when SIN1–Flag was immunoprecipitated from extracts of HEK-293 cells expressing AT₁R stimulated with AngII, in the presence or absence of PKC inhibition (Gö6983) (Fig. 5B; Figs S3, S4B, S5 and Tables S1 and S2). In both mass spectrometry experiments, the same three residues were reproducibly identified. S128 is in a canonical PKC recognition motif (K-x-S/T-x-K) and is located between the coiled-coiled (CC) and conserved region in the middle (CRIM) domains in the N-terminus of SIN1. The sequence surrounding S356 is a strong PKD recognition motif (L-x-x-R-S/T) and is found between the Raf-like Ras-binding (RBD) and the pleckstrin homology (PH) domains. The S315 site has characteristics of both PKC and PKD recognition motifs and is located within the RBD. All three residues are conserved in vertebrates (Fig. 5C). In support of our mass spectrometry data, overexpression of PKD led to a similar decrease in SIN1 mobility because PKCα overexpression and the decreased SIN1 mobility was completely blocked by PKD inhibition upon CID755673 treatment (Fig. S6A). G-protein-coupled receptors (GPCRs) and receptor tyrosine kinases (RTKs) activate PKD via two mechanisms: (1) binding to the second messenger diacylglycerol (DAG) followed by (2) conventional or novel PKC phosphorylation at the activation loop and autophosphorylation at the C-terminus (Steinberg, 2012). Consistent with PKD requiring activation by cPKC, co-expression of PKD with PKCαD463N, prevented the SIN1 mobility shift (Fig. S6B). Since overexpression of PKD was sufficient to induce the SIN1 mobility shift (Fig. S6A), we speculated that phosphorylation of a PKD consensus motif at S315 or S356 was likely to cause the delayed SIN1 mobility. To answer this question, we co-expressed PKCα or PKD1 with WT SIN1–V5 or the non-phosphorylatable SIN1 alanine mutants S315A, S356A or the double mutant S315A/S356A. PMA treatment triggered the delay in SIN1 mobility when WT or the S315A SIN1–V5 was expressed. In contrast, decreased SIN1 mobility was not observed when S356A or S315A/S356A SIN1–V5 was expressed (Fig. 5D), indicating that S356 phosphorylation is required for the observed SIN1 mobility shift.

To confirm phosphorylation of SIN1 by PKD at S356 and to determine whether the other sites identified by mass spectrometry were phosphorylated in response to PKC or PKD, we examined SIN1 phosphorylation induced by expression of PKC or PKD with anti-PKC or anti-PKD phospho-substrate antibodies. When PKD and WT SIN1–V5 were co-transfected in SIN1^−/− cells, PMA stimulated SIN1 phosphorylation (as detected by an anti-L-x-x-R-pS/T antibody) and this phosphorylation was prevented by PKD inhibition with CRT0066101 (a specific PKD inhibitor) (Fig. 5E). Mutation of S356 to alanine (S356A) alone did not prevent SIN1 phosphorylation suggesting another site on SIN1 was also phosphorylated by PKD (Fig. 5E). Accordingly, when SIN1^−/− cells were reconstituted with S315A SIN1, phosphorylation of SIN1 in response to PMA was reduced. Expression of the S315A/S356A double alanine mutant reduced SIN1 phosphorylation further, suggesting that PKD phosphorylates SIN1 at both the S315 and S356 sites (Fig. 5E). We also assessed phosphorylation of SIN1 by PKC using an anti-R/K-x-pS-x-R/K (pPKC substrate) antibody. Treatment of SIN1^−/− cells expressing both PKCα and WT SIN1 with PMA induced phosphorylation of SIN1 and this phosphorylation was blocked by PKC inhibition with Gö6976 (which blocks the activity of conventional PKC isoforms) (Fig. 5F). In SIN1^−/− cells reconstituted with the S128A SIN1 mutant, PMA-induced PKC phosphorylation was completely blocked, whereas expression of the S315A mutant exhibited comparable phosphorylation to WT SIN1. This suggests that PKC phosphorylates SIN1 exclusively at S128 and demonstrates the specificity of this antibody for SIN1 phosphorylated at this residue. Thus, based on peptide sequence data and western blot analysis, it is likely that PKC directly phosphorylates SIN1 at S128 and indirectly regulates phosphorylation at S315 and S356 through activation of PKD.

We next determined whether AngII or insulin signaling induces phosphorylation of SIN1 by using the anti-R/K-x-pS-x-R/K antibody which, as was shown in Fig. 5F, recognizes phosphorylation at S128. SIN1 S128 phosphorylation was observed at 5, 15, 30 and 60 min post-AngII stimulation (Fig. 6A). To assess whether cPKC activity was required for AngII-induced SIN1 phosphorylation we co-treated cells with both AngII and LY333531. LY333531 completely prevented the AngII-induced SIN1 S128 phosphorylation. Insulin also stimulated SIN1 S128 phosphorylation (Fig. 6B). Insulin-stimulated SIN1 phosphorylation peaked at 15 min post-treatment and, similar to AngII, was prevented by inhibition of cPKC. Thus, both AngII and insulin treatment elicit cPKC-dependent SIN1 phosphorylation at S128. It is notable that the AngII-induced SIN1 phosphorylation is more sustained than that induced by insulin.

As PKC-dependent SIN1 phosphorylation occurred on a timescale consistent with regulation of mTORC2-dependent SGK1 HM phosphorylation, we hypothesized that PKC-induced SIN1 phosphorylation might be required for AngII-stimulated SGK1 activity. To determine whether PKC exerts its effects on SGK1 HM phosphorylation directly through phosphorylation of SIN1, we examined SGK1 phosphorylation in SIN1^−/− cells reconstituted with WT SIN1, SIN1 with a non-phosphorylatable serine-to-alanine mutation (S128A) or a phospho-mimetic serine-to-aspartate mutation (S128D). We initially focused on the PKC consensus site, S128, since specific inhibition of PKD activity with CRT0066101 did not prevent AngII-induced SGK1 S422 phosphorylation (data not shown). Surprisingly, we observed a similar increase in AngII-stimulated SGK1 S422 phosphorylation when SIN1^−/− cells were reconstituted with either the S128A or S128D SIN1 mutant (Fig. 6C–F). We found similar results for SGK1 S422 phosphorylation in SIN1^−/− cells reconstituted with triple alanine (S128/S315/S356A) or aspartate (S128/S315/S356D) SIN1 mutants (data not shown). Thus, cPKC-dependent phosphorylation of SIN1 does not play a primary role in regulation of SGK1 S422 phosphorylation and is not the mechanism underlying inhibition of SGK1 S422 phosphorylation upon PKC inactivation.

cPKCs have been implicated in the spatial control of signal transduction in cells due to coupling of their membrane binding with kinase activation (Rosse et al., 2010). Importantly, the CRIM domain within SIN1 directly binds mTORC2 targets from the AGC kinase family, including PKC, SGK1 and Akt (Cameron et al., 2011; Lu et al., 2011; Tatebe et al., 2017). Previously, PKCα was implicated in mediating the localization of SIN1, RICTOR and mLST8 with syndecan-4 in lipid rafts of endothelial cells, which is required for proper growth-factor-induced Akt S473 phosphorylation (Partovian et al., 2008). Therefore, we asked whether cPKC activation controls SIN1 and SGK1 subcellular localization in HEK-293 cells and thereby influences SGK1 S422 phosphorylation. Through indirect immunofluorescence and confocal microscopy, we found that, in serum-starved SIN1^−/− cells reconstituted with WT SIN1–V5, SIN1 was localized prominently to the nucleus and to a lesser extent to the cytoplasm and plasma membrane. Activation of exogenously expressed WT PKCα with PMA led to the re-localization of SIN1 from the nucleus to a perinuclear compartment, the cytoplasm and the plasma membrane (Fig. S7). Stimulation of HEK-293-AT₁R cells with AngII also led to redistribution of exogenously expressed SIN1–V5 and Flag–SGK1 from the nucleoplasm to the cytoplasm and the plasma membrane (Fig. 7A,D). Strikingly, in serum-starved cells expressing GFP–PKCαD463N, nuclear localization of SIN1–V5 and Flag–SGK1 was significantly reduced and both proteins were found prominently localized at the plasma membrane (Fig. 7C,F; Fig. S8). Stimulation of these cells with AngII or insulin did not alter the localization of SIN1–V5 or Flag–SGK1 further. RICTOR was also re-localized from the cytoplasm to the plasma membrane in serum-starved cells when co-expressed with SIN1 and PKCαD463N (Fig. 8A). However, when mTOR was co-expressed with SIN1 and PKCαD463N, only a fraction of mTOR re-localized to the plasma membrane, consistent with the existence of two mTOR complexes (Fig. 8B). Interestingly, it has been reported that inhibition of cPKC catalytic activity causes pre-targeting of the inactive protein to the plasma membrane and accelerated agonist-induced translocation compared to that observed for WT PKC (Antal et al., 2014). We also observed reduced SIN1 and SGK1 localization in the nucleoplasm when cPKC activity was inhibited with LY333531, although the plasma membrane targeting was less-pronounced than with overexpression of PKCαD463N (Fig. 7B,E). Collectively, these data suggest that cPKC activation by PMA or AngII triggers the redistribution of SIN1 and SGK1 from a predominantly nuclear localization to a perinuclear compartment, cytoplasm and plasma membrane. However, premature re-localization of SIN1 and SGK1 away from the nucleus or perinuclear compartment through inhibition of cPKC catalytic activity selectively prevents SGK1 S422 but not Akt S473 phosphorylation and is consistent with a model where mTORC2 target specificity is achieved through subcellular partitioning of mTORC2 activity.

How pleiotropic master signaling molecules, such as mTOR, receive specific signaling inputs and respond selectively to modulate distinct physiological processes represents a general problem in signaling biology. In this report, we show that the highly related but functionally distinct kinases, SGK1 and Akt are phosphorylated by mTORC2 at different cellular membranes. cPKC, a critical downstream mediator of AT₁R signaling, regulates SIN1 and SGK1 subcellular localization. Inhibition of cPKC catalytic activity leads to re-localization of SIN1 and SGK1 from nuclear and peri-nuclear subcellular domains to the plasma membrane and inhibits SGK1, but not Akt, HM phosphorylation. Collectively, our results demonstrate that mTORC2-dependent SGK1 phosphorylation occurs at a perinuclear compartment, not the plasma membrane, and suggests that mTORC2 achieves specificity in SGK1 and Akt activation through phosphorylation of their hydrophobic motif sites at distinct subcellular locations.

AngII, a primary component of the renin–angiotensin–aldosterone system (RAAS), is responsible for the physiological control of blood pressure, thirst and Na⁺ balance (Hunyady and Catt, 2006). The physiological and pathophysiological actions of AngII are predominantly mediated via activation of the AT₁R, a typical seven-transmembrane GPCR. In response to AngII, AT₁R couples primarily through G_q/11-mediated inositol phosphate/Ca²⁺ signaling (Hunyady and Catt, 2006). Through both pharmacological and genetic means, we demonstrate that AngII signaling triggers SGK1 S422 phosphorylation through mTORC2, not mTORC1, activity. AngII signaling can in some cases trigger transactivation of growth factor receptor tyrosine kinases (RTKs) and induce Akt S473 phosphorylation (Higuchi et al., 2007; Daub et al., 1996). However, this cross-communication does not occur in HEK-293T cells (Higuchi et al., 2007; Hunyady and Catt, 2006; Shah et al., 2004). Surprisingly, we find that RTK-independent AT₁R activation does not trigger phosphorylation of Akt S473 despite mTORC2 activation and phosphorylation of SGK1 at its comparable HM site, S422. It is now known that many GPCRs generate signals from intracellular compartments, including the nucleus, ER and endosomes (Irannejad et al., 2015). AT₁R has been shown to undergo AngII-mediated internalization and translocation to the nucleus and endosomes (Hunyady and Catt, 2006; Lu et al., 1998). Thus, activation of AT₁R signaling at a localized domain, such as the nucleus, ER or mitochondria-associated ER membranes (MAMs), could underlie the specificity in AngII activation of SGK1 in HEK-293T and OKP cells.

How mTORC2 activation is regulated and why mTORC2 is localized to various subcellular compartments are major unanswered questions in the mTOR signaling field. Several mechanisms for mTORC2 activation have been proposed which include binding of the complex to ribosomes (Zinzalla et al., 2011), interaction of mTOR with the small GTPase Rac1 (Saci et al., 2011) and binding of the complex to PIP₃ at the plasma membrane through the PH domain of SIN1 (Gan et al., 2011; Liu et al., 2015b). In addition, post-translational modifications such as acetylation and phosphorylation of mTORC2 subunits have been shown to modulate mTOR activity (Gaubitz et al., 2016; Humphrey et al., 2013; Liu et al., 2013; Yang et al., 2015). In this study, we identify three serine residues on SIN1 (S128, S315 and S356) that are phosphorylated in response to cPKC activation. However, we find that SIN1 phosphorylation at these sites is not required for AngII-induced SGK1 S422 or Akt S473 phosphorylation. We do observe a trend for enhanced SIN1 T86 and SGK1 S422 phosphorylation in response to AngII when SIN1^−/− cells are reconstituted with the phospho-mimetic (S128D) SIN1 mutant. Notably, no trend was observed for Akt S473 phosphorylation. As SIN1 T86 phosphorylation has been reported to enhance growth factor-stimulated mTORC2 activity towards Akt (Yang et al., 2015), it is possible that AngII-induced phosphorylation of SIN1 at S128 facilitates phosphorylation at SIN1 T86 to potentiate SGK1-associated mTORC2 activity towards SGK1. Future work is needed to determine the precise role for PKC-mediated SIN1 phosphorylation.

SIN1 is the only mTORC2 subunit that contains a lipid-binding domain and has been shown to target mTORC2 to cellular membranes (Ebner et al., 2017; Liu et al., 2015a). mTORC2 has been detected in the cytosol, nucleus, early and late endosomes and plasma membrane in various cell lines including HEK-293T cells (Betz and Hall, 2013; Ebner et al., 2017). Interestingly, a recent study has shown that SIN1 subcellular localization does not change in response to growth factor stimulation or PI3K inhibition, suggesting that SIN1 interacts with cellular membranes independently of phosphatidylinositol (3,4,5)-trisphosphate (PIP₃) (Ebner et al., 2017). Here, we detect a similar pattern of SIN1 localization and also observe that SIN1 localization does not markedly change in response to insulin stimulation (Fig. S8). However, we find that PMA-activated PKCα and AT₁R signaling, which strongly activates PKC, induces translocation of SIN1 and SGK1 from the nucleus to the cytosol, perinuclear membranes and plasma membrane. The relocation of SIN1 from the nucleus to cellular membranes in response to AT₁R signaling suggests that mTORC2 localization is uniquely regulated in response to growth factor- or GPCR-stimulated signaling pathways.

While there is compelling evidence that the PH domain of SIN1 can target mTORC2 to the plasma membrane and other cellular membranes, we (this study) and another group have demonstrated that PKCα plays an important role in the subcellular localization of SIN1 and RICTOR (Partovian et al., 2008). Partovian et al. reported that in syndecan-4-deficient mice, Akt S473 phosphorylation is reduced due to a decrease in the amount of SIN1, RICTOR and PKCα in lipid rafts of endothelial cells (Partovian et al., 2008). The authors were able to restore SIN1 and RICTOR levels and Akt S473 phosphorylation by overexpressing a plasma membrane-targeted form of WT PKCα (myristoylated-WT PKCα) in syndecan-4-knockout endothelial cells. Consistent with our results, these authors found that: (1) PKCα can target SIN1 and RICTOR to the plasma membrane, and (2) Akt S473 phosphorylation occurs predominantly at the plasma membrane. However, when this study was published it was unclear how PKCα mediates relocation of SIN1 and RICTOR. From our previous work and others, it is now known that SIN1 interacts directly with PKC and SGK1 through its CRIM domain (Cameron et al., 2011; Lu et al., 2011; Tatebe and Shiozaki, 2017). Thus, we find that SIN1 can recruit mTORC2 and SGK1 to cellular membranes, not only through its PH domain but additionally through its interaction with cPKC. One limitation of our study is the use of SIN1 and SGK1 overexpression since overexpression of these proteins might influence their subcellular localization. However, we see similar results when SIN1 is reconstituted in SIN1^−/− cells at levels that reflect endogenous SIN1 expression (see Fig. S8) and with other mTORC2-specific proteins (RICTOR) (Fig. 8A). An interesting direction for future work is understanding how localization of mTORC2 is coordinately regulated by direct binding of lipids to the SIN1 PH domain and indirectly through SIN1 interaction with PKC.

cPKCs interact with cellular membranes through their DAG-binding cysteine-rich zinc finger (C1) domains and Ca²⁺-dependent phospholipid binding C2 domain (Rosse et al., 2010). cPKC requires three sequential priming phosphorylations that stabilize the protein in an active conformation and, importantly, mask the DAG-binding C1 domains (Rosse et al., 2010). Kinase-dead cPKC mutants do not fold correctly, allowing exposure of the C1 domains and pre-targeting of the kinase to the plasma membrane in the absence of stimulation (Antal et al., 2015). We find that inactivation of cPKC catalytic activity redistributes SIN1 and SGK1 to the plasma membrane in the absence of hormonal stimulation. Aberrant pre-targeting to the plasma membrane likely explains the effect of PKCαD463N expression on the dramatic SIN1 and SGK1 re-localization. Unexpectedly, when SIN1 and SGK1 are pre-targeted by cPKC to the plasma membrane, SGK1 phosphorylation by mTORC2 is prevented. In contrast, Akt S473 phosphorylation is only slightly reduced. Why mTORC2 does not phosphorylate SGK1 when localized in advance at the plasma membrane is a remaining question. One possibility is that catalytically inactive cPKC relocates SIN1, RICTOR and SGK1 to the plasma membrane in the absence of mTOR. Interestingly, Partovian et al. observed that mTOR localization in lipid rafts is not mediated by PKCα suggesting that mTORC2 could form locally at cellular membranes (Partovian et al., 2008). Alternatively, SGK1-associated mTORC2 might require an activating signal that is not present at the plasma membrane. Consistent with this hypothesis, a recent study has shown that plasma membrane-localized mTORC2 is constitutively active whereas endosomal and ER-localized mTORC2 activity requires PI3K activity (Ebner et al., 2017).

Although SGK1 and Akt share considerable similarity within their catalytic domains, their N-terminal domains diverge markedly: Akt isoforms preferentially bind to PIP₃ found at the plasma membrane through their N-terminal pleckstrin homology (PH) domain (Pearce et al., 2010). Akt is also found at the ER, mitochondria and MAMs. However, it is generally thought that T308 phosphorylation by PDK1 occurs at the PM followed by translocation of Akt to internal membranes (Hemmings and Restuccia, 2012). SGK isoforms do not contain a PH domain and instead bind PI(3)P, PI(4)P and PI(5)P through a Phox homology (PX)-like domain in their N-terminus (Pao et al., 2007). A significant consequence of the lack of a PH domain is that, in contrast to Akt, SGK1 requires mTORC2 S422 phosphorylation to create a docking site for PDK1 and the subsequent phosphorylation of its activation loop site residue T256 (Collins et al., 2003). SGK1 has also been found in multiple subcellular compartments including the nucleus (Buse et al., 1999), mitochondria (Engelsberg et al., 2006), endosomes (Engelsberg et al., 2006) and ER (Arteaga et al., 2006; Bogusz et al., 2006; Soundararajan et al., 2005). Consistent with our results, SGK1 has been shown to shuttle between the nucleus, cytoplasm and plasma membrane in response to hormones and other stimuli. Notably, mTORC2 is also reported to shuttle between the nucleus and the cytoplasm in response to rapamycin and the cell cycle (Rosner and Hengstschläger, 2008, 2011). There is evidence that the N-terminal region of SGK1 is responsible for its targeting to cellular membranes, but whether this is through direct binding of SGK1 to phosphoinositides is not clear (Pao et al., 2007). Moreover, it is not known whether SGK1 activation requires localization to a particular membrane domain or organelle. We find that SGK1 localization is influenced by cPKC, presumably through interaction with SIN1. In addition, our findings strongly suggest that, as opposed to Akt, SGK1 is subject to mTORC2 phosphorylation and activation at a perinuclear compartment.

SGK1 is unique among protein kinases in that its activity is regulated at both the transcriptional and enzymatic levels. SGK1 mRNA expression is acutely induced in response to a variety of hormones (Lang et al., 2006). Moreover, SGK1 protein abundance is also regulated post-transcriptionally by rapid ubiquitin-mediated proteolysis involving ER localization and the ER-associated degradation (ERAD) pathway (Arteaga et al., 2006; Brickley et al., 2002; Soundararajan et al., 2005). One explanation for the tight multi-level regulation of SGK1 expression and activation is its key role in the control of blood pressure and K⁺ balance through activation of Na⁺ reabsorption and K⁺ excretion. Our results suggest an intriguing possibility that SGK1 activation occurs in close proximity to its translation, coupling the acute increase in SGK1 protein level with its activation.

In conclusion, our results uncover an important distinction in regulation of the highly related AGC-kinases Akt and SGK1: mTORC2-dependent Akt S473 phosphorylation occurs at the plasma membrane whereas SGK1 S422 phosphorylation occurs at a perinuclear compartment. Moreover, we are able to uncouple activation of Akt and SGK1 by insulin through inhibition of cPKC activity and mis-localization of SGK1 to the plasma membrane. These findings provide new mechanistic insight into how the signaling pathways that control electrolyte balance and metabolic homeostasis are uniquely regulated and will provide a framework for further studies aimed at identifying novel therapies for hypertension-associated metabolic disease.

Dulbecco's modified Eagle medium (DMEM), fetal bovine serum (FBS) (JRS), sodium pyruvate and non-essential amino acid supplement were purchased from the UCSF Cell Culture Facility. RIA-grade BSA was purchased from Sigma. L-Glutamine and trypsin were from Corning. G418 was from Gibco. Polyethyleneimine ‘MAX’ (molecular mass 40,000 Da) transfection reagent was from Polysciences (Cat. #24756). PP242 and PIK90 were kind gifts from Kevan Shokat, UCSF, CA. Gö6986 and Gö6976 were from Calbiochem. CID755653, CRT0066101 and GSK650394 were from Tocris. U0126 was from Cell Signaling Technology. LY333531 was from Enzo Life Sciences. Lambda phosphatase was from New England Biolabs (NEB). Losartan, Human Angiotensin II and PMA were from Sigma. Humulin (human insulin) was purchased from the San Francisco General Hospital pharmacy. Bradford Assay Reagent was purchased from ThermoFisher Scientific (Cat. #1856209). 4′,6′-diamidino-2-phenylindole (DAPI) was from ThermoFisher Scientific (Cat #D1306). Fluoromount-G mounting medium was from Southern Biotech (Cat. #0100-01).

Antibodies used for western blotting were against the following: phosphorylated (p)SGK1 S422 (1:1000, sc-16745, Santa Cruz Biotechnology); total SGK1 (1:2000, 5188, Sigma); pAkt S473 (1:1000, 4060, Cell Signaling Technology; CST); total Akt (1:1000, 4691, CST); pERK1/2 T202/Y204 (1:1000, 4370, CST); Total ERK1/2 (1:1000, 4695, CST); pS6K T389 (1:1000, 9234, CST); total S6K (1:1000, 2708, CST); SIN1 (1:1000, 05-1044, Millipore); pSIN1 T86 (1:1000, 14716, CST); V5–HRP (1:2500, MA5-15253, Invitrogen); pPKD S744/S748 (1:1000, 2054, CST); pPKD S916 (1:1000, 2051, CST); Total PKD (1:1000, 90039, CST); pPKC substrate (1:1000, 2261, CST); pPKD substrate (1:1000, 4381, CST); GFP (1:10,000, 8371-2, Clontech); HA–HRP (1:5000, PA1-29751, ThermoFisher Scientific); and tubulin (1:5000, T9026, Sigma).

Antibodies used for immunoprecipitation were against the following: anti-Flag M2 agarose affinity gel (5-10 µl agarose/1000 µg protein, A2220, Sigma); anti-V5 (∼1 µg/1000 µg protein, 37-7500, Invitrogen); and anti-V5 agarose affinity gel (5-10 µl agarose/1000 µg protein, A7345, Sigma).

Antibodies used for immunofluorescence were against the following: SIN1 (1:100, 05-1044, Millipore); SGK1 (1:2000, 5188, Sigma); Alexa-Fluor-647-conjugated donkey anti-rabbit IgG (1:100, A35173, ThermoFisher Scientific); Alexa555-conjugated goat anti-mouse IgG (1:100, A32727, ThermoFisher Scientific); and Myc (1:2000, M4439, Sigma).

Flag-SGK1, SIN1-V5, and SIN1-Flag in pMO Vector were generated as described previously (Lu et al., 2011). SIN1-V5 (S128A), (S315A), (S356A), (S315A/S356A), (S128A/S315A), (S128A/S315A/S356A) and (S128D/S315D/S356D) were generated using the Quikchange II site-directed mutagenesis kit (Agilent) with pMO-SIN1-V5 as template according to the manufacturer's instructions. AT₁R was purchased from Origene. GFP–PKCα and GFP–PKCα(D463N) were kind gifts from Peter Parker, Francis Crick Institute, London, UK. HA–PKD1 and HA-PKD(S738E/S742E) were Addgene plasmids #10808 and #10810 (deposited by Alex Toker). Myc–RICTOR and Myc–mTOR were Addgene plasmids #1860 and #1861 (deposited by David Sabatini) (Sancak et al., 2010; Sancak et al., 2008; Sarbassov et al., 2004).

A single guide (sg)RNA with the sequence, 5′-GTATTCCAAATTTGATGGCA-3′, was designed for targeting the 3′ end of exon 3 of the human MAPKAP1 genomic locus. The sgRNA sequence driven by a U6 promoter was cloned into the plentiCRISPR V2 vector (Addgene plasmid #52961; deposited by Feng Zhang) that also expresses Cas9 using standard subcloning techniques. The lentiviral plasmid DNA was then packaged into lentivirus by co-transfection with Virapower (Invitrogen) in HEK293FT cells. Supernatant containing lentivirus was used to infect HEK293T cells and then infected cells were selected in puromycin (3 μg/ml). Single colonies were selected by fluorescence-activated cell sorting (FACS) into a 96-well plate, and expanded and tested for SIN1 expression by western blotting.

HEK293T, HEK293-AT₁R, and HEK293T-SIN1^−/− cells were maintained in DMEM with 2 mM L-glutamine and 10% FBS. HEK293-AT₁R cells (a kind gift from Tamas Balla, Intramural Research Program, NIH NICHD, Baltimore, MD) stably express double HA- and Flag-tagged AT₁R. HEK293FT cells (ATCC) were used to produce high-titer lentiviral particles and were maintained in DMEM with sodium pyruvate, non-essential amino acids, 2 mM L-glutamine, 500 µg/ml G418 and 10% FBS. The opossum kidney proximal tubule cell line (OKP) (a kind gift from Orson Moe, UTSW, Dallas, TX) was maintained in DMEM with 2 mM glutamine and 10% FBS. These cell lines were routinely tested for mycoplasma contamination. All cell lines were acquired from reputable sources but have not recently been authenticated. All cell lines were transfected using polyethyleneimine ‘MAX’ (molecular mass 40,000 Da). For all experiments, cells were serum-starved in DMEM containing 0.1% BSA overnight. For inhibitor assays, cells were treated for 15–30 min with 10 µM losartan, 300 nM PP242, 25 nM rapamycin, 5 µM Gö6976, 5 µM Gö6983, 25 µM CID655763, 5 µM CRT0066101, 200 nM LY333531, 1 µM PIK90, 10 µM U0126 or vehicle (DMSO), followed by 200 nM angiotensin II (AngII) or 200 nM insulin as indicated in figure legends.

Cells were rinsed once with ice-cold PBS, lysed in 1% Triton X-100 buffer (40 mM HEPES pH 7.5, 1 mM EDTA pH 8, 10 mM sodium pyrophosphate, 10 mM glycerophosphate, 50 mM sodium fluoride, 120 mM sodium chloride and 1% Triton X-100), and centrifuged at 10,000 r.p.m. for 10 min. Supernatant (cell extract) was removed and protein content estimated by performing a Bradford assay. Cell extract (15–40 μg protein) was separated by SDS-PAGE and transferred to PVDF membranes. To immunoprecipitate Flag–SGK1, 0.1–1 mg of cell extract protein was rotated overnight at 4°C with 10–20 µl 50% slurry anti-FLAG affinity gel. The agarose beads were collected by centrifugation, washed three times with 1% Triton X-100 cell lysis buffer, boiled and denatured in 1× Laemmli sample buffer, separated by SDS-PAGE and transferred to PVDF membrane. To immunoprecipitate SIN1–V5, 50–200 µg of cell extract protein was rotated overnight at 4°C with ∼1 µg anti-V5 antibody or 10–20 µl 50% slurry anti-V5-affinity gel. For immunoprecipitations with unconjugated antibody, 10 µl 50% slurry protein A/G-conjugated agarose beads (Santa Cruz Biotechnology) was added and lysates were rotated at 4°C for an additional hour. The agarose beads were collected by centrifugation (8,000 r.p.m. for 30 s), washed three times with cell lysis buffer, boiled and denatured in 1× Laemmli sample buffer, separated by SDS-PAGE and transferred to PVDF. Membranes were blocked in TBS plus 0.1% Tween-20 (TBS-T) containing 5% skim milk and incubated with the relevant primary antibodies for either 1–2 h at room-temperature (total SGK1, pAkt S473, tubulin, GFP, V5, and HA) or overnight at 4°C in blocking buffer containing 5% BSA (phospho-antibodies) or 5% milk (all other antibodies). After incubation with primary antibodies, membranes were washed in TBS-T and then incubated with HRP-labeled secondary antibodies for 1 h. Membranes were washed again in TBS-T, incubated with ECL reagent (GE Healthcare) and exposed to film. Quantification of western blots was performed on scanned films with NIH ImageJ Software.

Cells were fixed with 3.7% formaldehyde in modified Brinkley Buffer 1980 (80 mM PIPES pH 6.8, 1 mM CaCl₂ and 1 mM MgCl₂) for 20 min at room temperature. The fixation solution was removed by washing in PBS. The fixed cells were permeabilized and blocked by incubation in TBS containing 3% BSA and 0.1% Triton X-100 for 20 min. Primary antibodies were diluted in TBS containing 3% BSA and 0.1% Triton X-100. Cells were co-stained with DAPI to identify nuclei. Coverslips were mounted on slides in Fluoromount-G. Z-stack images were collected on a Yokagawa CSU22 spinning disk confocal microscope with a 40×, 0.95 NA, plan APO dry objective. These are all adjectives describing the objective. The middle slice of the z-stack was used for all representative images. Quantification of images was performed on average intensity z projections using ImageJ software (http://rsb.info.nih.gov/ij). For each image, the percentage of nuclear-localized SIN1 and SGK1 was calculated by first selecting the nuclear region of interests (ROIs) in the DAPI channel and measuring fluorescence intensity within those ROIs in either the SIN1 (Alexa Fluor 555) or SGK1 (Alexa Fluor 647) channels. Then, ROIs were created for total SIN1 or SGK1 and fluorescence intensity was measured in the appropriate channel. The percentage nuclear localization was determined by dividing nuclear fluorescence intensity by total SIN1 or SGK1 fluorescence intensity. Analysis of SIN1 and SGK1 intensity profiles was carried out using the ImageJ Plot Profile function.

HEK-293T cells were transiently transfected with pMO empty vector, SIN1-flag and WT GFP–PKCα or SIN1–Flag and GFP PKCα(D463N). After overnight serum starvation, cells were stimulated with 1 µM PMA for 15 min. Cells were lysed and SIN1 immunoprecipitated from 5 mg protein extract by overnight incubation with anti-Flag Affinity agarose. In separate experiments, HEK-293-AT₁R cells were transiently transfected with empty pMO vector or SIN1–Flag. After overnight serum starvation, cells were stimulated with 200 nM Ang II either in combination with vehicle (DMSO), 5 µM Gö6983 or 5 µM Gö6976. Cells were lysed and SIN1 immunoprecipitated from 5 mg protein extract by overnight incubation with anti-Flag-conjugated agarose beads (Sigma). Immunoprecipitated SIN1 was separated on SDS-PAGE and visualized by staining with Coomassie Blue (BioRad).

Gel pieces containing SIN1–flag were excised and digested in-gel as described previously (Rosenfeld et al., 1992). The extracted digests were vacuum-evaporated and resuspended in 10 µl 0.1% formic acid in water, and analyzed by nanoflow ultra-performance liquid chromatography tandem mass spectrometry (UPLC-MS/MS). Peptides were separated by nano-flow liquid chromatography using a 75-µm×150-mm reverse phase C18 column PepMap^® C18 EasySpray column (ThermoFisher Scientific) packed with 3 µm particles, at a flow rate of 300 nl/min in a NanoAcquity UPLC system (Waters). Mobile phase A was 0.1% formic acid in water, and mobile phase B was 0.1% formic acid in acetonitrile. Following equilibration of the column in 2% solvent B, an aliquot of each digest (5 µl) was injected, then the organic content of the mobile phase was increased linearly to 30% over 60 min, and then to 50% in 2 min. The liquid chromatography eluate was coupled to a hybrid linear ion trap-Orbitrap mass spectrometer (LTQ-Orbitrap Velos, ThermoFisher Scientific). Peptides were analyzed in positive ion mode and in information-dependent acquisition mode to automatically switch between MS and MS/MS acquisition. MS spectra were acquired in profile mode using the Orbitrap analyzer in the m/z range between 350 and 1500. For each MS spectrum, the six most-intense multiple charged ions over a threshold of 1000 counts were selected to perform collision-induced dissociation experiments. Product ions were analyzed in the linear ion trap in centroid mode. A dynamic exclusion window of 0.5 Da was applied that prevented the same m/z from being selected for 60 s after its acquisition.

Peak lists were generated using PAVA in-house software (Guan et al., 2011), based on the RawExtract script from Xcalibur v2.4 (ThermoFisher Scientific). The peak lists were searched against the human and murine subset of the SwissProt database of March 21 2012, containing 36775 entries, using in-house ProteinProspector version 5.10.17 (a public version is available on line) (Clauser et al., 1999). A randomized version of all entries was concatenated to the database for estimation of false discovery rates in the searches. Peptide tolerance in searches was 20 ppm for precursor and 0.6 Da for product ions, respectively. Enzyme specificity was set to trypsin or GluC as appropriate. Peptides containing two miscleavages were allowed. Carboxymethylation of cysteine was allowed as constant modification; acetylation of the N terminus of the protein, pyroglutamate formation from N terminal glutamine, oxidation of methionine, and phosphorylation of serine, theonine and tyrosine were allowed as variable modifications. The number of modifications was limited to two per peptide. A minimal ProteinProspector protein score of 20, a peptide score of 15, a maximum expectation value of 0.004 for protein and 0.02 for peptides (for tryptic searches), or 0.001 for protein and 0.005 for peptides (for GluC searches), and a minimal discriminant score threshold of 0.0 were used for identification criteria. The false discovery rate (FDR) was below 1%.in all cases

For relative quantitation of the phosphopeptide abundances between samples, the total intensity for precursor ions on the MS traces was integrated during the elution time using the Xcalibur Qual browser (ThermoFisher Scientific) or Skyline (Schilling et al., 2012). Levels were normalized between samples using non-phosphorylated peptides of SIN1, and expressed as ratios to the un-stimulated sample.

Statistical analysis was performed in Microsoft Excel using the Analysis Toolpak Add-In. Sample sizes (biological replicates) are listed in the figure legends. One-way ANOVA was used to determine differences between groups followed by unpaired two-tailed t-tests.

We thank David Bryant (Beatson Institute, Glasgow, Scotland, UK) for reagents and for critical review of the manuscript. We also thank Kaveh Ashrafi (UCSF, San Francisco, CA) for critical review of the manuscript. We thank Brad Heller for reagents and methods used in generation of CRISPR-mediated HEK-293T^SIN1−/− cells. We also thank Roshanak Irannejad and Jacob Bendor for methods and helpful discussions regarding immunostaining and quantification of confocal images.