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Phosphorylation of eIF2α triggered by mTORC1 inhibition and PP6C activation is required for autophagy and is aberrant in PP6C-mutated melanoma

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Science Signaling  10 Mar 2015:
Vol. 8, Issue 367, pp. ra27
DOI: 10.1126/scisignal.aaa0899

Abstract

Amino acid deprivation promotes the inhibition of the kinase complex mTORC1 (mammalian target of rapamycin complex 1) and activation of the kinase GCN2 (general control nonrepressed 2). Signaling pathways downstream of both kinases have been thought to independently induce autophagy. We showed that these two amino acid–sensing systems are linked. We showed that pharmacological inhibition of mTORC1 led to activation of GCN2 and phosphorylation of the eukaryotic initiation factor 2α (eIF2α) in a mechanism dependent on the catalytic subunit of protein phosphatase 6 (PP6C). Autophagy induced by pharmacological inhibition of mTORC1 required PP6C, GCN2, and eIF2α phosphorylation. Although some of the PP6C mutants found in melanoma did not form a strong complex with PP6 regulatory subunits and were rapidly degraded, these mutants paradoxically stabilized PP6C encoded by the wild-type allele and increased eIF2α phosphorylation. Furthermore, these PP6C mutations were associated with increased autophagy in vitro and in human melanoma samples. Thus, these data showed that GCN2 activation and phosphorylation of eIF2α in response to mTORC1 inhibition are necessary for autophagy. Additionally, we described a role for PP6C in this process and provided a mechanism for PP6C mutations associated with melanoma.

INTRODUCTION

Intracellular amino acid deprivation results in the activation of cellular adaptive mechanisms. Two responses to amino acid deprivation are inhibition of the mammalian target of rapamycin (mTOR) kinase and phosphorylation of eukaryotic initiation factor 2α (eIF2α), and these responses play key roles in the induction of macroautophagy, hereafter referred to as autophagy. During autophagy, organelles and misfolded or aggregated cytoplasmic proteins are targeted for lysosomal degradation, thus ridding the cells of potentially deleterious proteins and generating free amino acids in an attempt to maintain metabolic homeostasis. Because autophagy can provide a prosurvival mechanism for cells growing in a hostile tumor microenvironment, elucidating the pathways that regulate autophagy, and specifically identifying druggable targets such as kinases and phosphatases, could lead to new therapies for cancer and other diseases in which cellular stress and autophagy play important roles.

mTOR is a serine/threonine protein kinase that exists as two complexes: mTORC1, which contains raptor, and mTORC2, which contains rictor [reviewed in (1)]. mTORC1 inhibition by amino acid deprivation or treatment with the bacterial product rapamycin attenuates protein synthesis and suppresses cell growth through the dephosphorylation of the eukaryotic initiation factor 4E binding protein 1 (4E-BP1). It also induces autophagy in part by enabling the kinase ULK1 [uncoordinated family member (unc)-51–like kinase 1] to form an autophagy initiation complex with ATG13 (2). The phosphorylation of eIF2α, which also occurs with amino acid deprivation as well as with endoplasmic reticulum (ER) and other cellular stresses, similarly attenuates global protein synthesis [reviewed in (3)]. However, eIF2α phosphorylation also paradoxically increases the translation of select mRNAs, including activating transcription factor 4 (ATF4), which encodes a transcription factor that transactivates the autophagy genes LC3B and ATG5 (47).

Although amino acid deprivation inhibits mTOR, it does not induce autophagy in the absence of eIF2α phosphorylation (8, 9), suggesting that the inhibition of mTOR with amino acid deprivation is not sufficient to induce autophagy and that there could be coordination between these two amino acid–sensing mechanisms. Indeed, both TORC1 inhibition and eIF2α phosphorylation increase 4E-BP1 expression (10) and co-coordinately regulate the translation of select mRNAs (11). Whereas crosstalk of the mTOR and eIF2α signaling pathways has not been demonstrated in mammalian cells, in Saccharomyces cerevisiae, TORC1 inhibition leads to the activation of the mRNA encoding the phosphatase suppressor of initiation of transcription 4 (SIT4), which dephosphorylates and activates the kinase general control nonrepressed 2 (GCN2), which in turn phosphorylates eIF2α (12). GCN2 is also activated by TORC1 inhibition in Schizosaccharomyces pombe (13). The mammalian homolog for SIT4 is PP6C, which encodes the catalytic subunit of serine/threonine protein phosphatase 6; in contrast to other members of the mammalian PP2A-like phosphatases, PP6C rescues SIT4 deletion in S. cerevisiae and S. pombe (14, 15). PP6C, but not PP2A or PP4, can individually associate with three regulatory proteins, PP6R1, PP6R2, and PP6R3, which bridge the phosphatase to substrate proteins (16), a feature that may contribute to its specific effect. Although two independent groups have identified driver PP6C mutations in about 10% of all melanomas, the biological consequences of these mutations are unknown (17, 18).

Despite the similarities between the SIT4 and PP6C genes, the role of PP6C in mTORC1-induced phosphorylation of eIF2α has not been established. We thus investigated whether mTOR inhibition contributes to eIF2α phosphorylation and whether this potential relationship affects the induction of autophagy. We specifically addressed whether PP6C could play a role in linking these two amino acid–sensing mechanisms and whether this signaling pathway is affected by the PP6C mutations identified in melanoma.

RESULTS

mTOR inhibition leads to phosphorylation of eIF2α

In various transformed cell lines treated with the mTORC1 inhibitor rapamycin, eIF2α phosphorylation was increased, an effect similar to that seen in cells treated with the N-linked glycosylation inhibitor tunicamycin, an inducer of ER stress and activator of the eIF2α kinase PERK (Fig. 1A). Rapamycin treatment also led to a time-dependent increase in eIF2α phosphorylation in primary mouse and human fibroblasts (Fig. 1B). To confirm that mTORC1 inhibition was responsible for the induction of eIF2α phosphorylation, we used U2OS osteosarcoma cells depleted of a necessary component of TORC1 (raptor) or of TORC2 (rictor), as a control. As expected, depletion of rictor decreased the phosphorylation of Akt, whereas both raptor depletion and treatment with the mTORC1 inhibitor rapamycin decreased the phosphorylation of 4E-BP1 (Fig. 1C). Raptor depletion and rapamycin treatment, but not rictor depletion, also led to phosphorylation of eIF2α. Together, these findings suggest that the phosphorylation of eIF2α is a general response to mTORC1 inhibition.

Fig. 1 mTORC1 inhibition induces the phosphorylation of eIF2α.

(A) Cell lines were treated with 100 nM rapamycin (Rap) or tunicamycin (Tm) for 6 hours, and eIF2α phosphorylation was assessed by immunoblot. (B) Primary MEFs and BJhTERT cells were treated with rapamycin for the indicated times or tunicamycin, and eIF2α phosphorylation was assessed. (C) U2OS cells were treated with rapamycin or tunicamycin or depleted of rictor or raptor, and eIF2α phosphorylation was assessed. In (A) to (C), representative blots and graphs showing the average and SE for three biological replicates are displayed. CTL or C, control. P, phosphorylated.

mTORC1-regulated autophagy requires phosphorylation of eIF2α

We examined the effect of rapamycin in eIF2α+/+ mouse embryonic fibroblasts (MEFs) and in MEFs in which wild-type eIF2α had been replaced with a mutant that cannot be phosphorylated (eIF2αS51A/S51A) (19). We noted that the rapamycin-induced expression of multiple ATF4 downstream transcriptional targets (ATF3, CHOP, and the CHOP target gadd34) and of transcripts important for the induction of autophagy [LC3B and ATG5 (4, 6, 7)] was higher in eIF2α+/+ MEFs than in eIF2αS51A/S51A MEFs (Fig. 2A), although the protein abundance of ATF4 was similar in both types of MEFs (Fig. 2B).

Fig. 2 Phosphorylation of eIF2α is necessary for rapamycin-induced autophagy.

(A) eIF2α+/+ and eIF2αS51A/S51A MEFs were treated with rapamycin for the indicated times, and the expression of ATF4 transcriptional targets was assessed by real-time polymerase chain reaction (PCR). (B) eIF2α+/+ and eIF2αS51A/S51A MEFs stably expressing GFP-LC3 were treated with 300 nM rapamycin for the indicated times (hours) or tunicamycin for 24 hours. Cell lysates were immunoblotted for GFP and phosphorylated eIF2α (Ser51). (C) eIF2α+/+and eIF2αS51A/S51A MEFs were treated with 300 nM rapamycin for 8 hours, and endogenous LC3 foci were visualized and counted (scale bar, 25 μm). (D) ATF4+/+and ATF4−/− MEFs stably expressing GFP-LC3 were treated with 300 nM rapamycin for the indicated times (hours) or tunicamycin for 24 hours, and the lysates were immunoblotted for GFP. (E) ATF4+/+and ATF4−/− MEFs were treated and analyzed as in (B) (scale bar, 25 μm). (F) eIF2α+/+and eIF2αS51A/S51A MEFs were treated with 100 nM rapamycin in the absence or presence or 60 μM chloroquine (CQ) or chloroquine alone, and immunoblots were performed. Total eIF2α serves as a loading control. (G) TSC+/+ (top) and TSC−/− (bottom) MEFs, expressing either gadd34 or a control, were treated and analyzed as in (F). (H) ULK1/2+/+ (top) and ULK1/2−/− (bottom) MEFs, expressing either gadd34 or a control, were treated as in (G), and immunoblots were performed. (I) eIF2α+/+ (top) and eIF2αS51A/S51A (bottom) MEFs were treated with 300 nM rapamycin for 72 hours, and cell size was measured by forward scatter flow cytometric analysis. Representative analyses are displayed along with changes in fluorescence intensities from two biological replicates are shown. (J) eIF2α+/+and eIF2αS51A/S51A MEFs were treated with 300 nM rapamycin for the times indicated, and proliferation was determined by crystal violet staining and optical density (OD) measurement. The graphs in (A) and (J) show the average and SE from three biological replicates. In (C) and (E), P values were determined by Wilcoxon rank sum test from three independent experiments with more than 200 cells counted in each experiment. Representative blots from two biological replicates are displayed in (B), (D), (F), and (H). In (G), P values were determined by Student’s t test from n = 3 biological replicates.

We then assessed autophagy using several complementary standard assays including the formation of the faster migrating LC3II from LC3I, the formation of free green fluorescent protein (GFP) from an LC3-GFP fusion protein in autophagosomes, and the migration of endogenous LC3 from a diffuse cytoplasmic distribution to punctate intracellular foci (20). Although rapamycin induced phosphorylation of eIF2α and autophagy in eIF2α+/+ MEFs (as documented by increased conversion of LC3I to LC3II, generation of free GFP, and endogenous LC3 foci), it did not induce autophagy in eIF2αS51A/S51A MEFs (Fig. 2, B and C). Confirming the importance of ATF4 in mTORC1-mediated regulation of autophagy, there was minimal conversion of LC3I to LC3II, free GFP generation, or endogenous LC3 foci induced in rapamycin-treated ATF4−/− MEFs compared to ATF4+/+ MEFs (Fig. 2, D and E). Autophagy was blunted in rapamycin-treated eIF2αS51A/S51A MEFs even in the presence of an inhibitor of lysosomal acidification (chloroquine), indicating that the phosphorylation of eIF2α is necessary to induce autophagy and does not simply suppress the clearance of autophagosomes (Fig. 2F). Consistent with these findings, expression of gadd34, which encodes an eIF2α phosphatase, decreased both p62 degradation and the conversion of endogenous LC3I to LC3II in rapamycin-treated MEFs, in contrast to the expression of a phosphatase-dead control (21) (Fig. 2G, top). In cells with constitutive activation of mTOR because of the absence of the inhibiting tuberous sclerosis complex (TSC), rapamycin did not induce autophagy, confirming that suppression of mTOR activity is necessary for rapamycin’s regulation of autophagy (Fig. 2G, bottom). The expression of gadd34 in TSC2−/− MEFs still decreased autophagic flux in response to chloroquine treatment, although to a lesser degree than in TSC2+/+ MEFs, indicating that even in cells with increased activity of mTOR, suppressing the phosphorylation of eIF2α could diminish basal autophagy.

Active mTORC1 inhibits autophagy by phosphorylating the kinases ULK1 and ULK2, the mammalian homologs of yeast ATG1, which is essential for autophagy. Although the ULK1/2 complex is required for the induction of autophagy in response to amino acid deprivation, autophagy is potently induced by glucose deprivation and ammonia and more mildly with rapamycin treatment in ULK1/2-deficient cells, suggesting that autophagy regulation and mTOR signaling include a ULK-independent pathway (20, 2224). We therefore investigated whether mTORC1 regulation of autophagy through eIF2α phosphorylation was independent of ULK1/2 activity. Dephosphorylation of eIF2α by expression of gadd34 suppressed autophagy induction in response to rapamycin treatment in wild-type MEFs, although mild autophagic flux was still present as revealed by an increase in the conversion of LC3I to LC3II and in the generation of free GFP with concomitant treatment with rapamycin and chloroquine (Fig. 2H, top). As previously reported (23, 24), ULK1/2 deficiency only modestly blunted basal autophagy, as demonstrated by the continued conversion of LC3I to LC3II and the generation of free GFP in response to chloroquine treatment of these cells (Fig. 2H, bottom). In addition, rapamycin treatment also led to an induction of autophagy even in the absence of ULK1/2, as demonstrated by free GFP generation and conversion of LC3I to LC3II with rapamycin or rapamycin and chloroquine treatments. Both basal autophagy and rapamycin-induced autophagy were diminished by overexpressing gadd34 and blunting eIF2α phosphorylation in ULK1/2−/− cells, confirming that autophagy can be regulated by mTORC1 in a ULK1/2-independent pathway.

Rapamycin inhibited other mTORC1-regulated cellular functions, namely, cell size (Fig. 2I) and proliferation (Fig. 2J), to similar extents in eIF2αwild-type and eIF2αS51A/S51A MEFs. Thus, phosphorylation of eIF2α was necessary to induce autophagy triggered by mTORC1 inhibition, and in contrast to other mTOR-regulated events, autophagy may be affected by rapamycin’s ability to affect the phosphorylation status of eIF2α.

GCN2 is activated by inhibition of mTORC1

We next sought to identify the kinase(s) responsible for the phosphorylation of eIF2α in response to mTORC1 inhibition. Although there are multiple eIF2α kinases, each responsive to specific cellular stresses, we first focused on GCN2 because inhibition of mTOR leads to the activation of GCN2 in yeast (12, 13). GCN2 contains a region homologous to the histidyl–transfer RNA (tRNA) synthetase, which binds to uncharged tRNAs during periods of amino acid deprivation and triggers a conformational change of GCN2 that leads to autophosphorylation at Thr889 and activation of the kinase (2527). Rapamycin treatment did not promote the phosphorylation of eIF2α in GCN2−/− MEFs (Fig. 3A). In contrast, eIF2α was phosphorylated in PERK−/− cells treated with rapamycin (fig. S1A). In addition, ER stress was not induced by rapamycin treatment, as demonstrated by a lack of XBP1 splicing (fig. S1B). Furthermore, rapamycin treatment induced the autophosphorylation of GCN2 at Thr889 to a similar extent as did leucine deprivation (as expected, tunicamycin treatment did not induce GCN2 autophosphorylation) (Fig. 3B). Thus, inhibition of mTORC1 activates GCN2, and GCN2 is responsible for the phosphorylation of eIF2α.

Fig. 3 mTORC1 inhibition results in the phosphorylation of eIF2α through PP6C-mediated dephosphorylation of GCN2.

(A) GCN2+/+ and GCN2−/− MEFs were treated with 300 nM rapamycin for the indicated times (hours) or with tunicamycin for 12 hours or deprived of amino acids for 12 hours. Cell lysates were immunoblotted for GFP. (B) HeLa cells were treated with 300 nM rapamycin for the indicated times, deprived of leucine for 2 hours, or treated with tunicamycin for 4 hours, and cell lysates were immunoblotted. (C and D) TSC+/+ and TSC−/− MEFs were incubated in leucine-deficient media (10.5 mg/liter) (C) or histidinol (D) for the indicated times, and cell lysates were immunoblotted. (E) HCT-116 and HeLa cells stably expressing a control retroviral vector or a vector expressing PP6C were assessed for GCN2 and/or eIF2α phosphorylation. (F) U2OS cells stably expressing either a control or shPP6C lentivirus were treated with 300 nM rapamycin for the times indicated (hours) or tunicamycin for 12 hours. Protein lysates were immunoblotted for phosphorylated eIF2α and other noted proteins. (G) HeLa cells stably expressing either a control or shIGBP1 lentivirus were treated and analyzed as in (F). (H) Scramble (con) and PP6C-depleted cells were treated with rapamycin for 6 hours, and phosphorylated and total GCN2 were assessed. (I and J) Control or GCN2 S551A/S551A cells were treated with rapamycin or tunicamycin, and eIF2α phosphorylation (I) and conversion of LC3I to LC3II (J) were assessed. In (A), (C), (D), (F), (G), and (H), representative blots and graphs showing the quantification from two biological replicates are displayed. In (B), (E), and (I), representative blots of two biological replicates are displayed.

We next determined the relative contribution of mTOR inhibition to the phosphorylation of eIF2α. The absence of TSC leads to the constitutive activation of mTOR that is not suppressed by amino acid deprivation (28). Thus, when TSC2−/− cells are deprived of amino acids, mTOR inhibition cannot contribute to the activation of GCN2 or the phosphorylation of eIF2α, as demonstrated by a lack of decrease in the phosphorylation of S6K1 (Fig. 3C). Phosphorylation of eIF2α under baseline conditions was generally similar in TSC2−/− and TSC2+/+ MEFs (Fig. 3, C and D). However, in response to low leucine concentrations, the time-dependent increase in the phosphorylation of eIF2α seen in TSC2+/+ MEFs was diminished in TSC2−/− MEFs (Fig. 3C). Similarly, treatment with the histidinyl-tRNA synthetase inhibitor histidinol, which induces His-tRNA deacylation to activate GCN2 and inhibit mTOR (29), led to decreased phosphorylation of eIF2α in TSC2−/− MEFs compared to identically treated TSC2+/+ cells (Fig. 3D). Together, these data suggest that mTOR inhibition increases the activation of GCN2 and the phosphorylation of eIF2α during amino acid deprivation.

mTORC1 inhibition leads to phosphorylation of eIF2α and induction of autophagy through activation of PP6C

mTOR inhibition leads to rapid dephosphorylation of its direct targets, including 4E-BP1 and S6K, whereas the phosphorylation of eIF2α after mTOR inhibition takes longer [2 to 6 hours, depending on the cell line (Figs. 1 to 3)], suggesting a mechanism with multiple intermediaries. In S. cerevisiae, TORC1 inhibition activates the phosphatase SIT4, which dephosphorylates Ser577 in GCN2, leading to threonine autophosphorylation and activation of the kinase at modest concentrations of uncharged tRNAs (12, 30). We therefore investigated whether the mammalian homolog of SIT4, PP6C, promotes the phosphorylation of eIF2α in response to mTORC1 inhibition.

The overexpression of PP6C induced the phosphorylation of eIF2α in HeLa cells and various colon cancer, cervical cancer, melanoma, glioblastoma, and osteosarcoma cell lines (Fig. 3E and fig. S2A). As expected, in cells overexpressing PP6C, rapamycin treatment did not lead to further increases in the phosphorylation of eIF2α, although the phosphorylation of eIF2α was increased in these cells in response to tunicamycin treatment and severe amino acid deprivation (fig. S2A). Conversely, rapamycin treatment no longer led to the phosphorylation of eIF2α in cells depleted of PP6C (Fig. 3F), but not in those depleted of PP2A (fig. S2B). PP6C depletion did not affect PERK-induced phosphorylation of eIF2α in tunicamycin-treated cells (Fig. 3F). The depletion of PP6C led to not only a decrease in eIF2α phosphorylation with mTORC1 inhibition but also a decrease in autophagy, as demonstrated by decreased conversion of LC3I to LC3II and generation of free GFP (Fig. 3F).

The activity of PP6C and other members of the PP2A family are inhibited by the protein encoded by immunogloublin (CD79A) binding protein 1alpha 4 (IGBP1/α4) (31, 32). In S. cerevisiae, TOR inhibition activates SIT4 by dissociating the phosphatase from the yeast homolog of IGBP1/α4, TAP42 (12, 32, 33). When PP6C activity was increased by depleting IGBP1/α4, basal phosphorylation of eIF2α was increased compared to control cells and was not further increased by rapamycin treatment (Fig. 3G). Furthermore, autophagy was increased in rapamycin-treated cells depleted of IGBP1/α4, as demonstrated by increased conversion of LC3I to LC3II and generation of free GFP (Fig. 3G). Conversely, IGBP1/α4 overexpression, which diminishes PP6C activity, blunted phosphorylation of eIF2α at baseline as well as in response to rapamycin treatment, but not in response to complete amino acid deprivation or tunicamycin treatment (Fig. 2C). In PP6C-depleted cells, rapamycin did not lead to a substantial increase in the phosphorylation of Thr899 in GCN2 (Fig. 3H), consistent with a role for GCN2 in regulating the phosphorylation of eIF2α in response to mTORC1 suppression (Fig. 3A). Together, these data suggest that PP6C activity is necessary for phosphorylation of eIF2α and autophagy induced by mTORC1 inhibition.

mTORC1 inhibition induces autophagy through dephosphorylation of Ser551 in GCN2

In S. cerevisiae, TORC1 inhibition leads to SIT4-mediated dephosphorylation of Ser557 in GCN2 (12). There is little peptide homology between yeast and mammalian GCN2 (12, 30). Thus, to determine the phosphorylation status of mammalian GCN2 and how this changes with mTOR activity, we performed mass spectrometry (MS) on Flag-tagged GCN2 immunoprecipitated from 293T cells treated with either rapamycin or vehicle. GCN2 was phosphorylated at either Ser258 or Ser261 or Ser2544 (could not be resolved), Ser230, Ser551, and either Ser688 or Ser695 (could not be resolved) (fig. S3). However, the only site that became dephosphorylated with rapamycin treatment was Ser551. We then used CRISPR technology to mutate both GCN2 alleles so that they encoded an S551A mutant. GCN2 S551A cells had high basal eIF2α phosphorylation that was not responsive to rapamycin (Fig. 3I) and high basal autophagy with a blunted response to rapamycin (Fig. 3J). Thus, dephosphorylation of Ser551 in GCN2 plays a critical role in mTOR-regulated phosphorylation of eIF2α and autophagy.

PP6C, PP6Rs, and GCN2 form a complex that is necessary for the phosphorylation of eIF2α

The PP6C regulatory subunits PP6R1, PP6R2, and PP6R3 bridge PP6C to its substrates. Some substrates only bind to a specific subunit, whereas other substrates are more promiscuous (16, 34). We found that endogenous PP6R1, PP6R2, and PP6R2 immunoprecipitated both endogenous GCN2 and endogenous PP6C, suggesting that all three regulatory subunits can participate in a GCN2-PP6C complex (Fig. 4A). Depletion of PP6R1, PP6R2, or PP6R3 led to a partial decrease in the basal phosphorylation of eIF2α that correlated with the efficiency of knockdown, confirming the functional overlap between these regulatory subunits (Fig. 4B). The co-depletion of all three subunits led to a decrease in basal (Fig. 4C) and rapamycin-induced (Fig. 4D) phosphorylation of eIF2α. These observations confirm the role of PP6C in promoting the phosphorylation of eIF2α and suggest that a PP6C-GCN2 complex may be responsible.

Fig. 4 PP6C, PP6Rs, and GCN2 form a complex that is necessary for eIF2α phosphorylation, and PP6C mutants found in melanoma do not bind to PP6Rs.

(A) HeLa cell lysates were immunoprecipitated with protein G beads conjugated to antibodies targeting GFP, PP6R1, PP6R2, or PP6R3. Whole-cell extract input (WCE) and immunoprecipitated samples were immunoblotted for endogenous PP6Rs, GCN2, and PP6C. (B) Lysates from U2OS cells stably expressing Flag-tagged PP6R1, PP6R2, or PP6R3 and stably expressing a scramble (SCR) or corresponding shRNA (short hairpin RNA) were immunoblotted. (C) Lysates from U2OS cells stably expressing either a control or three shRNAs targeting PP6R1, PP6R2, and PP6R3 were immunoblotted for PP6C, phosphorylated eIF2α, and other noted proteins. (D) U2OS cells stably expressing either a control or three shRNAs targeting PP6R1, PP6R2, and PP6R3 were treated with 300 nM rapamycin for the times indicated, and cell lysates were immunoblotted. (E) 293T cells were cotransfected with vectors expressing a Myc-tagged PP6C mutant and Flag-PP6R1, Flag-PP6R2, or Flag-PP6R3. Lysates were immunoprecipitated (IP) with Sepharose beads conjugated to an anti-Flag antibody, and lysates were immunoblotted. WT, wild type. (F) Lysates from HCT-116 cells stably expressing a Myc-tagged WT or mutant PP6C were immunoblotted. (G) U2OS cells stably expressing a Myc-tagged PP6C expression retrovirus were treated with cycloheximide (CHX; 100 μg/ml) for the times indicated. Cell lysates were immunoblotted for PP6C. The graphs shows the abundance of exogenously expressed PP6C graphed as a function of time from two biological replicates. (H) Dot plot of the half-life of five PP6C mutants that bind to the PP6Rs compared to four PP6C mutants that do not bind to regulatory subunits. (I) Half-lives were calculated for each of eight PP6C mutant and correlated with its ability to bind to the PP6Rs. The Pearson correlation coefficient (R2) is displayed. (J) Lysates from U2OS cells stably expressing a Myc-tagged PP6C mutant or D84N catalytic mutant and a control or shPP6C (targeting the 3′UTR) lentivirus were immunoblotted for PP6C and phosphorylated eIF2α. (K) Dot plot of the half-life of endogenous PP6C in U2OS cells expressing one of four stable or three unstable PP6C mutants. In (C) and (D), representative blots and graphs showing quantification of two biological replicates are displayed. In (A), (B), (E), (F), and (J), blots representative of two biological replicates are displayed.

PP6C mutations found in melanoma can decrease PP6C-PP6R binding and PP6C stability

Unbiased searches for melanoma driver mutations by two independent groups have identified PP6C mutations in about 10% of all melanomas (17, 18). Up to 60% of the mutations found in PP6C cluster within a highly conserved region that encodes the predicted interface for PP6Rs (16, 17, 35). Because PP6C binding to PP6R was necessary to promote the phosphorylation of eIF2α, we further studied several of these mutants. We identified three melanoma-associated PP6C mutants with diminished capacity to bind to PP6R1, PP6R2, and PP6R3 (Fig. 4E and fig. S4A). The most frequent mutation, R264C, did not alter PP6C binding to regulatory subunits. As expected, expression of these PP6C mutants did not interfere with the ability of PP6R to bind to GCN2.

When stably expressed in several cell lines, the abundance of PP6C mutants that were unable to bind to PP6Rs was lower than that of wild-type PP6C or PP6C mutants that retained the ability to bind to PP6Rs (Fig. 4F and fig. S4B). In addition, we noted decreased PP6C abundance in cells depleted of PP6Rs (Fig. 4C), leading us to theorize that PP6C-PP6R interactions are required for PP6C stability. To test this hypothesis, we assessed the half-life of endogenous PP6C protein and ectopically expressed wild-type and mutated PP6C in transfected human embryonic kidney–293 cells, where similarly high expression of all constructs could be achieved. The half-lives of endogenous PP6C, overexpressed wild-type PP6C, and PP6C mutants that bound to PP6Rs averaged 2 to 4 hours, and were greater than the half-lives of those PP6C mutants that do not bind to PP6Rs, which all averaged about 0.8 hour (Fig. 4, G and H, and fig. S4C). The ability of PP6C constructs to bind to PP6Rs strongly correlated to their half-lives (Fig. 4I), suggesting that PP6C mutants that do not bind to regulatory subunits are destabilized.

Nonstable PP6C mutants found in melanoma increase the stability of wild-type PP6C and induce the phosphorylation of eIF2α

We predicted that expression of PP6C mutants that do not bind to PP6R would not promote, or might even blunt, the phosphorylation of eIF2α. Unexpectedly, however, we noted increased phosphorylation of eIF2α in cells expressing mutated PP6C constructs that did not bind to PP6R (Fig. 4J) (36). These seemingly contradictory findings were clarified when we observed that the protein abundance of endogenous PP6C was decreased by the expression of wild-type PP6C and mutants that bound to PP6R and was increased by that of PP6C mutants that did not bind to PP6Rs (Fig. 4, F and G). This increase in the abundance of endogenous PP6C was associated with an increase in endogenous PP6C protein half-life from 3 to 7 hours with expression of a PP6C mutant that does not bind PP6Rs, but not with that of a PP6C mutant that binds to PP6Rs (Fig. 4K). When we overexpressed PP6C mutants and at the same time depleted endogenous PP6C [with a shRNA directed against the 3′ untranslated region (3′UTR) of endogenous PP6C not contained in our constructs], the phosphorylation of eIF2α phosphorylation was diminished (Fig. 4J). Thus, PP6C mutants that do not bind to PP6Rs and are destabilized promote the stability of endogenous wild-type PP6C and lead to the phosphorylation of eIF2α.

Nonstable PP6C mutants found in melanoma induce autophagy in cells and are associated with increased autophagy in melanoma

We next examined how autophagy was affected by unstable PP6C mutants, which promote the phosphorylation of eIF2α although they cannot bind to regulatory subunits. First, we generated isogenic cell lines expressing an LC3-GFP fusion protein alone or with wild-type PP6C or PP6C mutants that do not bind regulatory subunits. Compared to empty vector, the overexpression of wild-type PP6C led to an increase in the basal conversion of LC3I to LC3II and in the generation of free GFP, even in the presence of chloroquine. Cells expressing PP6C mutants unable to bind regulatory subunits also demonstrated increased basal autophagy that, as expected, was minimally inhibited by rapamycin treatment (Fig. 5A).

Fig. 5 Autophagy is increased in cells expressing forms of PP6C with mutations that disrupt regulatory subunit binding.

(A) GFP-LC3 HCT116 cells expressing empty vector, WT PP6C, or mutated PP6C were treated with vehicle, 100 nM rapamycin for 6 hours, 60 μM chloroquine for 2 hours, or both. Immunoblots for GFP and tubulin were performed. Representative blots are displayed from two biological replicates. (B) Primary cell cultures of WT and PP6C-mutated melanoma tumors were treated as in (A) and immunoblotted for LC3 and p62. Representative blots from two biological replicates are shown. (C) Melanoma tumors were genotyped for PP6C and stained for LC3 and blindly evaluated for percentage of cells with LC3 foci. LC3 and phosphorylated eIF2α expression were assessed by immunofluorescence and immunohistochemistry, respectively, and expression was graded as 0 (<25% cells with expression), 1 (25 to 50% cells with expression), 2 (50 to 75% of cells with expression), and 3 (>75% cells with expression). Representative images, including both melanoma (M) and normal skin (N) with increasing phosphorylated eIF2α and LC3 foci, are displayed (scale bar, 150 μm). LC3 foci from tumors with WT PP6C (n = 12), harboring PP6C mutants that bind to regulatory subunits (n = 6) or PP6C mutants that do not bind to regulatory subunits (n = 6), are shown (scale bar, 40 μm). P values determined from Wilcoxon rank sum test. (D) Proposed model of mTORC1 regulation of eIF2α phosphorylation through PP6C-mediated activation of GCN2. Left: During periods of nutrient deprivation, mTORC1 inhibition activates PP6C, which associates with and dephosphorylates GCN2 in a complex with a PP6 regulatory protein. Dephosphorylation of GCN2 promotes its activation. Activated GCN2 then phosphorylates eIF2α, leading to the induction of autophagy. Right: Dissociation of PP6C from the PP6 regulatory proteins decreases its stability. Several PP6C mutants found in melanoma do not bind to the PP6Rs and are rapidly degraded, increasing the stability of WT PP6C and enhancing the induction of autophagy.

We then assessed autophagy in primary short-term cell cultures derived from tumors with wild-type PP6C and from tumors that harbor PP6C mutants that do not bind regulatory subunits. These cells are not isogenic and carry multiple mutations, any of which may differentially affect autophagy (17, 18). As expected, basal autophagy and autophagic flux greatly varied among these cell lines, as indicated by differing baseline LC3I/LC3II ratios and responses to the autophagy inhibition with chloroquine (Fig. 5B). However, autophagy in primary cells containing wild-type PP6C, but not in those containing PP6C mutants, was inhibited by rapamycin treatment (as indicated by reduced p62 abundance and increases in LC3I/LC3II ratios) (Fig. 5B), consistent with a model in which rapamycin-induced autophagy is at least partially dependent on PP6C.

Although autophagy in melanoma could be altered by mutations that affect not only the autophagy machinery but also vascularization and other aspects of the tumor microenvironment, we explored autophagy in paraffin-embedded melanoma samples genotyped for PP6C (37). The number of cells with LC3 foci was assessed in 6 tumors with PP6C mutants that did not bind to PP6C regulatory subunits, 6 tumors with PP6C mutants that bound to regulatory subunits, and 12 tumors expressing wild-type PP6C. Those tumors with PP6C mutants that do not bind to regulatory subunits had a significantly greater number of LC3 foci compared to those tumors with mutant PP6Cs that bind to regulatory subunits (Fig. 5C, bottom left). The phosphorylation of eIF2α was higher in those tumors with PP6C mutants that do not bind to regulatory subunits, although this increase was not statistically significant (Fig. 5C, bottom right). Together, these three complementary approaches suggest that PP6C mutants that do not bind regulatory subunits are associated with increased autophagy.

DISCUSSION

We determined that two amino acid–sensing systems were linked in mammalian cells, and specifically, that mTORC1 inhibition led to the phosphorylation of eIF2α through activation of the eIF2α kinase GCN2. Our data demonstrated that PP6C regulated the activation of GCN2 in response to mTORC1 inhibition and increased the phosphorylation of eIF2α in response to moderate amino acid deprivation (Fig. 5D). This system thus appears to be homologous to the system in yeast (12, 13). In addition, we showed that the activation of autophagy by mTORC1 inhibition depended on the crosstalk between these two pathways. Because the regulation of mTOR activity and eIF2α phosphorylation are involved in the cellular stress response, the link between these events provides insight into cellular adaptation in cancer and various other physiological and pathological conditions.

The induction of autophagy by mTOR inhibition may partially explain some of the failures of mTORC1 inhibitors as anticancer agents (38, 39). Alternative strategies to inhibit mTOR without inducing autophagy include the dual inhibition of mTORC1 and GCN2 or interference with GCN2 dephosphorylation by PP6C. Because GCN2 and PP6C interact, GCN2 is likely a direct PP6C substrate, and the identification of Ser551 as a key phosphorylation site that promotes the activation of GCN2 is a preliminary step in characterizing PP6C-PP6R-GCN2 interactions that may provide a molecular target for autophagy manipulation.

We found that forms of PP6C with mutations that disrupt PP6C-PP6R interactions are rapidly degraded, which paradoxically stabilizes wild-type PP6C. It is possible that some PP6C mutants in melanoma may have an increasing effect on the amount of wild-type PP6C generated from the nonmutated allele. Although most PP6C mutations reported in short-term cultures from melanomas were predicted to be loss-of-function mutations (17, 18), in direct sequencing of PP6C from more than 390 melanomas, we have found that many PP6C mutations are not associated with a loss of heterozygosity, and both PP6C-mutated and nonmutated mRNAs are found in single-cell clones derived from PP6C-mutated melanoma (37). PP6C is overexpressed in glioblastoma multiforme (40), supporting the potential for some PP6C mutations to serve as activating mutations. Mutated oncogenes have previously been shown to affect nonmutated alleles, such as the increased activation of wild-type Ras by oncogenic Ras (41).

Whereas many of the PP6C mutations found in melanoma cluster within a highly conserved region where PP6C interfaces with PP6Rs, mutations are found throughout the PP6C gene, and we found that several mutations, including the common R264C mutation, did not disrupt binding. This raises the possibility that distinct mutations in PP6C may have separate biochemical, molecular, and phenotypic implications, as has been noted for mutations in PTPN11, which encodes the phosphatase Shp2 (42). Mutations associated with Noonan syndrome are distributed throughout the coding region of PTPN11 and result in forms of Shp2 with increased or unregulated activity due to an inability of the phosphatase to maintain its autoinhibited conformation. In contrast, in Leopard syndrome, mutations occur in the catalytic core of the enzyme and produce catalytically impaired Shp2 variants. Both Noonan syndrome and Leopard syndrome are phenotypically similar, and it is unclear how mutations in the same gene that result in biochemically opposite characteristics result in similar phenotypes.

In addition to autophagy, PP6C has roles in the degradation of IκBε in response to TNFα, dephosphorylation of γ-H2AX after irradiation, and dephosphorylation of Aurora kinase A to regulate mitosis (34, 4346). In contrast to the gain-of-function activity shown by unstable PP6C mutants in the autophagy pathway, we and others have demonstrated that PP6C mutants that do not bind to regulatory subunits cannot dephosphorylate Aurora kinase A (37, 47). Autophagy appears to play a dual role in cancer in general and may function both as a tumor suppressor and as a promoter (4850). Further studies are required to better define the clinical, biological, and potentially therapeutic impact of PP6C mutations, GCN2 activation, and eIF2α phosphorylation on autophagy and/or melanoma initiation and progression.

MATERIALS AND METHODS

Cell culture

Cells were cultured as previously described (51) and treated with tunicamycin (2.5 μg/ml), 100 to 300 nM rapamycin, 60 μM chloroquine, or 10 μM histidinol, or depleted of amino acids by treating cells with low-leucine medium (10.5 mg/liter) supplemented with 10% dialyzed fetal bovine serum. MEFs deficient in PERK, GCN2, or TSC2 and eIF2α S51A/S51A MEFs and their wild-type controls have been previously described (19, 20, 5254). For the protein stability experiments, cells were treated with cycloheximide (100 μg/ml). Short-term primary cultures from PP6C-mutated melanoma have been previously described (18) and obtained from the Yale Dermatology Cell Culture Facility.

Plasmids

The pLKO.1 shPP6C (TRC0000002764), shIGBP1 (TRC0000039966), shPP6R1 (TRC0000129845), shPP6R2 (TRC0000144656), and shPP6R3 (TRC0000159679) lentiviral vectors were obtained from Sigma. The gadd34 (haA1.pBabepu) and phosphatase-dead control (Myd116.PFLAG.CMV2) (21) vectors were obtained from Addgene. The pLKO.1 shRictor and shRaptor lentiviral vectors were a gift from R. Schneider. The GFP-LC3 fusion, a gift from G. Kroemer, was cloned into the retroviral vector pQCXIN or pBabe. PP6R1, PP6R2, and PP6R3 were cloned into pCDNA. IGBP1/α4 was cloned into pLPC. PP6C was cloned into pQCXIN, pBABE-puro, and pLPC. Retroviruses were generated using standard retroviral generation and infection techniques (51, 55). 293T cells were transiently transfected by calcium phosphate precipitation.

CRISPR knock-in

The GCN2 S551A mutant was generated by cotransfecting the guide sequence CCAGCCAAAAATGCCTCTAG cloned into p458X (Addgene) along with a 200-nucleotide single-stranded oligonucleotide containing the sequence CTCTAGTaGAACAAgcTCCTGAAG, where the lowercase letters indicate mutations introduced to convert serine to alanine and to mutate the PAM sequence. Homozygous mutants were screened by PCR with mutation-specific primers, and sequences were confirmed by Sanger sequencing.

Immunoblots and immunoprecipitations

Immunoblots were done using fluorescent secondary antibodies and quantitated by LI-COR as previously described (55). Membranes were stained with antibodies directed against GFP (Cell Signaling, 2995), tubulin (Sigma, T9026), phosphorylated eIF2α (Epitomics, 1090), eIF2α (Santa Cruz Biotechnology, SC-11386), ATF4 (Santa Cruz Biotechnology, SC-200), S6 kinase (Cell Signaling, 9202), phosphorylated S6 kinase (Cell Signaling, 9234), rictor (Bethyl, A300-459A), raptor (Millipore, 09–217), Akt (Cell Signaling, 9272), phosphorylated Akt (Cell Signaling, 9271), 4E-BP1 (Cell Signaling, 9452), phosphorylated 4E-BP1 (Cell Signaling, 9456), GCN2 (Epitomics, 5417), phosphorylated GCN2 (Epitomics, 2425), PP2A (gift from C. Basilico), PP6C (Millipore, 07–1224), IGBP1 (Upstate, 05–930), PP6R1 (Bethyl, A300-968A), PP6R2 (Bethyl, A300-970A), PP6R3 (Bethyl, A300-972A), Flag tag (Sigma, F3165), hemagglutinin tag (Santa Cruz Biotechnology, SC-7392), and Myc tag (Cell Signaling, 2276). Blots were then washed in TBST (tris-buffered saline with Tween 20) and incubated in LI-COR IRDye-conjugated goat anti-rabbit (800 nm) or goat anti-mouse (680 nm) antibodies and imaged on the LI-COR Odyssey Infrared Imaging System. Immunoprecipitations were performed as previously described (56). Whole-cell extract (500 to 1000 μg) was mixed with protein G–Sepharose (GE Healthcare) as well as with immunoprecipitating antibody PP6R1 (Bethyl, A300-968A), PP6R2 (Bethyl, A300-970A), or PP6R3 (Bethyl, A300-972A). For the Flag tag immunoprecipitations, lysates were mixed with the Anti-Flag M2 affinity gel (Sigma, A2220). Samples were then incubated at 4°C with constant rotation overnight. Beads were then washed four times with lysis buffer and then resuspended in 4× Laemmli buffer and boiled for 10 min at 95°C.

Microscopy and assessment of eIF2α phosphorylation and LC3 foci

Cells were fixed with 3.7% paraformaldehyde for 20 min, stained with LC3 antibody (Novus, NB600-1384) and subsequently with a fluorescein isothiocyanate–conjugated secondary antibody (Molecular Probes, F2765). Both immunofluorescently labeled LC3 and GFP foci were visualized with confocal microscopy as previously described (51). GFP-LC3–expressing cells containing three or more foci were counted as positive and quantified as a proportion of all cells within a given field. Endogenous LC3 foci were quantified using the ImageJ software. Image intensity thresholds were set to a constant value to include particles with a high signal compared to background. Regions of interest were drawn around each cell in a field, and the number of foci was counted using the “analyze particles” function. Paraffin-embedded melanoma samples were obtained from the Institutional Review Board–approved NYU Interdisciplinary Melanoma Cooperative Group tumor bank. eIF2α immunohistochemistry and indirect immunofluorescence were performed as previously described (7, 55).

Mass spectrometry

293 cells were transfected with GCN2 and then treated with either rapamycin or vehicle for 4 hours. Cell lysates were then immunoprecipitated with an anti-GCN2 antibody as above and run on a polyacrylamide gel. The excised GCN2 gel bands were reduced with the addition of 0.2 M dithiothreitol at pH 8 for 1 hour at 57°C and subsequently alkylated using 0.5 M iodoacetic acid at pH 8 for 45 min in the dark at room temperature with gentle shaking. The samples were proteolytically digested with trypsin (Promega) at a 1:50 enzyme/substrate ratio overnight with gentle shaking at room temperature. The resulting peptide mixture was extracted from the gel pieces and desalted using a C18 Stage tip procedure, as previously described (57). The desalted peptide mixture was concentrated in a SpeedVac concentrator to remove organic solvents. A third of the peptide mixture was loaded onto an Acclaim PepMap 100 precolumn (75 μm × 2 cm, C18, 3 μm, 100 Å; Thermo Scientific) that was connected to an EASY-Spray PepMap RSLC column (75 μm × 25 cm, C18, 2 μm, 100 Å; Thermo Scientific) with a 5-μm emitter using the autosampler of an EASY-nLC 1000 instrument (Thermo Scientific). Peptides were gradient-eluted from the column directly into a Q Exactive mass spectrometer (Thermo Scientific) using a 120-min gradient from 2% solvent B to 40% solvent B. Solvent A was 2% acetonitrile in 0.5% acetic acid, and solvent B was 90% acetonitrile in 0.5% acetic acid. High-resolution MS1 spectra were acquired with a resolution of 70,000, an AGC target of 1e6 with a maximum ion time of 120 ms, and a scan range of 300 to 1500 m/z (mass/charge ratio). After each MS1, 15 data-dependent high-resolution HCD MS2 spectra were acquired. In addition, high-resolution MS2 spectra were collected of the +2 and +3 form of the MPLVEQSPEDSpEGQDYVETVIPSNR peptide in a targeted manner using an inclusion list. All MS2 spectra were collected using the following instrument parameters: resolution of 17,500, AGC target of 2e5, maximum ion time of 250 ms, one microscan, 2 m/z isolation window, fixed first mass of 150 m/z, 30-s exclusion list, normalized collision energy (NCE) of 27 for data-dependent MS2, and NCE of 24 for targeted MS2.

SUPPLEMENTARY MATERIALS

www.sciencesignaling.org/cgi/content/full/8/367/ra27/DC1

Fig. S1. mTORC1 inhibition does not lead to phosphorylation of eIF2α through the unfolded protein response.

Fig. S2. PP6C and IGBP1 overexpression regulate rapamycin-induced eIF2α phosphorylation.

Fig. S3. Analysis of GCN2 phosphorylation status by MS.

Fig. S4. PP6C binding to regulatory subunits affects PP6C stability.

REFERENCES AND NOTES

Acknowledgments: We gratefully acknowledge the gift of reagents from D. Ron, R. Kaufman, M. Sahin, D. Kwiatkowski, C. Koumenis, C. Thompson, G. Kroemer, and R. Schneider. MS was expertly performed by B. Ueberheide and J. Chapman-Lim of the NYU Proteomics Resource Center Core. Funding: This work was supported by T32GM066704 (J.W.), RO1DK081641 (L.B.G.), funds from the Lederman/Levin and Kwiat families, and the NYU Perlmutter Cancer Institute. The Proteomics Core is supported in part by P30CA016087 and UL1 TR00038. Author contributions: J.W. designed and performed most of the experiments, except where indicated. D.W. replicated the experiments in Fig. 1 and performed the experiments in Fig. 3 (B to D, I, and J) and Fig. 5 (A and B). H.Z. provided statistical review for the studies. S.W. and I.O. obtained the human melanoma samples. L.B.G. designed the study, supervised the data analysis, and wrote the paper. Competing interests: The authors declare that they have no competing interests.
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