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. Author manuscript; available in PMC: 2014 Jan 10.
Published in final edited form as: Mol Cell. 2012 Nov 8;49(1):172–185. doi: 10.1016/j.molcel.2012.10.003

Metabolic stress controls mTORC1 lysosomal localization and dimerization by regulating the TTT-RUVBL1/2 complex

Sang Gyun Kim 1, Gregory R Hoffman 1,6, George Poulogiannis 2, Gwen R Buel 1, Young Jin Jang 1,3, Ki Won Lee 3, Bo-Yeon Kim 4, Raymond L Erikson 5, Lewis C Cantley 2, Andrew Y Choo 1,*, John Blenis 1,*
PMCID: PMC3545014  NIHMSID: NIHMS414835  PMID: 23142078

Summary

The metabolism of glucose and glutamine, primary carbon-sources utilized by mitochondria to generate energy and macromolecules for cell growth, is directly regulated by mTORC1. We show that glucose and glutamine, by supplying carbons to the TCA cycle to produce ATP, positively feed back to mTORC1 through an AMPK-, TSC1/2-, and Rag-independent mechanism by regulating mTORC1 assembly and its lysosomal localization. We discovered that ATP-dependent TTT-RUVBL1/2 complex was disassembled and repressed by energy depletion, resulting in its decreased interaction with mTOR. The TTT-RUVBL complex was necessary for the interaction between mTORC1 and Rag, and formation of mTORC1 obligate dimers. In cancer tissues, TTT-RUVBL complex mRNAs were elevated, and positively correlated with transcripts encoding proteins of anabolic metabolism and mitochondrial function - all mTORC1-regulated processes. Thus, the TTT-RUVBL1/2 complex responds to the cell’s metabolic state, directly regulating the functional assembly of mTORC1, and indirectly controlling the nutrient signal from Rags to mTORC1.

Introduction

An essential requirement of cell growth is the availability of sufficient nutrient sources to meet biological demand. These nutrients include glucose and glutamine, which are two primary carbon sources for proliferating cells to supply substrates to the TCA cycle, to produce the fundamental building blocks for macromolecules production, and to generate sufficient ATP to drive energy-requiring cellular processes (Vander Heiden et al., 2009). At the epicenter of cellular metabolism is the mechanistic target of rapamycin complex 1 (mTORC1), whose activity is deregulated in many cancers and drives cell growth and nutrient uptake (Duvel et al., 2010; Edinger and Thompson, 2002; Fingar et al., 2004; Sipula et al., 2006). mTORC1 consists of mTOR, Raptor, and mLST8, and when active, promotes anabolic processes and suppresses catabolic processes to maximize cell growth and proliferation (Yecies and Manning, 2011). Considering the prevalence of mTORC1 activation in cancer and the requirement of nutrients for cell growth, understanding how mTORC1 activation coordinates nutrient levels and anabolic/catabolic processes is critical.

To ensure that anabolic signals are coupled with sufficient levels of nutrients, energy, and other resources required for growth, mTORC1 senses the cell’s environmental conditions (Fingar and Blenis, 2004; Zoncu et al., 2011). Deprivation of amino acids (AA), oxygen, mitogens, or energy – all of which are required for cell growth – inhibits mTORC1 activity, and in most cases, this inhibition is necessary to maintain cellular survival by balancing the supply and demand of the resource (Choo et al., 2010). The molecular details of these environment-sensing processes are poorly understood, although AA appear to activate the Rag GTPases to regulate mTORC1 localization to the lysosomes (Kim et al., 2008; Sancak et al., 2008); oxygen levels control HIF-1α and REDD1 expression to activate TSC2 (Brugarolas et al., 2004; Cam et al., 2010); mitogens activate kinases such as AKTs, RSKs, and MAPKs which phosphorylate and inhibit TSC2 (Inoki et al., 2002; Manning et al., 2002; Roux et al., 2004); and metabolic stress activates AMPK to phosphorylate TSC2 and Raptor, leading to increased TSC2 function and decreased Raptor function (Gwinn et al., 2008; Inoki et al., 2003). Thus, when any one of these essential resources is limited, mTORC1 activity and cell growth are decreased.

In proliferating cells, glucose and glutamine supply carbons to the TCA cycle (DeBerardinis et al., 2008). Glucose is metabolized through glycolysis, and the derived carbons could be secreted as lactate or be shunted to the pentose phosphate or hexosamine pathway (Vander Heiden et al., 2009). The glycolytic product, pyruvate, could also enter the TCA cycle to replenish intermediates that are used for macromolecule synthesis. In glutaminolysis, glutamine is first converted to glutamate and then to α-ketoglutarate, which replenishes the cycle, ultimately engaging in macromolecule synthesis or redox reactions (Wise and Thompson, 2010).

Considering the absolute importance of the TCA cycle in macromolecule synthesis and mitochondria respiration and mTORC1’s central role in cell growth, we investigated if and how the depletion of glucose and glutamine, the primary nutrients responsible for replenishing the TCA cycle, affected mTORC1 signaling. We discovered that glucose and glutamine depletion strongly inhibited mTORC1 in a TSC1/2-, AMPK-, and Rag-independent mechanism, but this inhibition was largely through energetic stress. Energetic stress prevented mTORC1 localization to the lysosomes, but this effect could not be recovered with expression of dominant-active mutant of RagB. By screening the mouse genome with an siRNA library, we uncovered that energetic stress disassembled and inhibited the Tel2-Tti1-Tti2 (TTT)-RUVBL1/2 complex, which is an AAA+ ATPase-containing complex that has been shown to regulate the assembly and stability of phosphoinositide-3-kinase-related protein kinases (PIKK) family proteins and to be necessary for various cellular processes, including the DNA damage response, nonsense-mediated decay (NMD), and telomerase assembly (Takai et al., 2007; Takai et al., 2010; Horejsi et al., 2010; Hurov et al., 2010; Izumi et al., 2010; Venteicher et al., 2008). Energetic stress decreased the interaction between mTOR and the TTT complex component Tel2 as well as the interaction between TTT components and RUVBL proteins. Importantly, the disassembly appeared to involve the inhibition of the ATPase activity of RUVBL1/2. Moreover, the inhibition of TTT-RUVBL1/2 disrupted the lysosomal localization of mTORC1 and inhibited the homodimerization of mTORC1 – a process that is necessary for phosphorylation of substrates (Yip et al., 2010). Finally, expression of the TTT-RUVBL1/2 complex was concertedly upregulated in several tumors and closely associated with mTORC1 signaling.

Results

Glucose and glutamine are essential, but redundant, regulators of mTORC1 activation through a TSC- and AMPK-independent mechanism

To investigate if the mTORC1 pathway senses the levels of glucose, glutamine, and/or TCA intermediates, we deprived Tsc2-/- mouse embryonic fibroblasts (MEFs) of glucose and/or glutamine for 12 hrs. Deprivation of either glucose or glutamine did not significantly inhibit mTORC1 in Tsc2-/- cells as measured by the phosphorylations of S6K1 and 4E-BP1 and the mobility shift of their total bands (Figure 1A). However, deprivation of both glucose and glutamine potently inhibited mTORC1 in Tsc2-/- and Tsc1-/- cells (Figure 1A and 1B). Consistent with mTORC1 inhibition, AMPK activation and Raptor phosphorylation were the most evident following the deprivation of both glucose and glutamine (Figure 1C); deprivation of other nutrients, such as branched chain amino acids (BCAA), failed to synergize with glucose deprivation to inhibit mTORC1 in Tsc-/- cells (Figure 1B). Therefore, glucose and glutamine, together, are essential regulators of mTORC1 activation, and their deprivation inhibits mTORC1 independently of TSC.

Figure 1. Glucose and Glutamine are essential, but redundant, regulators of mTORC1 activation through a TSC- and AMPK-independent mechanism.

Figure 1

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(A) Tsc2-/- MEFs were deprived of glucose, glutamine, or glucose and glutamine for 12 hrs. mTORC1 and AMPKα activities were then monitored by immunoblotting.

(B) Tsc2-/- and Tsc1-/- MEFs were deprived of the indicated nutrients for 12 hrs.

(C) The kinetics of the inhibitory effect of glucose/glutamine deprivation on mTORC1.

(D) Tsc2-/- MEFs were treated with AMPK inhibitor for 1 hr prior to cell lysis at the indicated concentrations following 12 hrs of glucose/glutamine deprivation.

(E) Tsc2-/- MEFs, which have the endogenous Raptor knocked down and have either the wild type (WT) or S722/792AA Raptor expressed, were deprived of the indicated nutrients and analyzed as in (A).

(F) Ampkα1,2+/+ and Ampkα1,2-/- MEFs deprived of the indicated nutrients for 10 hrs were analyzed as in (A).

We then investigated if AMPK activation or Raptor phosphorylation was causative or correlative with mTORC1 inhibition. Previous work showed that AMPK and LKB1 are necessary for the AMP analog AICAR to phosphorylate and inhibit Raptor (Gwinn et al., 2008). We utilized compound C, an ATP-competitive inhibitor of AMPK (Zhou et al., 2001), to determine if AMPK activation was required for mTORC1 inhibition by glucose/glutamine withdrawal. Treatment of compound C for 1 hr prior to lysis effectively inhibited Raptor phosphorylation dose dependently but did not recover mTORC1 inhibition (Figure 1D). To determine if the phosphorylation of Raptor was important, we utilized Tsc2-/- cells with endogenous Raptor depleted and the wild-type (WT) or S722/792A Raptor expressed. As shown in Figure 1E, glucose/glutamine deprivation led to robust phosphorylation of Raptor in WT Raptor-expressing cells, but not in the mutant Raptor-expressing cells. However, mTORC1 was severely attenuated in both cell types; thus Raptor phosphorylation is not required for glucose/glutamine deprivation to inhibit mTORC1.

Since AMPK could phosphorylate other currently unidentified substrate-targets to inhibit mTORC1, we utilized Ampkα and α2 deficient MEFs. The α subunit is the catalytic subunit and is required for AMPK activity (Viollet et al., 2009). As shown in Figure 1F, glucose/glutamine deprivation failed to increase Raptor phosphorylation in Ampkα-/- cells, but not in Ampkα+/+ cells. Surprisingly, glucose/glutamine deprivation potently inhibited mTORC1 in both Ampkα+/+ and Ampkα-/- cells, suggesting that although Raptor phosphorylation and AMPK activation are sufficient for mTORC1 inhibition (Gwinn et al., 2008; Inoki et al., 2003), neither is required for glucose/glutamine deprivation-induced mTORC1 inhibition.

Metabolism of glucose and glutamine and entry into the TCA cycle are required for mTORC1 activation

We next sought to determine if the metabolism of glucose and glutamine was involved in mTORC1 regulation. To ask if glutamate, which is produced from glutamine, could compensate for glutamine deprivation, glucose/glutamine-deprived Tsc2-/- cells were treated with glutamate. The treatment strongly recovered mTORC1 inhibition from glucose/glutamine deprivation (Figure S1A). Conversely, treatment with Epigallocatechin gallate (EGCG, an inhibitor of GDH), but not aminooxyacetate (AOA, an inhibitor of transaminases), inhibited mTORC1 only in glucose-deprived conditions (Figure S1B), suggesting that the metabolism of glutamine to α-ketoglutarate through GDH is required for mTORC1 activation. To evaluate glucose metabolism’s role, we added the glycolysis product pyruvate to the media. Both sodium pyruvate (SP) and methyl pyruvate (MP) sufficiently recovered the inhibition of mTORC1 via glucose/glutamine deprivation (Figure S1C).

Because the metabolism of both glucose and glutamine is critical for mTORC1 activation, we assessed if the replenishment of the TCA by these nutrients was involved in mTORC1 regulation. Treatment of Tsc2-/- cells with TCA intermediates succinate or oxaloacetate (OAA) significantly restored mTORC1 activity in the absence of glucose/glutamine (Figure S1D) – an effect also observed in Ampkα+/+ and Ampkα-/- MEFs (data not shown). Therefore, glucose and glutamine regulation of mTORC1 involves the replenishment of the TCA cycle.

Glucose/glutamine deprivation inhibits mTORC1 independently of Rags

Considering AA activate mTORC1 through activation of the Rag GTPases and recruitment of mTORC1 to lysosomal sites (Sancak et al., 2008), and various types of AA (including the BCAA) can provide carbons for the TCA cycle (Hutson et al., 2005), we asked if glucose/glutamine deprivation could also inhibit the lysosomal localization of mTORC1. Cells stimulated with AA exhibited mTOR localization to the lysosome, while those deprived of AA exhibited cytoplasmic localization (Sancak et al., 2008) (Figure 2A). Interestingly, glucose/glutamine deprivation also prevented mTOR localization to the lysosomes in various cell types including Tsc2+/+ and Tsc2-/- cells (Figures 2A, S2, and data not shown), but, unlike mTORC1 inhibition from AA deprivation, inhibition from glucose/glutamine deprivation was not recovered by the expression of a dominant-active form of RagB Q99L (Figure 2B). Consistently, while RagB Q99L expression rescued the lysosomal localization of mTOR in the absence of AA, it failed to recover localization in the absence of glucose/glutamine (Figure 2C), suggesting mTORC1 inhibition occurs either downstream of or in parallel with Rag activation.

Figure 2. Deprivation of glucose and glutamine inhibits mTORC1 independently of the Rag proteins.

Figure 2

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(A) Tsc2-/- MEFs were non-deprived (Con), or deprived of amino acids for 2 hrs, or of glucose and glutamine for 12 hrs. Cells were coimmunostained for mTOR (green) and the lysosomal protein LAMP1 (red), and then processed for images. See supporting data in Figure S2.

(B)3 Ampk α-/- MEFs stably expressing empty, RagB WT, or Q99L were starved of the indicated nutrients, or treated with rapamycin (20 nM). Phosphorylation of S6K1 was measured 10 hrs post deprivation.

(C) Tsc2-/- MEFs stably expressing RagB WT or Q99L were deprived of and processed as in (A)

Mitochondrial inhibitors block mTORC1 activation independently of AMPK and Rags

As shown in Figure 1, AMPK activation - a measure of energetic stress or a rise in AMP levels - was strongly associated with, but not required for, glucose/glutamine deprivation-induced mTORC1 inhibition. Thus, we next investigated if the depletion of TCA cycle intermediates by glucose/glutamine deprivation was inhibiting ATP production, and if ATP reduction was involved in mTORC1 inhibition. We reasoned that if glucose/glutamine deprivation-induced energetic stress is causing mTORC1 inhibition, mitochondrial inhibition should also inhibit mTORC1 independently of AMPK and Rags. Treatment of cells with phenformin (complex I inhibitor), rotenone (complex I inhibitor), or FCCP (mitochondrial uncoupling agent) inhibited mTORC1 activation in Ampkα+/+ and Ampkα-/- cells (Figure 3A). With all of the treatments, the inhibition correlated with reduction of ATP (Figure 3B). The effect of phenformin on mTORC1 in Ampkα-/- cells required higher concentration and took longer to observe than in Ampkα+/+ cells, despite similar effects on ATP in both cell types (Figure 3C-E). Together, these results suggest that besides AMPK activation, an AMPK-independent pathway inhibits mTORC1 following energetic stress. The expression of RagB Q99L failed to prevent mTORC1 inhibition from mitochondrial inhibition (Figure 3F) and also failed to re-localize mTOR to the lysosomes following mitochondria uncoupling (Figure 3G). Consistent with the notion that ATP depletion causes mTORC1 inhibition and mislocalization, deprivation of glucose/glutamine and mitochondrial inhibition decreased ATP levels, which appeared to correlate with mTORC1 inhibition (Figure S3A and 3B), and AICAR treatment failed to affect mTOR localization (Figure S3H). These results suggest that mitochondrial inhibition, likely through ATP depletion, blocks mTORC1 activation and localization through a Rag-independent mechanism.

Figure 3. Mitochondrial inhibitors suppress mTORC1 activity independently of AMPK and Rags.

Figure 3

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(A) Ampk+/+ and Ampk-/- MEFs were treated with phenformin (1 or 10 mM), rotenone (1 or 10 μg/ml) or FCCP (4 or 20 μM), for 1 hr. mTORC1 and AMPKα activity were measured.

(B) Ampk+/+ and Ampk-/- MEFs were treated as in (A) and ATP levels were measured. The data are represented as mean ± standard deviation.

(C-D) Ampk+/+ and Ampk-/- MEFs were treated with phenformin at the indicated concentrations for 1 hr (C) or treated with 5 mM phenformin for the indicated times (D).

(E) ATP levels were measured in Ampk+/+ and Ampk-/- MEFs after treatments as in (D). Each value represents the normalized mean ± standard deviation for n=3.

(F) Ampk-/- MEFs expressing either RagB WT or Q99L were deprived of amino acids (2 hrs) or glucose/glutamine (6 hrs), or treated with phenformin (5 mM), rotenone (10 μg/ml) or FCCP (20 μM), for 1 hr.

(G) Ampk-/- MEFs expressing either RagB WT or Q99L were treated as in (F) and coimmunostained for mTOR (green) and LAMP1 (red).

Glucose/glutamine deprivation-induced mTORC1 inhibition involves energetic stress or ATP depletion

If the loss of ATP from glucose/glutamine deprivation and mitochondrial inhibition was the cause for mTORC1 inhibition, we hypothesized that decreasing metabolic demand should recover the inhibition. Previously, we demonstrated that cycloheximide treatment (translation inhibitor) and total AA deprivation (thus no AA in DMEM) decreased the metabolic demand to allow cells to survive reduction of energy sources (Choo et al., 2010). As shown in Figure S3B and S3C, deprivation of glucose and glutamine strongly depleted cellular ATP levels and increased AMPK activation, but depletion of glucose/glutamine along with all other AA dramatically reduced the turnover of ATP and prevented AMPK activation from double-nutrient deprivation. Importantly, mTORC1 activity in the glucose/glutamine/AA-deprived cells was strongly inhibited, consistent with the role of AA in regulating mTORC1 activity (Figure S3C). Next, we deprived RagB Q99L-expressing cells of glucose/glutamine, AA, or glucose/glutamine/AA and measured mTORC1 activity; our rationale is that if ATP depletion, but not some non-energy-related effect from glucose/glutamine depletion, was causing mTORC1 inhibition, then RagB Q99L expression should recover mTORC1 activity in the glucose/glutamine/AA-deprived cells (RagB Q99L was needed to compensate for AA deprivation, which was needed to decrease metabolic demand). Conversely, if glucose/glutamine were providing an energy-independent effect that is required for mTORC1 activation, then RagB Q99L expression should not recover mTORC1 activity in the glucose/glutamine/AA deprived condition. As shown in Figures S3D and S3E, expression of RagB Q99L, but not WT RagB, significantly recovered mTORC1 activation from glucose/glutamine/AA deprivation, but not from glucose/glutamine deprivation, indicating energetic stress is causing mTORC1 inhibition. Finally, to verify further that energetic stress triggers mTORC1 inhibition, we treated glucose/glutamine-deprived cells with cycloheximide (CHX), which also decreases metabolic consumption, and observed a recovery in mTORC1 activity – something not seen with RagB Q99L expression, suggesting the CHX-mediated increase is not from translation inhibition-induced increase of cellular AA (Figure S3F). Since mTORC1 can also be regulated by other stress pathways, we determined if autophagy, ER stress, apoptosis, and the activation of stress kinases p38 and JNK can be induced by glucose and glutamine deprivation, and correlated with mTORC1 inhibition. We observed no clear association between glucose/glutamine deprivation-induced mTORC1 inhibition and activation of these pathways (Figure S3G). Together, these results suggest that energetic stress or ATP depletion is necessary for glucose/glutamine deprivation to inhibit mTORC1 through a TSC1/2-, AMPK-, and Rag-independent mechanism.

Energetic stress inhibits the assembly of the TTT-RUVBL1/2 complex and prevents the complex interaction with mTOR

To understand how energetic stress prevents mTORC1 activation and lysosomal localization, we performed a high-throughput siRNA screen to look for genes whose loss altered mTORC1 signaling in Tsc2-/- cells (Hoffman et al., 2010). We identified members of the TTT-RUVBL1/2 complex as one of the strongest regulators of mTORC1 in Tsc2-/- cells (Figure 4A-D). The multi-protein TTT complex consists of Tel2, Tti1, and Tti2, and was recently shown to regulate the assembly of PIKK-containing complexes, including ATM, ATR, and mTOR, by interacting with a host of proteins involved in protein folding, assembly, and stability, such as RUVBL1/2 (Horejsi et al., 2010; Hurov et al., 2010; Izumi et al., 2010; Kaizuka et al., 2010; Takai et al., 2007; Takai et al., 2010). Consistently, we observed that mTOR co-immunoprecipitated with Tel2, and that this interaction was reduced following glucose/glutamine deprivation and FCCP treatment, but not following AA-deprivation (Figure 4E).

Figure 4. Energetic stress regulates the assembly of the TTT-RUVBL1/2 complex and its interaction with mTOR.

Figure 4

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(A) List of the subunits of the TTT-RUVBL1/2 complex scored in the siRNA screen. Fold increase was calculated based on changes in phospho-rpS6 signals when the gene was knocked down.

(B-D) Tsc2+/+ and Tsc2-/- MEFs were transfected with the indicated siRNAs, and mTORC1 activity was measured 60 hrs post-transfection.

(E) Tsc2-/- MEFs were starved with glucose/glutamine (12 hrs) or amino acids (2 hrs), or treated with 30 μM FCCP (1 hr). Cells were subjected to immunoprecipitation with anti-mTOR antibody followed by immunoblotting with the indicated antibody.

(F) Tsc2-/- MEFs expressing either human RUVBL1 WT or D302N were transfected with non-targeting control or RUVBL1 siRNA for 48 hrs. Immunoprecipitation was analyzed as in (E).

(G) Tsc2-/- MEFs expressing HA-Tel2 along with either human RUVBL1 WT or D302N were transfected with siRNAs as in (F). Anti-HA immunoprecipitates from the cell lysates were analyzed by immunoblotting with the indicated antibodies.

(H) Cell lysates were prepared from Tsc2-/- MEFs expressing HA-Tel2 deprived of (12 hrs) or re-stimulated with 20 mM glucose and 4 mM glutamine (1 hr). Anti-HA immunoprecipitates were analyzed as in (G). See also Figure S4.

(I) Tsc2-/- MEFs expressing either human RUVBL1 WT or D302N were transfected with siRNAs as in (F). mTORC1 activity was addressed 48 hrs post-transfection.

In complex with the TTT proteins, the RUVBL1/2 ATPases were also strong regulators of mTORC1 (Horejsi et al., 2010; Izumi et al., 2010; Takai et al., 2010) (Figure 4B). Because the consequences of energetic stress include not only AMP accumulation, but also ATP depletion, we tested if the ATPase activity of RUVBLs was required for the mTOR-Tel2 interaction. To this end, we conducted rescue experiments by first stably expressing human RUVBL1 WT or its ATPase activity-deficient mutant (D302N) (Mezard et al., 1997) and then knocking down endogenous Ruvbl1 with siRNA in Tsc2-/- MEFs. It should be noted that the loss of function of either RUVBL proteins is sufficient to inactivate the entire complex (Jonsson et al., 2001), and that we expressed untagged RUVBL1 as the addition of tags alters the proper assembly of the complex (Cheung et al., 2010). While rescuing with WT RUVBL1 had no effect on mTOR-Tel2 interaction, rescuing with the activity-deficient mutant (D302N) prevented mTOR-Tel2 interaction, suggesting the ATPase activity of RUVBLs is necessary for mTOR to interact with Tel2 (Figure 4F). Moreover, rescue with D302N RUVBL1 also prevented Tel2 interaction with both Tti1 and RUVBL1 (Figure 4G). To determine if energetic stress is preventing the ATPase activity of the RUVBLs to inhibit the TTT-RUVBL complex and mTORC1 activity, we tested if energetic stress also disassembled the TTT-RUVBL complex. As shown in Figure 4H, deprivation of glucose/glutamine reduced Tel2 interaction with Tti1 and RUVBL1, and importantly, stimulation with glucose/glutamine for 1 hr (which significantly recovered ATP levels; Figure S4A) recovered complex formation. Moreover, the length of time of glucose and glutamine deprivation appeared to correlate tightly with TTT-RUVBL dissociation and mTORC1 inhibition (Figure S4B and S4C). We observed that mitochondrial inhibitors, phenformin and rotenone, also disrupted the TTT-RUVBL complex (Figure S4D), indicating energetic stress-induced disruption of the complex. Finally, we tested if RUVBL ATPase activity is required for energy-induced mTORC1 activation. Rescue with the D302N mutant prevented mTORC1 activation following glucose/glutamine starvation and stimulation (Figure 4I). These data suggest that energetic stress inhibits the assembly of the TTT-RUVBL complex, which is required for mTORC1 activation, and that the ATPase activity of RUVBLs is necessary for complex formation and its interaction with mTOR.

Energetic stress inhibits the activity of the TTT-RUVBL1/2 complex

Since energetic stress disassembled the TTT-RUVBL complex, we next tested if it also inhibited the activity of the complex. Although there are no assays that could directly measure the in vivo activity of RUVBLs, as an alternative, we deduced that energetic stress should also inhibit mTORC1-independent functions of the TTT-RUVBL complex. Accordingly, we measured the activity of mTORC2 (Kaizuka et al., 2010; Takai et al., 2010), the stability of other PIKK proteins (Horejsi et al., 2010; Izumi et al., 2010; Takai et al., 2007), and the DNA damage response (Hurov et al., 2010; Izumi et al., 2010) – three well-characterized processes. As shown in Figure S5A-E, both knockdown of RUVBL1 and energetic stress blocked AKT phosphorylation at serine 473, a mTORC2 site; reduced the protein levels of the PIKK family members ATM and ATR; and prevented etoposide-induced phosphorylation of CHK1, an event that requires the ATM/ATR proteins. For the protein turnover experiments, we performed the siRNA analysis after 60 hrs, at which time we observed significant protein reduction of PIKK enzymes, and energetic stress experiments after 24 hrs of starvation as mTOR levels did not change up to 12 hrs of deprivation (Figure 6B).

Figure 6. Energetic stress and loss of the TTT-RUVBL1/2 complex inhibit mTORC1 through mislocalization and prevention of mTORC1 dimerization.

Figure 6

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(A) Tsc2-/- MEFs expressing Myc-Raptor WT or Myc-Raptor-CAAX were deprived of glucose/glutamine (12 or 24 hrs) or of amino acids/serum (2 hrs), and mTORC1 activity was measured.

(B) Glucose/glutamine-deprived Tsc2-/- MEFs (12 or 24 hrs) were stimulated with glucose (20 mM)/glutamine (4 mM) for 1 hr, and mTOR levels and activity were measured.

(C) Myc-Raptor- or Myc-Raptor-CAAX-expressing cells were transfected with siRNAs against Tti1 or RUVBL1, and mTORC1 activity was measured 36 hrs post-transfection.

(D-E) Tsc2+/+ MEFs following 12 hrs glucose/glutamine deprivation (D) or 36 hrs siRNA transfection (E) were lysed with the lysis buffer containing the reduced salts. mTOR-Raptor interaction was measured using anti-Raptor immunoprecipitates followed by immunoblotting with the indicated antibodies. See also Figure S6.

(F) Tsc2-/- MEFs expressing HA-Raptor and Myc-Raptor were deprived of (12 hrs) and stimulated with glucose/glutamine (1 hr). Anti-HA immunoprecipitates were prepared from cells treated with DSP for 1 hr prior to lysis and analyzed by immunoblotting with the indicated antibodies.

(G) Tsc2-/- MEFs expressing HA-Raptor and Myc-Raptor were transfected with the indicated siRNAs, and treated and processed as in (F). Anti-HA immunoprecipitates were prepared 42 hrs post-siRNA transfection.

Loss of the TTT-RUVBL1/2 complex prevents mTOR localization to the lysosome and mTORC1-Rag association

Because energetic stress-induced mTORC1 inhibition involves mislocalization of mTOR and disassembly of the TTT-RUVBL complex, we tested if the loss of the complex could disrupt the localization of mTOR. Knockdown of either RUVBL1 or 2 was sufficient to disrupt lysosomal localization of mTOR (Figure 5A), and the effect was comparable to that achieved with glucose/glutamine-starvation (Figure 5B). We analyzed mTOR localization 36-42 hrs post-siRNA transfection, time-points at which we observed RUVBL knockdown and mTORC1 inhibition, but without either mTOR turnover or disassembly of mTOR-Raptor, indicating the knockdown-induced mTOR mislocalization was not from disruption of mTOR-Raptor interaction or general reduction of mTOR levels (data not shown; mTOR-Raptor interaction is discussed further in Figure 6).

Figure 5. Energetic stress and loss of RUVBL1/2 prevent mTORC1 recruitment to the lysosomes and mTORC1-Rag interaction.

Figure 5

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(A) Tsc2-/- MEFs were transfected with siRNA against RUVBL1 or RUVBL2 for 42 hrs, or deprived of glucose and glutamine for 12 hrs. Cells were then coimmunostained for mTOR and LAMP1, and processed for images.

(B) Quantitation of mTOR localized to the lysosomes was measured based on percentages of the mTOR area overlapped with the LAMP1 area using the Measure Colocalization function under the MetaMorph® program. At least three images processed as in (A) were analyzed per sample. The data are represented as mean ± standard deviation and p-value is indicated.

(C) Tsc2-/- MEFs expressing either Flag-RagB WT or Q99L were deprived of amino acids (2 hrs) or glucose/glutamine (12 hrs), and then stimulated with amino acids (30 min.) or glucose/glutamine (1 hr). After 1 hr treatment with 1 mg/ml DSP prior to lysis, anti-Flag immunoprecipitates were analyzed by immunoblotting with the indicated antibodies.

(D) Tsc2-/- MEFs expressing Flag-RagB WT were transfected with siRNAs against Tti1 or RUVBL1, and then starved (12 hrs) and stimulated with glucose and glutamine (1 hr). Anti-Flag immunoprecipitates were prepared 40 hrs post-siRNA transfection and analyzed as in (C).

As the expression of RagB Q99L was insufficient to recover mTORC1 activity or localization (Figure 2 and 3), we next investigated if the TTT-RUVBL complex was necessary for mTORC1-Rag interaction, as TTT-RUVBL regulates the assembly of various PIKK-containing complexes (Izumi et al., 2010). As shown in Figure 5C, glucose/glutamine starvation (as well as AA starvation) decreased the WT RagB-mTORC1 interaction, but unlike from AA starvation, glucose/glutamine starvation prevented RagB Q99L-mTORC1 interaction, highlighting the reason why RagB Q99L expression was insufficient to recover energetic stress-induced mTORC1 inhibition. Knockdown of Tti1 or RUVBL1 for 40 hrs prevented mTORC1-RagB interaction induced by glucose/glutamine stimulation without affecting mTOR turnover (Figure 5D); therefore, the TTT-RUVBL complex is required for in vivo Rag association with mTORC1.

Forced-localization of mTOR to Rheb-containing endomembranes is insufficient to recover mTORC1 inhibition from energetic stress or loss of the TTT-RUVBL1/2 complex

If mislocalization of mTOR is causative for energetic stress-induced mTORC1 inhibition, we reasoned that forcing mTOR localization to the lysosomes independently of the Rags should recover glucose/glutamine deprivation-induced mTORC1 inhibition. We expressed either Raptor WT or Raptor-CAAX and measured mTORC1 activity; the Raptor-CAAX mutant expresses the CAAX sequence from Rheb, forces mTORC1 localization to Rheb-containing endomembrane sites, and recovers mTORC1 activity following AA-deprivation (Sancak et al., 2010). While Raptor-CAAX expression almost completely restored mTORC1 activity following AA deprivation, it only partially recovered mTORC1 inhibition following 12-24 hrs of glucose/glutamine deprivation (Figure 6A). The incomplete recovery with Raptor-CAAX expression did not result from the overall reduction of mTOR levels seen after prolonged energetic stress as re-stimulation of 12- or 24-hr deprived cells with glucose/glutamine recovered mTORC1 activity, at least to a degree much greater than that observed in cells expressing Raptor-CAAX (Figure 6B). In addition, Raptor-CAAX expression failed to render the mTORC1 pathway insensitive to the loss of the TTT-RUVBL complex (Figure 6C). Thus, mislocalization of mTORC1 is insufficient to explain completely the mechanism of energetic stress-induced mTORC1 inhibition.

Energetic stress or loss of the TTT-RUVBL1/2 complex prevents mTORC1 dimerization

One possibility is that inhibition of the TTT-RUVBL complex does something more than just altering mTOR localization. As previously mentioned, the TTT complex has been reported to regulate mTOR-Raptor interaction (Kaizuka et al., 2010; Takai et al., 2010). But analyzing mTOR-Raptor interaction and the consequences of the interaction is difficult, as the more “active” mTORC1 interacts with less affinity, while the “less” active form interacts with greater affinity (Kim et al., 2002). Recently, Sancak et al. showed that reducing the salt concentration in the lysis buffer allows mTOR and Raptor to co-precipitate equally in active and inactive conditions (Sancak et al., 2007). Using this protocol, we measured the mTOR-Raptor interaction; as shown in Figure 6D-E and S6, the mTOR-Raptor interaction was not altered from energetic stress (12 hrs of starvation) or from knockdown of Tti1 or RUVBL1 after 36-42 hrs post-transfection (time-points at which we failed to observe mTOR protein reduction, but did observe mTORC1 inhibition). However, consistent with the previous reports (Kaizuka et al., 2010; Takai et al., 2010), analysis at longer time-points, in which mTOR levels are reduced, revealed reduced mTOR-Raptor interaction (data not shown). These data suggest that both energetic stress and loss of the TTT-RUVBL complex initially inhibit mTORC1 without disrupting mTOR-Raptor interaction. We next reasoned that earlier time-points in energetic stress or knockdown may not provide sufficient time to break apart the mTOR-Raptor interaction, but could alter other assembly-related effects. Recently, Yip et al. reported the EM structure of mTORC1, and proposed that two mTORC1s come together to form a rhomboid-shaped, obligate dimer to create a central cavity for substrate entry (Yip et al., 2010). We analyzed the dimerization of mTORC1 complexes by expressing two differently tagged Raptor constructs (Myc- and HA-tag) in the same cell and with aid of the intracellular cross-linker DSP. As shown in Figure 6F, glucose/glutamine deprivation significantly reduced mTORC1 dimerization, while acute stimulation recovered this interaction; the increased dimerization was concomitant with increased RUVBL1 and Tel2 association with mTORC1, and knockdown of RUVBL1 or Tti1 also reduced glucose/glutamine-induced dimerization of mTORC1 (Figure 6G). Therefore, these data suggest that energetic stress- or TTT-RUVBL inhibition-induced mTORC1 repression involves both the mislocalization and the reduced dimerization of the complex.

TTT-RUVBL1/2 complex genes and various metabolic genes downstream of mTORC1 are coordinately upregulated in breast carcinomas

Given that mTORC1 is hyperactive in many cancers, we examined if there was any link between expression of the TTT-RUBVL1/2-associated genes and mTORC1 signaling in cancer tissues. Using comprehensive meta-analysis of The Cancer Genome Atlas (TCGA) datasets, we found that the expression of many TTT-RUVBL complex genes was significantly higher in breast (Figure 7A) and colorectal (Figure 7B) carcinomas when compared to their normal tissue controls. Of the breast carcinomas, we identified a unique subgroup with high mRNA levels of the TTT-RUVBL complex genes (green) (Figure 7C), and the increase in mRNA levels of these genes correlated with each other (Figure 7D). Interestingly, we observed the highest correlation between mRNAs of genes that encode Tel2 and Tti1 and between those that encode RUVBL1 and RUVBL2 (Figure 7D, indicated in blue). We also observed a highly significant positive correlation between mRNAs of the TTT-RUVBL complex genes and the expression of genes involved in glycolysis (Figure 7E), pentose phosphate pathway (Figure 7F), lipid/sterol biosynthesis (Figure 7G and S7B-C), and mitochondrial function (Figure S7A) - all pathways that mTORC1 positively regulates via transcriptional control mechanisms (Duvel et al., 2010; Cunningham et al., 2007). Collectively, these data are consistent with the hypothesis that elevated expression of the TTT-RUVBL complex promotes mTORC1 signaling and metabolic processes that are necessary for tumor cell growth.

Figure 7. Bioinformatic analysis of the TTT-RUVBL1/2 complex genes in cancer.

Figure 7

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(A-B) Box-plots indicate significantly higher expression of the complex’s genes in breast (n=593) and colorectal (n=237) carcinomas compared to normal (P<0.05) based on TCGA database.

(C) Heat-map diagram of unsupervised hierarchical clustering of the complex’s genes mRNA levels indicates two clearly-defined groups of breast tumors with high and low expression of the complex’s genes.

(D) Spearman’s rank correlation coefficient matrix indicates high positive correlation (P<0.001) in the expression of the complex’s genes in breast carcinomas, in particular between TEL2-TTI1 and RUVBL1-RUVBL2 genes.

(E-G) Spearman’s rank correlation coefficient matrices indicate high positive correlation (P<0.001) between the expression of the complex’s genes and putative target genes of mTORC1 signaling encoding glycolytic (E), pentose phosphate pathway (F) and lipid/sterol biosynthesis enzymes (G). See also Figure S7.

(H) Model for glucose/glutamine (energy)-induced mTORC1 activation. Through glycolysis and glutaminolysis, glucose and glutamine provide carbons to the TCA cycle and consequently control ATP production. Upon availability of enough energy, the TTT-RUVBL1/2 complex gets stabilized and thus plays major roles in formation of the mTORC1 complex into the mature dimeric form, which may be necessary for Rag-dependent lysosomal localization of mTORC1.

Discussion

The mTORC1 signaling pathway integrates various extracellular cues to regulate cell growth; therefore, we investigated how glucose and glutamine – the two primary carbon sources for macromolecule synthesis, replenishment of the TCA cycle, and energy production – regulated mTORC1 activity. We discovered that glucose and glutamine – by supplying carbons to the TCA cycle and controlling ATP production – feeds back to regulate mTORC1 activation through an AMPK-, TSC1/2-, and Rag-independent mechanism, in part, by preventing mTOR localization to the lysosomes (Figure 7H). Recently, Kalender et al. reported that metformin inhibited mTORC1 independently of TSC1/2 and AMPK but instead through a Rag-dependent mechanism (Kalender et al., 2010). However, we could not observe significant recovery of energetic stress-induced mTORC1 inhibition or mislocalization in cells stably expressing dominant-active RagB. Their observation may have been a consequence of overexpression as increasing total mTORC1 recruitment to the lysosomes is likely to confound the readout of overall mTORC1 activity despite decreased activity per mTORC1 complex, much like what is often observed with overexpressing Rheb (Sancak et al., 2008). Although we intentionally utilized Tsc-/- and Ampk-/- cells to identify the mislocalization of mTOR following energetic stress, we believe this mechanism is critical in various cellular contexts; this effect is observed in the cell types containing TSC and AMPK function and controls Rag-mTORC1 interaction and mTORC1 dimerization, which are necessary for mTORC1 function (Figure 7H).

We have discovered that energetic stress inhibited the assembly of the TTT-RUVBL complex, which consists of Tel2, Tti1, Tti2, and the ATPases RUVBL1/2, as well as other components such as RPAP3, PIH1D1, and Hsp90 (Horejsi et al., 2010; Hurov et al., 2010; Izumi et al., 2010; Takai et al., 2010). Although first reported as a regulator of PIKK stability (Takai et al., 2007), Tel2 and its complex also control the assembly and activity of PIKK-containing complexes; for example, TTT-RUVBL regulates the ATR-ATRIP interaction as well as the assembly of mRNA surveillance complexes involved in NMD, and this regulation occurs prior to turnover of the PIKKs (Izumi et al., 2010). We also observed that prior to mTOR turnover or mTOR-Raptor dissociation, loss of the TTT-RUVBL complex or energetic stress induced mislocalization of mTOR, dissociation of Rag-mTORC1 interaction, and inhibition of mTORC1 dimerization, suggesting that these effects are a result of a specific, energy-regulated process, and not a result of global decrease in mTOR or mTOR-Raptor disassociation. Interestingly, the inhibition of TTT-RUVBL by energetic stress couples mTORC1 repression with a number of other cellular processes, such as NMD and ATM/ATR activation, ostensibly to eliminate all energetic events so as to prolong survival.

Our results raise several interesting points regarding the biology of mTORC1 and TTT-RUVBL. We have shown that the loss of RUVBL ATPase activity is sufficient to mimic the various effects of energetic stress, including disassembly of the TTT-RUVBL complex. Hsp90 has also been shown to associate with TTT-RUVBL and thus its ATPase activity may also contribute to energy regulation of mTORC1. However, 17-AAG treatment (Hsp90 inhibitor) did not alter mTORC1 localization (data not shown), suggesting a non-essential role of Hsp90 at least in the processes described here. Likewise, if dimerization of mTORC1 is a regulated process, do other nutrient and/or signaling pathways control this event? Our results suggest that AA levels do not affect mTORC1 dimerization (data not shown). But considering GTP-charged Rheb directly activates mTORC1 in vitro (Long et al., 2005; Sancak et al., 2007), an interesting question is if Rheb somehow alters dimerized mTORC1 to create the central cavity believed to be necessary for substrate access.

In conclusion, we have demonstrated that the TTT-RUVBL complex responding to the cell’s metabolic state controls mTORC1 homodimerization and lysosomal localization, integrating energy signaling input as a requirement for AA/Rag-mediated activation of mTORC1 (Figure 7H).

Experimental Procedures

Cell lines and culture media

Tsc2+/+, p53-/- MEFs, Tsc2-/-, p53-/- MEFs, Tsc1-/-, p53+/+ MEFs, Ampkα+/+ MEFs, Ampkα-/- MEFs, Raptor WT Tsc2-/-, and Raptor AA Tsc2-/- MEFs were cultured in Dulbecco’s modified Eagle’s medium (DMEM) with 10% fetal bovine serum (FBS). DMEM lacking glucose, amino acids, L-glutamine, or combinations were prepared based on the formulation provided by Cellgro. Extra energetic additives that are often added to some DMEM formulations such as sodium pyruvate and succinate were excluded. All DMEM deficient of certain nutrients were applied to the cell with 10% dialyzed FBS (Invitrogen) after washing once with DMEM free of all nutrients and serum.

Immunofluorescence assays

Cell images were taken with a 60x or 20x objective using spinning disk confocal microscopy (Nikon), processed, and analyzed using the MetaMorph® program. A detailed procedure for preparing the specimens is described in the Supplemental information.

In vivo cross linking assay

As described (Sancak et al., 2008) but with some modifications according to the manufacturer’s instruction, cells were intracellularly cross-linked with DSP (Pierce) prior to lysis. More details are provided in the Supplemental information.

Other experimental procedures

All other experimental procedures, including materials, plasmid constructs, immunoprecipitations, siRNA transfections, retroviral infections, metabolic assays, and bioinformatic analysis, are described in the Supplemental information

Supplementary Material

01

Highlights.

  • Glucose and glutamine feed back to promote mTORC1 signaling through ATP production

  • Energetic stress prevents mTOR lysosomal localization independently of AMPK and Rag

  • ATP-dependent TTT-RUVBL complex is disassembled and repressed by energetic stress

  • TTT-RUVBL is required for mTORC1 functional assembly and lysosomal localization

Acknowledgments

We would like to thank members of the Blenis laboratory for helpful discussions and critical reading of the manuscript. We also acknowledge the generous gifts of reagents from the following: Brendan Manning for Tsc2-/-, p53-/- MEFs, David Kwiatkowski for Tsc1-/-, p53+/+ MEFs, Reuben Shaw for Ampkα1,2+/+, Ampkα1,2-/-, WT Raptor Tsc2-/-, and AA Raptor Tsc2-/- MEFs, and Diane Fingar for the mTOR antibody. We are also grateful to the Nikon Imaging Center at Harvard Medical School for help with microscopy. This work was supported by grants from NIH RO1GM051405 (J.B.), the Tuberous Sclerosis Alliance (G.R.H.) and the World Class Institute research program of the National Research Foundation of Korea (Y.J.J., K.W.L., B.Y.K., R.L.E. and J.B.). J. B. is an Established Investigator of the LAM Foundation.

Footnotes

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