Exploiting macrophage autophagy-lysosomal biogenesis as a therapy for atherosclerosis

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Abstract

Macrophages specialize in removing lipids and debris present in the atherosclerotic plaque. However, plaque progression renders macrophages unable to degrade exogenous atherogenic material and endogenous cargo including dysfunctional proteins and organelles. Here we show that a decline in the autophagy–lysosome system contributes to this as evidenced by a derangement in key autophagy markers in both mouse and human atherosclerotic plaques. By augmenting macrophage TFEB, the master transcriptional regulator of autophagy–lysosomal biogenesis, we can reverse the autophagy dysfunction of plaques, enhance aggrephagy of p62-enriched protein aggregates and blunt macrophage apoptosis and pro-inflammatory IL-1β levels, leading to reduced atherosclerosis. In order to harness this degradative response therapeutically, we also describe a natural sugar called trehalose as an inducer of macrophage autophagy–lysosomal biogenesis and show trehalose's ability to recapitulate the atheroprotective properties of macrophage TFEB overexpression. Our data support this practical method of enhancing the degradative capacity of macrophages as a therapy for atherosclerotic vascular disease.

Abstract

Dysfunction of autophagy in plaque macrophages aggravates atherosclerosis. Here the authors show that induction of macrophage autophagy–lysosomal biogenesis either genetically by overexpression of the master transcriptional regulator of this process, TFEB, or pharmacologically with trehalose is atheroprotective.

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NLRP3 inflamasomes are required for atherogenesis and activated by cholesterol crystals that form early in disease

Peter Duewell, Hajime Kono, Katey Rayner … (2010)

The inflammatory nature of atherosclerosis is well established but the agent(s) that incite inflammation in the artery wall remain largely unknown. Germ-free animals are susceptible to atherosclerosis, suggesting that endogenous substances initiate the inflammation1. Mature atherosclerotic lesions contain macroscopic deposits of cholesterol crystals in the necrotic core but their appearance late in atherogenesis had been thought to disqualify them as primary inflammatory stimuli. However, using a novel microscopic technique, we revealed that minute cholesterol crystals are present in early diet-induced atherosclerotic lesions and that their appearance coincides with the first appearance of inflammatory cells. Other crystalline substances can induce inflammation by stimulating the caspase-1-activating NLRP3 inflammasome2,3, which results in cleavage and secretion of IL-1 family cytokines. Here, we demonstrate that cholesterol crystals also activate the NLRP3 inflammasome in phagocytes in vitro in a process that involves phago-lysosomal damage. Similarly, when injected intraperitoneally, cholesterol crystals induce acute inflammation, which is impaired in mice deficient in components of the NLRP3 inflammasome, cathepsin B, cathepsin L, or IL-1 molecules. Moreover, when low-density lipoprotein receptor (LDLR) deficient mice were reconstituted with NLRP3-, ASC-, or IL-1α/β-deficient bone marrow and fed a high cholesterol diet, they had markedly reduced early atherosclerosis and inflammasome-dependent IL-18 levels. Our results demonstrate that crystalline cholesterol acts as an endogenous danger signal and its deposition in arteries or elsewhere is an early cause rather than a late consequence of inflammation. These findings provide new insights into the pathogenesis of atherosclerosis and point to new potential molecular targets for the therapy of this disease. Cholesterol, an indispensable lipid in vertebrates, is effectively insoluble in aqueous environments and elaborate molecular mechanisms have evolved that regulate cholesterol synthesis and its transport in fluids4. Cholesterol crystals are recognized as a hallmark of atherosclerotic lesions5 and their appearance helps in the histopathological classification of advanced atherosclerotic lesions6. However, crystalline cholesterol is soluble in the organic solvents used in histology, so that the presence of large crystals is identifiable but only indirectly as so-called cholesterol crystal clefts, which delineate the space that was occupied before sample preparation. The large cholesterol crystal clefts in atherosclerotic plaques were typically only observed in advanced lesions and, therefore, crystal deposition was thought to arise late in this disease. However, given that atherosclerosis is intimately linked to cholesterol levels, we were interested to determine when and where cholesterol crystals first appear during atherogenesis. We fed atherosclerosis-prone Apo-E-deficient mice a high cholesterol diet to induce atherosclerosis7,8 and used a combination of laser reflection and fluorescence confocal microscopy3 to identify crystalline materials and immune cells. Many small crystals appeared as early as two weeks after the start of atherogenic diet within small accumulations of subendothelial immune cells in very early atherosclerotic sinus lesions (Fig. 1a, b and Supplementary Fig. 1, 2). The reflective material was identified as being mostly cholesterol crystals by fillipin staining (not shown). Crystal deposition and immune cell recruitment increased steadily with diet feeding and the appearance of crystals correlated with that of macrophages (r2=0.99, p<0.001) (Fig. 1a-d). Cholesterol crystals were not only detected in necrotic cores but also in subendothelial areas that were rich in immune cells. Confocal imaging revealed crystals to localize both inside and outside of cells (Fig. 1b), whereas in corresponding H&E stained sections that were treated with organic solvents during the staining process cholesterol crystal clefts were visible only after 8 weeks of diet and smaller crystals remained invisible (Fig. 1a). As expected, we failed to detect macrophages or accumulation of crystals in the aortic sinus sections in mice on a regular chow diet (Fig. 1a, b; bottom panel). Additional observations in human advanced atherosclerotic lesions showed that areas rich in immune cells also contained smaller crystals inside and outside of cells in addition to the larger crystals that would leave cholesterol crystal clefts in standard histology (Supplementary Fig. 3, 4). These studies establish that crystals emerge at the earliest time points of diet-induced atherogenesis together with the appearance of immune cells in the subendothelial space. Various crystals that are linked to tissue inflammation, as well as pore-forming toxins or extracellular ATP, can activate IL-1 family cytokines via triggering of NLRP39. Of note, NLRP3 inflammasome formation requires a priming step that can be provided by pattern recognition or cytokine receptors that activate NF-κB. Cellular priming leads to induction of pro-forms of IL-1 family cytokines and NLRP3 itself, a step, which is required for NRLP3 activation at least in vitro 10. To test whether cholesterol crystals could activate the release of IL-1β, we incubated LPS-primed human PBMCs with cholesterol crystals. Cholesterol crystals induced a robust, dose-responsive release of cleaved IL-1β in a caspase-1 dependent manner (Fig. 2a, b). Of note, cholesterol crystals added to unprimed cells did not release IL-1β into the supernatant indicating the absence of any contaminants that would be sufficient for priming of cells (Fig. 2a)10. IL-1 cytokines are processed by caspase-1, which can be activated by various inflammasomes9. Since the NLRP3 inflammasome has been reported to recognize a variety of crystals, we next stimulated macrophages from mice deficient in NLRP3 inflammasome components. Cholesterol crystals induced caspase-1 cleavage and IL-1β release in wild-type but not NLRP3- or ASC-deficient macrophages (Fig. 2c, d). Transfected dsDNA (dAdT), a control activator that induces the AIM-2 inflammasome11, activated caspase-1 and induced IL-1β release in an ASC-dependent yet NLRP3-independent manner, as expected (Fig. 2c, d). In addition, mouse macrophages also produced cleaved IL-18, another IL-1 family member that is processed by inflammasomes (Fig. 2e). We also found that chemically pure synthetic cholesterol crystals activated the NLRP3 inflammasome providing further evidence that cholesterol crystals themselves rather than contaminating molecules were the biologically active material (Supplementary Fig. 5a). Notably, priming of cells for NLRP3 activation could be achieved by other pro-inflammatory substances such as cell wall components of Gram-positive bacteria (Supplementary Fig. 5b). Moreover, minimally modified LDL also primes cells for NLRP3 activation (Supplementary Fig. 5c)12. Together, these data establish that crystalline cholesterol leads to NLRP3 inflammasome activation in human and mouse immune cells. Several hypotheses regarding the molecular mechanisms of NLRP3 inflammasome activation have been formulated3,13. To further elucidate the mechanisms involved in cholesterol crystal recognition, we pharmacologically inhibited phagocytosis with cytochalasin D or lantriculin A and found that these agents inhibited NLRP3 inflammasome activation by crystals (Fig. 3 and Supplementary Fig. 6 a, c, d). In contrast, these inhibitors did not block the activation of the NLRP3 inflammasome by the pore-forming toxin nigericin or the AIM2 activator dAdT (Fig. 3a and Supplementary Fig. 6a, c, d). To follow the fate of the internalized particles, we performed combined confocal reflection and fluorescence microscopy in macrophages incubated with cholesterol crystals. This analysis revealed that cholesterol crystals induced profound swelling in a fraction of cells (Fig. 3b) as observed for other aggregated materials3,14. Phago-lysosomal membranes contain lipid raft components15, which allowed us to stain the surface of cells with the raft marker choleratoxin B labeled with one fluorescent color and additionally label internal phago-lysosomal membranes after cell permeabilization with differently fluorescing choleratoxin B. Indeed, in macrophages that had previously ingested cholesterol crystals this staining revealed that some cholesterol crystals lacked phago-lysosomal membranes and resided in the cytosol of a fraction of cells, thus indirectly indicating crystal-induced phago-lysosomal membrane rupture (Fig. 3c). This finding was further supported by crystal-induced translocation of soluble lysosomal markers into the cytosol (see below). Additionally, in mouse macrophages cholesterol crystals dose-responsively led to a loss of lysosomal acridine orange fluorescence further confirming lysosomal disruption (Fig. 3d). These studies suggest that cholesterol crystals induced lysosomal damage in macrophages leads to the translocation of phago-lysosomal content into the cytosol. In further experiments we found that the inhibition of lysosomal acidification or cathepsin activity blocked the ability of cholesterol crystals to induce IL-1β secretion (Fig. 3e). Likewise, analysis of cells from mice deficient in single cathepsins (B or L) also showed that cholesterol crystals led to a diminished IL-1β release when compared to wild-type cells. However, the dependency of cholesterol crystal-induced IL-1β release on single cathepsins was less pronounced at higher doses suggesting functional redundancy of cathepsin B and L or potentially additional proteases (Fig. 3f). Together, these experiments suggest that cholesterol crystals induce translocation of lysosomal proteolytic contents, which can be sensed by the NLRP3 inflammasome by as yet undefined mechanisms. It has previously been demonstrated that oxidized LDL, a major lipid species deposited in vessels, has the potential to damage lysosomal membranes16. We found that macrophages incubated with oxidized LDL internalized this material and nucleated crystals in large, swollen, phago-lysosomal compartments (Fig. 3g); and in some cells these compartments ruptured with translocation of the fluorescent marker dye into the cytosol (Fig. 3g, arrows). A time course analysis revealed that small crystals appeared as early as one hour after incubation with oxidized LDL (not shown) and larger crystals were visible after longer incubation times (Fig. 3h). It is likely that cholesterol crystals form due to the activity of acid cholesterol ester hydolases, which transform cholesteryl esters supplied by oxidized LDL into free cholesterol. As indicated above, minimally modified LDL can prime cells for the NLRP3 inflammasome activation (Supplementary Fig. 5c). Recent evidence suggests that this priming proceeds via the activation of a TLR4/6 homodimer and CD3612. This, together with the propensity of minimally modified LDL to form crystals and to rupture lysosomal membranes, suggests that these LDL species could be sufficient to provide both signals 1 and 2 needed to activate IL-1β release from cells. Indeed, after 24 h incubation we observed spontaneous release of IL-1β in the absence of further NLRP3 inflammasome stimulation (Fig. 3i). In murine atherosclerotic lesions we identified not only macrophages and dendritic cells but also neutrophils accumulated within the intima space (see Supplementary Fig. 2). IL-1β plays a key role in the recruitment of neutrophils, and the IL-1-dependent intraperitoneal accumulation of neutrophils has frequently been used as an in vivo assay for inflammasome activation and IL-1 production2,17,18. Using this acute inflammation model we found that cholesterol crystals induced a robust induction of neutrophil influx into the peritoneum (Fig. 4a). Neutrophil influx into the peritoneum after cholesterol crystal deposition was markedly reduced in mice lacking IL-1 or the IL-1 receptor (IL-1R), indicating that IL-1 production is indeed induced and essential for cholesterol crystal-induced inflammation in vivo. Moreover, mice lacking NLRP3 inflammasome components or cathepsins B or L also recruited significantly fewer neutrophils into the peritoneum after cholesterol crystal injection than wild-type mice. However, the reduction in neutrophilic influx observed after cholesterol crystal deposition was more pronounced in mice lacking IL-1 related genes than in mice lacking NLRP3 inflammasome related genes (Fig. 4a), presumably because of the contribution of IL-1α signaling and/or caspase-1-independent processing of IL-1β19 in vivo. In any case, these data confirm that cholesterol crystals trigger NLRP3 inflammasome-dependent IL-1 production in vivo. To test whether the NLRP3 inflammasome is involved in the chronic inflammation that underlies atherogenesis in vessel walls, we tested whether the absence of NLRP3, ASC or IL-1 cytokines might modulate atherosclerosis development in LDLR-deficient mice20, a model for familial hypercholesterolemia. We reconstituted lethally irradiated LDLR-deficient mice with bone marrow from wild-type or NLRP3-,ASC- or IL1α/β-deficient mice and subjected these mice to 8 weeks of a high cholesterol diet. In these radiation bone marrow chimeras, the LDLR-deficiency radioresistant parenchyma causes the animals to become hypercholesterolemic when placed on a high fat diet, while their bone-marrow derived macrophages and other leukocytes lack the NLRP3-inflammasome or IL-1 pathway components needed to respond to cholesterol crystals. We found that the different groups of mice had similar levels of elevated blood cholesterol (not shown). However, mice reconstituted with NLRP3-, ASC-, or IL-1α/b-deficient bone marrow showed significantly lower plasma IL-18 levels, an IL-1 family cytokine whose secretion is dependent on inflammasomes and a biomarker known to be elevated in atherosclerosis21 (Fig. 4b). Additionally, and most importantly, mice whose bone marrow-derived cells lacked NLRP3 inflammasome components or IL-1 cytokines were markedly resistant to developing atherosclerosis (Fig. 4c, d). The lesional area in the aortas of these mice was reduced on average by 69% compared to chimeric LDLR-deficient mice that had wild-type bone marrow. These data demonstrate that activation of the NLRP3 inflammasome by bone marrow derived cells is a major contributor to diet-induced atherosclerosis in mice. The molecules that incite inflammation in atherosclerotic lesions have presented a long-standing puzzle. While the lesions are absolutely dependent on cholesterol, this abundant, naturally occurring molecule has been viewed as inert. Here, we show that the crystalline form of cholesterol can induce inflammation. The magnitude of the inflammatory response and the mechanism of NRLP3 activation appear identical to that of crystalline uric acid, silica and asbestos2,3,13. All these crystals are known to provoke clinically important inflammation as seen in gout, silicosis and asbestosis, respectively. The chronic inflammation in gout, silicosis and asbestosis is thought to derive from the inability of cells to destroy the ingested aggregates leading to successive rounds of apoptosis and reingestion of the crystalline material22. In the same way, immune cells cannot degrade cholesterol and depend, instead, on exporting the cholesterol to HDL particles, which carry the cholesterol to the liver for disposal. The success of this or any cellular mechanism in clearing crystals may thus depend on the availability of HDL. Low blood HDL levels are among the most prominent risk factors for atherosclerotic disease23, and pharmacologic means for increasing HDL are being actively pursued as treatments. Even though cholesterol cannot be degraded by peripheral cells it may be transformed to cholesteryl ester by the cellular enzyme, acylcoenzyme A:cholesterol acyltransferase (ACAT). Cholesteryl esters form droplets rather than crystals and are considered a storage form of cholesterol4. On the assumption that reduced cholesterol storage would be beneficial for reducing atherosclerosis, ACAT inhibitors were tested in large clinical trials. Studies with two such inhibitors showed not a decrease but an increase in the size of the coronary atheroma24,25. This apparent paradox may be reconciled by our findings that the crystalline form of cholesterol, which would be expected to be increased after inhibition of ACAT, may be key in driving arterial inflammation. Indeed, murine studies of ACAT-deficiency show enhanced atherogenesis with abundant cholesterol crystals26. Based on our findings, therapeutic strategies that would reduce cholesterol crystals or block the inflammasome pathway would be predicted to have clinical benefit by reducing the initiation or progression of atherosclerosis. In this context our findings also point to novel molecular targets for the development of therapeutics to treat this disease. Methods summary Mice Mice were kindly provided as follows: NLRP3−/− and ASC−/− (Millenium Pharmaceuticals); Caspase-1−/− (R. Flavell, Yale University, New Haven, CT). Cathepsin B−/− (T. Reinheckel, Albert-Ludwigs-University, Freiburg, Germany), Cathepsin L−/− (H. Ploegh, Whitehead Institute, Cambridge, MA), IL-1α−/− IL-1β−/−, IL-1α−/−β−/− (Yoichiro Iwakura, The Institute of Medical Sciences, The University of Tokyo, Tokyo, Japan). B6-129 (mixed background), C57BL/6, IL-1R−/−, ApoE−/− and LDLR−/− mice were purchased from The Jackson Laboratories. Animal experiments were approved by the UMass and Massachusetts General Hospital Animal Care and Use Committees. Cell culture media and reagents Immortalized macrophage cell lines and bone-marrow derived cells were cultured as described3 and primed with 10 ng/ml LPS for 2h prior to addition of inflammasome stimuli. Inhibitors were added 30 min prior to stimuli. Crystals and dAdT were applied 6h, ATP (5 mM) and nigericin (10 µM) 1h before supernatant was collected. Poly(dA:dT) was transfected using Lipofectamine 2000 (Invitrogen). Human PBMCs were freshly isolated by Ficoll-Hypaque gradient centrifugation, grown in RPMI medium (Invitrogen), 10% FBS (Atlas Biologicals) 10µg/ml ciprofloxacin (Celgro) at 2 × 105 cells per 96 well and primed with 50 pg/ml LPS for 2 hours before addition of inflammasome stimuli. Supernatants were assessed for IL-1β by ELISA and western blot. Neutrophil recruitment to peritoneal cavity Mice were intraperitoneally injected with 2 mg of cholesterol crystals in 200 µl PBS or PBS alone. After 15 hours, peritoneal lavage cells were stained with fluorescently conjugated mAbs against Ly-6G (Becton Dickinson, clone 1A8) and 7/4 (Serotec) in the presence of mAb 2.4G2 (FcgRIIB/III receptor blocker). The absolute number of neutrophil (Ly-6G+ 7/4+) was determined by flow cytometry. Methods Reagents Bafilomycin A1, cytochalasin D and zYVAD-fmk were from Calbiochem. ATP, acridine orange and poly(dA:dT) sodium salt were form Sigma-Aldrich and ultra-pure LPS was purchased from Invivogen. Nigericin, Hoechst dye, DQ ovalbumin and fluorescent choleratoxin B were purchased from Invitrogen. MSU crystals were prepared as described17. Cholesterol crystal preparation Tissue-culture grade or synthetic cholesterol was purchased from Sigma, solubilized in hot acetone and crystallized by cooling. After 6 cycles of recrystallizations, the final crystallization was performed in the presence of 10% endotoxin-free water to obtain hydrated cholesterol crystals. Cholesterol crystals were analyzed for purity by electron impact GC/MS and thin layer chromatography using silica gel and hexane-ethyl acetate (80:20) solvent. Crystal size was varied using a microtube tissue grinder. Fluorescent cholesterol was prepared by addition of the DiD or DiI dyes (Invitrogen) in PBS. ELISA and Western Blot ELISA measurements of IL-1β (Becton Dickinson) and IL-18 (MBL International) were made according to the manufacturer’s directions. Experiments for caspase-1 Western blot analysis were performed in serum-free DMEM medium. After stimulations, cells were lysed by the addition of a 10X lysis buffer (10% NP-40 in TBS and protease inhibitors), and post-nuclear lysates were separated on 4–20% reducing SDS-PAGE. Anti murine caspase-1 pAb was kindly provided by P. Vandenabeele (University of Ghent, The Netherlands). Anti-human cleaved IL-1β (Cell Signaling) from human PBMCs was analyzed in serum-free supernatants as above without cell lysis. Confocal microscopy ApoE−/− mice that were maintained in a pathogen-free facility were fed a Western-type diet (Teklad Adjusted Calories 88137; 21% fat (wt/wt), 0.15% cholesterol (wt/wt) and 19.5% casein (wt/wt); no sodium cholate) starting at 8 weeks of age and continued for 2, 4, 8 or 12 weeks (three mice in each group). Mice were euthanized and hearts were collected as described27. Hearts were serially sectioned at the origins of the aortic valve leaflets, and every third section (5 µm) was stained with hematoxylin and eosin and imaged by light microscopy. Adjacent sections were fixed in 4% paraformaldehyde, blocked and permeabilized (10% goat serum / 0.5% saponin in PBS) and stained with fluorescent primary antibodies against macrophages (MoMa-2, Serotec), DCs (CD11c, Becton Dickinson) or neutrophils (anti-Neutrophil, Serotec) for 1 h at 37 °C for imaging by confocal microscopy. Human atherosclerotic lesions were obtained directly after autopsy, serially sectioned at 2- to 3-mm intervals and frozen sections (5 mm) were prepared as above. Parallel sections were stained with Masson’s trichrome stain. Tissues were prepared for microscopy as above. Macrophages were stained with anti-CD68 (Serotec), smooth muscle cells were visualized with fluorescent phalloidin (Invitrogen). Human and mouse samples were counterstained with Hoechst dye to visualize nuclei. The atherosclerotic lesions were imaged on a Leica SP2 AOBS confocal microscope where immunofluoroscence staining was visualized by standard confocal techniques and crystals were visualized utilizing laser reflection using enhanced transmittance of the acousto-optical beam splitter as described3. Of note, laser reflection and fluorescence emission occurs at the same confocal plane in this setup. The mean lesion area, amount of crystal deposition and monocyte marker presence was quantified from three digitally captured sections per mouse (Photoshop CS4 Extended). For the quantification of crystal mass and macrophages present, the sum of positive pixels (laser reflection or fluorescence, respectively) was determined and the area calculated from the pixel size. Confocal microscopy of mouse macrophages was performed as described3. DQ ovalbumin only fluoresces upon proteolytic processing and marks phagolysosomal compartments in macrophages. Acridine orange lysosomal damage assay This assay was performed by flow cytometry as described3. Bone marrow transplantation and atherosclerosis model Eight weeks-old female LDLR−/− mice were lethally irradiated (11 Gy). Bone marrow was prepared from femurs and tibias of C57BL/6, NLRP3−/−, ASC−/− and IL-1α−/−b−/− donor mice and T cells were depleted using complement (Pel-Freez Biologicals) and anti-Thy1 mAB (M5/49.4.1, ATCC). Irradiated recipient mice were reconstituted with 3.5 × 106 bone marrow cells administered into the tail vein. After 4 weeks, mice were fed with a Western-type diet (Teklad Adjusted Calories 88137; 21% fat [wt/wt], 0.15% cholesterol [wt/wt] and 19.5% casein [wt/wt]; no sodium cholate) for 8 weeks. Mice were euthanized and intracardially perfused with formalin. Hearts were embedded in OTC (Richard-Allen Scientific, Kalamazoo, MI) medium, frozen, and serially sectioned through the aorta from the origins of the aortic valve leaflets and every single section (10 µm) throughout the aortic sinus (800 µm) was collected. Quantification of average lesion area was done from 12 stained with hematoxylin eosin or Giemsa per mouse by two independent investigators with virtually identical results. Serum cholesterol levels were determined by enzymatic assay (Wako Diagnostics), and serum IL-18 was measured by SearchLight protein array technology (Aushon Biosystems, Billerica, MA). Statistical analyses The significance of differences between groups was evaluated by one–way analysis of variance (ANOVA) with Dunnett’s post- comparison test for multiple groups to control group, or by Student’s t test for 2 groups. R squared was calculated from the Pearson correlation coefficient. Analyses were done using Prism (GraphPad Software, Inc.). 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A lysosome-to-nucleus signalling mechanism senses and regulates the lysosome via mTOR and TFEB

Carmine Settembre, Roberto Zoncu, Diego L Medina … (2012)

Introduction The lysosome maintains cellular homeostasis and mediates a variety of physiological processes, including cellular clearance, lipid homeostasis, energy metabolism, plasma membrane repair, bone remodelling, and pathogen defense. All these processes require an adaptive and dynamic response of the lysosome to environmental cues. Indeed, physiologic cues, such as ageing and diet, and pathologic conditions, which include lysosomal storage diseases (LSDs), neurodegenerative diseases, injuries, and infections may generate an adaptive response of the lysosome (Luzio et al, 2007; Ballabio and Gieselmann, 2009; Saftig and Klumperman, 2009). Our understanding of the mechanisms that regulate lysosomal function and underlying lysosomal adaptation is still in an initial phase. A major player in the regulation of lysosomal biogenesis is the basic Helix-Loop-Helix (bHLH) leucine zipper transcription factor, TFEB (Sardiello et al, 2009). Among the identified TFEB transcriptional targets are lysosomal hydrolases that are involved in substrate degradation, lysosomal membrane proteins that mediate the interaction of the lysosome with other cellular structures, and components of the vacuolar H+-ATPase (v-ATPase) complex that participate in lysosomal acidification (Sardiello et al, 2009; Palmieri et al, 2011). TFEB is also a main player in the transcriptional response to starvation and controls autophagy by positively regulating autophagosome formation and autophagosome–lysosome fusion both in vitro and in vivo (Settembre et al, 2011). TFEB activity and its nuclear translocation correlate with its phosphorylation status (Settembre and Ballabio, 2011; Settembre et al, 2011). However, it is still unclear how the cell regulates TFEB activity according to its needs. An intriguing hypothesis is that the lysosome senses the physiological and nutritional status of the cell and conveys this information to the nucleus to drive the activation of feedback gene expression programs. A ‘sensing device', which is responsive to the lysosomal amino acid content and involves both the v-ATPase and the master growth regulator mTOR complex 1 (mTORC1), was recently identified on the lysosomal surface (Zoncu et al, 2011a). The interaction between amino acids and v-ATPase regulates Rag guanosine triphosphatases (GTPases), which in turn activate mTORC1 by translocating it to the lysosomal surface (Sancak et al, 2008, 2010; Zoncu et al, 2011a). According to this mechanism, the lysosome participates in the signalling pathways regulated by mTOR, which controls several cellular biosynthetic and catabolic processes (Zoncu et al, 2011b). We postulated that TFEB uses the v-ATPase/mTORC1 sensing device on the lysosomal surface to modulate lysosomal function according to cellular needs. Consistent with this hypothesis, we found that TFEB interacts with mTOR on the lysosomal membrane and, through this interaction, it senses the lysosomal content. Therefore, TFEB acts both as a sensor of lysosomal state, when on the lysosomal surface, and as an effector of lysosomal function when in the nucleus. This unique lysosome-to-nucleus signalling mechanism allows the lysosome to regulate its own function. Results TFEB responds to the lysosomal status We postulated that TFEB activity was regulated by the physiological status of the lysosome. Therefore, we tested whether disruption of lysosomal function had an impact on TFEB nuclear translocation. TFEB subcellular localization was analysed in HeLa and HEK-293T cells transiently transfected with a TFEB–3 × FLAG plasmid and treated overnight with several inhibitors of lysosomal function. These treatments included the use of chloroquine (CQ), an inhibitor of the lysosomal pH gradient, and Salicylihalamide A (SalA), a selective inhibitor of the v-ATPase (Xie et al, 2004), as well as overexpression of PAT1, an amino acid transporter that causes massive transport of amino acids out of the lysosomal lumen (Sagne et al, 2001). Immunofluorescence analysis showed a striking nuclear accumulation of TFEB–3 × FLAG in treated cells (Figure 1A and B). We repeated this analysis using an antibody detecting the endogenous TFEB (Supplementary Figure S1). Similarly to their effect on exogenously expressed TFEB, both amino acid starvation and lysosomal stress induced nuclear translocation of endogenous TFEB (Figure 1C). These observations were confirmed by immunoblotting performed after nuclear/cytoplasmic fractionation (Figure 1D). Immunoblotting also revealed that TFEB nuclear accumulation was associated with a shift of TFEB–3 × FLAG to a lower molecular weight, suggesting that lysosomal stress may affect TFEB phosphorylation status (Figure 1D). mTORC1 regulates TFEB subcellular localization Based on the observation that mTORC1 resides on the lysosomal membrane and its activity is dependent on both nutrients and lysosomal function (Sancak et al, 2010; Zoncu et al, 2011a), we postulated that the effects of lysosomal stress on TFEB nuclear translocation may be mediated by mTORC1. Consistent with this idea, chloroquine or SalA inhibited mTORC1 activity as measured by level of p-P70S6K, a known mTORC1 substrate (Figure 2A; Zoncu et al, 2011a). The involvement of mTOR appears in contrast with our previous observation that Rapamycin, a known mTOR inhibitor, did not affect TFEB activity. However, recent data indicate that Rapamycin is a partial inhibitor of mTOR, as some substrates are still efficiently phosphorylated in the presence of this drug (Thoreen et al, 2009). Therefore, we used kinase inhibitors Torin 1 and Torin2, which belong to a novel class of molecules that target the mTOR catalytic site, thereby completely inhibiting mTOR activity (Feldman et al, 2009; Garcia-Martinez et al, 2009; Thoreen et al, 2009). We stimulated starved cells, in which TFEB is dephosphorylated and localized to the nucleus, with an amino acid rich medium supplemented with Torin 1 (250 nM), Rapamycin (2.5 μM), or ERK inhibitor U0126 (50 μM). Stimulation of starved cells with nutrients alone induced a significant TFEB molecular weight shift and re-localization to the cytoplasm (Figure 2B). Nutrient stimulation in the presence of the ERK inhibitor U0126 at a concentration of 50 μm induced only a partial TFEB molecular weight shift, suggesting that phosphorylation by ERK partially contributes to TFEB cytoplasmic localization. Treatment with 2.5 μM Rapamycin also resulted in a partial molecular weight shift but did not affect TFEB subcellular localization (Figure 2B), consistent with our previous observations (Settembre et al, 2011). However, Torin 1 (250 nM) treatment entirely prevented the molecular weight shift induced by nutrients and, in turn, resulted in massive TFEB nuclear accumulation. This conclusion is in contrast with a recent study that showed that mTOR-mediated TFEB phosphorylation promoted, rather than inhibited, its nuclear translocation (Pena-Llopis et al, 2011). Instead our data indicate that mTOR is a potent inhibitor of TFEB nuclear translocation and that TFEB is a rapamycin-resistant substrate of mTORC1. In a previous study, we showed that ERK2 phosphorylates TFEB and that starvation and ERK2 inhibition promote TFEB nuclear translocation (Settembre et al, 2011). We tested whether lysosomal stress caused TFEB nuclear translocation also via ERK inhibition. Overnight treatment of HeLa cells with either chloroquine or SalA did not have any effect on ERK activity (Figure 2A), suggesting that mTOR-mediated regulation is predominant. To quantify the effects of ERK and mTOR on TFEB subcellular localization, we developed a cell-based high content assay using stable HeLa cells that overexpress TFEB fused to the green fluorescent protein (TFEB–GFP) (see Materials and methods for details). We tested 10 different concentrations of each inhibitor (U0126, Rapamycin, Torin 1, and Torin 2) ranging from 2.54 nM to 50 μM. Figure 2C and D shows the TFEB nuclear/cytoplasmic distribution for each concentration of each compound in duplicate represented as dose–response curves using a non-linear regression fitting (see Materials and methods for details). Consistent with the above-described data, the most potent compounds that activate TFEB nuclear translocation were Torin 1 (EC50; 147.9 nM) and its analogue Torin 2 (EC50; 1666 nM). ERK inhibitor U0126 showed only a partial effect, while Rapamycin had no effects at any of the concentrations that are routinely used (10 nM–10 μM). Furthermore, Torin 1 treatment potently induced nuclear accumulation of endogenous TFEB in HEK-293T cells (Figure 2E), confirming the observations obtained with the TFEB–GFP construct. As Torin 1 inhibits both mTORC1 and mTORC2 complexes, we next evaluated the contribution of each complex to TFEB regulation. Three main observations suggest that TFEB is predominantly regulated by mTORC1: (1) stimulation of starved cells with amino acids, which activate mTORC1 but not mTORC2, induced an extensive TFEB molecular weight shift, which is highly suggestive of a phosphorylation event (Supplementary Figure S2); (2) knockdown of RagC and RagD, which mediate amino acid signals to mTORC1, caused TFEB nuclear accumulation even in cells kept in full nutrient medium (Figure 2F); (3) in cells with disrupted mTORC2 signalling (Sin1−/− mouse embryonic fibroblasts (MEFs)) (Frias et al, 2006; Jacinto et al, 2006; Yang et al, 2006) TFEB underwent a molecular weight shift and nuclear translocation upon Torin 1 treatment that were similar to control cells (Figure 2G). Together, these data indicate that mTORC1, not mTORC2, regulates TFEB by preventing its nuclear translocation. Finally, co-immunoprecipitation assays in HEK-293T cells expressing TFEB–3 × FLAG showed that TFEB binds both to mTOR and to the mTORC1 subunit raptor but not to the mTORC2 subunits rictor and mSin1, indicating that TFEB and mTORC1 interact both functionally and physically (Figure 2H). mTORC1 controls TFEB subcellular localization via phosphorylation of S142 We previously identified phosphorylation at Serine 142 as a key event for TFEB nuclear translocation during starvation (Settembre et al, 2011). To test whether mTORC1 phosphorylates TFEB at S142, we generated a phosphospecific antibody that recognizes TFEB only when phosphorylated at S142. No signal was detected by this antibody in cells that overexpress the S142A mutant version of TFEB, thus confirming its specificity (Supplementary Figure S3). Using this antibody, we observed that TFEB was no longer phosphorylated at S142 in HeLa cells stably overexpressing TFEB–3 × FLAG and cultured in nutrient-depleted media, consistent with our previous results (Figure 3A). Subsequently, we analysed the levels of S142 phosphorylation in starved cells supplemented with normal media with or without either Torin 1 or Rapamycin. While Torin 1 clearly blunted nutrient-induced S142 phosphorylation, rapamycin did not, suggesting that S142 represents a rapamycin-resistant mTORC1 site (Figure 3A). Indeed, an mTOR kinase assay revealed that mTORC1 phosphorylates highly purified TFEB in vitro with comparable efficiency to other known mTORC1 substrates, and this phosphorylation dropped dramatically when mTORC1 was incubated with the S142A mutant version of TFEB (Figure 3B). These results clearly demonstrate that TFEB is an mTOR substrate and that S142 is a key residue for the phosphorylation of TFEB by mTOR. Recent findings suggest that mTORC1 phosphorylates its target proteins at multiple sites (Hsu et al, 2011; Peterson et al, 2011; Yu et al, 2011). To identify additional serine residues that may be phosphorylated by mTOR, we searched for consensus phosphoacceptor motif for mTORC1 (Hsu et al, 2011) in the coding sequence of TFEB (Figure 3C and D). We mutagenized all TFEB amino acid residues that were putative mTORC1 targets into alanines. We then tested the effects of each of these mutations on TFEB subcellular localization and found that, similarly to S142A, a serine-to-alanine mutation at position 211 (S211A) resulted in a constitutive nuclear localization of TFEB (Figure 3E). Mutants for the other serine residues behaved similarly to wild-type TFEB (Figure 3E; Supplementary Figure S4; Settembre et al, 2011). Together, these data indicate that, in addition to S142, S211 also plays a role in controlling TFEB subcellular localization and suggest that S211 represents an additional target site of mTORC1. mTORC1 and TFEB interact on the lysosomal surface Based on the observations that TFEB is a substrate for mTORC1 (Figure 3A and B) and that the two proteins physically interact (Figure 2H), we tested whether the interaction of TFEB and mTORC1 occurs on the lysosomal membrane. Careful examination of HeLa cells that express TFEB–GFP showed that, while under normal growth conditions the majority of cells displayed a predominantly cytoplasmic TFEB localization, a subset of cells showed clearly discernible intracellular puncta of TFEB–GFP fluorescence, suggesting a lysosomal localization (Supplementary Figure S5). These observations were confirmed in MEFs that transiently express TFEB–GFP along with the late endosomal/lysosomal marker mRFP–Rab7 (Figure 4A). In a subset of cells, TFEB–GFP clearly colocalized with mRFP–Rab7-positive lysosomes and this association persisted over time as lysosomes trafficked inside the cell (Figure 4A and B; Supplementary Movie S1). We reasoned that the partial localization of TFEB to lysosomes may be due to a transient binding to mTORC1, followed by mTORC1-dependent phosphorylation and translocation of TFEB to the cytoplasm. To test this idea, we treated TFEB–GFP HeLa cells with Torin 1, as a way to ‘trap' TFEB in its bound state to inactive mTORC1. Confirming our hypothesis, Torin 1 caused a massive and dramatic accumulation of TFEB–GFP on lysosomes (Supplementary Figure S5). Similarly, Torin 1 treatment of MEFs resulted in a time-dependent accumulation of TFEB–GFP on lysosomes within minutes of drug delivery, followed by a more gradual accumulation into the nucleus (Figure 4C; Supplementary Movies S2 and S3). Interestingly, we also noticed that Torin 1 treatment caused a significant accumulation of endogenous mTOR on lysosomes compared with untreated cells (Figure 4D). Thus, two mechanisms contribute to clustering of TFEB on lysosomes upon Torin 1 treatment: (1) trapping of the mTORC1–TFEB complex in the inactive state and (2) increase of the amount of mTORC1 bound to the lysosomal surface. The accumulation of inactive mTORC1 on the lysosomal surface may reflect a feedback mechanism through which mTORC1 regulates its own targeting to lysosomal membranes via its kinase activity (Zoncu et al, 2011b). To investigate the lysosomal trapping of TFEB in a dynamic and quantitative way, we performed Fluorescence Recovery After Photobleaching (FRAP) experiments on TFEB–GFP-positive lysosomes (Figure 4E and F; Supplementary Movie S4). In control cells, photobleaching of TFEB–GFP-positive lysosomes was followed by a rapid (t 1/2=0.35 min) and substantial (60%) recovery of the initial fluorescence. Conversely, in Torin 1-treated cells, where TFEB–GFP-positive lysosomes were much more prominent and numerous, the fluorescence recovery was slower (t 1/2=0.57 min) and smaller (30% recovery of the initial fluorescence). Thus, a large fraction of TFEB was trapped onto the lysosomal surface through binding to inactive mTORC1 and was no longer able to exchange with the cytoplasm. In conclusion, these data indicate that TFEB and mTORC1 bind to each other on the lysosomal surface, where phosphorylation of TFEB by mTORC1 occurs. mTORC1 regulates TFEB via the Rag GTPases The observation that TFEB is regulated by mTORC1 prompted us to determine whether the activation state of the Rag GTPases, which together with the v-ATPase mediate mTORC1 activation by amino acids, played a role in the control of TFEB subcellular localization. Point mutants of the Rags are available, which fully mimic either the presence of amino acids (‘RagsCA') or their absence (‘RagsDN') (Sancak et al, 2008). We took advantage of these mutants to directly test the requirement for mTORC1 in sequestering TFEB to the lysosome and we asked whether the RagsDN mutants, which cause loss of mTORC1 from the lysosomal surface (Sancak et al, 2010), were able to suppress Torin 1-induced lysosomal accumulation of TFEB as well as TFEB-mTORC1 binding. In co-immunoprecipitation assays, Torin 1 clearly boosted the binding of both raptor and mTOR to TFEB–3 × FLAG (Figure 4G). However, co-expression of the RagsDN mutants reduced the binding of TFEB–3 × FLAG to mTORC1 components down to background levels, both in control and in Torin 1-treated cells (Figure 4G). Consistent with these results, immunofluorescence experiments in HEK-293T co-expressing TFEB–3 × FLAG and the RagsDN mutants showed that TFEB failed to cluster on lysosomes following Torin 1 treatment (Figure 4H). Together, these data strongly suggest that TFEB and mTORC1 only interact when they are both found on the lysosomal surface. Next, we tested whether the activation status of the Rags controlled TFEB nuclear translocation. In HEK-293T cells that co-express TFEB–3 × FLAG and a control small GTPase (Rap2A), amino acid withdrawal caused a massive translocation of TFEB to the nucleus (Figure 5A and D), as previously reported (Settembre et al, 2011). Consistent with mTORC1 re-activation, a brief (20 min) re-stimulation of starved cells with amino acids drove TFEB out of the nucleus in the majority of cells (Figure 5A). In contrast, in cells that co-express both TFEB–3 × FLAG and the RagsCA mutants, TFEB localization was always and completely cytoplasmic, regardless of the nutrient state of the cells (Figure 5B and D). Finally, in cells that co-express both TFEB and the RagsDN mutants, TFEB was almost exclusively found in the nucleus and did not translocate to the cytoplasm upon amino acid stimulation (Figure 5C and D). Thus, the activation state of the Rags completely overrides the nutritional status of the cells and is sufficient to determine TFEB localization. It was previously shown that the RagsCA rescue the inhibitory effect of various lysosomal stressors on mTORC1 activation (Zoncu et al, 2011a). Thus, we asked whether the RagsCA mutants were able to prevent the TFEB nuclear translocation promoted by these stressors (Figures 1A–D and 5E). In cells that co-express both TFEB–3 × FLAG and the RagsCA mutants, TFEB remained entirely cytoplasmic upon treatment with Chloroquine and SalA (Figure 5F and G), while it was nuclear in the vast majority of cells that express a control GTPase and were subject to the same drug treatments (Figure 5E and G). Importantly, treatment of cells co-expressing TFEB–GFP and RagCA with Torin 1 reverted the RagCA-induced cytoplasmic localization of TFEB and massively drove TFEB to the nucleus, further demonstrating that the action of the Rag mutants on TFEB is mediated by mTORC1 (Supplementary Figure S6). In summary, these results demonstrate that TFEB localization is directly regulated by the amino acid-mTORC1 signalling pathway via the activation state of Rag GTPases. The lysosome regulates gene expression via TFEB As the interaction of TFEB with mTORC1 on the lysosomal membrane controls TFEB nuclear translocation, we tested whether the ability of TFEB to regulate gene expression was also influenced by this interaction. The expression of several lysosomal/autophagic genes that were shown to be targets of TFEB (Palmieri et al, 2011) was tested in primary hepatocytes from a conditional knockout mouse line in which TFEB was deleted in the liver (Tcfebflox/flox; alb-CRE), and in a control mouse line (Tcfebflox/flox). Cells were treated with either chloroquine or Torin 1, or left untreated. These treatments inhibited mTOR as measured by the level of p-S6K, whereas the levels of p-ERK were unaffected (Figure 6A). Primary hepatocytes isolated from TFEB conditional knockout mice cultured in regular medium did not show significant differences in the expression levels of several TFEB target genes compared with control hepatocytes (Supplementary Figure S7). However, while the expression of TFEB target genes was upregulated in hepatocytes from control mice after treatment with chloroquine, this upregulation was significantly blunted in hepatocytes from TFEB conditional knockout mice (Figure 6B). Similarly, the transcriptional response upon Torin 1 treatment was significantly reduced in hepatocytes from TFEB conditional knockout mice (Figure 6C). Together, these results indicate that TFEB plays a key role in the transcriptional response induced by the lysosome via mTOR. Discussion Our study demonstrates that TFEB, a master gene for lysosomal biogenesis, is regulated by the lysosome via the mTOR pathway. mTORC1 and TFEB meet on the lysosomal membrane where mTORC1 phosphorylates TFEB. We previously reported that ERK2 phosphorylates TFEB and, in cells treated with an MEK inhibitor, the TFEB nuclear fraction was increased (Settembre et al, 2011). In the same study, we reported that the mTOR inhibitor rapamycin had little or no effects on TFEB subcellular localization. Here, we compared three different types of kinase inhibitors—MEK inhibitor U0126 and mTOR inhibitors rapamycin, Torin 1, and Torin 2—in their ability to cause a shift in TFEB molecular weight and to induce TFEB nuclear translocation. As shown in Figure 2, Torin 1 and Torin 2 induced TFEB nuclear translocation more efficiently compared to U0126. The more pronounced shift of TFEB molecular weight, which was observed in cells treated with Torin 1, suggests that mTORC1 induces TFEB phosphorylation at multiple sites, either directly or indirectly. In a recent high throughput mass spectrometry study, TFEB was predicted to be phosphorylated at 11 different residues, thus suggesting a complex regulation of its activity with several phosphorylation sites potentially involved (Dephoure et al, 2008). Here, we have used an mTORC1 in-vitro kinase assay and a phosphoantibody to demonstrate that serine S142, which we previously found to be phosphorylated by ERK2, is also phosphorylated by mTOR and that this phosphorylation has a crucial role in controlling TFEB subcellular localization and activity. In addition, we have mutated 12 different serines, which were candidate mTOR phosphorylation sites, into alanines, thus abolishing the corresponding TFEB phosphorylation sites. Testing the effects of each of these mutations on TFEB subcellular localization led to the identification of an additional residue, serine S211, which plays a role in TFEB subcellular localization, confirming the predicted complexity of TFEB regulation by phosphorylation. Phosphorylation of TFEB by mTOR had already been reported in a previous study (Pena-Llopis et al, 2011). However, in that study the authors concluded that mTOR promoted, rather than inhibited, TFEB activity. Several lines of evidence indicate that mTOR inhibits TFEB activity. First, TFEB is entirely nuclear when cells are either starved or treated with Torin1, both conditions in which mTOR activity is profoundly inhibited. Second, treatment of starved cells with nutrients, a condition that boosts mTORC1 activity, resulted in TFEB cytoplasmic accumulation, with TFEB being undetectable in the nuclear fraction. Third, treatment with drugs such as chloroquine or SalA, which inhibit mTORC1 function, induced TFEB nuclear accumulation. Fourth, transfection of mutant Rag proteins that inhibit mTORC1 resulted in nuclear accumulation of TFEB and, conversely, mutant Rags that constitutively activate mTORC1 prevented TFEB nuclear accumulation upon starvation, chloroquine and SalA treatment. Fifth, TFEB is in the nucleus in its low-phosphorylated form, an observation that is consistent with a model in which inhibition, rather than activation, of a kinase induces TFEB nuclear translocation. It is difficult to explain the discrepancy between our observations and those reported by Pena-Llopis et al. We considered the possibility that the TSC2-deficient cells that were used in that study may behave differently to other cellular systems in the assays performed. To test this possibility, we analysed TFEB regulation by amino acids, chloroquine and Torin 1 in TSC2−/− cells but obtained the same results that we observed in other cell types both on exogenous TFEB–GFP and on endogenous TFEB (Supplementary Figures S8 and S9, respectively). Our data indicate that mTORC1 negatively regulates TFEB via the amino acid/Rag GTPase pathway. The phosphorylation status of TFEB and its subcellular localization were entirely determined by the activation state of the Rag GTPases, which regulate mTORC1 activity downstream of amino acids (Kim et al, 2008; Sancak et al, 2008). In particular, constitutively active Rags rescued nuclear translocation of TFEB caused by starvation and lysosomal stress, while inactive Rag mutants caused TFEB to accumulate in the nucleus even in fully fed cells. These results imply that, among the many regulatory inputs to mTORC1, the amino acid pathway is particularly important in controlling TFEB activity and plays not only a permissive but also an instructive role. This idea is further supported by our observation that constitutive activation of the growth factor inputs to mTORC1 that occurs in TSC2−/− cells cannot prevent TFEB nuclear accumulation caused by starvation and lysosomal stress. Future work will be required to address how each upstream input to mTOR contributes to TFEB regulation. Nonetheless, compounded with recent evidence showing that amino acid sensing by the v-ATPase/Rag GTPase/mTORC1 may begin in the lysosomal lumen (Zoncu et al, 2011a) our findings substantiate the role of TFEB as the end point of a lysosome-sensing and signalling pathway. Our data shed light into the logic that underlies the control of TFEB localization. In fully fed cells, a fraction of TFEB could always be found on lysosomes, although the majority appeared to freely diffuse in the cytoplasm. The lysosomal localization of TFEB is associated with its ability to physically bind mTORC1, as shown by co-immunoprecipitation assays. Moreover, time-lapse analysis of TFEB–GFP in cells treated with Torin 1 showed that TFEB clustered on lysosomes shortly after the onset of drug treatment, and then progressively appeared in the nucleus (Supplementary Movies S2 and S3). Together, these results suggest the following model of control of TFEB subcellular localization and activity (Figure 7). At any given time, a fraction of TFEB rapidly and transiently binds to the lysosomal surface, where it is phosphorylated by mTORC1 and thus kept in the cytoplasm. Nutrient withdrawal, v-ATPase inhibition, and lysosomal stress inactivate the Rag GTPases, causing loss of mTORC1 from the lysosome and resulting in failure to re-phosphorylate TFEB. Unphosphorylated TFEB progressively accumulates in the nucleus, where it activates lysosomal gene expression programs aimed at correcting the defective nutrient and/or pH status of the lysosome. In this model, the lysosome represents a bottleneck where mTORC1 tightly regulates the amount of TFEB that is allowed to reach the nucleus. mTORC1 may regulate a yet undiscovered TFEB function at the lysosome. This possibility is supported by the observation that blocking mTORC1 activity with Torin 1 resulted in a dramatic accumulation of TFEB not only in the nucleus but also on lysosomes, which was visible as increased binding to mTORC1 in co-IP assays, as well as reduced mobility in FRAP experiments. Future work will address what function, if any, TFEB performs on the lysosomal surface. Interestingly, recent evidence indicating that TFEB regulates multiple aspects of lysosomal dynamics, including the propensity of lysosomes to fuse with the plasma membrane (Medina et al, 2011), suggests that the range of biological functions of TFEB still needs to be fully elucidated. Our data further emphasize the emerging role of the lysosome as a key signalling centre. In particular, a molecular machinery that connects the presence of amino acids in the lysosomal lumen to the activation of mTORC1 indicates a new role for the lysosome in nutrient sensing and cellular growth control (Rabinowitz and White, 2010; Singh and Cuervo, 2011; Zoncu et al, 2011a). It also suggests that mTORC1 participates in a lysosomal adaptation mechanism that enables cells to cope with starvation and lysosomal stress conditions (Yu et al, 2010). This mechanism responds to a wide range of signals that relay the metabolic state of the cell, as well as the presence of various stress stimuli. For instance, loss of lysosomal proton gradient, caused by either energy depletion or pathological conditions, may suppress mTORC1 activity, either by blocking the proton-coupled transport of nutrients to and from the lysosome, or by directly affecting the v-ATPase (Marshansky, 2007). Similarly, lysosomal membrane permeabilization observed in certain LSDs and neurodegenerative diseases may result in nutrient leakage and suppression of mTORC1 (Dehay et al, 2010; Kirkegaard et al, 2010). We found that the transcriptional response of lysosomal and autophagy genes to starvation and mTOR inhibition by Torin 1 was hampered in hepatocytes from mice carrying a liver-specific conditional knockout of TFEB, demonstrating that TFEB is a main mediator of this response. Therefore, TFEB translates a lysosomal signal into a transcriptional program. This lysosome-to-nucleus signalling mechanism, which operates a feedback regulation of lysosomal function, presents intriguing parallels with the sterol sensing pathway in the endoplasmic reticulum, where cholesterol depletion and ER stress cause the nuclear translocation of the Sterol Responsive Element Binding Protein (SREBP) transcription factor, which then activates gene expression programs that enhance cholesterol synthesis and ER function (Wang et al, 1994; Peterson et al, 2011). Another example is represented by the mitochondria retrograde signalling pathway, in which mitochondrial dysfunction activates factors such as NFκB, NFAT, and ATF, through altered Ca2+ dynamics (Butow and Avadhani, 2004). Finally, as TFEB overexpression was able to promote substrate clearance and to rescue cellular vacuolization in LSDs (Medina et al, 2011), the identification of a lysosome-based, mTOR-mediated, mechanism that regulates TFEB activity offers a new tool to promote cellular clearance in health and disease. Materials and methods Cell culture HeLa and HEK-293T cells were purchased from ATCC and cultured in DMEM supplemented with 10% fetal calf serum, 200 mM L-glutamine, 100 mM sodium pyruvate, penicillin 100 units/ml, streptomycin 100 mg/ml, 5% CO2 at 37°C. Primary hepatocytes were generated as follow: 2-month-old mice were deeply anaesthetized with Avertin (240 mg/kg) and perfused first with 25 ml of HBSS (Sigma H6648) supplemented with 10 mM HEPES and 0.5 mM EGTA and after with a similar solution containing 100 U/ml of Collagenase (Wako) and 0.05 mg/ml of Trypsin inhibitor (Sigma). Liver was dissociated in a petri dish, cell pellet was washed in HBSS and plated at density of 5 × 105 cells/35 mm dish and cultured in William's medium E supplemented with 10% FBS, 2 mM glutamine, 0.1 μM Insulin, 1 μM Dexamethasone and pen/strep. The next day, cells were treated as described in the text. Sin1−/− and control MEFs were generated as previously described (Jacinto et al, 2006) and maintained in DMEM supplemented with 10% FBS, glutamine and pen/strep. TSC2+/+ p53−/− and TSC2−/− p53−/− MEFs, kindly provided by David Kwiatkowski (Harvard Medical School), were maintained in DMEM supplemented with 10% heat-inactivated FBS, glutamine and pen/strep. Generation of a Tcfebflox mouse line We used publicly available embryonic stem (ES) cell clones (http://www.eucomm.org/) in which Tcfeb was targeted by homologous recombination at exons 4 and 5. The recombinant ES cell clones were injected into blastocysts, which were used to generate a mouse line carrying the engineered allele. Liver-specific KO was generated crossing the Flox/Flox mice with a transgenic line expressing the CRE under the Albumin promoter (ALB-CRE) obtained from the Jackson laboratory. All procedures involving mice were approved by the Institutional Animal Care and Use Committee of the Baylor College of Medicine. Plasmids and cell transfection Cells were transiently transfected with DNA plasmids pRK5-mycPAT1, pRK5-HAGST-Rap2A, pRK5-HAGST-RagB and its Q99L (CA) and T54N (DN) mutants, pRK5-HAGST-RagD and its Q121L (DN) and S77L (CA) mutants; pTFEB-GFP, and pCMV-TFEB-3 × FLAG using lipofectamine2000 or LTX (Invitrogen) according to the protocol from manufacturer. Site-direct mutagenesis was performed according to the manufacturer instructions (Stratagene) verifying the correct mutagenesis by sequencing. Drugs and cellular treatments The following drugs were used: Rapamycin (2.5 μM, otherwise indicated) from Sigma; Torin 1 and Torin 2 (250 nM, otherwise indicated) from Biomarine; U0126 (50 μM) from Cell Signaling Technology; Chloroquine (100 μM) from Sigma; Salicylihalamide A (2 μM) was a kind gift from Jeff De Brabander (UT Southwestern). Immunoblotting and antibodies The mouse anti-TFEB monoclonal antibody was purchased from My Biosource catalogue No. MBS120432. To generate anti-pS142 specific antibodies, rabbits were immunized with the following peptide coupled to KLH: AGNSAPN{pSer}PMAMLHIC. Following the fourth immunization, rabbits were sacrificed and the serum was collected. Non-phosphospecific antibodies were depleted from the serum by circulation through a column containing the non-phosphorylated antigene. The phosphospecific antibodies were next affinity purified using a column containing the phosphorylated peptide. Cells were lysed with M-PER buffer (Thermo) containing protease and phosphatase inhibitors (Sigma); nuclear/cytosolic fractions were isolated as previously described (Settembre et al, 2011). Proteins were separated by SDS–PAGE (Invitrogen; reduced NuPAGE 4–12% Bis-tris Gel, MES SDS buffer). If needed, the gel was stained using 20 ml Imperial Protein Stain (Thermo Fisher) at room temperature for 1 h and de-stained with water. Immunoblotting analysis was performed by transferring the protein onto a nitrocellulose membrane with an I-Blot (Invitrogen). The membrane was blocked with 5% non-fat milk in TBS-T buffer (TBS containing 0.05% Tween-20) and incubated with primary antibodies anti-FLAG and anti-TUBULIN (Sigma; 1:2000), anti-H3 (Cell Signaling; 1:10 000) at room temperature for 2 h whereas the following antibodies were incubated ON in 5% BSA: anti-TFEB (My Biosource; 1:1000), anti-P TFEB (1:1000) ERK1/2, p-ERK1/2, p-P70S6K, P70S6K (Cell Signaling; 1:1000). The membrane was washed three times with TBS-T buffer and incubated with alkaline phosphatase-conjugated IgG (Promega; 0.2 mg/ml) at room temperature for 1 h. The membrane was washed three times with TBS buffer and the expressed proteins were visualized by adding 10 ml Western Blue® Stabilized Substrate (Promega). In-vitro kinase assays FLAG–S6K1, TFEB–3 × FLAG, and TFEBS142A–3 × FLAG were purified from transiently transfected HEK-293T cells treated with 250 nM Torin 1 for 1 h and lysed in RIPA lysis buffer. The cleared lysates were incubated with FLAG affinity beads (Sigma) for 2 h, washed four times in RIPA containing 500 mM NaCl, and eluted for 1 h at 4°C using a competing FLAG peptide. mTORC1 was purified from HEK-293T cells stably expressing FLAG raptor in 0.3% CHAPS using FLAG affinity beads. Kinase assays were preincubated for 10 min at 4°C before addition of ATP, and then for 30 min at 30°C in a final volume of 25 μl consisting of kinase buffer (25 mM HEPES, pH 7.4, 50 mM KCl, 10 mM MgCl2) active mTORC1, 250–500 nM substrate, 50 μM ATP, 1 μCi [γ-32P]ATP, and when indicated 250 nM Torin 1. Reactions were stopped by the addition of 6 μl of sample buffer, boiled for 5 min, and analysed by SDS–PAGE followed by autoradiography. Immunoprecipitation assays HEK-293T cells that express FLAG-tagged proteins were rinsed once with ice-cold PBS and lysed in ice-cold lysis buffer (150 mM NaCl, 40 mM HEPES (pH 7.4), 2 mM EGTA, 2.5 mM MgCl2, 0.3% CHAPS, and one tablet of EDTA-free protease inhibitors (Roche) per 25 ml). The soluble fractions from cell lysates were isolated by centrifugation at 13 000 r.p.m. for 10 min in a microfuge. For immunoprecipitations, 35 μl of a 50% slurry of anti-FLAG affinity gel (Sigma) was added to each lysate and incubated with rotation for 2–3 h at 4°C. Immunoprecipitates were washed three times with lysis buffer. Immunoprecipitated proteins were denatured by the addition of 35 μl of sample buffer and boiling for 5 min, resolved by 8–16% SDS–PAGE, and analysed by immunoblotting. Immunofluorescence assays on HEK-293T cells HEK-293T cells were plated on fibronectin-coated glass coverslips in 35 mm tissue culture dishes, at 300 000 cells/dish. In all, 12–16 h later, cells were transfected with 100 ng of TFEB–3 × FLAG, along with 200 ng Rap2A or Rag GTPase mutants. The next day, cells were subjected to drug treatments or starvation, rinsed with PBS once and fixed for 15 min with 4% paraformaldehyde in PBS at RT. The slides were rinsed twice with PBS and cells were permeabilized with 0.05% Triton X-100 in PBS for 5 min. After rinsing twice with PBS, the slides were incubated with primary antibody in 5% normal donkey serum for 1 h at room temperature, rinsed four times with PBS, and incubated with secondary antibodies produced in donkey (diluted 1:1000 in 5% normal donkey serum) for 45 min at room temperature in the dark, washed four times with PBS. Slides were mounted on glass coverslips using Vectashield (Vector Laboratories) and imaged on a spinning disk confocal system (Perkin-Elmer). High content nuclear translocation assay TFEB–GFP cells were seeded in 384-well plates, incubated for 12 h, and treated with 10 different concentrations of ERK inhibitor U0126 (Sigma-Aldrich) and mTOR inhibitors Rapamycin (Sigma-Aldrich), Torin 1 (Biomarin), and Torin 2 (Biomarin), ranging from 2.54 nM to 50 μM. After 3 h at 37°C in RPMI medium, cells were washed, fixed, and stained with DAPI. For the acquisition of the images, 10 pictures per each well of the 384-well plate were taken by using confocal automated microscopy (Opera high content system; Perkin-Elmer). A dedicated script was developed to perform the analysis of TFEB localization on the different images (Acapella software; Perkin-Elmer). The script calculates the ratio value resulting from the average intensity of nuclear TFEB–GFP fluorescence divided by the average of the cytosolic intensity of TFEB–GFP fluorescence. The results were normalized using negative (RPMI medium) and positive (HBSS starvation) control samples in the same plate. The data are represented by the percentage of nuclear translocation at the different concentrations of each compound using Prism software (GraphPad software). The EC50 for each compound was calculated using non-linear regression fitting (Prism software). Live cell imaging and photobleaching protocol MEFs were transiently transfected with TFEB–GFP and mRFP–Rab7 by nucleofection (Lonza). Cells were plated on glass bottom 35 mm dishes (MatTek Corp.) at a density of 300 000 cells/dish. The next day, cells were transferred to a physiological imaging buffer (130 mM NaCl, 5 mM KCl, 2.5 mM CaCl2, 2.5 mM MgCl2, 25 mM HEPES) supplemented with 5 mM glucose and imaged on a spinning disk confocal microscope (Andor Technology) with a 488-nm and a 561-nm laser through a × 63 objective. To achieve photobleaching of individual TFEB–GFP-positive lysosomes, areas of interest were drawn around selected spots, and movie acquisition was started. Sixty seconds later, the spots were photobleached with a high power (50 mM) 488 nm pulse (100 μs/pixel illumination) using the Andor FRAPPA unit. FRAP analysis The fluorescence recovery of photobleached TFEB-GFP-positive lysosomes was analysed using custom-written plugins in ImageJ (National Institutes of Health). Circular areas of interest were drawn around the spots to be analysed, and the integrated fluorescence within these areas was measured throughout the movie. Fluorescence intensity traces from 5 to 10 spots per condition were normalized to the initial value and time aligned, and their mean and s.d. were calculated using Microsoft Excel. Final plots and curve fitting were made with Prism (GraphPad). RNA extraction, quantitative PCR, and statistical analysis Total RNA was extracted from cells using TRIzol (Invitrogen). Reverse transcription was performed using TaqMan reverse transcription reagents (Applied Biosystems). Lysosomal and autophagic gene-specific primers were previously reported (Settembre et al, 2011). Fold change values were calculated using the DDCt method. Briefly, GAPDH and Cyclophillin were used as ‘normalizer' genes to calculate the DCt value. Next, the DDCt value was calculated between the ‘control' group and the ‘experimental' group. Lastly, the fold change was calculated using 2(-DDCt). Biological replicates were grouped in the calculation of the fold change values. Unpaired T-Test was used to calculate statistical significance. Asterisks in the graph indicate that the P-value was <0.05. mTORC1 phosphosite prediction In order to identify possible phosphosites that may be targeted by mTORC1, we developed a simple method that quantifies the agreement between regions around serine or threonine sites in TFEB and the mTORC1 phosphorylation motif (Hsu et al, 2011). The method calculates the score according to a position-specific score matrix for an amino acid at given distance from the phosphosite of interest. The position starts from −5 and runs to +4. The phosphosite is set at position 0. If there is another serine or threonine in this interval, that residue's score is skipped in the sum. We used MyDomains tool in prosite/expasy.org to sketch the functional domains of TFEB. Domain information was retrieved from UniProt/SwissProt database. Human TFEB and its orthologue sequences were aligned by ClustalW (version 2.0.12), using the default parameters. Supplementary Material Supplementary Movie 1 Supplementary Movie 2 Supplementary Movie 3 Supplementary Movie 4 Supplementary Information Review Process File

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Trehalose, a novel mTOR-independent autophagy enhancer, accelerates the clearance of mutant huntingtin and alpha-synuclein.

Sovan Sarkar, Janet Davies, Zebo Huang … (2007)

Trehalose, a disaccharide present in many non-mammalian species, protects cells against various environmental stresses. Whereas some of the protective effects may be explained by its chemical chaperone properties, its actions are largely unknown. Here we report a novel function of trehalose as an mTOR-independent autophagy activator. Trehalose-induced autophagy enhanced the clearance of autophagy substrates like mutant huntingtin and the A30P and A53T mutants of alpha-synuclein, associated with Huntington disease (HD) and Parkinson disease (PD), respectively. Furthermore, trehalose and mTOR inhibition by rapamycin together exerted an additive effect on the clearance of these aggregate-prone proteins because of increased autophagic activity. By inducing autophagy, we showed that trehalose also protects cells against subsequent pro-apoptotic insults via the mitochondrial pathway. The dual protective properties of trehalose (as an inducer of autophagy and chemical chaperone) and the combinatorial strategy with rapamycin may be relevant to the treatment of HD and related diseases, where the mutant proteins are autophagy substrates.

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Author and article information

Journal

Journal ID (nlm-ta): Nat Commun

Journal ID (iso-abbrev): Nat Commun

Title: Nature Communications

Publisher: Nature Publishing Group

ISSN (Electronic): 2041-1723

Publication date (Electronic): 07 June 2017

Publication date Collection: 2017

Volume: 8

Electronic Location Identifier: 15750

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[1 ]Department of Medicine, Cardiovascular Division, Washington University School of Medicine , Campus Box 8086, 660 S Euclid Avenue, St Louis, Missouri 63110, USA

[2 ]Department of Medicine, Division of Endocrinology, Metabolism, and Lipid Research, Washington University School of Medicine , 660S Euclid Avenue, St Louis, Missouri 63110, USA

[3 ]Telethon Institute of Genetics and Medicine , Via Campi Flegrei 34, 80078 Pozzuoli, Naples, Italy

[4 ]Department of Pathology and Immunology, Washington University School of Medicine , Campus Box 8086, 660S. Euclid Avenue, St Louis, Missouri 63110, USA

[5 ]Peter Munk Cardiac Center, University Health Network , Toronto, Ontario, Canada M5G 2C4

[6 ]Department of Neurology, Washington University School of Medicine , Campus Box 8111, 660S. Euclid Avenue, St Louis, Missouri 63110, USA

[7 ]Department of Cell Biology and Anatomy, University of South Carolina School of Medicine , Columbia, South Carolina 29209, USA

[8 ]Department of Surgery, Washington University School of Medicine , 660S. Euclid Avenue, St Louis, Missouri 63110, USA

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[a ] brazani@ 123456im.wustl.edu

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Babak Razani http://orcid.org/0000-0002-7172-9240

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Publisher Item ID: ncomms15750

DOI: 10.1038/ncomms15750

PMC ID: 5467270

PubMed ID: 28589926

SO-VID: 92fd2038-fbd3-47e7-b19b-fbab7fb3c46c

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Open Access This article is licensed under a Creative Commons Attribution 4.0 International License, which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license, and indicate if changes were made. The images or other third party material in this article are included in the article's Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the article's Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder. To view a copy of this license, visit http://creativecommons.org/licenses/by/4.0/

History

Date received : 11 October 2016

Date accepted : 25 April 2017

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