Recent insights into the function of autophagy in cancer

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In this review, Amaravadi et al. discuss recent developments in the role of autophagy in cancer, in particular how autophagy can promote cancer through suppressing p53 and preventing energy crisis, cell death, senescence, and an anti-tumor immune response.

Abstract

Macroautophagy (referred to here as autophagy) is induced by starvation to capture and degrade intracellular proteins and organelles in lysosomes, which recycles intracellular components to sustain metabolism and survival. Autophagy also plays a major homeostatic role in controlling protein and organelle quality and quantity. Dysfunctional autophagy contributes to many diseases. In cancer, autophagy can be neutral, tumor-suppressive, or tumor-promoting in different contexts. Large-scale genomic analysis of human cancers indicates that the loss or mutation of core autophagy genes is uncommon, whereas oncogenic events that activate autophagy and lysosomal biogenesis have been identified. Autophagic flux, however, is difficult to measure in human tumor samples, making functional assessment of autophagy problematic in a clinical setting. Autophagy impacts cellular metabolism, the proteome, and organelle numbers and quality, which alter cell functions in diverse ways. Moreover, autophagy influences the interaction between the tumor and the host by promoting stress adaptation and suppressing activation of innate and adaptive immune responses. Additionally, autophagy can promote a cross-talk between the tumor and the stroma, which can support tumor growth, particularly in a nutrient-limited microenvironment. Thus, the role of autophagy in cancer is determined by nutrient availability, microenvironment stress, and the presence of an immune system. Here we discuss recent developments in the role of autophagy in cancer, in particular how autophagy can promote cancer through suppressing p53 and preventing energy crisis, cell death, senescence, and an anti-tumor immune response.

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Autophagy: renovation of cells and tissues.

Noboru Mizushima, Masaaki Komatsu (2011)

Autophagy is the major intracellular degradation system by which cytoplasmic materials are delivered to and degraded in the lysosome. However, the purpose of autophagy is not the simple elimination of materials, but instead, autophagy serves as a dynamic recycling system that produces new building blocks and energy for cellular renovation and homeostasis. Here we provide a multidisciplinary review of our current understanding of autophagy's role in metabolic adaptation, intracellular quality control, and renovation during development and differentiation. We also explore how recent mouse models in combination with advances in human genetics are providing key insights into how the impairment or activation of autophagy contributes to pathogenesis of diverse diseases, from neurodegenerative diseases such as Parkinson disease to inflammatory disorders such as Crohn disease. Copyright © 2011 Elsevier Inc. All rights reserved.

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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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The ubiquitin kinase PINK1 recruits autophagy receptors to induce mitophagy

Michael Lazarou, Danielle Sliter, Lesley A Kane … (2015)

Protein aggregates and damaged organelles are tagged with ubiquitin chains to trigger selective autophagy. To initiate mitophagy, PINK1 phosphorylates ubiquitin to activate Parkin, which builds ubiquitin chains on mitochondrial outer membrane proteins where they act to recruit autophagy receptors. Using genome editing to knock out five autophagy receptors, we find that two previously linked to xenophagy, NDP52 and Optineurin, are the primary receptors for PINK1/Parkin-mediated mitophagy. The ubiquitin kinase PINK1 recruits NDP52 and Optineurin, but not p62, to mitochondria to directly activate mitophagy independent of Parkin. Once recruited to mitochondria, NDP52 and Optineurin recruit ULK1, DFCP1 and WIPI1 to focal spots proximal to mitochondria revealing a function for these autophagy receptors upstream of LC3. This supports a new model that PINK1 generated phospho-ubiquitin serves as the autophagy signal on mitochondria and that Parkin amplifies it. This work also suggests direct and broader roles for ubiquitin phosphorylation in other autophagy pathways. Selective autophagy clears intracellular pathogens and mediates cellular quality control by engulfing cargo into autophagosomes and delivering it to lysosomes for degradation. Autophagy receptors bind ubiquitinated cargo and LC3-coated phagophores to mediate autophagy 1,2 . Damaged mitochondria are removed by autophagy following activation of the kinase PINK1 and the E3 ubiquitin ligase Parkin 3,4 . Upon loss of mitochondrial membrane potential or accumulation of misfolded proteins, PINK1 is stabilized on the outer mitochondrial membrane 3 , where it phosphorylates ubiquitin at Ser65 to activate Parkin ubiquitin ligase activity 5–7 . Although the autophagy receptors p62 and Optineurin (OPTN) have been shown to bind ubiquitin chains on damaged mitochondria, their roles, and the roles of the other autophagy receptors in mediating mitophagy is unclear 8–11 . Autophagy receptors in mitophagy To clarify autophagy receptor function during mitophagy, genome editing was used to knock out five autophagy receptors in HeLa cells (pentaKO), which do not express endogenous Parkin. DNA sequencing (Supplementary Table 1) and immunoblotting of TAX1BP1, NDP52, NBR1, p62 and OPTN (Fig. 1a, lane 6) confirmed their knockout. We analyzed mitophagy in pentaKOs by measuring the degradation of cytochrome C oxidase subunit II (CoxII), a mtDNA encoded inner membrane protein, following mitochondrial damage with oligomycin and antimycin A (OA). After OA treatment, CoxII was degraded in WT cells expressing Parkin, but not in pentaKOs or ATG5 KO HeLa cells, indicating a block in mitophagy (Fig. 1b, c, Supplementary Table 1 and Extended Data Fig. 1a). As a second indicator of mitophagy, mitochondrial DNA (mtDNA) nucleoids were quantified by immunofluorescence (Extended Data Fig. 1b). After 24 h OA treatment, WT cells were nearly devoid of mtDNA, whereas pentaKOs and ATG5 KOs retained mtDNA (Fig. 1d, e). Parkin translocated to mitochondria (Extended Data Fig. 1c) and Mfn1 and Tom20 were degraded via the proteasome comparably in WT and pentaKOs (Fig. 1b, Extended Data Fig. 1d). mtDNA nucleoids clump following OA treatment in ATG5 KO cells but not in pentaKOs, consistent with a reported role of p62 10,11 . The five endogenous receptors in WT cells (Extended Data Fig. 1c) and each receptor re-expressed in pentaKOs (Extended Data Fig. 1e, f) translocated to mitochondria after OA treatment. However, in pentaKOs only GFP-NDP52, GFP-OPTN and to a lesser extent, GFP-TAX1BP1, rescued mitophagy (Fig. 1f, g). Another recently reported autophagy receptor, Tollip 12 , neither recruited to mitochondria nor rescued mitophagy following OA treatment (Extended Data Fig. 1g–i). We generated single OPTN, NDP52 KO and NDP52/OPTN double KO (N/O DKO) and NDP52/OPTN/TAX1BP1 triple KO (N/O/Tx TKO) cell lines (Supplementary Table 1, Fig. 1a) and found no compensatory change in the expression of the remaining receptors. NDP52 or OPTN KO alone caused no defect in mitophagy, whereas NDP52/OPTN DKO and to a greater extent, NDP52/OPTN/TAX1BP1 TKO inhibited mitophagy (Fig. 2a–d, Extended Data Fig. 2a, b). The robust mitophagy observed in OPTN KOs contrasts with a report indicating loss of mitophagy using RNAi-mediated knockdown of OPTN in HeLa cells 9 . Although NDP52 and OPTN redundantly mediate mitophagy, they function non-redundantly in xenophagy 13 . Their expression levels in human tissues indicate that OPTN or NDP52 may function more prominently in different tissues (Extended Data Fig. 2c). Mutations in autophagy receptors can lead to diseases such as primary open angle glaucoma (POAG, OPTN; E50K) 14 , ALS (OPTN; E478G and Q398X) 15 and Crohn's disease (NDP52; V248A) 16 . Defects in xenophagy occur when OPTN is mutated to block its phosphorylation by TANK-binding kinase 1 (TBK1; S177A) or ubiquitin binding (D474N) 13,17 . In pentaKOs, the UBAN-domain disrupting mutants OPTN-Q398X, OPTN-D474N and OPTN-E478G (Extended Data Fig. 2d) failed to translocate to mitochondria (Extended Data Fig. 2e, f) or rescue mitophagy (Fig. 2e, Extended Data 2g). OPTN-S177A weakly rescued mitophagy and minimally translocated to mitochondria, whereas OPTN-E50K robustly translocated and substantially rescued mitophagy (Fig. 2e, Extended Data Fig. 2e–g). NDP52-V248A fully recruited to mitochondria and rescued mitophagy, but a mutant lacking the ZF ubiquitin-binding domains (NDP52-ΔZF) 18 did not (Extended Data Fig. 2h–k, Fig. 2f). Thus, ubiquitin binding by OPTN and NDP52 is necessary for mitophagy and some disease-causing mutations prevent mitophagy. TBK1 and OPTN cooperate in mitophagy TBK1 phosphorylation of OPTN at S177 increases its association with LC3 during xenophagy 13 , and the OPTN E50K mutation increases TBK1/OPTN binding 19 . TBK1 auto-phosphorylation at Ser172 is indicative of TBK1 activation 20 and occurs in a Parkin-dependent manner following 3 h OA treatment, but only in cells expressing OPTN (Extended Data Fig. 3b, lanes 4 and 10). Prolonged OA treatment induces moderate TBK1 phosphorylation in the absence of Parkin but still requires PINK1 (Extended Data Fig. 3c). To investigate TBK1 function during mitophagy, we generated TBK1 KO, TBK1/NDP52 (T/N) DKO and TBK1/OPTN (T/O) DKO HeLa cells (Extended Data Fig. 3d, Supplementary Table 1). Parkin translocated to mitochondria in all lines, however, only TBK1/NDP52 DKOs displayed defective mitophagy (Extended Data Fig. 3e, Fig. 2g–j). Mitophagy in TBK1/NDP52 DKOs was rescued by WT-TBK1 or phospho-mimetic OPTN (OPTN-S177D), but not by kinase-dead TBK1 (TBK1-K38M) (Extended Data Fig. 3g–i). Thus, in the absence of NDP52, TBK1 is critical for effective mitophagy via OPTN. Ubiquitin phosphorylation in mitophagy Since many autophagy receptors recruit to mitochondria following Parkin activation, why do only some function in mitophagy? Parkin-mediated mitophagy is driven by PINK1's phosphorylation of Ser65 of both ubiquitin 5–7,21,22 and the UBL domain of Parkin 23 . Since Ser65 phospho-ubiquitin is structurally unique, it may differentially interact with ubiquitin binding proteins 22 . To determine whether OPTN is directly recruited to phospho-ubiquitin on mitochondria, we conditionally expressed PINK1 on undamaged mitochondria 10 in HeLa cells lacking Parkin (Fig. 3a, Extended Data Fig. 4a). When PINK1Δ110-YFP-2xFKBP is cytosolic, mCherry-OPTN, mCherry-NDP52 and mCherry-p62 are also cytosolic (Extended Data Fig. 4b, c). When PINK1Δ110-YFP-2xFKBP is localized to FRB-Fis1 expressing mitochondria with rapalog, where ubiquitin on surface proteins 24 (Extended Data Fig. 4a) can be phosphorylated 5,21,25,26 , OPTN and NDP52 are recruited (Fig. 3a, b), but p62 remains cytosolic (Fig. 3b, Extended Data Fig. 4c). OPTN/NDP52 recruitment requires PINK1 kinase activity (Fig. 3a, b) and receptor-ubiquitin binding, as OPTN-D474N and NDP52-ΔZF fail to recruit following rapalog treatment (Fig. 3b, Extended Data Fig. 4d, e). Therefore, PINK1 ubiquitin kinase activity recruits OPTN/NDP52 via ubiquitin binding domains to mitochondria in the absence of Parkin. To determine whether the observed autophagy receptor recruitment to mitochondria in the absence of Parkin can induce mitophagy, we developed a sensitive FACS based mitophagy assay. We expressed mitochondrial-targeted mKeima (mt-mKeima, see Online Methods) in WT and pentaKOs also expressing mitochondrial FRB-Fis1 and PINK1Δ110-YFP-2xFKBP. mt-mKeima engulfment into lysosomes results in a spectral shift due to low pH. Only 1% (range 0.89–1.15) of WT or pentaKO cells display mitophagy when PINK1 is cytosolic. However, when PINK1 is recruited to mitochondria with rapalog, mitophagy increases ~7-fold in WT cells and ~8-fold with overexpressed OPTN (Table 1, Extended Data Fig. 5a). PentaKOs showed no increase in mitophagy after targeting PINK1 to mitochondria (Table 1, Extended Data Fig. 5b). When rescued with FLAG/HA-OPTN or FLAG/HA-NDP52, pentaKOs displayed an increase in mitophagy of more than 5-fold and 4-fold, respectively (Table 1, Extended Data Fig. 5b–e). Rescue with FLAG/HA-p62 or ubiquitin-binding mutants (OPTN-Q398X, OPTN-D474N and NDP52ΔZF) failed to increase mitophagy above baseline, but other mutants (OPTN-E50K, OPTN-S177A and NDP52-V248A) rescued mitophagy (Table 1, Extended Data Fig. 5c–f). OPTN-E50K and S177A restored mitophagy as well as or better than WT OPTN (Table 1), differing from their response in the presence of Parkin (Fig. 2e) likely due to the lack of robust TBK1 activation in the absence of Parkin (Extended Data Fig 3b). Here, enhanced OPTN-E50K binding to TBK1 19 may become advantageous by allowing OPTN phosphorylation by TBK1 in the absence of Parkin thus improving mitophagy. In the absence of TBK1 activation, WT OPTN is likely not phosphorylated at S177 and thus is functionally similar to S177A OPTN. Importantly, ubiquitin kinase activity of PINK1 is required, as kinase-dead (KD) PINK1 did not induce mitophagy (Table 1, Fig. 3c, Extended Data 5g). Parkin expression dramatically increased mitophagy in FLAG/HA-OPTN expressing pentaKOs (Table 1, Extended Data Fig. 5h), supporting the model that PINK1-phosphorylated ubiquitin recruits receptors for mitophagy and Parkin ubiquitination of mitochondrial substrates amplifies this ubiquitin signal. Comparing mitophagy induced by OA treatment in WT relative to PINK1KO cells confirmed that endogenous PINK1 mediates mitophagy in the absence of Parkin (Extended Data Fig. 6a, b). Re-expressing PINK1 in PINK1 KO cells rescued OA-induced mitophagy (Extended Data Fig. 6c, d). Furthermore, mCherry-OPTN is recruited to mitochondria in the absence of Parkin in a PINK1-dependent manner following prolonged exposure to OA (Extended Data Fig. 6e, f). Given that PINK1 ubiquitin kinase activity can recruit OPTN and NDP52, we investigated autophagy receptor binding to phospho-mimetic (S65D) HA-ubiquitin in HeLa cells. Endogenous OPTN and NDP52 preferentially co-immunoprecipitate (co-IP) with HA-ubiquitinS65D (Extended Data Fig. 7a). Conversely, p62 was present at equal levels in all co-IPs (Extended Data Fig. 7a). Ubiquitin-modified and unmodified forms of OPTN and NDP52 were present in co-IPs, and HA-ubiquitinS65D induced or preserved this modification (Extended Data Fig. 7a). Co-IP samples treated with the deubiquitinase USP2 removed the ubiquitin-modified bands on OPTN and NDP52, yet OPTN and NDP52 retained HA-ubiquitinS65D binding (Extended Data Fig. 7b). Binding of endogenous receptors in HeLa cell cytosol to in vitro phosphorylated strep-tagged ubiquitin (Extended Data Fig. 7c) showed that OPTN, but not p62, bound better to phospho-ubiquitin (Extended Data Fig. 7d, e). However, recombinant GST-OPTN did not bind better to in vitro phosphorylated K63 linked ubiquitin chains 27 indicating that OPTN may need additional factors or modification in vivo to preferentially bind Ser65 phosphorylated ubiquitin. OPTN/NDP52 recruit upstream machinery Autophagy receptors are thought to primarily function by bridging LC3 and ubiquitinated cargo 1,2 . In mCherry-Parkin WT cells, GFP-LC3B accumulated in distinct puncta adjacent to mitochondria after OA treatment (Extended Data Fig. 8a). Although OA also induced GFP-LC3B puncta in pentaKOs, they were fewer and not near mitochondria (Extended Data Fig. 8a). Conversely, GFP-LC3B in ATG5 KOs was near mitochondria, but not in puncta (Extended Data Fig. 8a). LC3B lipidation is retained in pentaKOs, but lost in ATG5 KOs (Extended Data Fig. 8b). This indicates that ATG5 is activated downstream of PINK1, but independently of autophagy receptors, and that LC3 lipidation and mitochondrial localization are independent steps of mitophagy. OPTN and NDP52 interact with LC3B and LC3C, respectively, for Salmonella clearance 13,28 . Beyond that, little is known about the specificity of LC3 family members toward autophagy receptors 29 or their involvement in mitophagy. We examined the recruitment of all LC3/GABARAP family members to mitochondria in WT, pentaKO and NDP52/OPTN DKO cells. The OA-induced mitochondrial localization of GFP-LC3s in WT cells was absent in pentaKOs, while only GFP-LC3B recruitment was inhibited in NDP52/OPTN DKOs (Fig. 4a, Extended Data Fig. 8c). GFP-LC3C recruitment was inhibited in NDP52/OPTN/TAX1BP1 TKOs (Extended Data Fig. 8d, e), indicating that TAX1BP1 can recruit LC3C during mitophagy. GABARAPs did not recruit to mitochondria, indicating they likely play no substantial role in mitophagy (Extended Data Fig. 9a). We also examined the involvement of WIPI1 and DFCP1, two proteins that mediate phagophore biogenesis upstream of LC3 30 , in mitophagy. In WT cells, OA induced foci of both GFP-WIPI1 and GFP-DFCP1, mostly localized on or near mitochondria (Fig. 4b, c, Extended Data Fig. 9b, c). In NDP52/OPTN DKOs, GFP-WIPI1 and GFP-DFCP1 foci were reduced and were almost undetectable in pentaKOs (Fig. 4b, c, Extended Data Fig. 9b, c). Despite this, phosphorylation of Beclin1 31 was normal in both pentaKOs and NDP52/OPTN DKOs (Extended Data Fig. 9d), indicating that failure to recruit WIPI1/DFPC1 was not due to defective Vps34 complex. GFP-DFCP1 recruitment in pentaKOs was rescued by expression of FLAG/HA-OPTN or FLAG/HA-NDP52, but not by FLAG/HA-p62 (Extended Data Fig. 10a). Though autophagy receptors are thought to function late in autophagy with LC3 32 , the deficit in WIPI1 and DFCP1 recruitment to mitochondria indicates a defect upstream in autophagosome biogenesis. ULK1 phosphorylation by AMPK at S317 and dephosphorylation at S757 33 , required for activation, occurs comparably in WT, NDP52/OPTN DKO and pentaKO cells (Fig. 4d). Despite this, ULK1 localization to mitochondria 34 following OA is diminished by half in the NDP52/OPTN DKOs and more than 80% in pentaKOs (Fig. 4e, f). FLAG/HA-OPTN or FLAG/HA-NDP52, but not FLAG/HA-p62, rescued GFP-ULK1 localization in pentaKOs (Extended Data Fig. 10b). Overall, these data indicate that NDP52 and OPTN recruit ULK1 to initiate mitophagy. We next assessed if ubiquitin phosphorylation, independent of Parkin, is also sufficient to recruit ULK1 to mitochondria. Rescue of pentaKOs expressing FRB-Fis1 and PINK1Δ110-YFP-FKBP with myc-OPTN or myc-NDP52 resulted in mitochondrial ULK1 puncta following rapalog treatment (Fig. 4g, h). Myc-OPTN-E50K also rescued ULK1 recruitment to mitochondria, but ALS-associated mutant myc-OPTN-Q398X did not (Fig. 4i, Extended Data Fig. 10d). ULK1 recruitment was restored by myc-OPTN-F178A (Fig. 4i, Extended Data Fig. 10d), a mutation that disrupts OPTN association with LC3 12 , indicating that ULK1 recruitment is not through LC3 interaction and occurs upstream of LC3. Taken together, our data show that PINK1 ubiquitin-kinase activity is sufficient to recruit the autophagy receptors and upstream autophagy machinery to mitochondria to induce mitophagy. Conclusions Through genetic knockout of five autophagy receptors we have defined their relative roles in mitophagy and identified their unanticipated upstream involvement in autophagy machinery recruitment. p62 and NBR1 are dispensable for Parkin-mediated mitophagy; OPTN and NDP52 are the primary, yet redundant, receptors. We also uncovered a new and more fundamental role for PINK1 in mitophagy: to directly induce mitophagy through phospho-ubiquitin-mediated recruitment of autophagy receptors. We posit that PINK1 generates the novel and essential signature (phospho-ubiquitin) on mitochondria to induce OPTN and NDP52 recruitment and mitophagy; Parkin acts to increase this signal by generating more ubiquitin chains on mitochondria, which are subsequently phosphorylated by PINK1. Our findings clarify the role of Parkin as an amplifier of the PINK1-generated mitophagy signal, phospho-ubiquitin, which can engage the autophagy receptors to recruit ULK1, DFCP1, WIPI1 and LC3 (see model in Extended Data Fig. 10e). Online Content Methods, along with any additional Extended Data display items, are available in the online version of the paper; references unique to these sections appear only in the online paper. METHODS Cell Culture, Antibodies and Reagents HEK293T, HeLa and PINK1 KO 35 cells were cultured in Dulbecco's modified eagle medium (Life Technologies) supplemented with 10% (v/v) Fetal Bovine Serum (Gemini Bio Products), 10 mM HEPES (Life Technologies), 1 mM Sodium Pyruvate (Life Technologies), nonessential amino acids (Life Technologies) and GlutaMAX (Life Technologies). HeLa cells were acquired from the ATCC and authenticated by the Johns Hopkins GRCF Fragment Analysis Facility using STR profiling. All cells were tested for mycoplasma contamination bimonthly using the PlasmoTest kit (InvivoGen). Transfection reagents used were: Effectene (Qiagen), Lipofectamine LTX (Life Technologies), Avalanche-OMNI (EZ Bio-systems), X-tremeGENE HP (Roche) and X-tremeGENE 9 (Roche). Rabbit monoclonal and polyclonal antibodies used: Beclin, pULK1-S317, pULK1-S757, TBK1, pTBK1-S172, NDP52, TAX1BP1, ATG5, Actin, and HA (Cell Signaling Technologies); GAPDH and LC3B (Sigma); ULK1 and Tom20 (Santa Cruz Biotechnology); Optineurin (OPTN) (Proteintech); GFP (Life Technologies); pSer65 ubiquitin (Millipore) and Mfn1 was generated previously 36 . Mouse monoclonal antibodies used: NBR1 and p62 (Abnova), Cytochrome C oxidase subunit II (CoxII, Abcam), Parkin (Santa Cruz Biotechnology), DNA (Progen Biotechnik), ubiquitin (Cell Signaling). Chicken anti-GFP (Life Technologies) was also used. For catalog numbers see Supplementary Table 1. Human tissue panel blots were purchased (NOVUS Biologicals). Generation of knockout lines using TALEN and CRISPR/Cas9 gene editing To generate knockout cell lines, TALENs and CRISPR gRNAs were chosen that targeted an exon common to all splicing variants of the gene of interest (listed in Supplementary Table 1). Transcription activator-like effector nuclease (TALEN) was used to generate the OPTN KO HeLa cell line. The TALEN constructs were generated by sequential ligation of coding repeats into pcDNA3.1/Zeo-Talen(+63), as previously described 37–39 . The CRISPR/Cas9 system generated by the Church lab 40 , was used to knockout ATG5, NDP52, TAX1BP1, NBR1, p62 and TBK1. Oligonucleotides (Operon) containing CRISPR target sequences were annealed and ligated into AlfII-linearized gRNA vector (Addgene) 40 . For CRISPR/Cas9 gene editing, HeLa cells were transfected with gRNA constructs, hCas9 (Addgene) and pEGFP-C1 (Clontech), or for TALEN gene editing HeLa cells were transfected with OPTN TALEN constructs and pEGFP-C1. Two days after transfection, GFP-positive cells were sorted by fluorescence activated cell sorting and plated in 96-well plates. Single colonies were expanded into 24-well plates before screening for depletion of the targeted gene product by immunoblotting. As a secondary screen of some knockout lines, genomic DNA was isolated from cells and the genomic regions of interest were amplified using PCR followed restriction enzyme digestion analysis (primers listed in Supplementary Table 1). Sequencing of targeted genomic regions of knockout lines was also conducted to confirm the presence of frameshifting indels in the genes of interest (Supplementary Table 1). To generate multiple gene knockout cell lines, parental cell lines were transfected sequentially with one or multiple gRNA constructs to generate desired knockout lines. Parental cell lines are outlined in Supplementary Table 1. Cloning and generation of stable cell lines pMXs-puro-GFP-WIPI1 and pMXs-puro-GFP-DFCP1 were a kind gift from Dr. N. Mizushima (University of Toyko, Japan) and pMXs-IP-GFP-ULK1 was purchased from Addgene (#38193). To generate pBMN-mEGFP-C1, mEGFP-C1 (Addgene #36412) was PCR amplified (together with the multiple cloning site) and cloned into pBMN-Z at BamHI/SalI sites using the Gibson Cloning kit (New England BioLabs) according to manufacturer's instructions. The BamHI and SalI sites used to insert mEGFP-C1 were not regenerated. The following GFP-tagged plasmids were generated by PCR amplification of open reading frames followed by ligation into pBMN-mEGFP-C1: OPTN, NDP52, p62, TAX1BP1, NBR1, LC3A, LC3B, LC3C, GABARAP, GABARAPL1, GABARAPL2. The Gateway Cloning (Invitrogen) system was used to generate GFP-, mCherry-, myc- and FLAG/HA-constructs. Briefly, TBK1, TBK1-K38M, NDP52, OPTN, p62, DFCP1, WIPI1 and ULK1 were cloned into pDONR2333. Mutations in cDNA sequences were introduced using PCR site directed mutagenesis in the pDONR2333 vector, (sequences of mutagenesis primers used are available upon request) then recombined into pHAGE-N-FLAG/HA, pHAGE-N-GFP, pHAGE-N-mCherry and/or pDEST-N-myc using LR Clonase (Invitrogen) as per the manufacturer's protocol. All constructs generated in this study were verified by sequencing. To generate stably transfected cell lines, retroviruses (for pBMN-mEGFP-C1 constructs, pBMN-mCherry-Parkin, pBMN-puro-P2A-FRB-Fis1, pCHAC-mt-mKeima-IRES-MCS2) and lentiviruses (for pHAGE- and pDEST- constructs) were packaged in HEK293T cells. HeLa cells were transduced with virus for 24 h with 8 μg/ml polybrene (Sigma) then optimized for protein expression via selection (puromycin or blasticidin) or fluorescence sorting. Translocation and mitophagy treatments Cells were either left untreated or treated with 10 μM Oligomycin (Calbiochem), □ μM Antimycin A (Sigma) (referred to as OA) in fresh growth medium for different periods of time as indicated in the figures. Some experiments were performed with 10 μM Carbonyl cyanide m-chlorophenyl hydrazine (CCCP) as indicated (Sigma-Aldrich). We chose to use OA to depolarize mitochondria in most of our experiments, as they are specific mitochondrial respiratory complex inhibitors and less toxic. Long treatment time points of both OA and CCCP were also supplemented with the apoptosis inhibitor 20 μM QVD (ApexBio) to prevent cell death. Immunoblotting and Phos-Tag gels HeLa cells seeded into 6-well plates were either untreated or treated with 10 μM Oligomycin (Calbiochem), □ μM Antimycin A (Sigma) and 20 μM QVD (ApexBio) in fresh growth medium for different periods of time as indicated in figure legends. Cells were lysed in 1X LDS sample buffer (Life Technologies) supplemented with 100 mM dithiothreitol (DTT, Sigma) and heated to 99 °C with shaking for 7–10 minutes. 25–50 μg of protein per sample was separated on 4–12% Bis-Tris gels (Life Technologies) according to manufacturer's instructions and then transferred to polyvinyl difluoride membranes and immunoblotted using antibodies as indicated in figure legends. To assess mitophagy, CoxII quantification was conducted using ImageLab software (BioRad). For uncropped images of all immunoblots, see Supplementary Information. To dephosphorylate samples, cells were collected as above and lysed in 1X NEB Buffer 3 (New England BioLabs) supplemented with 1% Triton X-100 and passed through a 26.5 gauge needle. Calf intestinal phosphatase (CIP, New England BioLabs) was added to half the cell lysate and the other half was used as an untreated control. Both samples were incubated for 1 h at 37 °C and analyzed by SDS-PAGE and immunoblotting. To analyze Beclin phosphorylation, lysates were prepared in sample buffer lacking EDTA and run on 8% Tris-Glycine gels containing 20 μM Phos-Tag (Wako) and 40 μM MnCl2 as described previously 31 . Gels lacking Phos-Tag were run simultaneously as a negative control. Electrophoresis and western transfer were carried out using standard protocols with the exception that Phos-Tag gels were incubated in 10 mM EDTA for 10 min to remove excess Mn2+ prior to transfer. Immunoprecipitation WT or PINK1 KO HeLa cells were transiently transfected with HA-tagged ubiquitin WT, S65A or S65D with or without mCherry-Parkin for 24 h. Cells were harvested, lysed and the HA-ubiquitin was immunoprecipitated as reported previously 5 , using anti-HA conjugated beads (Pierce). To deubiquitinate the bound proteins, after binding the HA-ubiquitin, beads were washed three times and incubated in 50 mM Tris-Cl (pH 7.5), 150 mM NaCl, 5 mM DTT and 1.47μg USP2 (Boston Biochem) at 37 °C for 1 h. The reaction was stopped and the remaining bound were proteins were washed 5 times with 1 mL of buffer (50 mM Tris-Cl (pH 7.5), 150 mM NaCl), then eluted by boiling with 1X LDS sample buffer. In vitro phosphorylation Strep-tagged ubiquitin was incubated with either TcPINK1 WT or kinase-dead as previously reported 5 . This ubiquitin was then incubated with cytosol from WT HeLa cells in 20 mM HEPES-KOH, pH 7.6, 220 mM mannitol and 70 mM sucrose at 4°C for 1 h. Strep-Tactin beads (Qiagen) were then added to bind the strep-ubiquitin for an additional 1 h at 4°C. The ubiquitin and bound proteins were then eluted with 50 mM biotin in 50 mM Tris for 15 min at room temperature (RT). Samples were then diluted in LDS sample buffer prior to SDS-PAGE and immunoblot analysis. Immunofluorescence microscopy HeLa cells, seeded in 2-well chamber slides (Lab-Tek), were treated as indicated in the figures legends. Following treatment, cells were rinsed in PBS and fixed for 15 min at RT with 4% paraformaldehyde. Cells were then permeabilized and blocked with 0.1% Triton X-100, 3% goat serum in PBS for 40 minutes at RT. For immunostaining, cells were incubated with antibodies (as indicated in figure legends) diluted in 3% goat blocking serum overnight at 4 °C, then rinsed with PBS and incubated with either anti- rabbit or mouse Alexa Fluor- 488 and 633 conjugated secondary antibodies (Life Technologies), or anti-chicken Alexa Fluor 488 conjugated antibody (Life Technologies) for 1 h at RT. Cells were washed 3 times for 5 min each with 1% Triton X-100, PBS. During the final wash step, cells were incubated with DAPI (10 μg/mL DAPI, Sigma) in PBS for 5 min. To measure mitophagy by mitochondrial DNA (mtDNA) immunostaining; images were collected from samples stained with DAPI and immunostained for DNA using a plan-Apochromat 63×/1.4 oil DIC objective on an LSM 510 microscope (Zeiss). Four image slices were collected through the Z plane encompassing the top and bottom of the cells. Image analysis was performed on all images collected in the Z plane using Volocity software (Perkin Elmer v6.0.1). The percent mtDNA stain remaining was calculated using the following formula: (cDNAv-nDNAv)/n, where cDNAv= the total cellular DNA volume determined by staining using anti-DNA antibodies and, nDNAv = the total nuclear DNA stain volume determined using DAPI, and n= the number of cells. The mtDNA stain volume in untreated cells was normalized to 100% and the amount of mtDNA stain remaining after drug treatment was subsequently determined. Final values represent data acquired from 50–200 cells from three independent experiments. To analyze LC3/mitochondria protein colocalization; cells were treated, fixed and immunostained as above. Between 5–8 slices were imaged through the Z plane using either a plan-Apochromat 63× or 100×/1.4 oil DIC objective on a CW STED confocal microscope (Leica). Volocity software (Perkin Elmer, v6.0.1) was used to measure intensity of the GFP signal representing LC3 in the volume occupied by mitochondria (as defined by Tom20 positive region) and the cytosol (as defined by Tom20 negative region). “Normalized mitochondrial LC3” was calculated using the following formula: Normalized mitochondrial LC3 = (mi/mv)/(ci/cv), where mi = mitochondrial GFP intensity, mv = mitochondrial volume, ci = cytosolic GFP intensity and cv= cytosolic volume. The resulting Normalized mitochondrial LC3 is equal to 1 if the intensity of GFP is equal per volume in the cytosolic and mitochondrial volumes (no translocation) and is above one if the mitochondrial intensity is higher per volume (translocation). Final values for Normalized mitochondrial LC3 represents data acquired from 50–105 cells from three independent experiments. For GFP-DFCP1, GFP-WIPI1 and GFP-ULK1 puncta analysis; cells were treated, prepared and imaged on the CW STED as above with the addition of immunofluorescence using either rabbit or chicken GFP antibodies to enhance the signal in the green channel. For GFP-DFCP1, puncta were quantified using Volocity software (Perkin Elmer v6.0.1) and for GFP-WIPI1 and GFP-ULK1 puncta were quantified manually. Colocalization of autophagy receptors with GFP-DFCP1 or GFP-ULK1 was assessed with line scans using LAS AF software (Leica, v.2.6.0.7266). Heterodimerization The C-terminal Fis1 tail of human Fis1 (amino acids 92–152) was cloned into pC4-RhE vector (ARIAD) at SpeI/BamHI sites to make FRB-Fis1 construct, the insert of which was then PCR amplified and cloned into pBMN-Z vector together with Puro-P2A sequence at HindIII, XhoI and NotI sites by In-Fusion kit from Clontech to make pBMN-puro-P2A-FRB-Fis1. For receptor translocation assays, WT HeLa cells stably expressing FRB-Fis1 were generated using retroviral transduction as described above. Previously generated PINK1Δ110-YFP-2xFKBP 41 WT and KD and individually each mCherry-tagged autophagy receptor were transfected into FRB-Fis1 stable HeLa cells for 24 h. Cells were then treated with 0.5 μM rapalog (Clontech) for 8 h as previously described 41 . Cells were then fixed and stained as described above. Cells were manually counted for translocation of mCherry-tagged autophagy receptors to mitochondria. Final values represent data collected from 100–150 cells for three independent experiments. For ULK1 and DFCP1 rescue analysis, pentaKO HeLa cells stably expressing FRB-Fis1 were transiently transfected with PINK-1Δ110-YFP-2xFKBP, mCherry-ULK1 and one of the autophagy receptors: myc-OPTN, myc-NDP52, myc-OPTN-F178A, myc-OPTN-E50K or myc-OPTN-Q398X for 18–24 h. Cells were treated with 0.5 μM Rapalog or 100% ethanol (vehicle) for 7 h and imaged live on a Zeiss 780 in a humidified 37°C/5% CO2 chamber. To visualize mitochondria in vehicle -treated controls, cells were pre-incubated for 10 minutes in 75 nM Mitotracker Deep Red (Invitrogen) prior to imaging. Fields of PINK1-YFP positive cells were imaged blindly to the mCherry-containing channel. The images were then blinded and counted manually for translocation of mCherry-tagged ULK1 to mitochondria. Final values represent >75 cells counted over at least two independent experiments. Mito-Keima mitophagy assay mt-mKeima 42 (a gift from A. Miyawaki, Brain Science Institute, RIKEN, Japan) was cloned into pCHAC-MCS-1-IRES-MCS2 vector (Allele Biotechnology). PINK1(Δ110)-YFP-2xFKBP WT and KD were PCR amplified from the original pC4 and cloned into pRetroQ-AcGFP-C1 at NheI/XhoI sites by Gibson assembly kit. HA-tag was removed and stop codon is introduced. WT and 5KO pentaKO HeLa cells stably expressing FRB-Fis1, mt-mKeima and either WT or KD PINK1Δ110-YFP-2xFKBP were generated using retroviral transduction as described above. FLAG/HA-receptors were stably-expressed in these cells by lentivirus transduction as described above then treated with 0.5 μM rapalog for 24 h. Cells were then resuspended in sorting buffer (145 mM NaCl, 5 mM KCl, 1.8 mM CaCl2, 0.8 mM MgCl2, 10 mM HEPES, 10 mM glucose, 0.1% BSA) containing 10 μg/mL DAPI. Analysis was performed using Summit software (v6.2.6.16198) on a Beckman Coulter MoFlo Astrios cell sorter. Measurements of lysosomal mt-mKeima were made using dual-excitation ratiometric pH measurements at 488 (pH 7) and 561 (pH 4) nm lasers with 620/29 nm and 614/20 nm emission filters, respectively. For each sample, 50,000 events were collected and subsequently gated for YFP/mt-mKeima double-positive cells that were DAPI-negative. Data were analyzed using FlowJo (v10, Tree Star). Statistical calculations All statistical data were calculated and graphed using GraphPad Prism 6. To assess statistical significance, data from three or more independent experiments were analyzed using one-way ANOVA and Tukey's post-test with a confidence interval of 95%. All error bars are expressed as mean ± standard deviation (s.d.). In Fig.4h, i outliers were removed using ROUT in GraphPad Prism 6 with a Q=1%, 1–2 values from each condition were removed. Extended Data Extended Data Figure 1 Analysis of knockout cell lines and characterization of autophagy receptor translocation to damaged mitochondria a, ATG5 KO cell line confirmed by immunoblotting. b, Representative images of mitochondrial DNA nucleoids in HeLa cells immunostained with an α-DNA antibody (green) confirming colocalization with the mitochondrial marker Tom20 (red) (n=3). c, Mitochondrial fractions from mCherry-Parkin (mCh-Parkin) expressing pentaKO and WT cells were assessed by immunoblotting. d, mCh-Parkin expressing WT, pentaKO and ATG5 KOs were treated with OA or OA and MG132. Cell lysates were assessed by immunoblotting. e, Expression levels of GFP-tagged OPTN, NDP52, p62, NBR1 and TAX1BP1 re-expressed in pentaKOs by immunoblotting. f, Representative images of mCh-Parkin expressing pentaKOs from e immunostained for Tom20 (n=3). g, Expression of GFP-Tollip in mCh-Parkin pentaKOs. h, pentaKOs mCh-Parkin and with or without GFP-Tollip expression were immunoblotted. i, Representative images of mCh-Parkin pentaKOs expressing GFP-Tollip immunostained for Tom20 (n=3). Scale bars, 10 μm. Extended Data Figure 2 OPTN, NDP52 and TAX1BP1 triple knockout analysis and disease-associated mutations a, KO cell lines with or without mCherry-Parkin (mCh-Parkin) expression were immunoblotted and b, CoxII levels were quantified. c, A panel of human tissue lysates was immunoblotted. d, Expression of WT or mutant GFP-OPTN in mCh-Parkin pentaKOs. e, Quantification of cells in f. >100 cells per condition. f, Representative images of mCh-Parkin pentaKOs expressing GFP-OPTN mutants immunostained for Tom20 (n=3). g, pentaKOs expressing mCh-Parkin were rescued with WT or mutant GFP-OPTN, analyzed by immunoblotting. See Fig. 2e for quantification of CoxII. h, Expression of WT or mutant GFP-NDP52 in mCh-Parkin pentaKOs. i, Quantification of cells in j. >100 cells per condition. j, Representative images of mCh-Parkin pentaKOs expressing WT or mutant GFP-NDP52 were immunostained for Tom20 (n=3). k, pentaKOs expressing mCh-Parkin rescued with WT or mutant GFP-NDP52 were analyzed by immunoblotting. See Fig. 2f for quantification of CoxII. Quantification in b and i are displayed as mean ± s.d. from 3 independent experiments using one-way ANOVA tests (***P<0.001, ns, not significant) and in e as mean from 2 independent experiments. Scale bars, 10 μm. Extended Data Figure 3 TBK1 in activates OPTN in PINK1/Parkin mitophagy a, Representative images of untreated mCherry-Parkin (mCh-Parkin) cells and merged images of treated cells as indicated immunostained for DNA. See Fig. 2a for anti-DNA/DAPI images of treated samples (n=3). b, Cell lysates from WT, N/O (NDP52/OPTN) DKO, OPTN KO and NDP52 KO cells with or without mCh-Parkin expression were immunoblotted for TBK1 activation. c, Cell lysates from WT and PINK1 KO cells without Parkin expression were immunoblotted for TBK1 activation (S172 phosphorylation). d, Confirmation of T/N (TBK1/NDP52) DKO, T/O (TBK1/OPTN) DKO and TBK1 KO by immunoblotting. e, KO cell lines from d were immunostained for Tom20 (n=3). f, Representative images of untreated mCh-Parkin WT and KO cells, and merged images of treated cells as indicated were immunostained for DNA. See Figure 2g for anti-DNA/DAPI images treated samples (n=3). g, T/N DKO cells rescued with GFP-TBK1 WT or K38M, or GFP-OPTN S177D and were assessed by immunoblotting. h, Cells in g were assessed by immunoblotting. i, Quantification of CoxII levels in h displayed as mean ± s.d. from 3 independent experiments and use one-way ANOVA tests (**P<0.005, ns, not significant). OA, Oligomycin and Antimycin A. Scale bars, 10 μm. Extended Data Figure 4 Parkin-independent recruitment of receptors to mitochondria through PINK1 activity a, Isolated mitochondria from WT and pentaKOs with or without FRB-Fis1 and with WT or kinase-dead (KD) PINK1Δ110-YFP-2xFKBP were immunoblotted. b–e, Representative images of pentaKOs expressing FRB-Fis1, WT (PINK1-WT) or kinase-dead (PINK1-KD) PINK1Δ110-YFP-2xFKBP and either (b) mCherry-OPTN or mCherry-NDP52, (c) mCherry-p62, (d) mCherry-OPTN-D474N or (e) mCherry-NDP52-ΔZF. Cells were (b) untreated or (c–e) treated with rapalog then immunostained for Tom20. All images are representative of three independent experiments. See Figure 3b for quantification. Scale bars, 10 μm. Extended Data Figure 5 PINK1 directly stimulates mitophagy in the absence of mitochondrial damage a, b, Cells were treated with rapalog and analyzed by FACS for lysosomal positive mt-mKeima. Representative data for WT HeLa (a) and pentaKO (b) without or with FLAG/HA-OPTN. c, d, Cell lysates from pentaKOs expressing FRB-Fis1, PINK1Δ110-YFP-2xFKBP, mt-mKeima and (c) WT FLAG/HA-OPTN or mutants, (d) FLAG/HA-p62, WT FLAG/HA-NDP52 or NDP52 mutants as indicated were assessed for receptor expression by immunoblotting. e, f, Cells from c and d were rapalog treated analyzed by FACS for lysosomal positive mt-mKeima. Representative data of two experiments is presented. g, Cell lysates from pentaKOs expressing FRB-Fis1, with or without FLAG/HA-OPTN and WT or kinase-dead (KD) PINK1Δ110-YFP-2xFKBP were assessed for OPTN by immunoblotting. h, FLAG/HA-OPTN pentaKOs expressing FRB-Fis1, PINK1Δ110-YFP-2xFKBP, mt-mKeima transfected and either vector or untagged Parkin were analyzed by FACS. Representative data of two experiments is presented. Extended Data Figure 6 PINK1 directly stimulates mitophagy upon mitochondrial damage Representative data of mt-mKeima-expressing a, WT, PINK1 KO or c, PINK1 KO rescued with PINK1-WT cells treated with OA then analyzed by FACS. b, d, Average percent mitophagy for two replicates of a and c, respectively. e, Representative images of WT HeLa cells expressing mCherry-OPTN and treated with OA as indicated were immunostained for Tom20 (n=3). f, Quantification of mCherry-OPTN translocation from cells in e. Data displayed as mean ± s.d. from 3 independent experiments and using one-way ANOVA tests (***P<0.001, ns, not significant). Extended Data Figure 7 OPTN and NDP52 preferentially bind phospho-mimetic ubiquitin a, HeLa cells expressing mCherry-Parkin (Parkin) and HA-ubiquitin (HA-UB) WT, S65D or S65A were treated with CCCP. HA-UB was co-immunoprecipitated and the bound fraction was analyzed by immunoblotting. Quantification of the total bound fraction of OPTN, NDP52 and p62 are shown. b, HA-ubiquitin transfected into HeLa cells with mCherry-Parkin were treated with CCCP. HA-ubiquitin was immunoprecipitated. The bound fraction was treated with the deubiquitinase USP2 and washed to remove all unbound protein following deubiquitination. Quantification of the total bound fraction of OPTN, NDP52 and p62 are shown in the right panel. c,d, Strep-tagged ubiquitin (Strep-UB) was incubated with either WT or kinase-dead (KD) PINK1 in an in vitro phosphorylation reaction, immunoblotted with an anti-phosphoS65 ubiquitin antibody (c) and was then incubated with cytosol harvested from untreated, WT HeLa cells. The ubiquitin was then pulled down using Strep-Tactin beads and (d) analyzed by immunoblotting. e, Quantification of bound OPTN and p62 normalized to total ubiquitin. Data displayed in a, b and e as mean ± s.d. from 3 independent experiments and use one-way ANOVA tests. (***P<0.001, **P<0.005, *P<0.05). †, non-specific band. a.u., arbitrary units. Extended Data Figure 8 Analysis of LC3 family members and their translocation to damaged mitochondria in autophagy receptor KO cell lines a, Representative images of WT, pentaKO and ATG5 KO HeLa cells expressing mCherry-Parkin (mCh-Parkin) and GFP-LC3B were immunostained for Tom20 (n=3). b, Cell lysates from mCh-Parkin expressing WT, pentaKO and ATG5 KO cells were immunoblotted. c, Representative images of WT, N/O (NDP52/OPTN) DKO and pentaKOs expressing mCh-Parkin and either GFP-tagged LC3A, LC3B or LC3C were immunostained for Tom20 (n=3, see Figure 4a for quantification). d, Representative images of WT and N/O/Tx (NDP52/OPTN/TAX1BP1) TKO cells expressing mCh-Parkin and GFP-LC3C were immunostained for Tom20 (n=3) and e, quantified for GFP-LC3C translocation to mitochondria. Quantification in e is displayed as mean ± s.d. from 3 independent experiments and use one-way ANOVA tests (***P<0.001). OA, Oligomycin and Antimycin A. Scale bars, 10 μm. Extended Data Figure 9 GABARAPs do not translocate to damaged mitochondria and early stages of autophagosome biogenesis mediated by WIPI1 and DFCP1 are inhibited in autophagy receptor deficient cell lines Representative images of WT, N/O (NDP52/OPTN) DKO and pentaKOs expressing mCherry-Parkin (mCh-Parkin) and either (a) GFP-tagged GABARAP, GABARAPL1 or GABARAPL2, (b) GFP-WIPI1 or (c) GFP-DFCP1 immunostained for Tom20 (n=3 for each condition, see Figure 4b, c for quantification of b and c). d, mCh-Parkin cell lines as indicated were subjected to either Phos-Tag SDS-PAGE or standard SDS-PAGE followed by immunoblotting. Arrows indicate the position of phosphorylated Beclin species. e, Representative images of untreated WT, N/O (NDP52/OPTN) DKO and pentaKO cell lines expressing mCh-Parkin and GFP-ULK1 were immunostained for Tom20 and GFP (n=3). OA, Oligomycin and Antimycin A. Scale bars, 10 μm. Extended Data Figure 10 OPTN and NDP52 rescue DFCP1 and ULK1 recruitment deficit in pentaKOs a, Representative images of pentaKOs expressing mCherry-Parkin (mCh-Parkin), GFP-DFCP1 and the indicated FLAG/HA-tagged autophagy receptors immunostained for HA (n=2). Right-hand panels display co-localization of FLAG/HA-tagged constructs and GFP-DFCP1 by fluorescence intensity line measurement. b, Representative images of pentaKOs expressing mCherry-Parkin and GFP-ULK1 were rescued with FLAG/HA-OPTN, FLAG/HA-NDP52, and FLAG/HA-p62, and immunostained for HA and GFP. Arrows indicate HA-tagged receptor puncta (n=2). Right panels display colocalization of HA and GFP by fluorescence intensity line measurement. c, d, Representative images of pentaKOs stably expressing FRB-Fis1 and transiently expressing PINK1Δ110-YFP-2xFKBP and vector or myc-tagged receptors, were (c) untreated or (d) treated with rapalog and imaged live (n=3, see Figure 4h, i for quantification of c, d). OA, Oligomycin and Antimycin A. Scale bars, 10 μm. e, Old and new models of PINK1/Parkin mitophagy. The old model is dominated by Parkin ubiquitination of mitochondrial proteins. Here PINK1 plays a small initiator role whose main function is to bring Parkin to the mitochondria. The new model depicts Parkin-dependent and independent pathways leading to robust and low-level mitophagy, respectively. Based on our data, PINK1 is central to mitophagy both before and after Parkin recruitment by phosphorylating UB to recruit both Parkin and autophagy receptors mitochondria, to induce clearance. In the absence of Parkin (right panel), this occurs at a low level due to the relatively low basal UB on mitochondria. When Parkin is present it serves to amplify the PINK1 generated UB-PO4 signal, allowing for robust and rapid mitophagy induction. Supplementary Material 1 2 supp_table

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Journal ID (nlm-ta): Genes Dev

Journal ID (iso-abbrev): Genes Dev

Journal ID (hwp): genesdev

Journal ID (pmc): genesdev

Journal ID (publisher-id): GAD

Title: Genes & Development

Publisher: Cold Spring Harbor Laboratory Press

ISSN (Print): 0890-9369

ISSN (Electronic): 1549-5477

Publication date (Print): 1 September 2016

Volume: 30

Issue: 17

Pages: 1913-1930

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[1 ]Abramson Cancer Center, Perelman School of Medicine, University of Pennsylvania, Philadelphia, Pennsylvania 19104, USA;

[2 ]Department of Medicine, Perelman School of Medicine, University of Pennsylvania, Philadelphia, Pennsylvania 19104, USA;

[3 ]Perlmutter Cancer Center, New York University Langone Medical Center, New York, New York 10016, USA;

[4 ]Department of Radiation Oncology, New York University Langone Medical Center, New York, New York 10016, USA;

[5 ]Rutgers Cancer Institute of New Jersey, New Brunswick, New Jersey 08903, USA;

[6 ]Department of Molecular Biology and Biochemistry, Rutgers University, Piscataway, New Jersey 08854, USA

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Corresponding author: eileenpwhite@ 123456gmail.com

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Medline ID: 8711660

DOI: 10.1101/gad.287524.116

PMC ID: 5066235

PubMed ID: 27664235

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Funded by: National Institute of Health http://dx.doi.org/10.13039/100000002

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