Autophagy is a major catabolic pathway by which eukaryotic cells degrade and recycle macromolecules and organelles. This pathway is activated under environmental stress conditions, during development and in various pathological situations. In this study, we describe the role of reactive oxygen species (ROS) as signaling molecules in starvation‐induced autophagy. We show that starvation stimulates formation of ROS, specifically H2O2. These oxidative conditions are essential for autophagy, as treatment with antioxidative agents abolished the formation of autophagosomes and the consequent degradation of proteins. Furthermore, we identify the cysteine protease HsAtg4 as a direct target for oxidation by H2O2, and specify a cysteine residue located near the HsAtg4 catalytic site as a critical for this regulation. Expression of this regulatory mutant prevented the formation of autophagosomes in cells, thus providing a molecular mechanism for redox regulation of the autophagic process.
Autophagy is a major pathway for delivery of proteins and organelles to lysosomes/the vacuole, where they are degraded and recycled. Autophagy plays an essential role in differentiation and development, as well as in cellular response to stress. It is activated during amino‐acid deprivation and has been associated with neurodegenerative diseases, cancer, pathogen infections and myopathies (reviewed in Cuervo, 2004; Shintani and Klionsky, 2004). Autophagy is initiated by the surrounding of cytoplasmic constituents by the crescent‐shaped isolation membrane/phagophore, which forms a closed double‐membrane structure, called autophagosome. Finally, the autophagosome fuses with a lysosome to become an autolysosome, and its content is degraded by lysosomal hydrolases.
The molecular mechanism underlying autophagy has been extensively researched in the past decade, and the genes participating in this process, denoted ATGs (autophagy‐related; Klionsky et al, 2003), were found to be conserved from yeast to man (Ohsumi, 2001; Huang and Klionsky, 2002). Yet, the roles played by the different gene products and their modes of action are to be resolved. A hallmark event in the autophagic process is the reversible conjugation of the Atg8 family of proteins to the autophagosomal membrane (Ichimura et al, 2000). This ubiquitin‐like (UBL) protein family, which has been implicated in a variety of cellular processes, includes Atg8p in yeast (Lang et al, 1998; Ichimura et al, 2000) and GATE‐16 (Golgi‐associated ATPase enhancer of 16 kDa; Sagiv et al, 2000), LC3 (light‐chain 3; Mann and Hammarback, 1994) and GABARAP (GABA receptor‐associated protein; Wang et al, 1999) in mammals. All Atg8 homologues (hereafter termed Atg8s) serve as substrates for the Atg4 family of cysteine proteases (Kirisako et al, 2000; Hemelaar et al, 2003; Marino et al, 2003; Scherz‐Shouval et al, 2003). Atg4s cleave Atg8s near the C‐terminus, downstream of a conserved glycine. This cleavage allows the conjugation of Atg8 to phosphatidylethanolamine (PE) through the exposed glycine, a process mediated through a ubiquitination‐like mechanism. Atg8‐PE serves as another substrate for Atg4, which cleaves Atg8 and releases it from the membrane. Notably, this conjugation process must be preceded by another ubiquitination‐like process, conjugating Atg12 to Atg5 (Tanida et al, 2004b). In mammalian cells, amino‐acid deprivation induces lipidation of LC3 (Kabeya et al, 2000) as well as of GATE‐16 and GABARAP (Kabeya et al, 2004). Lipidated Atg8s associate with phagophaores and autophagosomes and remain there until fusion with lysosomes, at which point intra‐autophagosomal Atg8 is probably degraded (Kabeya et al, 2000, 2004).
At least four Atg4 mammalian homologues have been reported based on sequence homology to the yeast Saccharomyces cerevisiae (Sc)Atg4. Two of the homologues, HsAtg4A and HsAtg4B, were shown to cleave the three mammalian Atg8s with different efficiencies: HsAtg4A cleaves mainly GATE‐16, whereas HsAtg4B cleaves all three homologues (GATE‐16, GABARAP and LC3), with the highest efficiency for LC3 (Kabeya et al, 2004).
Atg4s act both as conjugating and deconjugating enzymes and therefore their activity is expected to be tightly regulated. Hence, in the process of autophagy, following the initial cleavage of Atg8‐like proteins, Atg4 must become inactive so as to ensure the conjugation of Atg8 to the autophagosomal membrane. Later on, as the autophagosome fuses with the lysosome, Atg4 might be locally re‐activated in order to delipidate and recycle Atg8. How is this regulation achieved? We have previously shown that recombinant HsAtg4A is active as a cysteine protease of GATE‐16 only in the presence of the reducing agent DTT (Scherz‐Shouval et al, 2003). This could implicate that, in vivo, the delipidating activity of Atg4 is regulated through changes in redox potentials that take place under different conditions and at specific subcellular microenvironments. Such regulation could be inflicted, among other factors, by reactive oxygen species (ROS). At high levels, ROS are deleterious to cells, leading to programmed cell death (PCD) (Jabs, 1999; Lee et al, 2003; Macip et al, 2003). At low levels, however, ROS can serve as signaling molecules, by oxidizing factors in a variety of pathways that lead to growth and survival. Here, we report for the first time the involvement of ROS as signaling molecules in nutrient starvation‐induced autophagy, which is essentially a survival pathway. We show that under starvation conditions, cells form ROS, specifically H2O2, which is essential for autophagosome formation and autophagic degradation. The oxidative signal is partially PI3K dependent and leads to inhibition of Atg4. Using an in vitro assay, we could demonstrate that H2O2 directly regulates HsAtg4A. Moreover, we identified Cys81, situated four amino acids downstream of the active site, as an essential residue for the redox regulation of HsAtg4A. Expression of HsAtg4A mutated in Cys81, or the corresponding mutant of HsAtg4B, inhibited the formation of GATE‐16‐ or LC3‐labeled autophagosomes in cells.
Starvation induces formation of ROS
ROS serve as signaling molecules in a variety of cellular processes, including growth, differentiation, adhesion and PCD. To test whether ROS play a role in autophagy, we measured ROS production under starvation using a fluorescent probe, dihydroethidum (DHE). DHE reacts with peroxides to form a compound that upon binding to DNA fluoresces and can be visualized by confocal fluorescent microscopy (Vanden Hoek et al, 1997). Indeed, upon nutrient starvation, DHE staining was evident, indicating that cells accumulate peroxides under these conditions (Figure 1A). We then used another probe, 2′,7′‐dichlorofluorescin diacetate (DCF‐DA), which reacts mainly with H2O2 to form a fluorescent compound at the point of interaction (Cathcart et al, 1983; Vanden Hoek et al, 1997). DCF staining was already evident 15 min after the induction of starvation (Figure 1Bd and Supplementary Figure 1A, see text below). The pattern of DCF staining was punctated as well as cytosolic, and it was also apparent following overnight starvation (Figure 1Bh). These results indicate that H2O2 is most likely the oxidative agent accumulating in starved cells. To rule out the possibility that the oxidative signal associated with starvation is leading to PCD, we tested the viability of cells grown under nutrient starvation conditions. As shown in Supplementary Figure 1B, 5 h of nutrient starvation did not affect the viability of the cells, and the percent of cell death was equally low in control and starved cells. Taken together with the finding that the oxidative signal appears minutes after induction of starvation, these results suggest that H2O2 serves as a signaling molecule in the autophagic pathway, and not as part of an oxidative stress leading to PCD.
The pattern of DCF staining resembled mitochondrial staining, and therefore we double stained the cells with DCF and MitoTracker Red, a marker for functional mitochondria. We found significant overlap between the two markers (Figure 1Bf and j; enlargements in g and k, respectively). All the DCF puncta either colocalized with or were in close vicinity to MitoTracker‐stained structures, suggesting mitochondria as the source of ROS generated during starvation. This finding coincides with our observation that phagophores, identified by a double staining with Atg5 and Atg16 (Mizushima et al, 2003), tend to colocalize with MitoTracker Red‐stained mitochondria (Supplementary Figure 1C), suggesting a functional link between ROS generated in the mitochondria during starvation and autophagosome formation. This result is consistent with recently reported findings (Kissova et al, 2006).
In order to assess the rise in ROS in a more accurate manner, we measured the increase in DCF fluorescence using a fluorimeter set at 485 nm excitation and 535 nm emission in two different cell lines (CHO and HeLa). The cells were either deprived of serum, completely starved (deprived of serum and nutrients) or grown in control medium for 2 h, after which they were treated with DCF and the fluorescent signal of the whole population (3 × 104 cells) was monitored. As shown in Figure 1C, complete starvation of the cells resulted in a dramatic increase in the fluorescent signal over time, as compared to the control, whereas serum starvation had only a mild effect on the fluorescent signal, indicating a slight increase in ROS production. These measurements were further processed and summarized as the increase in DCF fluorescence over 20 min of incubation with the reagent (Figure 1D).
Class III PI3K is involved in starvation‐dependent ROS formation
In the attempt to understand where ROS production fits in the autophagic signaling pathway, we monitored the effect of two known autophagy inhibitors, wortmannin and 3‐methyladenine (3MA), on starvation‐dependent DCF staining (Petiot et al, 2000). Starvation of cells for a 2‐h period in the presence of either drug reduced the level of DCF fluoresence by 25–30% (Figure 2A, left panel). As expected, the autophagic activity in the treated cells, tested by the rate of degradation of long‐lived proteins (known to increase during autophagy, see Mizushima et al, 2001), was significantly reduced (Figure 2A, right panel). Wortmannin and 3MA are phosphatidyl inositol 3 kinase (PI3K) inhibitors. Class III PI3K plays a critical role in the early stages of autophagosome formation in mammals through formation of an essential complex with Beclin 1 and, hence, inhibition of its activity inhibits the autophagic process (Petiot et al, 2000; Tassa et al, 2003). This finding, therefore, indicates that ROS production is partially PI3K dependent. Indeed, partial silencing of Beclin 1 (previously shown to cause a partial autophagic defect; Reef et al, 2006) or the PI3K hVps34 resulted in 15 and 20% (respectively) reduction in starvation‐induced ROS production (Supplementary Figure 2A). Notably, partial silencing of hVps34 caused 30% reduction in the number of autophagosomes per cell (Supplementary Figure 2B, lower right panel).
Next, we assessed the level of DCF staining in cell lines deleted for the Atg5 gene, whose product is essential for autophagosome formation, acting downstream of the class III PI3K (Suzuki et al, 2001; Tanida et al, 2004b). DCF staining in the mutant cells did not decrease, as compared with control cells (Figure 2B), but rather increased significantly (40–50%), indicating that ROS production takes place upstream of the Atg5‐dependent step of autophagosome formation, and suggesting that in the absence of a functional autophagic pathway, cells accumulate ROS.
ROS formation is essential for autophagy
The results described above suggest that starvation induces ROS formation. But are ROS essential for autophagy? To address this question, we tested the effect of N‐acetyl‐l‐cysteine (NAC), a general antioxidant, and catalase, which specifically decomposes H2O2, on the formation of autophagosomes using GATE‐16 and LC3 as markers. LC3 and GATE‐16 were shown to label autophagosomes under starvation conditions (Kabeya et al, 2004; Supplementary Figure 3A), and as shown in Supplementary Figure 3B, the two proteins exhibit significant colocalization under these conditions. Addition of either NAC or catalase to the growth medium (Sakurai and Cederbaum, 1998; Preston et al, 2001; Xu et al, 2003) of the cells for 10 min before a 2‐h starvation period resulted in reduced lipidation of GATE‐16 and LC3 and abolished the formation of GATE‐16‐ and LC3‐labeled autophagosomes (Figure 3A and Supplementary Figure 3C). As expected, the same concentration of antioxidative reagents significantly reduced the level of ROS in the treated cells (Figure 3B). These results were further validated by testing the effect of antioxidants on starvation‐induced protein degradation (Figure 3C). Treatment with NAC caused 60% inhibition in starvation‐induced degradation, and catalase caused 25% inhibition in starvation‐induced degradation. Taken together, we conclude that starvation conditions induce formation of ROS, which are required for induction of autophagy. Notably, treatment of cells with H2O2 alone did not result in formation of autophagosomes (data not shown).
Atg4 is attenuated in response to starvation in a redox‐dependent manner
Next, we searched for a cellular target of the starvation‐induced oxidative signal. The amount of the lipidated form of Atg8s increases significantly during starvation (Kabeya et al, 2004). As shown in Figure 4A, this elevation, in mammalian Atg8s, does not result from increased transcription, as previously shown for ScAtg8 (Kirisako et al, 1999). Consistently, we could not observe changes in the protein levels of mammalian Atg8s under these conditions (Figure 4B and data not shown). We therefore deduce that the increase in lipidated mammalian Atg8 proteins in response to nutrient starvation is due to post‐translational regulation of the processing enzymes. To test this hypothesis, cells were starved for 3 or 13 h, after which they were lysed and the lysates were tested for their ability to cleave GATE‐16. In this assay, recombinant GATE‐16, tagged at both termini (His6‐GATE‐16‐HA), is used so that cleaved GATE‐16 can be detected by its faster mobility in SDS–PAGE. As shown in Figure 4C, incubation of GATE‐16 in a lysate prepared from cells that were starved for a short period of 3 h resulted in a decrease of up to 40% in the cleavage activity as compared with control cells. Starvation of cells for longer periods of 13 h (during which the cells are still viable) resulted in further inhibition in cleavage activity. However, addition of 1 mM DTT to the lysates brought about recovery of the cleavage activity, suggesting that the inhibition of Atg4 was caused by oxidation of the protein. Consistently, treatment of cells with 1 mM H2O2 for 1 h before lysis resulted in strong inhibition of the cleavage activity of Atg4 as compared with control cells. Addition of DTT to the lysate restored most of the lost activity, confirming that the inhibition by oxidation is reversible. Notably, the protein levels of the GATE‐16‐specific protease, HsAtg4A, did not change under any of the tested conditions (Supplementary Figure 4A), supporting the idea that the regulation of Atg4 activity is post‐translational. Specific anti‐HsAtg4A antibodies were utilized to confirm that the cleavage activity shown in Figure 4C is mediated by HsAtg4A (for details see Supplementary Figure 4B), indicating that Atg4 is a target for redox regulation under starvation, and suggesting that accumulation of lipidated Atg8s under starvation might result from inhibition of Atg4 activity. Indeed, we found that treatment with 1 mM H2O2 resulted in accumulation of lipidated LC3 (Supplementary Figure 4C).
Atg4 acts both in the initial processing of Atg8s (priming) and in the delipidation step. The priming step was previously shown to occur immediately after translation under nutrient‐rich conditions (Kabeya et al, 2000). To test whether this step is also inhibited during starvation, we transfected cells with GATE‐16 tagged with Myc at its N‐terminus and HA at the C‐terminus, starved them for 30 min (a time period in which autophagosomes are expected to form; Fass et al, 2006) or 13 h and subjected them to a [35S]methionine pulse–chase experiment as detailed in Materials and methods (Figure 4D). The control experiment shows that two bands at approximately 19 kDa are detected immediately after pulse labeling in the transfected cells (Figure 4D, left panel). A 1 h chase resulted in disappearance of the upper band, suggesting that this band corresponds to unprimed GATE‐16 (Myc‐GATE‐16‐HA), whereas the lower band corresponds to primed GATE‐16 (Myc‐GATE‐16). Short starvation does not induce accumulation of the unprimed form (Figure 4D, middle panel), and this form accumulates only in cells that were starved for a long period of 13 h.
These results indicate that HsAtg4A is redox regulated during starvation. As the priming activity is not affected by short starvation, we conclude that the delipidation activity is the main target of this regulation.
To characterize the redox regulation of HsAtg4, we expressed HsAtg4A as a recombinant protein and tested its activity in the cell‐free cleavage assay. As depicted in Figure 5A, the protease was gradually activated upon reduction of the reaction mixture and the activity saturated at 1 mM DTT. Next, we tested the inhibition of HsAtg4A activity by H2O2. The protein was preincubated in a low concentration of DTT for activation followed by addition of H2O2 for 5 min before incubation with GATE‐16 for the cleavage assay. The activity of HsAtg4A decreased as the H2O2 concentration increased above 100 μM (Figure 5B). In some experiments, a slight increase in the protease activity was detected upon addition of 30 μM H2O2 to the reaction mixture. Finally, we tested the reversibility of HsAtg4 inhibition by H2O2. In this experiment, the protein was preincubated in a low concentration of DTT (Figure 5C, lane 2) followed by addition of H2O2 (lane 3) and then addition of a high concentration of DTT (lane 4; each incubated for 5 min before addition of the next reagent) before incubation with GATE‐16. Under these conditions, HsAtg4A completely regained its activity. Taken together, these results show that reducing conditions activate HsAtg4A, whereas an oxidizing environment inhibits its activity. Moreover, these results indicate that H2O2 alone is sufficient to reversibly inhibit the activity of HsAtg4A in vitro, and therefore suggest that H2O2 might also act directly on the protease in vivo.
Cys81 is a target for the redox regulation of HsAtg4A
HsAtg4A contains 12 cysteines, seven of which are highly conserved among tetrapod homologues of Atg4A and Atg4B. The latter share an overall high conservation level with respect to the whole family (Figure 6). One of these conserved residues is the putative active residue of the protease Cys77 (Kirisako et al, 2000; Tanida et al, 2004a), the only cysteine conserved also in the yeast S. cerevisiae (Figure 6A, large arrow, numbering is according to HsAtg4B). Another conserved residue (Cys81 in HsAtg4A) is located four sites downstream of the active residue. We first sought to determine whether Cys77 is the catalytic residue, as suggested by homology to ScAtg4 and HsAtg4B. As shown in Figure 7A, recombinant HsAtg4A harboring a mutation of cysteine to alanine at position 77 (His6‐HsHsAtg4AC77A) did not cleave GATE‐16 in vitro, neither in the absence nor in the presence of DTT. Based on this result and the sequence homology described above, we conclude that Cys77 is part of the active site of HsAtg4A.
To study the mechanism of redox regulation of HsAtg4A, we mutated the conserved cysteine residues 81 and 92 (Figure 6A, small arrows), which are closest to the active residue, to serine (His6‐HsAtg4AC81S and His6‐HsAtg4AC92S, respectively) and tested their activity in the cell‐free cleavage assay of GATE‐16 under different redox conditions. As shown in Figure 7A, mutation in Cys81 significantly reduced the redox sensitivity of HsAtg4A so that His6‐HsAtg4AC81S was partially active even in the absence of DTT. Mutation in Cys92, however, had no effect on the regulation and the protease remained under the same conditions as active as His6‐HsAtg4AWT (data not shown). Next, we tested the effect of oxidation by H2O2 on the activity of His6‐HsAtg4AC81S. We found that the mutant protein retained significant activity in the presence of H2O2 (Figure 7B) under conditions in which the WT was completely inhibited (Figure 5C). Similar to the WT, the partial inhibition of His6‐HsAtg4AC81S by H2O2 could be reversed by addition of DTT. From these results, we conclude that Cys81 is important for redox regulation of HsAtg4A in vitro.
HsAtg4A and HsAtg4B are both redox regulated during autophagy
To examine whether Cys81 plays a role in the redox regulation of HsAtg4A in vivo, we monitored the formation of GFP‐GATE‐16‐labeled autophagosomes in cells transfected with HsAtg4A mutants. Expression of HsAtg4AC81S significantly impaired the formation of GATE‐16‐labeled autophagosomes in starved cells (Figure 8A), as compared with cells expressing HsAtg4AWT (Figure 8A) or HsAtg4AC92S (data not shown) at similar levels. The average number of autophagosomes formed in HsAtg4AC81S‐expressing cells was 70% lower than in HsAtg4AWT‐expressing cells (Figure 8A, lower panel). One could suggest that in the presence of HsAtg4AC81S, autophagosomes are formed, but then rapidly degraded, and therefore cannot be seen. To exclude this possibility, we inhibited the degradation of autophagosomes in the lysosome using bafilomycin A (Mousavi et al, 2001). This treatment did not affect the mutant phenotype, indicating that autophagosomes do not form in cells transfected with HsAtg4AC81S (data not shown). The finding that GATE‐16‐labeled autophagosomes do not form in HsAtg4AC81S‐expressing cells was further supported by GATE‐16 lipidation measurements. As the HsAtg4AC81S mutant is less sensitive to oxidation, we expect the delipidation activity to be higher in starved cells transfected with this mutant as compared with cells transfected with HsAtg4AWT. Indeed, we found nearly 40% less of the lipidated form of GATE‐16 in cells transfected with HsAtg4AC81S as compared with cells transfected with HsAtg4AWT (Figure 8B). It should be mentioned that expression of HsAtg4AC81S did not affect the pattern of LC3 staining in the cells (data not shown), as expected, as HsAtg4A is specific for GATE‐16 and does not process LC3. Taken together, our findings support the notion that Cys81 takes part in the redox regulation of HsAtg4A during autophagy.
In order to extend our study to other Atg4 family members, we decided to test the LC3‐specific protease, HsAtg4B. To that end, we transfected cells with GFP‐HsAtg4BWT, GFP‐HsAtg4BC78S or with GFP as control and tested the effect on lipidation of LC3 and formation of LC3‐labeled autophagosomes. We found that overexpression of both GFP‐HsAtg4BWT and GFP‐HsAtg4BC78S reduced autophagosome formation and lipidation of LC3 (data not shown). This phenomenon was previously reported for overexpression of HsAtg4BWT (Tanida et al, 2004a). To overcome this problem, we added H2O2 to the starvation media. Under these conditions, HsAtg4BWT was inhibited and a normal autophagic phenotype was observed (Figure 9). HsAtg4BC78S, however, was resistant to H2O2 and the cells expressing this construct failed to form lipidated LC3 and LC3‐labeled autophagosomes. These results indicate that HsAtg4B, similar to HsAtg4A, is redox regulated.
Autophagy is a unique pathway of intracellular trafficking activated in response to extracellular signals. Although many of the proteins involved in this process have been identified, the signal transduction pathway leading to activation of autophagy is not fully solved. Here we demonstrate, for the first time, the involvement of ROS as signaling molecules in starvation‐induced autophagy. We show that starvation triggers accumulation of ROS, most probably H2O2, which is necessary for autophagosome formation and the resulting pathway of degradation. The oxidative signal is partially PI3K dependent, located upstream of the Atg5‐dependent conjugation machinery. Furthermore, we identify a direct target for oxidation by H2O2—the protease Atg4.
Based on our results, we propose the following model, illustrated in Figure 10: nutrient starvation induces complex formation between class III PI3K and Beclin 1, which in turn leads, together with other signals, to a local rise in H2O2 in the vicinity of mitochondria. This oxidative signal leads to inactivation of Atg4 at the site of autophagosome formation, thereby promoting lipidation of Atg8, an essential step in the process of autophagy. As the autophagosome matures towards fusion with the lysosome, its localization apparently changes to an H2O2‐poor environment where Atg4 is active and can delipidate and recycle Atg8.
How does H2O2 regulate Atg4? Atg4s are cysteine proteases that share several conserved cysteine residues. One of these is the catalytic residue, found previously in other members of the Atg4 family and shown here to be Cys77 in HsAtg4A. Another conserved motif is a cysteine residue located four amino acids from the catalytic residue, Cys81 in HsAtg4A. Remarkably, we found this residue to be crucial for the redox regulation of HsAtg4A. Mutation of this residue to serine significantly impaired the sensitivity of the protein to H2O2 in vitro, and abolished the formation of GATE‐16‐labeled autophagosomes in cells. The essentiality of this residue for proper redox regulation was further confirmed by the finding that the corresponding HsAtg4B mutant (HsAtg4BC78S) exhibited a similar autophagic defect. Our in vitro study shows that H2O2 directly inactivates Atg4. This could be accomplished either by binding of H2O2 to Cys77 or to Cys81, forming reversible sulfenic acid that shields Cys77, or by oxidation, which leads to other modifications such as formation of a disulfide bond between Cys77 and Cys81, again shielding Cys77. Such a disulfide bond, however, was not detected in our experiments (unpublished data). Alternatively, H2O2 might be affecting a different site, which is altered when Cys81 is mutated. The active site, Cys77, is conserved from yeast to man, whereas Cys81 is conserved only in tetrapod homologues of Atg4A and Atg4B. Lower species harbor a conserved Ser or Thr in this residue and hence the choice of a C81S mutant. We thus propose that the redox regulation may be mediated via Cys77 in lower species, whereas tetrapods evolved a more complex mechanism of regulation that requires both Cys77 and Cys81, where the latter provides tetrapods with higher sensitivity to ROS than lower species. The recently solved three‐dimentional structure of HsAtg4B (Sugawara et al, 2005; Kumanomidou et al, 2006) supports our model, as it shows Cys77 and Cys81 to be situated at the beginning of an α‐helix, both facing the same plane of the helix. With such close proximity between the two cysteines, modifications on Cys81 could indeed affect the active site. Notably, both crystals were prepared under reducing conditions, and consequently no disulfide bridges or other modifications were reported for any of the cysteine residues.
Cysteine‐harboring proteins serve as redox sensors. They can undergo rapid, variable and, most importantly, reversible post‐translational modifications in response to changes in the oxidative conditions of the environment (Sitia and Molteni, 2004). Several cysteine proteases, including cathepsin D, cathepsin B and the cytosolic caspase‐3 (Chandler et al, 1998) and calpains (Glading et al, 2001), have been shown to undergo redox regulation through a direct modification of the active site (reviewed in Giles et al, 2003). Redox regulation of Atg4 can provide rapid activation and inactivation of this protease, as part of the signaling pathway, leading to induction of the autophagic process.
A growing body of evidence suggests that autophagy plays a dual role, both in survival and in death‐related pathways (Cuervo, 2003; Codogno and Meijer, 2005; Reef et al, 2006). The data available till now, linking ROS with autophagy, describe the death‐related pathway, known as type II PCD, which is induced by high levels of ROS (i.e. oxidative stress; Yu et al, 2004; Kiffin et al, 2006). Recently, it has been shown that caspase inhibition leading to cell death through autophagy involves accumulation of ROS, owing to catalase‐specific autophagic degradation. In that study, however, starvation did not lead to a similar effect (Yu et al, 2006). In our system, the rise in ROS is both local and reversible; it is not deleterious to cells and serves to oxidize a specific target. Therefore, our study provides the first indication for the involvement of ROS in starvation‐induced autophagy as signaling molecules in a survival pathway.
Materials and methods
All experiments presented in the Results section were repeated at least three times unless indicated otherwise.
Expression and purification of recombinant proteins
Recombinant His6‐HsAtg4A (WT and the mutants described in Supplementary data) and His6‐GATE‐16‐HA were expressed from the pQE‐30 plasmid and purified as described previously (Scherz‐Shouval et al, 2003). Purification of His6‐HsAtg4A (WT and mutants) was carried out in the presence of 10 mM βME to prevent formation of nonspecific disulfide bonds; however, before its use, the protein was diluted to at least 30‐fold or dialyzed in a buffer lacking βME.
Cell culture, transfection, reagents and immunofluorescence
All cell lines were grown in α‐MEM medium (Biological Industries, Israel) supplemented with 10% fetal calf serum and referred to as control medium. Atg5−/− MEFs and anti‐Atg16 polyclonal antibodies were a kind gift from the laboratory of Dr Mizushima (The Tokyo Metropolitan Institute of Medical Science). Stable clones of GFP‐GATE‐16‐, GFP‐LC3‐ or YFP‐Atg5‐transfected CHO cells were selected in 1 mg/ml geneticin (G418). Starvation was carried out in EBSS. Details regarding the use of different reagents for tissue culture treatments are described in Supplementary data. Transfections of CHO cells were preformed with lipofectamine (Invitrogen) according to the manufacturer's protocol. Transfections of HeLa and HEK 293 cells were performed by the standard calcium phosphate technique. For immunofluoresence, unless described otherwise, cells were fixed in methanol, permeabilized with acetone and then incubated with the indicated antibodies and visualized by a Nikon eclipse TE300 fluorescent microscope or by an Olympus IX‐70 confocal microscope.
Hydrogen peroxide measurements
H2O2 was measured using 50 μMDHE (Molecular Probes) and 30 μM DCFDA (Molecular Probes). See Supplementary data for details. The fluorescent signal was detected by an Olympus IX‐70 confocal microscope. Each experiment was repeated at least three times and representative images are shown. The DCFDA signal was also measured by a SPECTRAmax gimini fluorimeter, set to 485 excitation and 535 emission and kept at 37°C, for 40 min. Data collected from the fluorometric measurements were analyzed as detailed in Supplementary data. The results presented for all fluorimetric measurements are the means±s.d. of at least three experiments in duplicate or triplicate.
Protein degradation assay
The assay was preformed on CHO cells according to the protocol described by Fass et al (2006). When required, 10 mM 3MA, 100 nM wortmannin, 1000 μ/ml catalase or 10 mM NAC were added. Notably, NAC was added for 10 min before starvation induction and catalase was added 14 h before starvation. [14C]valine release was calculated as a percentage of the radioactivity in the TCA‐soluble supernatant relative to the total cell radioactivity.
Preparation of cell lysates
Cells were homogenized in Ripa buffer (0.1 M NaCl, 5 mM EDTA, 0.1 M Na2HPO4/NaH2PO4, pH 7.5, 1% Triton, 0.5% DOC, 0.1% SDS and protease inhibitors) by vortexing on ice. The homogenates were centrifuged for 15 min at 20 000 g and the supernatant containing the lysate was collected.
RNA was isolated by Tri‐reagent (Molecular Research Center Inc.) as detailed in Supplementary data.
HeLa cells were transiently transfected with Myc‐GATE‐16‐HA or with an empty vector. Cells were then kept in full medium for 24 h, after which they were either incubated in methionine‐free medium for 10 min or starved in EBSS for 30 min (2 × 10 cm plates). For long starvation, the cells (three plates) were shifted to EBSS 10 h post‐transfection for 13 h, after which the medium was replaced with fresh EBSS. All cells were then pulse‐labeled with 0.2 mCi of [35S]methionine for 10 min at 37°C, and either chased in full medium for 1 h at 37°C or lysed immediately on ice, in Ripa buffer supplemented with 0.5 mM N‐ethylmaleimide (sigma). Lysates were immunoprecipitated with anti‐Myc antibodies and analyzed on 15% SDS–PAGE.
Supplementary data are available at The EMBO Journal Online (http://www.embojournal.org).
Supplementary Figure 1
Supplementary Figure 2
Supplementary Figure 3
Supplementary Figure 4
We thank Vladimir Kish for invaluable guidance with the confocal microscope and Frida Shimron for her devoted assistance. We also thank Avihai Danon for his insights. This work was supported in part by grants from the Israel Science Foundation, Binational Science Foundation and by the Minerva center.
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