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H2S-Induced Sulfhydration of the Phosphatase PTP1B and Its Role in the Endoplasmic Reticulum Stress Response

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Science Signaling  13 Dec 2011:
Vol. 4, Issue 203, pp. ra86
DOI: 10.1126/scisignal.2002329


Although originally considered toxic, hydrogen sulfide (H2S) has been implicated in mediating various biological processes. Nevertheless, its cellular targets and mode of action are not well understood. Protein tyrosine phosphatases (PTPs), which regulate numerous signal transduction pathways, use an essential cysteine residue at the active site, which is characterized by a low pKa and is susceptible to reversible oxidation. Here, we report that PTP1B was reversibly inactivated by H2S, in vitro and in cells, through sulfhydration of the active-site cysteine residue. Unlike oxidized PTP1B, the sulfhydrated enzyme was preferentially reduced in vitro by thioredoxin, compared to glutathione or dithiothreitol. Sulfhydration of PTP1B in cells required the presence of cystathionine γ-lyase (CSE), a critical enzyme in H2S production, and resulted in inhibition of phosphatase activity. Suppression of CSE decreased H2S production and decreased the phosphorylation of tyrosine-619 in PERK [protein kinase–like endoplasmic reticulum (ER) kinase], thus reducing its activation in response to ER stress. PERK, which phosphorylates the eukaryotic translational initiation factor 2, leading to attenuation of protein translation, was a direct substrate of PTP1B. In addition, CSE knockdown led to activation of the nonreceptor tyrosine kinase SRC, previously shown to be mediated by PTP1B. These effects of suppressing H2S production on the response to ER stress were abrogated by a small-molecule inhibitor of PTP1B. Together, these data define a signaling function for H2S in inhibiting PTP1B activity and thereby promoting PERK activity during the response to ER stress.


Gasotransmitters are a class of gaseous signaling molecules that freely permeate membranes and, therefore, unlike many other regulators of signal transduction, act independently of transmembrane receptors (1). Gasotransmitters are generated enzymatically, in a regulated manner, and act through specific molecular targets. The classic example is nitric oxide (NO), a well-established signaling molecule generated by NO synthases. NO was identified as an endothelium-derived relaxing factor (EDRF) that functions by activating guanylyl cyclase, thereby stimulating the production of guanosine 3′,5′-monophosphate (cGMP). Subsequently, NO was shown to influence many cellular processes in various tissues (2). The study of NO paved the way for identification of other gaseous signaling molecules, including carbon monoxide (CO) and hydrogen sulfide (H2S) (1, 35). Although known for its toxic properties, at subtoxic concentrations, H2S regulates a broad spectrum of physiological events (68).

The ability to synthesize H2S is found in representatives of all evolutionary kingdoms (911), suggesting an ancient metabolic capability. In mammals, H2S biosynthesis is catalyzed predominantly by two pyridoxal phosphate–dependent enzymes, cystathionine β-synthase (CBS) and cystathionine γ-lyase (CSE), which together constitute a transulfuration pathway that provides a route for converting dietary methionine to cysteine. CBS predominates in the central nervous system, whereas both enzymes are found in peripheral tissues. Their activity is accompanied by production of H2S at micromolar concentrations (up to ~300 μM) in various tissues (12). As in the case of NO, the original function characterized for H2S was vasorelaxation; in fact, it has been proposed as a new EDRF, an effect mediated through activation of an adenosine 5′-triphosphate (ATP)–sensitive potassium channel (KATP), which lowers blood pressure (13). H2S has been implicated in the control of cell proliferation and survival in cardiomyocytes (12). It is also produced at sites of inflammation; indeed, drugs with the potential to trigger release of H2S have been suggested as potential anti-inflammatory agents (11, 14, 15). Intriguingly, treatment with H2S induces a suspended animation–like state in mice by decreasing metabolic rate and core body temperature, effects that depend on its reversible inhibition of cytochrome c oxidase (16). The effects of H2S are mediated through its sulfhydration of specific Cys residues in target proteins (17). For example, in the liver, which produces abundant H2S, 10 to 25% of actin, tubulin, and glyceraldehyde-3-phosphate dehydrogenase (GAPDH) are sulfhydrated in the basal state, a modification that enhances actin polymerization and GAPDH activity (17). Nevertheless, the identification of additional specific targets that mediate the various physiological functions of H2S and clarification of its roles in signaling remain important quests. Here, we tested the hypothesis that the protein tyrosine phosphatase (PTP) family of enzymes may provide a group of targets for H2S.

The PTPs comprise a large, structurally diverse family of transmembrane receptor–like and intracellular enzymes that are specific regulators of signal transduction and, in conjunction with the protein tyrosine kinases (PTKs), exert exquisite control over various biological functions (18). Members of the PTP family are characterized by a highly conserved catalytic domain containing a signature motif, His-Cys-(X)5-Arg-(Ser/Thr) (where X is any amino acid), in which the invariant cysteine has an unusually low pKa. This favors its function as a nucleophile to attack the phosphotyrosine substrate (18). The presence of this cysteine at the active site underlies an important aspect of PTP regulation. Covalent modification of the active-site cysteine abrogates its nucleophilic function and thereby inactivates the enzyme. The production of H2O2, in response to various growth factors, hormones, and cytokines that act through tyrosine phosphorylation, leads to the transient oxidation and inactivation of those PTPs that normally exert an inhibitory constraint on the signaling pathways. This transient PTP inactivation enhances the kinase-dependent tyrosine phosphorylation of components of the pathway and thereby enables fine-tuning of the response (19). Other agents that modify the active-site cysteine would also be expected to inhibit PTP activity. Indeed, NO inactivates PTPs by S-nitrosylation of the active-site cysteine, a modification that is proposed to provide a mechanism to prevent permanent inactivation of PTPs by irreversible oxidation (20). Here, we examined whether H2S-induced sulfhydration of PTP1B may also provide a mechanism for regulating tyrosine phosphorylation–dependent signaling.

We focused on PTP1B, which is the prototypic member of the PTP family (21) and is biomedically important because of its roles in inhibiting insulin and leptin signaling (21, 22), promoting HER2-mediated breast tumorigenesis (23, 24), and regulating other tyrosine phosphorylation–dependent signaling pathways (25). PTP1B is located on the cytoplasmic face of the endoplasmic reticulum (ER) and has been implicated in ER stress signaling (26). Three transmembrane ER proteins mediate distinct branches of the unfolded protein response (UPR) triggered by ER stress: inositol requiring-1 (IRE1), activating transcription factor 6 (ATF6), and double-stranded RNA–activated protein kinase–like ER kinase (PERK) (2730). Here, we demonstrate that PTP1B is reversibly sulfhydrated after production of H2S during the ER stress response. Sulfhydration inhibited PTP1B-mediated dephosphorylation of PERK specifically, without affecting the other arms of the UPR. Our study identifies sulfhydration as a redox modification that regulates PTP1B activity and, thus, the ER stress response, and illustrates a further aspect of the control of signaling in response to ER stress.


Inactivation of PTP1B by H2S

By measuring PTP1B phosphatase activity as a function of time and of H2S concentration, we observed an apparent second-order rate constant of 22.4 ± 1.8 M−1 s−1 for inactivation of PTP1B by H2S (Fig. 1A). The active-site cysteine of PTP1B, Cys215, undergoes reversible oxidation by H2O2 and reversible S-nitrosylation by NO (20); therefore, we compared the kinetics of PTP1B inactivation by H2O2 and NO with the rate of its inactivation by H2S (figs. S1 and S2). The rate of PTP1B inactivation by H2O2 was 10 ± 1.4 M−1 s−1 and that by NO was 2.1 ± 0.5 M−1 s−1; the overall extent of reversible inactivation was similar in all three treatments (~90%). Given the physiological roles of H2O2 and NO in regulating PTP1B (14, 15), this comparison suggested that inactivation by H2S may also be physiologically relevant. Furthermore, the data suggest that the active-site cysteine of PTP1B may display a degree of specificity in its reaction with oxidants.

Fig. 1

Redox-dependent inactivation and reactivation of PTP1B. (A) Time-dependent inactivation of PTP1B by H2S. The phosphatase activity of PTP1B was monitored in the presence of the following concentrations of H2S: 10 μM (•), 20 μM (▪), 50 μM (▴), 100 μM (▾), and 200 μM (♦). (Inset) The concentration dependence of the rate of inactivation was used to derive the second-order rate constant, 22.4 ± 1.8 M−1 s−1. (B) Time-dependent reactivation of PTP1B by DTT. The following concentrations of DTT were used: 5 mM (•), 10 mM (▪), 15 mM (▴), and 20 mM (▾). (Inset) The concentration dependence of the rate of reactivation was used to derive the second-order rate constant, 0.24 ± 0.1 M−1 s−1. For (A) and (B), the data were derived from three independent determinations. (C and D) Binding of fluorescently labeled phosphopeptide substrate (5′-FAM-ENDpYINASL) to wild-type (•), C215S (▪), or D181A (▴) mutant forms of PTP1B. The assays were performed in the absence of H2S (C) or after incubation with 100 μM H2S for 10 min (D).

We also used substrate-trapping mutant forms of PTP1B to examine the effects of H2S on substrate binding in an in vitro fluorescence polarization assay. In one of these mutants (D181A PTP1B), the invariant catalytic acid residue Asp181, which protonates the tyrosyl leaving group in the substrate, is mutated to Ala. This mutation impairs catalysis without affecting the affinity for substrate (31); therefore, the mutant forms stable complexes with tyrosine-phosphorylated substrates. A mutant in which the active-site Cys residue is changed to Ser (C215S PTP1B) also displays substrate-trapping properties (32). The fluorescence polarization assay assesses substrate binding on the basis of the difference in the rate of rotation of small fluorescently labeled species, such as the peptide substrate, and the larger complex between the PTP1B trapping mutant and the peptide substrate. When a fluorescent molecule is excited with plane-polarized light, it emits light in the same polarized plane only if it is stationary during the excitation of the fluorophore. As the molecule tumbles in solution, the polarization of the emitted light is lost. The large complex of the trapping mutant and peptide tumbles more slowly than the free peptide and so yields a higher fluorescence polarization signal than does the free peptide. Both C215S PTP1B and D181A PTP1B, but not the wild-type enzyme, formed a complex with a fluorescently labeled peptide substrate (Fig. 1C); however, only the peptide complex with the D181A mutant was disrupted by treatment with H2S, highlighting the importance of the active-site Cys for the inhibitory effects of H2S.

Reactivation of PTP1B by DTT, GSH, and TR/TRR

To examine the reversibility of PTP1B inactivation by H2S, H2O2, and NO, we compared the effects of varying concentrations of the reducing agents dithiothreitol (DTT), reduced glutathione (GSH), or the combination of thioredoxin and thioredoxin reductase (TR/TRR) on the rate of recovery of activity of inactivated PTP1B. In each case, the rate of PTP1B reactivation varied linearly as a function of the concentration of the reducing agent (Fig. 1B and figs. S3 and S4) and showed a second-order rate constant (Table 1). The reactivation of H2S-inactivated PTP1B was ~190-fold faster with TR/TRR than with DTT; however, such a difference was not observed for reactivation of oxidized or nitrosylated PTP1B (Table 1). The rates of reactivation of H2O2- and NO-treated PTP1B were similar, although with both H2O2- and NO-treated PTP1B, the rate of reactivation with glutathione was at least 10-fold slower than with DTT or TR/TRR. In all cases, >90% of the phosphatase activity was recovered with all three reducing agents. Therefore, the reversibility of H2S-treated PTP1B induced by TR/TRR is kinetically the most favorable, consistent with a potential physiological preference for this reducing agent.

Table 1

Rate of PTP1B reactivation by different reducing agents after inactivation with H2O2, H2S, and NO. The data are derived from three independent determinations.

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Change in mass of PTP1B after H2S treatment

To define more precisely the nature of the modification to PTP1B induced by incubation with H2S, we used a high-resolution quadrupole time-of-flight (QTOF) mass spectrometer to analyze a truncated form of PTP1B (residues 1 to 321) that made up the catalytic domain of the protein. The intact masses of H2S-treated and untreated PTP1B determined from electrospray ionization mass spectrometry (ESI-MS) spectra were 38,407.01 and 38,374.36, respectively, indicating a modification with a mass addition of 32.65 (Fig. 2A, upper panel). H2S treatment did not induce such a mass shift with the C215S mutant form of PTP1B (38,358.85) (Fig. 2A, lower panel). Furthermore, the increase in mass of 32.65 observed for wild-type PTP1B was abolished by treatment with DTT, which suggested that it involved a reversible cysteine modification.

Fig. 2

Analysis of H2S-modified PTP1B by MS. (A) Direct ESI-MS analysis of wild-type (WT) and C215S mutant forms of PTP1B after H2S treatment. WT or C215S mutant PTP1B (20 μg) was incubated with H2S (100 μM) for 10 min at 25°C and the mass of the protein was determined by ESI-MS analysis. H2S-treated WT PTP1B (upper panel) was resolved into two peaks corresponding to unmodified (38,374.36 daltons) and H2S-modified (38,407.01 daltons) protein (an increase in mass of 32 daltons). The deconvoluted spectrum for the active-site mutant C215S PTP1B showed a single peak of 38,358.85 daltons. (B) LC-MS/MS analysis of the tryptic peptide containing Cys215 of PTP1B. Purified PTP1B (20 μg) was treated with H2S (100 μM) or H2O2 (1 mM) for 10 min at 25°C, trypsinized overnight, and analyzed by LC-MS/MS. (Upper panel) Untreated peptide {[M + 3H]3+ at mass/charge ratio (m/z) of 725.692} corresponds to a free Cys215 sulfhydryl group; (middle panel) the H2S-treated peptide ([M + 3H]3+ at m/z 736.347, mass error −2.7 ppm) corresponds to sulfhydrated Cys215; (lower panel) the H2O2-treated peptide ([M + 3H]3+ at m/z 736.354, mass error −1.4 ppm) corresponds to the sulfinic acid modification of Cys215.

The active-site Cys215 of PTP1B can be oxidized to cysteine sulfinic acid (SO2H, 64.9697), which has a monoisotopic mass close to that of sulfhydrated cysteine (S-SH, 64.9520). To distinguish between these subtle mass differences, we trypsinized native, H2O2-treated, and H2S-treated PTP1B separately and analyzed the triply charged tryptic peptide containing the catalytic cysteine by MS. The three forms were readily distinguishable at the MS1 level by accurate mass measurements as the native (725.692, upper panel), sulfhydrated [736.347, mass error −2.7 parts per million (ppm), middle panel], and sulfinic acid (736.354, mass error −1.4 ppm, lower panel) forms (Fig. 2B). These three precursors were then selected for collision-induced dissociation (CID) to generate signature fragment ions. All three peptides generated confirmatory b ions (fragment ions from the N terminus) and y ions (fragment ions from the C terminus) for sequence identification. Notably, the H2S-treated peptide displayed a unique increase of 32 daltons compared to the native peptide for the y ion series for residues after Cys215 (y7), suggesting a stable modification of Cys215 with mass addition of 32. The tandem MS (MS/MS) spectrum for the same tryptic peptide after H2O2 treatment was different, with a hallmark neutral loss of 66 daltons for H2SO2 observed for y ions beyond y7, confirming the sulfinic acid modification of the cysteine. Together, the evidence from the intact protein mass measurements, accurate peptide masses at the MS1 level, and signature MS2 fragmentations suggest that the active-site cysteine (Cys215) of PTP1B undergoes sulfhydration upon treatment with H2S.

Measurement of sulfhydration of PTP1B with a thiolate anion–directed probe in HEK293T cells

To demonstrate sulfhydration of PTP1B within cells, we treated human embryonic kidney (HEK) 293T cells with varying amounts of a saturated solution of H2S and then lysed the cells in an anaerobic workstation to avoid post-lysis oxidation of cysteines. Sulfhydration was followed by a cysteinyl labeling assay that uses a biotinylated iodoacetic acid probe, which reacts through a nucleophilic substitution of the halide group by the PTP-reactive thiol group, resulting in a stable thioether bond (fig. S5) (33). After treating the cells with different concentrations of H2S for 30 min, we observed minimal labeling in untreated cells, representing the basal amount of reversibly oxidized PTP1B present before H2S addition, and saturation of the response at ~25 μM H2S (Fig. 3A). We detected PTP1B sulfhydration within 2 min of exposure of the cells to 100 μM H2S and peaked at 10 min (Fig. 3B). Finally, we used selected ion monitoring (SIM) MS to examine the tryptic peptide containing Cys215 from PTP1B. We observed sulfhydration of Cys215 in PTP1B from cells treated with H2S (Fig. 3C) but not in untreated cells. Thus, PTP1B was susceptible to H2S-induced sulfhydration in a cellular context.

Fig. 3

Detection of H2S-modified PTP1B with a thiolate anion–reactive probe. (A) HEK293T cells were serum-starved and then treated with the indicated concentrations of H2S for 30 min at 37°C. (B) HEK293T cells were treated with 100 μM H2S for the indicated times at 37°C. In both (A) and (B), cells were lysed, and the upper panel shows the modification of PTP1B monitored in 1.0 mg of cell lysate with a sulfhydryl-reactive IAP-biotin probe. The lower panel shows total PTP1B in 0.1 mg of cell lysate. Graphs to the right represent quantitation of the blots on the left and illustrate the ratio of Cys215 persulfide modification relative to total PTP1B, at different concentrations of H2S (upper) and different times at a fixed concentration of H2S (lower). (C) PTP1B was immunopurified from HEK293T cells that had been treated with 50 μM H2S for 30 min at 37°C, trypsinized, and analyzed by LC-MS/MS. This is an MS-MS spectrum of the peptide containing the active-site cysteine residue Cys215, illustrating that this residue was in the sulfhydrated form.

PTP1B sulfhydration under ER stress in control, but not in CSE-depleted, HeLa cells

We hypothesized that production of H2S may provide a mechanism for enhancing tyrosine phosphorylation, and thereby modulating cell signaling, through the sulfhydration and inactivation of PTP1B. PTP1B is localized to the ER and has been implicated in the control of signaling during ER stress (26). Furthermore, H2S production, mediated by CSE, has been linked to the ER stress–induced UPR (34). Therefore, we investigated whether H2S modified PTP1B function and cellular signaling under conditions of ER stress. We exposed HeLa cells to tunicamycin, an inhibitor of protein N-glycosylation, to trigger ER stress and observed a significant increase in H2S production (Fig. 4A). We immunoblotted cell lysates for PTP1B, CSE, and the ER chaperone BiP (binding immunoglobulin protein), which is a marker of ER stress that is sequestered by unfolded proteins, promoting its dissociation from the transmembrane ER proteins that trigger the stress response, thereby leading to activation of the response pathways. We detected no apparent change in the abundance of PTP1B, but we observed a small increase in the abundance of CSE within 2 hours of induction of ER stress. There was a marked increase in the abundance of BiP, which confirmed the induction of the ER stress response (fig. S6).

Fig. 4

H2S produced in response to tunicamycin-induced ER stress. (A) ER stress was induced by tunicamycin (10 μg/ml), and the production of H2S was measured in control (black bars) or CSE-deficient cells generated with two distinct shRNAs (open bars). Data are derived from three independent experiments and presented as means ± SEM. (B) The effect of tunicamycin-induced ER stress on the redox status of PTP1B Cys215 was followed by LC-MS/MS to quantify the unmodified Cys215 tryptic peptide ([M + 3H]3+ 725.692) and the sulfhydrated counterpart ([M + 3H]3+ 736.347). A time-dependent increase in sulfhydration (-SSH) was observed for PTP1B immunopurified from control (▪) but not CSE-deficient (•) cells. (C and D) PTP1B was immunopurified from control (C) and CSE-deficient (D) cells with the PTP1B-specific antibody DH8, and the enzymatic activity was monitored with p-NPP as substrate in the absence (open bars) or presence (black bars) of 5 mM DTT, after tunicamycin-induced ER stress. Data are derived from three independent experiments and presented as means ± SEM.

To suppress H2S production, we generated stable HeLa cell lines expressing short hairpin RNAs (shRNAs) that targeted the CSE-encoding mRNA and examined the effects on sulfhydration of PTP1B under ER stress. Immunoblot analysis indicated that both of the shRNAs we used decreased CSE abundance by >90% (fig. S7). We subjected both control and CSE-depleted cells to ER stress induced by tunicamycin for varying lengths of time and measured H2S production. Although H2S accumulated in control cells after exposure to tunicamycin, this was prevented in both of the stable cell lines expressing CSE shRNAs (Fig. 4A).

To characterize the modifications of the active-site cysteine residue that accompanied these changes in H2S production, we used SIM to quantify the unmodified peptide containing Cys215 ([M + 3H]3+ 725.692) and the sulfhydrated counterpart ([M + 3H]3+ 736.347), which we resolved chromatographically. In the absence of ER stress, 95% of PTP1B was detected in the reduced, active form (table S1). PTP1B from the control cells, which expressed endogenous CSE, showed an increase in sulfhydration after exposure to tunicamycin that plateaued within 6 to 8 hours, with a large proportion (~43%) of PTP1B in the sulfhydrated form (table S1). In contrast, cells in which CSE was depleted showed a much smaller increase in PTP1B sulfhydration (from ~5 to ~10%) together with small increases in oxidized forms (SO2H and SO3H) of the enzyme.

In parallel with the MS analysis, we immunopurified PTP1B at various times after exposure to tunicamycin and measured phosphatase activity in the immunoprecipitates. Treatment with tunicamycin induced a decline in the phosphatase activity of PTP1B immunopurified from control cells. In addition, there was a negative correlation between the extent of inhibition and the extent of sulfhydration (Fig. 4, B and C). Furthermore, addition of the reducing agent DTT restored PTP1B activity in the immunoprecipitate, indicating a reversible, inhibitory redox modification (Fig. 4C). In contrast, we did not detect any change in PTP1B activity, with or without reducing agent, in CSE-depleted cells (Fig. 4D). These data are consistent with modification of the essential active-site cysteine of PTP1B by sulfhydration and suggest that sulfhydration provides a mechanism for inactivation of the enzyme under conditions of ER stress.

H2S-mediated UPR signaling

Our data illustrate that H2S production in cells can lead to inhibition of PTP1B. Consequently, one might anticipate that changes in the amount of H2S may lead to changes in the phosphorylation of tyrosyl residues in proteins. Therefore, we used an antibody that recognizes phosphotyrosine and immunoblotting to assess tyrosine phosphorylation in control and CSE-depleted HeLa cells after tunicamycin-induced ER stress (Fig. 5). We anticipated that in the control cells with CSE, H2S would be produced, thereby inhibiting PTP activity and promoting tyrosine phosphorylation, whereas in CSE-depleted cells, in which the production of H2S was decreased, the result would be increased PTP activity, and thus lower cellular phosphotyrosine, compared to the control cells. Instead, we detected an unexpected increase in phosphorylation of proteins of 30 to 60 kD in CSE-depleted cells upon tunicamycin-induced ER stress (Fig. 5A). PTP1B activates the protein tyrosine kinase SRC, a 60-kD protein, by dephosphorylating the C-terminal inhibitory phosphorylation site, Tyr527, thereby promoting autophosphorylation of SRC on Tyr416, a process that is required for robust kinase activity (35, 36). Therefore, we assessed the phosphorylation status of Tyr527 and Tyr416 in SRC by immunoblotting with antibodies specific for the phosphorylated forms of SRC (Fig. 5B). We noted that there was a marked decrease in phosphorylation of Tyr527, with concomitant increase in phosphorylation of Tyr416, in CSE-depleted cells upon tunicamycin-induced ER stress. These effects were abrogated by inclusion of MSI-1436, a small-molecule, noncompetitive inhibitor of PTP1B that inhibits the phosphatase in mouse models of obesity (37), consistent with the identification of SRC as a substrate of PTP1B in the context of obesity.

Fig. 5

H2S-mediated signaling in HeLa cells in response to ER stress. (A) Immunoblot showing the overall change in tyrosine phosphorylation induced by ER stress, visualized with the 4G10 phosphotyrosine-specific antibody. The two left lanes show tyrosine phosphorylation in the absence of ER stress; the two right lanes show tyrosine phosphorylation after 4 hours of treatment with tunicamycin. Control and CSE-depleted cells are as indicated. (B) SRC activation was monitored in control and CSE-depleted cells with phosphorylation-specific antibodies to the inhibitory C-terminal site (Tyr527, pY527) and the autophosphorylation site (Tyr527, pY416). The lower panel shows total SRC. ER stress was induced for 4 hours with tunicamycin. Cells were treated with PTP1B inhibitor MSI-1436 (5 μM) as indicated. (C) Immunoblot analyses to examine the activation of components of the UPR. The data show a time course to examine activation of IRE1 [anti–phospho-IRE1α (Ser724) compared to total IRE1 (anti-IRE1α)], the activation of PERK [anti–phospho-PERK (Thr980) compared to total PERK (anti-PERK)], and the abundance of ATF6 and PTP1B in control and CSE-deficient cells. (D) Immunoblot showing that the PTP1B inhibitor MSI-1436 (5 μM) restored PERK activation in response to tunicamycin-induced ER stress in CSE-depleted cells. The histograms illustrate the ratio of activated to total PERK at different time points after initiation of ER stress, without (left) and with (right) the PTP1B inhibitor MSI-1436 in control (black bars) and CSE-deficient (white bars) cells.

The phosphotyrosine immunoblot also revealed that an increase in tyrosine phosphorylation of a protein of ~150 kD was induced by ER stress in response to tunicamycin in control cells, which was not detected in CSE-depleted cells (Fig. 5A). This is consistent with attenuation of phosphatase activity by H2S in the presence of CSE under ER stress. To identify this protein, we examined the three main branches of ER stress–induced signaling in control and CSE-depleted HeLa cells. We observed that ER stress induced by tunicamycin (Fig. 5C) or by thapsigargin (fig. S8) increased the phosphorylation of PERK on Thr980, a marker of PERK activation (29, 38, 39), and that this response was delayed and attenuated in CSE-depleted cells. In contrast, changes in the abundance of ATF6 or phosphorylation of IRE1 were considerably less pronounced. With the small-molecule inhibitor MSI-1436 of PTP1B (37), we observed that, whereas MSI-1436 did not affect PERK activation in response to ER stress in control cells, it restored PERK activation in CSE-depleted cells to an extent similar to that observed in control cells (Fig. 5D). These data suggest that regulation of PERK activation depended on sulfhydration of PTP1B.

PTP1B dephosphorylation of PERK

The cytoplasmic portion of PERK contains a protein kinase domain that is activated by oligomerization, leading to enhanced autophosphorylation (30).The phosphorylation of Thr980, which was measured in Fig. 5, C and D, is a marker of its activation status; however, considering the specificity of PTP1B for tyrosyl residues in proteins, this particular site will not be the direct substrate of the phosphatase. Autophosphorylation of Tyr619, which is located in a highly conserved segment of the kinase domain, promotes phosphorylation of Thr980 and activation of PERK (39). Therefore, using the D181A substrate-trapping mutant form of PTP1B for in vitro and transfection studies, we investigated the possibility that PERK phosphorylated at Tyr619 (pTyr619-PERK) was a substrate of PTP1B.

We generated lysates from untreated cells or cells treated with tunicamycin and incubated them with wild-type PTP1B or D181A PTP1B. PTP1B was then immunoprecipitated, and its interaction with PERK was monitored with PERK-specific antibody. We observed that the D181A PTP1B substrate-trapping mutant, but not the wild-type enzyme, coprecipitated PERK only in lysates of tunicamycin-treated cells (Fig. 6A). Treatment of the PTP protein with pervanadate, which promotes oxidation of the active-site cysteine of PTP family members, also disrupted the interaction between D181A PTP1B and PERK (Fig. 6A); this highlights the importance of the active-site cysteine, consistent with an enzyme-substrate interaction. In addition, to examine the importance of phosphorylation of PERK for its interaction with the substrate-trapping mutant form of PTP1B, we treated the lysate with calf intestinal alkaline phosphatase (CIAP) to eliminate PERK phosphorylation. We observed that this treatment abrogated the association (Fig. 6B). We examined the complexes that coimmunoprecipitated with wild-type or substrate-trapping mutant PTP1B by liquid chromatography–MS/MS (LC-MS/MS) and identified the pTyr619 PERK peptide in complexes associated with D181A PTP1B but not wild-type PTP1B (Fig. 6C).

Fig. 6

Interaction of the D181A substrate-trapping mutant form of PTP1B and tyrosine-phosphorylated PERK. (A) Lysates were prepared from untreated control cells (no stress) or cells treated with tunicamycin for 4 hours (ER Stress) and were incubated with PTP1B or D181A PTP1B in the presence or absence of 1 mM pervanadate. PTP1B immunoprecipitates were subjected to SDS-PAGE and immunoblotted with an antibody recognizing PERK (upper panel). The blot was then stripped and reblotted with antibody FG6 recognizing PTP1B (lower panel). (B) Lysates prepared from untreated cells (no stress) or cells treated with tunicamycin for 4 hours (ER Stress) were incubated in the absence (−) or presence (+) of calf intestine alkaline phosphatase (CIAP). The treated lysates were incubated with wild-type PTP1B or D181A PTP1B; PTP1B immunoprecipitates were analyzed as described in (A). (C) PERK was identified as a PTP1B substrate by MS. Immunopurified protein complex from the substrate-trapping experiment was treated with vanadate to release the bound substrate, which was then subjected to trypsinization, and the peptides obtained were analyzed by MudPIT. This is an MS/MS spectrum of the peptide containing phosphorylated Tyr619 in PERK, which bound to the substrate-trapping mutant D181A form of PTP1B. (D) Wild-type PTP1B or D181A PTP1B were overexpressed in HEK293T cells and immunoprecipitated after tunicamycin treatment for 4 hours, resolved by SDS-PAGE, immunoblotted with rabbit polyclonal antibody against PERK (upper panel), and stripped and reblotted using FG6 antibody recognizing PTP1B (lower panel). (E) Wild-type and mutant (Y615F) myc-tagged forms of mouse PERK were overexpressed in HEK293T cells and immunoprecipitated with 9E10 antibody recognizing myc after tunicamycin treatment for 4 hours. PERK was eluted with myc peptide and then incubated with Ni-NTA bead–bound wild-type or D181A substrate-trapping mutant forms of PTP1B. Complexes were resolved by SDS-PAGE, immunoblotted with 9E10 to detect the myc tag, and then stripped and reblotted with FG6 antibody recognizing PTP1B (lower panel). (F) Wild-type and mutant, K618A (catalytically inactive), and Y615F forms of myc-tagged mouse PERK were overexpressed in HEK293T cells and immunoprecipitated after 4 hours of tunicamycin treatment. Mock represents the vector control. The immunoprecipitated proteins were incubated with wild-type or C215S catalytically inactive mutant forms of PTP1B, resolved by SDS-PAGE, and immunoblotted with 4G10 antibody recognizing phosphotyrosine. The lower panel is a loading control obtained by stripping and reblotting with FG6 antibody recognizing PTP1B.

To examine the association between PTP1B and PERK in a cellular context, we overexpressed wild-type and substrate-trapping mutant forms of PTP1B in HEK293T cells and subjected the cells to ER stress. Similar amounts of the wild-type and substrate-trapping mutant PTP1B were detected in the immunoprecipitates (Fig. 6D, lower panel); however, as seen in vitro, only the D181A PTP1B substrate-trapping mutant coprecipitated PERK (Fig. 6D, upper panel). The detection of such complex formation within the cell is most likely explained because the catalytically impaired, substrate-trapping mutant was overexpressed to an extent that exceeded the ability of H2S to modify it and thus formed a complex with the pTyr-PERK produced in response to ER stress. Considering the specificity of PTP1B for tyrosyl residues in proteins, we tested the importance of Tyr619 in PERK for the association with the substrate-trapping mutant. We expressed myc-tagged murine PERK (wild-type or Tyr615, which is equivalent to Tyr619 in the human protein, substituted with Phe) in HEK293T cells, treated the cells with tunicamycin, and incubated the cell lysates with either wild-type or D181A mutant PTP1B (Fig. 6E). Mutation of the Tyr619 site of phosphorylation in PERK abrogated its interaction with the D181A PTP1B substrate-trapping mutant. Wild-type PTP1B directly dephosphorylated myc-tagged PERK that had been isolated from tunicamycin-treated HEK293T cells by immunoprecipitation (Fig. 6F). Collectively, these data establish a direct enzyme-substrate interaction between PTP1B and pTyr619-PERK.


The essential role of the active-site cysteine residue in PTP-mediated catalysis provides a mechanism for redox-based regulation of PTP activity. The reversible oxidation and inactivation of PTPs in response to the regulated production of H2O2 provides a well-established mechanism for control of tyrosine phosphorylation–dependent signaling (18, 19). NO-induced nitrosylation of the active-site cysteine in PTPs has also been reported (20). Here, we demonstrate that PTP1B function is regulated by reversible sulfhydration in response to endogenously generated H2S and that this contributes to the ER stress response.

Consistent with such previously identified targets as actin, tubulin, and GAPDH (17, 40), we observed that H2S sulfhydrated a specific cysteine in PTP1B, in this case the cysteine residue at the active site that is essential for catalysis. However, whereas sulfhydration is generally thought to activate target proteins (6), we found that sulfhydration of Cys215 inhibited PTP1B activity. We observed sulfhydration of PTP1B in vitro at concentrations of H2S encountered in vivo (12) and found that it occurred exclusively on the catalytic cysteine, Cys215, leaving the other cysteine residues in the protein unmodified. This is consistent with the unique properties of this residue, which distinguish it from other cysteine residues in the protein. Cys215 is present as a thiolate, rather than as a free thiol, a feature conferred by the architecture of the PTP active site (41) that renders these enzymes exquisitely sensitive to inactivation by modification of the catalytic cysteine. The observation that PTP1B was sulfhydrated and inactivated following addition of H2S in vitro was nevertheless surprising because direct reaction of the cysteine with H2S is unlikely. This raises questions as to the underlying mechanism. High-throughput screens of chemical libraries have shown that certain inhibitors function as “peroxide generators” when included with the other constituents of a standard phosphatase activity assay and that any peroxide thus generated inhibits PTP activity (10). It is possible that a similar situation may apply here, and any peroxide could affect the form in which H2S is encountered by PTP1B. For example, any H2O2 in the assay could interact with H2S/HS to generate HSSH (42), which could react directly with the PTP active site. Alternatively, any oxidized PTP1B produced in the reaction, which would adopt the cyclic sulfenamide conformation (41), could also react directly with H2S. We observed selectivity in the sensitivity of sulfhydrated PTP1B to different reducing agents. Although oxidized and nitrosylated PTP1B displayed sensitivities similar to that of DTT, GSH, and thioredoxin, the sulfhydrated enzyme, which features an -SSH moiety on the active-site cysteine, displayed ~190-fold faster reduction and reactivation by thioredoxin, the major intracellular disulfide bond reductase, compared to DTT and GSH. Therefore, sulfhydration of PTP1B may play a role in controlling the mechanism for reduction and reactivation of the enzyme (fig. S9). Nevertheless, these mechanistic considerations do not change the conclusions of our study: The sulfhydrated form of PTP1B, which was produced to high stoichiometry in cells under conditions of ER stress, is inactive and, therefore, provides a potential physiological mechanism for regulating pTyr-dependent signaling.

The ER plays a major role in controlling the folding and modification of newly synthesized transmembrane and secreted proteins. The folding capacity of the ER is modulated in response to the environmental and physiological status of the cell; thus, a cell can match its capacity for protein folding to its physiological requirements. However, if that capacity is exceeded, resulting in a condition known as ER stress, then an integrated set of three signaling pathways, the UPR, is activated (30). These pathways are triggered by transmembrane sensor proteins that detect unfolded proteins in the ER and transduce a signal that initiates a response in the cytosol or nucleus, enabling the cell to fine-tune the rates of protein synthesis and the folding capacity to maintain homeostasis and, if that is not successful, to trigger cell death. PTP1B is resident in the ER and has been implicated in the control of ER stress (26). Furthermore, the production of H2S by CSE has been implicated as a facet of the regulation of the ER stress response (43). Therefore, we explored further the potential for integration of the effects of PTP1B and H2S in the UPR.

The three pathways that constitute the UPR are initiated and controlled by the transmembrane ER proteins IRE1, ATF6, and PERK (2730). IRE1 is a bifunctional protein kinase and site-specific endoribonuclease. When its luminal segment senses the accumulation of unfolded protein, it undergoes oligomerization in the ER membrane, which induces autophosphorylation of the cytoplasmic kinase domain and activation of the endoribonuclease activity. The latter promotes splicing of X-box binding protein (XBP) mRNA to generate XBP1, which is a transcriptional activator that induces expression of genes encoding both chaperones and proteins associated with degradation. Furthermore, binding of the adaptor TRAF2 (TNF receptor–associated factor 2) by phosphorylated IRE1 promotes activation of c-Jun N-terminal kinase (JNK), which controls apoptosis. The stress sensor ATF6 also presents a segment to the ER lumen that detects unfolded proteins, resulting in activation and transport to the Golgi where it is cleaved by proteases. This cleaved ATF6 is released from the membrane and imported into the nucleus where it directs the transcription of a subset of genes that regulate the UPR. Finally, PERK, like IRE1, is an ER transmembrane kinase that oligomerizes under conditions of ER stress, leading to autophosphorylation and activation. The effects of PERK are exerted through phosphorylation of Ser51 in eIF2α, which catalyzes the rate-limiting step of initiation of protein translation (38). This phosphorylation traps eIF2α in an inactive, guanosine diphosphate (GDP)–bound state, which decreases protein synthesis and limits the load of unfolded protein entering the ER. Under conditions of eIF2α phosphorylation, translation of the transcription factor ATF4 is also induced, which regulates expression of genes involved in the UPR. The phosphorylation status of eIF2α is a major point of signal integration, such that downstream events are referred to as the integrated stress response (30). This places considerable importance on maintaining tight control over the phosphorylation status of eIF2α. The reversibility of PERK phosphorylation is an important component of this regulatory mechanism. Our data demonstrated that PTP1B influenced the activity of PERK, but not IRE1 or ATF6, under conditions of ER stress. There is evidence to suggest crosstalk between these three pathways (4446); however, differential regulation of the pathways is also thought to circumvent functional redundancy (30). Our study reveals that PTP1B underlies a new aspect of differential regulation of UPR signaling and illustrates another example of specificity in the function of a member of the PTP family.

PERK belongs to the RNA-dependent protein kinase (PKR) family of proteins. Phosphorylation of a conserved tyrosine residue within the catalytic domain has been implicated in activation of PKR kinases (47). In human PERK, this residue is Tyr619, which corresponds to mouse PERK Tyr615. PERK is a dual-specificity kinase that can autophosphorylate this tyrosyl residue, coincident with autophosphorylation of Thr980 in its activation loop. Mutation of human Tyr619 compromises PERK activation and its phosphorylation of eIF2α (39). Our data reveal that Tyr619 is a substrate of PTP1B through which PTP1B can attenuate PERK activity and thereby the UPR. PTP1B has a preference for acidic residues N-terminal to the phosphorylated tyrosine (32, 48); the sequence flanking PERK Tyr619 resembles that of other PTP1B substrates, with two aspartic acid residues N-terminal to the phosphorylated tyrosine (48, 49). Our observations are consistent with a study showing that PTP1B overexpression suppressed tunicamycin-induced PERK signaling and eIF2α phosphorylation in MIN6 insulinoma β cells, whereas RNA interference (RNAi)–mediated PTP1B deficiency enhanced this ER stress response (50). A previous study comparing ER stress signaling in fibroblasts from wild-type and PTP1B-deficient mice, however, suggested that PTP1B was required for activation of IRE1 signaling, as measured by XBP1 splicing and JNK activation, but did not affect PERK phosphorylation of Ser51 in eIF2α (26). The reason for this discrepancy is unclear; however, there have been no further reports defining the mechanism underlying these effects of PTP1B on IRE1 signaling. The difference may be related to the chronic loss of PTP1B in these animals, which, in itself, could be interpreted by the cells as a stress that may affect the UPR.

In summary, we have identified PTP1B as a phosphatase that specifically inhibited the PERK arm of the UPR. The ability of PTP1B to dephosphorylate Tyr619 and inactivate PERK is fine-tuned by the production of H2S by CSE in response to ER stress. The resulting sulfhydration and inactivation of PTP1B facilitated PERK phosphorylation and activation, which would promote restoration of ER homeostasis. The activity of PERK would be kept in check by the reduction and reactivation of PTP1B, likely mediated by thioredoxin, which would ensure reversibility of PERK phosphorylation and flexibility in the response to ER stress (Fig. 7).

Fig. 7

Proposed model for the role of sulfhydration of PTP1B in regulating the response to ER stress. Induction of ER stress increases the production of H2S by CSE, which leads to sulfhydration of the essential active-site cysteine residue in PTP1B and concomitant inactivation of the phosphatase. This transiently protects pTyr619 of PERK from dephosphorylation, promoting PERK activation and its ability to inhibit global translation by phosphorylating eIF2α. Reduction and reactivation of sulfhydrated PTP1B restores phosphatase activity and its ability to dephosphorylate and inactivate PERK.

Even mild oxidative stress can rapidly convert the PTP active-site cysteine to irreversibly oxidized sulfinic and sulfonic acid forms (41). Thus, in addition to acute regulation of PTP1B function, sulfhydration may also act as a protective mechanism to prevent irreversible oxidation and inactivation. NO-induced S-nitrosylation of the active-site cysteine protects PTP1B from permanent inactivation by reactive oxygen species (ROS) (20). Given the interplay among the different gasotransmitters and their regulation of similar functions, it is conceivable that H2S may also be involved in preventing irreversible PTP oxidation. This could be particularly important in the context of the ER, which is a highly oxidizing environment as a result of the redox dependence of protein folding, and ER stress, which also leads to an increase in ROS production (51, 52).

Materials and Methods


Antibodies were purchased from Cell Signaling (IRE1, PERK, phospho–PERK-T980, eIF2α, phospho–eIF2α-S51, pY419-SRC, and SRC), Sigma (β-actin), Thermo Scientific (phospho–IRE1-S724), Millipore (phosphotyrosine 4G10 and ATF6), and Abcam (CSE). Chemicals were obtained from Sigma unless otherwise noted. The Y615F-PERK (murine, equivalent to Y619F in human) was a gift from A. Koromilas (McGill University).

Cell culture

shRNAs targeting luciferase and CSE were obtained from Open Biosystems and cloned into the pSVG vector between the restriction sites Eco RI and Xho I. The sequences of the CSE shRNAs were AGGAGCTGATATTTCTATGTATTAGTGAAGCCACAGATGTAATACATAGAAATATCAGCTCCC and CCCTCAAGAACCTAAAGCTATTTAGTGAAGCCACAGATGTAAATAGCTTTAGGTTCTTGAGGA. Stable plasmid expression was achieved via sequential retroviral transduction, followed by selection with puromycin (2 μg/ml). ER stress was induced with tunicamycin (10 μg/ml) or thapsigargin (1 μM) for 0 to 8 hours. MSI-1436 was used at a concentration of 5 μM to inhibit PTP1B. For transient transfection, 293T cells were plated in Dulbecco’s modified Eagle’s medium (DMEM) supplemented with 10% fetal bovine serum for 16 hours. The culture medium was replaced by Opti-MEM (Life Technologies) without serum, and then plasmid (5 μg per dish) was introduced using FuGENE (Roche) according to the manufacturer’s recommendations. The transfection efficiency was routinely 80%.

Examination of PERK recognition as a substrate by PTP1B

To demonstrate direct dephosphorylation of PERK by PTP1B, we overexpressed wild-type and mutant (K618A and Y615F) forms of myc-tagged mouse PERK in HEK293T cells for 36 hours, after which we lysed the cells (either untreated or treated with tunicamycin for 4 hours) on ice for 30 min. Overexpressed PERK was immunopurified with the antibody recognizing myc (9E10). After extensive washing, the immunoprecipitate was incubated with wild-type or C215S catalytically inactive mutant forms of PTP1B (10 ng) for 30 min at room temperature, after which the beads were centrifuged at 3000g for 10 min, resolved by SDS–polyacrylamide gel electrophoresis (SDS-PAGE), and immunoblotted with antibody recognizing phosphotyrosine (4G10).

For substrate trapping in vitro, lysates prepared from untreated control cells or cells treated with tunicamycin (10 μg/ml) for 4 hours were incubated overnight at 4°C with 20 μl of His-tagged wild-type PTP1B and D181A PTP1B fusion protein coupled to beads (10 μg/μl) in the presence and absence of 1 mM pervanadate. After several washes, complexes were analyzed by immunoblotting. To examine the importance of phosphorylation of PERK for the interaction, we eluted immunopurified myc-tagged wild-type and Y615F PERK from the immunoprecipitate with myc peptide (100 μg/ml) for 30 min at 4°C and then incubated Ni-NTA bead–bound His-tagged wild-type or D181A substrate-trapping mutant forms of PTP1B. Complexes were resolved by SDS-PAGE, and the components were identified by immunoblotting. For trapping in cells, wild-type and D181A mutant PTP1B were overexpressed in HEK293T cells for 36 hours, after which cells (either untreated or treated with tunicamycin for 4 hours) were lysed on ice for 30 min. The PTP was then immunopurified with anti-PTP1B antibody FG6. After extensive washing in lysis buffer, the precipitated proteins were examined by immunoblotting.

Rate of inactivation and reactivation of PTP1B

The rate of inactivation of PTP1B was measured by a standard spectrophotometric assay using p-nitrophenyl phosphate (p-NPP) as substrate. PTP1B (1 μM) was incubated at 25°C with varying concentrations of H2S (1 to 200 μM), H2O2 (1 to 5 mM), or NO donor SNAP (S-nitroso-N-acetylpenicillamine, 1 to 20 mM) in 50 mM Hepes (pH 7.0), 100 mM NaCl, and 0.1% bovine serum albumin (BSA). Phosphatase activities were measured immediately in a continuous assay that monitors the production of nitrophenol at 405 nm (18,000 M−1 cm−1). To calculate the rates of reactivation by DTT, GSH, or TR/TRR, we initially inactivated PTP1B (10 μM) by incubation with H2S (1 to 100 μM), H2O2 (0.5 to 2 mM), SNAP, or GSNO (S-nitrosoglutathione, 1 to 20 mM); aliquots were then diluted twofold into 50 mM Hepes (pH 7.5), 100 mM NaCl, 0.1% BSA, and 1 mM EDTA containing varying amounts of DTT (10 to 200 mM), GSH (50 to 200 mM), or TR/TRR (0.2 to 5 equivalents of TR to PTP1B). After incubation under reducing conditions for varying amounts of time (1 to 60 min), aliquots were assayed for phosphatase activity with p-NPP (2 mM). For assays with TR/TRR as the reductant, NADPH (reduced form of nicotinamide adenine dinucleotide phosphate, 25 to 300 μM) was also included with a TR-to-TRR ratio of 200:1.

Fluorescence polarization assay

Wild-type and mutant forms of PTP1B were incubated with or without 100 μM H2S for 10 min at room temperature, and binding was measured with 25 nM substrate carboxyfluorescein labeled Glu-Asn-Asp-pTyr-Ile-Asn-Ala-Ser-Leu (5′-FAM-ENDpYINASL) in 50 mM Hepes (pH 7.0) and 100 mM NaCl by means of the SpectraMax Gemini XPS fluorescence plate reader (Molecular Devices). Excitation was set at 494 nm and emission was collected at 522 nm. Fluorescence polarization is plotted as fluorescence intensity (vertical)/fluorescence intensity (horizontal). Because it is a ratio of intensities, the polarization value is dimensionless; it is expressed as arbitrary millipolarization units.

Cysteinyl labeling assay

H2S stock solution was prepared freshly by directly bubbling pure H2S gas into distilled water to generate a saturated solution (0.09 M H2S at 30°C) (53). H2S stock solution was diluted to different concentrations in cell culture medium or appropriate buffer. Cells were grown to confluence and serum-starved for 16 hours before stimulation with H2S. After stimulation, cells were placed in an anaerobic workstation where the partial pressure of oxygen (PO2) concentration was maintained at 1 to 2 ppm. After removal of DMEM, cells were rapidly lysed with ice-cold degassed lysis buffer [25 mM sodium acetate (pH 5.5), 1% NP-40, 150 mM NaCl, 10% (v/v) glycerol, aprotinin (25 μg/ml), and leupeptin (25 μg/ml)] and transferred to brown-colored tubes. Lysates were shaken for 1 hour at room temperature to allow complete alkylation of free thiols. Cell debris was then cleared by centrifugation at 12,000g for 3 min at room temperature. Protein concentrations were determined by the method of Bradford, and 1 mg of cell lysate was slowly applied to desalting columns that had been equilibrated with iodoacetic acid–free lysis buffer. Buffer exchange was performed by centrifuging at 2000g for 2 min at 4°C. IAA-cleared lysates were then supplemented with 1 mM DTT and allowed to incubate for 30 min on a shaker at room temperature. During this phase, the persulfide and other reversibly oxidized forms of the active-site Cys residue, which were protected from alkylation in the previous step, were reduced back to their thiolate states. The lysates were then incubated with biotinylated IAP probe (5 mM) for 1 hour on a shaker at room temperature. Biotinylated proteins were enriched by using streptavidin-Sepharose beads for 16 hours at 4°C on a rotating wheel, with sequential rounds of centrifugation (12,000g, 1 min, 4°C) using phosphate-buffered saline (PBS) to wash the beads. The beads were resuspended in 20 μl of 4× Laemmli sample buffer and heated at 90°C for 1 min. PTP1B was then identified by immunoblotting.

Measurement of PTP1B activity by immunophosphatase assay

Cells untreated or treated with tunicamycin (10 μg/ml) were lysed on ice with 20 mM tris-HCl (pH 7.5), 100 mM NaCl, and 0.5% NP-40 supplemented with aprotinin (5 μg/ml), leupeptin (5 μg/ml), and 0.2 mM phenylmethylsulfonyl fluoride. PTP1B was immunoprecipitated from 500 μg of cell lysate with 2 μg of antibody DH8 for 90 min at 4°C, followed by incubation with protein G–Sepharose beads for 1 hour. The immunoprecipitates were washed twice, first with lysis buffer and then with PTP assay buffer [50 mM Hepes (pH 7.0), 100 mM NaCl, 2 mM EDTA, and BSA (0.1 mg/ml)]. After immunoprecipitation, PTP1B was assayed using 10 mM p-NPP as substrate for 30 min at 25°C in the absence and presence of 5 mM DTT. The reaction was stopped by addition of NaOH to a final concentration of 1 M. The increase in absorbance at 405 nm was measured to determine the enzyme activity. Linearity of activity was demonstrated with different amounts of the immunoprecipitate. The values in the absence of enzyme were taken as background and were always subtracted for correction. Data are presented as percent phosphatase activity, in which the activity in the immunoprecipitate is expressed relative to that of an equal amount of purified recombinant PTP1B.

H2S measurement

H2S was measured as described previously (53, 54). Cells were lysed in 50 mM phosphate buffer (pH 7.5) with zinc acetate [1% (w/v)] to trap H2S. The reaction was stopped after 10 min by the addition of 20 μM N,N-dimethyl-p-phenylenediamine sulfate and 30 mM FeCl3, incubated in the dark for 30 min, and 10% trichloroacetic acid was added to precipitate any protein. Subsequently, the mixture was centrifuged at 10,000g for 10 min and absorbance of the supernatant at 670 nm was measured. H2S concentration was calculated from a calibration curve of standard H2S solutions. Data are presented as percent increase in H2S relative to untreated cells.

Mass spectrometry

Recombinant PTP1B proteins (~20 μg each of wild type, H2S-treated, and H2O2-treated) were digested overnight with trypsin [1:50 (w/w)] at 37°C. Tryptic peptides were acidified by the addition of formic acid [0.1% (v/v) final], and 2-μl aliquots of peptide samples were injected onto a microfluidic reversed-phase LC (RPLC) chip (43 mm × 75 μm C18 chip column with 40-nl trap column) and acquired on a nano-LC QTOF system (Agilent QTOF 6520). Both data-dependent and targeted acquisition modes were applied to PTP1B peptides. Peak list files were generated with MassHunter software (version B.02.00) and searched against the International Protein Index (IPI) human database (87,130 sequences) with Mascot v2.3 (55). Manual interpretation was also applied to validate peptide spectra. For the time course study in tunicamycin-treated cells, SIM was used to quantify specifically the native and modified Cys215-containing peptides. Chromatography peak detection, integration, and quantitation were performed with MassHunter software. For intact protein mass measurement, proteins were first buffer-exchanged into 20 mM triethylammonium bicarbonate buffer with a PD10 column. Protein samples (~1 μg) were injected onto a customized C8 protein chip (43 mm × 75 μm C8 chip column with 40-nl trap column) and acquired on the QTOF 6520. Acquisition parameters were the same as for peptide MS, except that data were acquired in profile mode and the fragmentor voltage was set at 200 V. Intact protein spectra were integrated and deconvoluted with the maximum entropy workflow in the BioConfirm software. For examination of sulfhydration of Cys215-containing peptides in PTP1B from H2S-treated 293T cells, the samples were analyzed with an Orbitrap Velos mass spectrometer (Thermo Scientific) using capillary reversed-phase high-performance LC and acquisition parameters essentially as described in (56).

PTP1B substrate identification

Immunopurified protein complexes from the substrate-trapping experiments were treated with 5 mM vanadate to release the bound substrates and subjected to trypsin digestion overnight at 37°C after reduction with DTT and alkylation with iodoacetamide (57). The digested peptides were then acidified by 0.1% (v/v) trifluoroacetic acid and desalted by C18 stage tip (58). Peptide samples were then dried and resuspended in loading buffer (5% acetonitrile and 0.1% formic acid). Peptides were identified by a four-step multidimensional protein identification technology (MudPIT) (59) at ammonium acetate salt concentrations of 10, 45, 60, and 100%, respectively, each followed by a 120-min RPLC gradient on a Proxeon nano-LC system. Data were acquired on an LTQ-Orbitrap XL with a survey scan at 30,000 resolution and followed by six data-dependent CID MS2 scans at a resolution of 7500, 10-ms activation time, normalized collision energy at 25, and q at 0.25. Peak list files were generated by Mascot Distiller (v2.3) and searched against the IPI human database (87,130 sequences) with Mascot v2.3. Search parameters are 15 ppm for MS1 and 0.6 dalton for MS2, carbamidomethyl (Cys) as fixed modification, with methionine oxidation and tyrosine phosphorylation as variable modifications. Peptide scores with P value below 0.05 are considered as confident identifications.

Statistical analysis

Immunoblotting was performed by standard methods. All blots are representative of experiments that were performed at least three times. In figures where blots were quantitated, we used ImageJ software and have represented the data as a bar graph (±SEM).

Supplementary Materials

Fig. S1. Time-dependent inactivation of PTP1B by H2O2.

Fig. S2. Time-dependent inactivation of PTP1B by NO.

Fig. S3. Time-dependent reactivation of PTP1B by TR/TRR.

Fig. S4. Time-dependent reactivation of PTP1B by GSH.

Fig. S5. Mechanism of PTP labeling by the IAP probe.

Fig. S6. Induction of ER stress by tunicamycin.

Fig. S7. Decrease in CSE by RNAi.

Fig. S8. Changes in components of the UPR after exposure to thapsigargin.

Fig. S9. Proposed mechanism for persulfide modification of PTP1B.

Table S1. Quantitation of the different redox forms of Cys215 in PTP1B observed after induction of ER stress with tunicamycin.

References and Notes

  1. We thank C. Ruse and the members of the Cold Spring Harbor Laboratory (CSHL) Proteomics Shared Resource for their invaluable assistance. Funding: This work was supported by NIH grants CA53840 and GM55989 (to N.K.T.) and the CSHL Cancer Center Support grant CA45508. We are also grateful for support from the Irving Hansen Foundation and the Masthead Cove Yacht Club Carol Marcincuk Fund. PTP1B inhibitor MSI-1436 was provided by Ohr Pharmaceuticals and Genaera Corporation. Author contributions: N.K. and N.K.T. designed the experiments and analyzed the data. N.K. performed the experiments. C.F. and D.J.P. were responsible for the MS analysis. N.K. and N.K.T. wrote the manuscript, with contributions from C.F. and D.J.P. Competing interests: The authors declare that they have no competing interests.
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