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Molecular Cell, Vol. 9, 387–399, February, 2002, Copyright 2002 by Cell Press

Reversible Oxidation and Inactivation of Protein Tyrosine Phosphatases In Vivo Tzu-Ching Meng, Toshiyuki Fukada, and Nicholas K. Tonks1 Cold Spring Harbor Laboratory 1 Bungtown Road Cold Spring Harbor, New York 11724

Summary We have investigated the regulation of protein tyrosine phosphatases (PTPs) by reactive oxygen species (ROS) in a cellular environment. We demonstrate that multiple PTPs were reversibly oxidized and inactivated following treatment of Rat-1 cells with H2O2 and that inhibition of PTP function was important for ROSinduced mitogenesis. Furthermore, we show transient oxidation of the SH2 domain containing PTP, SHP-2, in response to PDGF that requires association with the PDGFR. Our results indicate that SHP-2 inhibits PDGFR signaling and suggest a mechanism by which autophosphorylation of the PDGFR occurs despite its association with SHP-2. The data suggest that several PTPs may be regulated by oxidation and that characterization of this process may define novel links between specific PTPs and particular signaling pathways in vivo. Introduction Over the last fifteen years it has been established that the protein tyrosine phosphatases (PTPs) are a large, structurally diverse family of receptor-like and nontransmembrane enzymes which exhibit exquisite substrate specificity in vivo and are critical regulators of a wide array of cellular signaling pathways (Andersen et al., 2001; Tonks and Neel, 2001). An important area of investigation in the field remains the characterization of mechanisms by which the activity of the PTPs themselves may be regulated in vivo. Recently, the proposal that certain PTPs may be susceptible to oxidation and inactivation has introduced an additional tier of complexity to the regulation of this family of enzymes. It is now apparent that reactive oxygen species (ROS) are not merely a harmful byproduct of life in an aerobic environment. The importance of ROS in phagocytic cells, such as neutrophils, is well documented. Various stimuli lead to the assembly of a multicomponent NADPH oxidase complex which mediates a process known as the respiratory burst (DeLeo and Quinn, 1996). NADPH oxidase catalyses transfer of one electron from NADPH to molecular oxygen to generate superoxide anions, which in turn may yield hydrogen peroxide either via protonation of superoxide or through the action of superoxide dismutase (Thelen et al., 1993). The large quantities of such ROS produced in phagocytic cells have been implicated as microbicidal agents and in certain pathological situations can result in host cell dam1

Correspondence: [email protected]

age (Smith and Curnutte, 1991). However, many recent studies have revealed that the production of ROS is tightly regulated, engendering the concept that at lower levels than those generated for a microbicidal function ROS may also function in propagating a signaling response to extracellular stimuli (Finkel, 1998, 2000). Thus, in a manner analogous to reversible protein phosphorylation, the reversible oxidation of target proteins in a cell may regulate the function of those proteins in response to various agonists and thus elicit a cellular response to stimulation (Finkel, 1998). Several lines of investigation have implicated ROS in the regulation of mitogenic signaling in mammalian cells (Adler et al., 1999; Chen et al., 1995; Sundaresan et al., 1995). Mild oxidation can yield a stable sulfenic acid modification of cysteine residues (Cys-SOH) in selected proteins, including a variety of enzymes and transcription factors, which has the potential to regulate the function of those proteins (Claiborne et al., 1999). In order to understand the role of ROS and redox regulation in the control of signal transduction, it is particularly important to identify the targets of reversible oxidation in vivo. In this context, attention has been drawn to the PTPs, which together with the protein tyrosine kinases (PTKs) are responsible for maintaining a normal tyrosine phosphorylation status in vivo. The PTPs are characterized by a signature motif, I/V-H-C-X-X-G-X-X-R-S/T, which forms the base of the active site cleft and contains an invariant Cys residue (Barford et al., 1995). The catalytic mechanism involves a two-step process commencing with nucleophilic attack by the S␥ atom of the catalytic Cys on the phosphorus atom of the phosphotyrosyl substrate, resulting in formation of a phospho-Cys intermediate. In the second step, the transient phosphoenzyme intermediate is hydrolyzed by an activated water molecule (Barford et al., 1995). Due to the unique environment of the PTP active site, the pK␣ of the sulfhydryl group of this Cys residue is extremely low (ⵑ5.4 in PTP1B [Lohse et al., 1997] and ⵑ4.7 in YOP [Zhang and Dixon, 1993]) compared to the typical pK␣ for Cys (ⵑ8.5), which favors its function as a nucleophile but renders it susceptible to oxidation. It has now been shown that treatment of various PTPs (Lee et al., 1998), dual-specificity phosphatses (Denu and Tanner, 1998), and low molecular weight PTPs (Caselli et al., 1998) with H2O2 in vitro leads to oxidation of the active site Cys to sulfenic acid. Such oxidation results in inhibition of activity because the modified Cys can no longer function as a phosphate acceptor in the first step of the PTP-catalyzed reaction. Oxidation of Cys to sulfenic acid is reversible (Claiborne et al., 1999) and thus has the potential to form the basis of a mechanism for reversible regulation of PTP activity. In contrast, oxidation by the addition of two (sulfinic acid) or three (sulfonic acid) oxygens to the active site Cys is irreversible. Interestingly, glutathionylation of the sulfenic acid form of PTP1B has been reported (Barrett et al., 1999) and proposed as a mechanism to protect against further, irreversible oxidation and as an important step in the reverse, reduction mech-

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Figure 1. Use of the In-Gel Assay to Identify PTPs that Are Susceptible to StimulusInduced Oxidation Cells were triggered with the appropriate stimulus and harvested under anaerobic conditions in lysis buffer containing IAA. Those PTPs that had not encountered ROS in the cell became irreversibly inactivated by alkylation of their active site Cys with IAA. However, in contrast, any PTPs in which the active site Cys had been oxidized in response to the stimulus were resistant to alkylation. For the in-gel phosphatase assay, a 10% SDS-PAGE gel was cast containing a radioactively labeled substrate. An aliquot of cell lysate was subjected to SDS-PAGE, and proteins in the gel were sequentially denatured, then renatured in the presence of reducing reagents. Under these conditions, the activity of the PTPs in which the active site Cys had been subjected to stimulus-dependent oxidation to sulfenic acid was recovered, whereas those that were not oxidized in response to the initial stimulus and were irreversibly alkylated in the lysis step remained inactive. The reaction was then terminated by fixing, staining, and destaining the gel. Finally, the gel was dried and exposed to film. The presence of a PTP was visualized by substrate dephosphorylation as the appearance of a clear, white area on the black background of labeled substrate.

anism. Rhee’s laboratory has recently shown that stimulation of A431 cells with EGF leads to the production of H2O2 and concomitant inhibition of PTP1B (Bae et al., 1997). Increased production of intracellular oxidants may contribute to enhanced, tyrosine phosphorylationdependent signaling, for example in response to growth factors, (Bae et al., 1997, 2000; Sundaresan et al., 1995), by transiently suppressing the enzymatic activity of members of the PTP family, thereby promoting a burst of PTK activity (Finkel, 1998, 2000). However, it was unclear how broadly this phenomenon may apply across the PTP family in a cellular context. In the present study, we have explored the potential of reversible oxidation as a general mechanism for regulation of PTP function. A major obstacle to such studies is the requirement for a method by which the oxidized/ inactivated PTPs could be distinguished from reduced/ activated PTPs in a cellular context. We have modified an “in-gel” PTP activity assay to allow visualization of a profile of oxidized PTPs following a particular stimulus. Our results have illustrated that several PTPs were oxidized and inactivated reversibly in Rat-1 cells following stimulation with H2O2 and that this was important for peroxide-induced mitogenic signaling. Furthermore, we have demonstrated that PDGF induced the oxidation and inhibition of the SH2 domain-containing PTP, SHP-2, which facilitated mitogenic signaling in response to the growth factor. The potential for extending these analyses to identify and characterize other PTPs and their role in the control of a broad array of signal transduction pathways is discussed below.

Results Exposure to Hydrogen Peroxide Results in the Reversible Inactivation of PTPs in Rat-1 Cells We wished to test the hypothesis that ROS stimulated intracellular tyrosine phosphorylation through the oxidation and inhibition of cellular PTPs. To pursue this line of investigation, we developed a modified in-gel PTP activity assay. As described in Figure 1, the PTPs that registered as active in this assay would be those originally protected from post-lysis alkylation by a stimulusdependent modification at the active site Cys, which was reversed by DTT, consistent with oxidation of the

Cys to sulfenic acid. The data shown in Figure 2A illustrate that iodoacetic acid (IAA) in the lysis buffer effectively inactivated PTPs in a lysate of Rat-1 cells (lane 2, compare to lane 1) via irreversible alkylation of the invariant, active site Cys residue of these enzymes (Zhang and Dixon, 1993). However, when H2O2 was added to the culture media it gained rapid access to the intracellular environment and within 1 min the active site Cys residue of various PTPs was oxidized, thereby protecting them from alkylation by IAA (Figure 2A, lanes 3–7). Furthermore, we observed that 200 ␮M H2O2 was sufficient to oxidize all of the PTPs detectable in this assay format, but more extensive oxidation occurred at higher concentrations of H2O2 (Figure 2A). In exploring further whether there was a link between oxidation/inhibition of PTPs and enhanced tyrosine phosphorylation in Rat-1 cells, we observed that the tyrosine phosphorylation of proteins of ⵑ120 kDa and 70 kDa was induced in a dose-dependent fashion coincident with exposure of cells to H2O2 (Figure 2B). This stimulation also triggered the phosphorylation of ERK MAP kinases (MAPKs) (data not shown). N-acetyl cysteine (NAC), a widely used ROS scavenger, blocked PTP oxidation and inactivation induced by 200 ␮M H2O2, thus confirming that the effects on PTP activity shown in the in-gel assay were due to H2O2-induced intracellular oxidation (Figure 2C). In addition, depletion of the cellular pool of glutathione (GSH) by exposure of the cells to L-buthionine-SR-sulfoximine (BSO), a specific inhibitor of ␥-glutamylcysteine synthetase, markedly attenuated the recovery of PTP activity following removal of an H2O2 stimulus (Figure 2D). Stimulation with H2O2 led to oxidation of several PTPs (Figure 2D, lane 2) which were quickly reduced once H2O2 was removed (lanes 3–6). Recovery was essentially complete within 10–20 min of removal of H2O2. However, when the same analysis was performed on Rat-1 cells that had been subjected to pretreatment with BSO, oxidation persisted even 30 min after removal of H2O2 (Figure 2D, lanes 8–12). Therefore, glutathionylation in conjunction with the reducing function of thioltransferase may represent a mechanism for reactivation of ROS-inactivated PTPs in vivo, consistent with previous observations of glutathionylation of PTP1B in vitro (Barrett et al., 1999). However, it is important to note that other cellular reducing systems, such as thioredoxin (Lee et al., 1998), may also contribute to the reactivation of oxidized PTPs in vivo. The data presented in Figure 2 indicate that the

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Figure 2. Treatment of Rat-1 Cells with H2O2 Triggered Reversible Oxidation of Multiple PTPs Concomitant with Tyrosine Phosphorylation (A) Serum-deprived Rat-1 cells were exposed to various concentrations of H2O2 for 1 min, harvested, and lysed in the absence (lane 1) or presence (lanes 2–7) of 10 mM IAA. Aliquots of lysate were subjected to the in-gel PTP assay. (B) Tyrosine phosphorylated proteins immunoprecipitated from lysates of H2O2-treated cells with Ab PT-66, then immunoblotted with antipTyr Ab (G104). (C) After preincubation in the absence or presence of 30 mM NAC for 40 min and removal of excess NAC by two washes with fresh culture medium, Rat-1 cells were exposed to 200 ␮M H2O2 and lysed in the presence of 10 mM IAA at the indicated times. Lysates were subjected to in-gel PTP assay. (D) Rat-1 cells were serum starved in the absence or presence of 2.5 mM BSO for 16 hr. H2O2 (200 ␮M) was added for 2 min, then removed by washing the cells with fresh culture media. Incubation was then continued until harvesting in lysis buffer containing 10 mM IAA at the times indicated. Oxidized PTPs were visualized by in-gel assay. Arrowheads indicate PTPs whose reduction/reactivation displayed dependence on intracellular GSH.

intracellular balance between the activities of PTKs and PTPs was indeed altered by oxidative stress in Rat-1 cells, leading to enhanced tyrosine phosphorylation. To the best of our knowledge, this is the first demonstration that multiple PTPs may be oxidized and inactivated by ROS in a cellular environment. H2O2-Induced Mitogenic Signaling Is Associated with Inactivation of PTPs In order to explore the importance of oxidation and inhibition of PTP function for ROS-induced mitogenesis, we

tested the effects of H2O2 and the synthetic ROS t-butyl hydroperoxide (t-BHP). Initially, we compared the susceptibility of an activated mutant form of SHP-2 (E76A) to alkylation by IAA following treatment with either H2O2 or t-BHP. Using the modified in-gel PTP assay, we showed that SHP-2, which had been pretreated with PBS, was inactivated by IAA (Figure 3A, lane 2, compare to lane 1), whereas oxidation with H2O2 protected SHP-2 from alkylation (Figure 3A). It is interesting to note that even at 2 mM H2O2, SHP-2 was not irreversibly oxidized since its activity was recovered in the in-gel assay (Fig-

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Figure 3. H2O2-Induced Mitogenic Signaling Was Associated with Inactivation of PTPs (A) Purified SHP-2 (E76A mutant) was incubated with either PBS, H2O2, or t-BHP at 37⬚C for 5 min. Aliquots were then incubated at room temperature for a further 5 min either in the absence (⫺ IAA) or presence (⫹ IAA) of 4 mM IAA and subjected to in-gel assay (1 ng SHP-2/ lane). (B) Rat-1 cells were preloaded with 20 ␮M H2DCFDA in the dark for 20 min, then exposed to H2O2 and t-BHP (both 200 ␮M) for 5 min. Images of ROS-induced DCF fluorescence are shown at magnification 400⫻ (upper panels). Cells (1 ⫻ 105) that underwent the same treatment as above were harvested and resuspended in Hanks’ solution, then immediately subjected to flow cytometric analysis to measure ROS-induced DCF fluorescence. The basal peak indicates background fluorescence, whereas the rightward-shifted peak indicates ROS-induced DCF fluorescence (lower panels). (C) Cells were exposed to H2O2 and t-BHP (each at 200 ␮M) for the indicated times, lysed in the presence of 10 mM IAA and oxidized PTPs visualized in the in-gel assay. (D) After exposure to H2O2 and t-BHP (each at 200 ␮M), lysates were prepared and pTyr proteins were immunoprecipitated with Ab PT-66, then immunoblotted with anti-pTyr Ab G104 (upper panel). An aliquot of lysate from each treatment was immunoblotted with anti-phosphoMAPK Ab and subsequently with anti-MAPK Ab (lower panel).

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ure 3A). In contrast, t-BHP was unable to oxidize and inactivate SHP-2 in vitro and thus did not protect the invariant Cys residue of SHP-2 from alkylation (Figure 3A), consistent with a previous report which indicated that t-BHP did not inhibit PTP activity in vitro and suggested that the bulky nature of t-BHP may exclude it from the PTP active site cleft (Denu and Tanner, 1998). We then compared the effects of H2O2 and t-BHP on inactivation of PTPs and activation of MAPK signaling pathways in a cellular context. Initially, we showed by fluorescence microscopy of Rat-1 cells preloaded with H2DCFDA that treatment with either H2O2 or t-BHP led to rapid oxidation and the appearance of the fluorescent derivative, DCF (Figure 3B, upper panels). Furthermore, upon flow cytometric analysis no quantitative difference was observed between the H2O2- and t-BHP-induced shift of fluorescence (Figure 3B, lower panels). Therefore, similar levels of intracellular ROS were produced following treatment with either reagent. However, when we examined their ability to oxidize PTPs in the cells, we detected reproducible inactivation of PTPs in response to H2O2 but not in response to t-BHP (Figure 3C). Finally, we compared the effects of H2O2 and t-BHP on tyrosine phosphorylation of cellular proteins and activation of MAPKs. As shown in Figure 3D, we observed that the inactivation of PTPs by H2O2 was associated with enhanced tyrosine phosphorylation and mitogenic signaling. In contrast, t-BHP, which lacks the ability to oxidize and inactivate the PTPs, elicited less pronounced effects on tyrosine phosphorylation and was unable to activate MAPKs (Figure 3D). These results are consistent with a pivotal role of the inactivation of PTPs in the mitogenic effects of ROS. Oxidation of a 70k PTP Is Associated with PDGFInduced Mitogenic Signaling in Rat-1 Cells We have observed that treatment of Rat-1 cells with H2O2 led to inactivation of multiple PTPs (Figures 2 and 3). In the next phase of the study, we wished to explore whether the production of ROS in response to physiological stimuli also resulted in oxidation and inactivation of members of the PTP family and whether there was specificity in the response. Initially, we examined the effects of a peptide growth factor, PDGF, which has been shown to produce ROS in various cell types (Bae et al., 2000; Sundaresan et al., 1995). As expected, treatment of Rat-1 cells with PDGF resulted in a rapid increase in the tyrosine phosphorylation of cellular proteins and the enhanced phosphorylation of MAPKs (data not shown). Therefore, we conducted the modified ingel PTP activity assay on lysates of PDGF-stimulated Rat-1 cells. Our data demonstrated that PDGF stimulation induced a rapid and transient oxidation of a PTP of ⵑ70k. Oxidation of this 70k PTP was reversible, reaching a maximum at 5 min followed by marked reduction, almost to basal levels, within 20 min of PDGF treatment (Figure 4A). We investigated the possible role of oxidation/inactivation of the 70k PTP in regulating PDGFR-mediated signaling by testing the effects of the antioxidant NAC. Cells were incubated in the presence or absence of NAC prior to PDGF stimulation. Then we utilized the modified in-gel PTP assay to examine the effects of the growth

factor on the activity of the 70k PTP. We observed that when the levels of PDGF-induced ROS were reduced by pretreatment with NAC, oxidation of the 70k PTP was markedly attenuated (Figure 4B). Furthermore, the ligand-induced tyrosine phosphorylation of the PDGFR was greatly diminished, and the activation of MAPKs was completely eliminated, in NAC-treated cells (Figure 4C). These data suggest that the rapid, transient inactivation of 70k PTP may be important for concomitant PDGFR-mediated phosphorylation and mitogenic signaling. Identification of the 70k PTP as SHP-2 In attempting to identify the 70k PTP that was oxidized following PDGF stimulation, our attention was drawn to the SH2 domain-containing PTP, SHP-2. This PTP has been shown to be associated with tyrosine phosphorylated PDGFR (Lechleider et al., 1993). In addition, the apparent molecular weight of SHP-2 on SDS-PAGE is similar to that of the PDGF-responsive 70k PTP detected in Figure 4. Initially, we confirmed that SHP-2 could be recruited by the ligand-activated PDGFR in Rat-1 cells. As shown in Figure 5A (top panel), a tyrosine phosphorylated protein of ⵑ70 kDa by SDS-PAGE associated rapidly with the PDGFR in response to ligand activation. Furthermore, we showed by immunoblotting that SHP-2 comigrated with this 70k phosphoprotein (Figure 5A, lower panels). The complex between PDGFR and SHP-2 persisted up to 20 min after stimulation, then the level of association decreased (Figure 5A, lower panels). To test whether SHP-2 was the 70k PTP that was oxidized following PDGF stimulation, we immunodepleted SHP-2 protein from cell lysates with increasing amounts of antiSHP-2 antibody and subjected the supernatants to the modified in-gel PTP assay. As shown in Figure 5B, antiSHP-2 antibody depleted the 70k PTP from Rat-1 cell lysates, whereas an anti-SHP-1 antibody control did not. These data identify SHP-2 as a PTP that was rapidly oxidized and inactivated following PDGF stimulation. We examined the association of other SH2 domaincontaining proteins with activated PDGFR. It has been shown that SHP-2 dephosphorylates the PDGFR on the autophosphorylation sites that function as binding sites for GTPase-activating protein (GAP) and phosphatidylinositol 3 kinase (PI3K) (Klinghoffer and Kazlauskas, 1995). However, we observed that both GAP and the p85 subunit of PI3K were recruited by PDGFR rapidly after ligand stimulation, even though SHP-2 was associated with the receptor at this time (Figure 5A). These results suggest that oxidation and inactivation of SHP-2 in response to PDGF may be important for permitting recruitment of GAP and PI3K by the activated PDGFR. Interestingly, GAP and PI3K dissociated from the receptor by 10 min after PDGF stimulation (Figure 5A), coincident with dephosphorylation of PDGFR␤ (Figure 5A) and reactivation of SHP-2 (Figure 4A). Specificity in Production of ROS, Oxidation and Inactivation of SHP-2 in Response to Growth Factor Stimulation Although the potential for PTPs to inhibit signaling through antagonizing PTK function is intuitively obvious, it has also become apparent that certain PTPs have the

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Figure 4. PDGF-Induced Oxidation of a 70k PTP in Rat-1 Cells (A) Serum-starved Rat-1 cells were exposed to 50 ng/ml PDGF-BB for the times indicated. Lysates were prepared in the presence of 10 mM IAA and subjected to in-gel PTP assay. The arrowhead indicates a 70k PTP that was transiently oxidized following stimulation of Rat-1 cells with PDGF. The result shown is a representative of four independent experiments. (B) Cells were preincubated in the absence or presence of 30 mM NAC for 40 min. Excess NAC was removed prior to addition of PDGF (50 ng/ml). PDGF-induced oxidation of the 70k PTP, which was impaired in the presence of NAC (arrowhead), was visualized by the modified in-gel PTP assay. (C) Cells were treated with NAC and PDGF as described above. PDGFR was immunoprecipitated from lysates with Ab-X and immunoblotted with anti-pTyr Ab G104. The same filter was subsequently reprobed with Ab-X (upper panels). Aliquots of cell lysate from each treatment were immunoblotted with anti-phosho-MAPK Ab and reprobed with anti-MAPK Ab (lower panels).

potential to function positively in a coordinate manner with PTKs to promote signaling. SHP-2 was one of the first PTPs to be recognized as playing such a role, particularly in the context of EGF and FGF receptor signaling (Bennett et al., 1996; Saxton et al., 1997). In contrast, our data showing oxidation and inhibition of SHP-2 in response to PDGF would be indicative of a negative role in signaling. Therefore, we have explored this apparent conundrum further. Treatment of Rat-1 cells with PDGF triggered production of intracellular ROS (Figure 6A) concomitant with oxidation and inactivation of SHP-2 (Figure 6B). In contrast, we were unable to detect production of ROS in response to either EGF or FGF (Figure 6A). Furthermore, using our modified in-gel PTP assay, we did not observe oxidation and inhibition of SHP-2 in response to EGF or FGF (Figure 6B). Nevertheless, EGF, FGF, and PDGF all activated MAPK to a similar extent in Rat-1 cells (Figure 6C). These results indicate that of the stimuli we examined in Rat-1 cells, transient oxidation and inactivation of SHP-2 is a specific response to PDGF, consistent with differences in the function of SHP-2 in these distinct growth factor signaling pathways. The PDGFR-Associated Pool of SHP-2 Was Susceptible to Oxidation and Inactivation Recent studies have suggested that a Rac1-associated, plasma membrane-bound NADPH oxidase is responsible for PDGF-induced generation of ROS in nonphago-

cytic cells (Bae et al., 2000). In light of the short half-life of such ROS, it is possible that their influence on PTPs may be spatially restricted to the subcellular regions proximal to their production. We had observed that only ⵑ10% of the total population of SHP-2 was recruited into a complex with the PDGFR following ligand stimulation in Rat-1 cells (data not shown). To examine whether this recruitment was required for oxidation and inactivation of SHP-2 in response to PDGF, we tested the ability of mutant forms of the PDGFR, which were deficient in their association with SHP-2, to induce oxidation of the PTP in response to ligand. We constructed chimeric receptors that consisted of the extracellular segment of human granulocyte colony stimulating factor (G-CSF) receptor and the transmembrane and cytoplasmic segments of human PDGFR. In this way, we could study G-CSF-induced recruitment of SHP-2 to the chimeric receptors and signaling in Rat-1 cells, which do not express endogenous G-CSF receptor (G-CSFR), while avoiding activation of endogenous PDGFR. The autophosphorylation site at Y 1009 of human PDGFR has been shown to be the major docking site for the N-terminal SH2 domain of SHP-2 (Lechleider et al., 1993). Therefore, we constructed chimeric receptors comprising both wild-type (WT) and Y1009F forms of the PDGFR intracellular segment and transiently transfected the chimeric receptors into Rat-1 cells. Upon stimulation with G-CSF, both WT and Y1009F chimeric receptors were tyrosine phosphorylated (Figure

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PDGFR (Figure 7A). Using the modified in-gel PTP assay, we observed that in response to G-CSF stimulation, WT chimeric receptors triggered rapid oxidation and inactivation of SHP-2. In contrast, activation of Y1009F mutant receptors did not induce oxidation of SHP-2 (Figure 7B). These data illustrate that recruitment of SHP-2 by activated, chimeric PDGFR was required for oxidation of the PTP by ROS generated in response to ligand. The observation that PDGF induced the oxidation and inhibition of SHP-2 (Figures 4–6) is indicative of an inhibitory function for this PTP in PDGFR signaling. To explore this possibility further, we utilized the G-CSF:PDGF receptor chimeras. A time course of exposure to G-CSF illustrated that both WT and Y1009F SHP-2 docking site mutant receptors were rapidly tyrosine phosphorylated following ligand stimulation. However, whereas tyrosine phosphorylation of the WT receptor was transient, the mutant receptor was maintained at a higher level of phosphorylation throughout the time course (Figure 7C). The differences were particularly striking at the later time points following 20 and 30 min of ligand stimulation. To explore further the importance of SHP-2 in regulation of downstream signaling events, we focused on the phosphorylation status of MAPKs. We observed maximal phosphorylation of p42 and p44 ERKs following 20 min of stimulation (Figure 7D). However, we observed that both the extent and duration of ERK phosphorylation was higher in cells expressing the mutant receptor, which was deficient in binding of SHP-2, compared to those expressing the WT receptor (Figures 7D and 7E). These results indicate that recruitment of SHP-2 into the PDGF receptor-containing signaling complex is important for downregulation of both receptor tyrosine phosphorylation and activation of MAPK. The data are also consistent with the conclusion that oxidation and inhibition of SHP-2 in the early phase of the response to PDGF is important for establishment of the signaling response. Discussion Figure 5. Identification of the 70k PTP that was Susceptible to PDGF-Induced Oxidation as SHP-2 (A) Serum-starved Rat-1 cells were exposed to PDGF (50 ng/ml) for the indicated times. The PDGFR and associated proteins were immunoprecipitated with antibody Ab-X, and pTyr proteins were visualized by immunoblotting with anti-pTyr Ab G104 (upper panel). The same filter was reprobed with anti-PDGFR, anti-SHP-2, antiGAP, and anti-p85 PI3K Abs. The positions of PDGFR (solid arrowhead) and SHP-2 (open arrowhead) are indicated. (B) Rat-1 cells, either untreated (⫺) or stimulated with 50 ng/ml PDGF (⫹), were harvested in lysis buffer containing 10 mM IAA. Lysates were incubated with antibody to either SHP-2 or SHP-1 and subjected to an in-gel PTP assay (upper panel). The arrowhead denotes the position of the 70k PTP that was inactivated in response to PDGF and immunodepleted from cell lysates with antibodies to SHP-2. The lower panel illustrates an immunoblot to show the immunodepletion of SHP-2.

7A). Although both receptors were activated following treatment with G-CSF, only the WT recruited SHP-2, which was recovered in immune complexes precipitated with antibodies to the intracellular segment of the

Recently, a variety of studies have indicated that ROS may function as second messengers to promote mitogenic signaling in various cell types (reviewed in Adler et al., 1999). Although ROS have been shown to stimulate tyrosine phosphorylation-dependent signaling pathways, including the activation of various PTKs (Brumell et al., 1996), the precise molecular targets for these effects have yet to be identified. In the present study, we have demonstrated that multiple PTPs are oxidized and inactivated by ROS in vivo. Using a modified in-gel PTP assay, we have shown that several PTPs, ranging from 40–120 kDa, were oxidized rapidly and reversibly following stimulation of Rat-1 cells with H2O2 (Figure 2). Furthermore, a comparison of the effects of H2O2 and t-BHP on PTP inhibition and tyrosine phosphorylation revealed that oxidation of PTPs is important for mitogenic signaling induced by ROS (Figure 3). We observed that stimulation of Rat-1 cells with ⬍1 mM H2O2 in the extracellular media led to oxidation of multiple intracellular PTPs (Figure 2). Interestingly, it has been shown that physiological stimuli, such as PDGF in vascular smooth

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Figure 6. The Oxidation and Inactivation of SHP-2 Was Induced by PDGF but Not by EGF or FGF (A) Rat-1 cells were incubated with 20 ␮M CM-H2DCFDA in the dark for 20 min, then exposed to peptide growth factors (50 ng/ ml) for an additional 10 min. Images of ROSinduced DCF fluorescence are shown at 50⫻ magnification. The data are representative of four independent experiments. (B) Cells were exposed to peptide growth factors for the indicated times, lysed in the presence of 10 mM IAA, and oxidized PTPs were visualized by an in-gel assay. (C) Aliquots of cell lysate from each treatment were immunoblotted with anti-phosphoMAPK Ab and reprobed with anti-MAPK Ab.

muscle cells, can induce a transient increase in the intracellular concentration of H2O2 which is equivalent to that achieved by stimulating the cells with H2O2 at an extracellular concentration in the range of 0.1–1.0 mM (Sundaresan et al., 1995). Thus, the concentrations of H2O2 used in this study would be likely to reflect a physiologically relevant condition. Nevertheless, it was important to demonstrate that such oxidation occurred in response to a physiological stimulus. PDGF-stimulated production of ROS has been well documented in a variety of cells (Bae et al., 2000; Sundaresan et al., 1995). Furthermore, attenuation of the rise in intracellular H2O2 by ectopic expression of catalase inhibited PDGF-induced tyrosine phosphorylation and mitogenesis (Sundaresan et al., 1995). Consistent with previous reports (Lechleider et al., 1993), we observed that association between SHP-2 and the PDGFR was induced by ligand stimulation. The phosphatase was engaged in a complex with the receptor within 2 min of

addition of PDGF, which persisted until ⵑ20 min after stimulation (Figure 5). It has been shown that SHP-2 not only binds to the PDGFR but also recognizes the receptor as a substrate (Klinghoffer and Kazlauskas, 1995). Thus, one would anticipate that following binding of SHP-2, tyrosine phosphorylation of the PDGFR would be low. However, we noted that the peak of autophosphorylation of the PDGFR occurred during the time in which SHP-2 was associated with the receptor (Figure 5). Furthermore, although it has been reported that SHP-2 preferentially dephosphorylates pTyr residues 771 and 751 of PDGFR␤ (Klinghoffer and Kazlauskas, 1995), which correspond to binding sites of Ras-GAP and PI3K, respectively (Kazlauskas et al., 1992), we observed that both of these proteins were recruited into a complex with the phosphorylated PDGFR while SHP-2 was also present in that complex (Figure 5A). This apparent anomaly was noted previously by Klinghofer and Kazlauskas, who proposed that the activity of SHP-2

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Figure 7. The Pool of PDGFR-Associated SHP-2, which Is Oxidized and Inactivated in Response to PDGF, Is Also Involved in Downregulation of MAPK Signaling Rat-1 cells were transiently transfected with plasmids expressing WT or Y1009F mutant G-CSFR/PDGFR chimeric receptor or with a plasmid encoding green fluorescent protein (GFP) as a control for expression. (A) After exposure to 100 ng/ml G-CSF for 5 min, the chimeric receptors were immunoprecipiated from lysates with antibody Ab-X and immunoblotted with anti-pTyr Ab G104. Immunoprecipitation of the receptors was verified by immunoblotting with Ab-X. The same filter was stripped and reprobed with anti-SHP-2 Ab. Expression of the chimeric receptors was verified by immunoblotting an aliquot of each lysate with Ab-X, which recognizes the intracellular segment of the PDGFR, and subsequently with anti-G-CSFR Ab, which recognizes the extracellular segment of chimeric receptors. (B) Transfected Rat-1 cells were treated with G-CSF for the indicated times, lysed in the presence of 10 mM IAA, and the lysates were subjected to an in-gel assay. The arrow denotes the position of SHP-2. (C) The WT and mutant chimeric receptors were immunoprecipitated at the indicated times and immunoblotted with anti-pTyr Ab (G104). The same filter was reprobed with anti-PDGFR Ab-X. (D) Aliquots of lysate from each treatment were subjected to immunoblotting with anti-phosho-MAPK Ab, then reprobed with anti-MAPK Ab. (E) Densitometric analysis of the gel image illustrates the ratio of phosphorylated ([D], top panel) over total ([D], bottom panel) MAPK.

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must be inhibited immediately following receptor activation, possibly by phosphorylation of the phosphatase (Klinghoffer and Kazlauskas, 1995). Our data reveal that SHP-2 was rapidly oxidized in Rat-1 cells stimulated with PDGF (Figures 4–6). The time course of oxidation coincided with that of autophosphorylation of the PDGFR (Figures 4 and 5). Furthermore, we also demonstrated that the recruitment of SHP-2 into a complex with the activated PDGFR was essential for oxidation and inactivation of the PTP (Figure 7). Therefore, our results now reveal that the ligand-induced, transient oxidation and inactivation of SHP-2 in the PDGFR signaling complex may provide the missing regulatory link that is important for facilitating these initial autophosphorylation events. SHP-2 is a ubiquitously expressed cytosolic PTP comprising two SH2 domains at the N terminus, a PTP catalytic domain and a C-terminal segment containing a prolyl-rich motif and sites for tyrosine phosphorylation. The crystal structure of SHP-2 has been solved, illustrating that in the ground state the PTP adopts an inactive conformation in which the active site is occluded by the N-terminal SH2 domain (N-SH2) (Hof et al., 1998). Following engagement by specific phosphotyrosyl ligands—for example, autophosphorylation sites on RPTKs—at a site distinct from that which interacts with the PTP domain, the N-SH2 loses complimentarity for the active site. This promotes adoption of an open, active conformation in which the catalytic site of SHP-2 is now free to interact with substrates (Hof et al., 1998). One would predict that occlusion of the active site of SHP-2 by the N-SH2 domain in the ground state would impede access not only of substrates but also access of ROS. The data in Figure 7 illustrate that recruitment to the PDGFR, which would induce a change to the open, active conformation of SHP-2, was required for oxidation of the PTP, an observation that is consistent with these structural considerations. However, the question of whether it is only the receptor-bound pool of SHP-2 that is oxidized is more difficult to answer. Unfortunately, the lysis conditions that are necessary to preserve oxidation of SHP-2 also disrupt its association with the PDGFR, and therefore it was not possible to compare directly the extent of oxidation of receptorbound and soluble pools of the PTP. Nevertheless, we examined the distribution and extent of oxidation of SHP-2 between saponin-soluble and insoluble fractions of PDGF-stimulated Rat-1 cells. Although we observed a low, constant baseline in the soluble, cytosolic fraction, the majority of ligand-induced oxidation of SHP-2 (⬎95% at 2 min of PDGF stimulation) occurred in the membrane fraction, and we did not detect translocation of oxidized SHP-2 into the cytosol (data not shown). Thus, it is likely that the majority of the oxidized SHP-2 will be found in the PDGFR-signaling complex. Initially, it was suggested that SHP-2 functioned predominantly in a positive manner to promote signal transduction pathways. For example, genetic studies have indicated that csw, the Drosophila counterpart of mammalian SHP-2, is a positive transducer of signals in the Torso pathway, directing normal development of terminal structures in embryogenesis (Perkins et al., 1992). In addition, the C. elegans homolog PTP-2 is a positive regulator of the Let23/EGF receptor pathway (Gutch et

al., 1998). In both cases, the PTP appears to play a role in setting the intensity of the signaling response. SHP-2 has also been shown to play a positive role in mammalian systems, such as in regulating signaling downstream of the FGF and EGF receptors (Bennett et al., 1996; Saxton et al., 1997). As anticipated for a positive regulator of signaling, we observed that the activity of SHP-2 was preserved following stimulation with either FGF or EGF (Figure 6). However, our observation of PDGF-induced oxidation and inhibition of SHP-2 is more consistent with the PTP functioning as an inhibitor of signaling in response to this growth factor. Interestingly, there are now several examples of such negative, signalattenuating roles of SHP-2 (Marengere et al., 1996; Fukada et al., 1999). Targeted mutations in murine SHP-2 have also been reported. Exon 3 knockout mice, in which a truncated phosphatase lacking the N-SH2 is expressed at ⵑ25% of WT levels, display embryonic lethality in mid-gestation due to defects in gastrulation (Saxton et al., 1997). Strikingly, primary cell cultures from the mutant embryos display defects in receptor PTK signaling. Although stimulation of MAPK by FGF is attenuated in SHP-2 mutant cells, consistent with a positive role of SHP-2 in FGF-induced signaling, PDGF-induced activation of MAPK is actually enhanced, suggesting relief of an inhibitory constraint upon PDGF signaling by disruption of SHP-2 (Saxton et al., 1997). However, the concept that SHP-2 functions as an inhibitor of PDGF-induced signaling is somewhat controversial. In contrast to Saxton et al. (1997), Feng’s lab established SV40 Large T immortilized cell lines from the same mice and concluded that SHP-2 played a positive role in the activation of ERK MAPK in response to PDGF (Shi et al., 1998). One of the striking features of these T antigen immortilized cell lines is that the levels of PDGFR-␤ but not PDGFR-␣ are dramatically reduced due to decreased stability of the PDGFR-␤ mRNA (Lu et al., 1998). Thus, these cells will primarily reflect signaling responses of PDGFR-␣. There are several important distinctions between the signaling events mediated by the ␣ and ␤ forms of the PDGF receptor, including differential regulation of the Ras-MAPK pathway (Klinghoffer et al., 2001) and the observation that signaling through PDGFR-␣ has the capacity to anatgonize PDGFR␤-induced transformation (Yu et al., 2000). SHP-2 interacts with distinct sites on the two forms of the PDGFR, Tyr 760 in the kinase-insert motif of PDGFR-␣ (Bazenet et al., 1996) and Tyr 1009 in the C-terminal segment of PDGFR-␤ (Lechleider et al., 1993), and whereas it appears that PDGFR-␤ is a substrate for SHP-2 (Klinghoffer and Kazlauskas, 1995), PDGFR-␣ is not (Bazenet et al., 1996). Furthermore, and of particular importance to our study, it has been reported that PDGF-AA, which stimulates the PDGFR-␣ homodimer specifically, did not induce production of ROS in human aortic smooth muscle cells, whereas PDGF-AB and BB, which also recognize the ␤ form of the PDGFR, both stimulated production of superoxide (Marumo et al., 1997). Therefore, these discrepancies in the literature may actually reflect differential signaling responses of the ␣ and ␤ forms of the PDGF receptor and differential roles of SHP-2 in regulating those responses. It is an oversimplification to think of PTPs as acting either exclusively as negative or positive regulators of signal transduction. Rather, the

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role of the PTP, such as SHP-2, will depend upon both the cellular and signaling context. In conclusion, our study demonstrates the potential for ROS-induced oxidation and inactivation of multiple PTPs, illustrating a further aspect of the regulation of tyrosine phosphorylation-dependent signal transduction. We have also shown that PTPs respond not only to oxidative stresses in the environment but also to intracellular ROS generated in response to physiological activation of cell surface receptors. Interestingly, it is known that UV (Gross et al., 1999), ionizing irradiation (Kyprianou et al., 1997), and agonists of G protein coupled receptors (GPCRs) (Daub et al., 1997; Prenzel et al., 1999) may induce transactivation of receptor PTKs. The proteolytic processing of proHB-EGF by a metalloproteinase to yield a functional growth factor was proposed as one mechanism for GPCR-induced transactivation of the EGF receptor (Prenzel et al., 1999). However, a common characteristic of the stimuli listed above is their ability to promote the generation of intracellular ROS (Chen et al., 1995; Holland et al., 1998; Ushio-Fukai et al., 1999). Thus, it is plausible that the inhibition of PTP activity associated with a burst of intracellular ROS could contribute to this transactivation of RPTKs. It will be intriguing to pursue the strategies we have described in this study to utilize stimulus-induced oxidation as a means of “tagging” those PTPs that are integral to the regulation of the signal transduction pathways initiated by that stimulus. Alkylation with IAA can be used to subtract out the bulk of the PTPs that are unaffected by the stimulus, and following reduction the responsive PTPs can be isolated and identified. However, it is important to note that some PTPs, in particular the receptor-like forms, do not renature efficiently in the in-gel assay (Burridge and Nelson, 1995). Therefore, proteomics-based strategies, using immobilized, broadspecificity, active site-directed inhibitors (such as phenylarsine oxide coupled to agarose) as affinity matrices should represent a powerful and complimentary technique for the purification and identification of oxidationsensitive PTPs. By combining these approaches with the use of substrate-trapping mutant forms of the PTPs thus identified (Flint et al., 1997), it should be possible to characterize the physiological substrate specificity of these enzymes and thus gain insights into their signaling function. Hopefully, application of these strategies will provide further insights into the physiological function of this important family of signal transducing enzymes. Experimental Procedures Materials Antibodies were purchased from the following companies: Santa Cruz (SHP-1 [C-19], SHP-2 [C-18], and PI3K [Z-8]), Cell Signaling (phospho-MAPK and MAPK), Transduction Laboratories (GAP), and Sigma (pTyr Ab PT66). The anti-pTyr antibody G104 was described previously (Garton et al., 1997). Anti-PDGFR␤ antibody (Ab-X) was a gift from Dr. Daniel DiMaio at Yale University (Irusta and DiMaio, 1998). Anti-human G-CSF receptor (G-CSFR) antibody was provided by Dr. Toshio Hirano at Osaka University, Japan (Fukada et al., 1996). Construction of Expression Vectors for Chimeric Receptors Full-length cDNA encoding wild-type (WT) and Y1009F mutant forms of human PDGFR␤ was provided by Dr. Jonathan Cooper (Fred

Hutchinson Cancer Center, Seattle, WA; Kashishian and Cooper, 1993). The cDNA encoding the extracellular segment of human G-CSFR was a gift from Dr. Shigekazu Nagata (Osaka University, Japan; Fukada et al., 1996). Chimeric receptors comprising the extracellular segment of G-CSFR fused to the transmembrane and intracellular (WT and Y1009F) segments of PDGFR␤ were constructed in the pcDNA3.1A vector (Invitrogen) by standard PCR protocols, then inserted into a pRK5 expression vector for transient transfection experiments. The integrity of the constructs was confirmed by sequencing. Transient Transfection, Immunoprecipitation, and Immunoblotting Rat-1 fibroblasts were routinely maintained in DMEM supplemented with 10% FBS, 1% glutamine, 100 U/ml penicillin, and 100 ␮g/ml streptomycin. For stimulation with H2O2 and peptide growth factors, cells were plated in media containing 10% FBS for 48 hr, then serum starved for 16 hr before treatment. For transient transfection, Rat-1 cells were plated in DMEM medium supplemented with 10% FBS for 16 hr. The culture medium was replaced by OptiMEM (Life Technology) without serum, then plasmid (5 ␮g/dish) was introduced into cells by LipofectAMINE and PLUS reagents (Life Technology) according to the manufacturer’s recommendations. The transfection efficiency was routinely 40%. For immunoprecipitation, cells were rinsed with ice-cold PBS, then lysed in ice-cold 20 mM HEPES (pH 7.5), 1% NP-40, 150 mM NaCl, 10% glycerol, 200 ␮M Na3VO5, and protease inhibitors (25 ␮g/ ml of aprotinin and leupeptin). Lysate (400 ␮g) was incubated with 5 ␮g of antibody conjugated to protein A/G-Sepharose (Amersham Pharmacia) for 2 hr at 4⬚C. For immunoblotting, aliquots of total lysates (30 ␮g per sample) or immunoprecipitates were subjected to SDS-PAGE and transferred to nitrocellulose filters which were incubated with appropriate primary and secondary antibodies. The specific signals were visualized by the ECL detection system (Amersham Pharmacia). Modified In-Gel PTP Activity Assay As substrate, we used routinely poly (4:1) Glu-Tyr (Sigma) labeled with [␥-32P]ATP using the GST-FER fusion PTK, as described previously (Shen et al., 1998). The labeled substrates were used within 3 weeks to limit the variation of its specific activity from experiment to experiment. The lysis buffer (25 mM CH3COONa, 1% NP-40, 150 mM NaCl, 10% glycerol [pH 5.5]) was degassed at 4⬚C overnight before catalase and superoxide dismutase (both 100 ␮g/ml), protease inhibitors, and 10 mM iodoacetic acid (IAA) were added. Following stimulation, cells were lysed under anaerobic conditions in an argon chamber. Lysates (25 ␮g) were processed as described in the legend to Figure 1, and an in-gel phosphatase assay (Burridge and Nelson, 1995) was conducted using SDS-PAGE gels containing a radioactively labeled substrate (1.5 ⫻ 106 cpm/20 ml gel solution, approximately 2 ␮M p-Tyr). Measurement of Intracellular ROS Levels Intracellular ROS were measured using 2⬘,7⬘-dichlorofluorescein diacetate (H2DCFDA) and 5-(and-6)-chloromethyl-2⬘,7⬘-dichlorodihydrofluorescein diacetate (CM-H2DCFDA) (Molecular Probes) either by fluorescence microscopy, using a Zeiss Axiovert 405M inverted microscope equipped with a fluorescence attachment and digital camera, or by cell sorting, using a FACSCalibur System (Coulter), according to the manufacturer’s recommendations. Acknowledgments This work was supported by a grant from the National Institutes of Health (R01-GM55989) to N.K.T. We also acknowledge support from the Cold Spring Harbor Laboratory Cancer Center Grant P30CA45508 from the National Institutes of Health. T.F. was supported by the Naito Foundation and a Japan Society for the Promotion of Science Postdoctoral Fellowship for Research Abroad. We are grateful to colleagues from CEPTYR Inc., particularly Drs. Deborah Cool, Larry Kruse, and Ben Neel for helpful discussions. Received August 3, 2001; revised December 19, 2001.

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