R. Karisch, Campbell Family Cancer Research Institute, Ontario Cancer Institute and Princess Margaret Hospital, University Health Network, MaRS Centre, Toronto Medical Discovery Tower, 8th Floor Rm 8-602, 101 College Street, Toronto, Ontario M5G 1L7, Canada Fax: +1 416 581 7698 Tel: +1 416 581 7726 E-mail: email@example.com
Reactive oxygen species (ROS), particularly hydrogen peroxide (H2O2), act as intracellular second messengers in many signaling pathways. Protein-tyrosine phosphatases (PTPs) are now believed to be important targets of ROS. PTPs contain a conserved catalytic cysteine with an unusually low pKa. This property allows PTPs to execute nucleophilic attack on substrate phosphotyrosyl residues, but also renders them highly susceptible to oxidation. Reversible oxidation, which inactivates PTPs, is emerging as an important cellular regulatory mechanism and might contribute to human diseases, including cancer. Given their potential toxicity, it seems likely that ROS generation is highly controlled within cells to restrict oxidation to those PTPs that must be inactivated for signaling to proceed. Thus, identifying ROS-inactivated PTPs could be tantamount to finding the PTP(s) that critically regulate a specific signaling pathway. This article provides an overview of the methods currently available to identify and quantify PTP oxidation and outlines future challenges in redox signaling.
quantitative analysis of the oxidized classical PTPome
quantitative analysis of the classical PTPome
platelet-derived growth factor
oxidized form of PTP1B
4-(azidomethyl)phenyl ethenesulfonate azide
reactive oxygen species
receptor protein-tyrosine phosphatase
selected reaction monitoring
The phosphorylation of proteins on tyrosyl residues is a critical post-translational modification that regulates several fundamental cellular processes, including cell growth, proliferation and migration [1,2]. The levels of phosphotyrosine are regulated by protein-tyrosine kinases  and protein-tyrosine phosphatases (PTPs) [2,3]. Dysregulation of either protein family leads to abnormal levels of phosphorylation and can contribute to several human diseases, including cancer (reviewed in [4,5]).
The PTP superfamily consists of 107 genes, which are divisible into four families . The class I cysteine-based PTPs comprise the largest PTP family and can be subdivided into classical and dual-specificity PTPs . The classical PTPs include 21 receptor and 17 nonreceptor PTPs, which exclusively hydrolyze phosphotyrosyl residues . All classical PTPs contain a highly conserved ‘signature motif’, [I/V]HCSXGXGR[S/T]G, wherein the invariant cysteine is essential for catalysis . In most classical PTPs, this cysteinyl residue has a pKa between 4.5 and 5.5 because of its close contact with several nearby main-chain amide groups and a hydrogen bond with the side chain of the serine residue in the signature motif . This property allows the catalytic cysteine to remain in the thiolate (S−) state at physiological pH, facilitating nucleophilic attack on substrate phosphotyrosines, but it also renders PTPs highly susceptible to oxidation [8–10].
Recent studies suggest that cells capitalize on the exquisite sensitivity of PTPs to oxidation by employing reactive oxygen species (ROS), particularly hydrogen peroxide (H2O2), as intracellular second messengers in many signaling pathways (reviewed in [11–14]). ROS are transiently and locally generated within cells by NADPH oxidases (NOXs) following growth factor stimulation (reviewed in ) and are required for full receptor phosphorylation and activation of downstream signaling (Fig. 1A) [16,17]. Growth factor-induced ROS production leads to the reversible oxidation and inactivation of PTPs to the sulfenic acid (SOH) state (Fig. 1B) . The SOH state is labile and, in different PTP family members, rapidly rearranges to form a sulfenylamide [18–20] with the adjacent main-chain nitrogen or a disulfide bond [21,22] with a nearby cysteinyl residue. The sulfenylamide and disulfide states help to prevent ‘hyperoxidation’ to the biologically irreversible sulfinic (SO2H) and sulfonic (SO3H) acid states (Fig. 1B) [10,18,19]. Several studies have reported the oxidation of specific PTPs in response to different types of cell stimuli, including PTPN1 (PTP1B) in epidermal growth factor receptor signaling , PTPN11 (SHP2) in platelet-derived growth factor (PDGF) signaling , PTP1B and PTPN2 (TC-PTP) in insulin signaling  and PTPN6 (SHP1) in B-cell receptor signaling [26–28] (reviewed in ). In contrast to normal cells, cancer cells often produce high levels of ROS, leading to decreased basal PTP activity and enhanced tyrosyl phosphorylation [29–34]. Readers are directed to a few recent reviews for a detailed discussion of the role of PTP oxidation in pathological cell signaling [35a and Bohmer, Szedlacsek, Tabernero, Ostman & den Hertog, unpublished results].
Given the prominent role of ROS in regulating normal and pathological cell signaling, the identification of ROS-inactivated PTPs might be tantamount to finding the PTP(s) that critically regulate a specific signaling pathway. Several different approaches are available to monitor classical PTP oxidation, each of which exploits the biochemical properties of PTP catalysis or oxidation. This article provides a critical analysis for each of these methods with a particular emphasis on their applicability to a global proteomic approach. We also outline future challenges in improving the identification of redox-regulated PTPs.
Detection of reversible PTP oxidation
Basal or ligand-induced PTP oxidation results in two pools of PTPs: oxidized (SOH; inactive) and reduced (S−; active). Numerous techniques have been developed to identify reversibly oxidized PTPs; these can be classified as ‘indirect’ or ‘direct.’ Indirect methods are the most common and, because they exploit conserved biochemical properties of PTP catalysis, also can be applied to detect PTP expression. Direct approaches rely on detecting the oxidized form of PTPs (or structural changes that arise because of oxidation).
Indirect approaches share a similar experimental workflow, but they can be divided into two general categories based on whether they detect a decrease in active PTPs (S−; ‘negative’) or an increase in oxidized PTPs (SOH; ‘positive’). Both rely on the ability of active PTPs to react stoichiometrically and irreversibly with alkylating agents [PTP-S-Alkyl; i.e. iodoacetic acid (IAA) or N-ethylmaleimide (NEM)] (see Introduction) and the resistance of oxidized PTPs to alkylation. In negative techniques, cells are lysed in the presence of a ‘labeled’ (e.g. radioactive or biotin-tagged IAA) alkylating agent, and PTP oxidation is measured based on decreased detection of that probe (Fig. 2A). Cells also are lysed in the presence of an alkylating agent in positive approaches; however, oxidized PTPs are then converted to the active state (using a reducing agent) and captured and detected using a PTP-reactive probe (e.g. biotin-tagged IAA; Fig. 2B).
Indirect approaches can be ‘targeted’ or ‘global.’ In targeted methods, the PTP of interest is immunoprecipitated; by contrast, with global approaches, oxidation of the entire PTP family can be assessed.
Negative approaches were the first methods developed to detect and quantify PTP oxidation. Lee et al. used radioactively labeled IAA (14C) to monitor oxidation of the nonreceptor PTP PTP1B (Fig. 2A and Table 1) . In their experiments, oxidation was assessed by first lysing cells in the presence of radiolabeled IAA to alkylate and irreversibly label the catalytic cysteinyl residue of active PTPs, whereas oxidized PTPs remained unaffected . PTP1B was then immunoprecipitated, resolved on an SDS/PAGE gel and the radioactivity incorporated (into PTP1B) was quantified . A decrease in incorporation (compared with no treatment) indicated PTP1B oxidation . Using this procedure, Lee et al. demonstrated that PTP1B is oxidized reversibly following H2O2 treatment of recombinant PTP1B and EGF stimulation of A431 cells . The latter result was particularly important, because it provided the first direct evidence that PTPs are physiologically relevant targets of growth factor-evoked ROS, consistent with the earlier suggestion that PTPs must be inhibited by ROS to allow for full receptor tyrosine kinase phosphorylation and activation of downstream signaling [16,17].
Table 1. Comparison of the current methods to detect oxidized PTPs.
Lee et al. also provided a framework to begin to experimentally measure the level of oxidized PTPs following ligand stimulation . But despite its success in identifying PTP1B as a negative regulator of epidermal growth factor receptor signaling, this method is inherently insensitive because it relies on measuring decreases in labeling. Although Lee et al. reported a 40% increase in PTP1B oxidation following EGF stimulation, this is likely an overestimate based on recent measurements of growth factor-induced oxidation (see below). Furthermore, because this is a targeted method, each PTP must be tested individually, provided antibodies are available. Several variations of this method have been used to measure PTP oxidation in different contexts [35,36]. Rather than relying on radiolabeled IAA, these assays used a biotin- or fluorescently-tagged IAA probe [35,36]. Such approaches also are amenable to measuring the oxidation of any reactive cysteine-containing proteins .
Modified in-gel phosphatase (PTPase) assay
The modified in-gel PTPase assay was the first technique used to detect an increase in PTP oxidation (i.e. positive signal) [24,25,38]. It is based on the in-gel PTPase assay, developed in the mid-1990s by Burridge and Nelson to monitor PTP expression. In the in-gel PTPase assay, total cell lysates are resolved on a denaturing SDS/PAGE gel, and proteins are then renatured and reacted with a radioactively labeled PTP substrate, i.e. 32P-labeled poly(GluTyr), that is incorporated into the gel prior to polymerization . PTPs are detected by autoradiography as regions where 32P has been selectively removed (i.e. negative bands) . By applying this approach, Burridge and Nelson detected PTP expression in several cell lines and tissues .
The modified in-gel PTPase assay permits the identification of oxidized PTPs (Fig. 2B and Table 1): lysates are first alkylated with IAA followed by the in-gel PTPase assay, which enables reactivation of formerly oxidized PTPs and their detection based on reactivity with the radiolabeled substrate [24,25,38]. This assay, developed by Meng et al.  was first used to detect increases in PTP oxidation in Rat1 cells following H2O2 stimulation, and also identified PTPN11 as a negative regulator of PDGF signaling. It was subsequently applied to demonstrate reversible oxidation of PTP1B and PTPN2 during insulin signaling . By detecting oxidation of different PTPs in these signaling pathways, Meng et al. added to the growing body of evidence demonstrating that PTPs are critical ROS targets and that different PTPs may be oxidized following activation of different receptor tyrosine kinases.
The development and application of the modified in-gel PTPase assay was an important step forward because it enabled the global, positive identification of PTP oxidation, which could not be assessed previously, and opened the door for the development of more sensitive assays (see below). Yet despite its success, the modified in-gel PTPase assay has several limitations. It is biased toward nontransmembrane PTPs, because most receptor protein-tyrosine phosphatases (RPTPs) do not renature well . Furthermore, although this assay can detect whether a PTP is oxidized, this approach still requires identification of the oxidized PTP by depletion experiments, and because of the differential ability of PTPs to renature, is not quantitative .
Modified cysteinyl-labeling assay
The modified cysteinyl-labeling assay, a clear improvement from the modified in-gel PTPase assay, detects oxidized proteins based on their reactivity with the biotin-tagged alkylating agent, IAP-biotin (Fig. 2B,C and Table 1) , which can react with any reactive cysteinyl residue . In this assay, cells are lysed in the presence of IAA, followed by a desalting column/buffer exchange to remove IAA . Eluates are treated with dithiothreitol (to reduce reversibly oxidized proteins) and subsequently incubated with the IAP-biotin probe to capture reduced (formerly oxidized) proteins . These can then be purified with streptavidin–Sepharose beads and detected by immunoblotting with streptavidin or specific antibodies . To limit the background of nonPTP proteins, the assay can be performed under mildly acidic conditions (pH 5.5), which ensures that the majority of cysteine-containing proteins are in the thiol (SH) state and cannot react with the probe . At this pH, the catalytic cysteine of most PTPs (including classical PTPs and dual-specificity PTPs) along with other highly reactive cysteine-containing proteins, such as hydroxylases, peroxidases and thiol proteases, remains in the active thiolate (S−) state and are reactive .
The concept underlying the modified cysteinyl-labeling assay was conceived in the late-1990s to assess the oxidation of nonPTP proteins, such as p53 . PTPs were first assessed by Li and Whorton , who used a 3-(N-maleimido-propionnyl)biocytin (NEM)-biotin (MPB) probe, to measure the nitrosylation of PTP1B in response to the treatment of A431 and Jurkat cells with various nitrosylation agents (i.e. S-nitroso-N-acetylpenicillamine and S-nitrosoglutathione) . Boivin et al.  developed the modified cysteinyl-labeling assay to detect increased oxidation of a number of proteins in PDGF-transformed angiomyolipoma cells (compared with nontransformed cells), which was prevented when the cells were treated with the NOX inhibitor diphenyliodonium or the antioxidant N-acetyl cysteine. Boivin et al. also demonstrated that this assay could detect oxidation of PTP1B and the RPTPs PTPRF (LAR) and PTPRA (RPTPα), along with the lipid phosphatase PTEN and the MAPK phosphatase MKP-1 in the same cells . Subsequently, the authors published several papers to outline the method to the redox signaling field [43,44], which will facilitate its adaptation to a global MS-based approach. In fact, a modified IAP-biotin probe was recently used to identify reactive cysteinyl residues by MS. Although > 1000 cysteine-containing peptides were identified, only three were PTP-derived, and these did not include active site peptides . Notably, these assays were performed at physiological pH (7.5), which as previously discussed, would increase the background of abundant (nonPTP) peptides [37,40]. By performing these experiments under mildly acidic conditions, it is possible that this approach could assess PTP expression (and oxidation) by MS, along with other highly reactive cysteinyl-containing proteins. Without using MS, however, this assay is limited by its inability to identify the oxidized proteins. Furthermore, in its current format, unlike the modified in-gel PTPase assay, where only PTPs are measured, a large panel of nonPTP proteins would be detected. The identification of other proteins maybe beneficial depending on the intended experimental aim (i.e. assessment of all redox-regulated proteins or exclusively classical PTPs). However, probe reactivity with nonPTP proteins might reduce the assay’s sensitivity and result in decreased detection of classical PTPs, as was evident in the aforementioned study .
Although global proteomic applications of the modified cysteinyl-labeling assay are currently unavailable to assess PTP oxidation, targeted MS approaches have been reported. For example, Lou et al.  developed a MS-based technique whereby all PTP oxidation states can be measured. In their method, cell lysates prepared in the presence of iodoacetamide are applied to a gel-filtration column, and the eluate is then reduced. PTPs of interest are immunoprecipitated, resolved on an SDS/PAGE gel and stained with Coomassie brilliant blue . The region corresponding to the PTP is excised and subjected to in-gel trypsinization . Tryptic peptides are then detected by MALDI-MS, permitting the identification (but not the quantification) of all PTP oxidation states (i.e. S−, SOH, SO2H, SO3H) . The authors applied this procedure to show that PTP1B is reversibly (SOH) and irreversibly (SO2H and SO3H) oxidized in several cancer cells . For this PTP, quantification was possible because its S− and SOH forms migrate differently on SDS/PAGE compared with the SO2H and SO3H forms . These experiments demonstrated that > 25% of PTP1B can be reversibly oxidized basally in cancer cells . This study provided a seminal contribution to the field by providing the first quantification of all PTP oxidation states; however, it is unlikely to be applicable to a global proteomic approach since the oxidized forms of most, in not all PTPs are unlikely to migrate differently.
Subsequently, Held et al. developed a targeted proteomic approach that relies on selected reaction monitoring (SRM, also known as multiple reaction monitoring) to quantify protein oxidation . SRM is a label-free, tandem MS method that allows the relative abundance of characterized peptide ions to be quantified (reviewed in [46,47]). In their assay, which they term oxidized SRM, cells are lysed in the presence of trichloroacetic acid to quench all redox reactions . Proteins are then resuspended in a denaturing buffer containing unlabeled (d0) NEM to alkylate free cysteinyl residues, precipitated with trichloroacetic acid and resuspended in denaturing buffer containing Tris(2-carboxyethyl)phosphine and a stable isotopic version of NEM with five deuteriums (d5) to label formerly oxidized proteins . The protein of interest is then immunoprecipitated, digested with trypsin and the peptides corresponding to the alkylated d5 NEM (oxidized) and d0 NEM (reduced) cysteines, which differ by 5 Da are then monitored by SRM to determine their ratio (oxidized/reduced [d5/d0]) . Similar to the modified in-gel PTPase assay, this method is not PTP specific and can be applied to monitor the oxidation of any protein, provided antibodies are available . The authors applied this method to demonstrate that PTP1B is oxidized following H2O2 and diamide (a ROS-producing agent) stimulation of MCF7 cells and that p53 is oxidized following diamide treatment .
The oxidized SRM assay provides a reliable general procedure to assess protein oxidation by SRM . By using SRM for quantification, the authors can monitor the oxidation of any reactive cysteinyl residue provided the peptide can be initially detected by LC-MS/MS . One of the limitations of the method, as the authors discuss, is the overestimation of the fraction of reversibly oxidized proteins if a high proportion are in the SO2H or SO3H state. However, this is a limitation shared by most approaches to assess oxidation . Overall, this is a promising technique to monitor oxidation, particularly since the fraction of each protein oxidized can be quantified using different isotopes of NEM for alkylation.
Activity-based PTP probes
Activity-based probes (ABPs) are reagents designed to react with mechanistically related enzymes [48,49]. ABPs have been developed to target several protein families, including caspases , papains , glycosidases  and hydrolases . By exploiting the properties of PTP catalysis, Zhang and coworkers have developed ABPs targeting classical PTPs (Fig. 3A). All classical PTPs contain two residues that are critical for the catalytic mechanism: the catalytic cysteine and an aspartic acid (D) residue in the WPD loop [6–8]. Phosphotyrosine substrates bind to, and are stabilized in, the nonpolar PTP catalytic cleft. The WPD loop then undergoes a dramatic conformational change, closing over the phenol ring of the phosphotyrosine, thereby holding the substrate in place and positioning it for dephosphorylation [6–8]. The catalytic thiolate then executes a nucleophilic attack on the phosphate moiety of the phosphotyrosine substrate [6–8]. The aspartic acid residue acts as the general acid, donating a proton (H+) to the tyrosine, facilitating release of the dephosphorylated substrate and formation of a PTP–S–PO42− intermediate [6–8]. The same aspartic acid residue then acts as a general base, removing a proton from water, and promotes nucleophilic attack by the resulting hydroxyl ion (OH−) on the PTP–S–PO42− intermediate, releasing the reactivated PTP and PO42− [6–8].
Zhang et al. developed ABPs targeting the classical PTP family by synthesizing molecules that: (a) resemble phosphotyrosine, allowing ABP binding to the PTP catalytic cleft; (b) contain a reactive group, permitting the reaction and subsequent trapping of the PTP; and (c) have a linker connecting the ABP to a reporter/affinity tag, enabling purification of the PTP–ABP adduct [54–56]. Two of these ABPs are α-bromobenzylphosphonate (BBP)-biotin (Fig. 2C)  and 4-(azidomethyl)phenyl ethenesulfonate azide (PVSN-N3; Fig. 2C) . Both contain a phenyl ring to promote binding to the PTP catalytic cleft and initiation of a normal substrate reaction. Each, however, traps PTPs differently (Fig. 3B,C) [54,55]. The phosphonate moiety of BBP-biotin reacts with the PTP catalytic cysteine, generating a transient phosphorane-like intermediate, which leads to the formation and subsequent opening of a three-membered epoxide ring, release of bromide, and rearrangement to yield a PTP–S–BBP-biotin adduct (Fig. 3B) . PVSN-N3 reacts with the catalytic cysteine via a 1,4-conjugate addition, forming a stable thioether bond (PTP–S–PVSN-N3; Fig. 3C) . In experiments using BBP-biotin, PTPs are isolated by binding to streptavidin–Sepharose , whereas PVSN-N3 adducts must first undergo a click chemistry reaction with an alkyne-conjugated reporter/affinity tag (Fig. 3D) . Click chemistry is a modular reaction that permits the reaction of two stable and otherwise inert reactants (i.e. azide and alkyne) in an efficient and irreversible reaction [57,58], most often Cu-catalyzed azide–alkyne cycloaddition [59,60] or Staudinger ligation . PVSN-N3 lacks the bulky biotin group associated with BBP-biotin, which might increase PVSN-N3 reactivity with PTPs by avoiding steric hindrance by the biotin moiety. Furthermore, PVSN-N3 can readily cross cell membranes in cells and react with PTPs, unlike BBP-biotin, which is too bulky . Zhang et al. demonstrated that both of these probes can react with PTPs, first using the Yersinia PTP, YopH and then the human PTPs: PTP1B, PTPN11 and PTPRJ (DEP1) [54,55]. These ABPs also react with dual-specificity PTPs (i.e. Cdc14, VHR, PRL-3) and the low molecular mass protein-tyrosine phosphatase [54,55].
Boivin et al. demonstrated that BBP-biotin can be used to monitor PTP oxidation . Briefly, cells are lysed in the presence of IAA, applied to a gel-filtration column, reduced and then reacted with the probe, allowing purification of formerly oxidized PTPs (Fig. 2B and Table 1) . BBP-biotin detected oxidation of several PTPs in PDGF-transformed angiomyolipoma cells, including PTP1B, PTPRF and PTPRA . PVSN-N3 has not yet been applied to detect PTP oxidation; however, based on its ability to detect PTP expression, this should be possible. Furthermore, it might have improved reactivity compared with BBP-biotin, because of the absence of the bulky biotin group (see above).
A benefit of using ABPs to monitor PTP oxidation is the reduction of background (nonPTP) proteins because of the mechanism-based specificity of these probes [54,55]. As yet, neither of these ABPs has been applied to MS experiments. Furthermore, a complicated and lengthy series of reactions are needed to generate BBP-biotin, which might restrict its use by the general redox signaling field . By contrast, PVSN-N3 requires only a few synthetic steps, making it an ideal candidate to globally detect PTP expression and oxidation .
Oxidized PTP active site antibody (oxPTP Ab)
The Ostman group employed an antibody-based strategy to detect oxidation of classical PTPs by immunizing rabbits with a peptide comprising a portion of the signature motif of PTP1B hyperoxidized to the SO3H state (VHCSO3HSAG) [62,63]. The high conservation of the signature motif ([I/V]HCSXGXGR[S/T]G; see Introduction) allows this antibody to cross-react with other classical PTPs (see below). Persson et al. used this antibody in targeted experiments to show reactivity with several classical PTPs, including PTP1B, PTPN2, PTPN6, PTPN11, PTPRA and PTPRJ [62,64]. Using an approach analogous to the modified cysteinyl-labeling assay, the oxPTP Ab also could be used to measure PTP oxidation (Fig. 2B). In this procedure, cells are lysed in the presence of NEM, and the PTP of interest is immunoprecipitated, reduced, washed and then hyperoxidized to the SO3H state using pervanadate (PV). The wash step between the reduction and oxidation reaction is required to prevent the conversion of PV to vanadate (VO43−), a reversible PTP inhibitor. Using this targeted protocol, Persson et al. showed that PTPN11 is oxidized following PDGF stimulation , the D2 domain of PTPRA is highly sensitive to oxidation [62,64], and that PTPN6 and PTPN11 are susceptible to H2O2-induced oxidation .
Our group modified the antibody-based approach to enable global detection of classical PTP expression and oxidation. To measure PTP expression, we applied dithiothreitol-treated lysates to a gel filtration column and then treated them with PV (Fig. 4) . Combined with a newly available monoclonal oxPTP Ab, we used this method, termed qPTPome, to detect several PTPs in PV-treated lysates following immunoprecipitation and immunoblotting with the oxPTP Ab. Efficient PTP purification can be achieved only if lysates are denatured prior to immunopurification, suggesting that the oxPTP Ab cannot bind PTPs in their native PTP–SO3H conformation. We further modified this assay to an MS-based approach by generating tryptic digests of PV-treated lysates, immunoprecipitating PTP–SO3H active site peptides with the oxPTP Ab, and subsequently analyzing the peptides by LC-MS/MS. These modifications allowed us to identify 17 PTPs in NIH 3T3 cells, including the D1 and D2 domains for several RPTPs. In subsequent experiments, we demonstrated that the oxPTP Ab can isolate PTPs from mouse, human or rat cells provided that their active sites match the consensus sequence [I/V]HCS[A/S]G, permitting the detection of all catalytically active classical PTPs (36 of 38 classical PTPs). Notably, the oxPTP Ab can not react with other phosphatases, such as dual-specificity PTPs or low molecular mass protein-tyrosine phosphatase.
We then developed a multiplexed SRM protocol to enable, in a single MS assay, comprehensive quantification of PTP expression (and oxidation) of mouse and human classical PTPs using qPTPome-processed samples (Fig. 4). Applying this method, we demonstrated that both relative and absolute (when combined with absolute quantification) levels of PTP expression can be measured.
To measure PTP oxidation, we developed q-oxPTPome, wherein cells are lysed in the presence of NEM, applied to a gel filtration column, reduced, applied to a second gel filtration column and then treated with PV (Fig. 4 and Table 1). PV-treated lysates are treated with trypsin, and PTP–SO3H peptides are immunopurified by the oxPTP Ab and quantified by multiplexed SRM. By combining q-oxPTPome with qPTPome, the fraction of each PTP oxidized (q-oxPTPome/qPTPome) can be determined. Because the oxPTP Ab detects all RPTP D1 domains and most D2 domains, this approach can be used to monitor the differential oxidation of these two domains. Furthermore, modifications to the procedure allow quantification of all PTP oxidation states (i.e. S−, SOH, SO2H, SO3H). By applying this approach, we showed that q-oxPTPome could monitor differential changes in PTP oxidation: (a) following H2O2 stimulation of NIH 3T3 cells, (b) following the alteration of the intracellular redox environment by depleting glutathione, (c) in several different cancer cell lines, and (d) following inhibition of a driver oncogene. Taken together, these data suggested that PTP oxidation adds an additional layer of complexity to the cancer phenotype.
Although q-oxPTPome can quantify the above changes in PTP oxidation, we have been unable to identify the subset of PTPs oxidized in response to growth factor stimulation of normal cells. This likely reflects the complex procedure involved in q-oxPTPome and the small and localized subset of PTPs that are inactivated by normal levels of growth-factor-evoked ROS. Because the assay is quantitative, it provides an upper bound on growth factor-induced increases in PTP oxidation (< 5%). Studies are underway in our laboratory to increase the sensitivity of q-oxPTPome to circumvent this limitation. Without these improvements, the assay will be unable to begin to interrogate the critical questions confronting the redox signaling field. Furthermore, owing to the divergence of some of the PTP active site sequences, the D2 domains of a few RPTPs (Ptprc, Ptprk, Ptprm, Ptprt and Ptpru) escape detection by the oxPTP Ab. However, the oxPTP Ab can detect their D1 domains, allowing quantification of their expression (and oxidation).
Overall, qPTPome and q-oxPTPome provide a general workflow to begin to study PTP signaling at a systems level. For example, qPTPome can be combined with phosphotyrosine proteomics to suggest PTP substrates. A similar protocol could be used in other systems to identify key PTP substrates. It might also be possible to combine q-oxPTPome with phosphotyrosine proteomics to identify PTP substrates following stimulation with different levels of H2O2, by correlating the levels of PTP oxidation with increases in the phosphorylation of specific proteins. Modifications of qPTPome might be developed to facilitate the simultaneous identification of all PTP post-translational modifications: classical PTPs could be isolated at the protein level using the oxPTP Ab, followed by secondary enrichment of trypsinized peptides based on a specific post-translational modification [i.e. phosphotyrosine antibodies (i.e. pY-100) to detect tyrosyl phosphorylation] and MS analysis. Such approaches could help to bridge the gap in our understanding of PTP and protein-tyrosine kinase signaling.
Summary of indirect approaches
Indirect approaches to assess PTP expression and oxidation have improved since the development of the radiolabeled assay by Lee et al. . Although there currently is no ‘gold standard’ assay, the redox-signaling field is rapidly moving towards the development of more robust methods to monitor PTP oxidation. Indirect approaches provide the unique opportunity to probe both PTP expression and oxidation by exploiting the properties of PTP catalysis. Because these approaches can monitor the entire classical PTP family, it will soon be possible for the signaling field to begin to study the global involvement of PTPs in signaling, rather than its current targeted format. Despite the potential of each of these assays, particularly the modified cysteinyl-labeling assay and qPTPome/q-oxPTPome, additional improvements are required to begin to interrogate the role of PTPs in regulating growth factor signaling at a systems level.
Unlike indirect approaches, which rely on the reactivity of active PTPs, direct methods monitor PTP oxidation by exploiting the properties of oxidized PTPs. Currently, there are two methods to directly assess PTP oxidation. The first relies on dimedone (5,5-dimethyl-1,3-cyclohexanedione), a cell-permeable small molecule that selectively and irreversibly reacts with sulfenic acids (SOH; Fig. 5A). Because dimedone reacts with the oxidized thiol, it targets the entire redoxome (not just PTPs), in principle, making it an ideal approach to directly and globally assess protein oxidation. The second approach relies on conformation-sensing antibodies that recognize structural modifications that occur in oxidized PTPs (compared with the active enzyme) that can trap, purify and inhibit PTP reactivation.
Dimedone-based probes provided the first direct method used to assess protein oxidation. Their reactivity is based on the 1,3-cyclohexadione backbone, which undergoes keto–enol tautomerism and, in the enol state, can react with the electrophilic sulfur atom in protein–sulfenic acids, releasing water and forming a thioether (C–S–C) bond with the oxidized protein (Fig. 5A). Dimedone was used to measure protein oxidation as early as the late 1960s, when it was shown to react with oxidized papain  and glyceraldehyde-3-phosphate dehydrogenase . However, its application was limited to targeted approaches because of the absence of a reporter/affinity tag.
Poole et al. synthesized several dimedone-based probes consisting of the dimedone reactive group, a linker and a reporter/affinity tag or an azide group (Fig. 5B and Table 1) [69–74]. Direct conjugation of dimedone to a fluorophore or biotin tag allows the detection of oxidized proteins using a single reagent, but, as for ABPs, might limit reactivity with proteins because of steric constraints [69–74]. By contrast, dimedone probes with an azide moiety require click chemistry to connect the probe to an alkyne-linked reporter/affinity tag [69–74]. Poole et al. used AhpC, a cysteine-based peroxidase in Salmonella typhimurium, to test these probes [69–71], showing by MS that dimedone selectively labels oxidized AhpC, but not active AhpC or a Cys46Ser mutant [75,76]. Both the direct and ‘clickable’ probes also could react with oxidized AhpC using a targeted in vitro approach [69–71]. Subsequently, the same group demonstrated that these probes could detect an overall increase in biotin–dimedone (Dcp–Bio1; Fig. 5B) labeling following T-cell activation and that both PTPN6 and PTPN11 were oxidized reversibly . This probe was also used to detect PTP1B and PTPRJ (DEP1) oxidation following vascular endothelial growth factor stimulation of human umbilical vein endothelial cells, demonstrating that Dcp–Bio1 can react with both nonreceptor and receptor PTPs . Recently, Carrroll and colleagues also demonstrated that Dyn-2, a click chemistry-based dimedone probe could detect oxidation of PTP1B, PTPN11 and PTEN in response to EGF stimulation of A431 cells .
Dimedone probes were first applied to measure protein oxidation globally by Charles et al., who compared H2O2-stimulated and control rat ventricular mycocyte lysates treated with biotin–dimedone . Oxidized proteins were purified on streptavidin beads, and resolved by SDS/PAGE, subjected to in-gel trypsinization (in regions that showed increased labeling after H2O2 stimulation), followed by LC-MS/MS . Twenty-two proteins were oxidized inducibly; however, these corresponded to highly abundant proteins, such as myosin or ATP synthase . It is possible that the bulky nature of the biotin moiety prevented the probes from reacting with signaling proteins, such as PTPs, whose oxidized cysteines are located within sterically restrictive ‘pockets’ or ‘clefts.’
Using a click chemistry-based dimedone probe (DAz1; Fig. 5B), Leonard et al. identified 189 reversibly oxidized proteins in HeLa cells using a similar MS-based approach. Unlike the previous study, however, several signaling proteins were identified suggesting that these probes have increased reactivity . Despite this improvement, however, PTPs were not identified. A recent publication by Leonard et al. suggested that previous dimedone-based probes have limited reactivity with classical PTPs . To circumvent this limitation, the authors synthesized several dimedone-based probes containing a ‘PTP-binding module,’ such as a phenyl group, which increased probe binding within the PTP catalytic cleft and consequently increased its reactivity (Fig. 5B) . These probes showed significantly increased reactivity with oxidized YopH, a Yersinia PTP, compared with previous dimedone probes . Although these probes have yet to be applied to monitor PTP oxidation, they are promising candidates to apply to develop a global proteomic approach.
Dimedone-specific antibodies also are available to directly monitor protein oxidation (Table 1). To generate these antibodies, Seo et al. synthesized a keynote limpet hemocyanin-conjugated dimedone hapten by joining the dimedone moiety with cysteamine via a thioether bond followed by a five-carbon linker . Keynote limpet hemocyanin-conjugated dimedone was injected into rabbits to produce polyclonal dimedone-specific antibodies . In principle, these antibodies could be used to detect oxidized proteins by treating cells or lysates with dimedone, followed by immunoprecipitation and/or immunoblotting . Indeed, in a targeted approach, dimedone-specific antibodies reacted with oxidized, but not reduced glyceraldehyde-3-phosphate dehydrogenase, peroxiredoxin I and actin . Oxidized proteins also could be detected in cell lysates by immunoblotting or in cells using immunofluorescence . Accordingly, the authors applied these dimedone-specific antibodies to measure the differential protein oxidation in a panel of breast cancer cell lines by treating the cells with dimedone for 2 h, followed by cell lysis and immunoblotting . Cells that were not treated with dimedone showed very low reactivity with the antibody, in contrast to dimedone-labeled cells, from which several immunoreactive bands were detected . The intensity and pattern of bands detected across the panel of breast cancer lines differed, suggesting each breast cancer line has a unique profile of oxidized proteins .
One limitation of the approach used by Seo et al. is that oxidation was assessed after 2 h of dimedone treatment . Because dimedone irreversibly reacts with sulfenic acids, incubating cells with dimedone for such an extended period, probably alters the intracellular redox state, leading to increases of protein oxidation. It would be best to treat cells with dimedone for as short a time as possible or to treat lysates rather than cells with dimedone to limit the alteration of redox homeostasis. The later approach might lead to postlysis oxidation, though, making it crucial that the experiment is performed under anaerobic conditions.
A second dimedone-specific antibody was developed by Maller et al., who used keynote limpet hemocyanin that was reacted directly with dimedone as the immunogen . Unfortunately, these antibodies only detected oxidized glyceraldehyde-3-phosphate dehydrogenase in H2O2-stimulated rat myocytes, with very few additional immunoreactive bands, unlike the previous antibodies . The inability of this antibody to detect several immunoreactive bands will limit its application.
Although the antibody developed by Seo et al. appears to specifically react with a large panel of reversibly oxidized proteins, it is unclear whether it can detect growth factor-induced changes in protein oxidation. Furthermore, it will be essential to identify these dimedone-immunoreactive bands by MS. Given dimedone’s low reactivity with PTPs, it may be difficult to apply these antibodies to monitor PTP oxidation; however, this will be unclear until targeted approaches are used to directly assess PTP oxidation or it is applied to MS. Seo et al. also recently developed an iododimedone (d0) derivative (Fig. 5B) that can react with any reactive cysteinyl-containing protein, permitting the quantification of the fraction of a protein oxidized when combined with a stable isotope version of dimedone (d6; d6/do) . It will be interesting to see whether this can be combined with the dimedone-specific antibodies to develop a quantitative global proteomic approach to assess protein oxidation.
Conformation-sensing antibodies of oxidized PTP1B (PTP1B-OX)
Recently, the Tonks group developed conformation-sensing antibodies targeting the oxidized form of PTP1B (PTP1B-OX; Table 1) . Earlier crystallographic studies showed that in the presence of ROS, PTP1B is oxidized to the sulfenic acid (SOH) state and rapidly rearranges to form a sulfenylamide (see Introduction) [18,19]. Sulfenylamide formation dramatically alters the PTP1B active site, exposing Tyr46 [18,19]. Haque et al. identified a mutant of PTP1B (CA/SA) that adopts a similar conformation to PTP1B-OX . The authors then employed phage display combined with subtractive panning to develop antibodies that specifically detected PTP1B-CA/SA . The resulting antibodies also reacted with PTP1B-OX in vitro and could prevent its reactivation in the prescence of the reducing agent Tris(2-carboxyethyl)phosphine . Remarkably, this antibody did not bind to closely related PTPN2, which shares ∼ 50% identity with PTP1B . When expressed in cells as an ‘intrabody,’ it could be used to immunoprecipitate and image PTP1B-OX in response to H2O2 or insulin stimulation, demonstrating for the first time the existence of the sulfenylamide state in vivo . Using immunofluoerescence, the authors reported that ∼ 40% of PTP1B colocalized with PTP1B-OX (suggesting that upto 40% of PTP1B might be oxidized) following insulin stimulation, although, based on immunoprecipitation experiments, it appeared that only 5–10% was oxidized. This discrepancy might reflect the limited spatial resolution of the immunofluorescence measurements. The immunoprecipitation data probably represents the ‘true’ level of PTP1B oxidation. This might be an overestimate of the fraction ‘instantaneously’ oxidized, because the conformation-specific antibodies bind and prevent reactivation of PTP1B. Consequently, PTP1B-OX accumulates over time, leading to an ‘integral’ measurement of PTP1B-OX, rather than the instantaneous value measured by other methods. These data suggest that the fraction of PTP1B oxidized following insulin stimulation is < 5%, demonstrating the importance of localized PTP oxidation in growth factor signal transduction. It will be interesting to apply these antibodies to monitor PTP1B-OX in other signaling pathways.
Summary of direct approaches
Direct approaches to monitor PTP oxidation provide a unique opportunity to assess oxidation because they omit the complicated series of steps required by indirect methods. Although these steps are generally considered to go to completion, each step probably results in some losses that lead to an overall decrease in assay sensitivity. In principle then, direct approaches are preferable for quantifying PTP oxidation, but it has been difficult to develop direct methods that monitor oxidation of the entire classical PTP family. Dimedone-based approaches provide a promising technique to directly and globally measure PTP oxidation. Recent improvements, such as the PTP-specific probes developed by Seo et al.  might allow this approach to be applied to the entire PTP family.
Summary and future perspectives
Oxidation has emerged as an important mechanism for regulating PTP activity in both normal and oncogenic cell signaling. ROS levels are increased following growth factor stimulation and in several diseases, including cancer. Elevated levels of ROS lead to an increased fraction of oxidized PTPs, shifting the dynamic equilibrium between PTPs and protein-tyrosine kinases in favor of increased tyrosyl phosphorylation and downstream signaling. Detecting ROS-inactivated PTPs should help to identify PTP(s) that regulate specific signaling pathways and to delineate the physiological roles of redox regulation. Ideally identification of these oxidized PTPs would involve the application of global approaches that provide a systems-level understanding of PTP signaling.
We have outlined several currently available methods to assess PTP expression and oxidation. Initially, these techniques were exclusively targeted methods, which could only monitor oxidation of a specific PTP at any given time. However, our increased understanding of PTP catalysis and oxidation, combined with improvements to current proteomic technologies, now provide the framework for beginning to monitor PTPs at a systems level. Despite the success of recent approaches, improvements to these methods are required to routinely quantify changes in PTP oxidation. These changes will not only allow the quantification of PTP oxidation and expression, but will likely provide the opportunity to interrogate other PTP regulatory mechanisms, such as tyrosyl phosphorylation. Only time will tell if new innovations can help to bridge the gap in our understanding between PTP and protein-tyrosine kinase signaling.
Work in the Neel lab is supported by National Institutes of Health Grant R37CA49152 (BGN) and the International Human Frontiers Science Program Organization Grant RGP0039/2009C102 (BGN). Additional support was provided by the Ontario Ministry of Health and Long Term Care and the Princess Margaret Hospital Foundation. BGN is a Canada Research Chair, Tier 1. RK is the recipient of a graduate fellowship from the Canadian Institutes of Health Research.