Protein Tyrosine Phosphatase regulation by reactive oxygen species

Abstract

Protein Tyrosine Phosphatases (PTPs) help to maintain the balance of protein phosphorylation signals that drive cell division, proliferation, and differentiation. These enzymes are also well-suited to redox-dependent signaling and oxidative stress response due to their cysteine-based catalytic mechanism, which requires a deprotonated thiol group at the active site. This review focuses on PTP structural characteristics, active site chemical properties, and vulnerability to change by reactive oxygen species (ROS). PTPs can be oxidized and inactivated by H₂O₂ through three non-exclusive mechanisms. These pathways are dependent on the coordinated actions of other H₂O₂-sensitive proteins, such as peroxidases like Peroxiredoxins (Prx) and Thioredoxins (Trx). PTPs undergo reversible oxidation by converting their active site cysteine from thiol to sulfenic acid. This sulfenic acid can then react with adjacent cysteines to form disulfide bonds or with nearby amides to form sulfenyl-amide linkages. Further oxidation of the sulfenic acid form to the sulfonic or sulfinic acid forms causes irreversible deactivation. Understanding the structural changes involved in both reversible and irreversible PTP oxidation can help with their chemical manipulation for therapeutic intervention. Nonetheless, more information remains unidentified than is presently known about the precise dynamics of proteins participating in oxidation events, as well as the specific oxidation states that can be targeted for PTPs. This review summarizes current information on PTP-specific oxidation patterns and explains how ROS-mediated signal transmission interacts with phosphorylation-based signaling machinery controlled by growth factor receptors and PTPs.

Introduction

Cellular signaling events during proliferation, adhesion, and migration are initiated and transmitted through the phosphorylation of protein tyrosines (Hunter, 1995). Two families of signaling enzymes—Protein tyrosine Kinases (PTKs), which phosphorylate tyrosine residues, and Protein Tyrosine Phosphatases (PTPs), which reverse them—balance this reversible post-translational modification of target proteins (Lalima G. Ahuja, 2018; Almo et al., 2007; M. J. Chen, Dixon, & Manning, 2017; Manning, Whyte, Martinez, Hunter, & Sudarsanam, 2002). Approximately 95 PTKs are counteracted by 107 PTPs such that mutations in both are linked to many human diseases including cancers(Jannik N. Andersen et al., 2004; Cohen, 2001). While PTPs were frequently dismissed as housekeeping enzymes that simply counteract the action of PTKs, it is now recognized that PTPs can have both negative and positive regulatory impacts on the signaling cascade(C. L. Welsh, Allen, & Madan, 2023; Colin L. Welsh, Pandey, & Ahuja, 2021). PTPs have recently regained reach as pharmacological targets for a variety of malignancies, thanks to increased patient data availability and clever designing of allosteric inhibitors(C. L. Welsh et al., 2023; Z.-Y. Zhang, 2017). These efforts are also supported by recent developments in mass spectrometry and peptide chemistry, as well as the convenience of working with purified PTP catalytic domains. PTP regulation occurs at the level of gene expression, through protein–protein interactions, allosteric intradomain interactions, and/or through phosphorylation of their serine/threonine or tyrosine residues(Lalima G. Ahuja, 2018; L. G. Ahuja & Gopal, 2014; Colin L. Welsh et al., 2021). In recent years, oxidation–reduction events, also known as redox reactions, have emerged as a significant mechanism for the regulation of cellular PTP activity by chemical agents(Boivin & Tonks, 2015; Hay et al., 2020; Londhe et al., 2020; Ostman, Frijhoff, Sandin, & Böhmer, 2011; Tonks, 2005; Tonks, 2006, 2013; Tsutsumi et al., 2017). These reactions are centered on the catalytic cysteine residue of these enzymes. This review briefly covers details on PTP oxidation and its regulation in response to oxidative stress and is intended to update and complement other reviews that have been published on this subject in the past(Boivin & Tonks, 2015; Chiarugi & Cirri, 2003; L. E. S. Netto & Machado, 2022; Tanner, Parsons, Cummings, Zhou, & Gates, 2011).

The Protein Tyrosine Phosphatase (PTP) catalytic domain

PTPs are the largest protein phosphatase enzyme superfamily and are mechanistically split into four types(Lalima G. Ahuja, 2018; M. J. Chen et al., 2017). Classes I-III contain cysteine-based PTPs with a nucleophilic cysteine in the signature active site HC-X_5-R motif (Z.-Y. Zhang et al., 1994). The evolutionarily variable Class IV PTPs Eya (Eyes Absent) are related to Haloacid Dehalogenases (HAD) and use aspartate instead of cysteine at the enzyme active site(Burroughs, Allen, Dunaway-Mariano, & Aravind, 2006). Cysteine-based Class I has classical PTP domains and dual-specificity DUSPs. The domain architecture of class I PTPs is highly conserved and these are strictly selective for pTyr substrates while DUSPs have wider substrate specificity. About 37 classical PTP genes in humans code for 35 active PTP enzymes (Jannik N. Andersen et al., 2001). Furthermore, based on their cell location, traditional PTPs can be split into two groups: membrane-anchored receptor protein tyrosine phosphatases (RPTPs) and intracellular PTPs (Figure 1A).

Figure 1: The PTP Superfamily of Enzymes:

A. Illustration of human PTPs with membrane-bound (receptor) and cytosolic (non-receptor) subtypes. B. (right) Domain architecture of the conserved PTP domain, indicating the positions of ten sequence motifs that are conserved across the superfamily. (left) An alternative view of the domain showing the active site’s position between numerous mobile loops. PTP1B crystal structures have been used for both the illustrations; open structure PDB ID: 5K9V, closed structure PDB ID: 1PTV.

PTP1B, the first PTP isolated from the human placenta and examined for its molecular features by protein crystallization, exemplifies the conserved PTP catalytic domain(Barford, Flint, & Tonks, 1994; Tonks, 2003; Tonks, Diltz, & Fischer, 1988). Figure 1B shows that the catalytic domain’s core is made up of twisted β-sheets surrounded by α-helices. Andersen et. al, have defined ten motifs that define the sequence conservation on the defined domain architecture (Figure 1B)(Jannik N. Andersen et al., 2001). Of these, four motifs include the PTP active site loops while the other six motifs maintain the domain’s structural integrity. The Cα-regiovariation score analysis reveals that the most conserved motif is the structural motif-4 (F/Y)IAxQGP, which forms the hydrophobic core around the PTP loop. Motif-4 and motif-3 DYINA(N/S) form a parallel-antiparallel β-sheet at the center of the PTP domain. The PTP domain’s hydrophobic cluster also comprises residues from motif-2 DxxR(V/I)xL, motif-5 TxxDFWx(M/L/V)x(W)(E/Q), motif-6 (I/L/V) (V/I)MxT, and motif-7 KCxxYWP. The aromatic side chains of residues from motif 5 and motif 7 (Phe95, Trp96, Tyr124, and Trp125 in PTP1B) help stabilize the core of the PTP domain through π-stacking interactions. Because a protein’s hydrophobic core is responsible for its thermostability, mutations in these motifs has a significant impact on the PTP’s folding properties(Muise, Vrielink, Ennis, Lemieux, & Tremblay, 1996).

The PTP active site is encased within numerous loops that contain catalytic residues in four conserved sequence motifs (Figure 1B). The pTyr-loop, also known as the pTyr-recognition loop or substrate binding loop, is responsible for confining PTP’s substrate specificity to phosphorylated tyrosines. This loop is a part of motif-1 Nxx(K/R)NRY, which guards the catalytic site and defines the depth of the active site crevice (L. L. Madan & Gopal, 2011). The motif has an 84% conserved tyrosine/phenylalanine that works as a causeway to the active site binding pocket, giving it a depth of ~9 Å. As a result, only a pTyr can access the binding pocket’s bottom. Furthermore, this tyrosine/phenylalanine aids in hydrophobic packing of the phosphorylated substrate by making π-π stacking interactions with the substrate’s phosphotyrosine residue, facilitating substrate binding into the binding site(L. L. Madan & Gopal, 2011). An asparagine (or aspartate) two residues downstream from the pTyr-loop creates a bipartite hydrogen bond with the substrate, stabilizing it in the active site(Peti & Page, 2015). The WPD loop is a part of motif-8 ((Y/F)xxWPDxGxP) and includes the general acid/base aspartate required for catalysis. PTPs with variations at the catalytic aspartate (as seen in the second PTP domains of bidomain RPTPs) are either inactive or have extremely poor catalytic activity (L. G. Ahuja & Gopal, 2014; L. L. Madan, Goutam, & Gopal, 2012; Lalima L. Madan et al., 2011; Colin L. Welsh et al., 2021). Mutations in the general acid-base aspartate form a clever method of trapping substrates at the active site (Blanchetot, Chagnon, Dubé, Hallé, & Tremblay, 2005). In the apo (substrate-unbound) form, the WPD-loop alternates between open and closed conformations, whereas in the substrate-bound form, the loop primarily samples a closed conformation. The tryptophan residue in the WPD loop acts as a hinge and mediates the loop’s mobility. The P-loop, also called the PTP-loop or phosphate-binding loop, is a component of motif-9 VHCSXGXGR(T/S)G. This contains the PTP’s signature HC-X₅-R motif’s active site cysteine and invariant arginine. The invariant arginine of this motif, combined with the P-loop’s backbone nitrogen atoms, generates a strong positively charged milieu in the active site and increases the PTP’s affinity for negatively charged phosphotyrosine substrates(John M Denu & Dixon, 1998). This arginine also aids in the proper placement of the substrate in the active site and stabilizes the cysteinyl-phosphate intermediate during catalysis. The histidine, serine/threonine, and invariant arginine contribute to decreasing the pKa of active site cysteine to around pH 6.5 and allows for it to maintain a deprotonated “thiol” state (Cys-S⁻)(Z. Y. Zhang & Dixon, 1993). Lastly, the Q-loop, which is part of motif-10 ((V/I/L)QTxxQYXF) comprises of two conserved glutamine residues that are essential for PTP active site function. Gln262 and Gln264 restricts the transfer of the phosphoryl group from the phosphoenzyme intermediate to a water molecule rather than other nucleophilic acceptors, preserving their hydrolase activity and preventing them from acting as kinase-like phosphotransferases (Z. Y. Zhang & Dixon, 1993; Zhao, Wu, Noh, Guan, & Zhang, 1998). Also, this loop creates hydrogen bonds with the P-loop and interacts with the aspartate of the WPD-loop to maintain the active site conformation required for maximum catalytic activity.

The E-loop is another conserved motif in PTPs, located close to the PTP-loop and WPD-loop (Figure 1B). This loop is made up of a 100% conserved glutamate residue and two 85–90% conserved lysine residues, which play crucial roles in coordinating the dynamics of the WPD loop and organizing the active site(J. N. Andersen et al., 2005; Jannik N. Andersen et al., 2001). The glutamate residue creates a bipartite hydrogen bond with the arginine of the P-loop, stabilizing the guanidium group in a location that increases its accessibility to the incoming phosphate. One of the conserved lysine residues is located close to the glutamate and interacts with the WPD loop’s catalytic aspartate via electrostatic interactions. This helps the WPD-loop to maintain its closed conformation in the substrate bound state. Indeed, changing this lysine to alanine reduces PTP’s catalytic performance significantly . The second lysine residue is located at the rim of the active site and aids in lowering the pKa of catalytic cysteine while also stabilizing the incoming pTyr substrate by giving a surface positive charge. Although the E-loop can take various forms (e.g., β-hairpin, fully disordered), the interaction of glutamate with PTP-loop arginine and the role of the two lysine residues are consistent across all PTPs (Asante-Appiah et al., 2006; Scapin, Patel, Patel, Kennedy, & Asante-Appiah, 2001).

A Cys-based catalytic mechanism and sensitivity to oxidation

All PTPs have one catalytic mechanism for phosphate monoester hydrolysis that utilizes their invariant nucleophilic (deprotonated thiol) cysteine and an aspartic acid as the general acid/base. The reaction follows a two-step double-displacement mechanism that makes a covalent intermediate at the PTP active site(Zhong Yin Zhang, Wang, & Dixon, 1994). The pTyr recognition loop (Tyr 46 in PTP1B) and E-loop (Glu 115 in PTP1B) residues help to admit and place the substrate at the active site (Stage I, Figure 2). A phenylalanine (Phe182 in PTP1B) downstream from the aspartate in the WPD loop generates π-π stacking contacts with the phosphotyrosine substrate, which helps align the substrate at the active site. In the first chemical step, the invariant cysteine (Cys215 in PTP1B) functions as a nucleophile, attacking the phosphorus on the phosphotyrosine substrate. This triggers the breakage of the phosphorus-oxygen link, resulting in the ejection of the product tyrosine, which is simultaneously protonated by the aspartate (Asp181 in PTP1B) of the WPD-loop (Stage II, Figure 2). At the end of this step, the active site is covalently modified and contains a cysteinyl-phosphate group at the P-loop (at Cys215)(John M Denu & Dixon, 1998; J. M. Denu, Lohse, Vijayalakshmi, Saper, & Dixon, 1996; Guan & Dixon, 1991)(Stage III, Figure 2). The conserved arginine (Arg 221 in PTP1B) from the P-loop interacts with the substrate’s phosphate group, assisting in the stability of the cysteinyl-phosphate intermediate. In the second (and rate-limiting) chemical step, glutamines (Gln 262, and Gln 266 in PTP1B) from the Q-loop sequester a water molecule, which is then attacked by deprotonated aspartate from the WPD loop(Stage IV, Figure 2). Simultaneously, catalytic aspartate attacks the intermediate’s phosphorus-sulfur link, facilitating hydrolysis of the covalent enzymatic intermediate and the release of free phosphate.

Figure 2: The PTP catalytic mechanism:

PTPs employ a conserved nucleophilic cysteine nested between a conserved HC-X₅-R active site motif. Stage I: The active site cysteine has a low pKa and operates in its thiolate form. Stage II: The pY-loop facilitates the entry of phosphotyrosine into the active site. The cysteine in the active site carries out a nucleophilic attack on the reactive phosphate group. This is facilitated by the aspartate residue in the WPD loop, which acts as a general acid/base. Stage III: A covalent intermediate is formed at the active site in the form of a cysteinyl phosphate group. Stage IV: glutamine from the Q loop activates water molecules to aid hydrolysis of the cysteinyl phosphate intermediate and regenerates the active site for subsequent catalytic cycles.

One of the factors influencing PTP catalytic rate is the dynamics of the WPD-loop, which varies between open and closed conformations during the catalytic cycle. Mutations affecting the flexibility of the WPD-loop have been shown to majorly impact PTP catalytic activity(Cui, Lipchock, Brookner, & Loria, 2019; Moise et al., 2018), indicating that motions of the WPD loop are correlated to the reaction rate (Whittier, Hengge, & Loria, 2013). More recent studies, however, indicate that the rate of catalysis of certain PTPs like PTP1B and HePTP is separate from their WPD loop’s dynamic equilibrium and that the internal protein dynamics of these PTPs, in regions both around and away from the active site influences catalytic activity(Choy et al., 2017; Crean, Biler, van der Kamp, Hengge, & Kamerlin, 2021; Cui et al., 2019). Therefore, it is more plausible that the process of catalysis is controlled by coherent and coordinated movements inside the active site, either in reaction to the opening of the loop or when the active site is in a completely closed condition(Choy et al., 2017; Torgeson, Clarkson, Kumar, Page, & Peti, 2020).

Oxidized PTPs have been reported in cells at physiological conditions indicating that redox processes underlie signaling events in cells(Tonks, 2005). Evidence suggests that redox signaling is especially prevalent in cancer cells where hyperactive PTKs mediate oxidative inhibition of PTPs(Chiarugi & Cirri, 2003). As the oxidation of cysteine in the active site of PTP results in the formation of an enzyme containing a nonfunctional catalytic center; it allows for uninterrupted phosphorylation signaling by PTKs in the absence of a counteractive phosphatase action (Chiarugi & Buricchi, 2007). PTP1B is reported to undergo oxidation in MEF thioredoxin−/− cells in response to PTK signaling(Lou et al., 2008). It is then reactivated by the thioredoxin system as is seen in NIH3T3 cells (Markus Dagnell et al., 2013). Oxidation of PTP1B by H₂O₂ is also reported in hepatoma (HepG2), adipose (3T3-L1), and human cancer A431cells in response to insulin receptor signaling(Lou et al., 2008; Mahadev, Zilbering, Zhu, & Goldstein, 2001). PTPs including SHP2, PTEN, DUSP1, and LAR are reported to be oxidized in angiomyolipoma tumor cells as a result of platelet-derived growth factor (PDGF) signaling (Boivin, Zhang, Arbiser, Zhang, & Tonks, 2008). Angiotensin II signaling in vascular smooth muscle cells produces oxidants that hinder SHP2 activity and stimulate Akt signaling(Frijhoff, Dagnell, Godfrey, & Östman, 2013). Oxidization of SHP1 and SHP2 proteins have been reported in EOL-1 myeloid leukemia and human embryonic kidney HEK293 cells(Weibrecht et al., 2007). SHP2 undergoes oxidation in fibroblasts during cell adhesion in response to PDGF stimulation, as well as in T-cells during cell migration in response to T cell receptor signaling(Frijhoff et al., 2013). RPTPε is reported to dimerize and undergo oxidation during EGF signaling and because of oxidative stress induced by treatment of HEK-293 cells with H₂O₂(Toledano-Katchalski et al., 2003).

Sources of Reactive Oxygen Species (ROS) in cellular signaling

All highly reactive oxygen (O₂) derivatives are referred to as Reactive Oxygen Species (ROS)(Bardaweel et al., 2018). This group consists of nonradical peroxides like hydrogen peroxide (H₂O₂), hypochlorous acid (HOCl), and singlet oxygen (¹O₂) that all contain a volatile O-O linkage and are therefore prone to becoming sources of free radicals. ROS also includes species of radicals like superoxide (O₂^•−), hydroxyl (^•OH), peroxyl (RO₂^•), and alkoxyl (RO^•) that have unpaired valence shell electrons(Bedard & Krause, 2007). According to several studies, hydrogen peroxide (H₂O₂) is the dominant ROS agent in cells(M. Dagnell, Cheng, & Arnér, 2021; M. Dagnell et al., 2019; Di Marzo, Chisci, & Giovannoni, 2018). The tendency of the highly reactive superoxide (O₂^•−) to rapidly undergo dismutation, both spontaneously and enzymatically, results in the formation of H₂O₂ which easily diffuses through membranes and across the cytosol. The O-O bond in H₂O₂, although less strong than the bond in dioxygen (O₂), makes the molecule relatively stable compared to other radical species. This stability allows H₂O₂ enough time to react with specific targets that it can oxidize. While H₂O₂ has been considered a toxic molecule since its discovery, in more recent times, it has become evident that the production of low concentrations of H₂O₂, under strict physiological control, plays a significant role in the regulation of several important signaling pathways including those involved in cell proliferation, differentiation, metabolism, and cell migration(Heo, Kim, & Kang, 2020; Lennicke, Rahn, Lichtenfels, Wessjohann, & Seliger, 2015; Prasad, Gupta, & Tyagi, 2017; Sies, 2017; Vilchis-Landeros, Matuz-Mares, & Vázquez-Meza, 2020).

The NADPH oxidase (NOX) family, which was initially identified as a source of microbiocidal oxidants produced in response to lipopolysaccharides, is recognized as the major producer of cellular ROS(Bedard & Krause, 2007; Brown & Griendling, 2009) (Figure 3A). The family includes seven different kinds of NOX complexes, known as NOX1, NOX2 (gp91phox), NOX3, NOX4, NOX5, DUOX1, and DUOX2 that are widely distributed in various cellular locations including the cell membranes, mitochondria, peroxisomes, and the endoplasmic reticulum. Their catalytic subunit is a transmembrane protein that binds electron-transferring FAD and heme prosthetic groups which can take an electron from NADPH. This electron is then used to reduce molecular O₂, leading to the production of O₂^·−, that is eventually converted to H₂O₂ by dismutases (Figure 3A). Research suggests that NOX -dependent ROS generation is linked with oncogenic signaling of RAS(J. A. Choi, Kim, Song, Kim, & Kim, 2008) and various growth factors including TGF-b1, interleukin-1, TNFa, insulin, PDGF, EGF, angiotensin II, thrombin, and lysophosphatidic acid (Mahadev et al., 2001; Mesquita et al., 2010; Ohba, Shibanuma, Kuroki, & Nose, 1994; Sundaresan, Yu, Ferrans, Irani, & Finkel, 1995; Svegliati et al., 2005; Weng, Chang, Hung, Yang, & Chien, 2018). Also, growth-factor stimulation activates the PI3K pathway, leading to Rac1 activation and triggers NADPH-oxidase activity (Bae et al., 2000; Svegliati et al., 2005).

Figure 3: Intracellular sources of ROS:

A. NADPH oxidase (NOX) enzymes are the primary producers of ROS. NOX enzymes are localized to several organelles and cellular membranes where they create superoxide (O₂^•−) radicals as a byproduct of NADPH oxidation. Lipoxygenases that convert arachidonic acid (2AA) to hydroperoxyeicosatetraenoic acids (HPETEs) also release superoxide (O₂^•−) into the cytosol. Subsequently, these superoxide molecules are transformed into hydrogen peroxide (H₂O₂) by the various superoxide dismutases (SOD). B. The mitochondrial respiratory chain is an incidental source of ROS. Superoxide (O₂^•−) radicals are created as a byproduct of ATP production and the movement of electrons through the electron transport chain. Included in the inner mitochondrial membrane is the respiratory chain complex comprised of NADH dehydrogenase (complex I), the succinate-coenzyme Q reductase complex (complex II), the cytochrome b-c1 complex (complex III), the cytochrome oxidase complex (complex IV) and the ATP synthase complex. Mitochondrial dismutases including MnSOD (manganese superoxide dismutase) and CuZnSOD (copper/zinc superoxide dismutase) convert superoxide (O₂^•−) radicals into the membrane permeable hydrogen peroxide (H₂O₂).

ROS are also produced as a byproduct of lipid metabolism. Several oxygenases, including lipoxygenase (Lox) and cyclooxygenase (Cox) enzyme families produce ROS during fatty acid metabolism as well as hormone and inflammatory mediator biosynthesis (Figure 3B). Eicosanoid biosynthesis occurs by the oxidation of fatty acids, such as arachidonic acid, catalyzed by the LOX enzymes. Alternatively, the COX enzymes can also catalyze the oxidation of fatty acids, resulting in the generation of prostaglandins. Both processes necessitate the introduction of molecular oxygen (O₂) to arachidonic acid, resulting in the generation of ROS molecules as a byproduct of this oxidative process. Additionally, leukotrienes, which are the final products of the LOX pathways, or intermediates such as hydroperoxyeicosatetraenoic acids (HPETEs) can induce the generation of ROS molecules from NOX and dismutatses(Boivin & Tonks, 2015; J. A. Choi et al., 2008).

An unintentional, yet critical source of H₂O₂ is the mitochondrial respiratory chain(Brand, 2016; Mailloux, 2015). ROS are produced as a byproduct of the electron transport chain’s activity during cellular aerobic respiration in the mitochondrion when O₂ is prematurely reduced by one electron and forms superoxide (O₂^•−) (Figure 3B). The amount of O₂^·− that is created during the transfer of electrons from NADH to O₂ and the simultaneous generation of ATP is determined by the relationship between the pool of accessible NADH electron donors, the local O₂ concentration alongside the efficiency of the ATP synthase. O₂^·− is converted into membrane-permeable H₂O₂ by the mitochondrial manganese superoxide dismutase (MnSOD) located in the matrix and copper/zinc superoxide dismutase (CuZnSOD) localized in the mitochondrial intermembrane space(Bedard & Krause, 2007). Hydrogen peroxide diffuses through mitochondrial membranes and into the cytosol via specific aquaporins (AQP) called peroxiporins(Sies, 2017). Oxidative stress caused by conditions such as hypoxia, injury to the mitochondria, or decreased levels of these free radical scavenging enzymes, enhance the “leakage” of ROS from the mitochondria and can trigger cell death(Prasad et al., 2017; Reczek & Chandel, 2015).

Oxidation of PTPs I: Cellular mechanisms

PTP’s cysteine-based active site makes them extremely sensitive to ROS action. The thiol chain of cysteine undergoes oxidation forming a reversible sulfenic acid (SOH) intermediate that can transform into a disulfide bond (S-S) with nearby cysteines or a sulfenyl amide (S-N) bond with a neighboring backbone nitrogen. These can be reversed to the “active” thiol form by antioxidants such as glutathione, thioredoxin, and their associated thiol-transferases like glutaredoxin(Ostman et al., 2011). Further oxidation of the sulfenic acid (SOH) form creates the hyperoxidized sulfinic (SO₂H) and sulfonic (SO₃H) states that are resistant to regeneration and lead to irreversible PTP inactivation (Figure 4). An increasing number of studies show that ROS are not a simplistic tool for the broad oxidation of random targets. The properties of the oxidizing agents produced have implications for the particular signaling response that is activated. Juarez et al. have demonstrated that PTP1B is suppressed by H₂O₂, but not by O₂^•−(Juarez et al., 2008). Similarly, Choi et al. show that specificity at the level of oxidant-targeting exhibits preferences for the target’s downstream effectors(H. K. Choi, Kim, Jhon, & Lee, 2011). Their studies how that SHP1 is inactivated in hematopoietic progenitors by ROS production caused by macrophage colony stimulating factor, but its close homologue SHP2 is not affected. Also, the oxidative inactivation of SHP1 specifically impacts the PI3K/Akt survival pathway, while the mitogen-activated protein kinase (MAPK) survival signaling remains unaffected, despite both pathways being regulated by SHP1.

Figure 4: PTP oxidation pathways:

In the presence of H₂O₂, the PTP catalytic cysteine’s thiolate ion is reversibly oxidized to sulfenic acid. Sulfenic acid’s instability causes it to quickly transform into a thiolate ion or reversibly oxidized intramolecular disulfide or sulfenyl amide state. Agents like glutathione and thioredoxin can reduce these to a thiol group. However, further oxidation of the sulfenic acid by H₂O₂ creates irreversibly oxidized sulfinic acid and sulfonic acid forms that permanently inactivate PTP active sites.

Kinetics wise, oxidation of PTPs with H₂O₂ is much slower (~10¹ M⁻¹ s⁻¹) compared to other reactive cysteine containing enzymes like Peroxiredoxins (Prxs) (~10⁷ M⁻¹ s⁻¹)(Winterbourn & Metodiewa, 1999). These Prx enzymes serve as sensors of ROS in cells and react specifically with hydroperoxides, but not with other electrophiles such as iodoacetamides or chloroamines (Peskin et al., 2007). Moreover, as peroxiredoxins are highly abundant in cells, and have a faster rate of oxidation, it is understood that they outcompete PTPs for reaction with H₂O₂. At present three non-mutually exclusive mechanisms of PTP oxidation have been outlined (Figure 5)(L. E. Netto & Antunes, 2016). The first model is of direct reaction of H₂O₂ with the PTP active site cysteine. The second model details an indirect, yet specific, oxidation method following a “redox relay mechanism” involving sequential transfers of oxidizing equivalents between Prxs and PTPs. At first, catalytic cysteines on Prx enzymes are oxidized by H₂O₂ to sulfenic acids (CysSOH) that subsequently converted to intra- or intermolecular disulfides (Cys-SS-Cys). Oxidized Prx enzymes use thiol-disulfide exchange reactions to transfer oxidizing equivalents to PTPs that they select by specific protein-protein interactions(Marinho, Real, Cyrne, Soares, & Antunes, 2014; L. E. Netto & Antunes, 2016; Perkins, Nelson, Parsonage, Poole, & Karplus, 2015). Support for this model comes from studies on Ras-induced breast cancer in mammary epithelial cells where Prx1 is seen to be critical for supporting the tumor suppressive function of PTEN phosphatase(Cao et al., 2009). Here, physical interaction between Prx1 and PTEN, and thiol exchange between the two proteins is reported to be critical for protecting PTEN from oxidation-induced inactivation. The third model of H₂O₂ sensing includes thioredoxin (Trx) enzymes that also function as redox sensors and can perform the “redox relay” with either PTPs or Prx enzymes. Studies show that while Trx is a slower redox sensor than Prx, it is extremely selective in its protein-protein interactions under different redox conditions (Berndt, Lillig, & Holmgren, 2007; Palde & Carroll, 2015). Subtle structural changes in Trx modify its protein interaction surfaces such that certain Trx-interactor complexes only exist when Trx is in its oxidized form(Berndt et al., 2007; Psenakova et al., 2020). For example, in vitro- and cell-based assays indicate that Trx1 binds and reduces oxidized PTP1B protein but not the SHP2 phosphatase(Markus Dagnell et al., 2013). Cells lacking Trx reductase TrxR1 (that regenerates reduced Trx1) show increased oxidation of PTP1B with no corresponding change in SHP2 oxidation. Hence models 2 and 3 indicate that redox sensing by PTPs is specified by critical protein-protein interactions that are a maneuver around the slow direct oxidation rates of these proteins. Under high H₂O₂ concentrations when Prxs are hyperoxidized to sulfinic or sulfonic states, localized accumulation of H₂O₂ allows for direct oxidation of PTPs despite their low oxidation rates. This is called the “floodgate hypothesis” (Rhee & Woo, 2011). However, this hypothesis is controversial because several studies show Prxs to be much more abundant than PTPs(Perkins et al., 2015), and that they use several alternate pathways for rapid regeneration(Rhee & Woo, 2011). Also, when Prxs are permanently inactivated by oxidation, cells have other antioxidants like GSH and GAPDH that can easily outcompete PTPs for chemical reactions with H₂O₂₍Winterbourn & Hampton, 2008_).

Figure 5: Cellular mechanisms of PTP oxidation:

PTPs are oxidized by H₂O₂ directly, or indirectly through the oxidation of peroxiredoxin (Prx) and thioredoxin (Trx) proteins that contain reactive thiols with higher affinity for hydrogen peroxide (H₂O₂) that PTPs. In the indirect mechanism, a redox relay of thiol-disulfide exchange reactions engages several proteins that are eventually reduced by glutathione and thioredoxin.

Oxidation of PTPs II: Structural aspects

Crystal structures of several PTPs have allowed for a thorough analysis of their H₂O₂ concentration-dependent oxidation (Table 1). PTP1B has been crystallized in all oxidized states of its catalytic cysteine (Figure 6) (Salmeen et al., 2003; van Montfort, Congreve, Tisi, Carr, & Jhoti, 2003). The reversible sulfenyl amide form consists of a covalent bond between the active site cysteine and the backbone nitrogen of the neighboring serine residue in the P-loop. Initially, it was suggested that sulfenyl amide formation in PTP1B proceeded via direct S_N2 mechanism when the backbone nitrogen of the Ser216 nucleophilically attacked the Sγ atom of the active site Cys215 SOH present in the sulfenic acid form(van Montfort et al., 2003). This was supported by the protonation of the active site His214 which also maintains the thiol form of Cys215(Z. Y. Zhang & Dixon, 1993). More recently, molecular dynamics simulations and high-level hybrid quantum mechanics/molecular mechanics-based studies show that sulfenyl amide formation occurs through an iminol-type intermediate that requires the participation of Glu115 (of the E loop) alongside His214(Dokainish & Gauld, 2015).

Table 1:

List of crystal structures of PTPs exhibiting their active sites cysteine in different oxidized states..

Protein Name	Active site cysteine	PDB ID	Reference
PTP1B	Reduced	5K9V	(Choy et al., 2017)
PTP1B	Oxidized to sulfenyl amide	1OEM	(Salmeen et al., 2003)
PTP1B	Oxidized to sulfenyl amide	1OES	(van Montfort et al., 2003)
PTP1B	Oxidized to sulfonic acid	1OEO	(Salmeen et al., 2003)
PTP1B	Oxidized to sulfonic acid	1OET	(van Montfort et al., 2003)
PTP1B	Oxidized to sulfenic acid	1OEU	(van Montfort et al., 2003)
PTP1B	Oxidized to sulfinic acid	1OEV	(van Montfort et al., 2003)
PTPRU D1 domain	Reduced	6SUB	(Hay et al., 2020)
PTPRU D1 domain	Oxidized to sulfonic acid	6SUC	(Hay et al., 2020)
PTPσ D1D2	D1 oxidized to sulfenic acid, D2 reduced	4BPC	(Jeon et al., 2013)
LYP	Reduced	2QCJ	Unpublished, structure available in the PDB
LYP	Oxidized, disulfide bonded to backdoor cysteine	3H2X	(Tsai et al., 2009)
SHP2 (N308D mutant)	Reduced	4NWF	Unpublished, structure available in the PDB
SHP2 (N308D mutant)	Oxidized, disulfide bonded to backdoor cysteine	6ATD	(Machado, Critton, et al., 2017)
STEP	Modified to S-ACETYL-CYSTEINE	2BV5	(Eswaran et al., 2006)

Figure 6: Structural features of reduced/oxidized PTP1B:

A. The reduced functional PTP1B contains a central active site Cysteine in a Thiol form flanked by various other active site loops. The WPD loop faces away from the active site cysteine in the apo (left, green) and moves towards the cysteine when the substrate binds (center, pink). The inset on the right shows the relative movements of the active site loops upon phosphotyrosine binding. B. The reversibly oxidized sulfenyl amide form of PTP1B is formed when the active site cysteine makes an “S-N” bond with the backbone nitrogen atom of the neighboring serine. This structure shows a unique and distinct conformational change in its P-loop (inset, right). C. Higher oxidized forms of PTP1B include its cysteine’s oxidation to sulfenic acid (green, left), sulfinic acid (center, blue), and sulfonic acid (right, gray). The sulfinic and sulfonic acid forms cannot be reduced by antioxidants to regenerate the active form of the enzyme. The insets under each structure show the chemical groups of the P-loop and highlight the hydrogen bonds between the active site cysteine and its neighboring residues.

In the sulfenyl amide state, the active site of PTP1B is distorted and shows alterations in the positioning of its active site loops(Salmeen et al., 2003; van Montfort et al., 2003). Changes are most pronounced in the conformational of the pY loop such that Tyr46 residue flips and breaks its interactions with the Ser 216 of the P-loop. Gly218 of the P-loop pivots by around 7Å, and the P-loop switches to break interactions between the side chain of Ser216 and Glu115 of the E-loop. Gln262 of the Q-loop switches its conformations and faces away from the active site (Figure 6). These perturbations expose the otherwise Ser50 to the cytosol and promote its phosphorylation by AKT (Ravichandran, Chen, Li, & Quon, 2001). This oxidized and phosphorylated PTP1B is transiently stabilized by binding of 14–3-3ζ protein and promotes sustained EGFR signaling as is seen in HEK293 cells(Londhe et al., 2020). Distortion in the active site of PTP1B in the sulfenyl amide form has been used to generate conformation-specific antibodies that can allow for studying reversible oxidation of PTP1B in cellular systems(Haque, Andersen, Salmeen, Barford, & Tonks, 2011). Krishnan et al. have used intrabody scFv45 to trap the reversibly oxidized PTP1B-OX in the catalytically inactive form and studied its effect on enhancing insulin and leptin signaling in hepatic stellate cells(Krishnan et al., 2018). These studies reveal the intricate interplay between phosphorylation and redox based signaling. Efforts are underway to harness the intrabody approach as a novel therapeutic methodology for the treatment of diabetes and obesity.

When compared to the sulfenyl amide form, perturbations in PTP1B active site aren’t as pronounced in the higher oxidized sulfenic, sulfinic, and sulfonic acid states(Salmeen et al., 2003; van Montfort et al., 2003) (Figure 6). In these states, the active site loops maintain the overall conformation as seen in the reduced Cys215 of the active enzyme. In the sulfenic form, Cys215 SOH forms a hydrogen bond with the side chain and backbone amide of Ser222 and keeps the P-loop locked in a tight conformation. Similarly, in the sulfinic form, Cys215 SO₂H forms hydrogen bonds with Ser222 that are stabilized by backbone amide groups of Ala217 and Gly218, also of the P-loop. Lastly, in the sulfonic form, Cys215 SO₃H forms similar hydrogen-bonded interactions as seen in the sulfenic and sulfinic acid states. Sulfenic and sulfonic acid states have also been crystallized for the receptor PTPσ where its active site Cys1589 shows similar conformations as it were in the reduced active form (Table 1)(Jeon, Chien, Chun, & Ryu, 2013). Like in PTP1B, higher oxidized forms of PTPσ show an open conformation of its WPD-loop. Interestingly, PTPσ has two cytosolic PTP domains, but only its membrane proximal D1-PTP domain is oxidized while its membrane distal D2-PTP domain remains reduced. As the D2-PTP domain is a naturally occurring pseudo-enzyme (lacks phosphatase activity), its differential oxidation/reduction processes from the D1 domain suggest an allosteric interplay between the two PTP domains of PTPσ.

Some PTPs including LYP, SHP1/SHP2, STEP, PTPRR, and PTPρ use a conserved cysteine in their motif 7 (KCxxYWP) to make a reversible disulfide bond with their active site cysteines(C.-Y. Chen, Willard, & Rudolph, 2009; Cunnick, Dorsey, Mei, & Wu, 1998; Hay et al., 2020; Machado, Critton, Page, & Peti, 2017; Machado, Shen, Page, & Peti, 2017; Tsai et al., 2009). This partnering cysteine here is referred to as the “backdoor” cysteine because of its location in the backside of the PTP active site. Disulfide formation does not alter the conformation of the P-loop but is mediated by a simple rotation of the active cysteine to access the backdoor cysteine (Figure 7). In LYP, a disulfide bond between the active site Cys227 and backdoor Cys129 stabilizes a partially open active site with the WPD-loop in an intermediate conformation(Tsai et al., 2009). This intermediate conformation is stabilized by electrostatic interactions between Glu133 of the E-loop and Arg233 of the P-loop. Interestingly, LYP is proposed to have two more backdoor cysteines including Cys139 in the E-loop and Cys231 that is downstream of the active site cysteine in the P-loop. It is unclear how the active site Cys227 chooses to form disulfide bonds given that the three backdoor cysteines are easily accessible. Biochemical data shows that Cys129S and Cys231S mutants of LYP allow for reversible oxidation of the PTP, perhaps by engaging the third Cys139 from the E-loop(Tsai et al., 2009).

Figure 7: Reversible oxidation via disulfide binds with a backdoor cysteine:

Lyp (A, top) and SHP2 (B, bottom) use the backdoor cysteine in motif 7 (KCxxYWP) to make a reversible disulfide bond and prevent irreversible oxidization of the active site cysteine to sulfinic and sulfonic acids. The insets on the right show the active site structure and disulfide bond formation in the two PTPs.

In SHP2, the active site Cys459 engages the backdoor Cys367 in a disulfide bond, and its WPD-loop is seen in an intermediate conformation(Machado, Critton, et al., 2017) (Figure 7). In both SHP1 and SHP2, a conserved proline in motif 4 (YILTQGP) is naturally seen as a cysteine (Cys327 in SHP1 and Cys333 in SHP2) that functions as the second backdoor cysteine and supports reversible oxidation of SHP1/SHP2 (C.-Y. Chen et al., 2009). Removal of both backdoor cysteines (Cys327, Cys373 in SHP1 and Cys333, Cys367 in SHP2) is necessary for the irreversible oxidative inactivation of SHP1/SHP2 to sulfinic and sulfonic acid states. Mutations of the second backdoor cysteine to the canonically conserved proline residues enhances the thermal stability of SHP1/SHP2 catalytic domains(Yarnall, Kim, Korntner, & Bishop, 2022). As SHP1/SHP2 is allosterically regulated by the movements in their tandem SH2 domains(C. L. Welsh et al., 2023), the presence of a second backdoor cysteine hints towards an evolved redox protection of the PTPs at the cost of protein stability.

Given that the backdoor cysteine is a conserved residue in motif 7 (KCxxYWP) of all PTPs, it’s intriguing that only some of them employ it for an intramolecular disulfide bond. Furthermore, the extent of antioxidant protection afforded by the backdoor cysteine differs among related PTPs. Mutation of the backdoor cysteines to serine in STEP (Cys384) and PTPRR (Cys501) makes them more sensitive to H₂O₂ mediated oxidation when compared to the wild type proteins, but PTPRR-C501S is less sensitive than STEP-C384S mutant and more easily reactivated by chemical reducing agents such as Dithiothreitol (DTT) (Machado, Shen, et al., 2017). Perplexingly, these studies also report that in the closely related HePTP, the backdoor cysteine Cys183 is dispensable for its reversible oxidation. Unlike its related STEP and PTPRR proteins (collectively known as the Kinase Interaction Motif (KIM)-PTPs, which dephosphorylate mitogen-activated protein kinases (MAPKs), HePTP is reported to form intermolecular disulfide bonds between two protein monomers; using the active site Cys270 of one protomer and a surface exposed Cys116 on another protomer(Machado, Shen, et al., 2017). Also, the reactivation rates of HePTP by DTT were found to be similar to those observed for PTP1B and SHP1/SHP2, although STEP and PTPRR were reported to be extremely slowly reactivated by reducing agents. These findings imply that protein-specific, most likely dynamics-based indicators may play a role in determining each PTP’s oxidative proclivity and reactivation susceptibility.

Despite all the sequence and structural data available, it is impossible to predict how a PTP would react to or protect itself against oxidation. Nonetheless, the conserved backdoor cysteine has been explored as a plausible allosteric site that could be harnessed for PTP inhibition. Covalent modification of PTP1B’s backdoor Cys121 by 4-(aminosulfonyl)-7-fluoro-2,1,3-benzoxadiazole (ABDF) is reported to decrease its catalytic activity by several fold without affecting peptide substrate binding (Hansen et al., 2005). ABDF has also been shown to interact with and inhibit the catalytic activity of TC-PTP and LAR PTPs while having no effect on CD45, indicating some selectivity. ABDF treatment of CHO-hIR cells has been shown to suppress PTP1B activity and allowing for sustained insulin receptor phosphorylation and signaling. Similarly, covalent modification of Cys121 by 1,2-naphthoquinone is reported to decrease PTP1B’s catalytic activity and allow for persistent transactivation of EGFR in human epithelial A431 cells(Iwamoto et al., 2007). More recently, biconjugate PTP1B inhibitors have been designed to covalently modify Cys121 while simultaneously engaging the active site Cys215 in a non-covalent interaction(Khan, Bjij, & Soliman, 2019; Punthasee et al., 2017). These use electrophilic α-bromoacetamide or α-fluoroacetamide groups conjugated to 5-aryl-1,2,5-thiadiazolidin-3-one 1,1-dioxide inhibitors that selectively react with the distal Cys121 and provide for an allosteric mechanism of PTP1B inhibition.

Conclusions

Oxidation of PTPs adds another complexity to the already intricate landscape of phosphorylation and redox-based cellular signaling. Currently, there is a renewed interest in understanding the structural and pharmacological processes that regulate PTPs in order to target them for drug discovery. Understanding the protein-specific reversibly oxidized states of PTPs is crucial for accessing and targeting these proteins. Conformation-specific characteristics of oxidized PTP1B have been utilized to produce antibodies that may help target it for therapeutic intervention, but several key mechanistic concerns remain unexplained. For example, why does the active site cysteine in PTP1B form a sulfenyl amide bond with the adjacent serine rather than a disulfide link with its backdoor cysteine? What chemical characteristics of the active site and backdoor cysteines influence their interactions in a reversible oxidized state? The range of oxidized states observed in PTPs suggests a protein-specific preference for oxidation states, which may be connected to protein dynamics rather than conserved sequence and structure. Further research, particularly on the significance of protein dynamics in conjunction with PTP oxidation, is essential to answer these questions.