Chemical Probes for Redox Signaling and Oxidative Stress

Abstract

Significance: Cellular reactive oxygen species (ROS) mediate redox signaling cascades that are critical to numerous physiological and pathological processes. Analytical methods to monitor cellular ROS levels and proteomic platforms to identify oxidative post-translational modifications (PTMs) of proteins are critical to understanding the triggers and consequences of redox signaling.

Recent Advances: The prevalence and significance of redox signaling has recently been illuminated through the use of chemical probes that allow for sensitive detection of cellular ROS levels and proteomic dissection of oxidative PTMs directly in living cells.

Critical Issues: In this review, we provide a comprehensive overview of chemical probes that are available for monitoring ROS and oxidative PTMs, and we highlight the advantages and limitations of these methods.

Future Directions: Despite significant advances in chemical probes, the low levels of cellular ROS and low stoichiometry of oxidative PTMs present challenges for accurately measuring the extent and dynamics of ROS generation and redox signaling. Further improvements in sensitivity and ability to spatially and temporally control readouts are essential to fully illuminate cellular redox signaling.

Introduction

Reactive oxygen species (ROS) are produced endogenously during a variety of cellular processes, and primary among these is aerobic respiration. In eukaryotic systems, the majority of cellular ROS is believed to be generated during the transfer of electrons through components of the electron transport chain localized in the inner mitochondrial membrane. This electron transport process results in the reduction of molecular oxygen (O₂) to water, and it drives adenosine triphosphate (ATP) synthesis to fuel key physiological processes. However, partial reduction of O₂ can result in the production of numerous ROS, including superoxide (O₂^•−) and hydrogen peroxide (H₂O₂) (Fig. 1A). Further, controlled ROS production also occurs at reduced nicotinamide adenine dinucleotide phosphate (NADPH) oxidase (NOX) enzymes, which oxidize NADPH to nicotinamide adenine dinucleotide phosphate (NADP⁺), and reduce O₂ to produce O₂^•− as a primary product. In various physiological processes, including cellular proliferation and migration, cytosolic regulatory subunits are assembled on membrane catalytic NOX proteins to activate ROS production (60, 118). Importantly, localization of NOX subunits provides spatial regulation within cells, such as the assembly of regulatory and catalytic subunits at the leading edge of migrating cells (125).

FIG. 1.

ROS and oxidative PTMs of cysteine. (A) ROS are derived from molecular oxygen through excitation and reduction. (B) Oxidative PTMs of cysteine highlighted in this review. PTMs, post-translational modifications; ROS, reactive oxygen species. Color images are available online.

To counteract the damaging effects of excessive ROS generation, organisms possess a system of small-molecule antioxidants, including ascorbic acid and glutathione, as well as redox enzymes, including glutaredoxins (Grxs), thioredoxins (Trxs), and peroxiredoxins (Prxs) (126). In unicellular organisms, ROS sensors trigger activation of antioxidant systems when ROS concentrations exceed physiological levels. For example, the H₂O₂ sensors OxyR (e.g., prokaryotes) and Orp1 (e.g., yeast) directly sense peroxide levels and activate the production of antioxidant enzymes such as catalase (18). In multicellular organisms, antioxidant enzymes play important roles in spatiotemporal regulation of redox signaling. Evidence suggests that the subcellular localization of antioxidant enzymes restricts ROS to a particular cellular compartment (126). In addition, antioxidant enzymes such as Prxs are inactivated by oxidation or phosphorylation, and they form temporal “hot spots” for redox signaling through feedback inhibition (102, 133).

These highly controlled systems of cellular ROS generation and attenuation result in strict spatiotemporal regulation of cellular ROS levels. It was initially believed that the high reactivity of ROS resulted mainly in irreversible damage to biomolecules; however, it is now realized that ROS can play critical roles in cell signaling and regulation. This process, known as redox signaling, is a key component of numerous physiological and pathological processes, including growth-factor signaling and inflammation. One of the major effectors of ROS in redox signaling are thiol groups of cysteine residues in proteins, which can be transformed to form distinct oxidative post-translational modifications (PTMs) such as S-sulfenylation, S-sulfinylation, and S-sulfonylation, as well as disulfide-bond formation (Fig. 1B). S-sulfenylation (sulfenic acid [RSOH]) is a reversible PTM under cellular redox potentials, at which sulfenic acids can readily be reduced to the corresponding thiol by cellular antioxidants. Sulfenic acids can also be converted to intra- or intermolecular disulfide bonds with thiol groups of redox-active small molecules and proteins. Further oxidation of sulfenic acids results in S-sulfinylation and S-sulfonylation. Sulfinic acids (RSO₂H) are only reversed by sulfiredoxin (Srx) (14), and sulfonic acids (RSO₃H) are considered irreversible in cells. Among the oxidative PTMs of cysteine (Cys), the reversibility of S-sulfenylation renders it suitable for temporal signal transductions akin to phosphorylation. Well-studied examples of S-sulfenylation include protein tyrosine phosphatase 1B (PTP1B) and epidermal growth factor receptor (EGFR). PTP1B is inhibited by H₂O₂ through S-sulfenylation of the catalytic Cys215 on growth-factor stimulation (63), and EGFR is activated through S-sulfenylation of Cys797 (93). Both these S-sulfenylation events serve to further magnify phosphorylation events that are key to growth-factor signaling.

To elucidate the ubiquity and functional consequences of redox signaling resulting from cellular ROS generation, it is critical to be able to visualize ROS production and analyze subsequent oxidative PTMs of cysteine. ROS detection has been enabled by small-molecule and protein-based probes, which are coupled to spectroscopic readouts that provide sensitive ROS detection in living cells (in situ). Oxidative PTMs resulting from increased ROS levels can be interrogated by applying chemical probes coupled with proteomic technologies such as gel electrophoresis and mass spectrometry (MS). In this review, we summarize the available array of chemical probes for analyzing both ROS generation and downstream oxidative cysteine PTMs. Detailed mechanisms, advantages, and limitations of each method will be discussed where relevant.

Chemical Probes for ROS

ROS and the related reactive nitrogen species (RNS) typically display high reactivity with biomolecules (Fig. 1A). ROS include O₂^•−, H₂O₂, hydroxyl radical (^•OH), hypochlorite (⁻OCl), and singlet oxygen (¹O₂). RNS include peroxynitrite (ONOO⁻), which is generated via the reaction of nitric oxide (^•NO) with O₂^•−. Due to the distinct reactivity characteristics of each ROS and RNS, these species can display unique effects on physiological processes. For example, O₂^•− and H₂O₂ are relatively mild reactants and are typically implicated in redox signaling that is mediated through oxidative PTMs of cysteine. In contrast, ⁻OCl is a stronger oxidant that is produced by myeloperoxidase (MPO) in phagosomes, and it attacks invading microbes that are engulfed by neutrophils and macrophages. Due to the unique reactivity characteristics of each ROS, it is imperative to develop methods that are specific for each species. To this end, a number of chemical probes that are specific to distinct ROS have been developed and utilize electron paramagnetic resonance (EPR), chemiluminescence, fluorescence spectroscopy, and microscopy platforms to monitor ROS levels. In this review, we focus on chemical probes for O₂^•− and H₂O₂, which are considered the most relevant to redox signaling (18, 102).

Chemical Probes for O₂^•−

EPR probes for O₂^•−

EPR detects the absorbance of microwave energy by radicals with unpaired electrons in an applied magnetic field. Because ROS are very short lived and unable to be directly detected in biological samples, chemical probes that form stable adducts with ROS, termed spin traps, are used for biological ROS detection by EPR. A nitrone-based probe, 5,5-dimethyl-1-pyrroline-N-oxide (DMPO), is the most widely used EPR probe for O₂^•−. DMPO reacts with unstable free radicals, such as O₂^•− and ^•OH, to form relatively stable radical adducts, DMPO/^•OOH and DMPO/^•OH, respectively, which can be characterized by EPR (Fig. 2A). The DMPO probe has been applied to detect free radicals in a wide variety of cells, including endothelial cells (105, 144) and myocytes (128). One of the limitations of DMPO is the short half-life of DMPO/^•OOH (t_1/2 ∼45 s), which lowers the detection sensitivity in EPR measurements. To overcome this limitation, a derivative of DMPO, 5-(diethoxyphosphoryl)-5-methyl-1-pyrroline-N-oxide (DEPMPO), has been developed, which generates a more stable adduct with O₂^•− (t_1/2 ∼15 min) (29). Another limitation of DMPO is the potential decomposition of DMPO/^•OOH to DMPO/^•OH through reductive degradation by biological antioxidants. The susceptibility to reductive degradation hampers sensitive detection in situ, resulting in false negatives when O₂^•− levels are low (9). The possibility of degradation also requires control experiments with superoxide dismutase (SOD) mimics to distinguish O₂^•− from ^•OH (119). Further, nitrone-based probes are limited by the slow reaction rates displayed with O₂^•− (e.g., k = 74 M⁻¹·s⁻¹ for a nitrone-based derivative, EMPO) (22).

FIG. 2.

Chemical probes for superoxide. (A) EPR-based spin traps form stable radicals with superoxide. (B) HE and MitoHE react with superoxide to form fluorescent products. (C) Lucigenin releases a photon on reaction with superoxide. EPR, electron paramagnetic resonance; HE, hydroethidine. Color images are available online.

Cyclic hydroxylamine-based probes, including 1-hydroxy-3-carboxy-2, 2, 5-tetramethyl-pyrrolidine hydrochloride (CPH), provide an alternative method for O₂^•− detection with high reliability and sensitivity (59) (Fig. 2A). CPH reacts with O₂^•− with a fast reaction rate (k = 3.2 × 10³ M⁻¹·s⁻¹) to give a stable radical product (t_1/2 = 330 min in smooth muscle cells) (23). CMH (1-hydroxy-3-methoxycarbonyl-2,2,5,5-tetramethylpyrrolidine), a plasma membrane-permeable derivative of CPH, has been applied to O₂^•− detection in cultured cells and tissue samples (24, 59). Cyclic hydroxylamine-based probes have overcome drawbacks of nitrone-based probes, such as slow reaction with O₂^•− and reductive degradation in living organisms. However, the reaction rates of hydroxylamine-based probes are still slower than that of spontaneous dismutation of O₂^•− under physiological conditions (k = 10⁵–10⁶ M⁻¹·s⁻¹) (113). In addition, these probes can be oxidized by multiple ROS and auto-oxidation as well, which necessitates control experiments with specific ROS-scavengers, such as SOD and catalase.

Triarylmethyl (TAM) radical-based probes have also been reported; the TAM radicals are EPR active and lose the EPR signal through the reaction with O₂^•− at fast reaction rates (k = 3.1 × 10³ M⁻¹·s⁻¹ for a TAM derivative, OXO63) (103). The TAM radical-based probe, CT-02H, showed selectivity to O₂^•− and ROO^• among ROS and was applied to the detection of cellular O₂^•− release on menadione stimulation (70). A major limitation of TAM radical-based probes is the signal-decreasing (turn-off) response to O₂^•−, which lowers sensitivity of detection due to high background signals and hampers the wide application of TAM probes in living cells.

Fluorescent and chemiluminescent probes for O₂^•−

Fluorescence imaging is a facile and sensitive methodology to detect ROS generation by coupling ROS-reactive chemical probes with fluorescence spectroscopy. Hydroethidine (HE) is a widely used fluorescence probe for O₂^•− both in vitro and in situ (Fig. 2B). HE is oxidized by O₂^•−, to form the highly fluorescent 2-hydroxyethidium (2-OH-E⁺) with a high reaction rate (k = 2.6 × 10⁶ M⁻¹·s⁻¹) (138, 139). However, other ROS such as ONOO⁻, OH^•, and H₂O₂ oxidize HE to give a fluorescent product, ethidium (E⁺). High-performance liquid chromatography can be used to separate 2-OH-E⁺ from E⁺ and provide a quantitative and specific measure of O₂^•− in the system (28). E⁺ and 2-OH-E⁺ can also be distinguished by fluorescence spectroscopy by excitation at 405 nm instead of 480 nm, which significantly reduces the fluorescence signal from E⁺ (85). However, control experiments utilizing SOD are still required to confirm specific detection of O₂^•−. A mitochondria-targeted HE derivative, Mito-HE (also known as MitoSOX) contains a positively charged triphenylphosphonium (TPP) group, which allows Mito-HE to accumulate in mitochondria (142). Mito-HE has been applied to the detection of mitochondrial O₂^•− production in a wide range of cells, including aortic endothelial cells (86) and cultured cerebellar granule neurons (48).

Hydrocyanines are reported as probes for several ROS, including O₂^•− (57). The reduced cyanines are readily prepared by reduction of cyanine dyes with sodium borohydride, resulting in a drastic change in cyanine absorption and fluorescence. One derivative, hydro-carbocyanine 3 (Cy3), displays a fluorescence increase on treatment with O₂^•−, ^•OH, and t-butyl peroxyl radical (TBO^•). One advantage of hydrocyanines over HE is the increased stability in aqueous solutions; the half-lives of hydro-Cy3 and hydro-carbocyanine 7 (Cy7) in neutral aqueous solutions are ∼3 days, whereas that of HE is 30 min. Deuterated hydrocyanines have been reported to be more stable against auto-oxidation, which reduces background signals and allows for more sensitive detection of ROS (56). Hydro-Cy3 has been applied to live-cell imaging of rat aortic smooth muscle cells stimulated by angiotensin II. Another advantage of hydrocyanines is near-infrared absorption and fluorescence, which enables applications of hydrocyanines for in vivo imaging of ROS production in mice (35, 57).

Sulfonate-based protecting groups are utilized as a design strategy for O₂^•− probes. Screening a library of benzenesulfonyl fluorescein (BES) derivatives identified BES-So and BES-H₂O₂ as fluorescent probes for O₂^•− and H₂O₂, respectively (74, 75). In addition, HKSOX-1, a fluorescein protected by triflate groups, is reported to be responsive to O₂^•− and applied to in vivo O₂^•− imaging in zebrafish (44).

Chemiluminescent probes are widely used for O₂^•− detection. The most common chemiluminescent probe for O₂^•− is lucigenin (Fig. 2C). In the initial step, lucigenin is reduced by O₂^•− to a cation radical, which then reacts with a second O₂^•− molecule to form a dioxetane structure. A photon is emitted when the high-energy dioxetane structure is cleaved. Although lucigenin-based chemiluminescence is facile to monitor, lucigenin itself can produce O₂^•− through a process known as redox cycling. This redox cycling phenomenon leads to the overestimation of O₂^•− levels in the sample (120), and it is important to minimize artifactual production of O₂^•− from lucigenin by choosing appropriate experimental conditions (67). Another class of chemiluminescent probes for O₂^•− are derived from coelenterazine, which is a luciferin that is responsible for the bioluminescence of marine organisms. Coelenterazine has been reported as a chemiluminescent probe for O₂^•− with enhanced light emission relative to lucigenin (52). One advantage of coelenterazine is that it does not produce artificial O₂^•− through redox cycling (120). Derivatization of coelenterazine has resulted in the improved probes, 2-methyl-6-phenylimidazo[1,2-a]pyrazin-3(7H)-one (CLA) (116) and 6-(4-methoxyphenyl)-2-methyl-imidazo[1,2-a]pyrazin-3(7H)-one (MCLA) (89); the optimized derivative MCLA has a significantly enhanced emission (>100-fold) over coelenterazine. CLA and MCLA have been applied to O₂^•− detection in a variety of biological samples such as living cells (53, 54) and tissues (58, 130). It is reported that MCLA can emit light by the reaction with ROS other than O₂^•− (49, 90); thus, control experiments with SOD and other specific quenchers of ROS are required.

Chemical Probes for Hydrogen Peroxide

Small-molecule-based probes for H₂O₂

The most widely used chemical probes for H₂O₂ are reduced dyes such as dihydrodichlorofluorescein (DCFH₂) and dihydrorhodamine123. Oxidation of these reduced dyes by ROS generates a fluorescence increase. However, these reduced dyes have several critical drawbacks when considered as fluorescence probes for H₂O₂. First, DCFH₂ has poor reactivity and specificity to H₂O₂; DCFH₂ is even more responsive to highly ROS, including ^•OH, ONOO⁻, and ⁻OCl, than H₂O₂ (112). Thus, DCFH₂ should be regarded as an indicator of gross ROS amount rather than an H₂O₂-specific probe. In addition, DCFH₂ can be oxidized by light irradiation, which requires intensive protection from light throughout the experimental workflow. In addition, the irradiation during fluorescence readings can also trigger increased DCFH₂ fluorescence, rendering the interpretation of imaging results complicated and unreliable (5, 112).

In attempts to overcome these drawbacks, chemical reactions that are specific for H₂O₂ have been sought as the underlying basis for fluorescence-based H₂O₂ probes. One example is the deprotection of boronate esters through oxidation by H₂O₂. Peroxyfluor-1 (PF1) and peroxyresorufin-1 (PR1) contain boronate ester groups, which can be converted to phenols by H₂O₂ to produce the highly fluorescent molecules, fluorescein (λ_abs = 488 nm, λ_em = 520 nm) and resorufin (λ_abs = 571 nm, λ_em = 585 nm), respectively (82) (Fig. 3A). PF1 and PR1 show a large fluorescence increment (>1000-fold) with reaction rate constants k = 0.54 M⁻¹·s⁻¹, and 1.0 M⁻¹·s⁻¹, respectively. The response of PF1 is reported to be selective for H₂O₂, with relatively small fluorescence increments observed for ^•NO treatment in vitro. Peroxy Green 1 (PG1), a derivative based on TokyoGreen (124), has also been applied for fluorescence imaging of endogenous H₂O₂ produced by A431 cells and cultured rat hippocampal neurons under epidermal growth factor (EGF) stimulation (83). Similarly, a wide variety of fluorophores have been protected with boronates; for example, rhodol-based derivatives, Peroxy Yellow 1 (PY1) and Peroxy Orange 1 (PO1), were developed for multi-color imaging, generating products that emit yellow (λ_abs = 519 nm, λ_em = 548 nm) and orange fluorescence (λ_abs = 540 nm, λ_em = 565 nm), respectively (21). PY1 and PO1 have been applied to live-cell imaging of endogenous H₂O₂ in multiple types of cells, including RAW264.7 macrophages stimulated by phorbol 12-myristate 13-acetate (PMA), and EGF-stimulated A431 cells. A major limitation of boronate-based probes is the slow reaction rate with H₂O₂ (k = 0.1–1 M⁻¹·s⁻¹). This slow reaction rate results in low sensitivity under physiological conditions where H₂O₂ is rapidly removed by antioxidant enzymes such as Prxs (132). Another limitation is that boronate ester-based probes have a fluorescence response to ONOO⁻ as well. Multiple reports show the application of boronate-based probes to the detection of ONOO⁻ (114, 137, 143); thus, control experiments with ONOO⁻ scavengers are required to differentiate H₂O₂ and ONOO⁻ when using boronate-based probes.

FIG. 3.

Fluorescent probes for H₂O₂. (A) Boronate-protected fluorophores are deprotected by H₂O₂. (B) The benzil group is cleaved by H₂O₂ to form highly fluorescent 5-carboxyfluorescein. (C) HyPer enables ratiometric fluorescent measurements of H₂O₂ in situ. (D) Orp1 forms a disulfide bond on reaction with H₂O₂, which is relayed to the fused roGFP2, leading to a fluorescence change. H₂O₂, hydrogen peroxide; roGFP2, redox-sensitive green fluorescent protein 2. Color images are available online.

Benzil chemistry can also be used as a design strategy for H₂O₂ probes. Benzil (α-dibenzoyl) has been reported to generate benzoic acid by H₂O₂ in alkaline conditions through an acyclic (Baeyer-Villiger) mechanism (109). To exploit this reaction for H₂O₂ sensing, benzil groups were appended to fluorescein to generate 5-benzoylcarbonylfluorescein derivatives (3) (Fig. 3B). The benzil group acts as an H₂O₂-reactive group while also quenching fluorescence through a donor-excited photoinduced electron transfer (d-PeT) mechanism (123, 124). In the d-PeT process, an electron from the excited fluorophore transfers to the lowest unoccupied molecular orbital of the benzil group, which results in non-radiative relaxation of the excitation energy (123). A derivative with a nitro group on the benzil moiety, 5-(4-nitrobenzoyl)carbonylfluorescein (NBzF), displays a 150-fold fluorescence increase (λ_abs = 495 nm, λ_em = 519 nm) in the presence of H₂O₂ with a reaction rate constant k = 3.2 M⁻¹·s⁻¹. In addition to H₂O₂, NBzF shows a relatively small fluorescence increment in the presence of t-butyl hydroperoxide (t-BuOOH) and ONOO⁻, but it does not respond to ^•NO in vitro. NBzF was applied for live-cell imaging of endogenous H₂O₂ in PMA-stimulated RAW264.7 cells and EGF-stimulated A431 cells. However, NBzF has limitations that are similar to boronate-based probes, including the relatively slow reaction rate with H₂O₂ and the necessity of control experiments with ROS scavengers.

The enzyme-mediated reaction of a small molecule with H₂O₂ can provide a robust and sensitive strategy for H₂O₂ detection. For example, Amplex Red is oxidized by H₂O₂ in the presence of horseradish peroxidase (HRP) to produce the fluorescent product, resorufin (140). This Amplex Red assay is a reliable method to detect extracellular H₂O₂ with a limit of detection of ∼5 pmol. However, HRP is inactivated in the highly reducing cytosolic environment, likely due to the reduction of functionally essential disulfide bonds on HRP (43), rendering Amplex Red not applicable to cytosolic H₂O₂ detection.

Due to the distinct physiological roles of H₂O₂ in different subcellular locations, it is important to visualize H₂O₂ production in a particular organelle. To this end, organelle-targeted probes for H₂O₂ have been developed. One strategy to target small-molecule-based probes to a particular organelle is to utilize organelle-directing groups. Mitochondria peroxy yellow 1 (MitoPY1) is a boronate-based probe conjugated with a TPP group, which is well known to localize in mitochondria due to its positive charge (20). MitoPY1 successfully detected exogenously added H₂O₂ in the mitochondria. Another strategy for directing probes to distinct organelles is through the use of protein tags such as the SNAP-tag. The SNAP-tag is an engineered human DNA repair protein, human O⁶-alkylguanine-DNA alkyltransferase (hAGT), which covalently binds to small molecules with an O⁶-benzylguanine group (51). Boronate-based probes are conjugated with SNAP-tag substrates (SNAP PG1 and PG2) (115) and directed to SNAP-tagged proteins with distinct subcellular localizations. This approach has been used to target boronate probes to the cytosol, nucleus, the extracellular face of the plasma membrane, as well as the endoplasmic reticulum and mitochondrial inner membrane, for exogenous H₂O₂ detection in each specific subcellular location. Similarly, organelle-specific imaging of endogenous H₂O₂ has been achieved. An organelle-targeted benzil-based probe, NBzF-BG, which has an O⁶-benzylguanine group conjugated with NBzF, has been applied to detect endogenous H₂O₂ produced during phagocytosis (2). NBzF-BG has been presented on the extracellular side of the plasma membrane of RAW264.7 macrophages, localizing NBzF-BG on the inside of the phagosomal membrane on phagocytosis of opsonized microbeads. NBzF-BG localized inside phagosomes has enabled the visualization of H₂O₂ production in phagosomes.

Genetically encoded fluorescence probes for H₂O₂

Genetically encoded fluorescent protein-based H₂O₂ probes, such as HyPer and redox-sensitive green fluorescent protein 2 (roGFP2)-Orp1, are powerful imaging tools with several advantages such as high sensitivity, ratiometric measurements, and ability to localize to a particular organelle. HyPer consists of a circularly permutated yellow fluorescent protein (cpYFP) inserted into the regulatory domain of OxyR, a prokaryotic H₂O₂ sensor (11) (Fig. 3C). H₂O₂ oxidizes Cys199 of the OxyR regulatory domain to form a sulfenic acid, followed by disulfide-bond formation with Cys208. The disulfide formation changes the conformation and excitation properties of HyPer, which enables ratiometric measurements with two excitation wavelengths (488/405 nm). The fluorescence response of HyPer is relatively sensitive to H₂O₂ concentrations, and HyPer is not responsive to other ROS including O₂^•−, ^•NO, and ONOO⁻, and oxidized glutathione (GSSG). Several variants of HyPer have been developed with larger signal enhancement (HyPer-2 and HyPer-3) (13, 77) and different excitation and emission wavelengths (HyPer-Red) (27). The fast reaction rates (k = 1.2–5.0 × 10⁵ M⁻¹·s⁻¹) with H₂O₂ and the ratiometric response confer HyPer probes that are suitable for various biological applications. HyPer has been applied for live-cell imaging in bacterial and mammalian cells, and it has also been targeted to organelles such as the cytoplasm, mitochondria, nucleus, endoplasmic reticulum, peroxisomes, and plasma membrane (12). The HyPer probes have also been applied to imaging of whole animals, including Caenorhabditis elegans (55) and zebrafish (88). A limitation of HyPer probes is that the fluorescence signal of HyPer can be affected by pH; thus, control experiments with pH sensors such as SypHer (79) are required to interpret HyPer-based imaging results.

Another strategy for genetically encoded H₂O₂ probes is based on roGFPs (25, 41) coupled with a yeast peroxidase Orp1 (38). Orp1 is oxidized to form a sulfenic acid by H₂O₂ at the catalytic Cys36, which rapidly forms an intramolecular disulfide bridge with the resolving Cys82. When fused to roGFP2, the disulfide bridge of Orp1 is efficiently relayed to roGFP2, which results in a ratiometric fluorescence signal change (390/480 nm) (Fig. 3D). The response of roGFP2-Orp1 is specific for H₂O₂ among oxidants such as GSSG, cystine, and dehydroascorbic acid (38). One advantage of roGFP2-Orp1 over HyPer is insensitivity to pH changes within the physiological range. Applications of roGFP2-Orp1 include the visualization of physiological H₂O₂ production in Drosophila larvae and tissues (6, 8, 122). Although roGFP2-Orp1 and HyPer have been successfully applied to H₂O₂ imaging in vivo, reaction rates of their sensing moiety are still >100-fold slower than that of typical 2-Cys Prxs (k = ∼10⁷ M⁻¹·s⁻¹), which are abundant in the cytosol. Recently, improved roGPF2-based H₂O₂ probes, roGFP2-Tsa2ΔC_R and roGFP2-Tsa2ΔC_PΔC_R, have been reported (84). These probes utilize a typical 2-Cys Prx Tsa2, with mutations at the peroxidatic cysteine (C_P) and/or the resolving cysteine (C_R), as an H₂O₂-sensitive moiety. These probes are ∼50% oxidized under unstressed conditions, which allows for dynamic monitoring of both increases and decreases in basal H₂O₂. By using the roGFP2-Tsa2ΔC_R probe, it has been shown that cellular growth rate correlates with basal mitochondrial H₂O₂ levels in yeast cells.

Chemical Probes for Oxidative PTMs of Cysteine

Cysteine residues have unique chemical reactivity, including high nucleophilicity and redox sensitivity, and play important roles in many protein functions, such as nucleophilic and redox catalysis, structural-disulfide formation, and binding to metal ions (91). Oxidative PTMs of cysteine play central roles in redox signaling by regulating the activity of key proteins within signaling pathways. One well-characterized example of cysteine-mediated redox signaling is in EGF signaling. In A431 epidermoid carcinoma cells, PTP1B is inhibited (63) and EGFR is activated (93) through S-sulfenylation, resulting in augmentation of the phosphorylation signals resulting from EGF binding to EGFR. Therefore, methods for identifying and quantifying oxidative cysteine PTMs are critical to understanding redox signaling. Numerous chemical probes with a variety of reactive handles have been developed and applied to globally monitor oxidative cysteine PTMs.

Chemical probes for oxidative cysteine PTMs can be categorized into two groups: (i) cysteine-reactive probes to visualize general cysteine PTMs and (ii) probes that are specific for a particular oxidative PTM of cysteine. Probes in the first category are based on monitoring the intrinsic cysteine reactivity with electrophilic chemical probes that form stable covalent adducts with reduced cysteines. Oxidation of a particular cysteine residue is reflected by a loss in nucleophilicity and subsequent decrease in probe labeling. These methods allow for identification of any cysteine PTM that reduces nucleophilicity, and therefore can be applied to all oxidative cysteine modifications. However, secondary methods need to be employed to identify the exact modification that is causing the measured loss in cysteine reactivity. In the second category, PTM-specific reactions are harnessed to design chemical probes that are specific for each oxidative cysteine PTM, including S-sulfenylation, S-sulfinylation, and S-glutathionylation. Contrary to the general cysteine-reactive probes, PTM-specific probes can focus on a particular PTM, lower the complexity of proteome samples, and enhance sensitivity and reliability of analysis. However, probes to directly monitor inter- and intra-molecular disulfides, and sulfonic acids, are currently lacking, rendering these PTMs intractable to existing PTM-specific methods. To date, both general and specific methods rely on two-dimensional electrophoresis and MS-based analytical methods for identification and quantification of labeled proteins.

Chemical-Proteomic Methods for General Oxidative PTMs of Cysteine

Two-dimensional gel electrophoresis

Two-dimensional gel electrophoresis (2DE)-based methods have been commonly used in proteomic research. 2DE methods are also frequently applied to the analysis of oxidative cysteine PTMs for visualization with radioactive or fluorescent reporters (17). In typical procedures for autoradiography, non-oxidized cysteine residues are capped by iodoacetamide (IAM) or N-ethylmaleimide (NEM). Then, oxidized cysteines are reduced by reducing agents such as dithiothreitol (DTT) or tris(2-carboxyethyl)phosphine (TCEP). Subsequently, the newly reduced cysteine residues are labeled with radioactive ¹⁴C-IAM or ¹⁴C-NEM for visualization by autoradiography (61, 65). Due to non-specific reduction by DTT or TCEP (34), these methods visualize multiple forms of oxidative cysteine PTMs, including disulfide bonds, sulfenic acids, and nitrosothiols. To achieve more facile visualization of 2D gels, radioactive probes have been replaced by fluorescent cysteine-reactive probes such as 5-iodoacetamidefluorescein (10), monobromobimane (136), and boron dipyrromethene (BODIPY) FL C1-IA (42). The signal intensity of radiation or fluorescence from proteins can be normalized by the protein abundance estimated from Coomassie Brilliant Blue (CBB) staining to provide a qualitative estimate of the stoichiometry of oxidation (42, 65).

2DE-based methods can be applied to directly compare the degree of cysteine oxidation between two samples. In a method termed redox differential in-gel electrophoresis (redox-DIGE), a pair of cysteine-reactive probes conjugated with orthogonal fluorophores such as Cy3-maleimide and carbocyanine 5 (Cy5)-maleimide are used for labeling two samples, respectively (16, 30, 46). After blocking reduced cysteines, oxidized cysteines in two distinct cell lysates are labeled with two different fluorophores. The two labeled samples are combined and analyzed on the same 2D gel, enabling direct visualization of differences in oxidative cysteine PTMs across two samples. Redox-DIGE has been applied to identify mitochondrial proteins that are sensitive to oxidative modifications by endogenous ROS (46). However, this redox-DIGE analysis lacks information about protein abundance; thus, fluorescence signals cannot be normalized with the protein amounts. To overcome this limitation, another differential labeling strategy was developed, whereby both oxidized and reduced cysteine residues are differentially labeled with two distinct fluorophores (62). A ratio between the two fluorescence signals gives a quantitative measure of the stoichiometry of oxidation by normalizing for protein abundance.

The 2DE-based methods described earlier commonly have major drawbacks. The limited resolution of 2DE results in insufficient separation of proteins with similar physicochemical properties. Further, limited detection sensitivity of gel-imaging methods prohibits the detection of low abundant proteins. In addition, even if protein spots are found to be oxidized by 2DE, the identification of each protein has to be performed individually by MS. Time-consuming 2DE procedures and subsequent individual identification of each protein by MS make 2DE methods low-throughput. Further, given that a protein has multiple cysteines, a signal of the protein on 2DE can represent an average of multiple cysteine residues, and identifying the exact cysteine(s) that are oxidized can be challenging.

Isotope-coded affinity tags

To overcome the multiple drawbacks of 2DE, a method based on isotope-coded affinity tags (ICAT) has been developed. The ICAT method utilizes biotinylated IAM derivatives that are labeled with isotopically light and heavy linkers (39). Peptides tagged with the ICAT probes were enriched by streptavidin beads and analyzed by quantitative MS (Fig. 4A). Isotopically labeled ICAT probes enable relative quantification of oxidized cysteines between the light- and heavy-labeled samples. The ICAT strategy was applied to identify cysteines that are susceptible to oxidation on H₂O₂ treatment (111). A modified ICAT strategy termed OxICAT (oxidative isotope-coded affinity tag) has been developed for differential labeling of oxidized and reduced cysteine in the same sample (64). In this OxICAT strategy, reduced cysteines are labeled with a light ICAT reagent, and oxidized cysteine residues are reduced with TCEP and labeled with a heavy ICAT reagent (Fig. 4B). This differential labeling enables quantification of the ratio between reduced and oxidized forms of each cysteine, and this ratio can then be compared across multiple samples. To better monitor protein abundance changes in tandem with oxidation variations, isotopic dimethyl labeling has been coupled with ICAT, enabling protein abundance measurements before cysteine enrichment (32, 33). Another strategy for protein quantification is a method termed GELSILOX in which a stable O₂ isotope is incorporated during the trypsin digestion step (78). These ICAT, OxICAT, and GELSILOX methods overcome major limitations of 2DE by achieving both the identification and quantification of multiple oxidized proteins in a single experiment, and, in addition, provide higher sensitivity than gel-based methods for the detection of low abundance proteins. However, detection sensitivity can be decreased by low ionization efficiency of certain peptides, multiple cysteines on the same peptide can complicate site identifications, and a cysteine cannot be identified if the residue is on a very short or long tryptic peptide.

FIG. 4.

Chemical-proteomic platforms for general cysteine PTMs. (A) Structures of ICAT reagents. (B) OxICAT is a modified ICAT technology for oxidative cysteine PTMs. (C) A quantitative proteomic platform, isoTOP-ABPP, enables comprehensive identification and quantification of cysteine PTMs. (D) Caged electrophilic probes are developed to achieve in situ labeling of cysteine and analysis of cysteine PTMs within a physiological context. CBK1, caged α-bromomethylketone alkyne; IAA, idoacetamide alkyne; ICAT, isotope-coded affinity tags; isoTOP-ABPP, isotopic tandem orthogonal proteolysis activity-based protein profiling; LC-MS/MS, liquid chromatography-tandem mass spectrometry; OxICAT, oxidative isotope-coded affinity tag; TCEP, tris(2-carboxyethyl)phosphine; UV, ultraviolet. Color images are available online.

Isotopic tandem orthogonal proteolysis activity-based protein profiling

ICAT probes have a bulky biotin reporter that may inhibit labeling of sterically hindered cysteines. An alternative MS-based method, termed isotopic tandem orthogonal proteolysis activity-based protein profiling (isoTOP-ABPP), relies on a simpler iodoacetamide alkyne (IAA) probe (131). In isoTOP-ABPP, proteome samples are labeled with IAA, before incorporation of a biotin-azide tag via copper(I)-catalyzed alkyne-azide cycloaddition (CuAAC) (Fig. 4C). The biotin moiety facilitates subsequent enrichment of labeled proteins by streptavidin beads, after which proteins then undergo an on-bead tryptic digestion. The biotin-azide tag has cleavable linkers that are susceptible to cleavage by treatment with tobacco etch virus protease or sodium dithionite (98, 131), which releases the enriched peptides for MS analysis. By using isotopically light- or heavy-labeled biotin-azide tags, relative quantification across two samples is achieved. An isoTOP-ABPP ratio, R, is calculated from signal intensities of light- and heavy-tagged peptides for each identified cysteine. The R value provides a quantitative measure of changes in cysteine reactivity, and the isoTOP-ABPP method can identify and quantify cysteine reactivity changes of ∼1000 to 1500 cysteine residues. The isoTOP-ABPP method and its derivatives have been applied for identification of bacterial proteins that are susceptible to oxidation by H₂O₂ (19). This method was also applied to identify S-nitrosation sites on numerous proteins on treatment with transnitrosation donors such as S-nitrosoglutathione (GSNO), identifying S-nitrosation sites on proteins including 3-hydroxyacyl-CoA dehydrogenase type 2 (HADH2) and cathepsin D (CTSD) (141).

Although isoTOP-ABPP can quantify and identify more than 1000 cysteines in a single tandem liquid chromatography-tandem mass spectrometry (LC/LC-MS/MS) run, coverage of the whole proteome by this method can still be limited due to the high complexity and heterogeneity of the proteome samples. Organelle isolation strategies can be coupled with isoTOP-ABPP to improve coverage of reactive cysteines in a particular organelle such as mitochondria (7). Another limitation of isoTOP-ABPP has been the usage of cytotoxic IAA probe, which is not suitable for live-cell applications. Thus, applications of isoTOP-ABPP have been primarily performed in lysate samples. However, because oxidative PTMs of cysteine are highly labile to environmental redox changes caused by cell lysis, it is ideal to perform the initial labeling step directly in living cells. Photolabile caged electrophilic probes have been developed to achieve cysteine labeling in situ (1, 4). A caged electrophile named caged α-bromomethylketone alkyne, or CBK1, is not cysteine reactive and, therefore, can be loaded into live cells with negligible cytotoxicity (Fig. 4D). CBK1 is activated to be cysteine reactive by ultraviolet (UV) light irradiation in situ to trigger cysteine labeling, and the labeled cysteine residues are identified by LC/LC-MS/MS. An additional advantage of this labeling strategy with caged probes is the high temporal resolution of cysteine labeling, which is required for accurate analysis of transient and reversible cysteine oxidations in redox signaling. The practical usability of CBK1 has been demonstrated by analyzing redox signaling in A431 epidermoid carcinoma cells stimulated by EGF. CBK1 coupled with isoTOP-ABPP identified hundreds of cysteine residues with R values, indicating the degree of oxidative modification for each cysteine under EGF stimulation. More recently, an optimized caged probe, caged α-iodoketone alkyne 4 (CIK4), has been reported with improved cysteine reactivity and uncaging efficiency, which enables more efficient detection of cysteine residues by LC/LC-MS/MS (1). Though caged electrophilic probes enable cysteine labeling in situ with short exposure time to cytotoxic agents and high temporal resolution, the limitation is the usage of UV light, which may cause cytotoxic effects on live cells.

Chemical Proteomic Methods for a Specific Oxidative PTM of Cysteine

Chemical probes for sulfenic acids

The 2DE, ICAT, and isoTOP-ABPP methods discussed earlier all rely on changes to the intrinsic reactivity of a cysteine induced by oxidation, and, therefore, do not differentiate cysteine PTMs such as sulfenic acids, sulfinic acids, and disulfides. Complementary strategies focus on a specific cysteine PTM by utilizing chemical probes that selectively react with a single PTM. The reversibility of sulfenic acids renders this PTM the ability to mediate cellular signal transduction events that are akin to phosphorylation and acetylation. One strategy for the specific labeling of sulfenylation is based on the selective reduction of sulfenic acids by arsenite, which is analogous to the differential tagging strategies described earlier (108). In this method, sulfenic acids are reduced by arsenite under denaturing conditions, then labeled with a biotin-maleimide probe. Another strategy is the direct labeling with chemical probes that are specific for sulfenic acids, including 5, 5-dimethyl-1, 3-cyclohexanedione, also known as dimedone, as a reactive moiety (Fig. 5A). The enolate form of dimedone facilitates nucleophilic attack onto sulfenic acids to form a covalent adduct. Dimedone or other 1,3-diketone compounds are conjugated with fluorophores or biotin analogues for in-gel visualization and enrichment with avidin for MS-based proteomic experiments (37, 87, 95, 96).

FIG. 5.

Chemical probes for sulfenic acids and sulfinic acids. (A) The reaction between sulfenic acids and dimedone. (B) Structures of dimedone-based probes. (C) A strained alkyne-based probe, BCN, can be conjugated to sulfenic acids. (D) The reaction between sulfinic acids and 2-nitroso benzoic acid derivatives. (E) NO-Bio is a nitroso-based chemical probe for sulfinic acids. BCN, bicycle[6.1.0]nonyne; DAz, azide-tagged dimedone; DYn, alkyne-tagged dimedone; NO-Bio, biotinylated 2-nitroso terepththalic acid. Color images are available online.

The presence of bulky reporter groups such as fluorophores and biotin analogues are not ideal because they result in low labeling efficiency by blocking access to buried sulfenic acids, as well as limiting the plasma membrane permeability of the probes. Sulfenic acids are highly labile to redox changes, and, therefore, labeling in situ is important for accurate analysis. To this end, dimedone derivatives with minimal modifications have been developed; azide-tagged probes (azide-tagged dimedone [DAz]-1 and DAz-2) (66, 100) and alkyne-tagged probes (alkyne-tagged dimedone [DYn]-1 and DYn-2) (93) have chemical handles for incorporation of reporters via a Staudinger reaction or CuAAC in subsequent steps (Fig. 5B). These probes are membrane permeable and can be applied to the labeling of sulfenic acids directly in living cells.

A strategy for differential labeling of sulfenic acids has been developed by utilizing isotopically coded dimedone- and iododimedone-bearing probes, which are reactive with sulfenic acids and reduced cysteines, respectively (110). Based on this strategy, the degree of sulfenylation at a particular cysteine residue can be accurately quantified. In a similar manner, dimedone and IAM conjugated with lantanide-1,4,7,10-tetraazacyclododecane-1,4,7,10-tetraacetic acid (DOTA) have been applied to quantify sulfenylation using inductively coupled plasma-MS (26).

Dimedone-based probes have been applied to various biological systems to uncover sulfenylation sites. For example, a biotinylated dimedone probe has been utilized to reveal that Cys124 of protein kinase B β (Akt2) is sulfenylated under physiologically relevant platelet-derived growth factor (PDGF) stimulation (129). The sulfenylation of Cys124 inhibits the activity of Akt2, showing a novel regulatory mechanism for kinase regulation in addition to phosphorylation. As another example, a biotinylated dimedone probe has been applied to T cells to identify increased sulfenylation on protein tyrosine phosphatases such as Src homology domain 2-containing protein tyrosine phosphatase (SHP)-1 and SHP-2, and actin on T cell activation (81). This study demonstrated that reversible sulfenylation is an important regulatory factor for proliferation and function of cluster of differentiation 8 (CD8⁺) T cells. An alkyne-functionalized dimedone probe, DYn-2, revealed that Cys797 of EGFR is sulfenylated under EGF stimulation and sulfenylation augments the kinase activity of EGFR (93). Lastly, isotopically light- and heavy-labeled DYn-2 probes were coupled to advanced LC-MS/MS-based methods to globally identify and quantify the exact sites of sulfenylation within proteomes (134).

Although dimedone-based probes have been successfully applied to proteomic experiments, one limitation of dimedone is the demonstrated slow reaction rate with sulfenic acids, which requires the use of the chemical probes at high concentrations to achieve efficient labeling. To overcome this limitation, a library of ∼100 cyclic C-nucleophiles have been evaluated for reactivity with sulfenic acids to find several novel compounds with more than 200-fold enhanced reactivity than that of dimedone (36). Strained bicycle[6.1.0]nonyne (BCN) derivatives were also reported as a new reactive group with sulfenic acids (Fig. 5C). The BCN derivatives can react with sulfenic acids with a faster reaction rate than dimedone by over two orders of magnitude (97). It is noteworthy that BCN is reported to undergo thiol-yne addition and can be non-selectively consumed by thiols (121). These new chemistries can offer an alternative strategy for monitoring sulfenylation in addition to the widely used dimedone-based probes.

Chemical probes for sulfinic acids

S-sulfinylation (RSO₂H) was regarded an artifactual hyper-oxidation of cysteine that occurs during sample handling, but increasing evidence indicates that S-sulfinylation is not only physiologically relevant but also important in redox signaling (47, 71). It was revealed that ∼5% of cysteine residues are oxidized to sulfinic acids in a soluble fraction of rat liver (40). Among S-sulfinylated proteins are Prxs, which are a family of H₂O₂-scavanging enzymes. Due to high abundance in the cytosol and very fast reaction rates with H₂O₂ (k = ∼10⁷ M⁻¹·s⁻¹), Prxs remove H₂O₂ very rapidly to keep the concentration of cytosolic H₂O₂ relatively low (132). Therefore, it has been proposed that Prxs activity may be locally regulated under redox signaling through PTMs such as S-sulfinylation (135) and phosphorylation (133). Although sulfinic acids are not reduced by most biological antioxidants, the S-sulfinylation on Prxs can be reduced through an ATP-dependent reduction by Srx (14).

To capture sulfinic acids in a specific manner, the reported reaction between 2-nitroso benzoic acid derivatives and sulfinic acids has been adapted (72) (Fig. 5D). The reactive nitroso warhead covalently modifies sulfinic acids to form stable nitroso-benzene derivatives. This ligation reaction is shown to be selective to sulfinic acids among various nucleophilic and/or redox-active small molecules. An observed side reaction involves the oxidation of cysteine to cystine concomitant with the reduction of nitroso compounds. A nitroso compound, 2-nitroso terepththalic acid (NO-Ph), was shown to successfully label a sulfinic acid on a recombinant Gpx3 mutant; however, NO-Ph also formed a stable sulfonamide adduct with a reduced cysteine on Gpx3. Thus, the blocking of reduced cysteine with NEM before the labeling of sulfinic acids is required for efficient MS analysis. The reaction termed sulfinic-acid-nitroso ligation (SNL) has been utilized to design a sulfinic acid-specific probe, biotinylated 2-nitroso terepththalic acid (NO-Bio), in which a 2-nitroso benzoic acid derivative is conjugated with a biotin analogue (73) (Fig. 5E). The NO-Bio probe has been applied to the labeling of sulfinic acids on recombinant Parkinson disease protein 7 (DJ-1) oxidized by H₂O₂ and in cell lysates induced by 2,2′-dipyridyl disulfide (DPS), and to profile oxidation among normal and matched tumor cells. The SNL reaction can also be utilized to monitor S-nitrosation (76). Biotinylated probes with sulfinic acid (Biotin-SO₂H) and GSNO (Biotin-GSNO) have been applied to quantitative MS using stable isotope labeling with amino acids in cell culture (SILAC), revealing relative occupancy ratios of S-sulfinylation and S-nitrosation in 293T cells. The SNL chemistry has not yet been applied to a wide variety of biological samples; however, it opens the door to proteome-wide analyses focusing on S-sulfinylation and S-nitrosation.

Chemical probes for S-glutathionylation

S-glutathionylation is known to regulate important protein functions that are involved in various physiological processes such as gene expression, signaling, energy metabolism, and cell survival (92). For example, glutathionylation of nuclear factor-κB (NF-κB) inhibits DNA-binding activity and downstream transcriptional activation (94). Another key protein in this pathway, inhibitor of κB kinase (IKK), is also glutathionylated, rendering the NF-κB pathway highly susceptible to redox changes in the cellular environment (101). The extent of modification with glutathione depends on the ratio of reduced (GSH) and oxidized forms (GSSG) of glutathione.

Various chemical-proteomic strategies have been developed to analyze S-glutathionylation within a biological context. In early methods based on autoradiography, GSH tagged with ³H or ³⁵S was utilized for labeling in vitro, and ³⁵S-cysteine was applied for metabolic incorporation in situ (31). GSH is synthesized from glutamate, cysteine, and glycine through two subsequent ATP-dependent reactions catalyzed by γ-glutamylcysteine synthetase and glutathione synthetase (GS) (80). The method for metabolic incorporation of ³⁵S-cysteine requires treatment with cycloheximide to inhibit incorporation of ³⁵S-cysteine into proteins, ensuring that ³⁵S-cysteine is only incorporated into GSH. Proteome samples modified with ³⁵S-labeled GSH are analyzed by sodium dodecyl sulfate polyacrylamide gel electrophoresis and imaged by autoradiography. This method was applied to monitor S-glutathionylation in human monocytes (104), revealing that glyceraldehyde-3-phosphate dehydrogenase (GAPDH) is S-glutathionylated during a respiratory burst (99). However, this method has major drawbacks, including the necessity to inhibit protein synthesis, which greatly affects cellular metabolism, and the lack of a handle to enrich modified proteins for global MS identification.

To overcome these drawbacks of radioactive GSHs, biotinylated GSH (BioGSH) analogues have been developed. GSH and GSSG can be conjugated with a biotin group at the primary amine by treatment with sulfosuccinimidyl-6-(biotinamido)-hexanoate (sulfo-NHS-biotin) (15). A membrane-permeable precursor of GSH, glutathione ethyl ester (GEE), can also be modified by sulfo-NHS-biotin to form biotinylated glutathione ethyl ester (BioGEE) (117) (Fig. 6A). The biotin groups can be utilized for enrichment of glutathionylated proteins by streptavidin beads, followed by reductive cleavage of the enriched proteins from the beads and analysis by gel electrophoresis and MS. Non-reducing Western blotting enables sensitive detection of glutathionylated proteins in combination with streptavidin-HRP or anti-biotin antibodies. These BioGSH derivatives have been applied to the identification of glutathionylation in vitro (45, 127) and in cells. BioGEE was applied to live HeLa cells under oxidative stress by exogenous H₂O₂ or tumor necrosis factor-α (TNF-α) treatment to identify annexin A2 and Trx peroxidase 2 as glutathionylated proteins (117). Limitations of this method include the modification with a bulky biotin group that may perturb a subset of proteins tagged with these reagents, and the exogenous addition of redox-active GSH derivatives that affect the native GSH/GSSG ratio.

FIG. 6.

Chemical probes for S-glutathionylation. (A) Structures of biotinylated glutathione derivatives, BioGSH and BioGEE. (B) Glycine surrogates with bioorthogonal handles for metabolic incorporation. (C) The glycine surrogates can be incorporated into GSH catalyzed by mutated GSs in living cells, which enables proteomic analyses of native glutathionylation. BioGEE, biotinylated glutathione ethyl ester; BioGSH, biotinylated GSH; CuAAC, copper(I)-catalyzed alkyne-azide cycloaddition; GS, glutathione synthetase; GSH, reduced glutathione. Color images are available online.

An elegant method to overcome the limitations of BioGSH derivatives has been developed by utilizing a mutated GS that catalyzes coupling of an azide-tagged alanine derivative (azido-Ala) to γ-glutamylcysteine (γGlu-Cys) (106) (Fig. 6B, C). The GSH derivative containing an azide handle (azide-GSH) for CuAAC allows for identification of glutathionylated proteins by gel and MS analysis. The strategy was utilized for analysis of glutathionylation in living cells under exogenous H₂O₂ stimulation and glucose depletion (107). Alkene-bearing Gly surrogates (ally-containing glycine [Allyl-Gly], allyl-containing serine [Allyl-Ser], and cyclopropane-containing serine [Cyp-Ser]) can also be enzymatically incorporated into GSH by mutated GSs, which enable bioorthogonal tagging by tetrazine-based reporters (50) (Fig. 6B).

Selective reduction of glutathione-containing disulfides mediated by Grx has been applied to proteomic analysis of glutathionylation (68). In the Grx-based method, on blocking non-oxidized cysteine residues with NEM, glutathione-containing disulfide bonds are reduced by Grx. Freshly formed cysteines are labeled with a biotin-tagged NEM probe, resulting in subsequent enrichment of labeled proteins. The Grx-based method has been applied to analyze glutathionylation under oxidative stress induced by diamide (69).

Summary

Redox signaling is involved in various physiological processes such as antioxidant induction, cellular proliferation, and migration. Due to the highly reactive and unstable nature of ROS, it has been challenging to measure physiological ROS generation in a reliable and quantitative manner. Taking advantage of the distinct reactivities of different ROS, chemical probes have been developed and applied to detect and quantify biological ROS production. To date, various detection methods for O₂^•− are available and widely used with EPR, fluorescence, and chemiluminescence instrumentation. Fluorescence probes for H₂O₂ have been designed by utilizing H₂O₂-specific chemical reactions and engineered H₂O₂-sensitive proteins. Despite the low reactivity of H₂O₂, fluorescent protein-based H₂O₂ probes are highly sensitive to H₂O₂. Currently available ROS probes have several advantages and limitations in terms of sensitivity, selectivity, and usability; thus, it is important to choose the appropriate methods for a given application. Control experiments with ROS scavengers and a combination of multiple ROS-detection methods can increase the reliability of the measurements. Besides the detection of ROS generation, proteome-wide analysis of oxidative cysteine PTMs is also critical to uncover the mechanisms and downstream consequences of redox signaling. Increased evidence shows that cysteine PTMs such as sulfenylation, sulfinylation, and glutathionylation can transiently regulate protein activity with critical downstream consequences. The dynamic nature of oxidative PTMs of cysteine has made it a challenge to perform comprehensive and quantitative analysis of these PTMs within native biological environments. Recent advances in chemical probes for in situ labeling of cysteine and its PTMs will provide facile and reliable approaches to capture native PTMs within a physiological context. In parallel, advances in MS instrumentation and quantitative proteomic strategies will enhance the efficiency of identification and quantification of cysteine PTMs. Analytical platforms for ROS and cysteine PTMs highlighted in this review are complementary to each other, and their combined use will offer a robust and comprehensive strategy to dissect redox signaling. The detection of ROS and oxidative cysteine PTMs will benefit from analytical advances that can improve the sensitivity of fluorescence and MS-based analyses, and further, methods to provide spatial and temporal control over measurements will be instrumental in furthering our understanding of redox signaling.

Abbreviations Used

2DE

two-dimensional gel electrophoresis

2-OH-E⁺

2-hydroxyethidium

Akt2

protein kinase B β

ATP

adenosine triphosphate

BCN

bicycle[6.1.0]nonyne

BES

benzenesulfonyl fluorescein

BioGEE

biotinylated glutathione ethyl ester

BioGSH

biotinylated GSH

CBK1

caged α-bromomethylketone alkyne

CLA

2-Methyl-6-phenylimidazo[1,2-a]pyrazin-3(7H)-one

C_P

peroxidatic cysteine

CPH

1-hydroxy-3-carboxy-2, 2, 5-tetramethyl-pyrrolidine hydrochloride

C_R

resolving cysteine

CuAAC

copper(I)-catalyzed alkyne-azide cycloaddition

Cy3

carbocyanine 3

Cys

cysteine

DAz

azide-tagged dimedone

DCFH₂

dihydrodichlorofluorescein

DMPO

5,5-dimethyl-1-pyrroline-N-oxide

d-PeT

donor-excited photoinduced electron transfer

DTT

dithiothreitol

DYn

alkyne-tagged dimedone

E⁺

ethidium

EGF

epidermal growth factor

EGFR

epidermal growth factor receptor

EPR

electron paramagnetic resonance

Grx

glutaredoxin

GS

glutathione synthetase

GSH

reduced glutathione

GSNO

S-nitrosoglutathione

GSSG

oxidized glutathione

H₂O₂

hydrogen peroxide

HE

hydroethidine

HRP

horseradish peroxidase

IAA

iodoacetamide alkyne

IAM

iodoacetamide

ICAT

isotope-coded affinity tag

isoTOP-ABPP

isotopic tandem orthogonal proteolysis activity-based protein profiling

LC/LC-MS/MS

tandem liquid chromatography-tandem mass spectrometry

MCLA

6-(4-methoxyphenyl)-2-methyl-imidazo[1,2-a]pyrazin-3(7H)-one

MitoPY1

mitochondria peroxy yellow 1

MS

mass spectrometry

NADPH

reduced nicotinamide adenine dinucleotide phosphate

NBzF

5-(4-nitrobenzoyl)carbonylfluorescein

NBzF-BG

O⁶-benzylguanine-conjugated NBzF

NEM

N-ethylmaleimide

NF-κB

nuclear factor-κB

^•NO

nitric oxide

NO-Bio

biotinylated 2-nitroso terepththalic acid

NO-Ph

2-nitroso terepththalic acid

NOX

NADPH oxidase

O₂

oxygen

O₂^•−

superoxide

⁻OCl

hypochlorite

^•OH

hydroxyl radical

ONOO⁻

peroxynitrite

OxICAT

oxidative isotope-coded affinity tag

PF1

Peroxyfluor-1

PG1

Peroxy Green 1

PMA

phorbol 12-myristate 13-acetate

PO1

Peroxy Orange 1

PR1

Peroxyresorufin-1

Prx

peroxiredoxin

PTM

post-translational modification

PTP1B

protein tyrosine phosphatase 1B

PY1

Peroxy Yellow 1

redox-DIGE

redox differential in-gel electrophoresis

RNS

reactive nitrogen species

roGFP

redox-sensitive green fluorescent protein

ROS

reactive oxygen species

SHP

Src homology domain 2-containing protein tyrosine phosphatase

SNL

sulfinic-acid-nitroso ligation

SOD

superoxide dismutase

Srx

sulfiredoxin

Sulfo-NHS-biotin

sulfosuccinimidyl-6-(biotinamido)-hexanoate

TAM

triarylmethyl

TCEP

tris(2-carboxyethyl)phosphine

TPP

triphenylphosphonium

Trx

thioredoxin

UV

ultraviolet