
==== Front
Redox Biol
Redox Biol
Redox Biology
2213-2317
Elsevier

S2213-2317(24)00310-0
10.1016/j.redox.2024.103332
103332
Review Article
Oxidative stress-mediated protein sulfenylation in human diseases: Past, present, and future
Mu Baoquan a1
Zeng Yan a1
Luo Li luoli0812@126.com
bc⁎⁎
Wang Kui kuiwang@scu.edu.cn
a⁎
a West China School of Basic Medical Sciences & Forensic Medicine, State Key Laboratory of Biotherapy, West China Hospital, Sichuan University, Chengdu, 610041, China
b Center for Reproductive Medicine, Department of Gynecology and Obstetrics, West China Second University Hospital, Sichuan University, Chengdu, 610041, China
c Key Laboratory of Birth Defects and Related Diseases of Women and Children (Sichuan University), Ministry of Education, Chengdu, 610041, China
⁎ Corresponding author. kuiwang@scu.edu.cn
⁎⁎ Corresponding author. Center for Reproductive Medicine, Department of Gynecology and Obstetrics, West China Second University Hospital, Sichuan University, Chengdu, 610041, China. luoli0812@126.com
1 These authors contributed equally to this work.

30 8 2024
10 2024
30 8 2024
76 10333224 7 2024
28 8 2024
28 8 2024
© 2024 The Authors
2024
https://creativecommons.org/licenses/by-nc/4.0/ This is an open access article under the CC BY-NC license (http://creativecommons.org/licenses/by-nc/4.0/).
Reactive Oxygen Species (ROS) refer to a variety of derivatives of molecular oxygen that play crucial roles in regulating a wide range of physiological and pathological processes. Excessive ROS levels can cause oxidative stress, leading to cellular damage and even cell demise. However, moderately elevated levels of ROS can mediate the oxidative post-translational modifications (oxPTMs) of redox-sensitive proteins, thereby affecting protein functions and regulating various cellular signaling pathways. Among the oxPTMs, ROS-induced reversible protein sulfenylation represents the initial form of cysteine oxidation for sensing redox signaling. In this review, we will summarize the discovery, chemical formation, and detection approaches of protein sulfenylation. In addition, we will highlight recent findings for the roles of protein sulfenylation in various diseases, including thrombotic disorders, diabetes, cardiovascular diseases, neurodegenerative diseases, and cancer.

Keywords

Reactive oxygen species
Redox signaling
Cysteine oxidation
Sulfenylation
Oxidative stress
==== Body
pmc1 Introduction

ROS, a category of uniquely active molecules, are produced by cells during normal metabolism or when cells are distressed by diseases or toxins [1]. They primarily include hydrogen peroxide (H2O2), superoxide anion (O2•−), hydroxyl radical (•OH), organic peroxides (ROOH), hypohalous acids (HOX), as well as other free radicals and non-radicals. Due to their high biological reactivities, ROS not only react with nucleic acids, proteins, and lipids to cause cellular damage [2], but also function as secondary messengers to influence cell signaling and regulate various physiological and pathological processes [3]. Moreover, the redox balance of ROS contributes to the maintenance of cellular homeostasis [4]. Stress factors such as toxins and pathogens can elevate ROS levels in the humans, leading to the development of various diseases, including cancer, diabetes, cardiovascular diseases, etc. [1]. One of the primary mechanisms by which ROS exert their biological functions is the oxidative post-translational modifications (oxPTMs) of protein cysteine thiols. In a redox proteomics study, it was noted that 5–12% of detectable cysteine sulfhydryls in proteins are oxidized under normal conditions. This ratio can rise to 40% under oxidative conditions [5]. Certainly, redox signaling can also occur through methionine oxidation [6], selenoproteins [7], oxidation of protein metal centers [8] and lipids [9], but these topics will not be discussed here.

Although cysteine constitutes a relatively low percentage of amino acids in proteins, it frequently appears at functional sites [10,11]. Protein cysteine thiols are considered to maintain redox balance due to their high reactivity and reversible oxidation capability [12]. Currently, over 50 oxPTM types of cysteine thiols have been characterized, among which protein sulfenylation plays a crucial role in regulating cell signaling in response to oxidative stress. Specifically, cysteine sulfenylation serves as a precursor for several other oxPTMs, such as sulfinylation, sulfonylation, glutathionylation, or disulfide bond [13]. More importantly, cysteine sulfenylation can reversibly regulate the structure, interaction, and function of proteins either in a direct or indirect way, thus in turn impact diverse cellular processes associated with thrombotic disorders, diabetes, cardiovascular diseases, neurodegenerative diseases, and cancer [[14], [15], [16], [17], [18]]. However, cysteine sulfenic acid in proteins is scarce and typically has a short half-life, posing significant challenges for its detection. During the past decades, researchers have intensively dedicated efforts to develop various in vitro and in vivo analytical probes and methods for the measurement of protein sulfenylation. We here will review its historical discovery, chemical formation, and detection approaches, as well as its crucial roles in regulating the progression of various diseases.

2 ROS signaling in biology

In the study of redox biology and medicine, ROS are hard to bypass, which are a class of essential redox signaling molecules rather than hazardous byproducts. ROS, initially discovered in skeletal muscles [19], can be continuously generated in cells undergoing normal metabolism, and can be maintained at a low steady-state level [20]. Derived from molecular oxygen (O2), ROS encompass a group of molecules, with our main focus here on hydrogen peroxide (H2O2), superoxide anion (O2•−), and hydroxyl radical (•OH), each having extensive documented functions. Although ROS were initially regarded as harmful byproducts of cellular aerobic metabolism due to their detrimental roles in causing cellular damage [21], research over the last few decades has revealed various beneficial functions of ROS in cells, such as inducing mitogenic response, triggering and sustaining oncogenic phenotype [22,23]. Consequently, ROS are now widely acknowledged as crucial signaling molecules that regulate various physiological and pathological processes [24].

2.1 Generation of intracellular ROS

Cells employ various pathways to generate ROS, mainly involving four sources: mitochondria, plasma membrane, endoplasmic reticulum (ER), and peroxisome [25,26]. Among these, the majority of ROS generation occurs within mitochondria, where O2 obtains electrons from the mitochondrial electron transport chain (ETC) to form O2•− [27]. Within the mitochondrial ETC, complex I generates O2•− towards the mitochondrial matrix, while complex III releases O2•− into both the mitochondrial matrix and intermembrane space (IMS). O2•− subsequently undergoes a rapid dismutation to generate H2O2 and O2, which can be further enhanced by superoxide dismutases (SODs), such as SOD1 located in the mitochondrial IMS or SOD2 located in the mitochondrial matrix [28]. It was previously believed that mitochondrial ROS production results from ETC leakage, implying ROS production in mitochondria as a detrimental cellular behavior. However, emerging evidence suggests that mitochondrial ROS can be engaged in regulating redox signaling and various fundamental physiological responses. Hence, we have chosen not to employ the term “leakage”. Details of these processes have been covered in some outstanding review articles [29,30].

Additionally, situated on the plasma membrane (PM), NADPH oxidase 1 (NOX1), NADPH oxidase 2 (NOX2), NADPH oxidase 3 (NOX3), and superoxide dismutase 3 (SOD3) are also major sources for ROS generation [31]. NOX1, NOX2, and NOX3 catalyze the conversion of O2 to O2•− by utilizing electrons from NADPH. Afterwards, O2•− is converted into H2O2 non-enzymatically or by the catalytic action of SOD3, which is tethered to the extracellular face of the PM [32]. ROS generated by the PM are primarily released into the extracellular ROS pool, contributing to the extracellular ROS concentrations being two to three orders of magnitude higher than those present intracellularly [33]. What's more, due to the well-recognized roles of NOXs in generating high local ROS concentrations, NOX isoforms located in ER contribute to the production of ER ROS [34]. For example, NADPH oxidase 5 (NOX5) can generate O2•−, and NADPH oxidase 4 (NOX4) is believed to have the capability to directly produce H2O2 [35]; both are produced in the ER and then released into the cytoplasm. It is worth noting that NOX4 is also present on the nuclear membrane for H2O2 generation.

By measuring ROS production in resting C2C12 mouse myoblasts under basal cell conditions, researchers observed that approximately 40% of total ROS pools is generated by NOXs, and about 45% by the mitochondrial ETC (30% by complex III, 15% by complex I) [36]. The remaining portion was attributed to various other enzymes, such as ER oxidoreductin 1 (ERO1) and cytochrome P450 (CYP) family enzymes, both locating on ER. Specifically, protein disulfide isomerase (PDI) oxidatively folds the disulfide bonds of its client proteins, and then transfers the obtained electrons to ERO1. ERO1 further reduces O2 to H2O2 by utilizing these electrons [37]. It has also been reported that PDI can influence ROS production by regulating the activity of NOXs [38]. Moreover, ROS can be generated through CYPs during drug metabolism and the degradation of endogenous substances [39,40]. In addition, many enzymes located in peroxisome also contribute to ROS generation, such as acyl-CoA oxidase (ACOX), D-amino acid oxidase (DAAO) and D-aspartate oxidase (DDO) [25,41,42].

2.2 Balancing ROS via the antioxidant systems

The key to maintain the balance of oxidative stress lies in the robust antioxidant systems, which comprise a series of enzymes and small molecule antioxidants. The primary pathways for the clearance of ROS are as follows: (1) SOD1, SOD2, and SOD3 catalyze the conversion of O2•− to H2O2 and O2 [43]. (2) Catalase (CAT) dismutates two molecules of H2O2 into H2O and O2 [44]. (3) Within the glutathione (GSH) antioxidant system, glutathione peroxidases (GPXs) use electrons from reduced GSH to reduce H2O2 to H2O, concurrently oxidizing GSH to oxidized glutathione (GSSG). Subsequently, glutathione reductase (GR) restores GSSG to GSH, by consuming electrons from NADPH [45,46]. (4) In the thioredoxin (Trx) antioxidant system, peroxiredoxins (PRDXs) use electrons from reduced thioredoxin (TrxR) to reduce H2O2 to H2O, concurrently oxidizing TrxR to oxidized thioredoxin (TrxO). Subsequently, thioredoxin reductase (TrxR) restores TrxO to TrxR, during which NADPH provides the electrons [47,48]. The GSH redox cycle and Trx redox cycle empower GSH and TrxR to function as potent antioxidants for ROS neutralization. Through these antioxidant mechanisms, cells prevent the accumulation of intracellular ROS, thereby alleviating cellular oxidative stress and ultimately shielding cells from the detrimental effects of ROS.

Under normal physiological conditions, cells maintain a rough dynamic balance between the generation and clearance of ROS by various antioxidant systems mentioned above (Fig. 1). In this state, the levels of ROS remain relatively low for the regulation various physiological processes, including blood pressure, embryonic development, cognition, and immune function [[49], [50], [51], [52]]. However, in pathological conditions where mitochondria is damaged, or cells are exposed to toxins, radiation, pathogens, or anticancer drugs, the balance between ROS production and clearance within cells will be disrupted. In this case, ROS can modulate cell proliferation, migration, differentiation, apoptosis, and autophagy, etc. [[53], [54], [55], [56]].Fig. 1 The balance between ROS generation and clearance. Cells mainly generate ROS through mitochondria, plasma membrane, endoplasmic reticulum, and peroxisome, and eliminate ROS through various antioxidant systems. A dynamic balance between the generation and elimination of ROS is maintained under normal physiological conditions.

Fig. 1

3 ROS-mediated protein sulfenylation: an emerging post-translational modification

The flow of ROS towards specific protein targets leads to their reversible or irreversible oxidation to alter protein activity, localization or interactions, thereby impacting various processes in cells and organs, including thrombus formation, cell proliferation, differentiation, migration, and angiogenesis [14,[57], [58], [59]]. These dysregulation events are involved in thrombotic disorders, diabetes, cardiovascular diseases, cancer, and neurodegenerative diseases [60]. Specifically, ROS can react with cysteine residues in proteins to form, for instance, reactive sulfenic acid (-SOH). This oxPTM, termed sulfenylation, can lead to structural and/or functional changes of the target redox-sensitive proteins, thereby activating or deactivating diverse cellular signaling pathways [61].

3.1 The historical discovery of protein sulfenylation

The finding of cysteine sulfenylation can be traced back to 1933. During the investigation of cysteine peroxides, Pirie et al. presumed that sulfenic acid might be an intermediate in the formation of disulfides [62]. However, this research was limited to small-molecule thiols, the formation of sulfenic acid in biological systems remained unknown. At that time, it was widely believed that solvent-accessible cysteine thiols (Cys-SH) were particularly susceptible to oxidation by mild oxidants like H2O2, forming sulfenic acid (Cys-SOH). Furthermore, if specific conformational constraints exist in the spatial arrangement of proteins that prevent sulfenic acid from forming intramolecular or intermolecular disulfide bonds, this would create an ideal environment for the formation and stabilization of sulfenic acid. However, due to the high reactivity of sulfenic acid, it is typically unstable and has a very short half-life, making its detection challenging. This reactivity led to the slow progress in the study of protein sulfenylation. For a long time, the scientific community generally agreed that sulfenic acid is just an intermediate formed in cells under oxidative stress, and would eventually be oxidized to form disulfide bonds or other oxidized forms. Consequently, protein sulfenylation did not attract much attention by scientists at the very beginning.

With the advancement of detection techniques, it was indirectly demonstrated in the 1980s that the active cysteine thiols of glyceraldehyde-3-phosphate dehydrogenase (GAPDH) can be sulfenylated [63,64]. This was supported by (1) the formation of sulfenyl iodide (R-S-I) similar to sulfenylation; (2) the stoichiometry of oxidation of certain protein thiols aligns with protein sulfenylation rather than disulfide bonds; (3) the mild oxidation of critical cysteine thiol group affecting GAPDH catalytic activity introduces electrophilic center with properties similar to sulfenyl halides (R-S-X), which are highly electrophilic intermediates capable of undergoing chemical reactions with other compounds. However, due to the lack of direct detection techniques, there was no direct evidence at that time to prove that proteins could undergo sulfenylation. Direct evidence was obtained in a study which characterized the sulfenylation of the cysteine residue (Cys42) of nicotinamide adenine dinucleotide (NADH) peroxidase using nuclear magnetic resonance (NMR) [65].

However, whether sulfenylation merely acts as an intermediate or itself possesses specific biological functions remained largely unknown. As research on sulfenylation continued to advance comprehensively, this concern has been addressed. For example, the sulfenylation of the cysteine residue (Cys149) of GAPDH alters its substrate affinity, leading to the gain of function as an acyl phosphatase for GAPDH [[66], [67], [68]]. This discovery marked the first indication of the potential biological role of protein sulfenylation, sparking further in-depth investigations into this oxPTM.

3.2 The chemical formation of cysteine sulfenic acid

Whether in antioxidant enzyme active sites or at other sites in proteins, cysteine sulfenylation can be formed in response to different oxidants (Fig. 2). The cysteine sulfur atom can form a chemical bond with one oxygen atom from H2O2 through nucleophilic attack. This interaction leads to the oxidation of the cysteine sulfur group, resulting in the sulfenylation of redox-sensitive proteins (Fig. 2a). It is worth noting that it is not the cysteine thiol but the cysteine thiolate (the deprotonated form) that reacts with H₂O₂. Under acidic conditions, the reaction between H2O2 and cysteine thiolate occurs more readily. The acidic environment facilitates the conversion of the relatively poor leaving group (-OH) to a better leaving group (H2O), promoting the oxidation of cysteine thiolate to form sulfenic acid [69]. Of note, a higher pH and/or lower pKa of the thiol leads to more sulfur groups of the thiolate form, which also promotes cysteine sulfenylation, as described later. Meanwhile, cells use cysteine sulfenylation to reduce H2O2, aiding in the detoxification of high levels of ROS in cells. Similar to H2O2, organic peroxide (ROOH) can also lead to protein cysteine sulfenylation (Fig. 2b). This also occurs through nucleophilic attack of the sulfur group on one of the oxygen atoms in the organic peroxide. However, several organic peroxides oxidize cysteine thiolate in active sites of certain enzymes faster than H2O2, such as linoleic acid hydroperoxide (LAHP) [70,71]. In addition to H2O2 and organic peroxides, another oxidizing agent, hypohalous acid (HOX) can also induce protein sulfenylation (Fig. 2c). Hypohalous acid are produced by neutrophils through the reaction of halogenic acid (HX) with oxidants such as H2O2. Different hypohalous acids vary in their oxidizing potential, with hypochlorous acid (HOCl) as the most potential one. HOCl can react with cysteine thiolate to form sulfenyl halide (Cys-S-X), which then hydrolyzes to generate sulfenic acid (Cys-SOH) [72,73].Fig. 2 The formation of protein cysteine sulfenic acid. Cysteine sulfenylation occurs when the sulfur atom in the Cys-SH reacts with oxidants such as H2O2, ROOH, HOX, ONOOH, and NO, or through disulfide hydrolysis under alkaline conditions.

Fig. 2

In addition, reactive nitrogen species (RNS) can also induce the formation of protein sulfenic acid under certain conditions. For example, N-chloramines (R-NH-Cl) generated from the reaction of HOCl with compounds containing amino groups (NH2) can result in the formation of protein sulfenic acid [74]. Moreover, the peroxynitrous acid (ONOOH), which is the conjugate acid of peroxynitrite anion (ONOO−) formed by the combination of O2•− and nitric oxide (NO), can cause sulfenylation of protein cysteine residues (Fig. 2d). Furthermore, nitric oxide (NO) can react with protein cysteine thiyl radical to generate nitrosothiol (Cys-S-NO), which can be hydrolyzed to sulfenic acid (Cys-SOH), leading to protein sulfenylation (Fig. 2e) [75,76].

Moreover, disulfides can be hydrolyzed under alkaline conditions to form sulfenic acid (Fig. 2f) [77]. However, due to the reactivity of sulfenic acid, it can easily be further oxidized by oxidants to form sulfinic acid, and even sulfonic acid (Fig. 2g). It is worth mentioning that the oxidation of protein sulfenic acid to sulfinic acid can be reversible. For instance, sulfiredoxin (SRX), an ATP-dependent antioxidant protein, can catalyze the reduction of over-oxidized peroxiredoxins (PRDX-SO2H) back to its active form (PRDX-SOH), thus restoring the antioxidant activity of PRDXs [78]. In summary, despite varying reaction rates and mechanisms, many physiologically relevant oxidants can oxidize cysteine sulfur group due to the high nucleophilicity of cysteine thiol(ate).

3.3 Factors influencing the formation of protein sulfenic acid

The generation and distribution of intracellular oxidants (such as H2O2) are highly compartmentalized [1]. The compartmentalized production of oxidants may give rise to differences in oxidant levels among various cellular organelles and subcellular locales [79]. These differences can impact intracellular biochemical reactions, including the sulfenylation of cysteine residues. In this context, the diffusion distance of oxidants becomes a primary factor influencing the formation of cysteine sulfenic acid. Once oxidants (such as H2O2) are produced or released, their concentrations decrease with increased diffusion distance, even more steeply when active H2O2-degrading systems are present.

In addition, due to the secondary and tertiary structures of proteins, the pKa values of cysteine thiol group in proteins can vary within a certain range. As a typical example, the Trx superfamily shows a broad variation in the pKa values of cysteine thiol group crucial for protein activity, spanning from 3 to 14 [80]. The activities of cysteine thiol groups in proteins are influenced by their pKa values at distinct pH level [81]. If the pKa value of a cysteine thiol group is relatively low, it means that under neutral conditions, more thiol groups will be in the negatively charged ion form. Negatively charged thiol ions have stronger nucleophilicity, making them more prone for sulfenylation. It is worth noting that backbone amides involved in hydrogen bonds can also affect the pKa values of cysteine thiol group, thereby modulating their activities. Additionally, the presence of macrodipole moments in the secondary structure of proteins can significantly impact the pKa of cysteine thiol group and potentially affect the formation of sulfenic acid [82]. The presence of charged side chains (such as glutamic acid) and long-range electrostatic interactions also influences the pKa of cysteine thiol group, although their impact is limited [83]. These above-mentioned factors indirectly modify nucleophilicity and affect sulfenylation formation by influencing the pKa of cysteine thiol group. On the other hand, some factors, such as the polarization of cysteine thiol group and the presence of non-bonding electrons on adjacent atoms, can directly alter the nucleophilicity of cysteine thiol group, thereby affecting the formation of protein sulfenic acid.

3.4 Factors influencing the stability of protein sulfenic acid

Given the reactivity of sulfenic acid, it is prone to undergo subsequent reactions [84]. For instance, when an excess of oxidizing agent is present, sulfenic acid may undergo further oxidation to form sulfinic acid, or even sulfonic acid. Moreover, sulfenylated cysteine can react with vicinal thiol group to frequently form intracellular disulfide bond, which is more stable (less reactive) than sulfenic acid. Specifically, when a protein has only one free cysteine, the cysteine sulfenylation can persist longer without forming disulfide bond [85,86]. However, the formation of intramolecular disulfide bond may be prevented by alkylation or other modifications which can eliminate neighboring thiol, thereby increasing the stability of sulfenic acid [87]. Furthermore, it has been reported that the formation of intramolecular disulfide bond between sulfenylated cysteine and vicinal cysteine serves acts as a protective mechanism to prevent further oxidation of the cysteine at the active site of CDC25 phosphatase [88,89].

The formation of hydrogen bond also plays a crucial role in stabilizing sulfenylation. When cysteine residues undergo sulfenylation and are in proximity to the structural units capable of forming hydrogen bond, the interaction of the formed hydrogen bond with sulfenic acid will enhance the stability of sulfenic acid [90,91]. Moreover, sulfenylated cysteine residues situated in hydrophobic regions and located at the interior of proteins with limited solvent exposure would contribute to the stability of sulfenic acid. Besides, the pKa of sulfenylated cysteines can also influence the stability of sulfenic acid. Cysteine residues with a lower pKa are more prone to forming sulfenate anions. In this negative ion state, they are less likely to undergo self-condensation or form disulfide bond, thereby enhancing the stability of sulfenic acid [85].

All these factors mentioned above contribute to the stability of protein sulfenic acid. Understanding these factors helps to unveil the relationship between protein structure and function, as well as the regulatory mechanisms of protein cysteine sulfenylation in chemical reactions.

4 Detection approaches for protein sulfenylation

The low abundance, typically short half-life, and highly dynamic nature of cysteine sulfenylation pose challenges for its enrichment and detection [11,69]. During the past decades, considerable efforts have been invested in developing various analytical methods and probes for the measurement of protein cysteine sulfenylation. Direct detection of sulfenylation can be achieved through physical techniques, such as X-ray crystallography [92], and NMR [65]. However, many proteins are not suitable for these physical techniques [92,93]. Therefore, we mainly exemplify several chemical methods for sulfenylation detection, with discussion of their advantages and limitations. Researchers can select suitable detection methods according to their specific experimental goals.

4.1 In vitro sulfenylation detection

In 1974, dimedone was demonstrated to be a chemically selective probe for protein cysteine sulfenylation [94]. Due to its high specificity, dimedone and diketones have been widely employed to characterize cysteine sulfenylation in the protein repertoire [11,76,95]. Later in 1997, Ellis and colleagues developed 7-chloro-4-nitrobenz-2-oxa-1,3-diazole (NBD-Cl), which reacts with sulfenic acid to produce a characteristic absorption peak (347 nm), facilitating the distinction and detection of sulfenylation [96]. In comparison to dimedone, NBD-Cl allows for the preservation of the sulfenic acid oxygen, enabling the distinguishment between Cys-SH and Cys-SOH by mass spectrometry (MS) analysis. The binding of NBD-Cl to sulfenic acid is reversible and can be removed by dithiothreitol (DTT), thus restoring the activity of sulfenylated proteins. However, despite NBD-Cl primarily modifies cysteine at neutral pH, it may also react with other reactive groups (e.g., amino and tyrosine hydroxyl groups) under different conditions. This potential for non-specific labeling can result in the increase of background signals.

Subsequently, in 2007, King's group developed seven novel reagents for detecting and isolating sulfenylated proteins or peptides by attaching fluorescein, rhodamine, or biotin to 1,3-cyclohexadione as detectable tags [97]. The seven new reagents are: two fluorescein derivatives, DCP-FL1 and DCP-FL2; three biotin derivatives, DCP-Bio1, DCP-Bio2, and DCP-Bio3; and two rhodamine derivatives, DCP-Rho1 and DCP-Rho2. The sulfenylated proteins or peptides labeled with these detectable tags can be measured with high sensitivity using in gel analysis or identified by MS analysis. Notably, these conjugates are capable of releasing specific and detectable fragments for MS analysis to determine the exact sequence of the labeled peptide segment and sulfenylated cysteine site. However, these reagents may require specific pH and temperature conditions to ensure efficient and specific sulfenic acid labeling, often necessitating optimization in complex experimental setups.

Later in 2009, Seo et al. developed a dimedone-specific antibody able to detect sulfenic acid sites trapped by dimedone [98]. They used dimedone to selectively react with the sulfur atom in sulfenic acid, forming a stable thioether product. This conversion of sulfenic acid allowed its specific recognition by an α-hapten antibody, ensuring the accuracy and stability during detection. Compared with dimedone-biotin and azide-tagged dimedone analog, this sulfenic acid-specific antibody demonstrate advantages for immunohistochemical staining in tissues, and also provides higher sensitivity and selectivity for western blot analysis [97,[99], [100], [101]]. However, the drawbacks of this antibody include the high cost of antibody preparation, as well as the dependency of experimental results on the quality and batch consistency of the antibodies.

4.2 In situ sulfenylation detection

The research team led by Wood developed a genetically encoded probe (Yap1-cCRD) for the first time to detect protein sulfenylation in vivo. This probe is based on Yap1 reactivity toward sulfenic acid on the thiol peroxidase Orp1, and can be employed to capture sulfenylated proteins by forming mixed disulfides in any genetically traceable organism or cells in culture [102,103]. With relatively high specificity and sensitivity, it can capture and identify low-abundance Cys-SOH. Notably, the Yap1-cCRD sequence is non-conserved in higher eukaryotes, thus does not interfere with endogenous signaling pathways. Moreover, the Yap1-cCRD probe can be easily modified by altering its gene sequence to meet various experimental needs, providing high flexibility. However, the disulfide complex it forms might only be temporarily stable in vivo, requiring rapid sample processing to prevent reduction.

Subsequently, Klomsiri and colleagues proposed a DCP-Bio1-based in situ labeling strategy to detect protein sulfenylation in living cells [104]. DCP-Bio1 exhibits good stability under different experimental conditions and can work effectively across various pH levels. Labeled proteins can be detected with high sensitivity through gel analysis or MS, ensuring the reliability and accuracy of the results. However, the reaction speed of DCP-Bio1 with sulfenic acid highly depends on the labeling environment and the accessibility and stability of the sulfenic acid, potentially resulting in varying reaction efficiencies among different proteins. Later, Qian et al. reported that linear β-ketoester can function as chemical probes for labeling protein sulfenylation. They developed alkyne β-ketoester probes, which can penetrate cells without inducing ROS accumulation or causing cell death. Compared to DCP-Bio1, alkyne β-ketoester probes are more suitable for detecting and identifying sulfenylated proteins in living cells. Additionally, Tags such as biotin can be coupled through click reactions, and the introduced tags can be removed by NH2OH to facilitate subsequent MS analysis [95].

Over the past decades, Carroll's lab has progressively developed a series of chemoselective probes for sulfenylation measurement, including DAz-1, DAz-2, DYn-1, DYn-2, TD, PYD, PRD, and BTD. For these sulfenic acid-specific chemical probes, dimedone is connected with an azide chemical handle for click chemistry reaction to enable the direct detection of protein sulfenylation in cells within their native environment. DYn-1 and DYn-2 are alkyne-modified analogs with higher sensitivity for detecting sulfenylation compared with DAz-1 and DAz-2. Notably, cells treated with DYn-2 show no significant loss of cell viability, and the signal from DYn-2 labeling is 40% higher than DAz-2 under the same conditions [100,105,106]. TD, PYD, PRD, and BTD are four alkyne-tagged probes that selectively label sulfenylated proteins. Compared to DYn-2, they all demonstrate higher reaction rates towards Cys-SOH. Specifically, BTD exhibits a reaction rate over two orders of magnitude higher, with great potential to capture transient protein sulfenylation events [107,108]. Moreover, Niu et al. recently developed a high-performance probe for detecting protein sulfenylation, named biotin-benzoboroxole (Bio-ben). Even at low concentration, the Bio-ben probe exhibits high capture efficiency for protein sulfenylation, significantly shortening the labeling time compared with the commonly used dimedone-based probes [109].

In addition, Yin et al. designed and synthesized the fluorescent probe CPD by combining a dimedone analogue with a 7-aminocoumarin fluorophore. Even at a low concentration (5 μM), CPD can effectively label intracellular sulfenylated proteins, which minimizes interference with intracellular ROS levels. This probe allows for rapid assessment of protein sulfenylation in cells through fluorescence readouts, enabling visualization of the global distribution and dynamic changes of protein sulfenylation in situ. However, it requires the use of a comparison probe (such as CPDDM) to verify the specificity of the labeling [110]. Later, Tom and colleagues developed a ratiometric fluorescent probe, F-DiNap, by coupling α-fluoro-substituted dimedone with an aminonaphthalene fluorophore. The F-DiNap probe significantly enhances the selectivity for sulfenylated proteins without affecting its labeling rate, as α-fluorine substitution prevents side reactions with biological aldehydes [111]. This makes it more suitable for assessing the redox status in living cells. However, the probe concentration needs to be precisely controlled to avoid cellular toxicity.

Notably, Holmila and colleagues developed a bioimaging probe, DCP-NEt2-Coumarin (DCP-NEt2C), which is well-suited for detecting sulfenylation of mitochondrial proteins. Importantly, it operates independently of mitochondrial membrane potential and exhibits no obvious mitochondrial toxicity [112]. Shortly thereafter, the team led by Fox developed trans-cycloocten-5-ol (SAM-TCO) probe, which exhibits faster capture kinetics of sulfenic acid compared with conventional probes. The majority of in situ labeling probes retain activity even after cell lysis, and thus require the addition of specific reagents post-lysis for quenching. SAM-TCO, however, can be readily quenched in living cells using tetrazines, thereby avoiding the nonspecific labeling [113].

Recently, Furdui's group developed the first positron emission tomography (PET) [18F]Fluoro-DCP radiotracer ([18F]F-DCP), which can be used to detect protein sulfenylation in vivo. The [18F]F-DCP radiotracer has been successfully applied to distinguish radiation-resistant (with reduced protein sulfenylation level) and radiation-sensitive head and neck squamous cell carcinoma (HNSCC) tumor xenografts in mice, suggesting its potential in the diagnosis of radiation-resistant HNSCC patients [114]. However, the accumulation of [18F]F-DCP in the kidney and liver may affect imaging results and data interpretation, requiring further research to optimize its selective accumulation in tumors.

4.3 Global sulfenome mining using MS analysis

Among the various probes mentioned above, some of them have been employed for global proteomics screening of sulfenylated proteins. Charles and colleagues applied a synthesized biotinylated dimedone analogue combined with LC-ESI-MS analysis to enrich and detect proteins with sulfenylation. They identified 22 sulfenylated proteins in isolated rat cardiomyocytes following H2O2 treatment [99]. Notably, these proteins underwent sulfenylation in response to either H2O or O2 treatment, implying that protein sulfenylation can act as a mechanism for oxygen sensing. However, it is essential to note that two of the identified proteins did not contain cysteines, indicating potential false positives for this method. Considering the important role that sulfenylated proteins may play in tumors, Leonard et al. used the cell-permeable DAz-2 probe in combination with LC-MS/MS analysis to perform global screening analysis of sulfenylated proteins in HeLa cells. 193 sulfenylated candidates were identified, with 14 proteins further confirmed to undergo sulfenylation [105]. These sulfenylated proteins were involved in multiple biological events, including signal transduction, cellular metabolism, redox homeostasis, ER quality control, vesicle transport, protein synthesis, and DNA repair, suggesting protein sulfenylation is extensively involved in various cellular activities.

Previous studies on cysteine modifications were mostly conducted using cell lysates treated with high concentrations of labeling agents. However, the research team led by Liebler directly treated RKO colon cancer cells with the DYn-2 probe to enrich sulfenylated proteins for LC-MS/MS analysis [115,116]. They identified over 700 candidate proteins with more than 1000 sulfenylation cysteine sites. These sulfenylated proteins were present in all major cellular compartments and participate in diverse cellular processes and signaling pathways. A significant number of sulfenylated proteins were located in the nucleus and involved in key nuclear processes, such as RNA translation, transport, and cell cycle. Notably, eukaryotic translation elongation factor 1 alpha 1 (EEF1A1), N-terminal methyltransferase 1 (NTMT1), and Sirtuin 6 (SIRT6) were demonstrated to serve as potential H2O2 sensors that interacted with transcription factors through thiol-disulfide exchange to participate in redox regulation, particularly the sulfenylation of SIRT6 Cys18. Additionally, GO classification and KEGG pathway analysis of these candidate proteins indicated that sulfenylated proteins were enriched in several cellular metabolism processes, such as glycolysis, pentose phosphate pathway, and fatty acid metabolism. Notably, sulfenylation at Cys358 of pyruvate kinase M2 (PKM2) was identified in this proteomics study. It has been previous reported that oxidation of PKM2 at Cys358 inhibited PKM2 kinase activity, redirecting glucose towards the pentose phosphate pathway to generate sufficient NADPH for detoxifying ROS in human lung cancer cells [117]. However, there is no direct evidence demonstrating that Cys358 was sulfenylated in this functional study, which requires further investigation. In addition to nuclear proteins and metabolic proteins, they discovered sulfenylated sites on functional domains of protein kinases, phosphatases, acetyltransferases, deacetylases, and deubiquitinases, suggesting potential regulatory crosstalk between sulfenylation and other major post-translational modification events.

Gupta et al. applied five clickable carbon nucleophile probes (BTD, DYn-2, PYD, PRD, TD) to capture sulfenylated proteins in RKO cells for LC/MS analysis [107]. A total of 761 candidate proteins with 1283 sulfenylation sites were identified, with varying quantities of modification sites ranked in descending order: BTD, DYn-2, PYD, PRD, TD. Interestingly, only nine proteins were labeled by all five probes (ACLY, EEF2, GAPDH, HSPA8, PFN1, RPS20, RPS21, RPS27A, RPS3A). This proteomics study holds significant implications for developing targeted cysteine inhibitors for disease treatment. For instance, the favored target sites of PYD probe were PTP1B Cys215, PTPN7 Cys396, and PTPN11 Cys459, all these cysteine residues are crucial for protein catalytic activities. Inhibitors could be developed by coupling the PYD probe to elements capable of blocking cysteine reactions to inhibit the activities of these proteins. For example, the aforementioned PTP1B is a potential therapeutic target for type II diabetes [118], cancer [119], and neurodegenerative diseases [120]. Developing targeted inhibitors based on the PYD probe may represent a potential therapeutic strategy for these diseases.

Fu et al. used a benzothiazine-based probe (BTD) combined with LC-MS/MS analysis to screen protein sulfenylation in RKO cells upon H2O2 treatment, which required only a small amount of input material to efficiently identify and quantify sulfenylated cysteine sites [108]. They identified 1867 sulfenylation cysteine sites of 1202 candidate proteins, with over 60% meeting the criteria of pQuant interference fraction less than 0.5 for quantification. Interestingly, both this study and the DYn-2 probe-based proteomics screening in RKO cells under different oxidative conditions observed an increase in the sulfenylation level at PRDX6 Cys91, but a decrease at PRDX6 Cys47 [115]. Cys47 is well-known as the peroxide-reactive Cys in the active site of PRDX6; whether Cys91 sulfenylation affects PRDX6's peroxidase activity merits further investigation.

Fransen's group established a DD-DAO Flp-In T-REx 293 cell line-based approach to manipulate intracellular H2O2 production dependent on subcellular compartment, dosage, and time [121]. Subsequently, they screened peroxisomal H2O2-mediated sulfenylation targets in Flp-In T-REx 293 cells using IBD-SBP-YAP1C combined with LC-MS/MS analysis [122]. Over 400 candidate sulfenylation targets in peroxisome, cytoplasm, and mitochondria were identified. Among them, 91 proteins were previously reported to undergo sulfenylation according to the iCysMod database [123]. Notably, H2O2 derived from peroxisomes mediated sulfenylation of multiple protein families, including protein chaperones, antioxidant enzymes, membrane-associated proteins, components of the cytoskeleton, etc. These proteins play roles in proteasome function, carbon metabolism, calcium signaling, protein synthesis, and protein folding. These results suggest that alterations in peroxisomal H2O2 metabolism may have implications for the development of diseases. Interestingly, catalase deficiency is implicated in diseases associated with oxidative stress, such as cancer, atherosclerosis, diabetes, and age-related disorders [124]. However, the mechanisms by which H2O2-induced sulfenylation influences these diseases still require further exploration.

5 The roles of protein sulfenylation in human diseases

As mentioned earlier, certain proteins have intrinsic structures and the environment around cysteine residues that are conducive to the formation and stabilization of sulfenylation. Moreover, the active cysteine residues in many proteins can serve as redox switches, and sulfenylation can affect their activities and then regulate multiple cellular processes. In this regard, we will summarize how protein cysteine sulfenylation plays crucial roles in various diseases by modulating protein activities or interactions, including thrombotic disorders, diabetes, cardiovascular diseases, neurodegenerative diseases, and cancer (Fig. 3). The sulfenylated proteins described in this review are also shown in Table 1.Fig. 3 Cysteine sulfenylation in various diseases. The sulfenylation of cysteine thiols plays a crucial role in various diseases by modulating protein activities or interactions. (A) The sulfenylation of Src kinase and PDI can promote thrombus formation. (B) The sulfenylation of Drp1 and UCP1 promotes mitochondrial dynamics and thermogenic respiration, thereby affecting the development of diabetes. (C) the sulfenylation of proteins such as Kv1.5, PRMT1, AR, AAT1, AE2, Smad3, and CypD impacts their activity and regulates cardiovascular health. (D) The sulfenylation of GAPDH may accelerate the progression of AD, while the sulfenylation of MGL can regulate neurotransmitter release to provide neuroprotection. (E) The sulfenylation of IRE-1 initiates the p38/SKN-1 (Nrf2) antioxidant response, increasing stress resistance and extending lifespan. (F) BCAT1 affects the sulfenylation levels of global proteins in cells, and the sulfenylation of proteins such as AURKA, RAD51, PKM2, AHR, and EGFR alters their activities, ultimately impacting cancer cell survival, proliferation, and drug resistance.

Fig. 3

Table 1 Summary of protein sulfenylation in various diseases with experimental validation.

Table 1Proteins	Sulfenylated cysteine sites	Activation/inactivation	Diseases	References	
Src	C277 and C185	Inactivation	Thrombotic disorders	[131]	
PDI	Unknown	Activation	Thrombotic disorders	[135]	
UCP1	C253	Unknown	Diabetes	[142]	
Drp1	C644	Activation	Diabetes	[140]	
DLDH	Unknown	Inactivation	Diabetes	[143]	
AAT1	C192	Inactivation	Cardiovascular diseases	[160]	
AE2	Unknown	Activation	Cardiovascular diseases	[161]	
Smad3	C64	Inactivation	Cardiovascular diseases	[159]	
CypD	C104	Inactivation	Cardiovascular diseases	[164]	
Kv1.5	C581	Unknown	Cardiovascular diseases	[145]	
PRMT1	C101 and C208	Inactivation	Cardiovascular diseases	[146]	
AR	C298 and C303	Activation	Cardiovascular diseases	[152]	
GAPDH	Unknown	Unknown	Neurodegenerative diseases	[170]	
MGL	C201 and C208	Inactivation	Neurodegenerative diseases	[176]	
IRE-1	C663	Inactivation	Aging	[182]	
AURKA	C290	Activation	Cancer	[185]	
AHR	C300	Unknown	Cancer	[18]	
EGFR	C797	Activation	Cancer	[106]	
RAD51	C319	Inactivation	Cancer	[194]	
PKM2	C424	Inactivation	Cancer	[198]	

5.1 Cysteine sulfenylation in thrombotic disorders

Under pathological conditions, signaling molecules like ROS link oxidative stress to procoagulant reactions. The non-receptor tyrosine kinase Src is widely expressed in platelets and serves as a signaling mediator regulating platelet function [125]. It has been reported that Src kinase can activate platelets, rendering them procoagulant to rapidly respond to vascular injury [126]. Interestingly, cysteine sulfenylation of Src activates its kinase activity, thereby reducing the threshold for platelet activation. Initially, Giannoni et al. proposed a correlation between increased intracellular ROS and enhanced Src kinase activity [127]. Moreover, substantial evidence suggests that ROS can impact Src kinase activity by reversibly oxidizing the active cysteine residues [[128], [129], [130]]. Heppner et al. further elucidated the molecular mechanisms underlying ROS-mediated cysteine oxidation in promoting Src activity [131]. They observed that Cys277 sulfenylation of Src disrupts the stability of the A-loop helical fold to expose Tyr416 for phosphorylation, leading to enhanced Src kinase activity. Notably, the structural alteration of Src caused by Cys277 sulfenylation further enhances the accessibility of H2O2 to Cys185, thereby promoting sulfenylation at Cys185 [131]. Cys185 sulfenylation then destabilizes the binding of pTyr527 with the Src homology 2 (SH2) domain, prevents Src autoinhibition, and ultimately facilitates Src activation. In addition, the platelet scavenger receptor CD36 binds to oxidized low-density lipoprotein (oxLDL), a risk factor for atherothrombosis, promoting H2O2 production. In turn, the generated H2O2 leads to the sulfenylation and subsequent activation of Src kinase, leading to CD36 recruitment and platelet activation. The inhibition of CD36/oxLDL-mediated platelet activation and aggregation can be achieved by using CD36-blocking antibody, antioxidant enzymes reducing H2O2 level, or a carbon nucleophile (BTD) for blocking Src sulfenylation [132].

In addition, it has been reported that cysteine sulfenylation contributes to the prothrombotic potential of protein disulfide isomerase (PDI), an ER resident enzyme with well-known role in oxidative protein folding. It has been previously reported that PDI can escape from ER to support thrombus formation after activation of platelets and endothelial cells [133,134]. Furthermore, oxidants produced during the activation of vascular cells promote the sulfenylation of PDI, thereby enhancing its oxidase activity and promoting thrombus formation. Similarly, oxLDL also induces the sulfenylation of PDI, facilitating thrombus formation [135].

5.2 Cysteine sulfenylation in diabetes

In diabetes, mitochondrial rupture can lead to excessive production of mitochondrial ROS, causing endothelial dysfunction [136,137]. Notably, dynamin-related protein 1 (Drp1) can be recruited to the outer mitochondrial membrane for mitochondrial contraction and division [138,139]. Kim et al. found that protein disulfide isomerase A1 (PDIA1) binds to Drp1 and prevents it from being oxidized in endothelial cells, thereby maintaining normal mitochondrial dynamics and endothelial function. Loss of PDIA1 in endothelial cells induces sulfenylation of Drp1 at Cys644, enhances Drp1 activity, promotes mitochondrial fragmentation and mitochondrial ROS accumulation, resulting in endothelial cell senescence, impairment of endothelium-dependent vasodilation, and angiogenesis [140]. Moreover, it has been demonstrated that mitochondrial ROS can alter the redox state of protein cysteines in fat cells, thereby promoting the thermogenic respiration. It is noteworthy that brown and beige adipose cells can resist diabetes through thermogenic respiration. Uncoupling protein 1 (UCP1) plays a key role in regulating heat generation in brown and beige adipose tissues [141]. Particularly, UCP1 can be sulfenylated by ROS at Cys253, which then drives thermogenic respiration in adipocytes [142]. However, whether UCP1 sulfenylation can impart resistance toward diabetes requires further investigation. In addition to Drp1 and UCP1, mitochondrial dihydrolipoamide dehydrogenase (DLDH), a multifunctional redox-sensitive oxidoreductase, loses its activity due to sulfenylation [143]. Researchers have extensively discussed the inhibition of DLDH as a potential approach to combat type 2 diabetes [144]. Whether sulfenylation of DLDH plays a role in type 2 diabetes remains to be determined.

5.3 Cysteine sulfenylation in cardiovascular diseases

Protein sulfenylation induced by oxidative stress was also implicated in the regulation of cardiovascular diseases. Using a DAz chemical probe to detect sulfenic acid-modified protein, it was found that human patients with atrial fibrillation has a global increased level of sulfenic acid modification compared with healthy individuals. Specifically, the sulfenylation of Kv1.5 (KCNA5) at Cys581 redirects the Kv1.5 channel from the recycling pathway to degradation, which may trigger atrial fibrillation [145]. Furthermore, H2O2-mediated sulfenylation of protein arginine methyltransferase 1 (PRMT1) at Cys101 and Cys208 inhibits its activity, thereby suppressing protein methylation and reducing the production of asymmetric dimethylarginine (ADMA), a risk factor associated with cardiovascular diseases [[146], [147], [148], [149], [150], [151]]. Additionally, oxidative stress caused by ischemia in the heart sulfenylates aldose reductase (AR) at Cys298 and Cys303, thereby activating AR [152]. Activation of AR reduces 4-hydroxy-trans-2-nonenal accumulation in the ischemic heart and increases the infarct-sparing effect of late preconditioning [153]. Furthermore, since AR can catalyze the reduction of glucose, activation of AR also promotes myocardial ischemia-reperfusion injury by reducing glycolysis and increasing NADH/NAD+ ratio [154,155].

Endogenous SO2 has been proposed as a novel gas signaling molecule in the regulation of cardiovascular function [156]. In addition to protein sulfenylation mediated by ROS, sulfenylation of proteins can also be induced by endogenous SO2 and plays a key role in the cardiovascular system, such as regulation of vasodilation and myocardial function [157,158]. Notably, SO2 can rapidly react with H2O2 to generate (HOO)SO2−, a more potent oxidant. (HOO)SO2− may bind to or react with endogenous metal ions, forming even stronger oxidants. Both of them can lead to protein sulfenylation. In a nutshell, SO2 may act as a booster for H2O2-mediated cysteine sulfenylation [159]. Under physiological conditions, SO2 can induce the sulfenylation of aspartate aminotransferase 1 (AAT1) at Cys192 in umbilical vein endothelial cells (HUVECs), inhibiting its AAT activity. This, in turn, inhibits the generation of SO2, thereby maintaining SO2 at a low concentration range and preserving cardiovascular system homeostasis [160]. SO2 was also reported to promote the sulfenylation of anion exchanger 2 (AE2). Cysteine sulfenylation significantly enhances AE2 activity, resulting in decreased intracellular pH and reduced proliferation of vascular smooth muscle cells [161]. Moreover, SO2 induces the sulfenylation of mothers against decapentaplegic homolog 3 (Smad3) at Cys64, preventing Smad3 nuclear translocation and thereby inhibiting its DNA binding activity. This inhibition can reduce angiotensin II-induced collagen deposition and vascular remodeling [159]. Furthermore, ROS in vascular smooth muscle cells can induce the inhibition of Smad3, with sulfenylation potentially playing a role in this inactivation. Notably, the activation of Smad3 in vascular smooth muscle cells is considered as a key mechanism in many cardiovascular diseases, including hypertension, atherosclerosis, and coronary artery diseases [162,163]. In addition, cyclophilin D (CypD) was found to undergo sulfenylation at Cys104 in response to SO2 [164]. SO2-mediated sulfenylation inhibits CypD activity to prevents the opening of the mitochondrial permeability transition pore (mPTP), thereby protecting myocardial cells from apoptosis [165].

5.4 Cysteine sulfenylation in neurodegenerative diseases

Glyceraldehyde 3-phosphate dehydrogenase (GAPDH) is a classical enzyme involved in the glycolytic pathway. It has been reported that PTMs of GAPDH can confer non-glycolytic functions [[166], [167], [168], [169]]. For instance, sulfenylation of GAPDH results in the formation of interprotein disulfide bonds [94,170]. This exposes its non-catalytic cysteines to form more interprotein disulfide bonds, ultimately leading to the oligomerization and aggregation of GAPDH [171,172]. Interestingly, GAPDH aggregation has been observed in post-mortem brain extracts from Alzheimer's disease (AD) patients [173], and the disulfide-bonded form of GAPDH was observed in the aged brains of AD transgenic mice [174]. Oxidized GAPDH is also associated with amyloid-like proteins, which accelerates amyloid-β generation, a process highly correlated with AD [175]. In addition, sulfenylation of monoacylglycerol lipase (MGL) at Cys201 and Cys208 inhibits MGL activity, and blocks the degradation of 2-arachidonoyl glycerol (2-AG) in neurons, thereby increasing 2-AG levels. This elevated 2-AG binds to and activates presynaptic cannabinoid receptor 1 (CB1R), which in turn reduces the release of glutamate to inhibit presynaptic neurotransmitter secretion. Ultimately, this inhibits excessive excitatory activity in neurons, enhancing 2-AG-mediated endocannabinoid signaling [176]. Furthermore, in neurodegenerative diseases, 2-AG exerts its neuroprotective effects by inhibiting nuclear factor κB (NF-κB) mediated by CB1R, resulting in reduced expression of cytokines including IL-1β, TNFα, and IL-6, implying its potential as a therapeutic target for these diseases [[177], [178], [179]]. However, whether sulfenylation of MGL directly regulates neurodegenerative diseases through 2-AG awaits further investigation.

5.5 Cysteine sulfenylation in aging

In the ER, the accumulation of unfolded proteins leads to ER stress, which triggers a complex unfolded protein response (UPRER) [180]. Inositol-requiring enzyme 1 (IRE-1), the most ancestral transducer of the transcriptional UPRER, has both kinase and RNase activities [181]. In response to unfolded proteins, IRE-1 oligomerizes and activates its RNase activity through autophosphorylation, splicing the mRNA encoding the UPRER transcription factor XBP1 to regulate gene expression, thereby promoting the restoration of normal ER function. However, Hourihan et al. found that ROS can sulfenylate Cys663 in the IRE-1 kinase active site, inhibiting its kinase and RNase activities, thus suppressing IRE-1-mediated UPRER. Moreover, IRE-1 Cys663 sulfenylation promotes the recruitment and activation of p38 signaling at IRE-1 to initiate the p38/SKN-1 (Nrf2) antioxidant response, thereby increasing stress resistance and extending the lifespan of C. elegans [182]. This discovery reveals a crucial role of ROS-mediated cysteine sulfenylation in aging.

5.6 Cysteine sulfenylation in cancer

About a century ago, researchers first observed metabolic changes in cancer cells, and tumor metabolism has since become a significant area in cancer biology [183]. Metabolic abnormalities can lead to the accumulation of ROS in cancer cells, which, in turn, can act as signaling molecules to alter the activity or function of metabolic enzymes to regulate cellular metabolism. For example, branched-chain amino acid transaminase 1 (BCAT1), the overexpression of which predicts poor prognosis in many malignant tumors, harbors a CXXC amino acid motif that is sensitive to oxidation [184]. Oxidation of this CXXC motif can lead to a loss of up to 50% of its transaminase activity. Interestingly, a recent study by Francois and colleagues found that this motif of BCAT1 also influences its non-enzymatic function. In mitotic cells, mutating Cys335 and Cys338 of the CXXC motif to serine (BCAT1SXXS) leads to an increase in the sulfenylation of total protein cysteine residues [185].

BCAT1 knockout leads to an increased sulfenylation of cysteine residues of various proteins, such as protein phosphatase 1 (PP1) [185]. PP1 can counteract mitotic kinases, including cyclin-dependent kinase 1 (CDK1), and is essential for mitotic exit [186,187]. Furthermore, cells expressing BCAT1SXXS mutant reduce the localization of Aurora kinase B (AURKB) at the inner centromere, resulting in incomplete mitosis [185]. Additionally, Aurora kinase A (AURKA) belongs to the serine/threonine kinase family and serves as a key regulator of mitosis [188]. AURKA is often highly expressed in various cancers and is closely associated with the occurrence and development of cancer [189]. In actively cycling cancer cells, ROS can induce the sulfenylation of AURKA at Cys290. Sulfenylated AURKA subsequently dimerizes through formation of disulfide bond to promote its autophosphorylation at Thr288, ultimately activating AURKA during the onset of mitosis [[190], [191], [192]]. Notably, AURKA can regulate cell proliferation, apoptosis, migration, invasion, and epithelial-mesenchymal transition (EMT) by phosphorylating mitosis regulators and tumor suppressors such as p53, ras association domain family member 1A (RASSF1A), liver kinase B1 (LKB1), neurofibromin 2 (NF2), and large tumor suppressor kinase 2 (Lats2) [193].

RAD51 is a crucial recombinase enzyme involved in homologous recombination (HR) DNA repair. In breast cancer cells, sulfenylation of RAD51 at Cys319 inhibits its activity to suppress ionizing radiation (IR)-induced RAD51 foci formation, thereby inhibiting HR DNA repair and promoting DNA damage. Additionally, sulfenylation of RAD51 sensitizes cancer cells to inhibitors of poly (ADP-ribose) polymerase (PARP). However, PRDX1 can bind to RAD51 and maintain RAD51 in a reduced state, therefore promoting HR DNA repair and facilitating cancer cell survival [194].

Aryl hydrocarbon receptor (AHR), a transcription factor, undergoes sulfenylation at Cys300 in response to ROS in drug-resistant tumor cells. Sulfenylation of AHR results in losing its capacity to bind to heat shock protein 90 (HSP90). Instead, Sulfenylated AHR competitively interacts with protein phosphatase 1 regulatory subunit 3C (PPP1R3C) to block the dephosphorylation of glycogen phosphorylase (GP) and thus preserves the activity of GP. GP activation then promotes glycogen breakdown and PPP, leading to NADPH generation for ROS clearance and subsequent chemoresistance [18]. This study indicates that ROS-mediated protein sulfenylation may offer a new approach for resisting chemotherapy resistance.

PKM2, a rate-limiting enzyme in glycolysis, is responsible for catalyzing the conversion of phosphoenolpyruvate (PEP) to pyruvate, with the production of ATP. PKM2 plays a crucial function in the growth and metastasis of cancer cells [195,196]. The formation of PKM2 tetramers is essential for its activity. Structural analysis of PKM2 shows that both Cys424 and Cys358 can affect subunit interactions and enzyme activity. Cys424 is located at the subunit interface, while Cys358 is located in the β-barrel that contains residues essential for catalytic activity [197]. Research has shown that ROS can inhibit PKM2 activity by oxidizing Cys358 and dissociating its subunits, leading to glucose flux into the pentose phosphate pathway (PPP) and increased NADPH production. NADPH is necessary for the generation of reduced GSH in cells, which can effectively detoxify ROS and ultimately help cells survive under oxidative stress [117]. However, the specific mechanisms by which ROS affect PKM2 tetramer formation and activity are not fully understood. Irokawa et al. found that oxidation of PKM2 Cys358 promotes the formation of intramolecular polysulfide bonds and does not affect PKM2 tetramer formation, whereas sulfenylation of PKM2 at Cys424 leads to a decrease in its tetramer formation with reduced PKM2 activity, thereby promoting cancer cell growth under oxidative stress [198].

It is widely recognized that the aberrant activation of epidermal growth factor receptor (EGFR) plays a significant role in promoting cancer progression [199,200]. Notably, H2O2-induced reversible sulfenylation of EGFR at Cys797 can enhance its tyrosine kinase activity [106]. The Cys797 sulfenylation not only influences EGFR activity but also reduces the sensitivity of cancer cells to EGFR inhibitors [[201], [202], [203]]. For example, cancer cells which were originally sensitive to EGFR inhibitors (such as afatinib and PD168393) yet became insensitive under conditions of chronic oxidative stress [204,205]. On one hand, the sulfenylation of EGFR caused local conformational changes, thereby diminishing the binding affinity between EGFR and its inhibitors. On the other hand, sulfenylation of EGFR leads to weakened nucleophilicity, thereby inhibiting the reactivity of EGFR with its inhibitors.

6 Discussion

In this review, we summarize the generation and clearance of ROS, followed by the highlight of ROS-mediated protein cysteine sulfenylation, an essential PTM in regulating protein function. Although research in this area has emerged relatively recently, there is growing evidence that ROS can regulate protein conformation, activity, and interaction by sulfenylating cysteine thiols, thereby modulating protein function and playing crucial roles in various diseases, including thrombotic disorders, diabetes, cardiovascular diseases, neurodegenerative diseases, cancer, and aging. These findings highlight the potential of targeting sulfenylated proteins for therapeutic intervention. However, the current understanding of sulfenylated proteins and their functions is still limited. Therefore, it is essential to identify more proteins whose functions are altered by sulfenylation and elucidate their mechanisms in disease progression.

Currently, the detection methods for protein sulfenylation have made significant progress, despite challenges posed by its low abundance, typically short half-life, and highly dynamic nature. In vitro detection methods, such as dimedone and NBD-Cl, exhibit high specificity and sensitivity but require optimized experimental conditions to prevent nonspecific labeling. In vivo detection methods, such as genetically encoded probes (such as Yap1-cCRD) and chemical probes (such as DCP-Bio1), can detect sulfenylation in living cells, but necessitate rapid sample processing to suppress post-lysis artifacts. Additionally, probes like SAM-TCO can be quenched in living cells to avoid nonspecific labeling after cell lysis. Furthermore, large-scale sulfenylation proteomics analysis based on these existing probes has revealed the significant role of this modification in regulating cellular functions and signaling pathways. This has important implications for elucidating disease mechanisms, identifying potential therapeutic targets, and developing novel treatment strategies. However, advances in protein sulfenylation research rely heavily on the development and application of specific probes. The reactivity and typically short half-life of sulfenic acid pose major challenges for its detection. Future progress may focus on developing more stable and sensitive probes and improving real-time in vivo detection technologies. We believe that combining three-dimensional structural information of proteins with the chemical reactivity characteristics of sulfenylated sites could help scientists design probes that can precisely recognize and target sulfenylated proteins. Additionally, developing computational simulation and modeling techniques to predict and simulate the formation and stability of Cys-SOH under different conditions could help optimize detection conditions and improve the specificity and efficiency of detection methods.

Protein sulfenylation plays a critical role in cellular redox regulation and signaling pathways, making it a promising target for disease diagnosis and treatment. For example, the [18F]F-DCP radiotracer for detecting protein sulfenylation levels in animal models has been successfully applied to distinguish between radiation-resistant (with reduced protein sulfenylation levels) and radiation-sensitive HNSCC tumor xenografts in mice, indicating its potential in improving the diagnosis of radiation-resistant HNSCC patients. In addition, the PYD probe has shown potential in selectively labeling sulfenylated proteins, with preferred target sites being PTP1B Cys215, PTPN7 Cys396, and PTPN11 Cys459. Therefore, the PYD probe can be used to develop covalent inhibitors targeting the redox-active cysteines of these proteins. For instance, PTP1B is a potential therapeutic target for type II diabetes, cancer, and neurodegenerative diseases, and developing PTP1B-targeted inhibitors based on the PYD probe may represent a potential therapeutic strategy for these diseases. However, further efforts should be made to minimize the trapping of the PYD probe to other sulfenylated proteins. This reduces the risk of nonspecific inhibition of other protein activities, which may lead to side effects and toxicity. Despite the limitations of existing detection methods, ongoing research and technological advancements will offer hope for disease diagnosis and treatment based on sulfenylation.

CRediT authorship contribution statement

Baoquan Mu: Writing – original draft, Visualization. Yan Zeng: Visualization. Li Luo: Writing – review & editing. Kui Wang: Writing – review & editing.

Declaration of competing interest

None.

Data availability

Data will be made available on request.

Acknowledgements

This work was supported by the Chinese NSFC No. 82273122 (K.W.), No. 82073081 (K.W.), and No. 82002963 (L.L.); 10.13039/501100021171 Guangdong Basic and Applied Basic Research Foundation No. 2019B030302012 (K.W.).
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References

1 Sies H. Jones D.P. Reactive oxygen species (ROS) as pleiotropic physiological signalling agents Nat. Rev. Mol. Cell Biol. 21 2020 363 383 32231263
2 Jomova K. Raptova R. Alomar S.Y. Alwasel S.H. Nepovimova E. Kuca K. Valko M. Reactive oxygen species, toxicity, oxidative stress, and antioxidants: chronic diseases and aging Arch. Toxicol. 97 2023 2499 2574 37597078
3 Zhang J. Wang X. Vikash V. Ye Q. Wu D. Liu Y. Dong W. ROS and ROS-mediated cellular signaling Oxid. Med. Cell. Longev. 2016 2016 4350965
4 Dan Dunn J. Alvarez L.A. Zhang X. Soldati T. Reactive oxygen species and mitochondria: a nexus of cellular homeostasis Redox Biol. 6 2015 472 485 26432659
5 Go Y.M. Jones D.P. The redox proteome J. Biol. Chem. 288 2013 26512 26520 23861437
6 Aledo J.C. Aledo P. Susceptibility of protein methionine oxidation in response to hydrogen peroxide treatment-ex vivo versus in vitro: a computational insight Antioxidants 9 2020
7 Brigelius-Flohé R. Flohé L. Selenium and redox signaling Arch. Biochem. Biophys. 617 2017 48 59 27495740
8 Santos C.X. Hafstad A.D. Beretta M. Zhang M. Molenaar C. Kopec J. Fotinou D. Murray T.V. Cobb A.M. Martin D. Zeh Silva M. Anilkumar N. Schröder K. Shanahan C.M. Brewer A.C. Brandes R.P. Blanc E. Parsons M. Belousov V. Cammack R. Hider R.C. Steiner R.A. Shah A.M. Targeted redox inhibition of protein phosphatase 1 by Nox4 regulates eIF2α-mediated stress signaling EMBO J. 35 2016 319 334 26742780
9 Haghikia A. Landmesser U. Lipoproteins and cardiovascular redox signaling: role in atherosclerosis and coronary disease Antioxidants Redox Signal. 29 2018 337 352
10 Marino S.M. Gladyshev V.N. Cysteine function governs its conservation and degeneration and restricts its utilization on protein surfaces J. Mol. Biol. 404 2010 902 916 20950627
11 Paulsen C.E. Carroll K.S. Cysteine-mediated redox signaling: chemistry, biology, and tools for discovery Chem. Rev. 113 2013 4633 4679 23514336
12 Held J.M. Gibson B.W. Regulatory control or oxidative damage? Proteomic approaches to interrogate the role of cysteine oxidation status in biological processes Mol. Cell. Proteomics : MCP. 11 2012 013037 R111
13 Antelmann H. Helmann J.D. Thiol-based redox switches and gene regulation Antioxidants Redox Signal. 14 2011 1049 1063
14 Yang M. Smith B.C. Cysteine and methionine oxidation in thrombotic disorders Curr. Opin. Chem. Biol. 76 2023 102350
15 Holendova B. Plecita-Hlavata L. Cysteine residues in signal transduction and its relevance in pancreatic beta cells Front. Endocrinol. 14 2023 1221520
16 Hussain M. Ikram W. Ikram U. Role of c-Src and reactive oxygen species in cardiovascular diseases Mol. Genet. Genom. : MGG. 298 2023 315 328
17 Hwang I. Tang D. Paik J. Oxidative stress sensing and response in neural stem cell fate Free Radic. Biol. Med. 169 2021 74 83 33862161
18 Zhou N. Chen J. Ling Z. Zhang C. Zhou Y. Wang D. Zhou L. Wang Z. Sun N. Wang X. Zhang H. Tang K. Ma J. Lv J. Huang B. Aryl hydrocarbon receptor sulfenylation promotes glycogenolysis and rescues cancer chemoresistance J. Clin. Invest. 133 2023
19 Commoner B. Townsend J. Pake G.E. Free radicals in biological materials Nature 174 1954 689 691 13213980
20 Sies H. Chance B. The steady state level of catalase compound I in isolated hemoglobin-free perfused rat liver FEBS Lett. 11 1970 172 176 11945479
21 Baud L. Ardaillou R. Reactive oxygen species: production and role in the kidney Am. J. Physiol. 251 1986 F765 F776 3022602
22 Yang B. Chen Y. Shi J. Reactive oxygen species (ROS)-Based nanomedicine Chem. Rev. 119 2019 4881 4985 30973011
23 Milkovic L. Cipak Gasparovic A. Cindric M. Mouthuy P.A. Zarkovic N. Short overview of ROS as cell function regulators and their implications in therapy concepts Cells 8 2019
24 Kietzmann T. Cellular redox compartments Antioxidants Redox Signal. 30 2019 1 4
25 Forrester S.J. Kikuchi D.S. Hernandes M.S. Xu Q. Griendling K.K. Reactive oxygen species in metabolic and inflammatory signaling Circ. Res. 122 2018 877 902 29700084
26 Cheung E.C. Vousden K.H. The role of ROS in tumour development and progression Nat. Rev. Cancer 22 2022 280 297 35102280
27 Zhao R.Z. Jiang S. Zhang L. Yu Z.B. Mitochondrial electron transport chain, ROS generation and uncoupling Int. J. Mol. Med. 44 2019 3 15 (Review) 31115493
28 Wang Y. Branicky R. Noë A. Hekimi S. Superoxide dismutases: dual roles in controlling ROS damage and regulating ROS signaling J. Cell Biol. 217 2018 1915 1928 29669742
29 Shadel G.S. Horvath T.L. Mitochondrial ROS signaling in organismal homeostasis Cell 163 2015 560 569 26496603
30 Willems P.H. Rossignol R. Dieteren C.E. Murphy M.P. Koopman W.J. Redox homeostasis and mitochondrial dynamics Cell Metabol. 22 2015 207 218
31 Vermot A. Petit-Härtlein I. Smith S.M.E. Fieschi F. NADPH oxidases (NOX): an overview from discovery, molecular mechanisms to physiology and pathology Antioxidants 10 2021
32 Lee K.S. Kim B.Y. Park M.J. Deng Y. Kim J.M. Kim Y.H. Heo E.J. Yoon H.J. Lee K.Y. Choi Y.S. Jin B.R. Bee venom induces acute inflammation through a H(2)O(2)-mediated system that utilizes superoxide dismutase Toxins 14 2022
33 Forman H.J. Bernardo A. Davies K.J. What is the concentration of hydrogen peroxide in blood and plasma? Arch. Biochem. Biophys. 603 2016 48 53 27173735
34 Roscoe J.M. Sevier C.S. Pathways for sensing and responding to hydrogen peroxide at the endoplasmic reticulum Cells 9 2020
35 Pena E. El Alam S. Siques P. Brito J. Oxidative stress and diseases associated with high-altitude exposure Antioxidants 11 2022 267 35204150
36 Wong H.S. Benoit B. Brand M.D. Mitochondrial and cytosolic sources of hydrogen peroxide in resting C2C12 myoblasts Free Radic. Biol. Med. 130 2019 140 150 30389498
37 Smirnova O.A. Bartosch B. Zakirova N.F. Kochetkov S.N. Ivanov A.V. Polyamine metabolism and oxidative protein folding in the ER as ROS-producing systems neglected in virology Int. J. Mol. Sci. 19 2018
38 Gimenez M. Veríssimo-Filho S. Wittig I. Schickling B.M. Hahner F. Schürmann C. Netto L.E.S. Rosa J.C. Brandes R.P. Sartoretto S. De Lucca Camargo L. Abdulkader F. Miller F.J. Jr. Lopes L.R. Redox activation of Nox1 (NADPH oxidase 1) involves an intermolecular disulfide bond between protein disulfide isomerase and p47(phox) in vascular smooth muscle cells Arterioscler. Thromb. Vasc. Biol. 39 2019 224 236 30580571
39 Jiaxin S. Shengchen W. Yirong C. Shuting W. Shu L. Cadmium exposure induces apoptosis, inflammation and immunosuppression through CYPs activation and antioxidant dysfunction in common carp neutrophils Fish Shellfish Immunol. 99 2020 284 290 32058096
40 Cao M. Yuan W. Peng M. Mao Z. Zhao Q. Sun X. Yan J. Role of CyPA in cardiac hypertrophy and remodeling Biosci. Rep. 39 2019
41 He A. Dean J.M. Lodhi I.J. Peroxisomes as cellular adaptors to metabolic and environmental stress Trends Cell Biol. 31 2021 656 670 33674166
42 den Toom W.T.F. van Soest D.M.K. Polderman P.E. van Triest M.H. Bruurs L.J.M. De Henau S. Burgering B.M.T. Dansen T.B. Oxygen-consumption based quantification of chemogenetic H(2)O(2) production in live human cells Free Radic. Biol. Med. 206 2023 134 142 37392950
43 Fukai T. Ushio-Fukai M. Superoxide dismutases: role in redox signaling, vascular function, and diseases Antioxidants Redox Signal. 15 2011 1583 1606
44 Glorieux C. Calderon P.B. Catalase, a remarkable enzyme: targeting the oldest antioxidant enzyme to find a new cancer treatment approach Biol. Chem. 398 2017 1095 1108 28384098
45 Pei J. Pan X. Wei G. Hua Y. Research progress of glutathione peroxidase family (GPX) in redoxidation Front. Pharmacol. 14 2023 1147414
46 Muri J. Kopf M. Redox regulation of immunometabolism Nat. Rev. Immunol. 21 2021 363 381 33340021
47 Hasan A.A. Kalinina E. Tatarskiy V. Shtil A. The thioredoxin system of mammalian cells and its modulators Biomedicines 10 2022
48 Rhee S.G. Kil I.S. Multiple functions and regulation of mammalian peroxiredoxins Annu. Rev. Biochem. 86 2017 749 775 28226215
49 Gao H.L. Yu X.J. Zhang Y. Wang C.L. Lei Y.M. Yu J.Y. Zong D.M. Liu K.L. Zhang D.D. Li Y. Tian H. Zhang N.P. Kang Y.M. Astaxanthin ameliorates blood pressure in salt-induced prehypertensive rats through ROS/MAPK/NF-κB pathways in the hypothalamic paraventricular nucleus Cardiovasc. Toxicol. 21 2021 1045 1057 34537923
50 Engineer A. Saiyin T. Greco E.R. Feng Q. Say NO to ROS: their roles in embryonic heart development and pathogenesis of congenital heart defects in maternal diabetes Antioxidants 8 2019
51 Ionescu-Tucker A. Cotman C.W. Emerging roles of oxidative stress in brain aging and Alzheimer's disease Neurobiol. Aging 107 2021 86 95 34416493
52 Chávez M.D. Tse H.M. Targeting mitochondrial-derived reactive oxygen species in T cell-mediated autoimmune diseases Front. Immunol. 12 2021 703972
53 Nakamura H. Takada K. Reactive oxygen species in cancer: current findings and future directions Cancer Sci. 112 2021 3945 3952 34286881
54 Varisli L. Tolan V. Increased ROS alters E-/N-cadherin levels and promotes migration in prostate cancer cells Bratisl. Lek. Listy 123 2022 752 757 35913012
55 Agidigbi T.S. Kim C. Reactive oxygen species in osteoclast differentiation and possible pharmaceutical targets of ROS-mediated osteoclast diseases Int. J. Mol. Sci. 20 2019
56 Wang B. Wang Y. Zhang J. Hu C. Jiang J. Li Y. Peng Z. ROS-induced lipid peroxidation modulates cell death outcome: mechanisms behind apoptosis, autophagy, and ferroptosis Arch. Toxicol. 97 2023 1439 1451 37127681
57 Sies H. Berndt C. Jones D.P. Oxidative stress Annu. Rev. Biochem. 86 2017 715 748 28441057
58 Jones D.P. Sies H. The redox code Antioxidants Redox Signal. 23 2015 734 746
59 Zhang L. Wang X. Cueto R. Effi C. Zhang Y. Tan H. Qin X. Ji Y. Yang X. Wang H. Biochemical basis and metabolic interplay of redox regulation Redox Biol. 26 2019 101284
60 Yang S. Lian G. ROS and diseases: role in metabolism and energy supply Mol. Cell. Biochem. 467 2020 1 12 31813106
61 Roos G. Messens J. Protein sulfenic acid formation: from cellular damage to redox regulation Free Radic. Biol. Med. 51 2011 314 326 21605662
62 Pirie N.W. The oxidation of sulphydryl compounds by hydrogen peroxide: catalysis of oxidation of cysteine by thiocarbamides and thiolglyoxalines Biochem. J. 27 1933 1181 1188 16745209
63 Soukri A. Mougin A. Corbier C. Wonacott A. Branlant C. Branlant G. Role of the histidine 176 residue in glyceraldehyde-3-phosphate dehydrogenase as probed by site-directed mutagenesis Biochemistry 28 1989 2586 2592 2659073
64 Ehrenfeld M. Jeck R. Klatte W. Kühn N. Woenckhaus C. [Half-of-the-sites reactivity of glyceraldehyde-3 phosphate dehydrogenase from rabbit muscle with structural analogs of NAD (author's transl)] Zeitschrift fur Naturforschung Section C, Biosciences. 36 1981 545 551 7281897
65 Crane E.J. Vervoort J. Claiborne A. 13C NMR analysis of the cysteine-sulfenic acid redox center of enterococcal NADH peroxidase Biochemistry 36 1997 8611 8618 9214307
66 Ehring R. Colowick S.P. The two-step formation and inactivation of acylphosphatase by agents acting on glyceraldehyde phosphate dehydrogenase J. Biol. Chem. 244 1969 4589 4599 4309146
67 Allison W.S. Connors M.J. The activation and inactivation of the acyl phosphatase activity of glyceraldehyde-3-phosphate dehydrogenase Arch. Biochem. Biophys. 136 1970 383 391 4314108
68 Allison W.S. Benitez L.V. An adenosine triphosphate-phosphate exchange catalyzed by a soluble enzyme couple inhibited by uncouplers of oxidative phosphorylation Proc. Natl. Acad. Sci. U.S.A. 69 1972 3004 3008 4507619
69 Gupta V. Carroll K.S. Sulfenic acid chemistry, detection and cellular lifetime Biochim. Biophys. Acta 1840 2014 847 875 23748139
70 Little C. O'Brien P.J. Products of oxidation of a protein thiol group after reaction with various oxidizing agents Arch. Biochem. Biophys. 122 1967 406 410 6066248
71 Little C. O'Brien P.J. The effectiveness of a lipid peroxide in oxidizing protein and non-protein thiols Biochem. J. 106 1968 419 423 5637351
72 Peskin A.V. Winterbourn C.C. Kinetics of the reactions of hypochlorous acid and amino acid chloramines with thiols, methionine, and ascorbate Free Radic. Biol. Med. 30 2001 572 579 11182528
73 Folkes L.K. Candeias L.P. Wardman P. Kinetics and mechanisms of hypochlorous acid reactions Arch. Biochem. Biophys. 323 1995 120 126 7487057
74 Peskin A.V. Winterbourn C.C. Taurine chloramine is more selective than hypochlorous acid at targeting critical cysteines and inactivating creatine kinase and glyceraldehyde-3-phosphate dehydrogenase Free Radic. Biol. Med. 40 2006 45 53 16337878
75 Verde C. Giordano D. Bruno S. NO and heme proteins: cross-talk between heme and cysteine residues Antioxidants 12 2023
76 DeMaster E.G. Quast B.J. Redfern B. Nagasawa H.T. Reaction of nitric oxide with the free sulfhydryl group of human serum albumin yields a sulfenic acid and nitrous oxide Biochemistry 34 1995 11494 11499 7547878
77 Andersson L.O. Hydrolysis of disulfide bonds in weakly alkaline media. II. Bovine serum albumin dimer Biochim. Biophys. Acta 200 1970 363 369 4984557
78 Roussel X. Kriznik A. Richard C. Rahuel-Clermont S. Branlant G. Catalytic mechanism of Sulfiredoxin from Saccharomyces cerevisiae passes through an oxidized disulfide sulfiredoxin intermediate that is reduced by thioredoxin J. Biol. Chem. 284 2009 33048 33055 19801666
79 Sies H. Belousov V.V. Chandel N.S. Davies M.J. Jones D.P. Mann G.E. Murphy M.P. Yamamoto M. Winterbourn C. Defining roles of specific reactive oxygen species (ROS) in cell biology and physiology Nat. Rev. Mol. Cell Biol. 23 2022 499 515 35190722
80 Hofmann B. Hecht H.J. Flohé L. Peroxiredoxins Biol. Chem. 383 2002 347 364 12033427
81 Ferrer-Sueta G. Manta B. Botti H. Radi R. Trujillo M. Denicola A. Factors affecting protein thiol reactivity and specificity in peroxide reduction Chem. Res. Toxicol. 24 2011 434 450 21391663
82 Kortemme T. Creighton T.E. Ionisation of cysteine residues at the termini of model alpha-helical peptides. Relevance to unusual thiol pKa values in proteins of the thioredoxin family J. Mol. Biol. 253 1995 799 812 7473753
83 Roos G. Foloppe N. Messens J. Understanding the pK(a) of redox cysteines: the key role of hydrogen bonding Antioxidants Redox Signal. 18 2013 94 127
84 Peskin A.V. Dickerhof N. Poynton R.A. Paton L.N. Pace P.E. Hampton M.B. Winterbourn C.C. Hyperoxidation of peroxiredoxins 2 and 3: rate constants for the reactions of the sulfenic acid of the peroxidatic cysteine J. Biol. Chem. 288 2013 14170 14177 23543738
85 Salsbury F.R. Jr. Knutson S.T. Poole L.B. Fetrow J.S. Functional site profiling and electrostatic analysis of cysteines modifiable to cysteine sulfenic acid Protein Sci. : a publication of the Protein Society 17 2008 299 312
86 Turell L. Botti H. Carballal S. Ferrer-Sueta G. Souza J.M. Durán R. Freeman B.A. Radi R. Alvarez B. Reactivity of sulfenic acid in human serum albumin Biochemistry 47 2008 358 367 18078330
87 Miller H. Claiborne A. Peroxide modification of monoalkylated glutathione reductase. Stabilization of an active-site cysteine-sulfenic acid J. Biol. Chem. 266 1991 19342 19350 1918050
88 Sohn J. Rudolph J. Catalytic and chemical competence of regulation of cdc25 phosphatase by oxidation/reduction Biochemistry 42 2003 10060 10070 12939134
89 Buhrman G. Parker B. Sohn J. Rudolph J. Mattos C. Structural mechanism of oxidative regulation of the phosphatase Cdc25B via an intramolecular disulfide bond Biochemistry 44 2005 5307 5316 15807524
90 Witt A.C. Lakshminarasimhan M. Remington B.C. Hasim S. Pozharski E. Wilson M.A. Cysteine pKa depression by a protonated glutamic acid in human DJ-1 Biochemistry 47 2008 7430 7440 18570440
91 Blackinton J. Lakshminarasimhan M. Thomas K.J. Ahmad R. Greggio E. Raza A.S. Cookson M.R. Wilson M.A. Formation of a stabilized cysteine sulfinic acid is critical for the mitochondrial function of the parkinsonism protein DJ-1 J. Biol. Chem. 284 2009 6476 6485 19124468
92 Stehle T. Claiborne A. Schulz G.E. NADH binding site and catalysis of NADH peroxidase Eur. J. Biochem. 211 1993 221 226 8425532
93 Furdui C.M. Poole L.B. Chemical approaches to detect and analyze protein sulfenic acids Mass Spectrom. Rev. 33 2014 126 146 24105931
94 Benitez L.V. Allison W.S. The inactivation of the acyl phosphatase activity catalyzed by the sulfenic acid form of glyceraldehyde 3-phosphate dehydrogenase by dimedone and olefins J. Biol. Chem. 249 1974 6234 6243 4371119
95 Qian J. Wani R. Klomsiri C. Poole L.B. Tsang A.W. Furdui C.M. A simple and effective strategy for labeling cysteine sulfenic acid in proteins by utilization of β-ketoesters as cleavable probes Chem. Commun. 48 2012 4091 4093
96 Ellis H.R. Poole L.B. Novel application of 7-chloro-4-nitrobenzo-2-oxa-1,3-diazole to identify cysteine sulfenic acid in the AhpC component of alkyl hydroperoxide reductase Biochemistry 36 1997 15013 15018 9398227
97 Poole L.B. Klomsiri C. Knaggs S.A. Furdui C.M. Nelson K.J. Thomas M.J. Fetrow J.S. Daniel L.W. King S.B. Fluorescent and affinity-based tools to detect cysteine sulfenic acid formation in proteins Bioconjugate Chem. 18 2007 2004 2017
98 Seo Y.H. Carroll K.S. Profiling protein thiol oxidation in tumor cells using sulfenic acid-specific antibodies Proc. Natl. Acad. Sci. U.S.A. 106 2009 16163 16168 19805274
99 Charles R.L. Schröder E. May G. Free P. Gaffney P.R. Wait R. Begum S. Heads R.J. Eaton P. Protein sulfenation as a redox sensor: proteomics studies using a novel biotinylated dimedone analogue Mol. Cell. Proteomics : MCP. 6 2007 1473 1484 17569890
100 Reddie K.G. Seo Y.H. Muse Iii WB. Leonard S.E. Carroll K.S. A chemical approach for detecting sulfenic acid-modified proteins in living cells Mol. Biosyst. 4 2008 521 531 18493649
101 Seo Y.H. Carroll K.S. Facile synthesis and biological evaluation of a cell-permeable probe to detect redox-regulated proteins Bioorg. Med. Chem. Lett 19 2009 356 359 19081252
102 Takanishi C.L. Ma L.H. Wood M.J. A genetically encoded probe for cysteine sulfenic acid protein modification in vivo Biochemistry 46 2007 14725 14732 18020457
103 Takanishi C.L. Wood M.J. A genetically encoded probe for the identification of proteins that form sulfenic acid in response to H2O2 in Saccharomyces cerevisiae J. Proteome Res. 10 2011 2715 2724 21476607
104 Klomsiri C. Nelson K.J. Bechtold E. Soito L. Johnson L.C. Lowther W.T. Ryu S.E. King S.B. Furdui C.M. Poole L.B. Use of dimedone-based chemical probes for sulfenic acid detection evaluation of conditions affecting probe incorporation into redox-sensitive proteins Methods Enzymol. 473 2010 77 94 20513472
105 Leonard S.E. Reddie K.G. Carroll K.S. Mining the thiol proteome for sulfenic acid modifications reveals new targets for oxidation in cells ACS Chem. Biol. 4 2009 783 799 19645509
106 Paulsen C.E. Truong T.H. Garcia F.J. Homann A. Gupta V. Leonard S.E. Carroll K.S. Peroxide-dependent sulfenylation of the EGFR catalytic site enhances kinase activity Nat. Chem. Biol. 8 2011 57 64 22158416
107 Gupta V. Yang J. Liebler D.C. Carroll K.S. Diverse redoxome reactivity profiles of carbon nucleophiles J. Am. Chem. Soc. 139 2017 5588 5595 28355876
108 Fu L. Liu K. Ferreira R.B. Carroll K.S. Yang J. Proteome-wide analysis of cysteine S-sulfenylation using a benzothiazine-based probe Current protocols in protein science 95 2019 e76
109 Niu Y. Chen Z. Jiang Z. Yang Y. Liu G. Cheng X. Jiang Z. Zhang G. Tong L. Tang B. Detection of cysteine sulfenic acid on E. coli proteins with a biotin-benzoboroxole probe ACS Chem. Biol. 18 2023 1351 1359 37260364
110 Yin Q. Huang C. Zhang C. Zhu W. Xu Y. Qian X. Yang Y. In situ visualization and detection of protein sulfenylation responses in living cells through a dimedone-based fluorescent probe Org. Biomol. Chem. 11 2013 7566 7573 24097070
111 Tom C. Crellin J.E. Motiwala H.F. Stone M.B. Davda D. Walker W. Kuo Y.H. Hernandez J.L. Labby K.J. Gomez-Rodriguez L. Jenkins P.M. Veatch S.L. Martin B.R. Chemoselective ratiometric imaging of protein S-sulfenylation Chem. Commun. 53 2017 7385 7388
112 Holmila R.J. Vance S.A. Chen X. Wu H. Shukla K. Bharadwaj M.S. Mims J. Wary Z. Marrs G. Singh R. Molina A.J. Poole L.B. King S.B. Furdui C.M. Mitochondria-targeted probes for imaging protein sulfenylation Sci. Rep. 8 2018 6635 29703899
113 Scinto S.L. Ekanayake O. Seneviratne U. Pigga J.E. Boyd S.J. Taylor M.T. Liu J. Am Ende C.W. Rozovsky S. Fox J.M. Dual-reactivity trans-cyclooctenol probes for sulfenylation in live cells enable temporal control via bioorthogonal quenching J. Am. Chem. Soc. 141 2019 10932 10937 31246462
114 Solingapuram Sai K.K. Chen X. Li Z. Zhu C. Shukla K. Forshaw T.E. Wu H. Vance S.A. Pathirannahel B.L. Madonna M. Dewhirst M.W. Tsang A.W. Poole L.B. Ramanujam N. King S.B. Furdui C.M. [(18)F]Fluoro-DCP, a first generation PET radiotracer for monitoring protein sulfenylation in vivo Redox Biol. 49 2022 102218
115 Yang J. Gupta V. Carroll K.S. Liebler D.C. Site-specific mapping and quantification of protein S-sulphenylation in cells Nat. Commun. 5 2014 4776 25175731
116 Yang J. Gupta V. Tallman K.A. Porter N.A. Carroll K.S. Liebler D.C. Global, in situ, site-specific analysis of protein S-sulfenylation Nat. Protoc. 10 2015 1022 1037 26086405
117 Anastasiou D. Poulogiannis G. Asara J.M. Boxer M.B. Jiang J.K. Shen M. Bellinger G. Sasaki A.T. Locasale J.W. Auld D.S. Thomas C.J. Vander Heiden M.G. Cantley L.C. Inhibition of pyruvate kinase M2 by reactive oxygen species contributes to cellular antioxidant responses Science 334 2011 1278 1283 22052977
118 Agrawal N. Dhakrey P. Pathak S. A comprehensive review on the research progress of PTP1B inhibitors as antidiabetics Chem. Biol. Drug Des. 102 2023 921 938 37232059
119 Sharma B. Xie L. Yang F. Wang W. Zhou Q. Xiang M. Zhou S. Lv W. Jia Y. Pokhrel L. Shen J. Xiao Q. Gao L. Deng W. Recent advance on PTP1B inhibitors and their biomedical applications Eur. J. Med. Chem. 199 2020 112376
120 Olloquequi J. Cano A. Sanchez-López E. Carrasco M. Verdaguer E. Fortuna A. Folch J. Bulló M. Auladell C. Camins A. Ettcheto M. Protein tyrosine phosphatase 1B (PTP1B) as a potential therapeutic target for neurological disorders Biomedicine & pharmacotherapy = Biomedecine & pharmacotherapie 155 2022 113709
121 Lismont C. Nordgren M. Brees C. Knoops B. Van Veldhoven P.P. Fransen M. Peroxisomes as modulators of cellular protein thiol oxidation: a new model system Antioxidants Redox Signal. 30 2019 22 39
122 Lismont C. Revenco I. Li H. Costa C.F. Lenaerts L. Hussein M.A.F. De Bie J. Knoops B. Van Veldhoven P.P. Derua R. Fransen M. Peroxisome-derived hydrogen peroxide modulates the sulfenylation profiles of key redox signaling proteins in flp-in T-REx 293 cells Front. Cell Dev. Biol. 10 2022 888873
123 Wang P. Zhang Q. Li S. Cheng B. Xue H. Wei Z. Shao T. Liu Z.X. Cheng H. Wang Z. iCysMod: an integrative database for protein cysteine modifications in eukaryotes Briefings Bioinf. 22 2021
124 Góth L. Nagy T. Inherited catalase deficiency: is it benign or a factor in various age related disorders? Mutat. Res. 753 2013 147 154 24025477
125 Senis Y.A. Nagy Z. Mori J. Lane S. Lane P. Platelet Src family kinases: a tale of reversible phosphorylation Research and practice in thrombosis and haemostasis 5 2021 376 389 33870023
126 Senis Y.A. Mazharian A. Mori J. Src family kinases: at the forefront of platelet activation Blood 124 2014 2013 2024 25115887
127 Giannoni E. Chiarugi P. Redox circuitries driving Src regulation Antioxidants Redox Signal. 20 2014 2011 2025
128 Tonks N.K. Redox redux: revisiting PTPs and the control of cell signaling Cell 121 2005 667 670 15935753
129 Kemble D.J. Sun G. Direct and specific inactivation of protein tyrosine kinases in the Src and FGFR families by reversible cysteine oxidation Proc. Natl. Acad. Sci. U.S.A. 106 2009 5070 5075 19273857
130 Zhang H. Davies K.J. Forman H.J. TGFβ1 rapidly activates Src through a non-canonical redox signaling mechanism Arch. Biochem. Biophys. 568 2015 1 7 25585026
131 Heppner D.E. Dustin C.M. Liao C. Hristova M. Veith C. Little A.C. Ahlers B.A. White S.L. Deng B. Lam Y.W. Li J. van der Vliet A. Direct cysteine sulfenylation drives activation of the Src kinase Nat. Commun. 9 2018 4522 30375386
132 Yang M. Li W. Harberg C. Chen W. Yue H. Ferreira R.B. Wynia-Smith S.L. Carroll K.S. Zielonka J. Flaumenhaft R. Silverstein R.L. Smith B.C. Cysteine sulfenylation by CD36 signaling promotes arterial thrombosis in dyslipidemia Blood advances 4 2020 4494 4507 32946569
133 Reinhardt C. von Brühl M.L. Manukyan D. Grahl L. Lorenz M. Altmann B. Dlugai S. Hess S. Konrad I. Orschiedt L. Mackman N. Ruddock L. Massberg S. Engelmann B. Protein disulfide isomerase acts as an injury response signal that enhances fibrin generation via tissue factor activation J. Clin. Invest. 118 2008 1110 1122 18274674
134 Cho J. Furie B.C. Coughlin S.R. Furie B. A critical role for extracellular protein disulfide isomerase during thrombus formation in mice J. Clin. Invest. 118 2008 1123 1131 18292814
135 Yang M. Chiu J. Scartelli C. Ponzar N. Patel S. Patel A. Ferreira R.B. Keyes R.F. Carroll K.S. Pozzi N. Hogg P.J. Smith B.C. Flaumenhaft R. Sulfenylation links oxidative stress to protein disulfide isomerase oxidase activity and thrombus formation J. Thromb. Haemostasis : JTH. 21 2023 2137 2150 37037379
136 Kluge M.A. Fetterman J.L. Vita J.A. Mitochondria and endothelial function Circ. Res. 112 2013 1171 1188 23580773
137 Shenouda S.M. Widlansky M.E. Chen K. Xu G. Holbrook M. Tabit C.E. Hamburg N.M. Frame A.A. Caiano T.L. Kluge M.A. Duess M.A. Levit A. Kim B. Hartman M.L. Joseph L. Shirihai O.S. Vita J.A. Altered mitochondrial dynamics contributes to endothelial dysfunction in diabetes mellitus Circulation 124 2011 444 453 21747057
138 Westermann B. Mitochondrial fusion and fission in cell life and death Nat. Rev. Mol. Cell Biol. 11 2010 872 884 21102612
139 Friedman J.R. Lackner L.L. West M. DiBenedetto J.R. Nunnari J. Voeltz G.K. ER tubules mark sites of mitochondrial division Science 334 2011 358 362 21885730
140 Kim Y.M. Youn S.W. Sudhahar V. Das A. Chandhri R. Cuervo Grajal H. Kweon J. Leanhart S. He L. Toth P.T. Kitajewski J. Rehman J. Yoon Y. Cho J. Fukai T. Ushio-Fukai M. Redox regulation of mitochondrial fission protein Drp1 by protein disulfide isomerase limits endothelial senescence Cell Rep. 23 2018 3565 3578 29924999
141 Ježek P. Jabůrek M. Porter R.K. Uncoupling mechanism and redox regulation of mitochondrial uncoupling protein 1 (UCP1) Biochim. Biophys. Acta Bioenerg. 1860 2019 259 269 30414927
142 Chouchani E.T. Kazak L. Jedrychowski M.P. Lu G.Z. Erickson B.K. Szpyt J. Pierce K.A. Laznik-Bogoslavski D. Vetrivelan R. Clish C.B. Robinson A.J. Gygi S.P. Spiegelman B.M. Mitochondrial ROS regulate thermogenic energy expenditure and sulfenylation of UCP1 Nature 532 2016 112 116 27027295
143 Yan L.J. Sumien N. Thangthaeng N. Forster M.J. Reversible inactivation of dihydrolipoamide dehydrogenase by mitochondrial hydrogen peroxide Free Radic. Res. 47 2013 123 133 23205777
144 Yang X. Song J. Yan L.J. Chronic inhibition of mitochondrial dihydrolipoamide dehydrogenase (DLDH) as an approach to managing diabetic oxidative stress Antioxidants 8 2019
145 Svoboda L.K. Reddie K.G. Zhang L. Vesely E.D. Williams E.S. Schumacher S.M. O'Connell R.P. Shaw R. Day S.M. Anumonwo J.M. Carroll K.S. Martens J.R. Redox-sensitive sulfenic acid modification regulates surface expression of the cardiovascular voltage-gated potassium channel Kv1.5 Circ. Res. 111 2012 842 853 22843785
146 Morales Y. Nitzel D.V. Price O.M. Gui S. Li J. Qu J. Hevel J.M. Redox control of protein arginine methyltransferase 1 (PRMT1) activity J. Biol. Chem. 290 2015 14915 14926 25911106
147 Jiang J.L. Zhang X.H. Li N.S. Rang W.Q. Feng Y. Hu C.P. Li Y.J. Deng H.W. Probucol decreases asymmetrical dimethylarginine level by alternation of protein arginine methyltransferase I and dimethylarginine dimethylaminohydrolase activity Cardiovasc. Drugs Ther. 20 2006 281 294 16897158
148 Lüneburg N. Harbaum L. Hennigs J.K. The endothelial ADMA/NO pathway in hypoxia-related chronic respiratory diseases BioMed Res. Int. 2014 2014 501612
149 Wilcox C.S. Asymmetric dimethylarginine and reactive oxygen species: unwelcome twin visitors to the cardiovascular and kidney disease tables Hypertension 59 2012 375 381 22215715
150 Wang K. Luo L. Fu S. Wang M. Wang Z. Dong L. Wu X. Dai L. Peng Y. Shen G. Chen H.N. Nice E.C. Wei X. Huang C. PHGDH arginine methylation by PRMT1 promotes serine synthesis and represents a therapeutic vulnerability in hepatocellular carcinoma Nat. Commun. 14 2023 1011 36823188
151 Luo L. Wu X. Fan J. Dong L. Wang M. Zeng Y. Li S. Yang W. Jiang J. Wang K. FBXO7 ubiquitinates PRMT1 to suppress serine synthesis and tumor growth in hepatocellular carcinoma Nat. Commun. 15 2024 4790 38839752
152 Kaiserova K. Srivastava S. Hoetker J.D. Awe S.O. Tang X.L. Cai J. Bhatnagar A. Redox activation of aldose reductase in the ischemic heart J. Biol. Chem. 281 2006 15110 15120 16567803
153 Shinmura K. Bolli R. Liu S.Q. Tang X.L. Kodani E. Xuan Y.T. Srivastava S. Bhatnagar A. Aldose reductase is an obligatory mediator of the late phase of ischemic preconditioning Circ. Res. 91 2002 240 246 12169650
154 Ramasamy R. Trueblood N. Schaefer S. Metabolic effects of aldose reductase inhibition during low-flow ischemia and reperfusion Am. J. Physiol. 275 1998 H195 H203 9688914
155 Ramasamy R. Oates P.J. Schaefer S. Aldose reductase inhibition protects diabetic and nondiabetic rat hearts from ischemic injury Diabetes 46 1997 292 300 9000707
156 Tian H. Advances in the study on endogenous sulfur dioxide in the cardiovascular system Chin. Med. J. 127 2014 3803 3807 25382339
157 Du S.X. Jin H.F. Bu D.F. Zhao X. Geng B. Tang C.S. Du J.B. Endogenously generated sulfur dioxide and its vasorelaxant effect in rats Acta Pharmacol. Sin. 29 2008 923 930 18664325
158 Bian J.S. Olson K.R. Zhu Y.C. Hydrogen sulfide: biogenesis, physiology, and pathology Oxid. Med. Cell. Longev. 2016 2016 6549625
159 Huang Y. Li Z. Zhang L. Tang H. Zhang H. Wang C. Chen S.Y. Bu D. Zhang Z. Zhu Z. Yuan P. Li K. Yu X. Kong W. Tang C. Jung Y. Ferreira R.B. Carroll K.S. Du J. Yang J. Jin H. Endogenous SO(2)-dependent Smad3 redox modification controls vascular remodeling Redox Biol. 41 2021 101898
160 Song Y. Peng H. Bu D. Ding X. Yang F. Zhu Z. Tian X. Zhang L. Wang X. Tang C. Huang Y. Du J. Jin H. Negative auto-regulation of sulfur dioxide generation in vascular endothelial cells: AAT1 S-sulfenylation Biochem. Biophys. Res. Commun. 525 2020 231 237
161 Wang Y. Wang X. Chen S. Tian X. Zhang L. Huang Y. Tang C. Du J. Jin H. Sulfur dioxide activates Cl(-)/HCO(3) (-) exchanger via sulphenylating AE2 to reduce intracellular pH in vascular smooth muscle cells Front. Pharmacol. 10 2019 313 30971931
162 Wang W. Huang X.R. Canlas E. Oka K. Truong L.D. Deng C. Bhowmick N.A. Ju W. Bottinger E.P. Lan H.Y. Essential role of Smad3 in angiotensin II-induced vascular fibrosis Circ. Res. 98 2006 1032 1039 16556868
163 Iyer D. Zhao Q. Wirka R. Naravane A. Nguyen T. Liu B. Nagao M. Cheng P. Miller C.L. Kim J.B. Pjanic M. Quertermous T. Coronary artery disease genes SMAD3 and TCF21 promote opposing interactive genetic programs that regulate smooth muscle cell differentiation and disease risk PLoS Genet. 14 2018 e1007681
164 Lv B. Peng H. Qiu B. Zhang L. Ge M. Bu D. Li K. Yu X. Du J. Yang L. Tang C. Huang Y. Du J. Jin H. Sulphenylation of CypD at cysteine 104: a novel mechanism by which SO(2) inhibits cardiomyocyte apoptosis Front. Cell Dev. Biol. 9 2021 784799
165 Zhou B. Kreuzer J. Kumsta C. Wu L. Kamer K.J. Cedillo L. Zhang Y. Li S. Kacergis M.C. Webster C.M. Fejes-Toth G. Naray-Fejes-Toth A. Das S. Hansen M. Haas W. Soukas A.A. Mitochondrial permeability uncouples elevated autophagy and lifespan extension Cell 177 2019 299 314.e16 30929899
166 Hara M.R. Agrawal N. Kim S.F. Cascio M.B. Fujimuro M. Ozeki Y. Takahashi M. Cheah J.H. Tankou S.K. Hester L.D. Ferris C.D. Hayward S.D. Snyder S.H. Sawa A. S-nitrosylated GAPDH initiates apoptotic cell death by nuclear translocation following Siah1 binding Nat. Cell Biol. 7 2005 665 674 15951807
167 Ventura M. Mateo F. Serratosa J. Salaet I. Carujo S. Bachs O. Pujol M.J. Nuclear translocation of glyceraldehyde-3-phosphate dehydrogenase is regulated by acetylation Int. J. Biochem. Cell Biol. 42 2010 1672 1680 20601085
168 Molina y Vedia L. McDonald B. Reep B. Brüne B. Di Silvio M. Billiar T.R. Lapetina E.G. Nitric oxide-induced S-nitrosylation of glyceraldehyde-3-phosphate dehydrogenase inhibits enzymatic activity and increases endogenous ADP-ribosylation J. Biol. Chem. 267 1992 24929 24932 1281150
169 Grant C.M. Quinn K.A. Dawes I.W. Differential protein S-thiolation of glyceraldehyde-3-phosphate dehydrogenase isoenzymes influences sensitivity to oxidative stress Mol. Cell Biol. 19 1999 2650 2656 10082531
170 Zaffagnini M. Fermani S. Calvaresi M. Orrù R. Iommarini L. Sparla F. Falini G. Bottoni A. Trost P. Tuning cysteine reactivity and sulfenic acid stability by protein microenvironment in glyceraldehyde-3-phosphate dehydrogenases of Arabidopsis thaliana Antioxidants Redox Signal. 24 2016 502 517
171 Nakajima H. Amano W. Fujita A. Fukuhara A. Azuma Y.T. Hata F. Inui T. Takeuchi T. The active site cysteine of the proapoptotic protein glyceraldehyde-3-phosphate dehydrogenase is essential in oxidative stress-induced aggregation and cell death J. Biol. Chem. 282 2007 26562 26574 17613523
172 Samson A.L. Knaupp A.S. Kass I. Kleifeld O. Marijanovic E.M. Hughes V.A. Lupton C.J. Buckle A.M. Bottomley S.P. Medcalf R.L. Oxidation of an exposed methionine instigates the aggregation of glyceraldehyde-3-phosphate dehydrogenase J. Biol. Chem. 289 2014 26922 26936 25086035
173 Wang Q. Woltjer R.L. Cimino P.J. Pan C. Montine K.S. Zhang J. Montine T.J. Proteomic analysis of neurofibrillary tangles in Alzheimer disease identifies GAPDH as a detergent-insoluble paired helical filament tau binding protein Faseb. J. : official publication of the Federation of American Societies for Experimental Biology 19 2005 869 871
174 Cumming R.C. Schubert D. Amyloid-beta induces disulfide bonding and aggregation of GAPDH in Alzheimer's disease Faseb. J. : official publication of the Federation of American Societies for Experimental Biology 19 2005 2060 2062
175 Itakura M. Nakajima H. Kubo T. Semi Y. Kume S. Higashida S. Kaneshige A. Kuwamura M. Harada N. Kita A. Azuma Y.T. Yamaji R. Inui T. Takeuchi T. Glyceraldehyde-3-phosphate dehydrogenase aggregates accelerate amyloid-β amyloidogenesis in alzheimer disease J. Biol. Chem. 290 2015 26072 26087 26359500
176 Dotsey E.Y. Jung K.M. Basit A. Wei D. Daglian J. Vacondio F. Armirotti A. Mor M. Piomelli D. Peroxide-dependent MGL sulfenylation regulates 2-AG-mediated endocannabinoid signaling in brain neurons Chem. Biol. 22 2015 619 628 26000748
177 Panikashvili D. Mechoulam R. Beni S.M. Alexandrovich A. Shohami E. CB1 cannabinoid receptors are involved in neuroprotection via NF-kappa B inhibition J. Cerebr. Blood Flow Metabol. : official journal of the International Society of Cerebral Blood Flow and Metabolism 25 2005 477 484
178 Panikashvili D. Shein N.A. Mechoulam R. Trembovler V. Kohen R. Alexandrovich A. Shohami E. The endocannabinoid 2-AG protects the blood-brain barrier after closed head injury and inhibits mRNA expression of proinflammatory cytokines Neurobiol. Dis. 22 2006 257 264 16364651
179 Bajaj S. Jain S. Vyas P. Bawa S. Vohora D. The role of endocannabinoid pathway in the neuropathology of Alzheimer's disease: can the inhibitors of MAGL and FAAH prove to be potential therapeutic targets against the cognitive impairment associated with Alzheimer's disease? Brain Res. Bull. 174 2021 305 322 34217798
180 Wang M. Kaufman R.J. The impact of the endoplasmic reticulum protein-folding environment on cancer development Nat. Rev. Cancer 14 2014 581 597 25145482
181 Maly D.J. Papa F.R. Druggable sensors of the unfolded protein response Nat. Chem. Biol. 10 2014 892 901 25325700
182 Hourihan J.M. Moronetti Mazzeo L.E. Fernández-Cárdenas L.P. Blackwell T.K. Cysteine sulfenylation directs IRE-1 to activate the SKN-1/Nrf2 antioxidant response Mol. Cell 63 2016 553 566 27540856
183 Warburg O. Wind F. Negelein E. The metabolism of tumors in the body J. Gen. Physiol. 8 1927 519 530 19872213
184 Conway M.E. Coles S.J. Islam M.M. Hutson S.M. Regulatory control of human cytosolic branched-chain aminotransferase by oxidation and S-glutathionylation and its interactions with redox sensitive neuronal proteins Biochemistry 47 2008 5465 5479 18419134
185 Francois L. Boskovic P. Knerr J. He W. Sigismondo G. Schwan C. More T.H. Schlotter M. Conway M.E. Krijgsveld J. Hiller K. Grosse R. Lichter P. Radlwimmer B. BCAT1 redox function maintains mitotic fidelity Cell Rep. 41 2022 111524
186 Grallert A. Boke E. Hagting A. Hodgson B. Connolly Y. Griffiths J.R. Smith D.L. Pines J. Hagan I.M. A PP1-PP2A phosphatase relay controls mitotic progression Nature 517 2015 94 98 25487150
187 Holder J. Poser E. Barr F.A. Getting out of mitosis: spatial and temporal control of mitotic exit and cytokinesis by PP1 and PP2A FEBS Lett. 593 2019 2908 2924 31494926
188 Hirota T. Kunitoku N. Sasayama T. Marumoto T. Zhang D. Nitta M. Hatakeyama K. Saya H. Aurora-A and an interacting activator, the LIM protein Ajuba, are required for mitotic commitment in human cells Cell 114 2003 585 598 13678582
189 Miralaei N. Majd A. Ghaedi K. Peymani M. Safaei M. Integrated pan-cancer of AURKA expression and drug sensitivity analysis reveals increased expression of AURKA is responsible for drug resistance Cancer Med. 10 2021 6428 6441 34337875
190 Byrne D.P. Shrestha S. Galler M. Cao M. Daly L.A. Campbell A.E. Eyers C.E. Veal E.A. Kannan N. Eyers P.A. Aurora A regulation by reversible cysteine oxidation reveals evolutionarily conserved redox control of Ser/Thr protein kinase activity Sci. Signal. 13 2020
191 Lim D.C. Joukov V. Rettenmaier T.J. Kumagai A. Dunphy W.G. Wells J.A. Yaffe M.B. Redox priming promotes Aurora A activation during mitosis Sci. Signal. 13 2020
192 Tsuchiya Y. Byrne D.P. Burgess S.G. Bormann J. Baković J. Huang Y. Zhyvoloup A. Yu B.Y.K. Peak-Chew S. Tran T. Bellany F. Tabor A.B. Chan A.E. Guruprasad L. Garifulin O. Filonenko V. Vonderach M. Ferries S. Eyers C.E. Carroll J. Skehel M. Bayliss R. Eyers P.A. Gout I. Covalent Aurora A regulation by the metabolic integrator coenzyme A Redox Biol. 28 2020 101318
193 Du R. Huang C. Liu K. Li X. Dong Z. Targeting AURKA in Cancer: molecular mechanisms and opportunities for Cancer therapy Mol. Cancer 20 2021 15 33451333
194 Skoko J.J. Cao J. Gaboriau D. Attar M. Asan A. Hong L. Paulsen C.E. Ma H. Liu Y. Wu H. Harkness T. Furdui C.M. Manevich Y. Morrison C.G. Brown E.T. Normolle D. Spies M. Spies M.A. Carroll K. Neumann C.A. Redox regulation of RAD51 Cys319 and homologous recombination by peroxiredoxin 1 Redox Biol. 56 2022 102443
195 Dong G. Mao Q. Xia W. Xu Y. Wang J. Xu L. Jiang F. PKM2 and cancer: the function of PKM2 beyond glycolysis Oncol. Lett. 11 2016 1980 1986 26998110
196 Christofk H.R. Vander Heiden M.G. Harris M.H. Ramanathan A. Gerszten R.E. Wei R. Fleming M.D. Schreiber S.L. Cantley L.C. The M2 splice isoform of pyruvate kinase is important for cancer metabolism and tumour growth Nature 452 2008 230 233 18337823
197 Dombrauckas J.D. Santarsiero B.D. Mesecar A.D. Structural basis for tumor pyruvate kinase M2 allosteric regulation and catalysis Biochemistry 44 2005 9417 9429 15996096
198 Irokawa H. Numasaki S. Kato S. Iwai K. Inose-Maruyama A. Ohdate T. Hwang G.W. Toyama T. Watanabe T. Kuge S. Comprehensive analyses of the cysteine thiol oxidation of PKM2 reveal the effects of multiple oxidation on cellular oxidative stress response Biochem. J. 478 2021 1453 1470 33749780
199 Sigismund S. Avanzato D. Lanzetti L. Emerging functions of the EGFR in cancer Mol. Oncol. 12 2018 3 20 29124875
200 Sabbah D.A. Hajjo R. Sweidan K. Review on epidermal growth factor receptor (EGFR) structure, signaling pathways, interactions, and recent updates of EGFR inhibitors Curr. Top. Med. Chem. 20 2020 815 834 32124699
201 Truong T.H. Ung P.M. Palde P.B. Paulsen C.E. Schlessinger A. Carroll K.S. Molecular basis for redox activation of epidermal growth factor receptor kinase Cell Chem. Biol. 23 2016 837 848 27427230
202 Gupta V. Carroll K.S. Profiling the reactivity of cyclic C-nucleophiles towards electrophilic sulfur in cysteine sulfenic acid Chem. Sci. 7 2016 400 415 26819701
203 Gupta V. Carroll K.S. Rational design of reversible and irreversible cysteine sulfenic acid-targeted linear C-nucleophiles Chem. Commun. 52 2016 3414 3417
204 Li D. Ambrogio L. Shimamura T. Kubo S. Takahashi M. Chirieac L.R. Padera R.F. Shapiro G.I. Baum A. Himmelsbach F. Rettig W.J. Meyerson M. Solca F. Greulich H. Wong K.K. BIBW2992, an irreversible EGFR/HER2 inhibitor highly effective in preclinical lung cancer models Oncogene 27 2008 4702 4711 18408761
205 Fry D.W. Bridges A.J. Denny W.A. Doherty A. Greis K.D. Hicks J.L. Hook K.E. Keller P.R. Leopold W.R. Loo J.A. McNamara D.J. Nelson J.M. Sherwood V. Smaill J.B. Trumpp-Kallmeyer S. Dobrusin E.M. Specific, irreversible inactivation of the epidermal growth factor receptor and erbB2, by a new class of tyrosine kinase inhibitor Proc. Natl. Acad. Sci. U.S.A. 95 1998 12022 12027 9751783
