Widespread interest in phosphorylation PTM has led to the development of numerous covalent chemical strategies for its analysis. A common theme among these methods involve, the selective enrichment of phosphopeptides from a complex mixture of peptides through a sequence of chemical reactions, followed by the subsequent analysis of isolated phosphopeptides by mass spectrometry.

One of the first covalent chemical approaches was developed by Aebersold et al., in 2001. They developed a multistep chemical method consisting of six reaction steps (Figure 3A) and applied it on tryptic digests containing various phosphorylated peptides (Zhou et al., 2001). First, the amino groups of tryptic digested peptides were protected using tert-butyl-dicarbonate (tBoc) chemistry (step 1). This was done to prevent potential intermolecular and intramolecular reactions in subsequent steps. Once the amino groups were protected, the carboxylic groups were protected using a carbodiimide-catalyzed condensation reaction with excess amines to form amides, and phosphoramidates, resulting from the modification of phosphates by the coupling reaction (step 2). Free phosphate groups were regenerated by acid hydrolysis (step 3, carboxylic acids remain protected), followed by a subsequent carbodiimide-catalyzed condensation reaction of regenerated phosphate groups with cysteamine (step 4). Internal disulfides of cysteamine were reduced prior to solid-phase capture of peptides through the reaction between the free sulfhydryl group of cysteamine with the iodoacetyl group on the resin (step 5). Acid cleavage of the phosphoramidate bonds led to the isolation of free phosphopeptides which were subsequently analyzed using LC-MS/MS. It is noteworthy to state that the carboxylate groups remained blocked at the end of the reactions (step 2). By utilizing various phosphopeptides and yeast cell lysates, Aebersold and coworkers demonstrated that their method was equally applicable in isolating serine-, threonine- and tyrosine-phosphorylated peptides from a highly complex peptide mixture.

By far, the most prevalent method for phosphopeptide analysis involves β-elimination of phosphate moiety under basic conditions by appropriate affinity tags (Figures 3B,C). The central principle is based on the fact that phosphate moieties on serine or threonine containing peptide residues are labile at high pH values (Fadden and Haystead, 1995; Jaffe et al., 1998). Under basic conditions, the phosphate moiety on phosphoseryl or phosphothreonyl residues can undergo β-elimination to form dehydroalanine, an alpha-beta unsaturated Michael acceptor which can readily react with a nucleophile such as ethanedithiol. Thiol containing modified residues are coupled to biotin through a maleimide or iodoalkyl linked biotin moiety (Figure 3B). Biotinylated peptides are enriched using avidin column chromatography followed by the release of enriched peptide under acidic conditions for analysis by MS (Oda et al., 2001). One major drawback associated with the use of biotin as an affinity handle is the inefficient recovery of tagged residues after affinity column chromatography. To address this difficulty, Chait et al., utilized a thiol containing affinity reagent instead of biotin (McLachlin and Chait, 2003). Reduction with dithiothreitol led to the recovery of modified phosphorylated peptides (Figure 3C). While this approach was effective in addressing issues associated with recovery, it did lead to side reactions where unmodified cysteine residues were captured by the affinity tag (∼1–2%).

In 2005, Aebersold et al. described an improved strategy for phosphopeptide enrichment and subsequent analysis. This improvement arises simply from the capture of phosphopeptides using a one-step chemical reaction between soluble amine-functionalized polymer (Generation-5 polyamidoamine dendrimer) and phosphate moiety on peptides to generate phosphoramidates (Figure 3D). This reaction led to peptide conjugates that were physically larger than unmodified peptides, thus were readily separated from other peptides and excess reagents using a 5-KDa filtration spin column. Notably, this approach was applicable to all classes of phosphopeptides including phosphotyrosine containing peptides. Using this approach, Aebersold and coworkers identified over 150 phosphorylation sites (Tao et al., 2005).

Acetylation on both the lysine side chains and the N-terminus is a common protein modification that affects an estimated 80% of all human protein species to varying degrees. However, the enrichment of acetylated peptide residues is difficult because acetylated amines cannot be easily derivatized. Normally, the lysine acetylome is characterized and quantified by enrichment with antibodies against acetylated lysine residues and subsequent mass spectrometric analysis.

Chemical methods of profiling acetylation often rely on the stoichiometric lysine or N-termini acetylation occupancy (i.e., the proportion occupied [acetylated] versus unoccupied [unacetylated]). Recently, several groups have reported methods for determining the occupancy of lysine acetylation sites, which measure the ratio of endogenously acetylated lysine to unmodified lysine (i.e., stoichiometry or occupancy) (Meyer et al., 2016). In stoichiometric experiments, mass spectrometric recordings determine the ratio of endogenous light acetyl groups to isotope-labeled heavy acetyl groups, which are generated in vitro by quantitative acetylation of all unmodified lysines after the cells have been harvested (Figure 4A). In this workflow, protein lysates were first incubated three times with acetic anhydride-d₆ in order to acetylate unmodified lysine residues. Second, samples were digested with endoproteinase Glu-C, followed by fractionation of the proteolytic peptides by basic reverse phase chromatography. Finally, peptides were analyzed by LC-MS.

In 2003 Gevaert et al. described a diagonal procedure for the isolation of N-terminal acetylated peptides (Figure 4B). In this new process, free cysteine groups were first blocked by alkylation followed by acetylation of amino groups in proteins and then digestion with trypsin (Gevaert et al., 2003). After reverse phase fractionation (RP) of the generated peptide mixture, internal peptides were blocked using 2,4,6-trinitrobenzene sulfonic acid (TNBS). During the modification, internal peptides showed a strong hydrophobic shift and therefore segregated from the unchanged N-terminal peptides during a second identical separation step. N-terminal peptides were then collected specifically for further liquid chromatography (LC)-MS/MS analysis. Using this approach, Gavaert and coworkers were able to identify 264 proteins and 78 in vivo acetylated proteins in a cytosolic and membrane skeleton fraction of human thrombocytes.

Of the various types of post-translational modifications, glycosylation is the most chemically and biosynthetically complex. The complexity of glycosylation results from two factors: the diversity of the building blocks and the diverse ways in which oligosaccharides can be built up within the cell. Furthermore, the non-templated biosynthesis generates enormous diversity and heterogeneity within the cellular glycome.

Chemical methods have long been used as alternative tools for the detection of glycans. Many of these approaches are destructive (Rutland, 2008) involving the use of periodic acid and alcian blue to detect sialic acid and polysialic acid, respectively. Both the reagents oxidize sialic acid to decarbonylated sialic acid and cause permanent damage to the treated carbohydrate chains thus preventing downstream applications. Lately, there have been powerful approaches to glycan detection that are variations of originally developed methods (Kayser et al., 1992; Dube and Bertozzi, 2003). One widely used approach is based on chemoenzymatic glycan labeling (CeGL). In this approach, Bertozzi and coworkers used a recombinant glycosyltransferase to transfer a monosaccharide analog from a nucleotide sugar donor to a specific glycan acceptor directly in cell lysates or on the cell surface (Dube and Bertozzi, 2003). The transferred monosaccharide was equipped with a reactive handle that was further derivatized to incorporate a detection probe (Figure 5A).

Inspired by the work of Torres and Hart, who incorporated GalT1 radiolabeled Uridine diphosphate N-acetylglucosamine (UDP-[3H]-Gal) into N-acetylglucosamine (GlcNac) residues (Torres and Hart, 1984), Hsieh-Wilson and coworkers used a mutated form of β1,4-galactosyltransferase (GalT1 Y289L) to facilitate the transfer of the N-Acetylglucosamine (GlcNAc) with an unnatural chemical ketone handle (Khidekel et al., 2003) (Figure 5B). The ketone functional group was labeled with an aminooxy-biotin probe for enrichment by streptavidin conjugates followed by analysis using LC-MS/MS (Figure 5B). Subsequently, the substrate scope of this GalT1 mutant was expanded to include an azide-bearing UDP-GlcNAcs as a nucleotide sugar donor that led to the attachment of affinity tag by copper(I)-catalyzed azide-alkyne cycloaddition (CuAAC), thus reducing the unspecific labeling in crude cell lysates containing other aldehydes and ketones, a disadvantage in connection with the reactive ketone/aminooxy pair.

Recently, Hsieh-Wilson and colleagues developed a method to quantify O-GlcNAc glycosylation stoichiometry and to understand dynamics. In this procedure, GalT1 was used to incorporate a ketogalactose residue into O-GlcNAcylated proteins, which was then reacted with aminooxy-functionalized polyethylene glycol (PEG) mass tag (Rexach et al., 2010). In this way, tags with defined molecular weights were attached to O-GlcNAcylated proteins, and the stoichiometry of O-GlcNAc modifications was resolved by SDS-PAGE by simply examining mass-shifted bands.

Although CeGL methods are relatively new and have a wide range of applications, one of the greatest challenges for the future expansion of CeGL is the identification of additional glycosyltransferases that have strict acceptor specificity but also show promiscuity for modified nucleotide sugar donors that can be used for glycan labeling in a cellular system.

Protein methylation typically occurs on lysine or arginine and, to a lesser extent, on histidine amino acid residues in proteins (Figure 6A). Lysine can be methylated one, two or three times by lysine methyltransferases, with different degrees of methylation leading to different functions and locations within a cell (Han et al., 2019). Arginine can be methylated once (monomethylated arginine) or twice, either with both methyl groups on one terminal nitrogen (asymmetric dimethylarginine) or one on both nitrogen’s (symmetric dimethylarginine), by enzyme arginine methyltransferases (PRMTs) (Yang and Bedford, 2013). Histidine can be methylated at two different positions either 1st or 3^rd position and recently has been shown to play important role in biological processes (Jakobsson, 2021) (Figure 6A).

Compared to other PTMs, the development of chemical tools for the detection of methylated residues is quite complicated, since the addition of a methyl group does not lead to a significant change in the physicochemical properties (charge and size) of proteins or peptides. So far there is only one chemical method for the pan-selective covalent modification of monomethyllysines (KMe) developed by our group. In 2020, our laboratory developed a triazenation reaction, which enables chemoselective labeling of KMe and enrichment from a complex mixture followed by subsequent chemically triggered decoupling to generate the native, unchanged coupling partners under mild conditions (Figure 6B) (Nwajiobi et al., 2021). By using electron-donating groups (EDG) on arene diazonium salts, selective modification of KMe peptides to triazenation products was observed within 30 min at pH 7.0 under mild reaction conditions. Although arene diazonium salts have been reported to react with primary amines, the reaction is reversible thus does not allow the enrichment and analysis of KMe peptides (Sonousi and Crich, 2015; Sengupta and Chandrasekaran, 2019). Similarly, the reaction of arene diazonium salts with tyrosine generated azo-coupling product, but it requires electron-withdrawing group (EWG) on arene diazonium ring and a slightly basic pH (∼9.0) conditions for efficient labeling (Schlick et al., 2005). Under our reaction conditions at pH 7 with ERG on the arene diazonium ions, we observed the chemoselective, pan-specific and near-quantitative modification of KMe peptides only within 30 min. In an attempt to achieve a traceless release of triazene products, we observed that the incubation of modified peptides in 10% trifluoroacetic acid (TFA) in water led to the decoupling of triazene products to unmodified KMe peptides in a traceless manner. This observation led to the application of this strategy for solid-phase capture of monomethyl lysine KMe peptides and the release of highly pure peptide samples for proteomics (Figure 6C). To this effect, arene diazonium ion functionalized resin was used to selectively capture and release KMe peptides from a complex tryptic digest mixture of myoglobin, cytochrome c (cyt c), and insulin.

In contrast to monomethyl lysine KMe, there are no covalent chemical methods for selective labeling of dimethyl and trimethyl lysines (Figure 6D). Existing non-covalent methods include the use of antibodies, mass spectrometry, stable isotope labeling techniques such as SILAC, and biosynthetic methods. Similarly, there are no covalent chemical methods for labeling methylated arginine and histidine residues.

Citrullination, also known as deimination, is an irreversible reaction that converts the guanidinium group of arginine to an ureido group, which results in the loss of both a positive charge and two potential hydrogen bond donors (Slade et al., 2014). The citrullination reaction is catalyzed by the protein arginine deiminases (PADs) and inhibits the methylation of the same arginine residue (Slade et al., 2014). The PADs have been documented to play a role in eukaryotic gene expression and are implicated in human diseases such as rheumatoid arthritis (Schellekens et al., 1998), multiple sclerosis (Musse et al., 2008), cancer (Chang et al., 2009) and inflammatory diseases (Kinloch et al., 2008). The detection, isolation, and enrichment of citrullinated proteins are challenging using MS and MS/MS analysis because the small 1 Da increase in mass imparted to the citrullinated protein is negligible. However, the following chemical tools have been developed for studying protein citrullination.

The color development reagent (COLDER) assay is commonly used to monitor citrullination levels (Knipp and Vašák, 2000). Many research groups have used the same approach to study arginine mimetics, peptides, and even proteins (Chumanevich et al., 2011; Willis et al., 2011; Bicker et al., 2012; McElwee et al., 2012). In this cost-effective and fast assay, the ureido group reacts with 2,3-butanedione monoxime to form an imidazoline which is stabilized by thiosemicarbazide (TSC) and Fe³⁺ facilitating rapid color development at 95^OC (Knipp and Vašák, 2000) (Figure 7A). This method has a relatively high limit of detection (0.6 nM) thus unable to detect low abundant citrullinated proteins in the proteome. However, it has been used for evaluating PAD invitro activity (Slade et al., 2014). Proost et al. developed a chemically modified antibody-based method to specifically detect and quantify citrullination on proteins (Moelants et al., 2011). First, using antipyrine and 2,3-butanedione at low pH, the citrullinated proteins were chemically modified and subsequently detected and quantified by specific antibodies raised against a modified citrulline-containing peptide (Figure 7B). Citrulline-containing chemokine, i.e., CXCL8, was used as a model protein. This assay was developed into a commercially available Anti-Citrulline (Modified) Detection Kit (ACM kit; Millipore; Billerica, MA).

Fleckenstein and coworkers described the details of the chemistry behind the reaction of the ureido group of citrulline with 2,3-butanedione in the presence of antipyrine N-butyl urea as a model substance (Holm et al., 2006). They proposed that N-butyl urea reacts with 2,3-butanedione affording the reactive imidazoline derivative, which is further trapped by antipyrine as a nucleophile (Figures 7B,C). Also, they contended that acidic conditions are required not only to catalyze the nucleophilic addition of antipyrine but also to prevent arginine residues from reacting. Tutturen et al. demonstrated that phenylglyoxal modified citrulline under acidic conditions. They made a chemically cleavable phenylglyoxal derivative on PL-DMA resin and utilized it to enrich citrullinated proteins (Tutturen et al., 2010) from a complex mixture (Figure 7D). However, this approach is not chromophoric thus cannot be employed to visualize proteins. To surmount this challenge, Thompson and colleagues synthesized phenylglyoxal compounds with an azido group (Rh-PG) on the meta position for further reaction with alkyne modified rhodamine via click chemistry (Bicker et al., 2012) (Figure 7D). The Rh-PG labeling method was found to be better than the ACM kit: takes significantly less time (∼3 h versus ≥25 h), requires fewer steps (6 versus 12), and has simpler analysis (fluorescent imaging versus Western blotting). These studies showed that the Rh-PG probe could be a powerful chemical probe for detecting protein citrullination and for providing powerful insights into the role of specific PAD substrates in PAD-related diseases.

Lipidation is a common post-translational modification in which lipid moieties are covalently attached to proteins. It has been reported that at least six types of lipids, including fatty acids, isoprenoids, sterols, phospholipids, glycosylphosphatidylinositol (GPI) anchors and lipid-derived electrophiles (LDEs) can covalently modify proteins (Chen et al., 2018). These saturated and unsaturated fatty acids can attach to the lysine, serine or more commonly cysteine residues of proteins, in a process known as fatty acylation (Chen et al., 2018). Fatty acylation significantly increases the hydrophobicity of proteins leading to changes in their conformation, stability, membrane association, localization, trafficking, and binding affinity to their co-factors (Chen et al., 2018). Thus, the lipidation of proteins regulates protein-membrane interactions, protein-protein interactions, enzymatic activities, and protein stability (Jiang et al., 2018). However, aberrant protein lipidation is implicated in various diseases, including cancers, neurological disorders, and metabolic disorders (Chen et al., 2018). Hence, chemicals methods to detect lipidation PTM are essential to fully understand their roles in varying pathologies.

Drisdel and Green described a method popularly known as Acyl-Biotin Exchange (ABE). Acyl-biotin exchange (ABE) remains one of the earliest chemicals methods developed to detect S-acylation of cysteines, based on the high reactivity of the thioester bond, which can be readily removed by weak bases, such as hydroxylamine (Drisdel and Green, 2004). It involves three sequential steps: blockade of free thiols with N-ethylmaleimide (NEM) followed by hydroxylamine-mediated cleavage of the palmitoyl-thioester bond to generate free thiols which are biotinylated with sulfhydryl-reactive reagents and enriched using streptavidin affinity purification (Figure 8A). Wan et al., described a variation of this approach by using N-[6-(Biotinamido) hexyl]-3`-(2′-pyridyldithio) propionamide (Biotin-HPDP), instead of the 1-BiotinaMido-4-[4`-(MaleiMidoMethyl) cyclohexanecarboxaMido] butane (Biotin-BMCC) originally used by Drisdel and Green involving coupling of thiol with a maleimide moiety. Unlike Biotin-BMCC, Biotin–HPDP forms disulfide bonds with the thiols exposed following hydroxylamine treatment and facilitates an easy uncoupling of biotinylated proteins bound to the streptavidin–agarose affinity matrix, through β-mercaptoethanol (BME)-mediated cleavage of the biotin-Cys linkage (Wan et al., 2007) (Figure 8A). Also, Biotin–HPDP is more thiol specific as compared to maleimide of Biotin-BMCC that can react with the amine of lysine side chains. The ABE provides a highly sensitive method for the visualization and quantitative estimate of protein S-acylation thus enabling the global analysis of protein S-palmitoylation in yeast (Lei et al., 2022). S-acylated species usually do not move differently from the non-acylated proteins on protein gel electrophoresis since S-acylation does not alter the protein mass significantly and the electric charge remains the same on both species. Thus, it is difficult to directly detect the levels and the stoichiometry of acylation of a protein of interest by gel electrophoresis (Chen et al., 2018). To overcome this challenge and efficiently measure the endogenous levels of S-fatty acylation, Percher et al. developed a mass-tag labeling method termed acyl-PEG exchange (APE) involving cleavage of thioesters by hydroxylamine to generate free thiols followed by reaction with maleimide-functionalized polyethylene glycol reagents (Percher et al., 2016). The mass-tag induces a shift on the S-acylated proteins thus target proteins can be monitored by Western blot without the need for metabolic labeling or affinity enrichment of proteins (Figure 8B). With the APE approach, the endogenous levels of interferon-induced transmembrane protein 3 (IFITM3) S-fatty acylation were characterized. Equally, Forrester et al., found an alternative to the use of biotin and repeated protein precipitations in ABE, which detects S-acetylated proteins by using resin-assisted captured termed acyl-RAC (Forrester et al., 2011). In this approach, the free thiols were first blocked with MMTS. Secondly, thioesters were cleaved with neutral hydroxylamine (NH₂OH) to generate thiols followed by capturing on thiol-reactive Sepharose resin. Employing “on-resin” proteolysis on the captured proteins, individual sites of S-acylation were identified. The method is fast and amenable to mass spectrometry techniques for identifying S-acylation sites.

One of the limitations of both ABE and APE approaches is the inability to identify the original acyl groups attached to the proteins since they are lost during the exchange step. Several research groups have surmounted this challenge using reporter groups that can be taken up by the cell, converted to acyl-CoAs and serve as lipid donors in cells. Hang et al., employed ω-azido-fatty acids as probes for the rapid detection of protein fatty acylation in mammalian cells (Hang et al., 2007). After the metabolic labeling of mammalian proteins with the reporter groups, a bio-orthogonal reaction with a phosphine-biotin reagent via the Staudinger ligation was carried out to detect fatty-acylated protein. The labeled proteins were rapidly visualized with streptavidin-biotin affinity binding. Using the approach, monounsaturated fatty acylation, prenylation and N- or O-acylation, which could not be detected by acyl exchange methods were detected. Also, Charron et al. widened the applicability of this approach by using fluorescent chemical reporters that afforded a rapid biochemical analysis as well as imaging of protein fatty-acylation in mammalian cells (Charron et al., 2009).

S-nitrosylation is a posttranslational modification involving a covalent bond between a cysteine thiol and an NO equivalent. S-nitrosylation has been reported to play important functions in different biological processes including blood flow (Singel and Stamler, 2005), phosphorylation (Hess and Stamler, 2012), neuronal function (Lipton et al., 1993), transcriptional regulation (Marshall et al., 2000) and nitric oxide cellular signal transduction (Miersch and Mutus, 2005) while dysregulation of S-nitrosylation has been implication in cardiovascular diseases (Chung and David, 2010), cancers (Wang, 2012) and neurodegenerative diseases (Chung and David, 2010).

The detection of S-nitrosylation poses a challenge due to the labile nature of S-nitrosothiols. Wang and Xian developed a fast reductive one-step ligation reaction that targets SNO moieties converting the unstable S-nitrosothiols into stable sulfenamide conjugates by phosphine esters (Wang and Xian, 2008) (Figure 9A). The reaction mechanism is believed to be similar to the Staudinger ligation of azides (Lin et al., 2005). The same research group developed a traceless version of this reaction in which an amide bond is formed between S-nitrosothiols and phosphine ester/thioester conjugates without the phosphine oxide moiety (Zhang et al., 2009a) (Figure 9A). This approach was inspired by the well-known traceless Staudinger ligation (Nilsson et al., 2000). They also reported an unexpected bis-ligation reaction of RSNOs triaryl substituted phosphine-thioester substrates that converts unstable primary RSNOs to stable disulfide-iminophosphorane products in good yields under mild conditions (Zhang et al., 2009b) (Figure 9B). In contrast to the sulfenamide products from the reductive ligation of RSNOs, disulfide-iminophosphorane products were not sensitive to phosphine reagents.

They went further to develop a reductive ligation-mediated disulfide formation between RSNOs and phosphine-thioester substrates which selectively converts unstable RSNOs to stable disulfides (Zhang et al., 2010) (Figure 9C). This approach was used in identifying SNO proteins in cell extracts. To increase the utility of the reductive ligation to detect S-nitrosylation, Pan et al. developed the first generation of fluorescent probes for SNO, which was based on SNO-mediated oxidation of phosphine substrates in which a coumarin-phosphine compound was used to measure the concentration of S-nitrosothiols (Pan et al., 2009) (Figure 9D). The coumarin-phosphine serves as a fluorogenic dye that is activated by S-nitrosothiols. This assay showed greater sensitivity to SNO but other reactive sulfur species could undergo oxidation as well (Zhang et al., 2014). To circumvent this, Zhang et al. developed a more sensitive SNO specific fluorogenic probe based on reductive ligation of SNO called SNOP1 (Zhang et al., 2014) (Figure 9E). The probes were based on the well-known principle that acylation on many fluorophores can quench the fluorescence and de-acylation can reform the fluorescent species (Li et al., 2014). Thus, hydroxyl (-OH) sensitive fluorophore was acylated to the triaryl phosphine probe and upon reacting with SNO, the fluorophore is released serving as a more sensitive means of probing for SNO. This fluorogenic probe works via a similar pathway as mentioned in Figure 9A. The applicability of this approach was demonstrated for the detection of GSNO in diluted deproteinized bovine plasma thus excellent for sensitive analysis of GSNO in biological samples (Li et al., 2014).

King and coworkers detailed the first reaction of triarylphosphines with RSNOs in water (Bechtold et al., 2010) (Figure 9F). The water-soluble triarylphosphine TXPTS reacted directly with S-nitrosothiol residues and generated stable S-alkylphosphonium ions, TXPTS-derived aza-ylide as well as TXPTS oxide. The formation of stable S-alkylphosphonium ions is due to the presence of sulfonate groups on the aromatic ring of triaryl phosphine. They showed efficient labeling of S-nitrothiols, S-nitrosoglutathione and a mutated peroxiredoxin, S-nitrosated C165S alkyl hydroperoxide reductase (C165S AhpC-SNO). This represented the first covalent detection and labeling of S-nitrothiols on proteins in the buffer.

Shao et al., developed a fluorogenic probe, termed PSNO, for the in-situ imaging of protein S-nitrosylation in live cells based on phosphine-mediated reductive ligation. In this reaction, endogenous thiols were first blocked followed by conversion of SNO to free thiol by phosphine-mediated reductive ligation and labeling with maleimide PEG. S-nitrosoglutathione (GSNO) was employed as a model substrate and an array of phosphine-based reagents were tested for their reactivity towards it. They found that PSNO was highly sensitive and specific towards GSNO in buffer, and they successfully imaged protein S-nitrosylated GAPDH in live cells (Shao et al., 2017).

Li et al. took advantage of the finding that the sulfenamide intermediate produced from reductive ligation can be over-reduced in the presence of excess phosphine reagent resulting in the formation of free thiol from the starting S-nitrosothiols. They developed a strategy to target these free thiols (Li et al., 2012) (Figure 9G). To forestall cross-reactivity with endogenous thiols, N-ethyl maleimide (NEM) was used to block all free thiols. Following thiol blockade, triphenylphosphine ester derivatives were used to selectively reduce SNO bonds in proteins. They demonstrated that triphenylphosphine ester derivatives are specific reductants of SNO in complex biological samples without the reduction of protein disulfides or protein thiols modified by hydrogen peroxide. The thiols generated from the reduction step were tagged with biotin or fluorescently labeled maleimide reagents. Using this approach, they efficiently labeled protein S-nitrosylation in both extracts and whole fixed cells without reducing protein disulfides.

Seneviratne et al. strategically combined reductive ligation reaction with biotin switch strategy in a reaction they coined SNOTRAP (SNO trapping by triaryl phosphine) which enhanced identification of S-nitrosylated proteins by mass spectrometry (Seneviratne et al., 2016). The SNOTRAP probe is made up of a triphenylphosphine thioester linked to a biotin molecule with polyethylene glycol (PEG) spacer group serving as a linker (Figure 9H) The triphenylphosphine thioester probe reacted with SNO groups and afforded an azaylide intermediate, which through a well-positioned electrophile (thioester), rearranged to form a disulfide–iminophosphorane similar to Figure 9B. The assay is in three steps: blocking of free cysteine residues with iodoacetamide (IAA) followed by biotin-mediated affinity capture of disulfide–iminophosphorane and release of S-nitrosylated proteins and nanoflow liquid chromatography-MSMS (nLC-MS/MS) analysis (Figure 9H). They applied this approach in fully assessing protein S-nitrosylation in the neurodegenerating brain where they discovered that S-nitrosylation is increased during the early stages of neurodegeneration.

Reeves et al. examined in greater detail the conversion of S-nitrosothiols to thiosulfonates with the sodium salt of benzenesulfinic acid which was first reported by Hart in 1985 (Hart, 1985; Reeves et al., 2013; Reeves et al., 2014) (Figure 9I). While Hart carried out the reaction under very acidic conditions, the results showed that the reaction is feasible under modestly acidic conditions (pH 4.0) making it biocompatible. S-nitrosoglutathione (GSNO) and S-nitrosylated bovine serum albumin (SNO-BSA) were selectively trapped by benzenesulfinic acid sodium salt under acidic conditions (Figure 9I). Though PhSO₂Na was successfully employed as an S-nitrosothiol probe, the S-phenyl sulfonyl cysteine functional group, the resultant linkage formed at the point of conjugation underwent hydrolysis at neutral pH resulting in the diminished signal. To salvage this, Reeves et al. developed a protocol coined thiosulfonate switch that displaced S-phenylthiosulfonates with a thiol-containing fluorescent probe to generate a stable disulfide linkage. The protocol as described had three steps: blockage of free cysteines using S-phenylsulfonylcysteine (a thiol blocking reagent, that blocks free thiols at pH 4.0 in the presence of S-nitrosothiols); addition of PhSO₂Na to convert protein SNO into protein S-phenylthiosulfonates and subsequent introduction of rhodamine based fluorophore possessing a reactive thiol to facilitate a displacement reaction with the protein S-phenylthiosulfonates to afford a disulfide between the probe and the formerly S-nitrosylated cysteine residue (Figure 9I). The purified proteins BSA-SNO, and alcohol dehydrogenase regulator (AdhR*-SNO) were labeled using this technique without the generation of false positive characteristics of reductive techniques since it is not dependent on the reduction of S-nitrosothiol to a free thiol. Other methods of labeling S-nitrosylation including the famous biotin switch technique and other cysteine oxidation products are well covered in these reviews (Chung et al., 2013; Alcock et al., 2018).