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Exposure of biological molecules to oxidants is inevitable and therefore commonplace. Oxidative stress in cells arises from both external agents and endogenous processes that generate reactive species, either purposely (e.g. during pathogen killing or enzymatic reactions) or accidentally (e.g. exposure to radiation, pollutants, drugs, or chemicals). As proteins are highly abundant and react rapidly with many oxidants, they are highly susceptible to, and major targets of, oxidative damage. This can result in changes to protein structure, function, and turnover and to loss or (occasional) gain of activity. Accumulation of oxidatively-modified proteins, due to either increased generation or decreased removal, has been associated with both aging and multiple diseases. Different oxidants generate a broad, and sometimes characteristic, spectrum of post-translational modifications. The kinetics (rates) of damage formation also vary dramatically. There is a pressing need for reliable and robust methods that can detect, identify, and quantify the products formed on amino acids, peptides, and proteins, especially in complex systems. This review summarizes several advances in our understanding of this complex chemistry and highlights methods that are available to detect oxidative modifications—at the amino acid, peptide, or protein level—and their nature, quantity, and position within a peptide sequence. Although considerable progress has been made in the development and application of new techniques, it is clear that further development is required to fully assess the relative importance of protein oxidation and to determine whether an oxidation is a cause, or merely a consequence, of injurious processes.
Biological systems are exposed to a wide variety of oxidizing species—both free radicals and two-electron oxidants. These species are often termed “reactive oxygen species,” although this is a misleading term, as the reactivity of these species varies enormously (see below). Oxidants are generated both deliberately (e.g. to kill invading pathogens or as intermediates in enzymatic reactions) or unintentionally (e.g. via electron leakage from electron transport chains, metabolism of drugs, exposure to chemicals, pollutants, and radiation). These processes have been reviewed in Refs.
The formation of these oxidants and their reactions are limited by cellular and organismal defense systems, which include enzymes that remove oxidants or oxidant precursors (e.g. superoxide dismutases, peroxiredoxins, thioredoxin/thioredoxin reductase, GSH peroxidase isoforms, and catalases), and water- and lipid-soluble oxidant scavengers, including thiols (e.g. GSH and thioredoxin), ascorbic acid (vitamin C), urate, tocopherol isoforms (vitamin E), quinols (e.g. reduced coenzyme Q10), carotenoids, and polyphenols. Although these systems are efficient, and in many cases show redundancy (i.e. multiple processes remove the same species), they are not 100% effective, and a large body of data indicates that biological targets suffer resulting damage. These protective systems are therefore complemented by systems that either repair damage or remove the modified molecules (e.g. methionine sulfoxide reductases, disulfide reductases/isomerases, glutaredoxins, sulfiredoxins, proteasomes, lysosomes, proteases, phospholipases, and DNA repair enzymes) (
Despite this battery of preventative and repair systems, many studies have reported increased damage, and accumulation of this, in human, animal, and microbial and plant systems exposed to stress conditions (reviewed in Refs.
). A higher level of damage may arise from increased oxidant generation, a decrease or failure of defense systems, or (most commonly) a mixture of both processes, as many defense systems are themselves subject to damage or show reduced activity due to co-factor depletion. This concept of an altered balance between formation and removal gave rise to the term “oxidative stress” (
), although it is now apparent that this is an oversimplification of a complex picture, as limited stress (“eustress”) can be beneficial in priming and protecting a system against greater damage (“distress”) (
). Increasing age is often associated with a decrease in enzyme levels or activity, and in some cases decreased levels of co-factors and essential trace elements, such that increased levels of oxidants and modified products are formed (
). These changes can be accelerated by disease or environmental factors, despite the presence of feedback loops (e.g. antioxidant-response elements, including the Nrf-2 pathway; DNA damage–response element; OxyR; SoxRS) that up-regulate the synthesis of protective species (
). In this study, we review the basic chemistry and biochemistry of protein modification by oxidants, with a focus on methods available for the detection, identification, and quantification of these changes.
Proteins as targets of oxidative damage
Proteins are major components of most biological systems and constitute ~70% of the dry mass of cells and tissues. The rate of reaction of an oxidant with a biological target depends on the concentration of the target, multiplied by the rate constant for its reaction with the oxidant. Both of these factors result in proteins being major targets for damage as proteins are both present at high concentrations (up to 1–3 mm in plasma and 5–10 mm in cells) and have high rate constants for reaction with oxidants. Thus, oxidant damage in most biological systems is likely to be skewed toward protein modification (
). This is clearly an oversimplification of a complex situation, as other factors are known to play an important role, including localization of the generating system relative to the target and particularly membrane barriers, micro-environments, binding, or association of the oxidant system to a target, the occurrence of secondary reactions, and intra- and intermolecular transfer reactions (
). Thus, low levels of modification of a critical target may have much greater consequences than a high level of damage to noncritical materials.
Radicals (e.g. HO•, CO3-., NO2·, ROO•, RO•, R•, and many others), two-electron oxidants (e.g. peroxides, 1O2, O3, ONOOH, HOCl, and related species), and metal–oxo complexes can all modify proteins, although the reactivity and selectivity of these oxidants are highly variable (
). Together, these generate a wide variety of post-translational modifications that alter amino acid and protein composition and structure, charge, hydrophobicity/hydrophilicity, folding, and function (
In the following section, a brief overview is presented on the formation and reactivity of a number of key oxidant species relevant to mammalian systems. Although each of these species is described as single entities, it should be noted that nearly all of these species undergo further interconversion reactions as illustrated in Fig. 1 (although this is situation-dependent) that result in complex mixtures in most reaction systems (
Superoxide O2-. radicals are generated continuously by most organisms as a result of the use of O2 as a terminal electron acceptor in some electron transport chains, such as those of mitochondria, the endoplasmic reticulum cytochrome P450 system, and the plasma membrane NADH/NADPH oxidase systems (
). O2-. undergoes rapid spontaneous, and superoxide dismutase-catalyzed, disproportionation to give hydrogen peroxide (H2O2) and O2, with the former being a precursor of further oxidizing species, as well as being a reactive species in its own right (
In addition to formation from O2-. by dismutation, H2O2 is also generated directly by a number of enzymes (e.g. NADPH oxidase-4, monoamine oxidases, hexose oxidases, amino acid oxidases, other oxidoreductases, protein-disulfide isomerases, and Ero1p during protein synthesis and folding (
)). Direct oxidation of biological targets by H2O2 is both limited in extent and is usually slow. Thus, direct reaction is limited to Cys, selenocysteine, and Met residues with these typically occurring with very low rate constants (see also below), with the exception of reaction with some specialized enzymes (e.g. peroxiredoxins and GSH peroxidases), which have catalytic centers that facilitate rapid O–O bond cleavage. Such environments can elevate the rate constant for reaction by a million-fold (
). H2O2 is a substrate for a large family of peroxidase enzymes, with these reactions used both synthetically (e.g. in the formation of thyroid hormones by thyroid peroxidase and in the generation of collagen matrices by peroxidasins) and to kill invading pathogens (
). HOCl (which exists in equilibrium with Cl2 at low pH values) is a key component of the innate immune response against pathogens, with this generated at high concentrations in neutrophil phagolysosomes (
). Release of myeloperoxidase to the extracellular space (instead of into phagolysosomes) and subsequent reaction with H2O2 can, however, result in host tissue damage, with the level and activity of myeloperoxidase associated with tissue damage in acute and chronic inflammatory conditions (
). Comparison of HOCl and H2O2 illustrates a key point about oxidants and their reactivity: H2O2 is a strong oxidant (reduction potential 1.32 V) and a more powerful oxidant than HOCl (reduction potential 1.28 V), but this reacts very slowly when compared with HOCl (cf. the rate constant, k, for reaction of HOCl with the amino acid methionine is ~108-fold higher than for H2O2 (
)), so evolution has favored the use of HOCl over H2O2, for kinetic rather than thermodynamic reasons. The low reactivity of H2O2 is likely to be one of the major reasons why this species is used biologically as a messenger molecule: slow and selective reactivity allows for specificity in transmission of messages as only a limited number of select targets are likely to be activated. Highly-reactive species such as HOCl would not allow such specificity of transfer of information.
Nitric oxide (NO•)
NO• is a key vascular regulator and messenger molecule, with this generated from arginine by nitric-oxide synthase enzymes (NOSs)
). The concentration generated by constitutive NOS isoforms is low (pico- to nanomolar), consistent with a regulatory function, but higher levels (up to micromolar) are formed by the inducible NOS isoform of macrophages, at sites of inflammation (
), with fast reactions occurring primarily with transition metal ions (e.g. the iron center of heme proteins, including that of soluble guanylate cyclase, the major effector of NO• signaling) and with other radicals (
). Reaction with other radicals can be protective (e.g. by terminating lipid peroxidation chain reactions) but also damaging when the product is a powerful oxidant, as is seen in the formation of peroxynitrous acid (
Diffusion-controlled (i.e. k 109–1010m−1 s−1) reaction of NO• with O2-. gives ONOO−(peroxynitrite). This species exists in equilibrium with the corresponding conjugate acid peroxynitrous acid (ONOOH), with the pKa (6.8) favoring the anion at most biological pH values (
). However, the acid (ONOOH) form is typically the more reactive species with protein targets. Reaction with CO2 and some boronic acid probes are exceptions, with the rate constant for these targets being higher with ONOO− (
). The reactivity of ONOOH is therefore modulated by CO2 (which is in equilibrium with HCO3−) as a result of the formation of ONOOCO2−, which decomposes to give CO3-. and NO2· that can either diffuse out of the solvent cage or recombine (reviewed in Ref.
) that can give rise to both two-electron and one-electron oxidation products. The former arises from direct reactions of ONOOH, and the latter is formed from limited homolysis to give HO• and NO2·, and subsequent radical chemistry (
). Metal ion–catalyzed decomposition of organic and lipid hydroperoxides gives alkoxyl radicals (RO•) that are less powerful oxidants than HO•, but more reactive than peroxyl radicals (ROO•) that are typically generated from rapid (typically diffusion-controlled) addition of O2 to carbon-centered radicals (R•) (
). R• and ROO• are key intermediates in lipid peroxidation chain reactions, i.e. typically initiated by hydrogen abstraction from methylene groups of polyunsaturated fatty acids. ROO• are the key chain carriers in lipid peroxidation (
UV light, singlet oxygen (1O2), and other photochemically-generated species
UV light with very short wavelengths (UVC) is strongly absorbed by atmospheric molecules, such as ozone, and hence does not give rise to significant effects at the earth’s surface. In contrast, longer wavelength UV light, particularly UVB (λ 280–320 nm) and UVA (λ 320–400 nm) wavelengths, can oxidize molecules via light absorption by suitable chromophores and generation of either excited state species (i.e. species with an electron in a higher orbital, type 2 photoreactions) or radicals (type 1 photoreactions), as a result of photoejection of an electron (
). The excited state species (usually triplet species due to the short lifetime of singlet states) can either induce direct oxidation by, for example, electron or hydrogen atom abstraction or by energy transfer to molecular oxygen O2 to give singlet oxygen (1O2) (
). In addition to its formation by type 2 photoreactions, 1O2 is also formed by some enzyme-catalyzed reactions, via termination reactions of ROO•, and some metal ion-catalyzed processes; 1O2 is therefore an important intermediate in both light-induced and “dark” reactions (
), and these modifications have been linked to a wide range of pathologies, including some cancers, neurological disorders (Alzheimer’s and Parkinson’s diseases and amyotrophic lateral sclerosis), sepsis, hypertension, cardiovascular diseases, including atherosclerosis, ischemia-reperfusion injury to multiple organs, renal and ocular damage, cataracts, chronic obstructive pulmonary disease, cystic fibrosis, asthma, rheumatoid and osteoarthritis, motor neuron disease, irritable bowel syndrome, pancreatitis, hepatitis, sunburn and photodamage, and many more (
). Although it is likely that some of these are only associations (i.e. oxidation is not causative but merely a consequence of other injurious processes), oxidative damage appears, at least in some cases, to be a contributing factor (i.e. causative). Methodologies that can detect and quantify oxidative damage are therefore of considerable importance, not least as potential biomarkers to assess therapeutic strategies. Oxidation can also be a valuable tool in the treatment of some diseases, with radiation and photodynamic therapy of tumors (
). The following section outlines current knowledge of the modifications arising from oxidant interactions with proteins, and subsequently how these can be detected and quantified.
Protein modifications induced by reactive oxidants
The majority of radical reactions with proteins occur via three major pathways— hydrogen atom abstraction from C–H, S–H, N–H, or O–H bonds, electron abstraction from electron-rich sites, and addition to electron-rich centers (aromatic rings and sulfur species) (
). The first of these reactions results in the formation of carbon-centered species (R•), thiyl radical (RS•), nitrogen-centered species (primarily indolyl radicals from Trp), and oxygen-centered radicals (primarily phenoxyl radicals from Tyr). Most R• radicals, including those generated from oxidation of aliphatic side chains (Leu, Ile, Val, and Pro, etc.) react rapidly with O2 to give ROO• at diffusion-controlled rates (k ~109m−1 s−1) (
). Although these reactions are fast, the O2 concentration is low in most biological samples (5–100 μm), which may limit ROO• formation and result in dimers (R–R) when the R• concentration is relatively high (Fig. 2) (
). The chemical structures of some of the most abundant and commonly examined products are given in Fig. 3.
For ROO• generated on aliphatic side chains, hydrogen-atom abstraction is a major reaction with other X–H bonds, with this resulting in hydroperoxide (ROOH) formation (Fig. 2). These are major products with multiple different attacking species (
). Alternatively, RO• may undergo hydrogen atom abstraction radicals to generate alcohols (Fig. 2). With primary or secondary ROO•, dimerization reactions yield 1 mol of alcohol, carbonyl, and O2 (Fig. 2) (
) resulting in damage to a subset of amino acid side chains; these data are summarized in Table 1. These include the sulfur-containing amino acids Cys, Met, and cystine, and the aromatic residues, Trp, Tyr, Phe, and His. For both Cys and His, the reactions are pH-dependent, with the rate of oxidation occurring more rapidly at higher pH values (
). With species such as HO•, this can result in a large range of different cleavage sites along a protein backbone (often detected as a “smear” on protein gels), although some selectivity in fragmentation has been reported (
), including reaction with Cys, cystine, and Met residues to give oxygenated species (Table 1). Oxidation of metal-ion centers can also occur via two-electron pathways. However, these reactions occur in competition with homolysis of ONOOH to give radicals (and hence one-electron oxidation products), and reaction of the anion ONOO− with CO2 to give the corresponding carbonate adduct (although the identity of this species is disputed (
)) and to decompose to give NO2· and CO3-., and thereby generates radical-mediated products. As NO2· is formed from both ONOOH and ONOOCO2−, nitrated products are commonly detected, with these being mainly generated from Tyr and Trp residues via dimerization reactions of NO2· with the TyrO• and TrpN• species formed by HO• or CO3-. (
). With Trp, Tyr, and His, the initial products are endoperoxides formed by cycloaddition reactions, with these subsequently undergoing ring opening to give hydroperoxides, oxygenated products, and further cyclized materials (Table 2) (
). As with the radical chemistry of Trp and His, these reactions can result in ring opening reactions and a similar (complex) mixture of species. The products from these amino acids therefore do not allow the initial oxidant to be easily identified.
From the above discussion it is clear that different oxidants have very different chemical behaviors and reaction kinetics, and these differences can be magnified or decreased by a range of other factors that influence oxidant selectivity. This is discussed further in the following section.
Factors affecting oxidant selectivity
The extent of damage by a particular oxidant can be modulated by multiple factors, including the accessibility of the oxidant to the target residue (e.g. Trp residues are often buried within protein structures and have limited solvent accessibility), and also electrostatic interactions with residues on the protein surface (e.g. the presence of charge on oxidants such as O2-. and CO3-.). Neutral species may induce greater damage than charged species, and also at more remote locations, due to the greater propensity of such species to traverse membranes and hence diffuse away from their site of generation. The neutral species may also be better electrophiles and provide better leaving groups. This is exemplified by the greater reactivity of HOCl over −OCl and ONOOH over ONOO− (
). Ionization of side-chain residues on a protein increases their electron density, increasing their capacity to act as a nucleophile and be more readily oxidized; thus, the thiolate anion, RS−, is more reactive than the parent RSH (and similarly for the related selenium-containing amino acid, selenocysteine, Sec), and the Tyr phenolate anion is more readily oxidized than the neutral phenol (
). Such transfers can occur over long distances (e.g. in ribonucleotide reductase, DNA photolyase, and photosystem II), such that initial oxidation at one site can be rapidly transferred to another remote residue, with the electron transfer occurring through bonds or space (Fig. 5) (
). Consequently, the initial site of oxidation may not be the final site of modification. As the one-electron reduction potentials of Trp and Tyr are similar, radical formation at one residue can result in equilibration between residues, assuming suitable electron transfer pathways are available (
). One consequence of this is that radical termination reactions (e.g. dimerization of two TyrO•, two TrpN•, or cross-reaction of these species with NO2· to give nitrated products) may occur via the most accessible, or reactive, TyrO• or TrpN• rather than at the site of initial radical generation. Thus, cross-link formation involving Tyr and Trp radicals, and formation of products such as 3-nitro-Tyr, may be determined by the accessibility and reactivity of a particular residue, rather than the extent of initial reaction at that site (Fig. 5).
In the light of these data, the next section summarizes commonly used methods to detect protein alterations, starting with modifications at the intact protein level (i.e. changes that markedly affect protein mass and structure: “gross changes”) and then progressing to techniques that identify and quantify changes at an amino acid level, and at specific sites within a protein sequence.
Detection and quantification of protein oxidation
Gross modification of parent proteins
Oxidation of proteins can generate both fragmentation and aggregation of proteins. The latter can involve both covalent cross-linking as well as noncovalent interactions. Separation methods based on mass or charge (e.g. one- or two-dimensional electrophoresis and column chromatography) with subsequent detection methods (e.g. silver staining or immunoblotting) can provide limited information about such changes. This works best with purified proteins or limited mixtures, but it has severe limitations with complex samples and also when comparing healthy versus diseased samples, as the protein pools may be very different in such cases, even when two-dimensional gels are used to enhance resolution. Immunoblotting with specific antibodies can provide high sensitivity and specificity detection, but this approach is severely limited with regard to both the quantification and identification of modifications. Artifactual proteolysis or aggregation is also a serious concern. Both oxidant-mediated fragmentation and aggregation can be investigated using these approaches, but as fragmentation is often nonspecific or poorly-specific, discrete bands or spots (from 2D gels) are rare, with “band smears” being the usual outcome.
Aggregation or cross-linking is more readily analyzed, as dimers (for example) generated by any pairing of residues are likely to provide bands/spots of similar mass. Care clearly needs to be taken as multiple proteins are typically present in each band or fraction. Reduced antibody recognition of a specific native epitope can be used as a method of assessing modification to that site, in either immunoblotting studies or ELISA. These approaches are limited by the availability of specific antibodies but have been used successfully in a large number of studies ranging from isolated proteins to tissues, and they have the advantage of very-high sensitivity. Increased information can be obtained if antibodies against both parent protein epitopes, and specific products (see below), are available (
). An example of this approach are the studies that have examined HOCl-mediated damage to the extracellular matrix underlying endothelial cells. Binding of three specific antibodies (anti-fibronectin, anti-laminin, and anti-thrombospondin) was decreased on treatment with HOCl, implicating damage to these proteins (
). However, analysis of such data can be complex, as damage may also enhance antibody binding by exposing cryptic epitopes. Thus, low doses of HOCl appear to increase the affinity of anti-fibronectin antibodies to plasma fibronectin, whereas high concentrations have the opposite effect (
). This has been rationalized in terms of the generation of an extended fibronectin conformation at low HOCl doses, and aggregation with high concentrations.
An absence of positive data from ELISA or immunoblotting studies does not preclude the presence of damage, as epitopes may become inaccessible on protein oligomerization or as a result of other structural changes. Quantification is also challenging as this is very dependent on the sensitivity of the antibody: strong signals may be detected for low-abundance material, whereas abundant species may give a weak (or no) signal with a poor antibody. Separation of modified species by HPLC, for example, has been employed successfully with oxidants that are highly selective and that induce damage at a limited set of residues. An example is the separation of modified apolipoproteins AI and AII (
). Subsequent MS analysis of the fractions identified the modifications as loss of parent Met and generation of the corresponding sulfoxide.
Other biophysical techniques (e.g. CD, light scattering, small-angle neutron scattering, small-angle X-ray scattering, turbidity methods, X-ray crystallography, and NMR spectroscopy) can also yield information on protein structure, particularly the generation of fragments or aggregates, as these methods are sensitive to changes in protein mass, the size of particles, modified secondary structure, and altered charge and solubility. X-ray crystallographic studies have provided evidence for increased electron density between residues in aggregated proteins supporting the presence and identification of particular cross-links and their nature (exact site of linkage and intra- versus inter-molecular; see, for example, data for oxidized peroxiredoxin 5, thioredoxin 2, and γS-crystallin (
)). However, with the exception of X-ray crystallography and NMR, these methods do not provide definitive information as to the sites and modifications, and these techniques are (currently) limited to homogeneous samples (often single proteins) with high modification levels.
Total amino acid analysis
This methodology can provide important quantitative data on the consumption of parent species, arising from all potential modification reactions, and for some species the yields of products can also be assessed (e.g. methionine sulfoxide, see below). Such data are important with regard to obtaining a material balance, something that has been difficult to achieve even with the simplest systems. An overview of this approach is provided in Fig. 6. Differences between loss and total product formation can provide vital information with regard to the generation of alternative (known or unknown) species (see also below). Typically, proteins are isolated (e.g. by precipitation from homogenates/lysates using TCA or organic solvents), cleaned up (e.g. delipidation), and subsequently subjected to hydrolysis to give the free amino acids and products (Fig. 6) (
). Proteolysis can be achieved using acid conditions (with this resulting in loss of Cys, cystine, and some Trp species, although this depends on the acid), alkaline conditions (which preserves Trp species, but results in loss of other species such as the product DOPA), or nonspecific proteases, such as Pronase (
). The free amino acids (and any products) are then separated (e.g. by HPLC/UPLC) and quantified by mass spectrometry (MS), fluorescence (either directly, for some aromatic species, or by pre-column fluorescent tagging of free amino groups using reagents such as o-phthaldialdehyde, OPA), UV absorption, or electrochemical methods (Fig. 6) (
). The combination of acid hydrolysis (using methane sulfonic acid) and OPA tagging allows data to be obtained for all common amino acids with the exception of Cys/cystine (which are acid-sensitive), Asn and Gln (which are converted to Asp and Glu, respectively), and Pro (which does not react with OPA, being an imine) (
). Enzymatic hydrolysis results in lower levels of artifactual oxidation due to the mild conditions (typically overnight incubation at 37 °C, pH 7.4) and hence preservation of acid/base-sensitive materials, but quantification can be problematic due to self-digestion of the protease resulting in release of additional amino acids although this is often limited (
). Lys quantification can be problematic due to the second side-chain amino group, which results in multiple peaks if labeling (such as tagging with OPA or other amine-reactive tags) is incomplete. An internal standard (e.g. homo-Arg) allows sample recovery and derivatization efficiency to be assessed (
). The use of sacrificial oxidation targets (e.g. tryptamine), anoxic conditions, antioxidants, and other inhibitors are important to prevent significant losses, and data are typically normalized to nonmodified amino acids to compensate for any losses during processing (
). Comparison with expected amino acid compositions is recommended for studies using pure proteins, to ensure that materials are not lost (or the extent compensated for) during processing. Some modifications (e.g. Trp products during acid hydrolysis) are also known to be lost during processing (see below), which may result in an underestimation of the level of damage. MS analysis can also be readily used for studies on free amino acids, peptides, and for proteins. For the last of these, analysis is usually undertaken on loss of specific peptides after digestion using trypsin (or other enzymes) or at the intact protein level. Absolute quantification can, however, be tricky to achieve (see below).
The total amino acid analysis approach is limited in that it only provides overall levels and not data on the sites of modifications within a sequence, nor the data on what proteins they might be present on, when analyzing complex mixtures. However, these data are an important complement to other approaches, such as MS peptide mass mapping, where only a limited number of species are typically analyzed.
Loss of some amino acids can be quantified by alternative methods. Direct fluorescence (typically λex 280–285 nm and λem 340–345 nm) has been widely used to examine Trp residues (
)) and is difficult to use in the presence of other species that absorb or exhibit fluorescence at these wavelengths. This may include proteins with high concentrations of Tyr residues, heme proteins, species arising from Trp degradation, and some glycoxidation products. Lys and Arg can be quantified by reagents that give strongly fluorescent derivatives, such as fluorescamine (
) respectively. These methods are rapid, sensitive, and give limited artifacts during sample preparation, but neither reagent is entirely specific, and hence needs to be used with care.
A number of methods have been developed to allow quantification of the loss of the key redox-sensitive amino acid Cys on proteins. Detection and quantification of Cys products are covered below. Methods include spectrophotometric (e.g. using 5,5′-dithiobis(2-nitrobenzoic acid) (DTNB), Ellman’s reagent (
)), fluorometric (e.g. using ThioGlo 1, (10-(2,5-dihydro-2,5-dioxo-1H-pyrrol-1-yl)-9-methoxy-3-oxo-3H-naphthol[2,1-b]pyran-2-carboxylic acid methyl ester), a naphthopyranone maleimide derivative, and related species (
), this reaction can be exploited to determine levels of GSH and the total of GSSG and GSH spectrophotometrically. GSSG levels can then be determined by difference in the values. The GSH concentration can be determined as described above, and also via the consumption of NADPH, the co-factor for GSH reductase (
). GSSG levels can also be determined using genetically-encoded probes, such as roGFP fused to glutaredoxin; these probes are not responsive to thioredoxin-mediated changes probably for steric reasons. These probes allow real-time imaging of redox changes at specific and defined locations within living cells, although they also have some significant caveats; in particular, these are not direct oxidant probes, for they only report on the redox status of the specific environment being examined (
). Kits are available for such measurements. Direct measurements of both GSH and GSSG can be achieved after separation (e.g. by HPLC/UPLC) with various detection methods, including electrochemistry (e.g. Ref.
). The absolute amounts, and ratio, of protein-bound Cys and low-molecular-mass thiols can be individually assessed after separation of the high- and low-molecular-weight fractions (e.g. by use of spin filters or protein precipitation).
Reduced and oxidized thiols on proteins separated on 1D or 2D gels can be assessed by a number of methods (reviewed in Ref.
). Reversible thiol modifications can be examined by use of derivatization reagents after oxidant treatment, reduction of the reversible modification, and use of a second orthogonal tag (also see below and Ref.
Direct quantification of disulfides (e.g. cystine) is complex but can be achieved by mild protein digestion methods (e.g. using chemicals such as cyanogen bromide or enzymatic approaches), which cleave the polypeptide backbone between the half-cystinyl residues, under conditions that minimize thiol-disulfide exchange and disulfide reduction. Diagonal electrophoresis was used in early studies (reviewed in Ref.
Modification of proteins and peptides by oxidants can result in the generation of a number of intermediate species that have modest lifetimes and stabilities. As the assay of these materials requires specialized methods, these are discussed separately from the analysis of long-lived (“stable”) products, which are discussed later in this review. An overview of these methods is provided in Fig. 7.
Radicals, both initiating species and those formed on peptides and proteins, can be detected by a number of approaches including direct spectroscopic methods such as UV-visible, resonance Raman, conductivity, and electron paramagnetic resonance (EPR). Because of the short half-lives of most radicals, techniques with rapid response times are required, and of the above methods, only EPR is specific for radicals (
). Measurements using other techniques can be readily confounded by more abundant nonradical species, and hence they are only typically used with very clean systems coupled to rapid radical generation methods (e.g. pulse radiolysis, flash photolysis, and stopped flow) (
). Although this is a very powerful analytical technique for identification of radicals (the “gold standard”), quantification is very challenging due to the short lifetime of these species. This can be (partially) overcome by use of ancillary techniques such as rapid-flow methods, in situ photolysis or radiolysis, freeze-quenching, and spin trapping (e.g. using nitrone compounds, such as 5,5-dimethyl-1-pyrroline N-oxide, DMPO), although each of these has advantages and disadvantages (reviewed in Refs.
). In this technique, a compound (the spin trap, typically a nitroso or nitrone) is added, with the aim of generating more stable, detectable adducts (Fig. 7). Analysis of the resulting spectra can then yield information on the species present (
). The technique is artifact-prone and needs to be carried out with care. The resulting data can be definitive with regard to the species present, but also have a number of inherent caveats. In particular, the sensitivity of the method allows minor pathways to be detected and potentially misinterpreted, and the data do not provide information as to the absolute radical concentrations. Differences in the rates of trapping (or adduct decay) may make a minor species appear important when compared with a major pathway that yields very transient species. The long lifetimes of some protein–radical adducts have allowed this approach to be combined with other analytical methods, including MS, to provide detailed information (reviewed in Ref.
), with this method utilizing the decay of a nitroxide spin adduct to a nitrone species to generate an antigen that is recognized by an antibody. This then allows the sensitive and specific detection of the former radical species, on isolated proteins, in cells, and in animals (primarily rodents). Immunospin trapping system can also be combined with LC/MS/MS and with the anti-DMPO antibody used to screen separate fractions (e.g. from HPLC or size-exclusion columns) for the presence of adducted DMPO. This allows residues that previously contained a radical to be readily detected, as the mass addition arising from the presence of DMPO can be readily detected (
Hydroperoxides are major products of both radical- and 1O2-mediated damage to proteins in the presence of O2. These species are formed on many side chains in high yield by a wide range of insults, including oxygen-derived radicals, 1O2, activated white cells, ONOOH, and metal ion-catalyzed systems (Table 2) (reviewed in Ref.
). Decomposition of these species, which have lifetimes of minutes to many hours, by metal ions or UV light can give further radicals (RO•, R•, and ROO•) that propagate damage, including to lipids, other proteins, DNA, and RNA (
) gives the corresponding (stable) alcohols as products. In contrast, reaction of ROOH/H2O2 with critical Cys residues present on proteins or enzymes can inhibit enzyme activity and exacerbate damage (
)), consistent with the occurrence of this chemistry in vivo. Hydroperoxides can be quantified by multiple methods, including iodometric titration, the ferrous oxidation–xylenol orange (FOX) assay, and also by use of boronic acid probes (Fig. 7) (
). In the first of these, reaction of ROOH with iodide ions (I−) in the presence of acid generates triiodide (I3−) that can be quantified by its absorbance at 358 nm. This method is quantitative and has a well-defined 1:1 stoichiometry, but it needs be performed under strict anoxic conditions due to the sensitivity of acidified iodide solutions to O2; it is therefore technically demanding (
). The FOX (ferrous oxidation–xylenol orange) method assays hydroperoxide-mediated oxidation of a Fe(II)–xylenol orange complex to the Fe(III) form, with the latter quantified via its absorbance at 560 nm (
). This assay is generic for all hydroperoxides, and also H2O2 (so samples are typically pre-treated with catalase to remove this species), and has been adapted to allow quantification of both protein- and lipid-derived hydroperoxides (
). This method has a low sensitivity to O2, but it is not compatible with some buffers and has a poorly-defined stoichiometry; consequently, data are typically reported as H2O2 equivalents obtained by use of a standard curve generated using this species (
)), but these overlap with many other species in complex systems, so quantification is usually achieved by reaction with an added probe such as 5-thio-2-nitrobenzoic acid (TNB, which is oxidized to the corresponding dimer, DTNB (
)), with quantification achieved via the loss of absorbance from TNB at 412 nm. Oxidation of TNB to DTNB also occurs with many oxidants (e.g. HOCl, HOBr, HOSCN, H2O2, and other peroxides, ONOOH, 1O2, and many radicals), and hence it is not specific for any particular species. Iodometric titration can also be employed (see section on “Hydroperoxides”), but again this lacks specificity. Iodide ions have also been used to catalyze the oxidation of 3,3′,5,5′-tetramethylbenzidine and dihydrorhodamine, by chloramines, to optically absorbing or fluorescent products (Fig. 7) (
Sulfenic acids (RSOH, with the formation of these species often termed S-sulfenylation), sulfenyl chlorides (RSCl), and S-nitrosated species (RSNO, S-nitrosylated) are major intermediates formed by two-electron oxidants with Cys residues and related species (
). S-Nitrosylated and S-nitrated (RSNO2) species are also formed by one-electron mechanisms involving reaction of RS• with NO• and NO2·, respectively. RSOH and the corresponding sulfenyl amides RS-NHC(O)R′ (
), particularly as these are formed in a reversible manner and hence may provide protection against irreversible oxidation of critical Cys residues and provide a facile “on-off” switch for enzyme activity and act as redox switches (
). Most of these species (with the exception of RSNO) react rapidly with other thiols (and other nucleophiles and oxidants) to give disulfides, thiosulfinates, and sulfinic (RSO2H) or sulfonic acids (RSO3H) (
). Most methods for the detection of RSOH rely on chemical derivatization or trapping methods, with the prototypic species being 5,5-dimethyl-1,3-cyclohexanedione (dimedone), which reacts with RSOH (and other species (
). Reduction of sulfenic acids by arsenite has been utilized to develop a “biotin-switch” method for labeling protein RSOH, with the reduced amino acid subsequently labeled with biotin-maleimide. The adducts can then be detected using immunoblotting with streptavidin-horseradish peroxidase or separated using streptavidin-agarose (
). In its classical form, nonmodified thiols are first blocked, and following the removal of excess alkylating reagent, ascorbate is added to selectively reduce any S-nitrosothiols (but not other species) to free thiols that are then labeled and detected by immunoblotting or fluorescent tagging following separation. Multiple iterations and improvements of this method have been proposed to enhance its specificity and to allow more rigorous quantification of RSNO levels. Many drawbacks have been reported, and considerable care and appropriate controls need to be employed to minimize artifacts (
). As with other unstable intermediates, considerable care needs to be taken to avoid artifactual changes in the levels and sites of modification, as it is well-established that some RSNO species undergo ready transnitrosation reactions (
In biological systems, nitrosation is readily reversed, and this appears to be primarily driven by enzymatic reactions with reduction of low-molecular-mass species, such as S-nitrosated GSH (GSNO), being catalyzed by the widespread enzyme S-nitrosoglutathione reductase (GSNOR) (
). These species are formed, at least in part, via downstream reactions of H2S, a relatively recent addition to the family of a gaseous signal transmitter family, the other members being NO• and CO. Conversion of Cys to Cys-SSH occurs through sulfuration or persulfidation processes and may involve oxidized H2S species and particularly polysulfides (H2Sn), as well as other pathways. H2S is generated from Cys and homocysteine by widely-expressed enzymes, including cystathionine β-synthase, cystathionine γ-lyase, cysteine aminotransferase, and 3-mercaptopyruvate sulfurtransferase. As these species are expressed in the vascular wall, H2S has been proposed as a regulator of vascular tone, neuronal health, the integrity of endothelial cells barriers, smooth muscle cell proliferation and survival, angiogenesis, and as a modulator of inflammation (