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Laboratorio de Fisicoquímica Biológica, Facultad de Ciencias, Universidad de la República, 11400 Montevideo, UruguayCentro de Investigaciones Biomédicas (CEINBIO), Universidad de la República, Montevideo, Uruguay
Laboratorio de Fisicoquímica Biológica, Facultad de Ciencias, Universidad de la República, 11400 Montevideo, UruguayCentro de Investigaciones Biomédicas (CEINBIO), Universidad de la República, Montevideo, Uruguay
To whom correspondence should be addressed: Laboratorio de Enzimología, Facultad de Ciencias, Universidad de la República, Iguá 4225, 11400 Montevideo, Uruguay. Tel./Fax: 598-25250749; E-mail: [email protected]
Centro de Investigaciones Biomédicas (CEINBIO), Universidad de la República, Montevideo, UruguayLaboratorio de Enzimología, Facultad de Ciencias, Universidad de la República, 11400 Montevideo, Uruguay
The free radical nitric oxide (NO•) exerts biological effects through the direct and reversible interaction with specific targets (e.g. soluble guanylate cyclase) or through the generation of secondary species, many of which can oxidize, nitrosate or nitrate biomolecules. The NO•-derived reactive species are typically short-lived, and their preferential fates depend on kinetic and compartmentalization aspects. Their detection and quantification are technically challenging. In general, the strategies employed are based either on the detection of relatively stable end products or on the use of synthetic probes, and they are not always selective for a particular species. In this study, we describe the biologically relevant characteristics of the reactive species formed downstream from NO•, and we discuss the approaches currently available for the analysis of NO•, nitrogen dioxide (NO2•), dinitrogen trioxide (N2O3), nitroxyl (HNO), and peroxynitrite (ONOO−/ONOOH), as well as peroxynitrite-derived hydroxyl (HO•) and carbonate anion (CO3•−) radicals. We also discuss the biological origins of and analytical tools for detecting nitrite (NO2−), nitrate (NO3−), nitrosyl–metal complexes, S-nitrosothiols, and 3-nitrotyrosine. Moreover, we highlight state–of–the–art methods, alert readers to caveats of widely used techniques, and encourage retirement of approaches that have been supplanted by more reliable and selective tools for detecting and measuring NO•-derived oxidants. We emphasize that the use of appropriate analytical methods needs to be strongly grounded in a chemical and biochemical understanding of the species and mechanistic pathways involved.
Soon after the discovery of nitric oxide (NO•) as a physiological mediator in the vascular, nervous, and immune systems, it became evident that this moderately-reactive free radical can give rise to secondary species, many of which are oxidizing, nitrosating, or nitrating agents toward biomolecules (
The species formed downstream from NO• (i.e. NO•-derived oxidants) include nitrogen dioxide (NO2•), dinitrogen trioxide (N2O3), nitroxyl (HNO), and peroxynitrite (ONOO−/ONOOH), as well as hydroxyl (HO•) and carbonate anion (CO3•−) radicals formed from peroxynitrite (Fig. 1). These species are short-lived (half-lives are typically in the millisecond to microsecond range) and are frequently referred to in general as “reactive nitrogen species.” However, this term should be used with caution because in a similar way as “reactive oxygen species,” with which NO•-derived species are usually grouped, this term may give the inaccurate idea that there exists only one ill-defined species that has a particular type of reactivity and targets all biomolecules (
). In contrast, the different species have unique reactivities and, depending on the particular properties of each, they may lead to oxidation, nitrosation, or nitration. As a further argument against the use of the term “reactive nitrogen species,” some of the species formed downstream from NO• (e.g. CO3•−) do not contain nitrogen. Finally, researchers in the nitrogen fixation field might argue that the reactive nitrogen species are those formed in the activation of nitrogen in the nitrogenase-catalyzed process of ammonia formation. Thus, in line with proposals in the free radical research field (
), we suggest that the name of the identified species should be used whenever possible. When the species that are being referred to are unknown, we suggest using the term “NO•-derived oxidants.”
The preferential targets of NO•-derived oxidants in biological systems are typically located in close proximity (in the micrometer distance range) and determined by a combination of factors, including kinetic aspects of rate constants multiplied by target concentration, compartmentalization, and membrane permeability. Some of the NO•-derived oxidants are good one-electron oxidants that start oxygen-dependent chain reactions in both aqueous and lipidic compartments, which may amplify the effects (
In many cases, the formation of NO•-derived oxidants is linked to the presence of partially-reduced oxygen species, as exemplified by peroxynitrite, which is formed from the reaction of NO• with the superoxide radical (O2•−). Thus, the formation of NO•-derived oxidants is frequently related to inflammation, in which increased formation of NO• through the inducible nitric-oxide synthase converges with increased formation of O2•− and other oxidants. In fact, the high reactivity of some of the species derived from NO• make them part of the weaponry that immune cells use in their battles against microorganisms (
) have expanded the traditional view of oxidative stress as a misbalance between oxidant formation and antioxidant action to include the view of a disruption in regulatory pathways.
Because of their high reactivity, the species derived from NO• have relatively short half-lives that impede their detection in biological systems through direct spectroscopic techniques. Thus, the analytical strategies used to demonstrate the formation of a certain species in a particular biological context are based either on the measurement of downstream stable products or on the use of probes that react with the species. Because these strategies are not always specific for a certain species, the modulation of the formation or decay pathways of precursors and products provides complementary evidence. For example, the modulation of NO• and O2•− formation, which are the precursors of peroxynitrite, should accompany the results obtained through the detection of the stable product 3-nitrotyrosine or through the use of peroxynitrite probes. The modulation of NO• formation can be carried out using nitric oxide synthase inhibitors, among other strategies.
In the following sections, we briefly describe the reactive species derived from NO• in a biological milieu and NO• itself, as well as some of the stable end products (Fig. 1). We examine methodologies used for their detection and quantification, focusing on strategies aimed at assessing their involvement in biological processes.
The discovery of nitric oxide, a free radical, as an endogenously generated effector molecule, was a paradigm shift in biological signaling. Nitric oxide (NO•, IUPAC names nitrogen monoxide, oxidonitrogen(•), or oxoazanyl) is a diatomic free radical produced in animals mainly by the enzymes nitric oxide synthases (NOS)
). It does not abstract hydrogen atoms, and it does not add to unsaturated bonds. Importantly, NO• does not react directly with thiols (RSH). Among the main targets of NO• in biological systems are metal centers. Coordination to the ferrous heme in soluble guanylate cyclase is responsible for many physiological effects of NO• (
). Nitric oxide can also react with oxygen, and this is analyzed in the next section on autoxidation.
The effects of NO• are exerted either via direct reactions with biological targets or indirectly via NO•-derived oxidants. Dysregulation of NO• homeostasis has been linked to neurodegeneration, cardiovascular disease, cancer, and inflammation. Therefore, the detection and quantification of NO• and its derived oxidants in vitro and in vivo are relevant to understanding the molecular bases of physiological as well as pathological processes.
Nitric oxide autoxidation
The reaction of NO• with oxygen (O2), termed autoxidation, is a complex process that gives different products and intermediates relevant to the detection of NO• and several of its derived species. This process is considered to be too slow to be of relevance under most physiological conditions. In the first step, two molecules of NO• and one molecule of O2 give two molecules of NO2• (Equation 1). Next, NO2• reacts with NO• reversibly to give N2O3 (Equation 2) (
). In water, in the absence of other targets, N2O3 is subsequently hydrolyzed to two molecules of NO2− (Equation 3).
The rate of decomposition of NO• is second order in NO• and first order in O2 concentrations, and in water the final stoichiometry is four molecules of NO• per O2, so that the rate equation is expressed as in Equation 4.
The limiting reaction in the autoxidation of NO• is the reaction with O2 (Equation 1); trapping subsequent products has no effect on the overall rate (
The autoxidation of NO• is slow in vivo because the rate of NO• decay is second order in NO• concentration, which is expected to be in the nanomolar range under normal conditions. This reaction can be accelerated in hydrophobic environments such as lipid membranes, lipoproteins, and proteins (
). This so-called “lens effect” may be of relevance where NO• concentrations are significantly increased, especially in sites of inflammation.
Detection of nitric oxide
Nitric oxide is difficult to measure in vivo because of its short half-life (typically in the range of 0.1–10 s), reactivity, and low steady-state concentration (i.e. nanomolar to micromolar). Nonetheless, several strategies have been developed to measure NO• or its derived species in vitro or in vivo that involve the use of absorbance, fluorescence, electron paramagnetic resonance (EPR), and electrochemistry (Fig. 2). Furthermore, NO• can be measured by chemiluminescence, a methodology that can be adapted to also measure other species. These methods are described in the following sections.
The identification of the endothelial-derived relaxing factor as NO• back in 1987 (
) was in part made by the change in the UV-visible spectrum of deoxyhemoglobin to form nitrosyl hemoglobin, with a corresponding shift of the Soret band from 433 to 406 nm. Nonetheless, the reaction mostly used to quantify NO•in vitro is with oxyhemoglobin, which is stable in air.
Oxyhemoglobin (Fe(II)(Hb)O2) is oxidized by NO• to yield NO3− and methemoglobin (Fe(III)(Hb)), which can be measured spectrophotometrically (Fig. 2A). The absorption of the Soret band is strong; thus the sensitivity of this method is relatively high (submicromolar). The maximum absorbance changes are observed at 401 (increased by reaction with NO•) and 421 nm (decreased), with an isosbestic point at 411 nm. If multiple wavelengths can be measured, the reaction can be followed by the absorbance difference of 401–421 nm (Δϵ401–421 = 77 mm−1 cm−1) (
). If there is interference at these wavelengths, the absorbance at 577 nm can be used (Δϵ577 = 10 mm−1 cm−1) or even both absorbances at 577 and 630 nm to calculate oxy- and methemoglobin concentrations before and after addition of NO• (
), oxyhemoglobin is also often used as NO• scavenger. Nitrite can also oxidize oxy- to methemoglobin but at much lower rates (although autocatalytically), so when high concentrations of NO2− are expected, like with the use of NO• donors for long-time periods, the contribution of NO2− to hemoglobin oxidation should be considered. Peroxynitrite can also oxidize oxyhemoglobin; thus, addition of superoxide dismutase is recommended as a control to prevent peroxynitrite formation from NO• and O2•− so that the methemoglobin formed can be associated with NO•. In addition, a potential O2•−-dependent redox cycling of hemoglobin can be avoided.
Electrodes specific for NO• are commercially available, but many research laboratories make their own. Typically, they consist of a filament made of carbon or platinum and a coating to provide specificity that either attracts NO• (
). At the anode, NO• is oxidized by one-electron to nitrosonium cation (NO+), which is converted to NO2− (Fig. 2B). The current generated from NO• oxidation is directly proportional to NO• concentration with a 10 nm detection limit (a calibration curve should be run with each experiment). The electrode is covered with a gas-permeable membrane that allows diffusion of NO• but not NO2− or other charged species. Temperature should be kept constant considering that solubility of NO• gas is very temperature-sensitive. Microelectrodes have been designed (<1 mm) that allow direct detection in cells in real time (
Although NO• is a free radical, i.e. it has an unpaired electron, it is difficult to detect directly by EPR, and spin-trapping techniques have to be used. Nitronyl nitroxides (with nitrone and nitroxide functional groups) are used as NO• probes (
). They react with NO• to give an iminonitroxide (Fig. 2C) with a dramatic change in the EPR spectrum that can be followed in a continuous and quantitative way. For example, 2-phenyl-4,4,5,5-tetramethylimidazoline-1-yloxyl-3-oxide (PTIO) or its water-soluble analogue carboxy-PTIO react with NO• with second-order rates constants of 104m−1 s−1 and a change in the EPR spectrum from five to seven lines (
Due to its fast reaction with NO•, carboxy-PTIO is often used as a scavenger of NO•; however, care should be taken because NO2• is a product of the reaction and has its own reactivities. In addition, biological reductants like thiols, ascorbate, or O2•− can nonspecifically reduce the nitroxides. Encapsulation of PTIO in liposomes has been used to avoid reduction (
Hydrophobic and hydrophilic nitroxides are available that allow detection of NO• at different depths of a biological membrane. Collisions of NO• with spin labels located in water or in membranes alter both the linewidth and the spin-lattice relaxation time that can be used to qualitatively and quantitatively measure NO• (
Colloid iron diethyldithiocarbamate (Fe(DETC)2) or N-methyl-d-glucamine dithiocarbamate are reliable spin traps for NO• detection. They form iron nitrosyl complexes with characteristic three-line EPR spectra (gav = 2.04; aN = 1.27 mT) at room temperature that are stable in the presence of oxygen (
). Dinitrosyl iron complexes (DNIC) with thiol-containing ligands have been detected in animal and bacterial cells by EPR. These complexes are formed in vivo in the paramagnetic (EPR-active) mononuclear as well as diamagnetic (EPR-silent) binuclear forms. The amount of NO• can calculated from the EPR amplitude signal because the linewidth of the NO-Fe(DETC)2 EPR spectrum may vary considerably with variations in the amount of Fe(DETC)2 in membrane lipids and the amount of Fe(III) present (
In addition, deoxyhemoglobin or other hemeproteins in the reduced Fe(II) form react with NO• to form nitrosyl–heme complexes that besides having characteristic UV-visible spectra are paramagnetic and can be followed by EPR. However, because of the instability of the complexes, the EPR spectra should be run at low temperatures (77 K) (
Fluorogenic probes have been developed that specifically react with NO•-derived species (i.e. N2O3) to yield fluorescent products, such as diaminonaphthalene (DAN) and diaminofluorescein (DAF) derivatives (Fig. 3D). The most popular of this kind is 4,5-diaminofluorescein (DAF-2), where nitrosation results in the highly-fluorescent triazole DAF-2 T (λexc = 495 nm, λem = 515 nm). The esterified diacetate derivative (DAF-2-DA) is also commercially available. It is highly membrane-permeable and detects intracellular nitrosation of the probe.
The pH should be carefully controlled because DAF-2 T fluorescence is pH-sensitive (
). Multiple and long exposures to excitation light, instead of causing photobleaching of the dye, potentiate the fluorescence response. Thus, minimum periods of light exposure are recommended. The use of 4-amino-5-methylamino-2′,7′-difluorofluorescein (DAF-FM) is favored because of its increased photostability, stability to pH, and reactivity toward NO•-derived species (
). The sensitivity of DAF-FM is 1.4 times higher than that of DAF-2. This increase of sensitivity is thought to result from the higher rate of the reaction with nitrosating NO+ equivalents due to the electron-donating effect of the methyl group (
The proposed mechanism of triazole formation involves N2O3 reacting with an amine to form an intermediate N-nitrosamine that at neutral pH can diazotize and then react with the second amine to yield the triazole (Fig. 3D) (
). Alternatively, a radical intermediate of the diamino-probe is formed by NO2• or other strong oxidants (e.g. radicals derived from peroxynitrite or peroxidases/H2O2) that then react with NO• (Fig. 3D) (
). These probes show some specificity issues, in that the triazole is not an exclusive product of NO•, and fluorescent products can be derived from peroxynitrite, nitroxyl (HNO), and ascorbic acid, complicating the interpretation of results (
). For example, one sensor has a fusion between a fluorescent protein and a bacteria-derived NO• domain that selectively binds NO• via a nonheme Fe(II) center. Once NO• binds, the domain gets closer to the fluorescent protein and quenches its emission (
The sample is injected into a purge vessel containing a given reactant such as triiodide. This vessel has fritted glass at the base and is purged at a constant flow rate with nitrogen or helium gas. The NO• that was injected (or generated) in the vessel is carried by this inert gas to the detector (
). The NO• in the carrier gas passes first through a reaction cell where ozone is constantly introduced. The reaction with ozone (O3) generates NO2• in the excited state (NO2•*) that is then carried by the inert gas flow to the detection cell where red and near IR light emission from NO2•* decay to the basal state is measured (
). The intensity of emission is directly proportional to the amount of NO• (Fig. 2E).
This method is not only useful to the study of NO• but also of other oxidation products that can be converted to NO• through different methods, such as NO2•, NO2−, S-nitrosothiols, nitrosyl–metal complexes, and N-nitrosamines (
). In the case of a biological sample that contains a mixture of these species, several tubes are prepared that include the parent sample, then one with acidic sulfanilamide to trap NO2−, and another that also includes HgCl2 to decompose S-nitrosothiols (
). The difference in the measured NO• with the different treatments indicate how much NO2− and S-nitrosothiol were in the sample.
Additional methods for more selective chemical reduction of S-nitrosothiols include copper-based assays where the reactant in the purge vessel consists of a buffer at neutral pH and Cu(II) plus an excess of cysteine (
). Therefore, the rate of NO• formation can be estimated from the rate of citrulline formation from arginine and saturating concentrations of NOS cofactors (NADPH, FAD, FMN, tetrahydrobiopterin, and calcium/calmodulin). Radiolabeled arginine is used, and the reaction is stopped with EDTA, which binds calcium and inactivates the enzyme. The radiolabeled citrulline product is separated from arginine by cation-exchange chromatography (cationic arginine is retarded and zwitterionic citrulline is eluted) and measured in a liquid scintillation counter (
). Because citrulline in the cell could be derived from non-NOS pathways, controls should be performed with addition of a NOS inhibitor as well as omission of NADPH.
There are commercially available kits to follow NOS activity indirectly, by measuring the time course of NO2− formation spectrophotometrically using the Griess reaction described below.
Bioassays for nitric oxide
The production of NO• in mammalian cells can be detected indirectly by measuring its biological activities like vasodilation, platelet aggregation, and guanylate cyclase activation (Fig. 2G).
In the cellular context, cyclic GMP (cGMP) is formed not only by guanylate cyclases stimulated by NO• (NO-GC or soluble GC) but also by the membrane natriuretic peptide receptor-coupled guanylate cyclases (GC-A and GC-B). Therefore, to measure levels of cGMP as an indirect measurement of NO•, controls with inhibitors of NOS should be included. The different methods used to determine cGMP have been recently reviewed in Ref.
). This function is explained by the formation of NO• from endothelial NOS, subsequent diffusion to the underlying smooth muscle, and activation of soluble guanylate cyclase, which initiates a signaling cascade that ultimately leads to vasodilation and increased blood flow. Thus, a method amply used by physiologists and pharmacologists to detect production of NO• consists of measuring tension in isolated vascular preparations treated with agonist and antagonists of NO•-dependent signaling (
). Washed human platelets are equilibrated at 37 °C in a turbidometric platelet aggregometer in the absence and presence of a system that produces NO•. An activator like thrombin is added to induce aggregation, and turbidimetry is followed with time (
Nitrogen dioxide (NO2•) is a reddish-brown free radical gas that forms part of air pollution. In biological systems, there are different endogenous sources of NO2•. These include: (a) NO• autoxidation (see section above); (b) NO2− oxidation, a reaction that is catalyzed by different heme-dependent peroxidases (Equations 5–7) and Cu,Zn-superoxide dismutase (
A tale of two controversies: defining both the role of peroxidases in nitrotyrosine formation in vivo using eosinophil peroxidase and myeloperoxidase-deficient mice, and the nature of peroxidase-generated reactive nitrogen species.
); and (c) homolysis of the peroxo bond of peroxynitrous acid (ONOOH) or of short-lived adducts formed from the reaction of peroxynitrite (ONOO−) with carbon dioxide (CO2), with carbonyl-containing compounds, or with metal centers (
). However, under most physiological conditions where NO2• concentrations are low (<1 μm), dimerization, which is a reversible process with a Keq = 7 × 104m−1, is outcompeted by bimolecular reactions of NO2• with different targets, some of which are far more concentrated (
). The partition coefficients in organic solvents indicate that NO2• is slightly hydrophobic, although less than NO•, which suggests a minor “lens effect” for NO2• reaction kinetics in membranes or other hydrophobic biological systems (
). Finally, NO2• reacts at diffusion-controlled rates with other radical species, such as tyrosyl radicals in proteins, to form 3-nitrotyrosine (see section on 3-nitrotyrosine below). The reversible reaction with NO• leads to the formation of the nitrosating species N2O3 (Equation 2).
Detection of nitrogen dioxide
The UV-visible absorption spectrum of NO2• shows a broad band peak at ∼400 nm with an absorption coefficient of 200 m−1 cm−1 in aqueous solution (
). The low absorption coefficient, the low stability of the radical, and the need to make corrections for N2O4 and NO2− absorption limit the technique. Nitrogen dioxide is frequently detected and quantified by chemiluminescence methods, in which the intensity of the emitted light is proportional to the concentration of NO2•. Some of these methods rely on the reaction of NO• with ozone (see section above on ozone-based chemiluminescence detection of nitric oxide and related species) and therefore require that NO2• be reduced to NO• using particular catalytic converters. The latter are usually nonspecific due to reduction of other nitrogen-containing compounds (
). In addition, luminol (5-amino-2,3-dihydro-1,4-phthalazinedione) in alkaline solution reacts with NO2• giving rise to intense chemiluminescence, although other one-electron oxidants can also lead to light emission (
Detection of NO2• in cells and tissues requires different methodologies. Because NO2• is a strong one-electron oxidant, it can react with typical redox probes such as 2′,7′-dichlorodihydrofluorescein (DCFH2). In experimental designs, addition of NO2− may be useful to convert other one-electron oxidants to NO2•.
One strategy depends on the ability of NO2• to rapidly combine with free or protein tyrosyl radicals to form 3-nitrotyrosine (
). This is analyzed in the section below on 3-nitrotyrosine. Furthermore, nitration of green fluorescent protein (GFP) leads to a decrease in its intrinsic fluorescence and was used to evaluate NO2• formation. Although the decrease in fluorescence intensity is not specific for nitration, it can be utilized in combination with pharmacological modulation of NO• levels to indicate NO2• formation (
Finally, because of the radical nature of NO2•, EPR has been utilized either by direct detection of NO2• in salt matrices and low temperatures or by using spin traps such as nitromethane at alkaline pH or nitrone compounds (
Due to the short half-life of NO2• in aqueous solutions even in the absence of other targets, as a result of dimerization and hydrolysis of N2O4, the study of the kinetics of NO2• reactions requires the use of very fast methodologies that allow measurements to be made in the microsecond time scale, such as pulse radiolysis. Furthermore, the low absorption coefficient of NO2• limits its direct detection so that product monitoring or competition kinetics need to be used. Competition with 2,2′-azino-bis-(3-ethylbenzothiazoline-6-sulfonic acid) (ABTS) oxidation to ABTS+ is frequently employed due to the high extinction coefficient of the latter radical at 414 nm (3.6 × 104m−1 cm−1) (
), it was suggested that N2O3 formation should be accelerated in hydrophobic regions. However, the nitrosation of thiols buried in the hydrophobic regions actually decreases because the dissociation to the more reactive thiolate is disfavored (
), so that S-nitrosothiols are not necessarily good indicators of N2O3 formation (see section below on S-nitrosothiols).
Besides measuring NO2− and S-nitrosothiols, another method to detect N2O3 is to use fluorogenic probes such as diaminonaphthalene (DAN) and diaminofluorescein (DAF), which were discussed in the section on detection of nitric oxide and suffer from the same issues as S-nitrosothiols.
Nitrosyl–metal complexes and their detection
Intracellular dinitrosyl iron complexes (DNICs) are formed from NO•, a ligand such as GSH, and loosely bound iron, also called labile or chelatable iron pool (
). EPR has several advantages such as the ability to measure signals in optically opaque samples, a good sensitivity (200 nm), and the capacity to distinguish enzymatically generated NO• by the change in the spectrum using [15N]arginine (
DNICs made with GSH can also be analyzed by UV-visible spectrophotometry, provided there is a separation step such as HPLC. They show a spectrum with maximum absorbance below 200 nm and characteristic peaks at 310, 360, and 680 nm (ϵ = 9200, 7400, and 200 m−1 cm−1, respectively) for the diamagnetic binuclear form, or 390 nm (ϵ = 3900 m−1 cm−1) for the paramagnetic mononuclear form (
Cellular DNICs can also be quantified through ozone-based chemiluminescence, using the triiodide method. Care should be taken in quantification because the signal is time-sensitive and decays within hours, and also because DNICs are sensitive to HgCl2, analogously to S-nitrosothiols (
). To distinguish between signals from S-nitrosothiols and DNICs, it was proposed to stabilize S-nitrosothiols in the cell lysate using a buffer containing diethylenetriaminepentaacetic acid and N-ethylmaleimide and to analyze the sample immediately after extraction and 20 h later to ensure the full decay of DNICs (
Formation of protein nitrosyl–metal complexes is particularly relevant in red blood cells, because NO• can react with deoxyhemoglobin to yield nitrosyl-hemoglobin. The detection of this product predominates at low oxygen tensions (
). In red blood cells, nitrosyl-hemoglobin is difficult to quantify by spectrophotometry where there is a mixture of different forms of hemoglobin absorbing at the same wavelength. EPR, in contrast, is specific for nitrosyl-hemoglobin and allows its quantification in packed red blood cells. The limit of quantification was calculated to be 200 nm. Under normal conditions, the amount of nitrosyl-hemoglobin in human blood is below the detection limit. However, patients exposed to 80 ppm NO• inhalation treatment increased its nitrosyl-hemoglobin levels up to 2 μm (
The formation of S-nitrosothiols is undoubtedly linked to the formation of NO• in biological systems. Nevertheless, the exact chemistry is still under debate. In fact, thiols or rather thiolates can be nitrosated by the products of NO• autoxidation (see section above on autoxidation) in a direct mechanism by N2O3 or stepwise by NO2• and NO• (Equation 9-11) (
Several methods have been developed to quantify total S-nitrosothiols and also to identify proteins that are nitrosated. S-Nitrosothiols show a UV spectrum with a maximum at 335 nm. The absorptivity for S-nitrosoglutathione at 335 nm is 922 m−1 cm−1; therefore, the sensitivity of the spectrophotometric analysis is low (above 50 μm) and depends on having a purified sample or on chromatographic or capillary electrophoresis separation (
). The released NO2− is then measured by the Griess method. This method has micromolar sensitivity (see section below on detection of nitrite).
Antibodies against S-nitrosocysteine have been used in immunohistochemical assays, Western blotting, and immunoprecipitation. However, specificity issues and the advent of biotin switch techniques that also allow mapping the modified cysteine within a protein have discouraged their use (
The gold standard method to quantify S-nitrosothiols is ozone-based chemiluminescence that provides nanomolar sensitivity and is appropriate for most biological applications (Fig. 3). As discussed in the section on chemiluminescence, S-nitrosothiols can be quantified using the triiodide reaction and also using copper ions and reductants (
). The original method involved blocking free thiols in S-nitrosated proteins with methyl methanethiosulfonate, then specifically reducing the nitrosated thiols with ascorbate, followed by reaction of these thiols with N-[6-(biotinamido)hexyl]-3′-(2′-pyridyldithio)-propionamide. The biotinylated proteins could then be selectively captured by using the specific binding to immobilized streptavidin (
). Some issues with this method have been pointed out, namely that it is very difficult to ensure that all free thiols are effectively blocked in the first step, that ascorbate does not reduce S-nitrosothiols directly but through the intermediacy of metals in solution (
), and that no chemical trace is left to indicate that the thiol was effectively nitrosated.
Relative quantification of protein S-nitrosation can be achieved through different means, including isotope-coded affinity tags (ICAT) and stable isotope labeling by amino acids in cell culture (SILAC). Both methods are based on using a light and a heavy isotope-containing tag. In ICAT, samples to compare are processed in parallel and tagged with biotin derivatives that include either light or heavy isotope linkers, and then mixed and further processed (
). The cell lysates from both cell cultures can then be mixed and processed as in the biotin switch method. If the same peptide is enriched from both control and treated samples, it will elute at the same time in the LC-MS analysis, but the mass spectra will differ by a known number of Da, and the relative amounts can be calculated from the intensities in the MS peaks (
Nitrite (NO2−, IUPAC name dioxidonitrate(1−)) and nitrate (NO3−, IUPAC name trioxidonitrate(1−)) were considered for a long time to be rather inert products of NO• oxidation. The concentration of NO3− in plasma of fasting individuals is 20–40 μm, and it is considered to derive mostly from the reaction between NO• and oxyhemoglobin, but also from the diet (
). The reduction by deoxyhemoglobin is thought to be quantitatively the most important pathway for the generation of NO• from NO2− and responsible for the NO•-like effects of NO2− infusion in the presence of red blood cells (
). The NO2− reductase activity of deoxyhemoglobin leads to the formation of NO• and methemoglobin (Equation 12).
Detection of nitrite and nitrate
There are several methods to detect NO2− in biological samples. The simplest method to measure NO2− is the Griess method, which sensitivity is in the micromolar range. The method is based on the diazotization of sulfanilamide by NO2− in acidic pH and the subsequent reaction with N-(1-naphthyl)ethylenediamine to yield an intensely pink-colored product with absorption maximum at 540 nm (Fig. 4) (
). Lower concentrations of NO2− (down to 20 nm) can be quantified by the formation of the fluorescent triazole derivative of DAN (see section above on fluorescent detection). In this case, the reaction of NO2− with DAN is done at acidic pH (Fig. 3), and then the fluorescence of the product is measured at alkaline pH (
). For low concentrations such as those often encountered in biological samples, the ozone-based chemiluminescence method (see section on chemiluminescence and Fig. 3) offers the required nanomolar sensitivity. In this case, the purge vessel needs to be filled with the triiodide acidic solution that converts NO2− to NO• that is then carried to the detection cell by the carrier gas. Nitrate is measured by first reducing it chemically or enzymatically to NO2−. The contribution of other species such as S-nitrosothiols to the signal is controlled by running samples treated with acidic sulfanilamide to trap all free NO2−.
The product of the one-electron reduction of NO• is HNO (nitroxyl, azanone, nitrosyl hydride, and hydrogen oxonitrate). The reduction potential of this process, E0′ (NO•, H+/HNO) ∼−0.55 V at pH 7 (
), is quite low, but high enough to make endogenous HNO formation a possibility. Biological studies are usually performed using nitroxyl donors (e.g. Angeli's salt). The ground state of HNO is a singlet in which all the electrons are spin-paired, whereas that of NO− (nitroxyl anion, oxonitrate (1−)) is a triplet with two unpaired electrons (
). In vivo, the preferential reactions of HNO are with thiols and metal centers. For example, the reaction between HNO and GSH, which is present in millimolar concentrations inside cells, has a rate constant of 3.1 × 106m−1 s−1 (
Cobalt(III) porphyrins react with HNO and constitute the basis of an amperometric electrochemical sensor for HNO (Fig. 5D). In the resting state, the polarized electrode (0.8 V) contains Co(III) porphyrin. When the porphyrin reacts with HNO it forms a Co(III)–NO− complex that is oxidized releasing NO• and the Co(III) porphyrin, ready for another cycle. The current intensity is proportional to HNO, and the sensitivity is in the nanomolar range. The success of the electrode is based on the fact that HNO reacts with Co(III) and not with Co(II) porphyrins, whereas NO• reacts with Co(II) and not with Co(III). This is an advantage of Co(III) over Fe(III) porphyrins, which react both with NO• and HNO (
Nitroxyl can reduce Cu(II) to Cu(I) and NO•. This is the basis of a group of fluorogenic probes in which the reduction of the metal ion is concomitant with the release of fluorescence quenching (Fig. 5E). The probes should be used with caution for the potential reduction by other reductants, as well as interference from hydrogen sulfide (H2S), S-nitrosothiols and oxygen (
). Due to the complex chemistry and to the potential to react with other reductants and oxidants, the use of the nitroxide probes in biological systems is limited.
Nitroxyl reacts fast with thiols. The formation of GSH sulfinamide (GS(O)NH2) from the reaction of GSH with HNO can be used as footprint for HNO. An N-hydroxysulfenamide is formed as an intermediate, and the final sulfinamide can be separated and detected by HPLC or MS (Equation 13) (
A probe has been developed that consists of an ester of 2-mercapto-2-methylpropionic acid and a fluorescent compound. The reaction of HNO with the thiol forms an N-hydroxysulfenamide intermediate that cyclizes releasing the fluorophore (Fig. 5G) (
). The azaylides are indicative of the formation of HNO and can be detected by NMR and MS, although, depending on the phosphine used, they may hydrolyze to the corresponding phosphine oxide. Although arylphosphines are resistant to reductants, possible interference by S-nitrosothiols is a potential concern (
The azaylides are nucleophilic and can react with an adjacent electrophilic group such as an ester or a carbamate. When the azaylide attacks the carbonyl, alcohol is released, and a unique amide phosphine oxide product is formed (Fig. 5H). This product, as well as the alcohol, can serve as reporters for HNO. The hydrolysis of the probe should be controlled as well as possible interference from S-nitrosothiols (
Despite the progress in the development of methods to measure HNO, the potential limitations should be carefully addressed. More than one method should be used, preferentially in combination with HPLC or MS detection of HNO-specific products (
Peroxynitrite (ONOO−) and peroxynitrous acid (ONOOH) are formed through the diffusion-controlled reaction between O2•− and NO• (Equation 15). IUPAC names for ONOO− and ONOOH are (dioxido)oxidonitrate(1−) and (hydridodioxido)oxidonitrogen, respectively. In this text, the term peroxynitrite is used for the mixture of ONOO− and ONOOH, unless specified.
Peroxynitrite is a powerful one- and two-electron oxidant; the reduction potentials are E0′ (ONOOH, H+/NO2•, H2O) = 1.6 V and E0′ (ONOOH, H+/NO2−, H2O) = 1.3 V (
). Peroxynitrous acid can traverse membranes through simple diffusion, whereas ONOO− can use anion channels. The anion is a good nucleophile, and ONOOH can act as an electrophile. In buffer, ONOOH (pKa 6.8, Equation 16) can decay to nitric acid (HNO3) plus a 30% fraction of HO• and NO2• radicals (Equation 17), but this process (k = 0.9 s−1 at pH 7.4 and 37 °C) is relatively slow and has limited physiological significance. The most relevant biological targets for peroxynitrite are peroxiredoxins, GSH peroxidases, CO2, and metal centers (
). The peroxiredoxins are thiol-dependent peroxidases that constitute the most efficient peroxynitrite scavengers known to date, with rate constants of ∼0.1–10 × 107m−1 s−1 and high concentrations in different cellular compartments (
The reactions with metal centers are diverse. Peroxynitrite can be reduced by one electron yielding NO2• as the metal center is oxidized, or by two electrons yielding NO2−. In addition, some hemeproteins (e.g. methemoglobin) catalyze peroxynitrite isomerization to NO3−, whereas others (e.g. Fe(III) cytochrome c) do not react at all (
) has proven to be very useful for the quantification of stock solutions in the laboratory at alkaline pH, as well as for following ONOO− decay in stopped-flow kinetic experiments.
One analytical approach for the detection of peroxynitrite is the use of probes that react with peroxynitrite itself or with its downstream radicals (NO2•, CO3•−, and HO•). Because the specificity of the probes is not always straightforward, particularly for the latter, the modulation of O2•− and NO• formations, which are the precursors of peroxynitrite, should accompany the results obtained with probes. Another analytical approach to evidence the involvement of peroxynitrite in a certain biological process is the detection of nitrotyrosine, a stable product formed from the reaction of radicals derived from peroxynitrite with tyrosine residues. As in the case of the probes, confirmatory evidence is required. These approaches are described in the next sections.
A growing number of small fluorogenic organic molecules designed and synthesized to detect peroxynitrite are reported constantly, having different selectivity and sensitivity toward this oxidant. The basic common characteristic is to have weak basal fluorescence, which is largely increased upon oxidation (
). In terms of the reaction mechanism, fluorogenic probes can be divided in two main groups: 1) probes that react with the radicals derived from peroxynitrite and yield a fluorescent end product by a radical mechanism; and 2) probes that react directly through a nucleophilic attack by peroxynitrite anion (ONOO−) to a particular functional group of the electrophilic probe, releasing masked fluorescence. The probes that react directly with ONOO− are potentially more straightforward, specific, and quantitative. They must react fast (>105–106m−1 s−1) and outcompete other routes of decay. Besides, genetically-encoded fluorescent protein sensors for peroxynitrite have been described recently (
). They use similar principles as some of the chemical probes that lead to direct detection of this oxidant (i.e. boronate-based compounds, see below).
Importantly, detection methods based on probes reveal only a minor fraction of total peroxynitrite, because a large proportion reacts with other targets in the cell. Moreover, the fraction trapped by the probe may vary with cell type or metabolic state according to the abundance of alternative targets (
Probes that react with the radicals derived from peroxynitrite
Probes frequently used for oxidant detection in biological systems are reduced dyes like 2′,7′-dichlorodihydrofluorescein (DCFH2) and dihydrorhodamine (DHR-123). Although extensively used, they present a series of limitations and caveats (
) yielding a radical intermediate (DCF•−), which is afterward oxidized to highly resonant moieties responsible for the increase in fluorescence emission (DCF) (Fig. 6A). These probes do not react directly with peroxynitrite (
). Neither NO• nor O2•− are able to oxidize either probe at significant yields; however, these radicals may react with the radical intermediate (DCF•−) in termination reactions giving nonfluorescent products (
). In addition to peroxynitrite-derived radicals, other potent one-electron oxidants such as those produced from heme peroxidases and other metalloproteins in the presence of H2O2 can generate fluorescent DCF (