Open access

Oxidative Processes and Antioxidative Metaloenzymes

Written By

Janka Vašková, Ladislav Vaško and Ivan Kron

Submitted: November 15th, 2011 Published: October 3rd, 2012

DOI: 10.5772/50995

Chapter metrics overview

3,061 Chapter Downloads

View Full Metrics

1. Introduction

Oxidative processes are necessary for life. They provide the energy necessary for many cellular functions. Most chemical energy in the body exists as ATP, produced during aerobic respiration. Nutrient oxidation is carried out by reduced coenzymes in the mitochondria, which are oxidized in the respiratory chain. The electrons are transferred to the oxygen created proton gradient that allows for ATP generation. One-electron transmission leads to the formation of reactive oxygen species (ROS). A series of oxidation processes take place in the peroxisomes. Hydrogen peroxide arises as a by-product of the oxidation of very long, long and branched-chain fatty acids, amino acids, synthesis and deamination of biologically active molecules (hormones, neurotransmitters, etc.), and biotransformation of xenobiotics. Oxidation reactions are also used for the degradation of unneeded molecules to excrement form, e.g. purines to form uric acid by xantine oxidase reaction, where hydrogen peroxide is also produced. The body also makes use of ROS also against the invasion of microorganisms, as neutrophils produce hypochloric acid from superoxide radicals via NADPH oxidase. ROS and reactive nitrogen species (RNS) may also be of exogenous origin. They can be taken up through diet or ventilation, or sometimes due to ionising radiation. Given that highly reactive substances could damage the cells and the whole organism, they must be inactivated by an antioxidative defence system. Metaloenzymes, which contain transition metals, and other antioxidant enzymes have an important role in this stage. Various endogenous substances, which are necessary for enzyme activity, form part of the antioxidant defence. Reduced glutathione is the most important of these endogenous substances and is functional both as a cofactor of other enzymes and for its reducing effects on oxidized molecules. NADPH is needed for the reduction of glutathione. The whole system is often referred as the glutathione defence system. The antioxidant defence system is very complicated. It is influenced by a number of other factors and circumstances, by both synthesis of endogenous and by intake of exogenous anti- or pro-oxidant substances. It also includes the receipt, transport and binding of metals into organic compounds of the organism; not just those that are part of the redox phenomena but also toxic elements, e.g. those which show a high affinity for sulphur, such as Hg, Cd, Sb, As, and other. These can affect the whole defence system, but may also be disposed to integration into the metalotioneins, peptides with high content of cysteine (approximately 1/3 of amino acids).

It is necessary to realize that the whole system is inducible. The aim of this is to describe the important aspects of this system that are mediated via metaloproteins.


2. Sources of reactive oxygen species and free radicals in an organism

All aerobic organisms produce reactive oxygen species physiologically. The five most productive pathways are involved in regulating the production of ROS/RNS and the resulting effects on signalling cascades. The five mechanisms described produce ROS in a non-regulated mode. However, there are many sources within the cells that are only mentioned.

2.1. Regulated production of reactive oxygen and nitrogen species

2.1.1. Nitric oxide synthase (NOS)

Nitric oxide (NO) is produced from a guanidine nitrogen of L-arginine via electron transfer from NADPH in two successive steps. The enzyme responsible this exists in three isoforms: neuronal (nNOS, type I, NOS-I or NOS-1), endothelial (eNOS, type III, NOS-III or NOS-3) and inducible (iNOS, type II, NOS-II or NOS-2). nNOS and eNOS are constitutively expressed, but their activity is regulated by the intracellular Ca2+ concentration. nNOS exhibits NADPH-diaphorase (NADPH-d) activity. The NOS isoforms are homodimeric, bi-domain enzymes. Each monomer consists of a flavin-containing reductase domain linked to a heme-containing oxygenase domain by a calmodulin-binding sequence. Although it possesses very little structural resemblance to P450, the oxygenase domain of NOS is referred to as being “P450-like” due to the presence of iron protoporphyrin IX (heme), linked axially by a cysteine residue to the NOS protein, which carries out “P450-like” mono-oxygenation reactions [130]. The isoform iNOS is inducibly expressed in macrophages after stimulation by cytokines, lipopolysaccharides, and other immunologically relevant agents [21]. Expression of iNOS is regulated at the transcriptional and post-transcriptional level by signalling pathways that involve agents such as the redox-responsive transcription factor NF-κB or mitogen-activated protein kinases (MAPKs) [120]. NO is a reactive and unstable free radical gas that can cross cell membranes easily by diffusion independent of any release or uptake mechanism [86].

The rate of NO synthesis is affected to some extent by the availability of the substrate L-arginine and by the cofactor tetrahydrobiopterin (BH4). The physiological function of NO varies widely due to the diverse localization of isoforms within different cell populations of the body. In physiological concentrations, NO functions as an intracellular messenger [88]. In pathophysiological situations where iNOS is upregulated, the most common RNS generated are dinitrogen trioxide (N2O3) and peroxinitrite (ONOO-), both of which are able to induce nitrosative and oxidative stress [194]. Upon NOS activation in many inflammatory diseases nitrite (NO2-), the major oxidation product from NO, is produced. In activated neutrophils, this can be oxidized by the effect of myeloperoxidase (MPO) to form either nitryl ion (NO2+) or nitrogen dioxide (NO2) [30].

2.1.2. NADPH oxidase NADPH oxidase in phagocytic cells

Activated neutrophils and macrophages produce superoxide and its derivatives as cytotoxic agents forming part of the respiratory burst via the action of membrane bound NADPH oxidase on molecular oxygen. It is a heme-containing protein complex. Hydrogen peroxide (H2O2) is produced by activated macrophages in an inflammatory environment, at an estimated rate of 2-6 x 10-14 mol.l-1.cell-1 and may reach a concentration of 10-100 μM in the vicinity of these cells [46,106]. This multicomponent enzyme catalyzes the one-electron reduction of O2 to superoxide (O2•-), using NADPH as the electron donor through the transmembrane protein cytochrome b558. The transfer of electrons occurs from NADPH on the inner side of the plasma membrane to O2 on the outer side. During phagocytosis, the plasma membrane is internalized as the wall of the phygocytic vesicle, with what was once the outer membrane surface now facing the interior of the vesicle. This targets the delivery of O2•- and its reactive metabolites internally for localized microbicidal activity [11].

The massive production of antimicrobial and tumoricidal ROS in an inflammatory environment is called the “oxidative burst” and plays an important role as the first line of defence against environmental pathogens. The combined activities of NADPH oxidase and myeloperoxidase (MPO) in phagocytes leads to the production of hypochlorous acid (HClO), one of the strongest physiological oxidants and a powerful antimicrobial agent [76]. MPO is a heterodimeric, cationic and glycosylated heme enzyme. The enzyme is a 140-kDa dimer of identical halves, each consisting of two polypeptide chains of 108 and 466 amino acids. Each half contains a covalently attached heme [7]. Like the other heme peroxidases, MPO combines with hydrogen peroxide and in the presence of halide (chloride, bromide, or iodide) to form the highly reactive redox intermediate in the phagosomes of neutrophils.

HOCl ↔ H+ +OCl-
HOCl + Cl- → Cl2 + HO

The oxidation of iron-sulfur centres in micro-organisms by the myeloperoxidase-H2O2-halide system may contribute to the death of an organism. MPO also catalyzes the oxidation of tyrosine in organisms to form the toxic amino acid residue, tyrosyl radical (Tyr), involved in the activity of neutrophils. Activated neutrophils and macrophages also generate singlet oxygen (1O2) by reactions that involve either MPO or NADPH oxidase [23,46]. Neutrophils also produce RNS, which can react with superoxide (O2•-) to produce ONOO-, itself a powerful oxidant, which may decompose to form a hydroxyl radical (OH). The activation of phagocytic NADPH oxidase can be induced by microbial products such as bacterial lipopolysaccharide, by lipoproteins, or by the cytokines interferon-γ, interleukin-1β, and interleukin-8 [23]. Enzyme activation is mainly controlled by rac2 in neutrophils and rac1 in macrophages and monocytes [46,112]. NADPH oxidase in nonphagocytic cells

Fibroblasts, endothelial cells, vascular smooth muscle cells, cardiac monocytes and thyroid tissue nonphagocytic NAD(P)H oxidase (similar but not identical to phagocytic NADPH oxidase) produce O2•- and to regulate intracellular signalling cascades [66,208]. In most of these, rac1 is involved in the induction of NAD(P)H oxidase activity [91,210]. Muscle cells and fibroblasts account for the majority of O2•- produced in the normal vessel wall. The NAD(P)H oxidase isoforms of the cardiovascular system are membrane-associated enzymes that appear to utilize both NADH and NADPH [66]. The rate of O2•- production in non-phagocytic cells is only about one-third that of neutrophils. O2•- and H2O2 are mainly produced intracellularly in vascular smooth muscle cells, in contrast to neutrophils, endothelial cells, and fibroblasts. The cardiovascular NAD(P)H oxidase isoforms are induced by hormones, hemodynamic forces, or by local metabolic changes [66]. Mechanical forces stimulate NAD(P)H oxidase activity in endothelial cells and reoxygenation in cardiac myocytes. An NAD(P)H oxidase with low affinity for oxygen and high affinity for cyanide is believed to act as one of the sensors for oxygen tension in the carotid body, controlling the rate of ventilation [2]. The function of oxygen sensing is apparently shared by several proteins, including a nonmitochondrial cytochrome b558, a mitochondrial protein, and possibly a third heme protein [105,209].

A similar group of proteins was suggested to be involved as oxygen sensors in the regulation of erythropoietin production in human hepatoma cells [209]. A microsomal NADH oxidase was implicated as an oxygen sensor in bovine pulmonary and coronary arteries, where changes in oxygen tension regulate vascular relaxation through changes in O2•- production and cGMP formation [198]. Increased aortic adventitial O2•- production contributes to hypertension by blocking the vasodilatatory effects of NO [189]. There is a strong possibility that rac-like proteins also occur in plants [1,193], where they may be involved in the induction of NAD(P)H oxidase-like enzymes [167]. The oxidative burst in plants is an effective bactericidal mechanism.

2.1.3. Arachidonate cascade enzymes 5-lipoxygenase (5-LOX)

The enzyme 5-LOX has been identified as an inducible source of ROS production in lymphocytes [23,118,126], but the evidence for its physiological role in redox signalling is still scarce. There are several lipoxygenases which differ by substrate specificity and optimum reaction conditions. Lipoxygenases in plants and animals are heme containing dioxygenases that oxidize polyunsaturated fatty acids at specific carbon sites to give enantiomers of hydroperoxide derivatives with conjugated double bonds. The number in specific enzyme names such as 5-LOX, 12-LOX, or 15-LOX refers to the arachidonic acid site that is predominantly oxidized [202]. 5-LOX is best known for its role in biosynthesis of the leukotrienes A4, B4, C4, D4 and E4. The oxidized metabolites generated by 5-LOX were found to change the intracellular redox balance and to induce signal transduction pathways and gene expression. 5-LOX was shown to be involved in the production of H2O2 by T lymphocytes after ligation of the CD28 costimulatory receptor [118] and in response to interleukin-1β [23]. A lipid metabolizing enzyme in fibroblasts similar to 15-LOX has been shown to generate large amounts of extracellular O2.- [168]. Cyclooxygenase (COX-1)

Cyclooxygenase-1 has been implicated in ROS production through formation of endoperoxides, which are susceptible to scavenging by some antioxidants in cells stimulated with TNF-α, interleukin-1, bacterial lipopolysaccharide, or the tumor promoter 4-O-tetradecanoylphorbol-13-acetate [48]. Cyclooxygenase participation in redox signalling remains scarce.

2.2. Non-regulated production of reactive oxygen species

2.2.1. Mitochondrial respiration

The four-electron reduction of oxygen occurs within the mitochondrial electron transport system of all cells undergoing aerobic respiration. It is estimated that 2-3% of O2 consumed by mitochondria is incompletely reduced, yielding ROS [173] and 1-5% leads to H2O2 production [134]. It is well documented that mitochondria are a source of H2O2; however, the release of O2•- from mitochondria into the cytosol has yet to be definitively established [77]. ROS are only produced at complexes I and II in the mitochondrial matrix, while complex III is capable of producing ROS on both sides of the mitochondrial inner membrane [135,173]. It is generally thought that the two major sites of mitochondrial ROS production are complexes I and III. NADH-ubiquinone oxidoreductase (complex I) is composed of ̴ 45 subunits and is the site of NADH oxidation. The flavin mononucleotide (FMN) of complex I accepts the electrons from NADH and passes them through a series of eight iron-sulfur clusters to ubiquionone [84] to generate O2•- in the presence of NADH. Complex I also generates ROS after the oxidation of succinate at complex II via a process referred to as reverse electron transport (RET). It is also hypothesized that ROS production from complex I during RET occurs from FMN as well [103,122]. Ubiquinol:cytochrome c oxidoreductase (complex III) has 11 subunits and contains 3 hemes and an Fe-S cluster center. Complex III plays an intricate role in passaging electrons from the ubiquinol generated by complexes I and II to cytochrome c [116]. Upon binding with the Qo site, one electron from ubiquinol is transferred through the Rieske Fe-S cluster protein to the electron acceptor, cytochrome c. The resulting unstable semiquinone then donates the remaining electron to the heme groups on cytochrome b. The electron in cytochrome b is then used to re-reduce ubiquinone at the Qi site to produce ubiquinol. Two electrons from semiquinones in Qo are required for the reduction of ubiquinone to ubiquinol in the Qi site. This process is referred to as the Q-cycle because lone electrons remaining in semiquinone are reused to reduce ubiquinone back to ubiquinol [35].

The mechanism of mitochondrial production and release of H2O2 and O2•- takes place in two steps. Firstly, part of O2•- generated during mitochondrial electron transfer is vectorially released into the intermembrane space [78]. The mechanism underlying the release of O2•- into the intermembrane space covers the formation of ubisemiquinone at two sites in the ubiquinone pool: the Q1 site that lies near the matrix, and the Qo site in the vicinity of the intermembrane space [154]. Autooxidation of ubisemiqiunone at the Qo site (UQo•-) results in the release of O2•- through the cytosolic side of the mitochondrial inner membrane. O2•- cannot cross membranes, except in the protonated form, which represents only a small fraction of the O2•- pool at physiological pH. Taken together, H2O2 is formed both at the intermembrane space and the matrix from O2•- generated towards the respective compartments [77]. Second, the release of O2.- into the intermembrane space would be in a functional relationship to the localization of a superoxide dismutase (SOD) activity in this compartment. The intermembrane space contains several O2•- scavenging pathways besides SOD, such as cytochrome c [179] as well as pores for O2•- diffusion across the outer membrane into cytosol, in particular the voltage-dependent anion channel [77].

O2•- released into the cytoplasm from mitochondria could play an important role in cell signalling, as O2•- has been implicated in several signalling events. In addition, cytoplasmic aconitase and other cytoplasmic enzymes susceptible to O2•- may be targets of O2•- released from mitochondria [61]. Another important decay pathway of O2•- at a diffusion-controlled rate may involve the reaction with NO to yield ONOO- in the intermembrane compartment. This may be of some significance, as nitrosation of cytochrome c and proapoptotic caspases occurs prior to apoptosis [123].

2.2.2. Chloroplasts

The ability of phototrophs to convert light into biological energy is critical for life and therefore organisms capable of photosynthesis are especially at risk of oxidative damage, due to their bioenergetic lifestyle and the abundance of photosenzitizers and oxidable polyunsaturated fatty acids in the chloroplast envelope. The presence of O2 in the atmosphere enables respiratory metabolism and efficient energy generation systems which use O2 as final electron acceptor, leading to the formation of ROS in cells [166]. The presence of ROS producing centres such as triplet chlorophyll, and ETC in PS I and PS II make chloroplasts a major site of ROS production in plants and algae [145]. Atmospheric oxygen is relatively non-reactive. It has been estimated that 1-2 % of O2 consumed by plants is sidetracked to produce ROS in various subcellular loci [19].

Oxygen generated in chloroplasts during photosynthesis can accept electrons passing through the photosystems (PS). PS II is a multisubunit protein complex also present in cyanobacteria that use light energy for oxidation of water and reduction of plastoquinone [146]. Various abiotic stresses such as excess light, drought, salt stress and CO2 limiting conditions, enhance the production of ROS. Under normal conditions, the electron flow from the excited PS centres is directed to NADP+, which is reduced to NADPH. It then enters the Calvin cycle and reduces the final electron acceptor, CO2. In cases of ETC-overloading, a part of the electron flow is diverted from ferredoxin to O2, reducing it to O2•- via the Mehler reaction. The acceptor side of ETC in PS II also provides sides (QA, QB) with electron leakage to O2 producing O2•-. On the external “stromal” membrane surface, O2•- is enzymatically dismutated to H2O2 [50,65,163]. 1O2 is a natural byproduct of photosynthesis, mainly formed at PS II even under low-light conditions [29]. Generation takes place due to the excitation energy transfer from triplet chlorophyll formed by the intersystem crossing from singlet chlorophyll and the charge recombination of separated charges in the PS II antenna complex and reaction center of PS II [146]. In cases of insufficient energy dissipation, the chlorophyll triplet state becomes able to react with 3O2 to give up 1O2 [79].

2.2.3. Xanthine oxidoreductase (XOR)

XOR exists as either an oxidase (XO) which transfers reducing equivalents to oxygen, or as a dehydrogenase (XDH) that utilizes NAD or oxygen as the final electron acceptor [17,59]. The enzyme is derived from xanthine dehydrogenase by proteolytic cleavage. It contains molybdenum in the form of molybdopterine, and two clusters with iron and sulfur compounds of FAD cofactor in both subunits. The enzyme catalyzes the production of uric acid with co-production of O2•-. The physiological substrates, xanthine and hypoxanthine, bind with the oxidized enzyme and donate two electrons into the molybdenum cofactor reducing it from Mo6+ to Mo4+. Substrates are hydroxylated by H2O at the molybdenum site as the electrons travel via two iron-sulfide residues to flavine–adenine dinucleotide (FAD). Reduced FAD can be divalently reoxidized by oxygen to produce hydrogen peroxide, or univalently reoxidized in two steps to generate two equivalents of superoxide O2•- [17,82]. Under normal conditions, XOR accounts for only a minor proportion of total ROS production [46]. The release of O2•- results in the recruitment and activation of neutrophils and their adherence to endothelial cells, stimulating formation of XOR in the endothelium with further O2•- production. Therefore, it has been observed in TNF-treated endothelial cells [58] and has been implicated as a major source of oxidative stress under ischemia and reperfusion [46].

2.2.4. Dopamine (DA)

As a neurotransmitter, DA is stable in the synaptic vesicle. When an excess of cytosolic DA exists outside of the synaptic vesicle, DA is easily metabolized via monoamino oxidase (MAO) or by autooxidation to produce ROS, subsequently leading to the formation of neuromelanin [162]. During the oxidation of DA by MAO, H2O2 and dihydroxyphenylacetic acid are generated [67]. Spontaneously oxidized cytosolic DA produces O2•- and reactive quinones such as DA quinones or DOPA quinones. DA quinones are also generated in the enzymatic oxidation of DA by COX in the form of prostaglandin H synthase, LOX, tyrosinase and XOR. These quinones are easily oxidized to the cyclized aminochromes: DA-chrome and DOPA-chrome, and are then finally polymerized to form melanin, as reviewed in Miyazaki & Asanuma [132]. Although ROS from the autooxidation of DA show widespread toxicity not only in DA neurons but also in other regions, highly reactive DA quinone or DOPA quinone exert cytotoxicity predominantly in DA neurons and surrounding neural cells. It is thought that DA acts as an endogenous neurotoxin, contributing to the pathology of neurodegenerative disorders and ischemia-induced damage in the striatum [24,124,201].

2.2.5. Photosensitization reactions

Photosensitization reactions involve the oxidation of organic compounds by atmospheric oxygen upon exposure to visible light. The photoexcitated state, most often the triplet state of the sensitizer, is the key photoreactive intermediate and exerts photodamage through direct reaction with substrate molecules (type I photosensitization) or activation of molecular oxygen by energy transfer reactions (type II photosensitization) [199]. 1O2 is an excited state molecule formed by direct energy tranfer between the excited sensitizer and ground state 3O2. Less than 1% of triplet oxygen is converted in parallel to superoxide anion (O2-). The formation of O2- as a precursor of H2O2 occurs via electron transfer via production of a sensitizer radical cation, or after an intermediate reduction of the sensitizer with a substrate followed by the single electron reduction of O2 [38,99].

2.6. Other cellular ROS sources

The most studied producers of O2.- by oxidizing unsaturated fatty acids and xenobiotics are cytochrome P450 and the b5 family of enzymes [168]. Electrons leaking from nuclear membrane cytochrome oxidases and electron transport systems may give rise to ROS [75]. In addition to intracellular membrane-associated oxidases, aldehyde oxidase, dihydroorotate dehydrogenase, flavoprotein dehydrogenase and tryptofan dioxygenase can all generate ROS during catalytic cycling. pH-dependent cell wall peroxidases, germin-like oxalate oxidases and amine oxidases have been proposed as a source of H2O2 in the apoplast of plant cells [22]. Glycolate oxidase, D-amino acid oxidase, urate oxidase, flavin oxidase, L-α-hydroxy acid oxidase, and fatty acyl-CoA oxidase are important sources of total cellular H2O2 production in peroxisomes [168]. Auto-oxidation of small molecules such as epinephrine, flavins, and hydroquinones can also be an important source of intracellular ROS production [57].


3. Chemistry of reactive oxygen and nitrogen species

During plant photosynthesis and in analogous reactions of the respiratory chain, triplet oxygen is reduced to water (reaction 3). As a result of one-, two- and three- electron reduction, toxic forms of oxygen, free radicals and covalent compounds are produced as side products and oxidize additional biomolecules [181].

3O2 + 4ē → 2H2O

1O2 is the first excited electronic state of O2, and is an unusual ROS, as it is not related to electron transfer to O2. It is formed in photosensing reactions and is effectively quenched by β-carotene, tocoferols, plastoquinones and vitamin C. If not, 1O2 can lead to gene upregulation, involved in the molecular defence responses against photooxidative stress [102]. The lifetime of 1O2 in a cell has been measured to be approximately 3 μs [79] and in this time, a fraction of 1O2 may be able to diffuse over considerable distances of several hundred nanometers. Other studies have also found that 1O2 can last for nearly 4 μs in H2O and 100 μs in polar solvent [102].

The monovalent reduction of molecular oxygen, the one-electron reduction of 3O2 catalyzed by NADPH oxidases, gives rise to O2•- (reaction 4). O2•- has an approximate half-life of 2-4 μs and undergoes fast, non-enzymatic, one-electron reduction or dismutation in the Haber-Weiss reaction (reaction 5).

3O2 + ē → O2•-
O2•- + H2O2 → HO + HO- + 3O2

It has been noted that O2•- can undergo protonation to give up a strong oxidizing agent HO2 (reaction 6) which directly attacks the polyunsaturated fatty acids (PUFAs) in negatively charged membrane surfaces [65]. The hydrogen donor for the reduction of PUFAs may well be ascorbic acid, forming H2O2 and a radical of ascorbic acid. Enzymatic dismutation to H2O2 is the most effective quenching mechanism (reactions 7, 8).

O2•- + H+ → HO2
O2•- + ē → O22-
O22- + 2H+ → H2O2

The interaction of O2 with trace concentrations of redox-active transition metals leads to O2•- production (reaction 9) and the non-enzymatic reduction of O2•- in the presence of Fe forms 3O2 (reaction 10). At low pH, dismutation of O2•- is unavoidable, with one O2•- giving up its added electron to another O2•-, generating H2O2 following protonation (reaction 11) [181].

Mn+ + O2 → M(n+1)+1 + O2•-
O2•- + Fe3+3O2 + Fe2+
O2•- + H+ + HO2 → H2O2 + O2

H2O2 is produced by the two-electron reduction of 3O2 (reaction 12) and the univalent reduction of O2•-. H2O2 is moderately reactive and has a relatively long half-life (1 ms) [19]. It is broken down partially enzymatically by catalase or glutathione peroxidase to water or in case of substrate peroxides to corresponding alcohols and water. In cases where the speed of its decomposition is not sufficient, it may lead to its one-electron reduction (reaction 13).

3O2 + 2H + ē → H2O2
H2O2 + ē → HO.+ HO-

The reaction takes place similarly to the Haber-Weiss reaction in the presence of transition metals (Fenton reaction), producing the very reactive HO and HO- (reaction 14) [181]. Instead of O2•-, HO2 may arise, which is actually the H2O2 radical (reactions 15, 16). Common mechanisms involving the Fenton reaction, generation of the O2•- and HO appear to be involved for Fe, Cu, Cr, V, Co primarily associated with mitochondria, microsomes and peroxisomes. However, a recent discovery, that the upper limit of free pools of Cu is far less than a single atom per cell casts serious doubt on the role of Cu in Fenton-like generation of free radicals [178].

H2O2 + Fe2+ → HO+ HO-+ Fe3+
Fe2++ O2 + H2O → Fe3++ HO2 + HO-
Fe2++ HO2 + H2O → Fe3++ HO- + H2O2

HO is also generated by the three-electron reduction of 3O2 (reaction 17). It predominantly attacks the unsaturated fatty acids of membranes. The most effective protective mechanisms include reduction of HO2 by tocopherols, taking the form of tocopherol radicals, for which the retroactive reduction of the reaction requires the oxidation of ascorbic acid. Resulting hydroperoxides (R-O-OH) are released by phospholipase A2, which makes them available substrates for peroxidases. In the presence of suitable transition metals, especially Fe, HO can also be produced from O2•- and H2O2 at neutral pH and ambient temperatures by the iron-catalyzed Fenton reaction [187].

3O2 + 3ē + 3H+ → HO + H2O

NO is generated by specific NOSs, which metabolise arginine to citrulline via a five electron oxidative reaction [63]. NO reacts with O2•- (reaction 18) in a reaction with the highest rate constants known (7.0 x 109 m-1.s-1) [32]. ONOO- can be transformed into peroxynitrite acid and then to HO (reaction 19). NO binds certain transition metal ions; in fact, many effects of NO are exerted as a result of its initial binding to Fe2+ heme groups. The most commonly seen product of such a reaction is [Fe3+ NO-] [177].

NO + O2•- → ONOO-
ONOO- + H+ → ONOOH → HO + NO2 → NO3- + H+

4. Formation of radicals in biological systems and consequences of oxidation of biological molecules

4.1. Oxidation of lipids

This is considered to be the most damaging process known to occur in living organisms [62]. It includes a number of reactions leading to the development of oxidized lipids and fatty acids that give rise to free radicals. Oxidation products of lipids, particularly (2E)-4-hydroxyalk-2-enals and aldehydes such as malondialdehyde, as well as alkanes, lipid epoxides and alcohols, react with proteins and nucleic acids. The overall effects of lipid oxidation are a decrease in membrane fluidity, an increase in the leakiness of the membrane to substances that do not normally cross it except through specific channels and damage to membrane proteins, and inactivation of receptors, enzymes, and ion channels.

4.1.1. Oxidation by 3O2

The most common oxidation of fatty acids is by 3O2 from the air. Oxidation of unsaturated fatty acids only occurs in three stages at normal temperatures. In the initiation stage, free hydrogen H and fatty acid R emerge as the C-H covalent bond of the hydrocarbon chain is split. The energy required to split bonds can come from ultraviolet radiation, radioactivity, and also visible light. In the latter case, it is a two-electron oxidation of 1O2. A reaction also exists to break any binding with other free radicals or transition metals. During the second, propagation stage the reactive R quickly merges with O2, and produces a peroxyl radical (R-O-O). As the hydrogen atom splits from the hydrocarbon chain, another molecule of unsaturated fatty acid forms hydroperoxide (R-O-OH) and another R (reaction 20). The initiation rate of oxidation for the production of R-O-OH is slow (induction period) leading to a gradual accumulation of R-O-OH, followed by the creation of other radicals. As long as there is enough oxygen, the reaction takes place spontaneously, sharply rising to reach the maximum speed of reaction, in which reactive groups are diminished. The rate of this reaction then slows and starts to be overtaken by the degradation of R-O-OH. R-O-OH is very fragile and H splits from the molecule, leaving R-O-O (reactions 21, 22) or HO. According to the current knowledge, R-O-OH degradation with conjugated double bonds leads preferentially to formation of the alkoxyl radical (R-O) (reaction 23) [36].

R-H + R-O-O → R + R-O-OH
R-O-OH → R-O-O + H
R-O-OH → R-O-O + H2O
R-O-OH → R-O + HO

The reaction of R with O2 is much faster than with a hydrocarbon lipid chain. When the concentration of free radicals is high, it is likely that these will react together to form a nonradical product, which terminates the chain reaction. While R is prevalent in the reaction system, hydrocarbon radical recombination is the major termination reaction. If, however, there is a preponderance of R-O-O, the termination reaction leads either to recombination of R with R-O-O forming peroxide bridged dimers (reaction 24) or to the reciprocal recombination of R-O-O (reaction 25). In the case of unsaturated fatty acids, H+ splits from the methylene group near the double bond producing mainly R-O-OH. This reaction becomes easier as the number of double bonds increases. However, if the number of double bonds is unchanged, the double bond moves one carbon closer to the carboxyl or methylene end of the chain. By moving the double bonds, a double bond in the cis configuration is changed to more stable trans configuration.

R + R-O-O → ROOR

4.1.2. Oxidation by R-O-OH

R-O-OH of fatty acids and their radicals may react in three ways. In the first case, there is no change in the number of carbon atoms in the molecule. R-O-OH species from polyunsaturated fatty acids (PUFAs) containing three or more double bonds in a molecule are unstable, and they tend to pass in 1,4 cyclization to the six-member peroxides derived from 1,2-dioxanes, which are also unstable compounds and decompose to low molecular active products. R-O-OH molecules by 1,3 cyclisation pass to five-member peroxides, 1,2-dioxolanes and endoperoxides. The main malondialdehyde precursors emerge from 1,2 dioxolane-type peroxohydroperoxides. R-O-OH and R-O-O, react very easily with the double bond of unsaturated fatty acids to generate epoxides. The addition of R-O-O across a double bond can take place intermolecularly. R-O-OH is oxidized by the nonradical mechanism and the resulting epoxide is immediately hydrolyzed to dihydroxyderivatives. Epoxides can arise even with the addition of R-O to PUFA by intermolecular reactions. An accrued radical of epoxy acid reacts with oxygen to give HO2, from which R-O-OH is formed and subsequently, R-O. By the recombination of R-O with H, a competent hydroxyl acid or oxo acid arises by elimination of H+. In the second case, the molecule breaks and gives volatile and sensory active substances with less carbon atoms. Breaking the molecule takes place both due to the R-O created (reaction 26) and depending on the position of the double bond in relation to the hydroperoxide group. From this, saturated and unsaturated aldehydes, saturated and unsaturated hydrocarbons, and oxo acids are formed. The most reactive compounds formed are aldehydes, which are further oxidized and react with the proteins. Malondialdehyde is an important product of this oxidation [125]. The third mechanism is oxypolymerization, in which the number of carbons in the molecule is increased due to the reduction of two radicals. Concerning R-O, radicals are condensed by a -C-C- bond, which is not frequent, because R-O is less available. Therefore, the majority of radicals combine through ether-like -C-O-C- or peroxide-like -C-O-O-C- bonds.


4.1.3. Oxidation by 1O2

Excitation of the common 3O2 leads to a reactive 1O2 which may react with the double bond of unsaturated lipids and other unsaturated compounds. It reacts with the listed compounds because they are rich in electrons and are therefore able to fill its free molecular orbital [158]. The rate of reaction between common unsaturated acids and 1O2 is at least 1450-fold higher in comparison to the reaction with triplet oxygen. It has been found that the PUFAs (linoleic acid 18:2 and linolenic acid 18:3) are particularly susceptible to attack from 1O2 and HO [134]. Unstable cyclic peroxide compounds moloxides with four or six-member rings are formed by adduction across double bonds. Intermediate products of the reaction decompose rapidly and give rise to respective hydroperoxides.

By the reaction with an atom in methylene groups on the carboxyl end of fatty acids, R-O-OH arises in a similar process to peroxide oxidation by 3O2. However, the mechanism of primary production of hydroperoxides differs from the mechanisms of 3O2 oxidation, therefore producing a different ratio of constitutional isomers.

4.1.4. Oxidation catalyzed by metals

This type of oxidation is catalyzed by compounds of transition metals, especially Fe and Cu, which are present in tissues that are reduced by accepting an electron. They are involved directly or indirectly in initiation, propagation and termination reactions of radicals [181].

Metals in their higher oxidation state M(n+1)+ are responsible for initiating oxidation reactions. The electron transfer in the reactions leads to the formation of R (reaction 27)

M(n+1)+ + R-H → Mn+ + R + H+

The initial reaction is also indirectly catalyzed by metals in the lower oxidation state Mn+, producing a transient complex with the metal, oxygen and R-H before decaying to R, metals with higher oxidation state and ROS (reactions 28-32) [156,181].

Mn+ + O2 + R-H → [Mn+ O2 (R-H)] → R + M(n+1)+ + HO2-
Mn+ + O2 + R-H → [Mn+ O2(R-H)] → R + M(n+1)+ + HO2
Mn+ + O2 + R-H → [Mn+ O2 (R-H)] → RO + M(n+1)+ + HO-
Mn+ + O2+ R-H → [Mn+ O2(R-H)] → HO2 + [M(n+1)+ R-]
Mn+ + O2 → [Mn+O2] → M(n+1)+ + O2•-

Subsequently, the oxidation reaction is catalyzed by the ROS produced. The reaction of HO2 with unsaturated fatty acids is slow, while O2•- does not react at all. HO is more reactive (R-H + HO → R + H2O), and is generated by the Fenton reaction.

Metals in a lower oxidation state, such as Fe and Cu catalyze decomposition of R-O-OH to R-O (reaction 33) and, in their higher oxidation state, catalyze decomposition of R-O-OH to R-O-O (reaction 34). These emerging radicals increase the reaction rate by increasing the propagation phase rate, as the metal-catalyzed R-O-OH disintegration is faster than the emergence of new radicals.

R-O-OH + Mn+ → R-O + M(n+1)+
R-O-OH + M(n+1)+ → R-O-O + Mn+

Metals bound in complexes might or might not be effective depending on the environment. The addition of an iron complex to biological samples encouraged peroxidation by peroxide decomposition, generating R-O and R-O-O. The rate constant for this reaction when ferrous ions are involved, has been given as 1.5 x 103 mol-1.l-1.s-1, which is higher than the rate constant for the reaction of ferrous ions with H2O2 in the Fenton reaction 76 mol-1.l-1.s-1 [73]. The redox potentials of the metals Mn and Co are low and are therefore incapable of catalyzing the breakdown of R-O-OH in aqueous systems. In fats, however, they can catalyze the decomposition of R-O-OH through the transient hydroperoxide complexes to R-O-O. It is not yet known whether the oxidation of lipids can also be catalyzed by complexes of Fe with oxygen (Fe3+-O2-Fe2+) and hypervalent iron as ferryl cations FeO2+ and ferrate anions FeO42, which are the active forms in the enzymes containing heme cofactors, e. g. catalases and cytochrome P450. However, it is known that (ferric; Fe3+-Px) peroxidases mediate one-electron oxidation of organic compounds with the concomitant reduction of H2O2 to H2O. In this mechanism, peroxidase donates two electrons to H2O2 resulting in cleavage of H2O2 and formation of a redox intermediate of enzyme (I). This intermediate consists of an oxoferryl protein cation radical, in which one of the oxidation equivalents exists as the ferryl ion and the other as a porphyrin-centred cation radical (reaction 35). The enzyme intermediate reacts with reductants (R-H) to generate substrate free radicals and another redox intermediate (II), in which oxoferryl species remain intact but the cation radical is reduced. A one-electron reduction of II by a second molecule of reductant regenerates the ferric enzyme and forms a second equivalent of R (reaction 36). Another redox intermediate (III) is formed in the course of peroxidase catalytic cycle (reaction 37). It is catalytically inactive and exists as a resonance form between the Fe2+-O2 and Fe3+-O2- complexes [49,141].

Fe3+-Px + H2O2 → CI + H2O
CI + R-H → CII + R
CII + RH → Fe3+-Px + R + H2O

Also the perferryl [Fe5+] radicals are catalytically active in numerous biological processes, and these ferryl/perferryl moieties, whether as components of enzymes or simple iron complexes, can be very powerful oxidants capable of abstracting hydrogen atoms in lipid peroxidation [20]. Some metal ions with a fixed oxidation number can affect the rate of peroxidation, e.g. Ca2+, Al3+, and Pb2+ ions can accelerate peroxidation stimulated by iron salts under certain conditions [72].

Exposure to heavy metals can change the composition of the reaction products. High concentrations of free radicals may outweigh termination reactions, where the metals inhibit the oxidation. Inhibition of oxidation may occur with higher concentrations of metal ions. It is supposed that Fe and Cu ions oxidize and reduce hydrocarbon free radicals to their corresponding anions (reaction 38) and cations (reaction 39) together with the emergence of free radical complexes (reaction 40). Other complexes are formed with Co (reactions 40-42). All of them break the radical chain reaction.

R + Fe2+ → Fe3+ + R-
R + Fe3+ → Fe2+ + R+
R + Fe3+ → [Fe3+ R]
R + CoA3 → [R-CoA2]
R-O + CoA3 → [R-O-CoA2]
R-O-O + CoA3 → [R-O-O-CoA2]

4.2. Oxidation of proteins

The principal agents for protein oxidation are atmospheric O2, 1O2, O2•-, HO, R-O-OH and H2O2. Other agents that lead to protein oxidation include HOCl, xenobiotics, reduced transition metals, γ-irradiation in the presence of O2, activated neutrophils and oxidoreductase enzymes [153]. Free peroxyl radicals react with proteins and produce protein radicals, which then react with other free protein radicals to form dimers, or with free lipid radicals to form copolymers. A protein radical arises most frequently when the more labile hydrogen atom on Cα splits from the protein. A hydroxyl acid is obtained from an alkoxyl radical, and hydroperoxide from a peroxyl radical. Recombination of protein radicals leads subsequently to protein oligomeres.

Besides Trp and Tyr, sulfur-containing amino acids, Met (-S-CH3), Cys (-SH) are also quite oxidabile in proteins. O2 oxidation of thiol groups (–SH) leads to disulfide formation (–S-S-) and vice versa. Under normal conditions, dehydrogenases have the same effect in organisms, such as the oxidation of Cys to cysteine, for example. The first stage of oxidation is the emergence of alkylthiolate (RS-) in the presence of the hydroxyl anion (HO-) (reaction 44). Thiolate reacts with oxygen and produces a thyil radical (RS) (reaction 45) [85,94]. The second stage is the reactions with thiols and their emerging radicals (reaction 46, 47). As their quantity increases, the probability of them reacting to form a non-radical product also grows (2 RS → RSSR).

RSH + HO- ↔ RS- + H2O
RS- + O2 ↔ RS + O2•-
RSH + O2•- → RS + HO2-
RSH + HO → RS + H2O

H2O2 and R-O-OH are, however, more efficient oxidizing agents. In response to the reaction of protein thiols (PrS) with R-O-OH, atoms of sulfur are simultaneously oxidized (frequently those in Cys), forming corresponding monoxides (thiosulfinates) and, where appropriate, further oxidized products containing 2 sulfoxide groups (disulfoxide), sulfone moiety (dioxide, thiosulfonate), sulfoxide and sulfone miety (sulfoxido sulfone, trioxides), and 2 sulfone groups (disulfonates, tetraoxides) [181]. Reactions with hydro and hydrogen peroxides convert thiol proteins also into sulfenic acids (RSOH), which can be further oxidized to higher oxidation states such as sulfinic (RSO2H) and sulfonic (RSO3H) acids [85,169]. Oxidative modifications of critical amino acids within the functional domain of proteins may also occur by S-glutathionylation. Such alterations may alter the activity of an enzyme if the critical cysteine is located within its catalytic domain or the ability of a transcription factor to bind DNA if it is located within its DNA binding motif [14]. RS then reacts with a glutathionylate anion (GS-) to form a radical mixed disulfide (RSSG•-), which can lose an electron to oxygen to form O2•-, leaving a mixed disulfide (reaction 48) [155,195]. Another route to mixed disulfides is through the two electron oxidation of a thiol to RSOH, which will then react with a thiolate anion to displace HO- (reaction 49). Exposure to NO during pathological conditions can lead to the formation of ONOO-, which can oxidize thiols to either RS or RSOH and lead to protein glutathionylation. It is also possible that S-nitrosylation of PrSH to form PrSNO can lead to protein glutathionylation by the displacement of the NO- by glutathione (reaction 50) [26,55]. A study from Thannickal & Fanburg [168] confirms that cysteine modification involving S-glutathionylation is readily reversed to the active sulfhydryl group by thioltransferases. Met is oxidized to methioninsulfoxide. Further methioninsulfoxide oxidation produces methioninsulfone, which is unexploitable.

PrS + GS- → RS•--SG + O2 → O2•- + PrS-SG
PrSOH + GS- → PrS-SG + HO-
PrSNO + GS- → PrS-SG + NO-

Trp is a very oxylabile compound, especially in an acidic environment. It is easily oxidized by O2 on exposure to light, in a photooxidation reaction catalyzed by riboflavine. Oxidation occurs due to the action of sulfoxides, peroxyacids, H2O2, R-O-OH, but also undergoes autooxidation under γ-irradiation [90]. Autooxidation propagated by peroxy radicals is a chain reaction. The initial phase is the reaction of HO with tryptophan across the double C=C bonds, yielding Trp-OH adducts [93]. These adducts react with oxygen to produce the corresponding peroxy radicals. H reacts with Trp yielding the corresponding Trp-H adducts, while a small amount of the H-atoms react with oxygen yielding HO2•-. The following set of reactions according to Janković & Josimović [90] demonstrates initiation (reaction 51, 52), propagation (forming 2- and 3-adduct peroxy radicals) in reactions 53-56 and termination reactions (57) of Trp autooxidation:

Trp + HO → Trp-OH
Trp-OH + O2 → Trp(OH)OO
Trp + Trp(OH)OO + O2 → Trp(OH)OO-Trp
Trp(OH)OO-Trp+ O2 → Trp(OH)OO-TrpOO
TrpH(OH)OO-TrpOO → RR”O2 + HO
Trp(OH)OO-TrpOO → RR”O2 + HO2•-
2Trp(OH)OO →R2O2 + O2

Similarly, the preferred targets of radicals produced during γ-radiolysis in proteins are other hydrophobic amino acids such as Tyr, Phe, Val and Ile. In biological systems, the presence of HO during radiolysis leads predominantly to extensive protein-protein crosslinkage via tyrosine-tyrosine (dityrosine) bonding and possibly other amino acid cross-links as well [71]. The mechanism of dityrosine formation begins with the generation of a Tyr, radical isomerisation followed by diradical reaction, and finally enolization. The overall rate constant for this process was reported to be 4 x 108 M-1s-1 [69]. Tyr may dissipate by pathways other than those involving intermolecular diradical crosslinking of Tyr. Formation of Tyr oxidation products might involve cyclization, decarboxylation, and further oxidation steps on either the protein or fragments released from the protein [70].

The chelating amino acids in proteins, such as His, are most susceptible to oxidative attack due to their proximity to the radicals formed by binding transition metals [160]. Metal-catalyzed oxidation of histidine generally causes formation of oxo-His or Asp [16,114]. Other amino acyl moieties, especially Lys, Arg, Pro and Thr, incur formation of carbonyl groups (aldehydes and ketones) on the side chains [6,159].

ONOO- causes nitrosylation of Tyr residues and oxidative modification of other amino acid residues including Cys, Trp, Met and Phe [89] but it is a poor inducer of protein carbonyls [171]. The interaction of HOCl with Tyr, Trp, Lys and Met residues leads to the formation of chlorotyrosine, chloramines, aldehydes and methionine sulfoxide [80,97].

Metal-mediated formation of free radicals causes also various modifications to DNA bases, altered calcium and sulfhydryl homeostasis. Whilst Fe, Cu, Cr, V and cobalt Co undergo redox-cycling reactions, for a second group of metals, Hg, Cd and Ni, the primary route for their toxicity is depletion of glutathione and bonding to sulfhydryl groups of proteins. As is thought to bind directly to critical thiols, however, other mechanisms, involving formation of hydrogen peroxide under physiological conditions, have been proposed [178].

Indirect oxidative modification of protein amino acyl side chains occurs through the formation of adducts with products of oxidatively modified lipids, amino acids and sugars. Lipid peroxidation products such as hydroxynonenal, malondialdehyd and acrolein bind covalently to Lys, His and Cys residues, leading to the addition of aldehyde moieties to the protein [149,153,174]. α-β unsaturated alkenals may react with sulfhydryl groups of proteins to form stable covalent thioether adducts also containing carbonyl groups [69]. Products of free amino acid oxidation can also form covalent attachments to proteins [81]. Glutathiolation of Cys residues similarly, Schiff bases, obtained by the reaction of reducing sugars with an ε-amino group of lysyl residues in proteins may, upon Amadori rearrangement, yield ketoamine protein conjugates [71].

4.3. Oxidation of DNA

Reactions that alter DNA and other macromolecules in living systems are induced by oxidizing conditions resulting from normal metabolism or ionizing and ultraviolet radiation. The most basic reaction is one-electron oxidation, the result of which is essentially independent of the process by which it is oxidized [110]. The loss of an electron converts DNA to its radical cation (an electron “hole”), which migrates reversibly through duplex DNA by hopping until it is trapped in an irreversible chemical reaction to form a structurally modified base [95]. The dominant mechanism for radical cation migration in DNA is multi-step hopping [113,172] where charge resides on a single base or on small number of adjacent bases and thermal fluctuations precipitate its movement from one base to another [96]. Superexchange is possible, but less effective for long distances, whereby charge is transported coherently in one step by tunnelling [13,157] from a donor to an acceptor through intervening bridging nucleobases [92]. An incoherent, multi-step, random passage from donor to acceptor consists of short-distance tunnelling intervals linked by base sequences that serve as resting sites for charges [64,115].

With respect to DNA, HO oxidation is most prevalent. HO reacts with DNA by addition across double bonds of DNA bases at or near diffusion-controlled rates with rate constants of 3 to 10 x 109 M-1.s-1, the rate constant of H atom abstraction is 2 x 109 M-1.s-1 [186]. The addition of HO to the C4, C5, and C8 positions of purines generates OH adduct radicals. C4-OH and C5-OH adduct radicals of purines dehydrate and are converted to an oxidizing purine(-H) radical, which may be reduced and protonated to reconstitute the purine [142]. C4-OH adduct radicals possess oxidizing properties, whereas C5-OH and C8-OH adduct radicals are primarily reductants. On the other hand, different mesomeric structures of these radicals may have ambivalent redox states [182]. The rate constants of the dehydration of C4-OH adduct radicals of purines at neutral pH amount to 1.5 x 105 s-1 and 6 x 103 s-1. In contrast to C4-OH adduct radicals, the reaction of O2 with C8-OH adduct radicals of purines is diffusion-controlled [182]. The one-electron oxidation leads to the formation of 8-hydroxypurines (7,8-dihydro-8-oxopurines) in DNA [25]. However, 8-hydroxypurines are also formed in the absence of O2, but to a lesser extent. The oxidation of C8-OH adduct radicals competes with the unimolecular opening of the imidazole ring by scission of the C8-N9 bond at a rate constant of 2 x 105 s-1. The one-electron reduction of the ring-opened radical leads to the formation of 2,6-diamino-4-hydroxy-5-formamidopyrimidine from guanine and 4,6-diamino-5-formamidopyrimidine from adenine [25]. The one-electron reduction of C8-OH adduct radicals without ring-opening may also occur, resulting in the formation of 7-hydro-8-hydroxypurines, hemiorthoamides, which may be converted into formamidopyrimidines. The formation of 8-hydroxypurines is preferred in the presence of O2.

The observation that DNA oxidation occurs predominantly at guanines has been attributed to the fact that this base has the lowest Eox [31]. Similarly, it was found that GG steps are the preferred sites for reaction, with the 5´-G being especially reactive [147]. The relative reactivity of the guanines in a GG step is influenced by the surrounding bases. In particular, the reactivity of the 3´-G is reduced when flanked by pyrimidines, which has also been attributed to electronic effects [33]. The guanine radical cation (guanine•+) is formed by elimination of HO- from the C4-OH adduct radical of guanine and may deprotonate depending on pH to give guanine(-H). Guanine•+ does not hydrate to form the C8-OH adduct radical or go on to form 8-hydroxyguanine (8-oxoguanine, 8-OH-Gua) by oxidation; however, it may react with 2'-deoxyribose in DNA by H abstraction, causing DNA strand breaks [129]. On the other hand, the hydration of guanine•+ in double stranded DNA forms the C8-OH adduct radical, which gives rise to 8-OH-Gua upon oxidation [45]. The C4-OH adduct radical of guanine barely reacts with O2; however, O2 adds to guanine-(-H) with a rate constant of 3 x 109 M-1.s-1. The reaction of guanine(-H) with O2 leads to imidazolone and oxazolone derivatives [34,47].

In the case of adenine, at least two OH adduct radicals are formed: C4-OH and C8-OH. The C4-OH adduct radical of adenine reacts with O2 with a rate constant of 1.0 x 109 M-1.s-1, giving rise to as yet unknown products [182]. 2-hydroxyadenine is also formed from adenine in DNA by a possible mechanism, such as HO attack at the C2-position of adenine, followed by oxidation [136].

Addition of HO across the C5-C6 double bond of pyrimidines leads to C5-OH and C6-OH adduct radicals and H atom abstraction from thymine, resulting in the formation of the allyl radical. The redox properties of adduct radicals differ; C5-OH adduct radicals are reducing while C6-OH adduct radicals are oxidizing [161]. In the absence of O2, the oxidation of C5-OH adduct radicals, followed by addition of HO- (or addition of water followed by deprotonation), leads to cytosine glycol and thymine glycol [25,85]. C5-OH-6-peroxyl radicals are formed by addition of O2 to C5-OH adduct radicals at diffusion-controlled rates. C5-OH-6-peroxyl radicals eliminate O2•-, followed by a reaction with water by HO- addition to yield thymine and cytosine glycols [34,85]. Oxygen reacts with the allyl radical, producing 5-hydroxymethyluracil and 5-formyluracil. Thymine peroxyl radicals are reduced, and then protonated to give hydroxyperoxides [189], which break down to yield thymine glycol, 5- hydroxymethyluracil, 5-formyluracil, and 5-hydroxy-5-methylhydantoin [189].

The products of cytosine oxidation undergo deamination and dehydratation. Cytosine glycol deaminates to give uracil glycol, 5-hydroxycytosine, and 5-hydroxyuracil [25,44,188]. In the absence of O2, C5-OH adduct radicals may be reduced, and subsequently protonated to give 5-hydroxy-6-hydropyrimidines. 5-hydroxy-6-hydrocytosine readily deaminates into 5-hydroxy-6-hydrouracil. Similarly, C6-OH adduct radicals of pyrimidines may lead to the production of 6-hydroxy-5-hydropyrimidines. These products are typical of anoxic conditions because O2 inhibits their formation by reacting with OH adduct radicals. By contrast, pyrimidine glycols and 5-hydroxymethyluracil are formed under both oxic and anoxic conditions. Further reactions of C5-OH-6-peroxyl and C6-OH-5-peroxyl radicals of cytosine result in formation of 4-amino-5-hydroxy-2,6(1H,5H)-pyrimidinedione and 4-amino-6-hydroxy-2,5(1H,6H)-pyrimidinedione, respectively, which may deaminate to give dialuric acid and isodialuric acid. Dialuric acid is oxidized in the presence of O2 to alloxan [44]. C5-OH-6-hydroperoxide gives rise to trans-1-carbamoyl-2-oxo-4,5-dihydroxyimidazolidine as a major product of cytosine [34,188] but as a minor product from DNA [43].

4.4. Oxidation of saccharides

The functional group of carbohydrates is subject to oxidation. Auto-oxidation of carbohydrates is slow in neutral and faster in an acidic environments. D-glucose and D-fructose form unstable hydroperoxides via 1-en-1,2-diols, which break down to form D-arabinonic and formic acids [181].

Monosaccharide autooxidation is a metal-catalysed process. In the presence of transition metals, compounds with α-hydroxyaldehyde, such as glucose [196] and fructose [181] enolize and reduce transition metals and O2 sequentially, forming α-ketoaldehydes as major products in the following reaction. Hydroxyaldehyde → 1-en 1,2-diol (enediol) → enediol radical anion → α-dicarbonyl + O2•-. H2O2, formed by O2•- dismutation, regenerates the catalytic metal oxidation state and produces HO. In the presence of HO, hydroxyaldehyde hydrate is formed, which becomes a hydroxyalkyl radical (or alkyl radical, R). Because glucose is an aldose and fructose is a ketose, it is expected that the extraction of hydrogen from the two kinds of saccharides by HO would take place at different sites of the two kinds of saccharides, thus the alkyl/alkoxyl radicals they produce would have different properties [119]. Hydroxyalkyl radicals in the presence of O2 give rise to dicarbonyls and HO2. Alternativelly, peroxyl radicals and HO2 are formed, giving rise to α-hydroxyacids and further ketoaldehydes [196]. Similar products are also formed by oxidation of H2O2 alone. In the presence of Fe2+ ions, the decay of H2O2 generates free radicals and also oxidizes sugars to the glycosuloses.

The formation of R-O-O from lipid oxidation involves C-H bond cleavage at the C2 carbon of the carbohydrate. The resulting radical reacts with O2 and the resulting peroxyl radical spontaneously decays to the corresponding glyculosis and HO2, which is subject to further disproportionation to H2O2 and O2 (reaction 58). The mechanism of HO2 formation, which causes the transfer of H+ in five members structures of the intermediate structure is not the only like that of carbohydrates by R-O-O. The alternative is to induce the transfer of H+ in the six-member structure of the intermediate. This creates aldonic acid, H2O2, and glyoxal [181].

HO2 → H2O2 + O2

α-dicarbonyl compounds with the original number of carbon atoms are further oxidised and decomposed. The radicals that develop as intermediate products, α-dicarbonyl compounds and α-ketoaldehydes, react further with Lys and Arg in proteins and are involved in early glycosylation reactions. N-substituted 1-amino-1-deoxyfructose (Amadori product) appears as one of the early products of protein glycosylation [197]. Other important processes in these reactions are glycation, glycoxidation and the formation of advanced glycation by-products [191].

Reactions of HO with the sugar moiety of DNA by H abstraction give rise to sugar modification and strand breaks. A unique reaction of the C5'-centered sugar radical is addition to the C8-position of the purine ring of the same nucleoside. This reaction leads to intramolecular cyclization, followed by 8,5-cyclopurine-2'-deoxynucleosides after oxidation [41,42]. Both 5'R- and 5'S-diastereomeres of 8,5'-cyclo-2'-deoxyguanosine (cyclo-dG) and 8,5'-cyclo-2'-deoxyadenosine (cyclo-dA) are formed in DNA [41,42]. These compounds cause concomitant damage to both base and sugar moieties. O2 inhibits their formation by reacting with C5'-centered sugar radical before cyclization is possible [34].


5. Antioxidant metaloenzymes

5.1. Superoxide dismutase (SOD)

Superoxide dismutase (EC belongs to the group of oxido-reductases. SOD contributes significantly to protecting the organism from the toxic effects of O2•- [152]. In living systems, O2•- is capable of reacting with another molecule of O2•- (dismutation) or is also able to react with another radical, such as NO. Formation of HO from O2•- via the metal-catalyzed Haber-Weiss reaction has a reaction rate 10 000 times faster than that of spontaneous dismutation, so SOD provides the first line of defence against ROS [65].

2O2•- + 2 H+ → H2O2 + O2

These enzymes are present in almost all aerobic cells as well as in anaerobic organisms, in all subcellular compartments. The active site of the enzyme contains one or two different atoms of a transition metal in a certain oxidation state [148]. SODs are classified by their metal cofactors into known types: the Cu/ZnSOD and MnSOD, which are localized in different cellular compartments. Cu/Zn SOD is mainly extracellular and cytosolic, while MnSOD is a mitochondrial enzyme. Both types are also present in plants [131]. FeSOD isozymes, often not detected in plants, are usually associated with the chloroplast compartment [5]. The prokaryotic MnSOD, FeSOD and eukaryotic Cu/ZnSOD are dimers, whereas MnSOD of mitochondria are tetramers. NiSOD is the most recent class of SOD, which was discovered in Streptomyces [205] and cyanobacteria [144]. On the basis of amino acid sequence, metal ligand environment, and spectroscopic properties, NiSOD is distinct from other known SODs [28,87]. However, all SODs are known to have very similar catalytic rate constants, pH dependence, and catalytic functions [200]. Therefore, like its counterparts, the catalytic dismutation activity of NiSOD occurs through the oxidative and the reductive half-reactions, which can be described by the following two equations, where SOD-Ni2+ and SOD-Ni3+ represent the reduced and oxidized states of the metal center in the enzyme, respectively.

SOD-Ni2+ + O2- + 2H+ → SOD-Ni3+ + H2O2
SOD-Ni3+ + O2- → SOD-Ni2+ + O2

Formation of H2O2 in the absence of the enzyme, takes place with a dismutation rate k = 105-106 M-1.s-1. This reaction is accelerated 104-times by SOD [65].

5.2. Superoxide reductase (SOR)

(EC is the second enzyme responsible for the detoxification of O2•-, found only in prokaryotic cells, allowing them to survive in the presence of O2. SOR is a non-heme iron enzyme. The active site consists of a mononuclear ferrous ion in an unusual [Fe2+ (N-His)4(S-Cys)] square pyramidal pentacoordination complex [3,203]. The free, solvent-exposed, sixth coordination position is the site of O2•- reduction [104,139]. The reaction of SOR with O2•- may proceed through two reaction intermediates [117,140]. The first, presumably a Fe3+-peroxo species, is formed by the almost diffussion-limited binding of O2•- to the ferrous active site. This intermediate undergoes two sequential protonation reactions, first yielding a second intermediate, possibly a Fe3+-hydroperoxo species, and then the final reaction products, H2O2 and the ferric active site [140].

The active site of SOR binds ferrocyanide, also referred as hexacyanoferrate (II) or K4Fe(CN)6, at its sixth coordination position through a cyano bridge between the iron and the ferrocyanide molecule. The complex has both reduced and oxidized forms of iron in the active site [2]. Molina-Heredia et al. [133] proposed a mechanism for the reaction of SOR-Fe(CN)6 complex with O2•-, forming weakly reactive components in comparison to H2O2, as they cannot be involved in the formation of HO.

SOR-Fe2+-NC-Fe2+(CN)5 + O2•- → SOR-Fe2+-Fe3+(CN)5 + O22-
SOR-Fe2+-NC-Fe3+(CN)5 + HCOO- + 2H+ → SOR-Fe2+-NC-Fe3+(CN)5 + HCOOO- + H2O
SOR-Fe2+-NC-Fe3+(CN)5 → SOR-Fe3+-NC-Fe2+(CN)5

5.3. Indole-2,3-dioxygenase (IDO)

Indoleamine 2,3-dioxygenase (decyclizing) or indole:oxygen 2,3-oxidoreductase (EC uses O2•- as cofactor in the initial step during the degradation of the indole ring of Trp to form kynurenine. This step involves two different enzymes, tryptophan-2,3-dioxygenase (TDO) and indoleamine-2,3-dioxygenase (IDO). IDO is a heme-dependent cytosol enzyme present predominantly in monocytes, macrophages, and microglial cells within the brain parenchyma. In liver hepatocytes, however, TDO is predominantly expressed (for more details see [37]) and as a proenzyme [51,52]. IDO does not show substrate specificity exhibited by TDO, catalyzing the oxygenative ring cleavage of various indoleamine derivatives. Even though it catalyzes the same dioxygenation reaction as classical hepatic TDO, it differs from the latter with respect to molecular size substrate specificit y, cofactor requirements, and immunogenicity [18]. The enzyme scavenges O2•-, which increases only when SOD is inhibited. After conversion into 3-hydroxykynurenine, most of the kynurenine formed via IDO is metabolized into xanthurenic acid, rather than complete oxidation along the glutarate pathway or conversion into NAD [18]. IDO is stimulated by pro-inflammatory cytokines, especially IFN-γ [150,165], virus infection [204] and the administration of bacterial endotoxin [175]. The induction of IDO causes a marked increase in Trp catabolism in the body [164] causing kynurenine production and overall depletion of Trp in the cell. Trp is essential for the growth of bacteria and the growth of bacteria is suppressed by actively depleting Trp within infected cells and surrounding milieu [68,127]. IDO is down-regulated by NO as a consequence of the L-Arg metabolic pathway activation, which is also affected by IFN-γ [170]. Since IDO is expressed both in the periphery and in the central nervous system, it represents a possible link between the immune system and serotogenic pathway, as Trp availability controls the synthesis of serotonin [111]. Macrophages and dendritic cells, in particular plasma-cytoid cells, have been implicated in the IDO-mediated suppression of T-cells [10]. More recently, it has been established that IDO regulates maternal tolerance and possibly more general aspects of T-cell tolerance [128]. The findings of Scott et al. [151] suggest that IDO modulates inflammatory responses, in particular those driven by B-cells.

5.4. Catalase (CAT)

Catalase (H2O2:H2O2 oxidoreductase, EC is a heme-containing enzyme that is present in virtually all aerobic organisms tested to date [4,12]. In the cell, it is localized predominantly in the peroxisomes [15], where it is important in the removal of H2O2 generated by oxidases involved in β-oxidation of fatty acids, respiration, and purine catabolism [9]. CATs from many species are known to be tetramers of 60-65 kDa subunits with each subunit containing 1 Fe-protoheme IX moiety (4 heme groups per tetramer). Each tetrameric molecule of mammalian CATs contains four molecules of tightly bound NADPH, which does not seem to be essential for the enzymatic conversion of H2O2 to H2O and O2, but rather protects CAT against inactivation by H2O2 [98]. CAT has the highest turnover rate among all enzymes, one molecule of CAT can convert approximately 6 million molecules of H2O2 to H2O and O2 per minute [62] and the pH optimum obtained from different sources is 6.8-7.5. The enzyme can function in 2 ways: α and β phases [107,108].

The α-phase works catalytically (reactions 65, 66), breaking H2O2 down into H2O and O2 without the production of free radicals. The reaction takes place in two two-electron reactions. In the first, a H2O2 molecule oxidizes the heme to compound I (CI), removing one oxidation equivalent from the ferric iron, generating the oxoferryl species, and the other from the porphyrin ring, generating a porphyrin cation radical. The second H2O2 then reduces CI to regenerate the resting (ferric) enzyme while releasing H2O and molecular O2.

CAT(Por-Fe3+) + H2O2 → CI(Por+• -Fe4+=O) + H2O
CI(Por+• -Fe4+=O) + H2O2 → CAT(Por-Fe3+) + H2O + O2

At limiting H2O2 concentrations, catalases may undergo a one-electron reduction (reactions 67, 68) to an inactive intermediate, compound II (CII), which can be subsequently converted to another inactive form, compound III (CIII) [138].

CI(Por+• -Fe4+=O) + HA → CII(Por-Fe4+-OH) + A
CII(Por-Fe4+-OH) + H2O2 → CIII(Por-Fe2+=OOH) + H2O

The β phase works peroxidatively (reactions 69-71), by eliminating H2O2 with oxidizing alcohols, formate (RH2) or nitrate as described in Aksoy et al. [4], thereby releasing O2•- and the natural enzyme.

CI + HN3 (or RH2 + H2O2) → CII + N3 (or R + 2H2O)
CII + H2O2 → CIII + H2O
CIII → CAT-Fe3+ + O2•-

The CAT reaction has evolved in at least three phylogenetically unrelated protein types: the monofunctional or “classical” CAT, the bifunctional catalase-peroxidase (KatG; EC, and the non-heme, Mn-containing catalase [138]. Generally, rate constants for the formation of CI from peroxidases and catalases were calculated to be in the range of 106 to 108 M-1 s-1 [48]. A distal His-Asn pair has been shown to be essential for CI formation in classical CATs, while a distal His-Arg has the same funtion in peroxidases [48,54]. The main diference in the enzymatic mechanism between CAT and peroxidases is CI reduction. In a catalase cycle, a second H2O2 molecule is used as a reducing agent for CI. This two-electron reduction completes the cycle forming ferric-CAT and O2 (for details see [207]). With most substrates in a peroxidase cycle, CI is reduced back the the ferric enzyme in two consecutive one-electron steps via CII. KatGs can be viewed as a molecular fossil revealing the common phylogeny of catalytic and per-oxidative activity during evolution [206]. It has been proposed that KatG is responsible for the catalytic oxidation of H2O2 in a two-electron oxidation step with both oxygen atoms being derived from the same H2O2 molecule. This non-scrambling mechanism is independent of pH and is not affected by manipulation of highly-conserved and important catalatic residues. Principally, there are two possible mechanisms for the formation of O2 following this retention mechanism: an ionic mechanism, via initial proton abstraction with the help of an acid–base catalyst followed by a hydride-ion removal from H2O2 and release of O2; and hydrogen atom transfer from H2O2 to the ferryl species to yield a radical intermediate [185]. Until now, the complete gene sequences of KatGs were characterised only from prokaryotes (both from archaebacteria and eubacteria) although several reports describe the presence of KatGs in lower eukaryotes [56]. It was shown phylogenetically that the closest neighbours of KatGs are eukaryotic ascorbate peroxidases and yeast cytochrome c peroxidase [192]. So far, KatGs are the only peroxidases known with both catalase activity comparable with catalases and typical peroxidase activity with broad specificity.

The variable response of CAT activity has been observed under metal stress [65], while all ions of heavy metals are non-competitive inhibitors of CAT. Cyanides are strong inhibitors of CAT as they form a strong bond with the heme of CAT and stop its catalytic activity [184]. Some studies have shown that CAT is effective in the degradation of H2O2 present only in mmol.l-1, while glutathione peroxidase is effective in peroxide degradation at concentrations lower than 100 µmol.l-1 [39].

5.5. Glutathione peroxidase (GPx)

Glutathione peroxidise (EC has eight known izoenzymes that, in active positions, may contain co-factors, such as heme and residues of cysteine or selenocysteine. GPx1-4 and GPx 6 are selenoenzymes, which contain the non-metal selenium [27]. Selenocysteine participates directly in the transfer of electrons to a peroxide substrate, thereby oxidizing it. However, the pathophysiological role of these isoenzymes in antioxidant defence is of substantial importance [100,101,180]. CAT is found in many types of cells and scavenges H2O2 as its sole substrate, GPx scavenges various peroxides. The expression of CAT in most cells is lower than that of GPx, with the exception of hepatocytes and erythrocytes. The Km value of CAT for H2O2 is higher than that of GPx, implying the primary importance of GPx in most tissues [8].

Like all peroxidases, they mediate the one-electron oxidation of organic compounds (reduced glutathione, GSH) with a concomitant reduction of H2O2 (for more detailed mechanisms of peroxidase action see 4.1.4 and 5.4). The activity of GPx is affected by the presence of another important antioxidant enzyme, glutathione reductase, which continuously recycles the oxidised glutathione to the reduced state. Lawrence et al. [109] described that non-selenium dependent GPx activity contributes to glutathione-S-transferase B (G-S-T) activity in mechanisms analogous to the G-S-T mechanism. Thus, an enzyme bound to GSH may attack the electrophilic oxygen of the peroxide and a second molecule of GSH may react in a non-enzymatic fashion similar to the reaction with organic nitrates, or by another enzyme catalyzed step to yield the glutathione disulphide (GSSG). Non-selenium dependent GPx also has the ability to reduce phospholipid hydroperoxides, without G-S-T activity [53]. Kinetic analysis of GPx activity indicated a tert-uni ping-pong mechanism similar to that described for other GSH peroxidases [176].

2GSH + H2O2 → GSSG + 2H2O

Kinetic behaviour of the overall reaction is discussed in detail in Ng et al. [137].

GPxr + H2O2 + H+ → GPxo + H2O
GPxo + GSH → [GS-SG] + H2O
[GS-SG] + GSH → GPxr + GSSG + H+

Hall et al. [74] showed that an epididymis-specific, secretory GPx has very little activity towards H2O2 or organic hydroperoxides. Instead, it binds to lipid peroxides.

Virtually all known peroxidases are inactivated by H2O2 and other hydroperoxides at relatively high concentrations [83]. This substrate inactivation leads to modification of the heme prosthetic group and the formation of a verdohemoprotein as the final product. The existence of CIII as the peroxy iron prophyrin free radical resonance form can facilitate the transfer of electrons from the ferrous state to an extra H2O2 molecule, thereby generating HO. This highly reactive species has the propensity to attack the heme porphyrin ring and lead to irreversible inactivation [60].

5.6. Heme oxygenase (HO)

An iron-containing decyclizing oxygenase (EC can be legitimately considered a part of the phase 2 response [40]. HO catalyzes the first, rate-limiting step of heme degradation. HO cleaves the α-meso carbon bridge of b-type heme molecules via oxidation to yield equimolar quantities of biliverdin IXa, CO and free iron. Biliverdin is subsequently converted to bilirubin via the action of biliverdin reductase, while the free iron is promptly sequestered into ferritin. To date, three isoforms (HO-1, HO-2 and HO-3) have been identified. Under physiological conditions, HO activity is highest in the spleen where senescent erythrocytes are sequestered and destroyed [143]. HO-1, can be induced by a variety of non-heme products including ultraviolet irradiation, endotoxins, heavy metals, and oxidants as well as H2O2 [121,183]. The production of bilirubin/biliverdin and carbon monoxide from heme catabolism is capable of exerting protection against toxic compounds in the cell. Indeed, in a variety of cells and tissues, inducible oxidative stress represents part of an adaptive cellular response to inflammation.


6. Conclusion

Oxidative processes are essential to life, particularly for obtaining the energy needed for various metabolic processes, but they also serve as a source of ROS. Oxidation and reduction processes are inseparable. Given that transit metals readily accept or give away electrons, they play an important role in the oxidoreduction processes and are constituents of various proteins and enzymes. In fact metalloenzymes participate significantly in the antioxidant protection of the body as phase I and II antioxidants. It is important to understand and study the antioxidant defence system of the organism so that one can use this knowledge to prevent and treat diseases in which it has been proven to participate.


This work was supported by Slovak Grant Agency for Science VEGA 1/1236/12.


  1. 1. AboA.PickE.HallA.TottyN.TeahanC. G.SegalA. W.Activation of the NADPH oxidase involves the small GTP-binding protein 21rac1Nature 1991
  2. 2. AckerH.DufauE.HuberJ.SylvesterD.Indications to an NADPH oxidase as a possible pO2 sensor in the rat carotid body. FEBS Letters 1989
  3. 3. AdamV.RoyantA.NivièreV.Molina-HerediaF. P.BourgeoisD.Structure of superoxide reductase bound to ferrocyanide and active site expansion upon X-ray-induced photo-reduction. Structure 200412917291740
  4. 4. AksoyY.BalkM.ÖğüşH.ÖzerN.The Mechanism of Inhibition of Human Erythrocyte Catalase by Azide. Turk. Journal of Biology 20042865
  5. 5. AlscherR. G.ErtukN.HeathL. S.Role of superoxide dismutases (SODs) in controlling oxidative stress in plants. Journal of Experimental Botany 20025337213311341
  6. 6. AmiciA.LevineR. L.TsaiL.StadtmanE. R.Conversion of amino acid residues in proteins and amino acid homopolymers to carbonyl derivatives by metal-catalyzed oxidation reactions. Journal of Biological Chemistry 1989264633413346
  7. 7. Andrews PC & Krinsky NI.The reductive cleavage of myeloperoxidase in half, producing enzymically active hemi-myeloperoxidase. Journal of Biological Chemistry 1981256942114218
  8. 8. AsahiM.FujiiJ.SuzukiK.SeoH. G.KuzuyaT.HoriM.TadaM.FujiiS.TaniguchiN.Inactivation of glutathione peroxidase by nitric oxide. Implication for cytotoxicity. Journal of Biological Chemistry 1995270362103521039
  9. 9. Azevedo RA, Alas RM, Smith RJ & Lea PA.Response of antioxidant enzymes to transfer from elevatde carbon dioxide to air and ozone fumigation, in leaves nad roots of wild-type and catalase-deficient mutant of barley. Physiologia Plantarum 1991104280
  10. 10. BabanB.HansenA. M.ChandlerP. R.ManlapatA.BingamanA.KahlerD. J.MunnD. H.MellorA. L. A.minorpopulation.ofsplenic.dendriticcells.expressingC. D.mediatesI. D.O-dependentT.cellsuppression.viatype. I. I. F. N.signalingfollowing. B.ligationInternational Immunology 2005177909919
  11. 11. Babior BM. NADPH oxidase: an update. Blood199993514641476
  12. 12. BMBabiorCurnutte. J. T.OkamuraN.The Respiratory Burst Oxidase of the Human Neutrophil Radicals and Tissue Injury. In: B. Halliwell (ed.), Oxygen Radicals and Tissue Injury, Upjohn Company, Maryland, 43481988
  13. 13. RNBarnettCleveland. C. L.JoyA.LandmanU.SchusterG. B.Charge migration in DNA: Ion-gated transport. Science 20012945542567571
  14. 14. BarrettE. G.JohnstonC.OberdörsterG.FinkelsteinJ. N.Silica-induced chemokine expression in alveolar type II cells is mediated by THF-α-induced oxidant stress. American Journal of Physiology 1999Pt 1) L979L988.
  15. 15. BergJ. M.TymoczkoJ. L.StryerL.Biochemistry5th.EditionW. H.FreemanCompanyNew.Yorkpp.502002
  16. 16. Berlett BS, Levine RL & Stadtman ER.Comparison of the effect of ozone on the modification of amino acid residues in glutamine synthetase and bovine serum albumin. Journal of Biological Chemistry 1996271841774182
  17. 17. Berry CE & Hare JM.Xanthine oxidoreductase and cardiovascular disease: molecular mechanisms and pathophysiological implications. Journal of Physiology 2004Pt 3) 589-606.
  18. 18. BertazzoA.RagazziE.BiasioloM.CostaC. V.AllegriG.Enzyme activities involved in tryptophan metabolism along the kynurenine pathway in rabbits. Biochimica et Biophysica Acta 200115273167175
  19. 19. BhattachrjeeS.Reactive oxygen species and oxidative burst: roles in stress, senescence and signal transduction in plant. Current Science 2005891113
  20. 20. Bielski BH.Studies of hypervalent iron. Free radical research communications 1991Pt 2) 469-477.
  21. 21. BogdanC.RöllinghoffM.DiefenbachA.Reactive oxygen and reactive nitrogen intermadiates in innate and specific immunity. Current Opinion in Immunology 20001216476
  22. 22. BolwellG. P.WoftastekP.Mechanism for the generation of reactive oxygne species in plant defense-broad perspective. Physiological and Molecular Plant Pathology 199751347
  23. 23. BonizziG.PietteJ.MervilleM. P.BoursV.Cell type-specific role for reactive oxygen species in nuclear factor-kappaB activation by interleukin-1. Biochemical Pharmacology. 2000591711
  24. 24. BozziY.BorrelliE.Dopamine in neurotoxicity and neuroprotection: what do D2 receptors have to do with it? Trends in Neurosciences 2006293167174
  25. 25. Breen AP & Murphy JA.Reactions of oxyl radicals with DNA. Free Radical Biology and Medicine 199518610331077
  26. 26. BrennanJ. P.WaitR.BegumS.BellJ. R.MJDunnEatonP.Detectionmappingof.widespreadintermolecular.proteindisulfide.formationduring.cardiacoxidative.stressusing.proteomicswith.diagonalelectrophoresis.Journal of Biological Chemistry 2004279404135241360
  27. 27. Brigelius-FlohéR.WinglerR.MüllerC.Estimation of individual types of glutathione peroxidases. Methods in Enzymology 2002347101
  28. 28. Bryngelson PA, Arobo SE, Pinkham JL, Cabelli DE & Maroney MJ. Expression, reconstitution, and mutation of recombinant Streptomyces coelicolor NiSOD.Journal of the American Chemical Society 20041262460461
  29. 29. BuchertF.ForreiterC.Singlet oxygen inhibits ATPase and proton translocation activity of the thylakoid ATP synthase CF1CFo. FEBS Letters 20105841147152
  30. 30. BurnerU.FurmüllerP. G.KettleA.KoppenolW. H.ObingerC.Mechanism of reaction of myeloperoxidase with nitrite. Journal of Biological Chemistry 2000275272059720601
  31. 31. CadetJ.DoukiT.RavanatJ. L.Oxidatively generated damage to the guanine moiety of DNA: Mechanistic aspects and formation in cell. Accounts of Chemical Research 200841810751083
  32. 32. CarrA.Mc CallM. R.FreiB.Oxidationof. L. D. L.bymyeloperoxidase.reactivenitrogen.speciesreaction.pathwaysantioxidantprotection.Arteriosclerosis, Thrombosis, and Vascular Biology 200020717161723
  33. 33. ČekovićŽ.Reactions of sigma-carbon radicals generated by 15hydrogen transfer to alkoxyl radicals. Tetrahedron 2003
  34. 34. ChandraJ.SamaliA.OrreniusS.Triggering and modulation of apoptosis by oxidative stress. Free Radical Biology and Medicine 2000
  35. 35. ChoudhuryS. B.LeeJ. W.DavidsonG.YimY. I.BoseK.SharmaM. L.KangS. O.CabelliD. E.MJMaroneyExamination of the nickel site structure and reaction mechanism in Streptomyces seoulensis superoxide dismutase. Biochemistry 1999381237443752
  36. 36. ClevelandC. L.RNBarnettBongiorno. A.JosephJ.LiuC.SchusterG. B.LandmanU.Steric Effects on Water Accessability Control Sequence-Selectivity of Radical Cation Reactions in DNA. Journal of the American Chemical Society 20071292784088409
  37. 37. MSCookeEvans.MDDizdarogluM.LunecJ.OxidativeD. N. A.damagemechanisms.mutationdiseaseF. A. S. E. B.Journal2003171011951214
  38. 38. CroftsA. R.HollandJ. T.VictoriaD.KollingD. R.DikanovS. A.GilbrethR.LheeS.KurasR.KurasM. G.TheQ-cycle.reviewedHow.welldoes. a.monomericmechanism.ofthe.bc(complexaccount.forthe.functionof.dimericcomplex?.Biochimicaet.BiophysicaActa.2008
  39. 39. DaleW. E.DangY.BrownO. R.Tryptophan metabolism through the kynurenine patway in rat brain and liver slices. Free Radical Biology and Medicine 2000292191198
  40. 40. Davies MJ.Reactive species formed on proteins exposed to singlet oxygen. Photochemical and Photobiological Sciences 2004311725
  41. 41. DemelashA.KarlssonJ. O.NilssonM.BjörkmanU.Selenium has a protective role in caspase-3-dependent apoptosis induced by H2O2 in primary cultured pig thyrocytes. European Journal of Endocrinology 20041506841849
  42. 42. Dinkova-KostovaA. T.MAMassiahBozak. R. E.HicksR. J.TalalayP.Potency of Michael reaction acceptors as inducers of enzymes that protect against carcinogenesis depends on their reactivity with sulfhydryl groups. Proceedings of the National Academy of Sciences of the United States of America 200198634043409
  43. 43. DirksenM. L.BlakelyW. F.HolwittE.DizdarogluM.Effectof. D. N. A.conformationon.thehydroxyl.radical-inducedformation.of8,5’-cyclopurine-2’-deoxyribonucleoside.residuesin. D. N. A.International Journal of Radiation Biology 1988542195204
  44. 44. DizdarogluM.Free-radical induced formation of an 8,5’-cyclo-2’-deoxyguanosine moiety in deoxyribonucleic acid. Biochemical Journal 19862381247254
  45. 45. DizdarogluM.BaucheC.RodriguezH.LavalJ.Novel substrates of Escherichia coli nth protein and its kinetics for excision of modified bases from DNA damaged by free radicals. Biochemistry 2000391855865592
  46. 46. DizdarogluM.LavalJ.BoiteuxS.Substrate specificity of the Escherichia coli endonuclease III: excision of thymine- and cytosine-derived lessions in DNA produced by radiation-generated free radicals. Biochemistry 199332451210512111
  47. 47. DoetschP. W.ZasatawnyT. H.MartinA. M.DizdarogluM.Monomeric base damage products from adenine, guanine, and thymine induced by exposure of DNA to ultraviolet radiation. Biochemistry 1995343737742
  48. 48. DrögeW.2002Free radicals in the physiological control of cell function. Physiological Reviews 8214795
  49. 49. DuarteV.GasparuttoD.JaquinodM.CadetJ.2000In vitro DNA synthesis opposite oxazolone repair of this DNA damage using modified oligonucleotides. Nucleic Acids Research 28715551563
  50. 50. DunfordH. B.Horseradishperoxidase. I.Ligandbinding.redoxpotentials.formationof.itscompounds.someof.theirreaction.inHeme.Peroxidases-VWiley.NewC. H.Yorkpp.58911999
  51. 51. Dunford HB.How does enzyme work? Effect of electron circuits on transition state acid dissociation constant. Journal of Biological Inorganic Chemistry 200168819822
  52. 52. Elstner EF.Mechanism of oxygen activation in different compartments, In: E.J. Pell, K.L. Steffen (ed.). Active Oxygen/Oxidative Stress and Plant Metabolism, American Society of Plant Physiologists, Roseville, 13251991
  53. 53. FallarinoF.Asselin-PaturelC.VaccaC.BianchiR.GizziS.FiorettiM. C.TrinchieriG.GrohmannU.PuccettiP.Murine plasmacytoid dendritic cells initiate the immunosuppressive pathway of tryptophan catabolism in response to CD200 receptor engagement. Journal of Immunology 2004173637483754
  54. 54. FallarinoF.VaccaC.OrabonaC.BelladonnaM. L.BianchiR.MarshallB.KeskinD. B.MellorA. L.FiorettiM. C.GrohmannU.PuccettiP.Fuctional expression of indoleamine 2,3-dioxygenase by murine CD8 alpha(+) dendritic cells. International Immunology 20021416568
  55. 55. FisherA. B.DodiaC.ManevichY.ChenJ. W.FeinsteinS. I.Phospholipid hydroperoxides are substrates for non-selenium glutathione peroxidase. Journal of Biological Chemistry 1999274302132621334
  56. 56. FitaI.RossmannM. G.The active center of catalase. Journal of Molecular Biology 198518512137
  57. 57. Foster MW & Stamler JS.New insights into protein S-nitrosylation. Mitochondria as a model system. Journal of Biological Chemistry 2004279242589125897
  58. 58. Fraaije MW, Roubroeks HP, Hagen WR & Van Berkel WJ.Purification and characterization of an intracellular catalase-peroxidase from Penicillium simplicissimum. European Journal of Biochemistry 1996
  59. 59. Freeman BA & Crapo JD. Biology of disease: free radicals and tissue injury.Lab. Invest. 1989475412426
  60. 60. Friedl HP, Till GO, Ryan US & Ward PA.Mediator-induced activation of xanthine oxidase in endothelial cells. FASEB Journal 198931325122518
  61. 61. GalbuseraC.OrthP.FedidaD.SpectorT.Superoxide radical production by allopurinol and xanthine oxidase. Biochemical Pharmacology 2006711217471752
  62. 62. García-ArellanoH.Buenrostro-GonzalezE.Vazquez-DuhaltR.Biocatalytic transformation of petroporphyrins by chemical modified cytochrome C. Biotechnology and Bioengineering 2004857790798
  63. 63. GardnerP. R.RaineriI.EpsteinL. B.WhiteC. W.Superoxide radical and iron modulate aconitase activity in mammalian cells. Journal of Biological Chemistry 1995270221339913405
  64. 64. GargN.ManchandaG. R. O. S.generationin.plantsboon.orbane?.PlantBiosystems.20091438
  65. 65. GhafourifarP.CadenasE.Mitochondrial nitric oxide synthase. Trends in Pharmacological Sciences 2005264190195
  66. 66. GieseB.AmaudrutJ.KohlerA. K.SpormannM.WesselyS.Direct observation of hole transfer through DNA by hopping between adenine bases and by tunnelling. Nature 20014126844318320
  67. 67. GillS. S.TutejaN.Reactive oxygen species and antioxidant machinery in abiotic stress tolerance in crop plants. Plant Physiology and Biochemistry 20104812909930
  68. 68. GriendlingK.SorescuD.Ushio-FukaiM. N. A. D. P. H.oxidaserole.incardiovascular.biologydiseaseCirculation Research 2000865494501
  69. 69. GrimaG.BenzB.ParpuraV.CuénodM.DoK. Q.Dopamine-induced oxidative stress in neurons with glutathione deficit: implication for schizophrenia. Schizophrenia Research 2003623213224
  70. 70. GrohmannU.FallarinoF.PuccettiP.ToleranceD.Cstryptophanmuch.adoabout. I. D. O.Trends in Immunology 2003245242248
  71. 71. GuiliviC.DaviesK. J. A.Dityrosinea.markerfor.oxidativelymodified.proteinsselectiveproteolysis.Methods in Enzymology 1994233363
  72. 72. GuiliviC.DaviesK. J. A.Mechanism of the formation and proteolytic release of H2O-induced dityrosine and tyrosine oxidation products in hemoglobin and red cells. Journal of Biological Chemistry 2001276262412924136
  73. 73. GuiliviC.TraasethN. J.DaviesK. J. A.Tyrosineoxidation.productsanalysis.biologicalrelevance.Amino Acids 2003
  74. 74. GutteridgeJ. M.1993Free radicals in disease processes: a compilation of cause and consequences [Review]. Free radical research communications 193141158
  75. 75. Gutteridge JM.Lipid Peroxidation andAntioxidants as Biomarkers of Tissue Damage. Clinical Chemistry 1995Pt 2) 1819-1828.
  76. 76. HallL.WilliamsK.PerryA. C.FrayneJ.JuryJ. A.The majority of human glutathione peroxidase type 5 (GPX5) transcripts are incorrectly spliced: implications for the role of GPX5 in the male reproductive tract. Biochemical Journal 1998Pt 1) 5-9.
  77. 77. HalliwellB.GutteridgeJ. M. C.Free Radicals in Biology and Medicine. New York: Oxford University Press, 22851989
  78. 78. Hampton MB, Kettle AJ & Winterbourn CC.Inside the neutrophil phagosome: oxidants, myeloperoxidase, and bacterial killing. Blood 199892930073017
  79. 79. HanD.AntunesF.CanaliR.RettoriD.CadenasE.Voltage-dependent anion channels control the release of the superoxide anion from mitochondria to cytosol. Journal of Biological Chemistry 2003278855575563
  80. 80. HanD.WilliamsE.CadenasE.Mitochondrial respiratory chain-dependent generation of superoxide anion and its release into the intermembrane space. Biochemical Journal 2001Pt 2) 411-416.
  81. 81. HatzS.LambertJ. D. C.OgilbyP. R.Measuring the lifetime of singlet oxygen in a single cell: addressing the issue of cell viability. Photochemical and Photobiological Sciences 200761011061116
  82. 82. HazellL. J.van denBerg. J. J. M.StockerR.Oxidation of low-density lipoprotein by hypochlorite causes aggregation that is mediated by modification of lysine residues rather than lipid oxidation. Biochemical Journal 19943021297304
  83. 83. HazenS. L.GautJ. P.HsuF. F.CrowleyJ. R.d’AvignonA.Heinecke-HydroxyphenylacetaldehydeJ. W. p.themajor.productof.L-tyrosineoxidation.bythe.myeloperoxidase-HO2-chloride.systemof.phygocytescovalently.modifiesepsilon-amino.groupsof.proteinlysine.residueJournal of Biological Chemistry 1997272271699016998
  84. 84. HilleR.Molybdenum-containinghydroxylases.Archives of Biochemistry and Biophysics 20054331107116
  85. 85. HinerA. N.Hernández-RuizJ.Rodríguez-LópezJ.ArnaoM. B.VarónR.CánovasF. G.AcostaM.The inactivation of horseradish peroxidase isoenzyme A2 by hydrogen peroxide: an example of partial resistance due to the formation of a stable enzyme intermediate. Journal of Biological Inorganic Chemistry 2001
  86. 86. HirstJ.CarrollJ.FearnleyI. M.ShannonR. J.WalkerJ. E.The nuclear encoded subunits of complex I from bovine heart mitochondria. Biochimica et Biophysica Acta 200316043135150
  87. 87. HurdT. R.FilipovskaA.CostaN. J.DahmC. C.MurphyM. P.Disulphide formation on mitochondrial protein thiols. Biochemical Society Transactions 200533613901393
  88. 88. Ignarro LJ.Heam-dependent activation of guanylate cyclase and cyclic GMP formation by endogenous nitric oxide: a unique transduction mechanism for transcellular signaling. Pharmacology and Toxicology 199067117
  89. 89. B.Peroxynitrite-mediated oxidative protein modifications. FEBS Letters 19953643279282
  90. 90. Janković IA & Josimović LR.Autooxidation of tryptophan in aqueous solution. Journal of the Serbian Chemical Society 2001669571580
  91. 91. Jones SA, O’Donnell VB, Wood JD, Broughton JP, Hughes EJ & Jones OT.Expression of phagocyte NADPH oxidase components in human endothelial cells. American Journal of Physiology 1996Pt 2) H1626H1634.
  92. 92. JortnerJ.BixonM.AAVoityukRoschN.Superexchange mediated charge hopping in DNA. Journal of Physical Chemistry A 20021063375997606
  93. 93. Josimović LR, Janković IA & Jovanović SV.Radiation induced decomposition of tryptophan in the presence of oxygen. Radiation Physics and Chemistry 199341835
  94. 94. Jung CH & Thomas JA.S-glutathiolated hepatocyte proteins and insulin disulfides as substrates for reduction by glutaredoxin, thioredoxin, protein disulfide isomerase, and glutathione. Archives of Biochemistry and Biophysics 199633516172
  95. 95. KanvahS.JosephJ.SchusterG. B.RNBarnettCleveland. C. H. L.LandmanU.Oxidationof. D. N. A.Damageto.NucleobasesAccount of Chemical reserch 2010432280287
  96. 96. KawaiK.KoderaH.OsakadaY.MajimaT.Sequence-independent and rapid long-range charge transfer through DNA. Nature Chemistry 200912156159
  97. 97. Kettle AJ.Neutrofils convert tyrosyl residues in albumin to chlorotyrosine. FEBS Letters 19963791103106
  98. 98. KirkmanH. N.RolfoM.FerrarisA. M.GaetaniG. F.Mechanisms of protection of catalase by NADPH. Kinetics and stoichiometry. Journal of Biological Chemistry 1999274201390813914
  99. 99. KlotzL. O.KrönckeK. D.SiesH.Singlet oxygen-induced signaling effects inmammalian cells, Photochemical and Photobiological Sciences 2003228894
  100. 100. KocanL.FirmentJ.SimonováJ.VaskováJ.GuzyJ.Selenium supplementation in patients with severe acute pancreatitis. Rozhledy v Chirurgii 2010898518521
  101. 101. KocanL.VaškováJ.VaškoL.HokováH.MajerníkM.KrištofováB.ŠimonováJ.FirmentJ.Severe course of Still΄s disease with multiple organ failure with predominant liver failure. Anesteziologie a Intenzivní Medicína 2011226337342
  102. 102. Krieger-LiszkayA.FufezanC.TrebstA.Singlet oxygen production in photosystem II and related protection mechanism. Photosynthesis Research 2008
  103. 103. KudinA. P.Bimpong-ButaN. Y.VielhaberS.CEElgerKunzW. S.Characterization of superoxide-producing sites in isolated brain mitochondria. Journal of Biological Chemistry 2004279641274135
  104. 104. Kurtz DM Jr.Microbial detoxification of superoxide: the non-heme iron reductive paradigm for combating oxidative stress. Accounts of Chemical Research 20043711902908
  105. 105. LahiriS.AckerH.Redox-dependent binding of CO to heme protein controls O2sensitive chemoreceptor discharge of the rat carotid body. Respiratory Physiology 1999
  106. 106. LanderH. M.HajjarD. P.HempsteadB. L.MirzaU. A.ChaitB. T.CampbellS.QuilliamL. A. A.molecularredox.switchon.p21(rasStructural basis for the nitric oxide-21ras) interaction. Journal of Biological Chemistry 1997
  107. 107. Lardinois OM & Rouxhet PG.Peroxidatic degradation of azide by catalase and irreversible enzyme inactivation. Biochimica et Biophysica Acta 199612982180190
  108. 108. Lardinois OM, Mestdagh MM & Rouxhet PG.Reversible inhibition and irreversible inactivation of catalase in presence of hydrogen peroxide. Biochimica et Biophysica Acta 199612952222238
  109. 109. Lawrence RA, Parkhill LK & Burk RF.Hepatic cytosolic non selenium-dependent glutathione peroxidase activity: its nature and the effect of selenium deficiency. Journal of Nutrition 19781086981987
  110. 110. LeeY. A.DurandinA.DedonP. C.GeacintovN. E.SharirovichV.Oxidation of guanine in G, GG, and GGG sequence contexts by aromatic pyrenyl radical cations and carbonate radical anions: Relationship between kinetics and distribution of alkali-labile lesions. Journal of Physical Chemistry B 2008112618341844
  111. 111. LestageJ.VerrierD.PalinK.DantzerR.The enzyme indoleamine 2,3-dioxygenase is induced in the mouse brain in response to peripheral administration of lipopolysaccharide and superantigen. Brain, Behavior, and Immunity 2002165596601
  112. 112. LeusenJ. KleinA.HilariusP. M.AhlinA.PalmbladJ.SmithC. I.DiekmannD.HallA.VerhoevenA. J.RoosD.Disturbed interaction of 21with mutated p67-phox causes chronic granulomatous disease. Journal of Experimental Medicine 1996
  113. 113. LewisF. D.ZhuH.DaublainP.SigmundK.FiebigT.RaytchevM.WangQ.ShafirovichV.Gettingto.guaninemechanism.Dynamicsof.chargeseparation.chargerecombination.inD. N. A.revisitedPhotochemical and Photobiological Sciences 200875534539
  114. 114. Lewisch SA & Levine RL.Determination of 2-oxohistidine by amino acid analysis. Analytical Biochemistry 19952312440446
  115. 115. LiX. Q.ZhangH.YanY. A.superexchange-mediatedsequential.hoppingtheory.forcharge.transferin. D. N. A.Journal of Physical Chemistry A 20011059563
  116. 116. LiuS. S.MitochondrialQ.cycle-derivedsuperoxide.chemiosmoticbioenergetics.Annals of the New York Academy of Sciences 2010120184
  117. 117. LombardM.Houée-LevinC.TouatiD.FontecaveM.NivièreV.Superoxide reductase from Desulfoarchus baarsii: reaction mechanism and role of glutamate 47 and lysine 48 in catalysis. Biochemistry 2001401650325040
  118. 118. LosM.SchenkH.HexelK.BaeuerleP. A.DrögeW.Schulze-OsthoffK. I. L.geneexpression.K-kappaN.activationB.throughC. D.requiresreactive.oxygenproduction.by5-lipoxygenase. E. M. B.EMBO Journal 1995141537313740
  119. 119. LuoG-m.QiD-h.ZhengY-g.MuY.YanG-l.YangT-s.Shen-CJ.studiesE. S. R.onreaction.ofsacharide.thefree.radicalsgenerated.fromthe.xanthineoxidase/hypoxanthine.systemcontaining.ironF. E. B.FEBS Letters 2001
  120. 120. MacMicking. J.XieQ. W.NathanC.Nitric oxide and macrophage function. Annual Review of Immunology 199715323
  121. 121. MaeshimaH.SatoM.IshikawaK.KatagataY.YoshidaT.Participation of altered upstream stimulatory factor in the induction of rat heme oxygenase-1 by cadmium. Nucleic Acids Research 1996241529592956
  122. 122. Mailloux RJ & Harper ME.Uncoupling proteins and the control of mitochondrial reactive oxygen species production. Free Radical Biology and Medicine 201151611061115
  123. 123. MannickJ. B.SchonhoffC.PapetaN.GhafourifarP.SziborM.FangK.GastonB. S.S-Nitrosylation of mitochondrial caspases. Journal of Cell Biology 2001154611111116
  124. 124. MaragosW. F.YoungK. L.AltmanC. S.PocernichC. B.DrakeJ.ButterfieldD. A.SeifI.HolschneiderD. P.ChenK.ShihJ. C.Striatal damage and oxidative stress induced by the mitochondrial toxin malonate are reduced in clorgyline-treated rats and MAO-A deficient mice. Neurochemical Research 2004294741746
  125. 125. Marnett LJ. Lipid peroxidation-DNA damage by malondialdehyde. Mutat. Res.1999
  126. 126. Mc IntyreM.BohrD. F.DominiczakA. F.Endothelial function in hypertension: the role of superoxide anion. Hypertension 1999Pt 1) 539-545.
  127. 127. Mellor AL & Munn DH. IDO expression by dendrtic cells: tolerance and tryptophan catabolism.Nature Reviews Immunology 2004410762774
  128. 128. MellorA. L.MunnD.ChandlerP.KeskinD.JohnsonT.MarshallB.JhaverK.BabanB.Tryptophan catabolism and T cell responses. Advances in Experimental Medicine and Biology 200352727
  129. 129. MelvinT.BotchwayS.ParkerA. W.O’NeillP.Induction of strand breaks in single-stranded polyribonucleotides and DNA by photosensitisation: one electron oxidised nucleobase radicals as precursors. Journal of the American Chemical Society 199611810031
  130. 130. Miller RT. NOx and R-NOx: effects on drug metabolism.Current Drug Metabolism 200456535542
  131. 131. MittlerR.Oxidativestress.antioxidantsstresstolerance.Trends in Plant Science 200279405410
  132. 132. MiyazakiI.AsanumaM.Dopaminergic Neuron-Specific Oxidative Stress Caused by Dopamine Itself. Acta Med. Okayama 2008623141150
  133. 133. Molina-HerediaF. P.Houveé-LevinC.BerthomieuC.TouatiD.TremeyV. F.NivièreV.Detoxification of superoxide without production of H2O2: antioxidant acitivty of superoxide reductase complexed with ferrocyanide. Proceedings of the National Academy of Sciences of the United States of America 2002103401475014755
  134. 134. MollerI. M.JensenP. E.HanssonA.Oxidative modifications to cellular components in plants. Annual Review of Plant Physiology 200758459
  135. 135. MullerF. L.LiuY.Van RemmenH.ComplexI. I. I.releasessuperoxide.toboth.sidesof.theinner.mitochondrialmembrane.Journal of Biological Chemistry 2004279474906449073
  136. 136. NackerdienZ.KasprzakK. S.RaoG.HalliwellB.DizdarogluM.NickelI. I.cobaltI.I)-dependentdamage.byhydrogen.peroxidethe. D. N. A.basesin.isolatedhuman.chromatinCancer Research 1991512158375842
  137. 137. Ng CF, Schafer FQ, Buettner GR & Rodgers VG.The rate of cellular hydrogen peroxide removal shows dependency on GSH: mathematical insight into in vivo H2O2 and GPx concentrations. Free Radical Research 2007411112011211
  138. 138. NichollsP.FitaI.LoewenP. C.Enzymology and structure of catalases. Advances in Inorganic Chemistry 20015151
  139. 139. NivièreV.FontecaveM.Discovery of superoxide reductase: an historical perspective. Journal of Biological Inorganic Chemistry 200492119123
  140. 140. NivièreV.AssoM.WeillC. O.LombardM.GuigliarelliB.FavaudonV.Houée-LevinC.Superoxide reductase from Desulfoarculus baarsii: identification of ptotonation steps in the enzymatic mechanism. Biochemistry 2004433808818
  141. 141. Olorunniji FJ, Iniaghe MO, Adebayo JO, Malomo SO & Adediran SA.Mechanism-Based Inhibition of Myeloperoxidase by Hydrogen Peroxide: Enhancement of Inactivation Rate by Organic Donor Substrates. Open Enzyme Inhibition Journal 2009228
  142. 142. O’NeillP.ChapmenP. W.Potential repair of free radical adducts of dGMP and dG by series of reductants. A pulse radiolytic study. International Journal of Radiation Biology & Related Studies in Physics, Chemistry & Medicine 19854717180
  143. 143. Otterbein LE & Choi AM. Heme oxygenase: colors of defense against cellular stress.American Journal of Physiology- Lung Cellular and Molecular Physiology 2000L10291037
  144. 144. PalenikB.BrahamshaB.LarimerF. W.LandM.HauserL.ChainP.LamerdinJ.RegalaW.EEAllen McCarren. J.PaulsenI.DufresneA.PartenskyF.WebbE. A.WaterburyJ.The genome of a motile marine Synechococcus. Nature 2003424695210371042
  145. 145. PfannschmidtT.Chloroplastredox.signalshow.photosynthesiscontrols.itsown.genesTrends in Plant Science 2003813341
  146. 146. PospíšilP.Molecular mechanisms of production and scavenging of reactive oxygen species by photosystem II. Biochimica et Biophysica Acta 201218781218231
  147. 147. PratF.HoukK. N.FooteC. S.Effect of guanine stacking on the oxidation of 8oxoguanine in B-DNA. Journal of the American Chemical Society 1998
  148. 148. RatnamD. V.AnkolaD. D.BhardwajV.SahanaD. K.KumarM. N.Role of antioxidants in prophylaxis and therapy: A pharmaceutical perspective. Journal of Controlled Release 20061133189207
  149. 149. Requena JR, Fu MX, Ahmed MU, Jenkins AJ, Lyons TJ, Baynes JW & Thorpe SR.Quantification of malondialdehyde and 4hydroxynonenal adducts to lysine residues in native and oxidized human low-density lipoprotein. Biochemical Journal 1997Pt 1) 317-325.
  150. 150. SaitoK.MarkeyS. P.HeyesM. P.Chronic effects of gamma-interferon on quinolinic acid and indoleamine-2,3-dioxygenase in brain of C57BL6 mice. Brain Research 19915461151154
  151. 151. Schafer FQ & Buettner GR.Redox environment of the cell as viewed through the redox state of the glutathione disulphide/glutathione couple. Free Radical Biology and Medicine 2001301111911212
  152. 152. Schaich KM.Metals and lipid oxidation. Contemporary issues. Lipids 1992273209218
  153. 153. Schuster GB.Long-range charge transfer in DNA: Transient structural distortions control the distance dependence. Accounts of Chemical Research 2000334253260
  154. 154. ScottN. G.DuHadaway. J.PigottE.RidgeN.PrendergastG. C.MullerA. J.Mandik-NayakL.TheImmunoregulatory.enzymeI. D. O.paradoxicallydrives. B.cell-mediatedautoimmunity.Journal of Immunology 20091821275097517
  155. 155. SentmanM. L.GranströmM.JakobsonH.ReaumeA.BasuS.MarklundS. L.Phenotypes of mice lacking extracellular superoxide dismutase and copper- and zinc-containing superoxide dismutase. Journal of Biological Chemistry 20062811169046909
  156. 156. ShacterE.Quantification and significance of protein oxidation in biological samples. Drug Metabolism Reviews 2000
  157. 157. Sharp RE, Moser CC, Gigney BR & Dutton PL.Primary Steps in the Energy Conversion Reactions of the Cytochrome bc1 Complex Qo Site. Journal of Bioenergetics and Biomembranes 1999313225233
  158. 158. SpitellerP.KernW.ReinerJ.SpitellerG.Aldehydic lipid peroxidation products derived from linoleic acid. Biochimica et Biophysica Acta 200115313188208
  159. 159. Stadtman ER & Berlett BS.Reactive oxygen-mediated protein oxidation in aging and disease. Chemical Research in Toxicology 1997105485494
  160. 160. Stadtman ER & Oliver NC.Metal-catalyzed oxidation of proteins. Journal of Biological Chemistry 1991266420052008
  161. 161. SteenkenS.Addition-elimination paths in electron-transfer reactions between radicals and molecules. Journal of the Chemical Society, Faraday Transactions I 198783113
  162. 162. SulzerD.BogulavskyJ.LarsenK. E.BehrG.KaratekinE.KleinmanM. H.TurroN.KrantzD.EdwardsR. H.GreeneL. A.ZeccaL.Neuromelanin biosynthesis driven by excess cytosolic catecholamines not accumulated by synaptic vesicles. Proceedings of the National Academy of Sciences of the United States of America 200097221186911874
  163. 163. TakahashiM.AsadaK.Superoxide production in a aprotic interior of chloroplast thylakoids. Archives of Biochemistry and Biophysics 19882672714722
  164. 164. TakikawaO.YoshidaR.KidoR.HayaishiO.Tryptophan degradation in mice initiated by indoleamine-2,3-dioxygenase. Journal of Biological Chemistry 1986261836483653
  165. 165. Taylor MW & Feng GS. Relationship between interferon-gamma, indoleamine 2,3-dioxygenase, and tryptophan catabolism. FASEB Journal199151125162522
  166. 166. Temple MD, Perrone GG & Dawes IW.Complex cellular responses to reactive oxygen species. Trends in Cell Biology 2005156319326
  167. 167. TenhakenR.LevineA.BrissonL. F.DixonR. A.LambC.Function of the oxidative burst in hypersensitive disease resistance. Proceedings of the National Academy of Sciences of the United States of America 1995921041584163
  168. 168. Thannickal VJ & Fanburg BL.Reactive oxygen species in cell signaling. American Journal of Physiology- Lung Cellular and Molecular Physiology 2000L10051028
  169. 169. ThomasJ. A.PolandB.HonzatkoR.PerspectivesProtein.sulfhydrylstheirrole.inthe.antioxidantfunction.ofprotein.S-thiolationArchives of Biochemistry and Biophysics 1995319119
  170. 170. ThomasS. R.MohrD.StockerR.Nitric oxide inhibits indoleamine 2,3-dioxygenase activity in interferon-gamma primed mononuclear phagocytes. Journal of Biological Chemistry 1994269101445714464
  171. 171. TienM.BSBerlettLevine. R. L.ChockP. B.StadtmanE. R.Peroxynitrite-mediated modification of proteins at physiological carbon dioxide concentration: pH dependence of carbonyl formation, tyrosine nitration, and methionine oxidation. Proceedings of the National Academy of Sciences of the United States of America 1999961478097814
  172. 172. TriberisG. P.DimakogianniM.Correlated small polaron hopping transport in 1Ddisordered systems u at high temperatures: a possible charge transport mechanism in DNA. Journal of Physics: Condensed Matter 2009
  173. 173. Turrens JF.Mitochondrial formation of reactive oxygen species. Journal of Physiology 2003Pt 2) 335-344.
  174. 174. UchidaK.KanematsuM.MorimitsuY.OsawaT.NoguchiN.NikiE.Acrolein is a product of lipid peroxidation reaction. Formation of free acrolein and its conjugate with lysine residues in oxidized low density lipoproteins. Journal of Biological Chemistry 1998273261605816066
  175. 175. UradeY.YoshidaR.KitamuraH.HayaishiO.Induction of indoleamine 2,3-dioxygenase in alveolar interstitial cells of mouse lung by bacterial lipopolysaccharide. Journal of Biological Chemistry 19832581066216627
  176. 176. UrsiniF.MaiorinoM.GregolinC.The selenoenzyme phospholipid hydroperoxide glutathione peroxidase. Biochimica et Biophysica Acta 198583916270
  177. 177. ValkoM.LeibfritzD.MoncolJ.CroninM. T. D.MazurM.TelserJ.Free radicals and antioxidants in normal physiological functions and human disease. International Journal of Biochemistry and Cell Biology 20073914484
  178. 178. ValkoM.MorrisH.CroninM. T.Metalstoxicity.oxidativestress.Current Medicinal Chemistry 2005121011611208
  179. 179. VanderHeiden. M. G.ChandelN. S.LiX. X.SchumackerP. T.ColombiniM.ThompsonC. B.Outer mitochondrial membrane permeability can regulate coupled respiration and cell survival. Proceedings of the National Academy of Sciences of the United States of America 200097946664671
  180. 180. VaškováJ.KocanL.FirmentJ.VaškoL.Positive correlation of selenium supplementation and principal antioxidant defenders at sepsis. Current Opinion in Cellular Host-Pathogen Interactions. A Current Opinion in Cell Biology Conference, PS1.18; 2010
  181. 181. VelíšekJ.HajšlováJ.Chemiepotravin.3rd editionOssis.Táborpp.2009
  182. 182. VieiraA. J. S. C.SteenkenS.Patternof. O. H.radicalreaction.withadenine.itsnucleosides.nucleotidesCharacterisation of two types of isomeric OH adduct and their unimolecular transformation reactions. Journal of the American Chemical Society 19901123669866994
  183. 183. Vile GF & Tyrrell RM.Oxidative stress resulting from ultraviolet A irradiation of human skin fibroblasts leads to a heme oxygenase-dependent increase in ferritin. Journal of Biological Chemistry 1993268201467814681
  184. 184. VlasitsJ.JakopitschC.BernroitnerM.ZamockyM.FurtmüllerP. G.ObingerC.Mechanisms of catalase activity of heme peroxidases. Archives of Biochemistry and Biophysics 201050017481
  185. 185. VlasitsJ.JakopitschC.SchwanningerM.HolubarP.ObingerC.Hydrogen peroxide oxidation by catalase-peroxidase follows a non-scrambling mechanism. FEBS Letters 20075812320324
  186. 186. VonSonntag. C.The Chemical Basis of Radiation Biology, in Enzymes, Taylor and Francis Ltd., New York-Philadelphia, 4294571987
  187. 187. VranováE.AtichartpongkulS.VillarroelR.Van MontaguM.InzéD.Van CampW.Comprehensive analysis of gene expression in Nicotiana tabacum leaves acclimated to oxidative stress. Proceedings of the National Academy of Sciences of the United States of America 200299161087010875
  188. 188. Wagner JR.Analysis of oxidative cytosine products in DNA exposed to ionizing radiation. Journal de Chimie Physique et de Physico-Chimie Biologique 1994911280
  189. 189. WagnerJ. R.Van LierJ. E.BergerM.CadetJ.Thymidinehydroperoxides.structuralassignments.conformationalfeatures.thermaldecomposition.inwater.Journal of the American Chemical Society 19941162235
  190. 190. WangH. D.PaganoP. J.DuY.CayatteA. J.QuinnM. T.BrecherP.CohenR. A.Superoxide anion from the adventitia of the rat thoracic aorta inactivates nitric oxide. Circulation Research 1998827810818
  191. 191. Waultier JL & Guillausseau PJ.Advanced glycation and products, their receptors and diabetic angiopathy. Diabetes and Metabolism 200127535
  192. 192. WelinderK. G.MauroJ. K.Nørskov-LauritsenL.Structure of plant and fungal peroxidases. Biochemical Society Transactions 1992202337340
  193. 193. WingeP.BrembuT.BonesA. M.Cloning and characterization of rac-like cDNAs from Arabidopsis thaliana. Plant Molecular Biology 1997354483495
  194. 194. Wink DA & Mitchell JB.Chemical biology of nitric oxide: insight into regulatory, cytotoxic and cytoprotective mechanisms of nitric oxide. Free Radical Biology and Medicine 1998
  195. 195. Winterbourn CC.Superoxide as an intracellular radical sink. Free Radical Biology and Medicine 19931418590
  196. 196. Wolf SP & Dean RT.Glucose autooxidation and protein modification. Biochemical Journal 19872451243250
  197. 197. Wolf SP & Dean RT.Aldehydes and dicarbonyls in non-enzymic glycosylation of proteins. Biochemical Journal Letters 1987248249
  198. 198. Wolin MS, Burke-Wolin TM & Mohazzab-H KM. Roles of NAD(P)H oxidases and reactive oxygen species in vascular oxygen sensing mechanisms.Respiratory Physiology 19991152229238
  199. 199. Wondrak GT, Jacobson MK & Jacobson EL. Endogenous UVA-photosensitizers: mediatorors of skin photodamage and novel targets for skin photoreception.Photochemical and Photobiological Sciences 200652215237
  200. 200. WuergesJ.LeeJ. W.YimY. I.YimH. S.KangS. O.DjinovicCarugo. K.Crystal structure of nickel-containing superoxide dismutase reveals another type of active site. Proceedings of the National Academy of Sciences of the United States of America 20041012385698574
  201. 201. XiaX. G.SchmidtN.TeismannP.FergerB.SchulzJ. B.Dopamine mediates striatal malonate toxicity via dopamine transporter-dependent generation of reactive oxygen species and D2 but not D1 receptor activation. Journal of Neurochemistry 20017916370
  202. 202. YamamotoS.Mammalianlipoxygenases.molecularstructures.functionsBiochimica et Biophysica Acta 1992
  203. 203. YehA. P.HuY.JenneyF. E.Jr AdamsM. W.ReesD. C.Structure of the superoxide reductase from Pyrococcus furiosus in the oxidized and reduced states. Biochemistry 2000391024992508
  204. 204. YoshidaR.UradeY.TokudaM.HayaishiO.Induction of indoleamine 2,3-dioxygenase in mouse lung virus infection. Proceedings of the National Academy of Sciences of the United States of America 197976840844086
  205. 205. YounH. D.YounH.LeeJ. W.YimY. I.LeeJ. K.HahY. C.KangS. O.Unique isoenzymes of superoxide dismutase in Streptomyces griseus. Archives of Biochemistry and Biophysics 19963342341348
  206. 206. ZámockýM.JanecekŠ.KollerF.Common phylogeny of catalase-peroxidases and ascorbate peroxidases. Gene 2000
  207. 207. ZámockýM.RegelsbergerG.JakopitschC.ObingerC.The molecular peculiarities of catalase-peroxidases. FEBS Letters 20014923177182
  208. 208. ZhaoW.DizD. I.MERobbinsOxidative damage pathways in relation to normal tissue injury. British Institute of Radiology 2007Spec 1) S23S31.
  209. 209. ZhuH.BunnH. F.Oxygen sensing and signaling: impact on the regulation of physiologically important genes. Respiratory Physiology 19991152239247
  210. 210. ZweierJ. L.BroderickR.KuppusamyP.Thompson-GormanS.LuttyG. A.Determination of the mechanism of free radical generation in human aortic endothelial cells exposed to anoxia and reoxygenation. Journal of Biological Chemistry 1994269392415624162

Written By

Janka Vašková, Ladislav Vaško and Ivan Kron

Submitted: November 15th, 2011 Published: October 3rd, 2012