Formation of Reactive Nitrogen Species during Peroxidase-catalyzed Oxidation of Nitrite
A POTENTIAL ADDITIONAL MECHANISM OF NITRIC OXIDE-DEPENDENT TOXICITY*

(Received for publication, October 2, 1996, and in revised form, December 17, 1996)

Albert van der Vliet , Jason P. Eiserich , Barry Halliwell § and Carroll E. Cross

From the Division of Pulmonary/Critical Care Medicine, Department of Internal Medicine, University of California, Davis, California 95616 and the § Neurodegenerative Diseases Research Center, Pharmacology Group, King's College, University of London, Manresa Rd., London SW3 6LX, United Kingdom

ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
FOOTNOTES
REFERENCES


ABSTRACT

Involvement of peroxynitrite (ONOO-) in inflammatory diseases has been implicated by detection of 3-nitrotyrosine, an allegedly characteristic protein oxidation product, in various inflamed tissues. We show here that nitrite (NO2-), the primary metabolic end product of nitric oxide (NO·), can be oxidized by the heme peroxidases horseradish peroxidase, myeloperoxidase (MPO), and lactoperoxidase (LPO), in the presence of hydrogen peroxide (H2O2), to most likely form NO·2, which can also contribute to tyrosine nitration during inflammatory processes. Phenolic nitration by MPO-catalyzed NO2- oxidation is only partially inhibited by chloride (Cl-), the presumed major physiological substrate for MPO. In fact, low concentrations of NO2- (2-10 µM) catalyze MPO-mediated oxidation of Cl-, indicated by increased chlorination of monochlorodimedon or 4-hydroxyphenylacetic acid, most likely via reduction of MPO compound II. Peroxidase-catalyzed oxidation of NO2-, as indicated by phenolic nitration, was also observed in the presence of thiocyanate (SCN-), an alternative physiological substrate for mammalian peroxidases. Collectively, our results suggest that NO2-, at physiological or pathological levels, is a substrate for the mammalian peroxidases MPO and lactoperoxidase and that formation of NO2· via peroxidase-catalyzed oxidation of NO2- may provide an additional pathway contributing to cytotoxicity or host defense associated with increased NO· production.


INTRODUCTION

Nitric oxide (NO·)1 is produced by a wide variety of cell types by both constitutive and inducible nitric oxide synthases (1) and has many physiological functions ranging from regulation of vascular tone to neurotransmission and modulation of inflammatory processes (2). Induction of NO· synthesis during inflammatory processes represents a defense mechanism against invading microorganisms, but excessive formation of NO· has also been implicated in host tissue injury (2-4). Although NO· is a free radical, it has selective reactivity and reacts predominantly with other paramagnetic species, including ferrous or ferric iron in heme proteins or iron-sulfur centers, and other radical species such as molecular oxygen (O2), superoxide anion (Obardot 2), and lipid or protein radicals (5-9). Autoxidation of NO· by reaction with O2 results in the formation of nitrite (NO2-) as the primary end-product (10), although at physiological concentrations of NO· and O2, this reaction may be too slow to be of major importance in vivo (4, 6). In the vascular system, NO· is rapidly oxidized by reaction with oxyhemoglobin (HbO2), resulting in formation of methemoglobin (Hb3+) and nitrate (NO3-) (11). NO· also reacts with Hb3+ to form a complex (Hb-NO), which can hydrolyze to Hb2+ and NO2- (reviewed in Ref. 12). NO· also reacts at a near diffusion-limited rate with Obardot 2 to yield peroxynitrite (ONOO-), a powerful oxidizing species, and many of the cytotoxic properties of NO· have in fact been attributed to the formation of ONOO- (4). Reaction of ONOO- or its conjugate acid, ONOOH, with a wide variety of biomolecules also results in concomitant production of NO2- (7). Hence, irrespective of the mechanisms involved, NO2- is a major oxidation product derived from NO·, and increased NO2- levels can often be detected in situations where NO· production is elevated.

In healthy human subjects, NO2- can be detected at levels of 0.5-3.6 µM in plasma (13, 14), ~15 µM in respiratory tract lining fluids (15), 30-210 µM in saliva, and 0.4-60 µM in gastric juice (16). Oral NO2- levels are increased dramatically to near millimolar levels after ingestion of nitrate (NO3-) because of NO3- reduction by the oral microflora (17). Extracellular NO2- levels are also markedly increased during inflammatory processes, reflecting increased NO· production. For instance, increased NO2- levels have been detected in synovial fluids of patients with rheumatoid arthritis (18), and serum NO2- levels of 36 µM have been reported in human immunodeficiency virus-infected patients with interstitial pneumonia (19), dramatically higher than normal serum NO2- levels. Moreover, increased NO2- levels have been detected in condensed exhalates from patients with asthma compared with those of healthy subjects (20), which is in accordance with increases in expired NO· by asthmatics compared with healthy control subjects (21).

Although NO2- is a major end product of NO· metabolism, it does not accumulate in vivo but is rapidly oxidized to NO3- (22). NO2- can be oxidized by HbO2 or oxymyoglobin to form methemoglobin or metmyoglobin and NO3- (23, 24), and catalase has also been demonstrated to contribute to NO2- oxidation (25, 26). However, NO2- can also be oxidized by inflammatory oxidants such as hypochlorous acid (HOCl) (27, 28), and we have recently discovered that oxidation of NO2- by HOCl results in the production of reactive nitrogen intermediates (29). Furthermore, NO2- can also be oxidized by heme peroxidases in the presence of hydrogen peroxide (H2O2) (30-32), and it has been suggested that reactive nitrogen intermediates are produced during such processes (32). Oxidation of NO2- by such mechanisms could be of importance at loci of inflammatory immune processes, when NO· and NO2- levels are enhanced and myeloperoxidase (MPO) and/or eosinophil peroxidase are secreted from activated granulocytes.

The purpose of the present study was to investigate NO2- oxidation by heme peroxidases and to assess its potential physiological importance. The results indicate that MPO and other peroxidases can catalyze oxidation of NO2-, to most likely form NO2· as an intermediate product. At physiological or pathological levels, NO2- can act as a substrate for MPO and lactoperoxidase (LPO), even in the presence of chloride (Cl-) and thiocyanate (SCN-), the proposed major physiological substrates for these peroxidases. Hence, formation of reactive nitrogen intermediates via peroxidase-catalyzed oxidation of NO2- could represent an important contributing mechanism to NO·-mediated toxicity.


EXPERIMENTAL PROCEDURES

Materials

Sodium nitrite (NaNO2), DL-tyrosine, 3-nitrotyrosine, 4-hydroxyphenylacetic acid (HPA), 4-hydroxy-3-nitrophenylacetic acid (3-NO2-HPA), 4-hydroxy-3-chlorophenylacetic acid (3-Cl-HPA), 4-methoxybenzoic acid, 4-methoxy-3-nitrobenzoic acid, 5,5'-dithiobis(2-nitrobenzoic acid) (DTNB), monochlorodimedon, hydrogen peroxide (30%), bovine serum albumin (essentially fatty acid-free), diethylenetriaminepentaacetic acid (DTPA), D-glucose, glucose oxidase (type V-S), catalase (from bovine liver; 25,000 units/mg protein), horseradish peroxidase, and lactoperoxidase (from bovine milk; 120 units/mg protein) were purchased from Sigma. Myeloperoxidase (from human leukocytes; 250-300 units/mg protein) was obtained from Alexis Corp. (San Diego, CA). All peroxidases were used without further purification. 3,3'-Dityrosine and 3,3'-bis(4-hydroxyphenylacetic acid) (3, 3'-diHPA) were synthesized by reaction of DL-tyrosine or HPA with horseradish peroxidase and hydrogen peroxide (H2O2) (33). All other reagents were of the highest purity commercially available.

Peroxidase-catalyzed Oxidation of NO2-

Initial studies were performed with horseradish peroxidase (HRP). DL-Tyrosine (1 mM) was dissolved in 50 mM sodium phosphate buffer (pH 7.4) containing 100 µM DTPA, 1 µM HRP, and various concentrations of NaNO2. DTPA was included in the reaction mixture to avoid interfering reactions with contaminating free metal ions. Reactions were initiated by addition of 1 mM hydrogen peroxide (H2O2) and allowed to proceed at 37 °C for various periods of time. Reactions were terminated at various time points by addition of 10 nM catalase, and tyrosine oxidation products were analyzed by HPLC. Similar experiments were performed in which H2O2 was generated in situ, using D-glucose and glucose oxidase, instead of adding reagent H2O2. Hereto, 10-50 milliunits of glucose oxidase was added to a solution containing 100 µM DL-tyrosine, 1 µM horseradish peroxidase (HRP), 280 µM D-glucose in the absence or presence of NO2- in 50 mM phosphate buffer (pH 7.4) containing 100 µM DTPA, and reaction mixtures were incubated at 37 °C. Reactions were terminated by centrifugation (15,000 rpm) on Microcon-3 concentrators (3,000 molecular weight cutoff) (Amicon Inc., Beverly, MA) to remove proteins. Tyrosine and its oxidation products in the filtrates were analyzed by HPLC using a 5-µm Spherisorb ODS-2 RP-18 column and 93% 50 mM potassium phosphate (pH 3.0), 7% methanol as mobile phase at 1 ml/min, and UV detection at 274 nm and fluorescence detection (excitation, 284 nm and emission, 410 nm). Identification and quantitation of tyrosine oxidation products were performed using external standards and by spectral matching using photodiode array detection (34).

Similar experiments were performed using the mammalian peroxidases myeloperoxidase (MPO) or lactoperoxidase (LPO), in the absence or presence of various concentrations of chloride (Cl-) and/or thiocyanate (SCN-). In these experiments, HPA (1 mM) was used as a substrate instead of tyrosine, to avoid interfering reactions of intermediate oxidants with the amino group in tyrosine. Incubations were terminated by filtration to remove proteins, and the filtrates were analyzed by HPLC. HPA and its oxidation products were separated on a 5-µm Spherisorb ODS-2 RP-18 column using 65% 50 mM potassium phosphate (pH 3.0), 35% methanol as mobile phase at a flow rate of 1 ml/min and analyzed by UV (274 nm) or fluorescence (excitation, 284 nm and emission, 410 nm) detection.

Analysis of Aromatic Hydroxylation

To study the potential formation of hydroxylating species during peroxidase-catalyzed oxidation of NO2-, phenylalanine (5 mM in 50 mM phosphate buffer (pH 7.4)) was used as a trap for aromatic hydroxylation and incubated for 60 min with 10 nM MPO, 700 µM glucose, and 25 milliunits/ml glucose oxidase (resulting in H2O2 production at 10 µM/min) in the absence or presence of 1 mM NO2-. After filtration to remove proteins, the filtrates were analyzed by HPLC for hydroxylation products, using a Spherisorb ODS-2 RP-18 column, 0.1% sodium chloride in 1% (v/v) acetonitrile in water adjusted to pH 3.0 with acetic acid as mobile phase at 1 ml/min, and fluorescence detection (excitation, 275 nm and emission, 305 nm) (35). Using this procedure, the detection limit for p-, m-, and o-tyrosine was approximately 20 nM.

Protein Nitration by Peroxidases and NO2-

Bovine serum albumin (1 mg/ml) in 50 mM sodium phosphate buffer (pH 7.4) was incubated with 10 nM MPO, 10 nM LPO, or 1 µM HRP in the presence of D-glucose/glucose oxidase and varying concentrations of NO2-. After incubation, 100-µl aliquots were mixed with 20 µl of sample loading buffer (20% glycerol, 10% beta -mercaptoethanol, 6% SDS in 125 mM Tris-HCl (pH 6.8)), heated (5 min, 95 °C), and loaded on 10% SDS-polyacrylamide gels for electrophoresis. After electrophoresis, proteins were transferred to polyvinylidene difluoride membranes (Sigma) and immunoblotted with a rabbit polyclonal antibody against 3-nitrotyrosine (Upstate Biotechnology, Lake Placid, NY). The antibody was detected using an anti-rabbit secondary antibody conjugated with HRP (Sigma) and stained using H2O2 and diaminobenzidine (Vector Laboratories, Burlingame, CA).

Oxidation of TNB by MPO or LPO

5-Thio-2-nitrobenzoic acid (TNB) was prepared by reduction of 1 mM DTNB in 100 ml of 50 mM sodium phosphate buffer (pH 7.4) with 4 µl of 2-mercaptoethanol (36). MPO or LPO (10 nM) was added to 50 mM sodium phosphate buffer (pH 7.4) containing 40 µM TNB in the absence or presence of various concentrations of NO2-, Cl-, or SCN-. Reactions were initiated by addition of 30 µM H2O2 and allowed to proceed at 20 °C. Oxidation of TNB to DTNB was followed spectrophotometrically at 412 nm (epsilon 412 = 27,200 M-1 cm-1; Ref. 36).

MPO-catalyzed Chlorination of Monochlorodimedon

Monochlorodimedon (MCD) is a substrate often used to study peroxidase-catalyzed chlorination (37). Chlorination of MCD to dichlorodimedon results in a decrease in absorbance at 290 nm. MCD (40 µM) was dissolved in 50 mM phosphate buffer containing 150 mM Cl- and mixed with 100 µM H2O2 in the absence or presence of NO2- (2-100 µM). Reactions were initiated by addition of 10 nM MPO, and the decrease in A290 was followed spectrophotometrically. Reactions were performed at pH ranging from 6.0 to 7.5, at 20 °C. Control experiments were performed in the absence of Cl- to study MCD oxidation in the presence of NO2- alone.

Other Biochemical Assays

Production of H2O2 by glucose/glucose oxidase was quantitated by oxidation of Fe(II) and formation of a Fe(III)-thiocyanate complex. Aliquots of 800 µl of the reaction mixture were mixed with 100 µl of 10 mg/ml bovine serum albumin, and proteins were precipitated by addition of 100 µl of 100% trichloroacetic acid, after which 800 µl of the supernatants were mixed with 200 µl of Fe(NH4)2(SO4)2 and 100 µl of 2.5 mM KSCN. Formation of the Fe(III)-thiocyanate complex was measured spectrophotometrically at 450 nm within 10 min. Linear standard curves were obtained with 1-50 µM H2O2 solutions treated in a similar manner.

MPO activity was measured spectrophotometrically at 470 nm, using guaiacol oxidation (38). One unit of MPO activity is defined as the amount of enzyme that utilizes 1.0 µmol of H2O2/min in the oxidation of guaiacol at 25 °C and pH 7.0.

NO2- was determined spectrophotometrically at 543 nm, using Griess reagent (1% sulfanylamide, 0.1% naphthylethylenediamine, and 2.5% H3PO4) (28).


RESULTS

Peroxidase-catalyzed Oxidation of NO2-

In the presence of H2O2, peroxidases such as horseradish peroxidase (HRP) are known to catalyze oxidation of the amino acid tyrosine to form tyrosyl radicals, as indicated by the production of the dimerization product 3,3'-dityrosine (34). In the presence of NO2-, 3-nitrotyrosine is formed as an additional product, suggesting that HRP/H2O2 can oxidize both tyrosine and NO2- (32). The identity of 3-nitrotyrosine was confirmed by photodiode array detection and spectral matching compared with authentic 3-nitrotyrosine. Although dityrosine levels reached a maximum after 15-30 min incubation of 1 mM tyrosine with 1 µM HRP, 1 mM H2O2 and varying concentrations of NO2-, formation of 3-nitrotyrosine was found to increase linearly over time during the incubation (Fig. 1A), the rate of tyrosine nitration being proportional to the concentration of NO2- (Fig. 1B). No detectable tyrosine oxidation or nitration was observed in the absence of HRP or H2O2. Qualitatively similar results were obtained using D-glucose/glucose oxidase to continuously generate H2O2 in situ. Tyrosine (100 µM), incubated with 1 µM HRP, 280 µM D-glucose, and 10 milliunits/ml glucose oxidase (generating H2O2 at 1.0 µM/min), was oxidized at a rate comparable with the rate of H2O2 production (Fig. 2). Dityrosine accumulated and eventually it became a substrate for HRP/H2O2 and was oxidized further to trityrosine or other polymeric products (39, 40). In the presence of NO2-, the yield of dityrosine was decreased, and 3-nitrotyrosine was found to accumulate during the incubation (Fig. 2), suggesting that NO2- competes with tyrosine for oxidation by HRP/H2O2.


Fig. 1. Tyrosine nitration by HRP/H2O2/NO2- as a function of NO2-. A, tyrosine (1 mM in 50 mM phosphate buffer (pH 7.4)) was incubated in the presence of 1 mM H2O2, 1 µM HRP, and NO2- at 0.1 (black-square), 0.5 (black-diamond ), or 1.0 mM (bullet ). At various time points, reactions were terminated by addition of 10 nM catalase, and reaction mixtures were filtered on Microcon-3 filters, and the filtrates were analyzed by HPLC for 3-nitrotyrosine. B, tyrosine (1 mM) was incubated in the presence of HRP/H2O2 and the indicated concentration of NO2- for 60 min, and the rate of 3-nitrotyrosine formation was calculated as a function of NO2- concentrations. Mean values of 2-3 different experiments are shown.
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Fig. 2. Time-dependent tyrosine oxidation by HRP/H2O2 and NO2-. Tyrosine (100 µM in 50 mM phosphate buffer containing 100 µM DTPA (pH 7.4)) was incubated in the presence of 1 µM HRP, 280 µM D-glucose, and 10 milliunits/ml glucose oxidase (generating 1.0 µM/min H2O2) in the absence (A) or presence (B) of 1 mM NO2-. At various time points aliquots were taken, filtered on Microcon-3 filters, and the supernatants analyzed by HPLC for tyrosine (black-square), dityrosine (black-diamond ), or 3-nitrotyrosine (bullet ). Data are represented as mean values from two separate experiments.
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The mammalian peroxidases myeloperoxidase (MPO) and lactoperoxidase (LPO) are also able to oxidize tyrosine in the presence of H2O2, to form dityrosine, and are reported to do so more efficiently than HRP (39). In the presence of NO2-, these peroxidases also generate 3-nitrotyrosine as an additional product. Using 1 mM tyrosine, 100 µM NO2-, and 4.0 µM/min H2O2, the yield of both oxidation products increased with the concentration of MPO or LPO and was maximal at enzyme concentrations between 10 and 20 nM. At higher enzyme concentrations the yield of dityrosine did not increase further, and production of 3-nitrotyrosine was found to decrease (not shown). For this reason, further studies were performed using 10 nM of MPO or LPO. Both tyrosine oxidation products accumulated rapidly during the first 30 min, and the rate of product formation declined dramatically thereafter, possibly because of enzyme inactivation or due to depletion of dissolved molecular oxygen necessary for H2O2 generation. Hence, in further experiments using glucose/glucose oxidase, reaction mixtures were incubated for 30 min. As demonstrated in Table I, the extent of tyrosine nitration by MPO or LPO was dependent on the initial NO2- concentration, although the production of dityrosine was not dramatically affected. The yield of both products was markedly lower when LPO was used instead of MPO, consistent with the notion that MPO more efficiently catalyzes oxidation of tyrosine (39) and perhaps NO2-. However, the results indicate that both peroxidases are capable of oxidizing NO2- in the presence of H2O2 to form a reactive intermediate that is capable of nitrating tyrosine.

Table I.

Tyrosine oxidation by MPO and LPO in the presence of NO2-

Tyrosine (1 mM in 50 mM phosphate buffer (pH 7.4) including 100 µM DTPA) was incubated with the indicated concentration of NO2-, 280 µM glucose, and 50 milliunits/ml glucose oxidase (generating 4.0 µM/min H2O2) and 10 nM MPO or 10 nM LPO for 30 min at 37 °C. Tyrosine oxidation products were measured by HPLC with UV detection or fluorescence detection. Mean values ± SD from three separate experiments are shown.
NO2- Myeloperoxidase
Lactoperoxidase
DiTyr NO2-Tyr DiTyr NO2-Tyr

mM µM µM µM µM
0 33.2  ± 1.7 15.7  ± 3.2
0.01 33.0  ± 1.2 0.27  ± 0.07 12.9  ± 1.3 0.09  ± 0.01
0.1 30.7  ± 1.6 2.21  ± 0.39 20.0  ± 1.4 0.63  ± 0.11
1.0 34.0  ± 1.8 8.60  ± 0.19 14.8  ± 1.6 1.34  ± 0.15

In the presence of NO2-, MPO/H2O2 was also found capable of nitrating tyrosine residues in proteins. As shown in Fig. 3, reaction of 10 nM MPO with 1 mg/ml albumin in the presence of 0.5 or 1.0 mM NO2- and 10 µM/min H2O2 resulted in nitration of tyrosine residues, as detected by SDS-polyacrylamide gel electrophoresis and Western blotting using a rabbit polyclonal antibody against 3-nitrotyrosine. Protein nitration was also observed when LPO or HRP were used instead of MPO (not shown).


Fig. 3. Protein nitration by MPO/H2O2/NO2-. Bovine serum albumin (BSA) (1 mg/ml in 50 mM sodium phosphate (pH 7.4)) was incubated in the presence of 10 nM MPO, 700 µM glucose, and 125 milliunits/ml glucose oxidase (producing 10 µM/min H2O2), and various concentrations of NO2-. After incubation for 30 min, 5 µg of protein was loaded on 10% SDS-polyacrylamide gels for electrophoresis, and proteins were transferred and immunostained using a rabbit polyclonal antibody against nitrotyrosine (A) or stained with Coomassie Blue (B). Lane 1, complete system without NO2-; lane 2, + 0.5 mM NO2-; lane 3, + 1.0 mM NO2-.
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Possible Oxidation Mechanisms

Peroxidases commonly oxidize aromatic substrates by a one-electron oxidation mechanism, although oxidation of halides by mammalian peroxidases has been generally thought to occur via two-electron oxidation (39-42). Peroxidase-catalyzed oxidation of NO2- therefore results in formation of either NO2· or an NO2+-like intermediate. Oxidation of NO2- by MPO/H2O2 did not result in aromatic hydroxylation of the amino acid phenylalanine, indicating that a hydroxylating species such as ONOOH (43) is not formed during MPO-catalyzed NO2- oxidation. Addition of 1 mM NO2+ (as the nitryl salt NO2BF4) to a solution of 1 mM tyrosine at pH 7.4 resulted in formation of 2 ± 1 µM 3-nitrotyrosine (mean ± S.D.; n = 3), which is markedly less than the extent of nitration by MPO/H2O2 in the presence of 1 mM NO2- (Table I). Although NO2+ is capable of nitrating aromatic rings by electrophilic aromatic substitution, up to 2 mM NO2BF4 did not induce detectable nitration of the aromatic substrate 4-methoxybenzoic acid (a substrate incapable of forming phenolic radicals) in aqueous solution, possibly because of rapid hydrolysis of NO2+. Hence, it is unlikely that phenolic nitration via peroxidase-catalyzed oxidation of NO2- is due to formation of NO2+.

NO2- as a Competing Substrate for MPO or LPO

The presumed major physiological substrate for MPO is chloride (Cl-), although other anions such as bromide (Br-) or thiocyanate (SCN-) are sufficiently abundant in biological fluids to act as alternative physiological substrates (44, 45). Similarly, SCN- and Br- have been proposed as a physiological substrates for eosinophil peroxidase (EPO) or LPO (46-48). We therefore investigated whether MPO or LPO could catalyze oxidization of NO2- in the presence of either Cl- or SCN-, using TNB as an oxidizable substrate. In the absence of other substrates, MPO/H2O2 slowly oxidizes TNB to form DTNB. The rate of TNB oxidation was, however, enhanced dramatically when either Cl-, NO2-, or SCN- was also present in the reaction mixture, suggesting that these anions act as substrates for MPO/H2O2 to form diffusible oxidation products which in turn oxidize TNB. Rates of TNB oxidation by MPO/H2O2 in the presence of these anionic substrates are shown in Fig. 4 and demonstrate that SCN- is used more efficiently as a substrate than NO2-, which is preferred as a substrate over Cl- for MPO. Similarly, NO2- is a substrate for LPO, but LPO/H2O2 preferentially utilizes SCN- in oxidation of TNB (not shown). Only minimal oxidation of TNB was observed by LPO/H2O2 in the presence of 100 mM Cl-. As LPO is unable to catalyze Cl- oxidation (see below), this is most likely due to contaminants, such as Br-.


Fig. 4. Increased MPO-catalyzed TNB oxidation by Cl-, NO2-, or SCN-. MPO (6 nM) was mixed with 40 µM TNB, and the indicated concentration of Cl- (bullet ), NO2- (black-diamond ), or SCN- (black-square). Reactions were initiated with 30 µM H2O2, and TNB oxidation was followed spectrophotometrically at 412 nm. Formation of DTNB was calculated using epsilon 412 = 27,200 M-1 cm-1. Results were corrected for TNB oxidation in the absence of anionic substrate (0.06 µM/min). Mean values of 2-3 measurements are shown.
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The rate of TNB oxidation by MPO/H2O2 and 100 mM Cl- was found to be markedly inhibited in the presence of NO2-. High concentrations of NO2- were found to inhibit TNB oxidation by MPO/H2O2/Cl- to rates more similar to those observed in the presence of NO2- alone (Fig. 5A), and dramatic (>75%) inhibition of Cl--dependent TNB oxidation was observed in the presence of NO2- at concentrations as low as 10 µM. Nearly identical results were obtained when 100 µM SCN- was used instead of 100 mM Cl- (not shown). As 10 µM NO2- is unlikely to inhibit oxidation of 40 µM TNB by the intermediately formed oxidants (HOCl or HOSCN) to such an extent, the results suggest that NO2- also competitively inhibits Cl- or SCN- oxidation by MPO/H2O2. In contrast to the results with MPO, SCN--catalyzed TNB oxidation by LPO/H2O2 was found to be enhanced by 10 µM NO2- (Fig. 5B), suggesting that NO2- also can act as a substrate for LPO in the presence of physiological levels of SCN-.


Fig. 5. NO2- as a peroxidase substrate in the presence of Cl- or SCN-. A, MPO (6 nM) was mixed with 40 µM TNB in 50 mM phosphate buffer (pH 7.4), and 100 mM Cl- and/or various concentrations of NO2-. Reactions were initiated by addition of 30 µM H2O2, and TNB oxidation was followed spectrophotometrically at 412 nm. bullet , 100 mM Cl-; square , 1 mM NO2-; black-square, 100 mM Cl- + 1 mM NO2-; diamond , 100 µM NO2-; black-diamond , 100 mM Cl- + 100 µM NO2-; triangle , 10 µM NO2-; and black-triangle, 100 mM Cl- + 10 µM NO2-. B, LPO (10 nM) was mixed with 40 µM TNB and various concentrations of SCN- and/or NO2-. Reactions were started by addition of 30 µM H2O2 and the decrease in A412 was followed. black-square, 1 mM SCN-; square , 1 mM SCN- + 10 µM NO2-; bullet , 100 µM SCN-; open circle , 100 µM SCN- + 10 µM NO2-; diamond , 10 µM NO2-. Mean values of 2-3 measurements are shown.
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Effect of NO2- on MPO-catalyzed Chlorination of MCD

Addition of MPO to a solution containing 40 µM MCD, 100 µM H2O2, and 150 mM Cl- was found to result in chlorination of MCD, as demonstrated by a decrease in A290 (37, 49). The rate of MPO-catalyzed MCD chlorination increased with decreasing pH (between pH 6.0 and 7.5), consistent with earlier reports (50). Reaction of MPO with excess H2O2 results in rapid formation of MPO compound I, which is then spontaneously reduced to compound II (50, 51). As MPO compound II is unable to oxidize Cl- (49), accumulation of compound II will result in a decreased rate of MCD chlorination, and this was indeed observed (Fig. 6). Addition of 2-10 µM NO2- to the reaction mixture did not significantly affect the initial rate of MCD oxidation but diminished the decrease in chlorination rate over time, and at NO2- concentrations of 5 µM or higher, the rate of MCD chlorination was constant over the course of the experiment (Fig. 6). The rate of MCD chlorination was not increased much further in the presence of higher NO2- concentrations (Fig. 6), possibly because high NO2- concentrations may scavenge the chlorinating species, thereby partly inhibiting MCD chlorination. The MCD concentration was only minimally affected by MPO/H2O2 and 100 µM NO2- in the absence of Cl- (minimal decrease in A290; Fig. 6), indicating that oxidation products of NO2- do not importantly contribute to the observed increased MCD oxidation/chlorination in the presence of Cl-.


Fig. 6. Effect of NO2- on MPO-catalyzed chlorination of monochlorodimedon (MCD). MCD (40 µM) in 50 mM phosphate buffer, containing 150 mM Cl- (pH 6.0) was mixed with 100 µM H2O2 in the absence (square ) or presence of 2 (black-square), 5(bullet ), 10 (black-triangle) or 100 (black-diamond ) µM NO2-. After addition of 6 nM MPO the decrease in A290 was followed immediately. As a control, MCD oxidation by MPO/H2O2 and 100 µM NO2- was measured in the absence of Cl- (diamond ). Results are expressed as the average of 2-3 measurements.
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Peroxidase-catalyzed Phenolic Nitration in the Presence of Cl- or SCN-

Modification of phenolic substrates by MPO-catalyzed oxidation of NO2- was also studied in the presence of alternative peroxidase substrates. MPO-catalyzed oxidation of Cl- is known to induce chlorination of aromatic substrates (37, 52, 53), which was confirmed in the present study using HPA. Incubation of 1 mM HPA with MPO/H2O2 and Cl- resulted in formation of 3-chloro-HPA, the extent of HPA chlorination being proportional to the concentration of Cl-. In the absence of Cl-, MPO/H2O2 catalyzes oxidation of HPA to 3,3'-diHPA (the dimerization product of HPA), and formation of 3,3'-diHPA was partly inhibited by Cl- (the yield of 3,3'-diHPA was decreased up to 30% in the presence of 50 mM Cl-, but to a lesser extent in the presence of higher Cl- concentrations). Thus, high concentrations of Cl- appear to outcompete HPA for oxidation by MPO/H2O2, but Cl- oxidation may also contribute to HPA oxidation and dimerization.

MPO-catalyzed HPA chlorination in the presence of physiological levels of Cl- (150 mM) was found to be enhanced by low concentrations of NO2-. As shown in Fig. 7, HPA chlorination was increased most dramatically in the presence of 2-10 µM NO2-. Increasing NO2- concentrations did not dramatically enhance further HPA chlorination and, in fact, HPA chlorination was partly inhibited in the presence of concentrations of NO2- above 200 µM (not shown). Addition of 2-10 µM NO2- also enhanced MPO/H2O2/Cl--induced formation of 3,3'-diHPA, and 3-NO2-HPA was formed as an additional product (Fig. 7), and the extent of HPA nitration and dimerization continued to increase in the presence of increasing concentrations of NO2-. These observations are consistent with our results with MCD and suggest that NO2- increases MPO-catalyzed Cl- oxidation by reducing MPO compound II to ferric MPO. At high concentrations, NO2- may also partly scavenge the intermediate chlorinating species (HOCl or Cl2; Ref. 54), thereby inhibiting aromatic chlorination by MPO/H2O2/Cl-.


Fig. 7. Effect of NO2- on aromatic modification by MPO/H2O2/Cl-. HPA (1 mM) was incubated at 37 °C in 50 mM phosphate buffer containing 150 mM Cl- and 100 µM DTPA (pH 7.4) with 10 nM MPO in the presence of the indicated concentration of NO2-. H2O2 was generated at 4.0 µM H2O2/min using 280 µM D-glucose and 50 milliunits/ml glucose oxidase, and reactions were initiated by addition of glucose oxidase. After 30 min reaction mixtures were filtered on Microcon-3 filters, and supernatants were analyzed by HPLC for 3-Cl-HPA (black-square), 3,3'-diHPA (black-triangle), and 3-NO2-HPA (bullet ). Average values of 2-3 separate experiments are shown.
[View Larger Version of this Image (17K GIF file)]


Nitration of HPA by MPO/H2O2/NO2- could be partially inhibited by a large excess of Cl-. When the inhibition of HPA nitration was plotted against the negative logarithm of the concentration of Cl-, a sigmoidal curve was obtained (Fig. 8), suggesting that Cl- competitively inhibits NO2- oxidation by MPO/H2O2, and the IC50 for Cl- increased when higher concentrations of NO2- were used. HPA nitration was inhibited maximally by 30-45% (Fig. 8), and nitration of HPA was still observed even when Cl- was present in 1,000-10,000-fold excess over NO2-.


Fig. 8. Inhibition of MPO/H2O2/NO2--induced aromatic nitration by Cl-. HPA (1 mM) was incubated for 30 min at 37 °C in 50 mM phosphate buffer containing 100 µM DTPA (pH 7.4) in the presence of 10 nM MPO, 4.0 µM H2O2/min, and 10 µM (black-square), 100 µM (black-diamond ), or 1 mM (bullet ) NO2- and the indicated concentration of Cl-. Reaction mixtures were filtered on Microcon-3 filters, and supernatants were analyzed by HPLC for 3-NO2-HPA. Average results from three separate experiments are shown.
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Extracellular fluids also contain significant levels of SCN- (20-120 µM in plasma but much higher in secretions such as milk or saliva, for example), which has been proposed as an alternative substrate for peroxidases. Both nitration and chlorination of HPA, observed after incubation of HPA with MPO/H2O2 in the presence of 150 mM Cl- and 100 µM NO2-, were found to be inhibited by SCN- (Fig. 9). However, chlorination of HPA was inhibited with an IC50 approx 5 µM, and no chlorination of HPA could be detected in the presence of >25 µM SCN- (<0.2 µM 3-Cl-HPA). On the other hand, nitration of HPA was inhibited much less efficiently by SCN- (IC50 approx 60 µM), and formation of 3-NO2-HPA was still detectable in the presence of 500 µM SCN- (0.05 µM 3-NO2-HPA after 60 min incubation), indicating that NO2- can act as a substrate for MPO/H2O2 even in the presence of physiological levels of Cl- and SCN-. Nitration of HPA by LPO/H2O2/NO2- was not affected by 150 mM Cl-, consistent with the notion that Cl- is not a substrate for LPO. Moreover, chlorination of HPA was not observed by LPO/H2O2/Cl- in the presence or absence of NO2-. HPA nitration by LPO/H2O2 and 100 µM NO2- was inhibited by SCN- (IC50 approx 10 µM), and no HPA nitration was detected in the presence of >100 µM SCN-.


Fig. 9. Inhibition of peroxidase-catalyzed nitration and chlorination by SCN-. A, MPO (10 nM) was incubated at 37 °C in 50 mM phosphate buffer (pH 7.4) containing 150 mM Cl-, 1 mM HPA, 100 µM NO2-, 280 µM D-glucose, and the indicated concentration of SCN-. Reactions were initiated by addition of 50 milliunits/ml glucose oxidase (resulting in production of 4.0 µM H2O2/min), and formation of 3-Cl-HPA (black-square) and 3-NO2-HPA (bullet ) was measured after 30 min. Mean values of 2-3 determinations are presented. B, similar experiment as above with 10 nM LPO instead of MPO.
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DISCUSSION

The results presented herein demonstrate that the peroxidases HRP, MPO, and LPO can oxidize NO2- in the presence of H2O2 or H2O2-generating systems to form reactive nitrogen intermediate(s) capable of nitrating phenolic compounds such as tyrosine. At acidic pH, NO2- is partly protonated and reacts with H2O2 directly to form peroxynitrous acid (ONOOH), which causes aromatic nitration (34). However, this did not occur to an appreciable extent at pH 7.4 under our reaction conditions. In addition to being oxidized to more reactive nitrogen species, NO2- was also found to catalyze MPO-mediated oxidation of TNB or chlorination of MCD or HPA. These results confirm and extend a recent study by Shibata et al. (32), who demonstrated that HRP and H2O2 oxidize NO2- to form an intermediate that is capable of causing chlorophyll degradation and tyrosine nitration. They postulated that HRP oxidizes NO2- to NO2· in the presence of H2O2 according to the following reaction scheme (Reactions 1-3):
<UP>HRP</UP>+<UP>H</UP><SUB>2</SUB><UP>O</UP><SUB>2</SUB> → <UP>Compound I</UP>+<UP>H</UP><SUB>2</SUB><UP>O</UP>
<UP>Compound I</UP>+<UP>NO</UP><SUP><UP>−</UP></SUP><SUB>2</SUB> → <UP>Compound II</UP>+<UP>NO</UP><SUP>·</SUP><SUB>2</SUB>
<UP>Compound II</UP>+<UP>NO</UP><SUP><UP>−</UP></SUP><SUB>2</SUB> → <UP>HRP</UP>+<UP>NO</UP><SUP>·</SUP><SUB>2</SUB>
<UP>R<SC>eactions</SC></UP> 1–3
Since MPO and LPO can catalyze both one- and two-electron oxidations (39-42), oxidation of NO2- by these peroxidases can result in formation of either NO2· or NO2+. We were unable to obtain evidence for intermediate formation of NO2+, and chemical studies have indicated that NO2+ is an inefficient nitrating species in neutral aqueous solutions. Hence, it seems more likely that NO2· is the nitrating intermediate during NO2- oxidation, although oxidation of NO2- by MPO/H2O2 may also result in formation of an enzyme bound NO2+ species, which may be a more efficient nitrating species than "free" NO2+. Peroxidase-catalyzed oxidation of NO2- also results in nitration of tyrosine residues in proteins, which are unlikely to act as direct substrates for MPO and LPO, because they contain a restricted active site and may not allow direct oxidation of large proteins (41, 55). Hence, NO2- is most likely oxidized to a diffusible nitrating species, such as NO2·. As peroxidases oxidize tyrosine to form tyrosyl radicals (33, 40), indirect oxidation of NO2- by intermediately produced tyrosyl radicals via electron transfer may also contribute to the observed results (Reaction R4).
<UP>Tyr-O</UP><SUP>·</SUP>+<UP>NO</UP><SUP><UP>−</UP></SUP><SUB>2</SUB> → <UP>Tyr-O</UP><SUP><UP>−</UP></SUP>+<UP>NO</UP><SUP>·</SUP><SUB>2</SUB>
<UP><SC>Reaction R4</SC></UP>
The fact that NO2- enhances MPO-catalyzed TNB oxidation or MCD chlorination in the presence of Cl- clearly demonstrates that NO2- is a direct substrate for MPO and other peroxidases. However, in the absence of other oxidizable substrates, reaction of NO2- with MPO/H2O2 did not result in significant depletion of NO2-, most likely because the oxidation product of NO2- reacts with MPO itself, either resulting in enzyme inactivation or in regeneration of NO2-. Oxidation of NO2- by MPO/H2O2 in the presence of tyrosine resulted in a small but significant depletion of NO2-, corresponding to the extent of tyrosine nitration (not shown), supporting formation of NO2· rather than NO2+ as the oxidation product, as NO2+ reacts extremely rapidly with water to NO3- (56). Because NO2- can be regenerated during oxidation of substrates by NO2· (Reaction R5), NO2- can act as a catalyst in the oxidation of other substrates by MPO/H2O2.
<UP>NO</UP><SUP>·</SUP><SUB>2</SUB>+<UP>RH</UP> → <UP>NO</UP><SUP><UP>−</UP></SUP><SUB>2</SUB>+<UP>H</UP><SUP><UP>+</UP></SUP>+<UP>R</UP><SUP>·</SUP>
<UP><SC>Reaction R5</SC></UP>
Recently, Floris et al. (57) have demonstrated that ONOO-/ONOOH reacts rapidly with MPO, resulting in formation of MPO compound II without intermediate detection of compound I. They proposed that reaction of ONOOH with MPO initially results in formation of a compound I-NO2- complex which then immediately converts to compound II and NO2·, consistent with one-electron oxidation of NO2- by MPO/H2O2. This mechanism may also explain the demonstrated ability of MPO to catalyze phenolic nitration by ONOOH (58), which may in part have been due to the presence of NO2- in the ONOO- stock solutions. The overall rate of compound II formation by reaction of ONOOH with MPO was reported to occur with a second order rate constant of 2 × 107 M-1 s-1 (57), which indicates that the oxidation of NO2- by MPO compound I occurs extremely rapidly.

Taken together, our results suggest that in the presence of H2O2, heme peroxidases can catalyze oxidation of NO2- to predominantly NO2·, and this could contribute to aromatic nitration in vivo at sites where NO2- levels are sufficiently high. Although tyrosine nitration by NO2· is rather inefficient because two NO2· molecules are needed for nitration of one tyrosine residue, and intermediately formed tyrosyl radicals undergo rapid reactions including dimerization, tyrosyl radicals in proteins are often more long-lived than free tyrosyl radicals, rendering tyrosyl radicals in proteins more susceptible targets for nitration by NO2·, via rapid radical-radical reaction (k = 3 × 109 M-1 s-1; Ref. 59). Moreover, tyrosyl radicals have been detected in various enzymes such as ribonucleotide reductase and prostaglandin H synthase (60), and formation of NO2· could result in nitration of these tyrosine residues and potentially affect enzyme activity.

Nitrite as a Physiological Substrate for Mammalian Peroxidases

Although mammalian peroxidases such as MPO and LPO can catalyze oxidation of NO2-, biological fluids contain alternative substrates that might be preferentially oxidized. For instance, intracellular and extracellular fluids contain 100-150 mM Cl-, which is commonly thought to be the physiological substrate for MPO (48). Eosinophil peroxidase (EPO) can also oxidize Cl-, but Br- and SCN- (plasma levels are 20-100 and 20-120 µM, respectively; Ref. 45) have been suggested as more important physiological substrates for EPO, because these anions are oxidized more easily. Similarly, SCN- has been proposed as the physiological substrate for LPO in saliva or milk, whereas SCN- levels are as high as 5 mM (47). Reported oxidation potentials of SCN-, Br-, Cl- and NO2- are summarized below in Reactions 6-9 (61, 62):
            E°(<UP>V</UP>)
   <UP>SCN</UP><SUP><UP>−</UP></SUP> → <UP>SCN</UP><SUP>·</SUP>+e<SUP><UP>−</UP></SUP>  <UP>−</UP>0.77 <UP>V</UP>
<UP>NO</UP><SUP><UP>−</UP></SUP><SUB>2</SUB> → <UP>NO</UP><SUP>·</SUP><SUB>2</SUB> +e<SUP><UP>−</UP></SUP> <UP>−</UP>0.99 <UP>V</UP>
<UP>Br</UP><SUP><UP>−</UP></SUP> → <UP>Br</UP><SUP>·</SUP> +e<SUP><UP>−</UP></SUP> <UP>−</UP>1.07 <UP>V</UP>
<UP>Cl</UP><SUP><UP>−</UP></SUP> → <UP>Cl</UP><SUP>·</SUP> +e<SUP><UP>−</UP></SUP> <UP>−</UP>1.36 <UP>V</UP>
<UP>R<SC>eactions</SC></UP> 6–9
The respective oxidation potentials predict that NO2- is oxidized more efficiently by MPO/H2O2 than Cl-, consistent with our studies with TNB. Reduction potentials of the couples compound I/compound II and compound II/ferric enzyme for HRP have been reported to be 0.90 and 0.87 V at pH 7.0 (63), indicating that compounds I and II of HRP are capable of oxidizing NO2- but will not efficiently oxidize Cl-. No reduction potentials have been reported for the couples of compound I/compound II and compound II/ferric enzyme for LPO, EPO, or MPO, but the fact that compound I of MPO and EPO is able to oxidize Cl- suggests that compound I of these enzymes is more strongly oxidizing than HRP compound I.

Our studies with TNB have indicated that NO2- acts as a competing substrate for MPO/H2O2 in the presence of Cl- or SCN-, as low concentrations of NO2- dramatically inhibited TNB oxidation by MPO/H2O2/Cl- or MPO/H2O2/SCN-. However, NO2- could also scavenge the oxidation products of Cl- or SCN-, thereby inhibiting TNB oxidation. In contrast, TNB oxidation by LPO/H2O2/SCN- was enhanced by NO2-, indicating that NO2- can act as a substrate for LPO/H2O2, even in the presence of excess SCN-. The opposing results obtained with both peroxidases may be due to different enzyme specificity or differences in oxidation mechanisms. The notion that NO2- can act as a substrate for MPO/H2O2 in the presence of physiological levels of Cl- is further supported by our studies with MCD chlorination or HPA oxidation, which demonstrated that low concentrations of NO2- markedly enhanced MPO-catalyzed chlorination of these substrates. Reducing agents such as ascorbate, urate, or Obardot 2 have been reported to enhance the chlorinating activity of MPO/H2O2/Cl- by reducing MPO compound II to the native ferric enzyme (50, 64, 65). As MPO compound II is unable to oxidize Cl- (49, 65), accumulation of MPO compound II would reduce the rate of aromatic chlorination, which can be prevented in the presence of reducing agents. We propose that NO2- may act similarly as a reducing substrate for MPO compound II, thereby recycling it to ferric MPO, which would again participate in oxidation of Cl-. In this process, NO2- is oxidized by a one-electron mechanism to NO2·, causing enhanced HPA dimerization and nitration by MPO/H2O2/Cl- in the presence of NO2-. In a recent study with water-soluble metalloporphyrins, oxidation of a Fe(III) porphyrin with m-chloroperoxybenzoate was found to produce the O=Fe(IV) species (equivalent to compound II), which could rapidly be reduced back to the Fe(III) porphyrin by addition of equivalent amounts of NO2- (66), which is consistent with the proposed reduction of MPO compound II by NO2-. The observed partial inhibition of MPO/H2O2/NO2--induced HPA nitration by Cl- is also consistent with this mechanism. Although Cl- is able to compete with NO2- for oxidation by MPO compound I, Cl- cannot reduce MPO compound II (65) and hence does not completely inhibit MPO/H2O2/NO2--mediated HPA nitration. Fig. 10 schematically summarizes the proposed mechanisms by which MPO can catalyze oxidation of NO2-.


Fig. 10. Possible mechanisms of NO2- oxidation by MPO/H2O2. MPO reacts with H2O2 to form compound I, which can further react with H2O2 to compound II. MPO compound I can oxidize Cl- and perhaps NO2- by a two-electron oxidation mechanism, with regeneration of ferric MPO. Alternatively, NO2- can oxidized by a one-electron oxidation to form compound II and NO2·. Furthermore, MPO compound II can oxidize NO2-, by one-electron oxidation, to form NO2· and regenerate ferric MPO.
[View Larger Version of this Image (15K GIF file)]


In addition to catalyzing MPO-mediated chlorination by reduction of MPO compound II, NO2- may also react with the initially formed chlorinating species (HOCl or Cl2; Ref. 54), to form a reactive intermediate similar to nitryl chloride (NO2Cl), which is capable of inducing aromatic chlorination and nitration (29). Hence, NO2Cl may contribute to the observed MCD chlorination or HPA chlorination and nitration by MPO/H2O2/Cl- in the presence of NO2-. However, this mechanism is unlikely to fully account for the observed increases in aromatic chlorination and nitration because (i) low concentrations of NO2- (2-10 µM) are unlikely to efficiently compete with the much higher concentrations of MCD or HPA for reaction with HOCl or Cl2, and (ii) the produced intermediates (including NO2Cl) are unstable and rapidly hydrolyze to NO2- and Cl-. In fact, studies with reagent HOCl have indicated that chlorination of HPA by HOCl was decreased in the presence of NO2-, with a concomitant increase in HPA nitration (29), which most likely explains why increasing concentrations of NO2- failed to dramatically further enhance aromatic chlorination by MPO/H2O2/Cl- (Figs. 6 and 7).

Both nitration and chlorination of HPA by MPO/H2O2 in the presence of Cl- and NO2- were inhibited by SCN-, an alternative physiological substrate for mammalian peroxidases. However, the fact that nitration of HPA was still detectable in the presence of physiological SCN- levels, whereas HPA chlorination was completely inhibited, further suggests that nitration is due to direct oxidation of NO2- by MPO/H2O2, rather than via oxidation by chlorinating intermediates such as HOCl or Cl2. Similarly, LPO/H2O2/NO2- also caused detectable HPA nitration in the presence of SCN- at concentrations similar to that of NO2-. SCN- may competitively inhibit oxidation of Cl- and NO2- by these enzymes but could also act by scavenging reactive intermediates formed by oxidation of Cl- or NO2-.

Physiological/Pathological Consequences

One important implication of the results presented herein is that oxidation of NO2- by mammalian peroxidases in the presence of H2O2 may represent an alternative or additional mechanism of aromatic nitration in vivo. We have recently demonstrated that oxidation of NO2- by HOCl also results in formation of reactive intermediates capable of inducing phenolic nitration (29), and both mechanisms of NO2- oxidation may contribute to tyrosine nitration observed in tissues undergoing inflammatory responses and where high levels of MPO are present. Interestingly, 3-nitrotyrosine and large amounts of active MPO have both been detected in atherosclerotic lesions (67-69). Moreover, high levels of MPO are also present in the rheumatoid joint of patients with rheumatoid arthritis, and synovial fluids of rheumatoid arthritis patients are also reported to contain increased levels of NO2- and 3-nitrotyrosine (18, 70). Furthermore, tyrosine nitration has also been detected in lung sections of patients with acute pulmonary inflammation (71, 72), a condition characterized by infiltration of neutrophils as well as increased production of NO· and NO2- (e.g. Refs. 20, 21). Detection of 3-nitrotyrosine in vivo, often regarded specific evidence for formation of ONOO-, may therefore also be due to formation of reactive nitrogen species by oxidation of NO2- by MPO or by HOCl. Hence, 3-nitrotyrosine is not a unique biomarker of ONOO- production but merely indicates formation of reactive nitrogen species derived from NO·, and formation of 3-nitrotyrosine in vivo may often largely depend on the presence of MPO. Interestingly, it was recently reported that NO· synthase is induced in cytokine-stimulated human neutrophils and found to be co-localized with MPO in primary granules (73). Induction of NO· synthase and activation of MPO are both likely to be involved in the observed tyrosine nitration around ingested bacteria (73).

We have performed preliminary studies with isolated human polymorphonuclear neutrophils (PMN), which demonstrated that stimulation of 2 × 106/ml PMN in PBS (150 mM Cl-) with 100 ng/ml phorbol myristate acetate in the presence of 10-100 µM NO2- and 1 mM HPA results in HPA nitration. Furthermore, HPA chlorination was enhanced in the presence of NO2-, similar to our studies with purified MPO. Stimulation of PMN with phorbol myristate acetate results in complete degranulation and secretion of MPO into the incubation medium. After 60-min incubations, the medium MPO activity was 14 ± 5 milliunits/ml with 2 × 106/ml resting PMNs and 175 ± 56 milliunits/ml (mean ± S.D.; n = 4) after stimulation of PMN with phorbol myristate acetate. The amount of MPO released from these PMNs was similar to that used in the studies with purified MPO; 10 nM MPO corresponds to an enzyme activity of 150 milliunits/ml. The same number of activated PMN produce Obardot 2 at a rate of about 5 µM/min (74), which would yield 2.5 µM H2O2/min upon dismutation, similar to the rate of H2O2 production used in our studies with purified MPO and glucose/glucose oxidase. Therefore, the NO2- oxidation mechanisms described in this study are likely to be relevant during conditions of acute inflammation, where PMNs are recruited and activated.

Peroxidase-catalyzed oxidation of NO2- or oxidation of NO2- by oxidants such as HOCl may result in underestimation of NO· production under inflammatory conditions, when NO2- is measured to quantitate NO· production. These oxidation mechanisms may significantly contribute to NO2- oxidation under inflammatory conditions when MPO and/or EPO are secreted. Interestingly, it has recently been demonstrated that the H2O2-scavenging activity of respiratory tract mucus is predominantly due to the presence of a peroxidase similar to LPO (75), which in the presence of sufficient concentrations of NO2- could lead to production of reactive nitrogen intermediates such as NO2· in the respiratory tract. Similarly, in situations when SCN- concentrations are sufficiently low, NO2- could be an important substrate for peroxidases in the oral cavity, especially when salivary concentrations of NO2- are elevated after ingestion of NO3-.

As NO2· is a powerfully oxidizing species (27), peroxidase-catalyzed NO2- oxidation could induce biomolecular modifications in host tissues and contribute to cell or tissue injury as a result of increased NO· production. Moreover, NO2- could also contribute to tissue injury by catalyzing MPO-mediated oxidation and chlorination reactions. Alternatively, NO2- has also been demonstrated to increase the bactericidal activity of MPO (28), and peroxidase-catalyzed NO2- oxidation could therefore also represent an additional host defense mechanism.

In conclusion, we have demonstrated that NO2- is a potential physiological substrate for heme peroxidases such as MPO and LPO, even in the presence of physiological concentrations of the alternative substrates Cl- or SCN-, and is able to catalyze peroxidase-mediated oxidation and chlorination of biological targets. Moreover, peroxidase-catalyzed oxidation of NO2- results in formation of NO2· or a related species, which can contribute to tyrosine nitration and could be involved in cell and tissue injury during situations of increased NO· production. This potential contributing role of NO2- to NO· biochemistry has been relatively overlooked, and further studies are needed to evaluate its physiological and/or pathological importance.


FOOTNOTES

*   This work was supported in part by National Institutes of Health Grant HL47628 and the Cystic Fibrosis Foundation. This work was done during the tenure of a research fellowship from the American Heart Association, California Affiliate (to A. v. d. V.), and a Research Training Fellowship from the American Lung Association, California Affiliate (to J. P. E.).The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
   A Parker B. Francis Fellow in Pulmonary Research. To whom correspondence should be addressed: University of California, Davis, Dept. of Internal Medicine, Pulmonary Research Laboratory, c/o Med. Sponsored Programs, TB 150, Davis, CA 95616. Tel.: 916-752-6305; Fax: 916-752-4374; E-mail: avandervliet{at}ucdavis.edu.
1   The abbreviations used are: NO·, nitric oxide; Obardot 2, superoxide anion; HbO2, oxyhemoglobin; HPA, 4-hydroxyphenylacetic acid; 3-NO2-HPA, 4-hydroxy-3-nitrophenylacetic acid; 3-Cl-HPA, 4-hydroxy-3-chlorophenylacetic acid; 3,3'-diHPA, 3,3'-bis(4-hydroxyphenylacetic acid; DTNB, 5,5'-dithiobis(2-nitrobenzoic acid; DTPA, diethylenetriaminepentaacetic acid; HRP, horseradish peroxidase; MCD, monochlorodimedon; PMN, polymorphonuclear neutrophils; HPLC, high performance liquid chromatography; MPO, myeloperoxidase; LPO, lactoperoxidase; EPO, eosinophil peroxidase; TNB, 5-thio-2-nitrobenzoic acid.

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