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Originally published In Press as doi:10.1074/jbc.M411905200 on January 28, 2005

J. Biol. Chem., Vol. 280, Issue 17, 16594-16600, April 29, 2005
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Xanthine Oxidase Catalyzes Anaerobic Transformation of Organic Nitrates to Nitric Oxide and Nitrosothiols

CHARACTERIZATION OF THIS MECHANISM AND THE LINK BETWEEN ORGANIC NITRATE AND GUANYLYL CYCLASE ACTIVATION*

Haitao Li{ddagger}, Hongmei Cui, Xiaoping Liu§, and Jay L. Zweier

From the Center for Biomedical Electron Paramagnetic Resonance Spectroscopy and Imaging, Davis Heart and Lung Research Institute and Division of Cardiovascular Medicine, Department of Internal Medicine, The Ohio State University College of Medicine, Columbus, Ohio 43210

Received for publication, October 20, 2004 , and in revised form, January 4, 2005.


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Organic nitrates have been used clinically in the treatment of ischemic heart disease for more than a century. Recently, xanthine oxidase (XO) has been reported to catalyze organic nitrate reduction under anaerobic conditions, but questions remain regarding the initial precursor of nitric oxide (NO) and the link of organic nitrate to the activation of soluble guanylyl cyclase (sGC). To characterize the mechanism of XO-mediated biotransformation of organic nitrate, studies using electron paramagnetic resonance spectroscopy, chemiluminescence NO analyzer, NO electrode, and immunoassay were performed. The XO reducing substrates xanthine, NADH, and 2,3-dihydroxybenz-aldehyde triggered the reduction of organic nitrate to nitrite anion (). Studies of the pH dependence of nitrite formation indicated that XO-mediated organic nitrate reduction occurred via an acid-catalyzed mechanism. In the absence of thiols or ascorbate, no NO generation was detected from XO-mediated organic nitrate reduction; however, addition of L-cysteine or ascorbate triggered prominent NO generation. Studies suggested that organic nitrite (R-O-NO) is produced from XO-mediated organic nitrate reduction. Further reaction of organic nitrite with thiols or ascorbate leads to the generation of NO or nitrosothiols and thus stimulates the activation of sGC. Only flavin site XO inhibitors such as diphenyleneiodonium inhibited XO-mediated organic nitrate reduction and sGC activation, indicating that organic nitrate reduction occurs at the flavin site. Thus, organic nitrite is the initial product in the process of XO-mediated organic nitrate biotransformation and is the precursor of NO and nitrosothiols, serving as the link between organic nitrate and sGC activation.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Organic nitrates such as glyceryl trinitrate (GTN)1 and isosorbide dinitrate (ISDN) have been used clinically in the treatment of cardiovascular disease for more than a century. The beneficial effects of organic nitrates result primarily from their vasodilator properties, which have been attributed to their metabolic product, nitric oxide (NO) (1–4). In 1998, the Nobel Prize was awarded for the discovery of "nitric oxide as a signaling molecule in the cardiovascular system." It is well known that organic nitrates require enzymatic metabolism to generate bioactive NO; however, the molecular mechanisms of their biotransformation have not been fully elucidated.

It has been reported that flavoenzymes play important roles in the bioactivation of organic nitrates (5–9), and the flavin inhibitor diphenyleneiodonium chloride (DPI) has shown strong inhibition of the bioactivation of organic nitrate (6, 10). Xanthine oxidase (XO) is a ubiquitous flavoenzyme in mammalian cells. XO plays a variety of important roles in normal physiology and disease, and it is the only identified enzyme that can mediate organic nitrate and inorganic nitrate/nitrite reduction (11–14). It has been reported that XO can catalyze organic nitrate reduction to nitrite anion (NO–2) under anaerobic conditions (13), but questions remain concerning the initial product, the precursor of NO, and the link of organic nitrate to the activation of soluble guanylyl cyclase (sGC). Thus, there is a need to characterize the process of XO-mediated organic nitrate bioactivation, and this should provide broader insights regarding bioactivation of organic nitrates by other enzymes as well.

Normally, NO–2 has been assumed to be the initial product of organic nitrate metabolism and the precursor of NO that accounts for organic nitrate bioactivity. However, controversies remain whether inorganic nitrite is an active intermediate in the vascular metabolism of organic nitrate to form NO (15–20). It has been questioned how the low micromolar to nanomolar levels of GTN in tissues can produce sufficient inorganic nitrite to serve as an intermediate to generate the levels of NO required to induce significant vasodilatory effects (21). Additional studies have shown that the magnitude and time course of GTN-mediated NO–2 formation in tissues make it impossible for GTN to act via liberation of nitrite (15, 19, 22–24). Thus, the link between organic nitrate and sGC activation remains elusive and is the most puzzling issue regarding the mechanism of organic nitrate biotransformation.

The literature is replete with conflicting observations concerning the role of thiols in the biotransformation of organic nitrate. Previous studies show that the bioactivation of organic nitrate involves thiols or sulfhydryl-containing compounds, and the repeated administration of organic nitrate causes sulfhydryl depletion and consequent tolerance to further vasodilation (25–28). However, other investigators have shown that there is no correlation between the concentration of endogenous thiols and the state of tolerance (29, 30). Therefore, the molecular mechanism of the interaction between organic nitrate and thiols to generate NO and S-nitrosothiol remains unclear.

To characterize the mechanism of XO-catalyzed organic nitrate reduction and elucidate the precise molecular mechanism of organic nitrate bioactivation in biological systems, electron paramagnetic resonance (EPR) spectroscopy, chemiluminescence NO analyzer, NO electrode, and immunoassay studies were performed under anaerobic conditions to characterize the initial product, the precursor of NO and nitrosothiol, and to determine the link between organic nitrate and sGC activation. These results demonstrate that XO catalyzes the bioactivation of organic nitrate to form NO and nitrosothiols in a thiol-dependent reaction.


   EXPERIMENTAL PROCEDURES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
RESULTS
 DISCUSSION
 REFERENCES
 
Materials—Xanthine oxidase from buttermilk (xanthine: oxygen oxidoreductase; EC 1.1.3.22 [EC] ), xanthine, oxypurinol, DPI, sodium nitrite, Griess Reagent, {beta}-NADH, bovine serum albumin (BSA), and 2,3-dihydroxybenz-aldehyde (DBA) were obtained from Sigma. Soluble guanylyl cyclase was obtained from Alexis Biochemical Corp. (San Diego, CA). Direct cGMP assay kit was obtained from Assay Designs, Inc. (Ann Arbor, MI), and cGMP production was quantified by immunoassay according to the protocol provided by the company. GTN was synthesized as described previously (9). N-Methyl-D-glucamine dithiocarbamate (MGD) was synthesized using carbon disulfide and N-methyl-D-glucamine (31, 32). Ferrous ammonium sulfate was purchased from Aldrich Chemical Co. (99.997%). Dulbecco's phosphate-buffered saline (PBS) was obtained from Invitrogen. Millipore ultra free centrifugal filter (nominal molecular weight limit 10,000) was obtained from Fisher.

EPR Spectroscopy—EPR measurements were performed using a Bruker EMX spectrometer with HS resonator operating at X-band. Measurements were performed at ambient temperature with a modulation frequency of 100 kHz, modulation amplitude of 2.5 G, and a microwave power of 20 milliwatts. NO generated by XO-catalyzed reduction of organic nitrate was purged out using argon to a vessel that contained the spin-trap Fe2+-MGD, as described previously (14). This setup was designed to isolate the reaction solution from spin trap and thus avoid any possible perturbation caused by the reaction of (MGD)2-Fe2+ with nitrite or with the enzyme. Fe2+-MGD complexes were prepared by adding solid ferrous ammonium sulfate and MGD (molar ratio, 1:5) to the deoxygenated (argon-purged) solution with a final concentration of 2 mM iron. Quantitation of NO formation and trapping were performed by double integration of observed EPR signal with comparison to that from the similar aqueous NO-Fe-MGD standard (14).

Chemiluminescence Measurements—The rate of the NO production was measured using a Sievers 270B nitric oxide analyzer interfaced through a DT2821 A to D board to a personal computer. In the analyzer, NO is reacted with ozone forming excited-state NO2, which emits light. Mixing of reagents and separation of NO from the reaction mixture were done at controlled temperature in a glass-purging vessel equipped with heating jacket. An ice-water cooling condenser was attached to the top of the vessel to reduce the outflow of vapors during purging. The release of NO was quantified by analysis of the digitally recorded signal from the photomultiplier tube using specially designed data acquisition and analysis software developed in our laboratory (33). After an initial 60 s of equilibration of flow from the purging vessel to the detector, the signal provides the rate of NO formation over time.

As described previously, nitrite in the solution is measured from its reduction to NO under conditions of acidic pH in the presence of glacial acetic acid and 1% KI (12). This method has been shown to be highly sensitive, enabling detection of nitrite down to picomole amounts (33). Neither GTN nor ISDN was found to generate any detectable NO under these conditions. XO-dependent nitrite generation was calculated by subtracting nitrite production without addition of XO (control). Calibration of the magnitude of nitrite and NO production was determined from the integral of the signal over time compared with that from nitrite concentration standards (12, 34).

Electrochemical Measurements—Electrochemical measurements of NO generation by XO under anaerobic conditions were carried out at 37 °C in a sealed electrochemical vial using a CHI 832 electrochemical detector (CH Instruments, Inc., Cordova, IN) and a Clark-type NO electrode (ISO-NOP; Word Precision Instruments, Sarasota, FL). A slow flow of argon gas was maintained in the space above the solution. The electrochemical detector continuously recorded the current through the working electrode, which is proportional to the NO concentration in the solution. The sensor was calibrated before and after experiments with known concentrations of NO, using NO equilibrated solutions.

Assay of S-Nitrosothiol—The concentrations of S-nitrosothiol were determined from the increase of nitrite concentration after treatment of samples with HgCl2. Spectrophotometric quantitation of nitrite with Griess Reagent was performed according to the protocol provided by Sigma, using a Varian Cary 300 UV-visible spectrophotometer. One milliliter of sample was supplemented with 50 µl of 10 mM HgCl2 and 1 ml of Griess Reagent for 10 min at room temperature. Background levels of nitrite in the samples were determined by using Griess Reagent without addition of HgCl2. The standard curve was obtained with a known amount of nitrite.

Immunoassay of Guanylyl Cyclase Activity—Activation of soluble sGC by XO-mediated organic nitrate reduction was investigated by employing enzyme-linked immunoassay. After incubation of XO (0.04 mg/ml) and GTN (10 µM)/or ISDN (100 µM) with 1 ml of reaction buffer (10 ng of sGC, 10% BSA, 5 mM EDTA, 2 mM MgCl2, 1 mM GTP, and 1 mM NADH/or 20 µM xanthine in 1 ml of PBS) at 37 °C under anaerobic conditions, the enzymes in the samples were removed using Millipore filter by centrifugation at 2000 x g for 10 min at 4 °C. The measurements of cGMP in the solution were performed using direct cGMP assay kit by immunoassay according to the manufacturer's product protocol (Assay Designs, Inc.). The standard curve was obtained with known amounts of cGMP.

Statistical Analysis—Values are expressed as the mean ± S.D. of at least three repeated measurements, and statistical significance of difference was evaluated by Student's t test. A p value of ≤0.05 was considered to indicate statistical significance.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
DISCUSSION
 REFERENCES
 
Inorganic Nitrite Generation from XO-mediated Organic Nitrate Reduction—It has been reported that XO can catalyze organic nitrate reduction to NO–2 (13). To investigate the mechanism of NO–2 generation, as well as its possible role as an intermediate of NO in the process of XO-mediated organic nitrate reduction, the rate of nitrite formation derived from XO-catalyzed GTN and ISDN reduction was measured under anaerobic conditions (Fig. 1, A and B, respectively). In the presence of GTN (10 µM) or ISDN (100 µM) and reducing substrates DBA (0.5 mM), xanthine (20 µM), or NADH (1.0 mM), after addition of XO, the reaction mixture was sampled every 2 min, and its nitrite concentration was determined by NO analyzer with reduction of nitrite to NO using 1% KI under acidic conditions. With FAD site-binding substrate NADH as an electron donor, nitrite concentrations in the reaction mixture linearly increase for the first 10 min and then gradually plateau by 30 min (Fig. 1, c). With xanthine or DBA that provides electrons to XO at the molybdenum site, the increase in nitrite concentrations plateaus within 20 min, and a quicker decrease in nitrite generation rates is seen (Fig. 1, a and b).



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  		FIG. 1.
Time course of nitrite generation from XO-catalyzed GTN or ISDN reduction. Measurements were performed under anaerobic conditions at 37 °C in PBS, pH 7.4, in the presence of XO (0.04 mg/ml) and (A) GTN (10 µM) with (a) xanthine (20 µM), (b) DBA (0.5 mM), or (c) NADH (1 mM) or (B) ISDN (100 µM) with (a) xanthine (20 µM), (b) DBA (0.5 mM), or (c) NADH (1 mM). The reaction mixture was sampled 0.1 ml every 2 min, and nitrite concentration was measured by NO analyzer as described under "Experimental Procedures."

 
For xanthine as reducing substrate, the initial rate of XO-mediated nitrite formation was determined as a function of GTN or ISDN concentration, and Michaelis-Menten kinetics were observed with correlation coefficient {gamma}2 > 99% (Fig. 2, A and B). The apparent Km and Vmax values are shown inside each curve. The apparent Km values of GTN (0.49 mM) and ISDN (1.64 mM) are far higher than the clinical levels of organic nitrates in tissues or blood. Thus, at pharmacological levels in the range of 10–100 µM, the rate of XO-mediated organic nitrate reduction would be expected to increase linearly with the given organic nitrate dose. From this kinetic data, it is possible to predict the magnitude of XO-mediated nitrite formation as a function of organic nitrate and to determine the quantitative importance of this mechanism of nitrite generation and its subsequent NO generation in a given biological system where these substrate levels are known.



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  		FIG. 2.
Kinetics of XO-mediated nitrite generation as a function of GTN or ISDN concentration using xanthine as reducing substrate. Measurements were performed under anaerobic conditions at 37 °C in PBS, pH 7.4, using 0.04 mg/ml XO and 20 µM xanthine. A shows the initial rate of nitrite generation in the presence of GTN (0.01–3 mM); B shows the initial rate of nitrite generation in the presence of ISDN (0.1–10 mM). For each graph, the corresponding fitting (solid lines), Km, and Vmax were obtained using data the Michaelis-Menten equation.

 
Under ischemic conditions, in addition to marked hypoxia, marked intracellular acidosis also occurs, and pH values in tissues, such as the heart, can fall to levels of 6.0 or below (35). In order to assess the nitrite formation under different physiological or pathological conditions and to further characterize the mechanism of XO-catalyzed organic nitrate reduction, experiments were performed to measure the effects of different pH values on the magnitude of nitrite generation. Because the apparent Km of organic nitrates is far higher than the pharmacological levels of organic nitrates in tissues or blood, we evaluated the effect of pH on XO-mediated organic nitrate reduction with a typical pharmacological level of 10 µM GTN or 100 µM ISDN. As shown in Table I, it was observed for each of the three reducing substrates (xanthine, NADH, and DBA) that under acidic conditions, increased XO-mediated nitrite generation occurs. In contrast, under alkaline conditions, decreased nitrite generation was seen.


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  		TABLE I
Effect of pH on nitrite generation rate

The initial nitrite generation rates were calculated as described in Fig. 1. Measurements were performed with XO (0.04 mg/ml) in the presence of GTN (10 µM) or ISDN (100 µM) with reducing substrate.

 
Nitric Oxide Generation from XO-mediated GTN or ISDN Reduction—It is well known that organic nitrates such as GTN and ISDN are prodrugs requiring metabolism to generate bioactive NO. In order to investigate whether XO can directly catalyze the reduction of GTN and ISDN to NO and quantitate the rates of NO generation, EPR spectroscopy was applied to measure GTN- and ISDN-mediated NO generation under anaerobic conditions. NO generated by organic nitrate was purged out using argon to a purging vessel that contained the spin-trap Fe2+-MGD. NO is paramagnetic and binds with high affinity to the water-soluble spin trap Fe2+-MGD forming the mononitrosyl iron complex NO-Fe2+-MGD with characteristic triplet spectrum at g = 2.04 and hyperfine splitting aN = 12.8. From the intensity of the observed spectrum, quantitative measurement of NO generation can be performed (36–38). With ISDN (100 µM) or GTN (10 µM), in the presence of XO (0.04 mg/ml) and NADH (1 mM), no signal was seen (Fig. 3, spectra A and B). This demonstrates that no significant NO formation occurs.



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  		FIG. 3.
EPR measurement of NO generated from XO under anaerobic conditions. XO (0.04 mg/ml) and 1 mM NADH were incubated in PBS buffer with 10% BSA, pH 7.4, at 37 °C with the following: spectrum A, ISDN (100 µM); spectrum B, GTN (10 µM); spectrum C, ISDN (100 µM) and 5 mM L-cysteine; spectrum D, GTN (10 µM) and L-cysteine (5 mM); and spectrum E, ISDN (100 µM), L-cysteine (5 mM), and DPI (100 µM). NO was continuously purged using argon into a trap vessel containing 5 ml of 2 mM (MGD)2-Fe2+. Samples were taken from the trap vessel after 30 min, and spectra of (MGD)2-Fe2+-NO adducts formed are shown.

 
It has been reported that sulfhydryl (–SH) compounds are needed for GTN activation and that repeated administration of GTN causes sulfhydryl depletion and consequent tolerance to further vasodilatation (25–28). To test whether sulfhydryl compounds play a role in XO-mediated transformation of organic nitrates, L-cysteine (5 mM) was added, and this triggered marked NO generation from 100 µM ISDN with 0.28 µM NO trapped over 30 min (Fig. 3, spectrum C) and from 10 µM GTN with 0.22 µM NO trapped over 30 min (Fig. 3, spectrum D). The XO molybdenum site inhibitor oxypurinol (100 µM) could not inhibit this NO generation from GTN or ISDN (data not shown); however, the FAD site inhibitor DPI (100 µM) effectively blocked XO-dependent NO formation (Fig. 3, spectrum E). This suggests that organic nitrate reduction takes place at the FAD site of XO.

In order to further investigate XO-mediated GTN and ISDN reduction and quantitate the rates of NO generation, studies were performed using a chemiluminescence NO analyzer. NO was purged from the solution by argon and then reacted with ozone in the analyzer to form an excited-state NO2, which emits light. This method provides direct measurement of the rate of NO generation as a function of time. With GTN (10 µM), in the presence of xanthine (20 µM) or NADH (1 mM) with XO (0.04 mg/ml), no measurable rate of NO formation was observed (Fig. 4, trace D). To test the effect of sulfhydryl compounds on GTN reduction, L-cysteine (5 mM) was added, and prominent NO generation was triggered (Fig. 4, traces A and B). In the absence of XO, cysteine alone reacted with GTN to produce NO, but only trace production was seen (Fig. 4, trace C).



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  		FIG. 4.
Measurement of the rate of NO generation from XO catalyzed GTN reduction. Measurement was performed using a chemiluminescence NO analyzer under anaerobic conditions at 37 °C in PBS (10% BSA), pH 7.4. Trace A shows the rate of NO generation by XO (0.04 mg/ml), xanthine (20 µM), GTN (10 µM), and L-cysteine (5 mM). Trace B shows the rate of NO generation by XO (0.04 mg/ml), NADH (1 mM), GTN (10 µM), and L-cysteine (5 mM). Trace C shows the rate of NO generation by GTN (10 µM) and L-cysteine (5 mM). Trace D shows the rate of NO generation by XO (0.04 mg/ml), NADH (1 mM), and GTN (10 µM).

 
XO-mediated ISDN reduction was similar to that of GTN reduction. In the presence of XO (0.04 mg/ml), ISDN (0.1 mM), and xanthine or NADH as reducing substrate, no detectable NO was generated (Fig. 5, trace C). However, after addition of L-cysteine, prominent NO generation was seen (Fig. 5, traces A and B). With ISDN and cysteine alone, no detectable NO production was seen (data not shown).



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  		FIG. 5.
Measurement of the rate of NO generation from XO-catalyzed ISDN reduction. Measurements were performed using a chemiluminescence NO analyzer under anaerobic conditions at 37 °C with XO (0.04 mg/ml) in PBS (10% BSA), pH 7.4. Trace A shows the rate of NO generation with xanthine (20 µM), ISDN (100 µM), and L-cysteine (5 mM). Trace B shows the rate of NO generation with NADH (1 mM), ISDN (100 µM), and L-cysteine (5 mM). Trace C shows the rate of NO generation with xanthine (20 µM) and ISDN (100 µM).

 
To further confirm these observations and quantitate the NO concentration that accumulates, NO generation was measured by electrochemical detection. Prior to the addition of XO, no detectable NO was seen. After the addition of XO (0.04 mg/ml), NO was triggered from L-cysteine (5 mM) with ISDN (1 mM or 0.1 mM) and NADH (1 mM) in the presence of L-cysteine (5 mM) (Fig. 6, traces A and B), whereas no NO generation was observed without L-cysteine (Fig. 6, trace C).



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  		FIG. 6.
Electrochemical measurement of NO generation from XO-mediated ISDN reduction. The time course of NO generation was measured using an electrochemical NO sensor under anaerobic conditions at 37 °C in PBS (10% BSA), pH 7.4. The arrow shows the time at which XO (0.04 mg/ml) was added. Trace A shows the data from ISDN (1 mM) in the presence of NADH (1 mM) and L-cysteine (5 mM); trace B shows the data from ISDN (100 µM) in the presence of NADH (1 mM) and L-cysteine (5 mM); trace C shows the data from ISDN (1 mM) in the presence of NADH (1 mM).

 
It is commonly known that the combination of ascorbate with GTN or ISDN reduces the occurrence of nitrate tolerance. To investigate the role of ascorbate in the activation of organic nitrate, NO generation from XO-mediated ISDN reduction was detected with or without ascorbate. Without XO, no detectable NO was generated with ascorbate (1 mM), ISDN (0.1 mM), and xanthine (20 µM) or NADH (1 mM) (Fig. 7, trace C). However, upon addition of XO (0.04 mg/ml), prominent NO generation was triggered from ISDN (100 µM) using xanthine or NADH as reducing substrates (Fig. 7, traces A and B).



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  		FIG. 7.
Measurement of the rate of NO generation from XO-catalyzed ISDN reduction in the presence of ascorbate. Measurements were performed using a chemiluminescence NO analyzer under anaerobic conditions at 37 °C in PBS (10% BSA), pH 7.4. Trace A shows the rate of NO generation by XO (0.04 mg/ml), xanthine (20 µM), ISDN (100 µM), and ascorbate (1 mM). Trace B shows the rate of NO generation by XO (0.04 mg/ml), NADH (1 mM), ISDN (100 µM), and ascorbate (1 mM). Trace C shows the rate of NO generation by ascorbate (1 mM), xanthine (20 µM), and ISDN (100 µM).

 
Assay of Nitrosothiol Formation from XO-mediated GTN or ISDN Reduction—It has been reported that thiols such as glutathione (GSH) react with labile organic nitrite esters, as could be formed from reduction of organic nitrates, to give rise to corresponding S-nitrosothiols either chemically or enzymatically, and these S-nitrosothiols, in turn, can serve as a precursor for NO formation (39–43). To investigate whether the initial product of XO-mediated reduction of organic nitrate is organic nitrite (R-O-NO) and whether this organic nitrite can be the nitrosothiol precursor, studies were performed to measure the S-nitrosoglutathione (GSNO) production from XO-catalyzed organic nitrate reduction in the presence of GSH (Fig. 8). Following incubation at 37 °C under anaerobic conditions (argon), the reaction mixture was sampled at the time indicated, and the detection and quantification of nitrosothiols in the solution were performed using the Saville Assay, which is based on mercury ion-mediated heterolysis cleavage of the S-NO bond. Nitrosonium ion is released, which is in turn measured by the Griess Reaction. The concentrations of GSNO were determined from the increase in the nitrite concentration induced by treatment of samples with HgCl2. Without XO, GTN (100 µM) or ISDN (1 mM) chemically react with GSH (5 mM) with formation of low levels of GSNO of 0.21 µM (Fig. 8A, d) or 0.11 µM (Fig. 8B, d), respectively, after a 30-min incubation. With the addition of XO (0.04 mg/ml), GSNO production from GTN quickly increased to 1.38 µM within 20 min and then gradually plateaued (Fig. 8A, a). GSNO production from ISDN and XO showed similar kinetics with 2.05 µM GSNO production after 20 min. Using the molybdenum site inhibitor oxypurinol, XO-dependent nitrosothiol production was not inhibited (Fig. 8, b). When the FAD site inhibitor DPI (100 µM) was used, it completely inhibited XO-dependent nitrosothiol production (Fig. 8, c). This suggests that organic nitrate reduction takes place at the FAD site of XO.



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  		FIG. 8.
Nitrosothiol production from XO-mediated GTN or ISDN reduction. GTN (A; 100 µM) or ISDN (B; 1 mM) was incubated in PBS (10% BSA), pH 7.4, under anaerobic conditions at 37 °C with the following: a, NADH (1 mM), GSH (5 mM), and XO (0.04 mg/ml); b, NADH (1 mM), GSH (5 mM), and oxypurinol; c, NADH (1 mM), GSH (5 mM), XO (0.04 mg/ml), and DPI (100 µM); or d, NADH (1 mM), GSH (5 mM). The reaction mixture was sampled at 10, 15, 20, 25, and 30 min. Detection and quantification of nitrosothiols were performed as described under "Experimental Procedures."

 
Effect of XO-mediated GTN or ISDN Reduction on sGC Activity—In order to determine whether XO-mediated organic nitrate biotransformation can induce sGC activation, enzyme-linked immunoassay was performed to determine the cGMP concentrations in the reaction mixture. After incubation of XO (0.04 mg/ml) with GTN (10 µM) or ISDN (100 µM) in reaction buffer (10 ng of sGC, 10% BSA, 5 mM EDTA, 2 mM MgCl2, 1 mM GTP, and 1 mM NADH or 20 µM xanthine in 1 ml of PBS) under anaerobic conditions for 10 min, measurements of the sGC product cGMP were performed. Without L-cysteine, with NADH or xanthine as XO-reducing substrates, no cGMP could be detected, suggesting that sGC was not activated (Fig. 9, A). However, for similar conditions with the addition of L-cysteine (5 mM), a significant amount of cGMP production was generated from XO-mediated reduction of GTN (Fig. 9, B) or ISDN (Fig. 9, C). Using the molybdenum site-specific inhibitor oxypurinol, XO-mediated sGC activation by organic nitrate was largely inhibited (>95%) when xanthine was used as the reducing substrate, but not when NADH was used (Fig. 9, D). When the FAD site inhibitor DPI was used, it inhibited XO-mediated sGC activation by GTN regardless of the type of reducing substrate present (Fig. 9, E), further confirming that organic nitrates are reduced at the FAD site.



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  		FIG. 9.
Soluble guanylyl cyclase activation by XO-mediated GTN or ISDN reduction. Activation of sGC was determined from the measurements of cGMP. The reaction buffer is PBS (1 ml; pH 7.4) with 10% BSA, EDTA (5 mM), MgCl2 (2 mM), sGC (10 ng), and GTP (1 mM). XO (0.04 mg/ml) with NADH (1 mM) or xanthine (20 µM) as reducing substrate was incubated for 10 min in the reaction buffer in the presence of the following: A, GTN (10 µM); B, GTN (10 µM) and L-cysteine (5 mM); C, ISDN (100 µM) and 5 mM L-cysteine; D, GTN (10 µM), 5 mM L-cysteine, and oxypurinol (100 µM); and E, GTN (10 µM), 5 mM L-cysteine, and DPI (100 µM).

 

   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
REFERENCES
 
Organic nitrates (R-O-NO2) such as GTN and ISDN have been widely used clinically in the treatment of myocardial ischemia. Over the last decade, their efficacy has been attributed to their metabolic conversion to NO. However, the molecular mechanism of this organic nitrate biotransformation has not been fully defined. Questions remain concerning the initial product, the precursor of NO, and the link of organic nitrate to the activation of sGC. There is further uncertainty regarding how this process occurs under the markedly hypoxic conditions in ischemic tissues. Recently, XO has been reported to be able to mediate organic nitrate and inorganic nitrate/nitrite reduction under anaerobic conditions (11–13). To investigate the mechanism of XO-mediated organic nitrate reduction and thus further elucidate the precise molecular mechanism of bioactivation, we performed a series of studies using EPR spectroscopy, chemiluminescence NO analyzer, immunoassay, and NO electrode techniques to measure the magnitude of various products (NO–2, NO, and R-S-NO) derived from XO and their effects on activation of sGC.

It was observed that each of the typical XO reducing substrates xanthine, DBA, and NADH could act as electron donors to support this XO-mediated GTN/ISDN reduction (Figs. 1 and 2). However, the time-dependent nitrite concentrations in the reaction mixture showed progressively decreased nitrite generation rates (Figs. 1 and 2). This is consistent with the previous report of suicide inhibition in the process of XO-mediated GTN/ISDN reduction (13).

For xanthine as a reducing substrate, the rate of XO-mediated nitrite formation was determined as a function of GTN or ISDN concentrations, and Michaelis-Menten kinetics were observed (Fig. 2, A and B). From these kinetic data, comparing the kinetics of XO-mediated inorganic nitrate (NO–3) reduction (12), with the same substrate levels, nitrite formation rates are about 5 times higher for ISDN reduction and about 30 times higher for GTN reduction than from nitrate anion (NO–3) reduction. Similar to NO–3 reduction (12), XO-mediated reduction of organic nitrates also takes place via an acid-catalyzed mechanism (Table I).

Traditionally, it has been assumed that NO–2 production is the first step in the process of organic nitrate biotransformation and that it is the precursor of NO or nitrosothiol. It has been assumed that organic nitrates are converted to NO–2 by reaction with sulfhydryls (–SH); nitrite then liberates NO via nitrous acid, and NO combines with thiol to generate a nitrosothiol that activates sGC (3).

(REACTIONS 1-5)

However, controversy remains regarding whether this nitrite can be an active intermediate in the vascular metabolism of organic nitrates to form NO (15–20). Normally, the concentrations of NO–2 in the serum or tissues range from 1 to 10 µM, and it has been questioned how micromolar to nanomolar levels of GTN, the clinically relevant concentrations, can produce sufficient NO–2 to serve as an intermediate for the generation of sufficient NO to exert vasodilatory effects (21). Studies in whole tissue or in isolated mitochondria have shown that clinically used GTN levels cannot produce sufficient NO–2 as intermediate of NO to exhibit vasodilatory effects (15, 19, 22–24). However, long term high dose administration of organic nitrates can highly elevate nitrate anion (NO–3) or NO–2 concentrations in tissues (44), and these can be important sources of NO during ischemia (11, 12, 14, 45–47).

It is well known that sulfhydryl (–SH) compounds are needed in GTN activation and that the repeated administration of GTN causes sulfhydryl depletion and consequent tolerance to further vasodilatation. In this study, it was shown that the presence of L-cysteine or ascorbate triggered significant NO generation from XO-mediated reduction of GTN/ISDN, whereas no detectable NO was generated without the addition of thiols or ascorbate (Figs. 3, 4, 5, 6, 7). Of note, at pH 7.4, the presence of sulfhydryls or ascorbate does not increase NO generation from XO-mediated NO–2 reduction. All these experimental results suggest that NO–2 is not the critical intermediate of NO in the process of XO-mediated reduction of GTN or ISDN.

It has been reported that thiols such as GSH can react with organic nitrites to form the corresponding S-nitrosothiols either chemically or enzymatically, and this S-nitrosothiol, in turn, can serve as a precursor to NO formation (39–43). In this study, besides observation of NO generation in the process of XO-mediated organic nitrate reduction, GSNO production was also observed in the presence of GSH. This XO-dependent GSNO production was inhibited in the presence of the FAD site inhibitor DPI (Fig. 8). All these results suggest that organic nitrite (R-O-NO) is the initial product in the process of XO-mediated organic nitrate reduction and the precursor of NO and nitrosothiol. The presence of L-cysteine or ascorbate can reduce organic nitrite further, triggering significant NO generation. Of note, even with this competitive reduction, the maximum rate of NO production is an order of magnitude lower than the rate of nitrite production from organic nitrite hydrolysis (Figs. 3, 4, 5, 6, 7). This suggests that under hydrophilic conditions, most of the organic nitrite (R-O-NO) will quickly hydrolyze to produce NO–2 (Figs. 1 and 2). However, under hydrophobic conditions, NO and nitrosothiols could be major products of the reaction of organic nitrite (R-O-NO) with sulfhydryl compounds either chemically (39) or enzymatically as catalyzed by glutathione S-transferase (GST) (41, 42, 48).

Activation of sGC by XO-mediated organic nitrate reduction was investigated by employing enzyme-linked immunoassay. Our results showed that sulfhydryl compounds or ascorbate is needed for the activation of sGC. Without sulfhydryl compounds or ascorbate, sGC is not activated by GTN or ISDN (Fig. 9). The flavin-binding XO inhibitor DPI inhibited XO-mediated organic nitrate reduction and sGC activation, indicating that organic nitrate reduction occurs at the flavin site.

The following reactions define the steps in the reaction mechanism of XO-mediated biotransformation of organic nitrate:

(REACTIONS 6-9)

Eox is the oxidized enzyme, and Ered is the reduced enzyme. Organic nitrite (R-O-NO) is the initial product in the process of organic nitrate biotransformation and the precursor of NO and nitrosothiol. Thus, organic nitrite (R-O-NO) is the link between organic nitrate and activation of sGC.

Several enzyme systems are involved in the biotransformation of organic nitrate, including GST (49, 50), cytochrome P450 systems (6, 51, 52), xanthine oxidoreductase (13), and mitochondrial aldehyde dehydrogenase (53). These enzymes can be classified into two groups: (a) flavoenzymes, including xanthine oxidoreductase and cytochrome P450/P450 reductase systems; and (b) enzymes that have sulfhydryl (–SH) groups in the active site, such as GST and mitochondrial aldehyde dehydrogenase.

It has been demonstrated previously that the activity of XO in the postischemic rat heart is 16.8 millunits/g protein (36), which corresponds to 0.013 mg XO/g protein or ~3.4 µg/g cell water. The total XO and xanthine dehydrogenase activity, however, is 10-fold above this value. During ischemia, xanthine levels rise from near zero to values on the order of 10–100 µM. Tissue or blood sulfhydryl compound concentrations are typically in the range of 0.5–2 mM under normal physiological conditions (54, 55). Thus, pharmacological levels of GTN (10–25 µM) or ISDN (100–250 µM) in the heart could generate about 1–2 nM/s NO from xanthine oxidoreductase-mediated reduction under anaerobic conditions, which is comparable to maximal NO production from tissue NO synthases. In the rat liver, XO activity is about 10-fold above the levels in heart (56), and thus NO generation from organic nitrate in liver could be 10-fold higher.

Previous studies have shown that free flavin can mediate organic nitrate reduction. This study further demonstrated that the flavin site of XO catalyzes reduction of organic nitrate to organic nitrite. Our results suggest that multiple flavoenzymes may be able to similarly catalyze this reaction and may also be involved in the biotransformation of organic nitrates to form NO.

Besides flavoenzymes, GST and mitochondrial aldehyde dehydrogenase also have been reported to play important roles in organic nitrate bioactivation. Both GST and mitochondrial aldehyde dehydrogenase have sulfhydryl (–SH) groups in the active site. Previous studies have shown that nucleophilic attack by the active site sulfhydryl groups of GST or mitochondrial aldehyde dehydrogenase (mtALDH) on organic nitrate (R-O-NO2) can result in the formation of E-S-NO2 (57–59), whereas compared with organic nitrate (R-O-NO2), organic nitrite (R-O-NO) would be predicted to much more readily react with sulfhydryl compounds to form nitrosothiols or NO.

(REACTION 10)

These two enzymatic systems may synergistically act in the process of organic nitrate biotransformation. First, flavoenzymes would be required to catalyze the reduction of organic nitrate to produce organic nitrite; then enzymes such as GST and mitochondrial aldehyde dehydrogenase could further catalyze reaction of organic nitrite and thiols to produce nitrosothiols.

Overall, this study demonstrates that XO can reduce GTN and ISDN to their respective organic nitrites (R-O-NO). These organic nitrites (R-O-NO) then react chemically or enzymatically with thiols or other reducing substrates to form nitrosothiols or NO. Furthermore, the depletion of thiols or inhibition of crucial enzyme(s) may cause nitrate tolerance. Nitrite anion (NO–2) is the hydrolysis product of organic nitrite, which is a byproduct rather than an important intermediate in the process of organic nitrate bioactivation. Thus, organic nitrite (R-O-NO) is the link between organic nitrate and activation of sGC.


   FOOTNOTES
 
* This work was supported in part by National Institutes of Health Grants HL63744, HL65608, and HL38324. The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact. Back

§ Supported in part by an American Heart Association scientist development award. Back

{ddagger} To whom correspondence may be addressed: 110G Davis Heart and Lung Research Institute, 473 West 12th Ave., Columbus, OH 43210-1252. E-mail: li-13{at}medctr.osu.edu. ¶ To whom correspondence may be addressed: 110G Davis Heart and Lung Research Institute, 473 West 12th Ave., Columbus, OH 43210-1252. E-mail: zweier-1{at}medctr.osu.edu.

1 The abbreviations used are: GTN, glyceryl trinitrate; XO, xanthine oxidase; ISDN, isosorbide dinitrate; sGC, soluble guanylyl cyclase; DBA, 2,3-dihydroxybenz-aldehyde; NO, nitric oxide; BSA, bovine serum albumin; GSH, glutathione; GST, glutathione S-transferase; MGD, N-methyl-D-glucamine dithiocarbamate; EPR, electron paramagnetic resonance; DPI, diphenyleneiodonium chloride; PBS, phosphate-buffered saline; GSNO, S-nitrosoglutathione. Back


   ACKNOWLEDGMENTS
 
We thank Dr. Frederick Villamena for kind help with GTN purification and Dr. Alexandre Samouilov for helpful comments.



   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 

  1. Arnold, W. P., Mittal, C. K., Katsuki, S., and Murad, F. (1977) Proc. Natl. Acad. Sci. U. S. A. 74, 3203–3207[Abstract/Free Full Text]
  2. Murad, F., Mittal, C. K., Arnold, W. P., Katsuki, S., and Kimura, H. (1978) Adv. Cyclic Nucleotide Res. 9, 145–158[Medline] [Order article via Infotrieve]
  3. Ignarro, L. J., Lippton, H., Edwards, J. C., Baricos, W. H., Hyman, A. L., Kadowitz, P. J., and Gruetter, C. A. (1981) J. Pharmacol. Exp. Ther. 218, 739–749[Free Full Text]
  4. Mellion, B. T., Ignarro, L. J., Myers, C. B., Ohlstein, E. H., Ballot, B. A., Hyman, A. L., and Kadowitz, P. J. (1983) Mol. Pharmacol. 23, 653–664[Abstract]
  5. Blehert, D. S., Knoke, K. L., Fox, B. G., and Chambliss, G. H. (1997) J. Bacteriol. 179, 6912–6920[Abstract/Free Full Text]
  6. McGuire, J. J., Anderson, D. J., McDonald, B. J., Narayanasami, R., and Bennett, B. M. (1998) Biochem. Pharmacol. 56, 881–893[CrossRef][Medline] [Order article via Infotrieve]
  7. Mohazzab, H. K., Kaminski, P. M., Agarwal, R., and Wolin, M. S. (1999) Circ. Res. 84, 220–228[Abstract/Free Full Text]
  8. Ratz, J. D., McGuire, J. J., Anderson, D. J., and Bennett, B. M. (2000) J. Pharmacol. Exp. Ther. 293, 569–577[Abstract/Free Full Text]
  9. Meah, Y., Brown, B. J., Chakraborty, S., and Massey, V. (2001) Proc. Natl. Acad. Sci. U. S. A. 98, 8560–8565[Abstract/Free Full Text]
  10. McGuire, J. J., Anderson, D. J., and Bennett, B. M. (1994) J. Pharmacol. Exp. Ther. 271, 708–714[Abstract/Free Full Text]
  11. Li, H., Samouilov, A., Liu, X., and Zweier, J. L. (2001) J. Biol. Chem. 276, 24482–24489[Abstract/Free Full Text]
  12. Li, H., Samouilov, A., Liu, X., and Zweier, J. L. (2003) Biochemistry 42, 1150–1159[CrossRef][Medline] [Order article via Infotrieve]
  13. Doel, J. J., Godber, B. L., Eisenthal, R., and Harrison, R. (2001) Biochim. Biophys. Acta 1527, 81–87[Medline] [Order article via Infotrieve]
  14. Li, H., Samouilov, A., Liu, X., and Zweier, J. L. (2004) J. Biol. Chem. 279, 16939–16946[Abstract/Free Full Text]
  15. Kowaluk, E. A., Chung, S. J., and Fung, H. L. (1993) Drug Metab. Dispos. 21, 967–969[Medline] [Order article via Infotrieve]
  16. Walters, C. L., Casselden, R. J., and Taylor, A. M. (1967) Biochim. Biophys. Acta 143, 310–318[Medline] [Order article via Infotrieve]
  17. Kozlov, A. V., Staniek, K., and Nohl, H. (1999) FEBS Lett. 454, 127–130[CrossRef][Medline] [Order article via Infotrieve]
  18. Walters, C. L., and Taylor, A. M. (1965) Biochim. Biophys. Acta 96, 522–524[Medline] [Order article via Infotrieve]
  19. Nohl, H., Staniek, K., Sobhian, B., Bahrami, S., Redl, H., and Kozlov, A. V. (2000) Acta Biochim. Pol. 47, 913–921[Medline] [Order article via Infotrieve]
  20. Mayer, B. (2003) Angew Chem. Int. Ed. Engl. 42, 388–391[CrossRef][Medline] [Order article via Infotrieve]
  21. Kleschyov, A. L., Oelze, M., Daiber, A., Huang, Y., Mollnau, H., Schulz, E., Sydow, K., Fichtlscherer, B., Mulsch, A., and Munzel, T. (2003) Circ. Res. 93, e104–e112[Abstract/Free Full Text]
  22. Romanin, C., and Kukovetz, W. R. (1988) J. Mol. Cell. Cardiol. 20, 389–396[Medline] [Order article via Infotrieve]
  23. Sharpe, M. A., and Cooper, C. E. (1998) Biochem. J. 332, Pt 1, 9–19
  24. Burger, I. H., and Walters, C. L. (1971) Biochem. J. 123, 9P[Medline] [Order article via Infotrieve]
  25. Hutter, J., Schmidt, M., and Rittler, J. (1988) Eur. J. Pharmacol. 156, 215–222[Medline] [Order article via Infotrieve]
  26. Needleman, P., and Johnson, E. M., Jr. (1973) J. Pharmacol. Exp. Ther. 184, 709–715[Abstract/Free Full Text]
  27. Needleman, P., Jakschik, B., and Johnson, E. M., Jr. (1973) J. Pharmacol. Exp. Ther. 187, 324–331[Abstract/Free Full Text]
  28. Winniford, M. D., Kennedy, P. L., Wells, P. J., and Hillis, L. D. (1986) Circulation 73, 138–142[Abstract/Free Full Text]
  29. Gruetter, C. A., and Lemke, S. M. (1986) Can. J. Physiol. Pharmacol. 64, 1395–1401[Medline] [Order article via Infotrieve]
  30. Ignarro, L. J. (2002) Proc. Natl. Acad. Sci. U. S. A. 99, 7816–7817[Free Full Text]
  31. Shinobu, L. A., Jones, S. G., and Jones, M. M. (1984) Acta Pharmacol. Toxicol. 54, 189–194[Medline] [Order article via Infotrieve]
  32. Zweier, J. L., Wang, P., and Kuppusamy, P. (1995) J. Biol. Chem. 270, 304–307[Abstract/Free Full Text]
  33. Samouilov, A., and Zweier, J. L. (1998) Anal. Biochem. 258, 322–330[CrossRef][Medline] [Order article via Infotrieve]
  34. Samouilov, A., Kuppusamy, P., and Zweier, J. L. (1998) Arch. Biochem. Biophys. 357, 1–7[CrossRef][Medline] [Order article via Infotrieve]
  35. Zweier, J. L., Samouilov, A., and Kuppusamy, P. (1999) Biochim. Biophys. Acta 1411, 250–262[Medline] [Order article via Infotrieve]
  36. Xia, Y., and Zweier, J. L. (1995) J. Biol. Chem. 270, 18797–18803[Abstract/Free Full Text]
  37. Zweier, J. L., Wang, P., Samouilov, A., and Kuppusamy, P. (1995) Nat. Med. 1, 804–809[CrossRef][Medline] [Order article via Infotrieve]
  38. Lancaster, J. R., Jr., Langrehr, J. M., Bergonia, H. A., Murase, N., Simmons, R. L., and Hoffman, R. A. (1992) J. Biol. Chem. 267, 10994–10998[Abstract/Free Full Text]
  39. Wong, P. S., and Fukuto, J. M. (1999) Drug Metab. Dispos. 27, 502–509[Abstract/Free Full Text]
  40. Lee, W. I., and Fung, H. L. (2003) Nitric Oxide 8, 103–110[CrossRef][Medline] [Order article via Infotrieve]
  41. Ji, Y., Akerboom, T. P., and Sies, H. (1996) Biochem. J. 313, Pt 2, 377–380
  42. Meyer, D. J., Kramer, H., and Ketterer, B. (1994) FEBS Lett. 351, 427–428[CrossRef][Medline] [Order article via Infotrieve]
  43. Feelisch, M., Schonafinger, K., and Noack, E. (1992) Biochem. Pharmacol. 44, 1149–1157[CrossRef][Medline] [Order article via Infotrieve]
  44. Schneeweiss, A., and Weiss, M. (1990) Advances in Nitrate Therapy, Springer-Verlag, New York
  45. Tiravanti, E., Samouilov, A., and Zweier, J. L. (2004) J. Biol. Chem. 279, 11065–11073[Abstract/Free Full Text]
  46. Zhang, Z., Naughton, D., Winyard, P. G., Benjamin, N., Blake, D. R., and Symons, M. C. (1998) Biochem. Biophys. Res. Commun. 249, 767–772[CrossRef][Medline] [Order article via Infotrieve]
  47. Godber, B. L., Doel, J. J., Sapkota, G. P., Blake, D. R., Stevens, C. R., Eisenthal, R., and Harrison, R. (2000) J. Biol. Chem. 275, 7757–7763[Abstract/Free Full Text]
  48. Meyer, D. J., Kramer, H., Ozer, N., Coles, B., and Ketterer, B. (1994) FEBS Lett. 345, 177–180[CrossRef][Medline] [Order article via Infotrieve]
  49. Tsuchida, S., Maki, T., and Sato, K. (1990) J. Biol. Chem. 265, 7150–7157[Abstract/Free Full Text]
  50. Nigam, R., Anderson, D. J., Lee, S. F., and Bennett, B. M. (1996) J. Pharmacol. Exp. Ther. 279, 1527–1534[Abstract/Free Full Text]
  51. McDonald, B. J., and Bennett, B. M. (1990) Can. J. Physiol. Pharmacol. 68, 1552–1557[Medline] [Order article via Infotrieve]
  52. Chung, S. J., and Fung, H. L. (1990) J. Pharmacol. Exp. Ther. 253, 614–619[Abstract/Free Full Text]
  53. Chen, Z., Zhang, J., and Stamler, J. S. (2002) Proc. Natl. Acad. Sci. U. S. A. 99, 8306–8311[Abstract/Free Full Text]
  54. Khramtsov, V. V., Grigor'ev, I. A., Foster, M. A., and Lurie, D. J. (2004) Antioxid. Redox Signal. 6, 667–676[Medline] [Order article via Infotrieve]
  55. Boyne, A. F., and Ellman, G. L. (1972) Anal. Biochem. 46, 639–653[CrossRef][Medline] [Order article via Infotrieve]
  56. Maia, L., and Mira, L. (2002) Arch. Biochem. Biophys. 400, 48–53[CrossRef][Medline] [Order article via Infotrieve]
  57. Bennett, B. M., McDonald, B. J., Nigam, R., and Simon, W. C. (1994) Trends Pharmacol. Sci. 15, 245–249[CrossRef][Medline] [Order article via Infotrieve]
  58. Habig, W. H., Pabst, M. J., and Jakoby, W. B. (1974) J. Biol. Chem. 249, 7130–7139[Abstract/Free Full Text]
  59. Mukerjee, N., and Pietruszko, R. (1994) J. Biol. Chem. 269, 21664–21669[Abstract/Free Full Text]

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