Regulation of xanthine oxidase by nitric oxide and peroxynitrite.

Xanthine oxidase (XO) is a central mechanism of oxidative injury as occurs following ischemia. During the early period of reperfusion, both nitric oxide (NO(*)) and superoxide (O-*(2)) generation are increased leading to the formation of peroxynitrite (ONOO(-)); however, questions remain regarding the presence and nature of the interactions of NO(*) or ONOO(-) with XO and the role of this process in regulating oxidant generation. Therefore, we determined the dose-dependent effects of NO(*) and ONOO(-) on the O-*(2) generation and enzyme activity of XO, respectively, by EPR spin trapping of O-*(2) using 5-(diethoxyphosphoryl)-5-methyl-1-pyrroline-N-oxide and spectrophotometric assay. ONOO(-) markedly inhibited both O-*(2) generation and XO activity in dose-dependent manner, while NO(*) from NO(*) gas in concentrations up to 200 microM had no effect. Furthermore, we observed that NO(*) donors such as NOR-1 also inhibited O-*(2) generation and XO activity; however, these effects were O-*(2)-dependent and blocked by superoxide dismutase or ONOO(-) scavengers. Finally, we found that ONOO(-) totally abolished the Mo(V) EPR spectrum. These changes were irreversible, suggesting oxidative disruption of the critical molybdenum center of the catalytic site. Thus, ONOO(-) formed in biological systems can feedback and down-regulate XO activity and O-*(2) generation, which in turn may serve to limit further ONOO(-) formation.

It has been demonstrated that oxygen free radical generation is a critical mechanism causing injury in postischemic cells and tissues. Xanthine oxidase (XO), 1 a metalloflavoprotein, is an important source of oxygen free radicals (1). The enzyme catalyzes the reduction of O 2 , leading to the formation of superoxide (O 2 . ) and H 2 O 2 , and it has been proposed as a central mechanism of oxidative injury (2,3). Both direct and spin trapping EPR techniques have demonstrated markedly increased oxygen free radical generation in tissues, such as heart, following postischemic reperfusion (4 -6). XO has been shown to be the primary source of the oxygen radical generation, with this process largely triggered by increased formation of the XO substrates, xanthine and hypoxanthine, due to ATP degrada-tion during ischemia (2,(7)(8)(9). During ischemia and reperfusion, increased nitric oxide (NO ⅐ ) formation also occurs, and this can interact with XO-derived O 2 . , leading to the formation of peroxynitrite (ONOO Ϫ ) (10). However, questions remain concerning the effects of NO ⅐ and ONOO Ϫ on XO itself. The free radical, NO ⅐ , is generated in biological tissues and is an important regulator of a wide range of biological functions (11). It is of critical importance in modulating vascular tone and was identified as the mediator of endothelium-derived relaxation (12)(13)(14). NO ⅐ also inhibits the enzyme activity of a number of enzymes including gluthathione peroxidase (15), cytochrome c oxidase (16), and NADPH oxidase (17,18). Major mechanisms attributed to explain NO ⅐ -mediated inhibitory effects involve heme binding or destruction of enzyme Fe-S centers to yield inactive Fe-S-NO derivatives and thiol oxidation (19,20). Recent reports have shown that O 2 . plays a critical role in NO ⅐ -induced toxicity, and previously proposed mechanisms for both O 2 . -and NO ⅐ -mediated tissue injury now include a role for their combined reaction product, ONOO Ϫ (21). The radicalradical reaction between O 2 . and NO ⅐ is extremely fast and almost diffusionally limited in rate (2 ϫ 10 9 M Ϫ1 s Ϫ1 ) (22). ONOO Ϫ is a potent oxidant that can attack a wide variety of biological molecules and is produced in diverse inflammatory and pathological processes including postischemic injury (10), septic shock (23), chronic tissue rejection (24), multiple sclerosis (25), amyotrophic lateral sclerosis (26,27), Alzheimer's disease (28), cardiomyopathy (29,30), and atherosclerosis (31,32). ONOO Ϫ can directly oxidize sulfhydryls (33) and also reacts by either one-or two-electron oxidation reactions with various biological target molecules (34). In acid conditions, the homolysis of peroxynitrous acid (HOONO) (pK a 6.8) gives reactive intermediates with OH ⅐ -like properties that can oxidize DNA and proteins (33,35,36). ONOO Ϫ directly oxidizes an active site methionine, resulting in inactivation of ␣ 1 -antiprotease (37).
During the early period of reperfusion, both NO ⅐ and O 2 .
generation are increased, leading to formation of ONOO - (10). Following the postischemic burst of ONOO Ϫ generation, it has been observed that XO activity is decreased in the early period of reperfusion in heart tissue (9). It has been suggested that either NO ⅐ or ONOO Ϫ could feedback and inhibit XO, however, controversy remains regarding the presence and nature of interactions of NO ⅐ or ONOO Ϫ with XO and the role of this process in regulating oxidant generation. NO ⅐ could inhibit and regulate endothelial cell xanthine dehydrogenase/XO activity (38,39), and a recent paper reported that XO and xanthine dehydrogenase are inactivated by NO ⅐ under anaerobic conditions (40). However, other studies reported that ONOO Ϫ inactivates the enzyme, while NO ⅐ has no effect (41 ONOO Ϫ Preparation-The ONOO Ϫ was synthesized from acidified nitrite and hydrogen peroxide according to Beckman et al. (34). Alternatively, similarly prepared ONOO Ϫ was obtained from Alexis Corp. (San Diego, CA). The concentration of ONOO Ϫ was checked by optical absorbance measurements at 302 nm at the time of synthesis and again just prior to each experiment, and only ONOO Ϫ that was Ͼ95% of the original concentration was used. To assure that there was no significant pH change upon the addition of alkaline ONOO Ϫ to the reaction mixture, we also monitored the pH and limited the amount of alkaline ONOO Ϫ stock used so that the final pH did not significantly change.
NO ⅐ Gas Solution-NO ⅐ was scrubbed of higher nitrogen oxides by passage through a trap with solid NaOH pellets and a second trap with 1 M deaerated (bubbled with argon for 30 min) NaOH solution. 500 ml of phosphate-buffered saline, pH 7.4 (PBS), was deaerated by bubbling with argon for 30 min and then bubbled with scrubbed NO ⅐ for 30 min (42,43). To further verify the precise NO ⅐ concentration from NO ⅐ gas-equilibrated solution, electrochemical measurements of NO ⅐ concentrations were carried out at 25°C using a CHI 832 electrochemical detector with a Faraday Cage (CH Instruments, Inc., Cordova, IN) and WPI NO ⅐ electrode.
Spectrophotometric Measurements-UV-visible absorption spectra of XO and assays of XO activity were performed with a Varian Cary 300 UV-visible spectrophotometer equipped with a temperature-controlled circulator. To remove the salicylate and other low molecular weight compounds, XO (0.1 M) was passaged through Sephadex G-25 preequilibrated with PBS, pH 7.4. XO activity was assayed at 25°C in PBS after the addition of xanthine (360 M) by measurement of uric acid production from the absorbance change at 295 nm (e ϭ 37,800 M Ϫ1 cm Ϫ1 ). We determined the effects of ONOO Ϫ , NO ⅐ , and NOR-1 on XO activity as follows. We confirmed that the various concentrations of the alkaline ONOO Ϫ stock used were neutralized to pH 7.4 and quantitated the ONOO Ϫ concentration spectrophotometrically at 302 nm. As previously reported, ONOO Ϫ rapidly decays at pH 7.4 with a half-life of Ͻ1 s (34), and after 1 min no detectable ONOO Ϫ remains. Therefore, ONOO Ϫ (0 -200 M) was added with a gas-tight syringe to the XO reaction mixture in PBS, and after 1 min the reaction mixture was transferred to the spectrophotometer cuvette, and xanthine (360 M) was added for measurement of enzyme activity. Dissolved NO ⅐ gas solutions were used to determine the effects of NO ⅐ on XO activity. The dissolved NO ⅐ gas (0 -200 M) was preincubated with XO for 10 min in 0.1 M PBS (pH 7.4), after which xanthine was added, and enzyme activity was measured. Electrode measurements confirmed that after a 10-min preincubation, no detectable NO ⅐ remained, ensuring that NO ⅐ would not significantly scavenge O 2 . generated from the XO-xanthine system. For the NO ⅐ donor NOR-1, the NOR-1 (0 -100 M) and xanthine were added simultaneously to the XO reaction mixture in 0.1 M PBS, and the rate of uric acid production was immediately measured. For the anaerobic experiments, ONOO Ϫ (100 M) and NO ⅐ (100 M) were added to the XO reaction mixture under argon for 10 min. After continued argon purging to remove any remaining NO ⅐ , the enzyme activity was measured with the addition of xanthine (360 M) and exposure to air. Anaerobic solutions of the reaction buffer with XO, ONOO Ϫ , and xanthine were prepared by purging with argon prior to use.
EPR Measurements-All EPR measurements were performed using a Bruker ER 300 spectrometer operating at X-band with a TM 110 cavity. generation from either control untreated or ONOO Ϫ (100 M)-pretreated XO measured by EPR spin trapping following the addition of xanthine at time 0 (a) and the kinetics of XO activity determined by uric acid production from spectrophotometric assay at 295 nm (b).
generation, EPR spin trapping studies were performed using the spin trap 5-(diethoxyphosphoryl)-5-methyl-1-pyrroline-N-oxide (DEPMPO) (44). The instrument settings used in the spin trapping experiments were as follows: modulation amplitude, 0.32 G; time constant, 0.16 s; scan time, 60 s; modulation frequency, 100 kHz; microwave power, 20 milliwatts; microwave frequency, 9.76 GHz. The samples were placed in a quartz EPR flat cell, and spectra were recorded at ambient temperature (25°C). The component signals observed in these spectra were identified and quantified as reported (46). The double integrals of DEPMPO-OOH experimental spectra were compared with those of a 1 M TEMPO sample measured under identical settings to estimate the concentration of O 2 . adduct.
Statistical Analysis-All of the experiments were performed in triplicate and repeated a minimum of three times. Results are expressed as means Ϯ S.E. Statistical analysis was performed by Student's t test or a one-way analysis of variance. Statistical significance was defined at a level of p Ͻ 0.05. DEPMPO. XO activity was assayed under these same experimental conditions. As reported previously (44), after the addition of xanthine to XO, we observe primarily a characteristic DEPMPO-OOH adduct spectrum with hyperfine splitting giving rise to 12 resolved peaks (Fig. 1A, a). adduct comprised 92% of the total intensity, and the DEPMPO-OH adduct comprised 8%. With ONOO Ϫ (100 M) pretreatment of XO and the subsequent addition of xanthine, the DEPMPOϪOOH signal was decreased by more than 5-fold, and a larger signal of DEPMPOϪOH was seen (Fig. 1A, b).

Effects of ONOO
These data indicate that ONOO Ϫ markedly inhibits O 2 . generation from XO and either forms OH ⅐ or hydroxylates DEPMPO in this system. With NO ⅐ treatment, however, no alterations in the EPR spectrum were seen with identical DEPMPOϪOOH (92%) and DEPMPOϪOH (8%) spin adducts as in the absence of NO ⅐ (Fig. 2A). Measurements of XO activity confirmed that ONOO Ϫ strongly inactivated the enzyme with decreased initial rates of uric acid formation (Fig. 1B, b), while NO ⅐ had no effect (Fig. 2B, b). The dose-dependent effects of ONOO Ϫ and NO ⅐ on O 2 . generation and XO activity were measured (Fig. 3) . In contrast to NO ⅐ gas, with which a rapid fall in NO ⅐ concentration to near zero within 10 min is observed, with NOR-1, NO ⅐ concentrations in solution initially increase and then persist for more than 15 min (Fig. 4). When NOR-1 was added to XO and this was immediately followed by the addition of xanthine, a dose-dependent inhibition of O 2 . generation and XO activity was observed (Figs. 5 and 6A). With NOR-1 concentrations of 100 M, more than 50% inhibition was seen. This inhibition by 100 M NOR-1 was completely blocked by superoxide dismutase (250 units/ml) (Fig. 6B), the ONOO Ϫ scavenger urate (10 M) (Fig. 7A), or the ONOO Ϫ decomposition catalyst FeTMPS (10 M) (Fig. 7B) (Fig. 8).
Effects of ONOO Ϫ on the Molybdenum Center-To examine the effects of ONOO Ϫ on the critical molybdenum center of XO, EPR measurements were performed on frozen enzyme at 77 K. In the oxidized state, the molybdenum is present as Mo(VI), which is EPR-silent; however, with reduction to Mo(V), various EPR signals have been reported (45, 46 -48). Studies of the Mo(V) EPR signal of XO have provided important information regarding the mechanism of enzyme catalysis and have shown that the oxidative hydroxylation of xanthine to uric acid takes place at the molybdenum center. As previously reported, we observe that native XO exhibits the rapid signal Mo(V) EPR spectrum (Fig. 9B) in Bicine buffer (pH 8.2), and the intensity of this signal further increases after the addition of xanthine (Fig. 9C) (45). Following the addition of ONOO Ϫ , however, the rapid signal of the Mo(V) EPR spectrum is almost totally abolished (Fig. 9D). The loss of this signal could not be reversed by the addition of xanthine (Fig. 9E) or dithionite (Fig. 9F). These observations suggest that ONOO Ϫ irreversibly oxidizes and disrupts the molybdenum center of XO. DISCUSSION XO is an important source of oxygen free radicals in biological cells and tissues and has a particularly important role in oxygen radical generation and pathogenesis of injury following postischemic reperfusion (2). More recently, we have also demonstrated that there is markedly increased NO ⅐ generation and accumulation in ischemic and postischemic tissues, such as the heart, and this NO ⅐ reacts with O 2 . generated during the early period of reperfusion, resulting in the formation of ONOO Ϫ  (Fig. 3).
We found that NO ⅐ does not directly inactivate XO but must first react with O 2 . to form ONOO Ϫ (Fig. 3). Since in the process of assaying XO activity with the addition of xanthine or other substrates the enzyme generates O 2 . , any NO ⅐ present at the time of enzyme activation can be converted to ONOO Ϫ . Although the NO ⅐ donor NOR-1 is reported to provide rapid release of NO ⅐ , this release was measured to persist for more than 10 min as determined by either polarigraphic electrode or EPR spin trapping methods. While NO ⅐ itself was ineffective  . When prolonged periods of time were allowed to assure full decomposition of NOR-1 and NO ⅐ decay prior to the addition of xanthine to the enzyme, no inhibition was seen. Thus, the NOR-1-mediated XO inhibition was associated with ONOO Ϫ formation from the reaction of O 2 . formed by the XOxanthine system and NO ⅐ released from NOR-1. We further investigated the effects of ONOO Ϫ on the structure of the critical molybdenum center. ONOO Ϫ treatment resulted in a near total and irreversible loss of the rapid signal Mo(V) EPR spectrum, indicating oxidative disruption of the molybdenum center (Fig. 9). The near total loss of this spectrum correlated with the loss of enzyme activity, indicating that oxidative disruption of the molybdenum was the basis for the loss of enzyme function.
A recent study reported that NO ⅐ reacts with reduced XO and converts the enzyme to the inactive desulfo-form under anaerobic conditions (40). Since NO ⅐ rapidly decomposes by reaction with oxygen under aerobic conditions (Fig. 4), this could limit the effect of NO ⅐ on XO if this reaction with the enzyme proceeds at a slow rate. We observed under aerobic conditions that little if any inhibition of XO occurred with NO ⅐ added at initial concentrations of up to 200 M. The inhibition of the reduced enzyme, however, could be in part the result of ONOO Ϫ formation upon exposure of the reduced enzyme to oxygen and xanthine at the time of the assay of XO activity. If NO ⅐ was present at the time of reoxygenation or the addition of xanthine, ONOO Ϫ would be formed as was the case with the NO ⅐ donor NOR-1.
The physiological or pathophysiological relevance of the modulation of XO function by ONOO Ϫ or NO ⅐ can be considered in view of the levels of ONOO Ϫ required to modulate XO function. It was observed that ONOO Ϫ concentrations greater than 10 M exerted prominent effects on the function of XO, while NO ⅐ concentrations of up to 200 M had no effect. Furthermore, with NO ⅐ donors that resulted in a low level flux of a more sustained production of ONOO Ϫ , it was observed that solution NO ⅐ concentrations of less than 1 M were sufficient to inhibit XO function when present at the time when XO is activated by its substrate xanthine to produce O 2 . . Thus, a low level flux of ONOO Ϫ formation was sufficient to inhibit the enzyme. Since NO ⅐ concentrations of 0.1 M or more occur in postischemic organs or following sepsis or inflammation (49,50), it is likely that ONOO Ϫ formation would feedback and regulate O 2 . generation from XO. With ONOO Ϫ generated from O 2 . released from the active site of XO, high local concentrations would be produced near the critical molybdenum and other catalytic centers. Thus, XO function could be particularly sensitive to feedback regulation by the ONOO Ϫ derived in part from the enzyme. This could serve a critical function to regulate oxidative tissue injury. It is interesting to also consider that the XO product, uric acid, is an effective ONOO Ϫ scavenger so that under conditions with loss of perfusion and stasis this XO product could scavenge ONOO Ϫ , limiting this feedback regulation. From prior data in the literature, one can consider if this ONOO Ϫ -mediated regulation of XO function may actually occur in biological tissues. We have previously observed in the isolated rat heart with reintroduction of flow following ischemia that there is increased generation of O 2 . from XO as well as NO ⅐ from nitric-oxide synthase or nitrite reduction leading to a burst of ONOO Ϫ formation during the early minutes of reflow (5,10,49,50). Interestingly, we previously observed that while XO activity is initially increased during ischemia, a 30% decline in XO activity occurs during reperfusion at a time following the burst of reperfusion-associated ONOO Ϫ generation (9,10). This suggests that ONOO Ϫ -mediated regulation of XO does occur in postischemic tissues.
In conclusion, we have demonstrated that ONOO Ϫ inhibits the O 2 . generation and activity of XO in a dose-dependent manner, while NO ⅐ only exerts significant inhibition in the presence of O 2 . under conditions in which ONOO Ϫ is formed.
This ONOO Ϫ -mediated XO inhibition could be suppressed by urate. ONOO Ϫ inhibited XO function primarily by oxidative disruption of the molybdenum catalytic site. Taken together, ONOO Ϫ in biological systems can feedback and down-regulate XO activity that in turn may serve to limit further ONOO Ϫ formation and oxidant-derived injury.