J Biol Chem, Vol. 275, Issue 13, 9369-9376, March 31, 2000
Regulation of Xanthine Oxidase by Nitric Oxide and
Peroxynitrite*
Chang-il
Lee
§,
Xiaoping
Liu, and
Jay L.
Zweier¶
From the Molecular and Cellular Biophysics
Laboratories, Department of Medicine, Division of Cardiology and the
Electron Paramagnetic Resonance Center, The Johns Hopkins Medical
Institutions, Baltimore, Maryland 21224 and the § Department
of Pharmacology and Electron Spin Resonance Center, Kanagawa Dental
College, 82 Inaoka-cho Yokosuka, Kanagawa, Japan 238-0003
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ABSTRACT |
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 O2 generation and enzyme activity of XO, respectively, by EPR spin trapping of O2 using
5-(diethoxyphosphoryl)-5-methyl-1-pyrroline-N-oxide and
spectrophotometric assay. ONOO markedly inhibited both
O2 generation and XO activity in dose-dependent manner, while NO· from NO· gas in concentrations up to
200 µM had no effect. Furthermore, we observed that
NO· donors such as NOR-1 also inhibited O2 generation
and XO activity; however, these effects were
O2-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 O2 generation, which in turn may
serve to limit further ONOO formation.
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INTRODUCTION |
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 O2, leading to
the formation of superoxide (O2) and
H2O2, 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 degradation during ischemia (2, 7-9). During ischemia and
reperfusion, increased nitric oxide (NO·) formation also occurs,
and this can interact with XO-derived O2, leading to the
formation of peroxynitrite (ONOO) (10). However,
questions remain concerning the effects of NO· and
ONOO on XO itself.
The free radical, NO·, 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 (12-14). NO· 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·-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 O2 plays a critical role in NO·-induced
toxicity, and previously proposed mechanisms for both O2- and
NO·-mediated tissue injury now include a role for their combined reaction product, ONOO (21). The radical-radical reaction
between O2 and NO· is extremely fast and almost
diffusionally limited in rate (2 × 109
M1 s1) (22).
ONOO 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 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) (pKa 6.8) gives reactive
intermediates with OH·-like properties that can oxidize DNA and
proteins (33, 35, 36). ONOO directly oxidizes an active
site methionine, resulting in inactivation of
1-antiprotease (37).
During the early period of reperfusion, both NO· and O2
generation are increased, leading to formation of ONOO-
(10). Following the postischemic burst of ONOO
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· or ONOO could feedback and inhibit
XO, however, controversy remains regarding the presence and nature of
interactions of NO· or ONOO with XO and the role
of this process in regulating oxidant generation. NO· 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· under anaerobic conditions
(40). However, other studies reported that ONOO
inactivates the enzyme, while NO· has no effect (41).
To resolve the controversy regarding the effects of NO· and
ONOO on XO activity, we have determined the
dose-dependent effects of NO· and ONOO
on XO activity and O2 generation from purified XO. EPR spin trapping techniques were applied to quantitate O2. We
demonstrate that ONOO induces a
dose-dependent loss of activity, while its precursor NO· does not. The mechanism of this inactivation is further
shown to be due to disruption of the critical molybdenum center of the enzyme.
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EXPERIMENTAL PROCEDURES |
Materials--
XO (grade III; from buttermilk,
chromatographically purified, in 2.3 M
(NH4)2SO4, 10 mM sodium
phosphate buffer (pH 7.8), containing 1 mM EDTA and 1 mM sodium salicylate) was obtained from Sigma. The
salicylate was removed chromatographically with Sephadex G-25 prior to
use. Xanthine, uric acid, and superoxide dismutase were also
obtained from Sigma. The ONOO decomposition catalyst
(5,10,15,20-tetrakis(2,4,6-trimethyl-3,3-disulfonatophenyl)porphyrinato Fe(III) (FeTMPS) was obtained from Monsanto Corp. (St Louis, MO). The
NO· donor NOR-1 was purchased from BIOMOL Research Laboratories
(Plymouth Meeting, PA).
ONOO Preparation--
The ONOO was
synthesized from acidified nitrite and hydrogen peroxide according to
Beckman et al. (34). Alternatively, similarly prepared
ONOO was obtained from Alexis Corp. (San Diego, CA). The
concentration of ONOO was checked by optical absorbance
measurements at 302 nm at the time of synthesis and again just prior to
each experiment, and only ONOO that was >95% of the
original concentration was used. To assure that there was no
significant pH change upon the addition of alkaline ONOO
to the reaction mixture, we also monitored the pH and limited the
amount of alkaline ONOO stock used so that the final pH
did not significantly change.
NO· Gas Solution--
NO· 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· for 30 min (42, 43). To further verify the precise
NO· concentration from NO· gas-equilibrated solution,
electrochemical measurements of NO· 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· 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 µM) 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 µM) by measurement of uric acid production from the
absorbance change at 295 nm (e = 37,800 M1 cm1). We determined the
effects of ONOO, NO·, and NOR-1 on XO activity as
follows. We confirmed that the various concentrations of the alkaline
ONOO stock used were neutralized to pH 7.4 and
quantitated the ONOO concentration spectrophotometrically
at 302 nm. As previously reported, ONOO rapidly decays at
pH 7.4 with a half-life of <1 s (34), and after 1 min no detectable
ONOO remains. Therefore, ONOO (0-200
µM) 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 µM) was
added for measurement of enzyme activity. Dissolved NO· gas
solutions were used to determine the effects of NO· on XO
activity. The dissolved NO· gas (0-200 µM) 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· remained, ensuring that NO· would not
significantly scavenge O2 generated from the XO-xanthine system. For the NO· donor NOR-1, the NOR-1 (0-100
µM) 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 (100 µM) and NO· (100 µM) were added to
the XO reaction mixture under argon for 10 min. After continued argon
purging to remove any remaining NO·, the enzyme activity was
measured with the addition of xanthine (360 µM) and
exposure to air. Anaerobic solutions of the reaction buffer with XO,
ONOO, 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
TM110 cavity. The microwave frequency was measured with an
EIP model 575 microwave counter (EIP Microwave, Inc., San Jose, CA). To
determine O2 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 µM TEMPO sample measured under identical settings to
estimate the concentration of O2 adduct.
Measurements of the Mo(V) EPR signal of XO were performed as follows.
The samples of native XO (10 µM), XO plus xanthine, ONOO-treated XO, ONOO-treated XO plus
xanthine (360 µM), and ONOO-treated XO plus
dithionite (1 mM) were prepared as described (45). The
reaction mixture (700 µl) in bicine buffer (pH 8.2) was frozen in a
3-mm quartz EPR tube and measured at 77 K. The instrument settings were
as follows: modulation amplitude, 5.0 G; time constant, 0.32 s;
scan time, 60 s; modulation frequency, 100 kHz; microwave power,
10 milliwatts; microwave frequency, 9.54 GHz.
Statistical Analysis--
All of the experiments were performed
in triplicate and repeated a minimum of three times. Results are
expressed as means ± 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 < 0.05.
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RESULTS |
Effects of ONOO and NO· on O2
Generation and XO Activity--
We investigated the effects of
ONOO and NO· on XO-mediated O2 generation
measured by EPR spin trapping with 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). In addition to the large signal of DEPMPO-OOH, a small signal of DEPMPO
OH was
observed. The O2-derived DEPMPO-OOH adduct comprised 92% of
the total intensity, and the DEPMPO-OH adduct comprised 8%. With
ONOO (100 µM) pretreatment of XO and the
subsequent addition of xanthine, the DEPMPOOOH signal was decreased
by more than 5-fold, and a larger signal of DEPMPOOH was seen (Fig.
1A, b). These data indicate that
ONOO markedly inhibits O2 generation from XO and
either forms OH· or hydroxylates DEPMPO in this system. With
NO· treatment, however, no alterations in the EPR spectrum were
seen with identical DEPMPOOOH (92%) and DEPMPOOH (8%) spin
adducts as in the absence of NO· (Fig.
2A). Measurements of XO
activity confirmed that ONOO strongly inactivated the
enzyme with decreased initial rates of uric acid formation (Fig.
1B, b), while NO· had no effect (Fig.
2B, b). The dose-dependent effects of
ONOO and NO· on O2 generation and XO
activity were measured (Fig. 3). It was
clearly observed that ONOO decreased both O2
generation and XO activity in a dose-dependent manner with
more than 25, 50, and 90% inhibition with ONOO levels of
10, 30, and 200 µM, respectively (Fig. 3A);
however, NO· in concentrations up to 200 µM did
not decrease either O2 generation or XO activity (Fig.
3B).
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Fig. 1.
Effect of ONOO on the
O2 generation and activity of XO. A, EPR spin
trapping measurement of O2 generation from XO (0.1 µM) and xanthine (360 µM) in 0.1 M PBS (pH 7.4). a, in the absence of
ONOO, a prominent spectrum of DEPMPOOOH with only a
small signal of DEPMPOOH is seen with relative intensities of 92 and
8%. b, with preexposure of XO to ONOO (100 µM), the DEPMPOOOH signal was decreased 5-fold, and the
DEPMPOOH adduct was increased with relative intensities of 44 and
56%. In B, are shown the time course of O2
generation from either control untreated or ONOO (100 µM)-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).
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Fig. 2.
Effect of NO· on the O2
generation and activity of XO. A, EPR spin trapping
measurement of O2 generation from XO (0.1 µM)
and xanthine (360 µM) in 0.1 M PBS (pH 7.4).
In a and b, measurements were performed either
with control XO or XO preexposed for 10 min to NO· (100 µM) added from gas-equilibrated solution, respectively.
With either control or NO·-pretreated enzyme, similar prominent
spectra of DEPMPOOOH with small signals of DEPMPOOH are seen with
relative intensities of 92 and 8%. B, the time course of
O2 generation measured by EPR spin trapping as in A
for control XO or the enzyme preexposed to NO· (100 µM) (a) and the kinetics of XO activity
determined by uric acid production from spectrophotometric assay at 295 nm (b).
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Fig. 3.
Dose-dependent effects of
ONOO and NO· on O2 generation and
activity of XO. ONOO (0200 µM)
(A) or NO· gas solution (0200 µM)
(B) was added to XO (0.1 µM) as described in
the legends to Fig. 1 and 2. O2 generation was measured by EPR
spin trapping using DEPMPO (solid bars), and XO activity was
measured from spectrophotometric assay of urate production monitored at
295 nm (open bars). Both were performed with the addition of
xanthine (360 µM) to the reaction mixture. Data are
presented as mean ± S.E. of triplicate experiments. *,
significance of p < 0.01 from the corresponding
control value.
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Effects of NO· Donors on O2 Generation and XO
Activity--
While NO· in aerobic solution has a short
concentration-dependent half-life of only a few seconds,
longer sustained NO· levels can be achieved with NO·
donors. It was previously reported that NO· released from
NO· donors can inhibit XO function (38, 39); however, it is
unknown if this effect is mediated by NO· or ONOO
generated from the reaction of NO· with XO-derived O2.
The time course of NO· release from NOR-1 was measured by the
NO· electrode (see "Experimental Procedures"). In contrast
to NO· gas, with which a rapid fall in NO· concentration
to near zero within 10 min is observed, with NOR-1, NO·
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 O2
generation and XO activity was observed (Figs. 5 and
6A). With NOR-1 concentrations
of 100 µM, more than 50% inhibition was seen. This
inhibition by 100 µM NOR-1 was completely blocked by
superoxide dismutase (250 units/ml) (Fig. 6B), the ONOO scavenger urate (10 µM) (Fig.
7A), or the ONOO
decomposition catalyst FeTMPS (10 µM) (Fig.
7B). In similar experiments in which NOR-1 was preincubated
with XO for 2 h, a time sufficient for total NO· release
and decay, no inhibition of subsequent O2 generation or XO
activity occurred (data not shown). Thus, the inhibition of XO function
was mediated by ONOO (Figs. 6 and 7), and EPR
measurements demonstrate that the inhibition by the NO· donor on
XO activity occurs only when NO· continues to be released at the
time of O2 generation from the XO-xanthine system (Figs. 4 and
5).
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Fig. 4.
Time course of decline in NO·
concentration in aqueous solutions after the addition of NO· gas
or the NO· donor NOR-1. NO· concentration was
measured by polarigraphic electrode. A, at time 0, either
200, 100, 30, 10, or 1 µM NO· (top
curve to bottom curve) from NO·
gas- equilibrated solutions was added to PBS (pH 7.4) at 25 °C.
B, at time 0, either 100, 50, or 10 µM (a, b, and c,
respectively) NOR-1 was added to PBS (pH 7.4) at 25 °C. While the
NO· concentration from NO· gas falls to near 0 (<25
nM) by 10 min, the NO· donor NOR-1 provides
sustained NO· release and measurable solution NO·
concentrations for more than 15 min.
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Fig. 5.
Effect of the NO· donor NOR-1 on the
O2 generation and activity of XO. A, EPR spin
trapping measurement of O2 generation from XO (0.1 µM) and xanthine (360 µM) in 0.1 M PBS (pH 7.4). a, in the absence of NOR-1 a
prominent spectrum of DEPMPOOOH with only a small signal of
DEPMPOOH is seen as in Fig. 1; b, with the addition of
NOR-1 (100 µM) the DEPMPOOOH signal was decreased about
3-fold. In B are shown the time course of O2
generation from XO and xanthine in the presence or absence of NOR-1
(100 µM) (a) and the XO activity assayed from
the kinetics of uric acid production monitored at 295 nm
(b).
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Fig. 6.
Dose-dependent effects of NOR-1
on O2 generation and XO activity (A) and
effect of superoxide dismutase (SOD) on NOR-1-mediated
inhibition (B). A, NOR-1 (0100
µM) and xanthine (360 µM) were added to XO
(0.1 µM) in 0.1 M PBS (pH 7.4) at 25 °C,
and O2 generation was then immediately measured by EPR spin
trapping with DEPMPO, while in matched experiments enzyme activity was
measured from urate production monitored at 295 nm. In B,
experiments were performed as in A, but superoxide dismutase
(250 units/ml) was added to the XO before the addition of NOR-1 (100 µM) and xanthine (360 µM). Data are
presented as mean ± S.E. of triplicate experiments. *,
significance of p < 0.01 for the difference from the
corresponding control value;  , significance of p < 0.01 for the difference from the value with NOR-1.
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Fig. 7.
Effect of the ONOO scavenger
urate and ONOO decomposition catalyst FeTMPS on
NOR-1-mediated inhibition of XO activity. NOR-1 (100 µM) and xanthine (360 µM) were added
simultaneously to the XO (0.1 µM) in 0.1 M
PBS, and XO activity was immediately measured. ONOO (100 µM) was preincubated with XO for 1 min, and then xanthine
was added. The assays of XO activity were performed by measurement of
uric acid production at 295 nm. XO was preincubated with urate (10 µM) (A) or with FeTMPS (10 µM)
(B) for 3 min before the addition of NOR-1 and xanthine or
the addition of ONOO. Data are presented as mean ± S.E. of triplicate experiments. *, significance of p < 0.01 for the decrease from the corresponding value of control; ,
significance of p < 0.01 for the increase from the
corresponding value of XO treated with NOR-1; ¶, significance of
p < 0.05 for the increase from the value of XO treated
with ONOO.
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Effects of ONOO and NO· on O2
Generation and XO Activity under Anaerobic Conditions--
Under
anaerobic conditions, NO· is stable and persists for longer
periods of time so that anerobic incubation of XO with a given
concentration of NO· could exert more pronounced effects than
those seen with aerobic exposure. Since NO· from
gas-equilibrated solution (
200 µM) did not
significantly decrease XO activity under aerobic conditions, we
examined effects of NO· and ONOO on enzyme
activity under anaerobic conditions. Similar to the results under
aerobic conditions, only ONOO-significantly decreased XO
activity, while there was no significant effect on enzyme activity with
NO· preexposure (Fig. 8).
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Fig. 8.
Effects of ONOO and NO·
on XO activity under anaerobic conditions.
ONOO (100 µM) or NO· (100 µM) from NO· gas solution were incubated with XO
(0.1 µM) under an argon-saturated atmosphere for 10 min
at 25 °C, and then, after continued argon purging to remove any
remaining NO·, the enzyme activity was measured with the
addition of xanthine (360 µM) and exposure to air.
A, shows the kinetics of XO activity measured from uric acid
production at 295 nm. The bar graph in
B shows the mean ± S.E. values observed in a series of
triplicate experiments. *, significance of p < 0.01 for the difference from the corresponding value of control.
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Effects of ONOO on the Molybdenum Center--
To
examine the effects of ONOO 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, 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 irreversibly oxidizes and disrupts the
molybdenum center of XO.
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Fig. 9.
Mo(V) EPR spectra of XO treated with
ONOO. A, 20 mM Bicine buffer
(pH 8.2). B, native XO (10 µM); the rapid
signal Mo(V) EPR spectrum is observed. C, native XO plus
xanthine (360 µM); the rapid signal Mo(V) EPR spectrum is
increased compared with B. D, XO treated with
ONOO (200 µM); the rapid signal of Mo(V)
EPR spectrum is abolished. E, with xanthine addition to
preparation in D; no restoration of the Mo(V) signal is
seen. F, as in D with the addition of dithionite
(1 mM); no restoration of the Mo(V) signal occurs. The
reaction samples in Bicine buffer (pH 8.2) were frozen in quartz EPR
tubes and measured at 77 K. The instrument settings were as follows:
modulation amplitude, 5.0 G; time constant, 0.32 s; scan time,
60 s; modulation frequency, 100 kHz; microwave power, 10 milliwatts; microwave frequency, 9.52 GHz.
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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· generation and accumulation in ischemic
and postischemic tissues, such as the heart, and this NO· reacts
with O2 generated during the early period of reperfusion, resulting in the formation of ONOO (10). However,
questions and controversy remain regarding whether NO· or
ONOO modulate XO function and XO-mediated free radical
generation. Most prior studies assessed the effects of NO· and
ONOO on XO activity measured by urate production;
however, there was a lack of direct monitoring of the concentrations of
NO· present along with the process of XO-mediated O2
generation. This has led to the present controversy and confusion
regarding the role of NO· and ONOO on XO function.
Therefore, we have performed studies to assess the effects of
NO· and ONOO on XO activity and O2
generation using electrochemical and EPR methods to monitor NO·
concentrations and O2 generation directly. Our studies provide direct evidence that ONOO down-regulates O2
generation from XO. ONOO was shown to decrease both XO
activity and O2 generation in a dose-dependent
manner, while NO· had no significant effect (Fig. 3).
We found that NO· does not directly inactivate XO but must first
react with O2 to form ONOO (Fig. 3). Since in
the process of assaying XO activity with the addition of xanthine or
other substrates the enzyme generates O2, any NO·
present at the time of enzyme activation can be converted to ONOO. Although the NO· donor NOR-1 is reported to
provide rapid release of NO·, this release was measured to
persist for more than 10 min as determined by either polarigraphic
electrode or EPR spin trapping methods. While NO· itself was
ineffective even at 200 µM concentrations, the much lower
but more sustained NO· concentrations, <5 µM,
provided by the NO· donor were effective in inhibiting
O2 generation and XO activity (Figs. 5 and 6). This
NOR-1-mediated XO inhibition was prevented by either superoxide
dismutase, the ONOO scavenger urate, or the
ONOO decomposition catalyst FeTMPS (Figs. 6 and 7). When
prolonged periods of time were allowed to assure full decomposition of
NOR-1 and NO· 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 formation from the reaction of
O2 formed by the XO-xanthine system and NO· released
from NOR-1.
We further investigated the effects of ONOO on the
structure of the critical molybdenum center. ONOO
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· reacts with reduced XO and
converts the enzyme to the inactive desulfo-form under anaerobic conditions (40). Since NO· rapidly decomposes by reaction with
oxygen under aerobic conditions (Fig. 4), this could limit the effect
of NO· 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· added at initial
concentrations of up to 200 µM. The inhibition of the
reduced enzyme, however, could be in part the result of ONOO formation upon exposure of the reduced enzyme to
oxygen and xanthine at the time of the assay of XO activity. If
NO· was present at the time of reoxygenation or the addition of
xanthine, ONOO would be formed as was the case with the
NO· donor NOR-1.
The physiological or pathophysiological relevance of the modulation of
XO function by ONOO or NO· can be considered in
view of the levels of ONOO required to modulate XO
function. It was observed that ONOO concentrations
greater than 10 µM exerted prominent effects on the
function of XO, while NO· concentrations of up to 200 µM had no effect. Furthermore, with NO· donors
that resulted in a low level flux of a more sustained production of
ONOO, it was observed that solution NO·
concentrations of less than 1 µM were sufficient to
inhibit XO function when present at the time when XO is activated by
its substrate xanthine to produce O2. Thus, a low level flux
of ONOO formation was sufficient to inhibit the enzyme.
Since NO· concentrations of 0.1 µM or more occur
in postischemic organs or following sepsis or inflammation (49, 50), it
is likely that ONOO formation would feedback and regulate
O2 generation from XO. With ONOO generated from
O2 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 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 scavenger so that under
conditions with loss of perfusion and stasis this XO product could
scavenge ONOO, limiting this feedback regulation.
From prior data in the literature, one can consider if this
ONOO-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 O2 from XO as well as
NO· from nitric-oxide synthase or nitrite reduction leading to a burst of ONOO 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 generation (9,
10). This suggests that ONOO-mediated regulation of XO
does occur in postischemic tissues.
In conclusion, we have demonstrated that ONOO inhibits
the O2 generation and activity of XO in a
dose-dependent manner, while NO· only exerts
significant inhibition in the presence of O2 under conditions
in which ONOO is formed. This ONOO-mediated
XO inhibition could be suppressed by urate. ONOO
inhibited XO function primarily by oxidative disruption of the molybdenum catalytic site. Taken together, ONOO in
biological systems can feedback and down-regulate XO activity that in
turn may serve to limit further ONOO formation and
oxidant-derived injury.
| |
ACKNOWLEDGEMENTS |
We thank Dr. P. Kuppusamy, Dr. A. Samouilov,
Dr. A. F. Vanin, and Dr. Russ Hille for helpful advice.
| |
FOOTNOTES |
*
This work was supported by National Institutes of Health
Grants HL-38324 and HL-52315.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.
¶
To whom correspondence should be addressed: The EPR Center,
Johns Hopkins Asthma and Allergy Center, 5501 Johns Hopkins Bayview Circle, Baltimore, MD 21224. Tel.: 410-550-0339; Fax: 410-550-2448; E-mail: jzweier@welch.jhu.edu.
| |
ABBREVIATIONS |
The abbreviations used are:
XO, xanthine
oxidase;
O2, superoxide;
NO·, nitric oxide;
ONOO, peroxynitrite;
DEPMPO, 5-(diethoxyphosphoryl)-5-methyl-1-pyrroline-N-oxide;
FeTMPS, (5,10,15,20-tetrakis(2,4,6-trimethyl-3,3-disulfonatophenyl)porphyrinato Fe(III);
PBS, phosphate-buffered saline;
Bicine, N,N-bis(2-hydroxyethyl)glycine.
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ABSTRACT
INTRODUCTION
