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INTRODUCTION |
Mammalian xanthine oxidase
(XO1; EC 1.1.3.22) and
xanthine dehydrogenase (XDH; EC 1.1.1.204), which are alternative forms of the same gene product (for a recent review, see Ref. 1), are complex
flavoproteins composed of two identical subunits of Mr 145,000; each subunit contains one molybdenum
center, two non-identical Fe2S2-type
iron-sulfur centers, and one FAD center. The enzymes catalyze oxidation
of xanthine to uric acid with concomitant reduction of NAD+
or molecular oxygen. The oxidative hydroxylation of xanthine to uric
acid takes place at the molybdenum center, and reducing equivalents
thus introduced into enzymes are transferred rapidly via intramolecular
electron transfer to FAD, where physiological oxidation occurs (2).
Both enzymes can reduce molecular oxygen to superoxide and hydrogen
peroxide (1), but XDH is characterized by high reactivity toward
NAD+, but low reactivity toward O2, whereas XO
has high reactivity toward O2, but negligible reactivity
toward NAD+ (2). Mammalian XDH can be converted to XO
either by the oxidation of sulfhydryl groups or by limited proteolysis
(3, 4). XO is thought to be one of the key enzymes for cellular injury
by superoxide and related active oxygen species (5). In particular, much attention has been paid to XO in connection with the pathogenesis of ischemia/reperfusion injury since it has been proposed that superoxide/H2O2 production by XO is enhanced by
the accelerated conversion of XDH to XO (6), the accumulation of ATP
degradation products (i.e. hypoxanthine and xanthine) that
are substrates for XO (7), and the up-regulation of XDH/XO mRNA
(8). However, the role of XDH/XO in the pathogenesis of such injury is
still controversial (9).
Nitric oxide (NO) is now recognized as a multifunctional molecule (10,
11), one function of which is to inactivate biologically important
enzymes such as aconitase (12), ribonucleotide reductase (13),
glutathione peroxidase (14), cytochrome c oxidase (15), and
NADPH oxidase (16, 17). Although inhibitory actions of NO are
attributed to its reaction with heme or non-heme iron or copper or to
S-nitrosylation or sulfhydryl oxidation (18), the precise
mechanisms remain to be established (12). Considering that NO
generation also increases under conditions where superoxide generation
increases, it is necessary to clarify whether NO inhibits superoxide-generating systems. We recently investigated two major sources of superoxide. We identified the site affected by NO in neutrophil NADPH oxidase (16) and showed that NO inhibits XO during
enzyme turnover under cell-free conditions (19). However, the exact
mechanism and kinetics of the reaction between NO and XO remained to be
examined. Subsequent reports demonstrated the NO-induced inactivation
of XDH and XO in interferon-
-stimulated macrophages (20) and
endothelial cells using exogenously and endogenously produced NO
(21, 22). Direct binding of NO to the enzyme iron-sulfur moiety or to
its sulfhydryl groups was postulated since NO did not significantly
affect the mRNA expression of XDH in these cellular systems (20,
22) and may modify XO directly at a post-translational level. The exact
mechanism of the inhibition still remains to be determined because of
the limitations of the cellular system. In this study, we examined the
kinetics of the reaction between XO and NO and identified the site
responsible for the inactivation. The inhibitory kinetics of XDH were
also examined to obtain basic data on the interaction of NO and the superoxide-generating system in the event of reduced oxygen supply, such as ischemia.
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EXPERIMENTAL PROCEDURES |
Materials--
XO was isolated from fresh bovine milk and
purified according to the method reported previously (23). The
activity/flavin ratio (AFR) of the prepared XO was >190 (fully active
enzyme has a value of 210) (23-25). The desulfoenzyme was prepared by
incubating the enzyme with 10 mM KCN for 2 h at
25 °C, followed by gel filtration to remove KCN as described by
Massey and Edmondson (25). Bovine milk XDH, prepared essentially
according to the method of Nakamura and Yamazaki (26) without folate
affinity chromatography, has an AFR of >100 and a
dehydrogenase/oxidase activity ratio of >7. The dehydrogenase/oxidase
activity ratio was determined as the ratio of the absorbance change at
295 nm under aerobic conditions in the presence of NAD+ to
that in the absence of NAD+. Desulfo-XDH was prepared using
the same procedure as described for XO, except that 5 mM
dithiothreitol was added to the incubation mixture to avoid conversion
to XO during preparation. Xanthine, allopurinol, and Hepes were
obtained from Sigma. A stock solution of oxymyoglobin (~0.7
mM) was prepared in 200 mM Hepes buffer (pH
7.0) by reducing metmyoglobin (horse heart, Sigma) with sodium dithionite, followed by gel filtration on a Sephadex G-25 column (medium) (27). The solution was stored frozen at
80 °C until use.
Spectrophotometric Determination--
Spectrophotometric
measurements were conducted on a Hitachi U-3200 spectrophotometer
equipped with a temperature-controlled circulator. The XO concentration
was determined spectrophotometrically using a molar absorption
coefficient of 37,800 M
1 cm
1 at
450 nm (28). The concentration of nitric oxide in reaction mixtures was
determined spectrophotometrically using oxymyoglobin. The principle of
this procedure is the same as that of the oxyhemoglobin method reported
previously (29). The standard oxymyoglobin concentration was determined
spectrophotometrically using a molar absorption coefficient of 10,700 M
1 cm
1 at 540 nm after
conversion to cyanometmyoglobin (30, 31). Nitric oxide solution was
mixed with 5-10 µM oxymyoglobin in 200 mM
Hepes buffer (pH 7.4), and the increase in absorbance at 406 nm due to
metmyoglobin formation was monitored. The standard curve for the assay
was obtained by oxidation of oxymyoglobin with titrated amounts of
potassium ferricyanide. The oxidation was complete at 15 min at room temperature.
Enzyme Assay--
All activity measurements were performed at
25 °C. For XO, xanthine-O2 activity was measured
spectrophotometrically in terms of the absorbance change at 295 nm (
= 9500 M
1 cm
1) in 50 mM sodium pyrophosphate buffer (pH 8.5) containing 0.2 mM EDTA and 0.15 mM xanthine under
air-saturated conditions. The electron transfer activity from xanthine
or allopurinol (0.15 mM) to 2,6-dichlorophenolindophenol
(DCPIP; 50 µM) was determined spectrophotometrically by
monitoring the absorbance of DCPIP at 600 nm (
= 16,100 M
1 cm
1) in 50 mM
potassium phosphate buffer (pH 7.8) under air.
For XDH, all activity measurements were conducted in 50 mM
sodium pyrophosphate buffer (pH 8.5) containing 0.2 mM
EDTA. Xanthine-O2 activity was measured as described above
for XO. Xanthine-NAD+ activity was measured with 0.15 mM xanthine and 0.5 mM NAD+. For
the measurements of electron transfer activities between xanthine and
DCPIP, NADH and DCPIP, NADH and ferricyanide, and NADH and methylene
blue, 0.15 mM xanthine or NADH and 50 µM
electron acceptor (DCPIP, potassium ferricyanide, or methylene blue)
were used.
Anaerobic Treatment of Enzymes with Nitric Oxide--
Each
solution below was prepared with 0.2 M Hepes buffer
containing 1 mM EDTA (pH 7.0). Anaerobic XO or XDH (40-80
µM) samples were prepared in an all-glass apparatus by
sequential evacuation and re-equilibration with oxygen-free argon.
Oxygen-free argon was prepared by passing commercially obtained pure
argon through a column of Oxyout (Osaka Sanso). The solution of an
electron donor such as xanthine, allopurinol, or NADH was sealed in a
glass vial tube with a rubber cap and made anaerobic by bubbling
oxygen-free argon. Saturated nitric oxide solution was prepared by
bubbling NO gas, which was passed through 5 M KOH to remove
NO2, into the anaerobic buffer.
To prepare nitric oxide-treated enzymes, XO and XDH were first reduced
by mixing enzyme solution into the electron donor solution anaerobically. The final concentrations of XO, XDH, and
xanthine/allopurinol were 4-15, 10, and 150-300 µM,
respectively. Then, saturated NO solution was added so that the final
concentration became 250-500 µM. The mixture was drawn
into a gas-tight syringe, incubated for 30 min at 25 °C, and
filtered through a small column of Sephadex G-25. To check the effects
of electron acceptors such as methylene blue (50 µM) and
potassium ferricyanide (50 µM) during NO treatments, xanthine and the electron acceptor were mixed and made anaerobic before
being mixed with enzyme solution.
To examine the kinetics of the reaction between XDH/XO and nitric
oxide, various amounts of saturated NO solution were added to enzyme
solutions that had been pre-reduced anaerobically with xanthine or NADH
(0.15 mM). The final concentrations of XO and XDH were 4 and 10 µM, respectively. The mixture was vigorously mixed
and immediately drawn into a gas-tight syringe to avoid escape of NO
into the gas phase and to maintain a constant NO concentration. The NO
concentration of the reaction mixture in the syringe was immediately
determined in duplicate by the oxyhemoglobin method. At various time
points, 10-µl aliquots of the mixture were used to measure
xanthine-O2 activity for XO and xanthine-NAD+
reductase activity for XDH.
Aerobic Treatment of Enzymes with Nitric Oxide--
XO (AFR > 200) or XDH (AFR = 109.00) in 0.2 M Hepes buffer
(pH 7.4) containing 1 mM EDTA and 1 mM NADH was
mixed with saturated NO solution, which was prepared by bubbling NO gas
into the anaerobic buffer, under air at 25 °C. The enzyme
concentration and initial NO concentration of the mixture were 5 and
470 µM, respectively. Control experiments were conducted
under the same conditions by mixing with anaerobic 0.2 mM
Hepes buffer (pH 7.4) containing 1 mM EDTA instead of
saturated NO solution. After having been incubated under air for 20 min, the mixture was gel-filtered, and aliquots were used for the
determination of xanthine-oxygen or xanthine-NAD+ reductase activity.
EPR Measurements--
EPR measurements were conducted on a Jeol
JES-FE2XG spectrometer. Sample temperature was controlled by a variable
temperature controller above 77 K and by a continuous flow cryostat
system (CT-470-esr-1, Research and Manufacturing Co., Inc.) below 77 K. The samples used to measure the Mo(V) signals of native XO, NO-treated
XO, and desulfo-XO were prepared by reducing the different forms of XO
based on reported procedures (32), and these EPR spectra were acquired
at 123 K. To obtain the "very rapid" signal, enzymes (15 µM) in 20 mM Ches buffer (pH 10.2) were
anaerobically reduced with 6-methylpurine (5 mM) for 1 min,
and 0.5 ml of the solution was frozen 20 s after having been
reoxidized by bubbling O2 through the solution. To obtain
the "slow" or "rapid type 1" signal, enzymes (15 µM) in 20 mM Bicine buffer (pH 8.2) were made anaerobic by bubbling argon gas for 20 min, and 0.5 ml of the solution
was frozen in an EPR sample tube 1 min after the addition of excess
sodium dithionite. The EPR spectra of iron-sulfur centers (33) were
obtained using the same sample as prepared for the slow signal and
measured at 22 K. The conditions of the EPR measurements were as
follows: microwave frequency, 9.2 GHz; response, 0.3 s; microwave
power, 10 milliwatts; field modulation width, 0.2 millitesla; and sweep
time, 2 or 4 min. The magnetic field was calibrated with the signal of
an external Mn2+ (in MgO) standard. At this microwave
frequency, the apparent g values of the third and fourth
Mn2+ signals are 2.033 and 1.981, respectively. EPR spectra
were acquired through an A/D converter board and processed with an
IBM-PC compatible computer.
Reactivation of NO-treated XO and Desulfo-XO--
Both
desulfo-XO (AFR = 0) and NO-treated XO (AFR = 4.6) (4.6 µM) were reactivated using sodium thiosulfate and
rhodanese as previously reported (34).
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RESULTS |
Effects of NO on Xanthine Oxidase--
NO completely inactivated
XO when XO was reduced with xanthine or allopurinol under anaerobic
conditions prior to the NO treatment, although no marked decrease in
the activity was observed when XO was not pre-reduced with xanthine or
allopurinol, as shown in Table I. This is
consistent with the previous report that XO was inactivated by NO
during enzyme turnover of the xanthine oxidase reaction (19). The fact
that the activity of NO-treated XO did not recover upon gel filtration
with Sephadex G-25 indicates that the inhibition is an irreversible
process. In the presence of an electron acceptor such as methylene blue
(50 µM) or potassium ferricyanide (50 µM),
XO was not markedly inhibited by NO even in the presence of xanthine
(data not shown), in accordance with the result that only reduced XO
can be inhibited by NO. As NO-treated XO lost both
xanthine-O2 and xanthine-DCPIP activities, it is likely
that NO reacted with the molybdenum center since DCPIP can directly
accept an electron from reduced molybdenum (35). It should be noted
that the extent of inhibition of XO pre-reduced with allopurinol is
lower than that of XO pre-reduced with xanthine, although NO can also
inhibit XO pre-reduced with allopurinol. This result is consistent with
the conclusion that NO attacks XO at the molybdenum center since a
small amount of oxypurinol, the oxidative product of allopurinol, might
partially form a tight-binding complex with the reduced molybdenum (24)
to protect the enzyme from inactivation by NO.
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Table I
Inactivation of XO by NO under anaerobic conditions
XO was treated anaerobically with 250 µM NO for 30 min at
25 °C, followed by gel filtration as described under "Experimental
Procedures." The activities (measured in duplicate), are based on the
rates of electron transfer from xanthine or allopurinol to O2
or DCPIP and are expressed as mean percentage of the control (before
treatment).
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Kinetics of the Reaction between XO and Nitric Oxide--
To
clarify further the characteristics of the reaction between XO and NO,
the reaction kinetics were determined. XO (4 µM) was
pre-reduced with xanthine (150 µM) and treated with
various concentrations of nitric oxide anaerobically at 25 °C, and
the time course of the xanthine-oxygen reductase activity was
determined. As shown in Fig.
1A, the XO activity was
inhibited dose-dependently by NO. We presumed that XO
reacts with NO according to second-order kinetics as follows (Equations
1 and 2),
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(Eq. 1)
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(Eq. 2)
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where XO denotes the active form, XO* denotes the inactive form,
and k is the second-order rate constant for the reaction. When [XO]
[NO], k' = k[NO] can be
assumed to be constant, and the reaction should follow pseudo
first-order kinetics. Since the time courses of enzyme inhibition
depicted in Fig. 1A showed a single exponential form, a
pseudo first-order rate constant (k') could be determined at
various NO concentrations. Fig. 1B shows the dependence of
k' on nitric oxide concentration. As expected, k'
was proportional to nitric oxide concentration, indicating that the
reaction of XO with nitric oxide follows second-order kinetics as
formulated in Equations 1 and 2. The second-order rate constant
(k) obtained from the slope in Fig. 1B is
14.8 ± 1.4 M
1 s
1
(25 °C).

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Fig. 1.
Inhibition of xanthine oxidase activity by
NO. A, XO (4 µM) was pre-reduced with
xanthine (150 µM) and treated anaerobically with NO at
25 °C. The vertical axis indicates remaining activity
expressed as a ratio to the original (0 time) activity for each plot.
Xanthine-oxygen reductase activity was measured by following the
absorbance change at 295 nm. The concentrations of NO (19, 72, and 131 µM) are indicated. For simplicity, only three experiments
at different NO concentrations are shown. B, shown are plots
of pseudo first-order rate constant (k') versus
NO concentration. The second-order reaction rate constant was
calculated to be 14.8 ± 1.4 M 1
s 1.
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Spectroscopic and Redox Properties of NO-inactivated XO--
The
initial spectrum obtained from the fully oxidized form of
NO-inactivated XO is very similar to that of the native enzyme (Fig.
2). Since the absorption in the visible
region is mainly attributable to the flavin and iron-sulfur centers, it
is suggested that these cofactors remain intact. Upon addition of
xanthine to the inactivated enzyme under anaerobic conditions,
negligible immediate bleaching of the visible absorbance at around 450 nm was observed, but gradual bleaching was observed after prolonged incubation; only <20% reduction occurred even after 4 h. Such a
biphasic reduction mode is very similar to that of the desulfo-type inactive enzyme (25). The second phase of reduction might be due to
slow reduction of the inactive enzyme by a small amount of the active
enzyme remaining in the sample. Although bleaching of the visible
absorbance at around 450 nm of the NO-inactivated enzyme was very slow,
it occurred readily upon addition of dithionite as in the case of the
native enzyme. As shown in Fig. 2, the flavin and iron-sulfur centers
in NO-treated XO can be reduced with dithionite. Moreover, the fully
reduced inactive enzyme was reoxidized readily when exposed to air, and
the spectrum returned to that of the initial oxidized form (data not
shown), indicating that the redox function of the flavin and
iron-sulfur centers in NO-treated XO was preserved. Fig.
3 shows the EPR spectra of iron-sulfur
centers of dithionite-reduced XO before and after treatment with NO. It has been shown that the fully reduced active enzyme exhibits EPR signals due to two distinct Fe2S2 centers (33).
Native and NO-treated XO showed essentially the same signals due to the
iron-sulfur centers (gav = 1.95 and 2.01, respectively),
and there was no indication that NO interacted with the iron-sulfur
center of NO-treated XO. These spectroscopic observations suggested
that NO reacted with neither flavins nor iron-sulfur centers, but
rather with the molybdenum center.

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Fig. 2.
Absorption spectra of NO-treated XO. XO
treated with NO as described in the legend to Table I was used for
experimentation. Trace 1, NO-treated XO (before addition of
xanthine) in 0.2 M Hepes buffer containing 1 mM
EDTA (pH 7.0); trace 2, immediately after addition of 50 µl of 1 mM xanthine in 0.01 M NaOH to 1 ml of
NO-treated XO under anaerobic conditions (the spectrum was not
corrected for dilution); trace 3, the same sample as
trace 2 after prolonged incubation (took spectrum at 4 h after addition of xanthine); trace 4, intermediate
spectrum after partial addition of sodium dithionite to the same
preparation of NO-treated XO; trace 5, NO-treated XO
completely reduced with sodium dithionite.
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Fig. 3.
EPR spectra of iron-sulfur centers of XO
reduced with dithionite. The EPR spectra of dithionite-reduced
iron-sulfur centers were obtained at 22 K for native XO (trace
a) and NO-treated XO (trace b) (15 µM) in
20 mM Bicine buffer (pH 8.2) under the conditions described
under "Experimental Procedures." mT, milliteslas.
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Molybdenum EPR Spectra of NO-treated XO--
All of the above
results indicate that the inactivation of XO by nitric oxide occurs at
the molybdenum center. To confirm this conclusion and to clarify what
kind of change occurs at the molybdenum center, the EPR spectrum of
NO-treated XO was measured and compared with those of the native and
desulfo-type enzymes. First, the EPR spectrum of NO-treated XO was
observed without treatment with reducing reagents. Although XO in the
resting state, which has Mo(VI) and two antiferromagnetically coupled
Fe(III)2S2 clusters, is EPR-silent, it can be
paramagnetic if the NO adduct of XO forms at either iron center to
produce Fe(II)-nitrosyl heme complexes. However, no paramagnetic signal
other than a weak signal due to a non-heme iron impurity at g = 4.3 was observed (data not shown). Thus, no NO adduct was observable in
the oxidized state of NO-treated XO. Fig.
4 shows the EPR spectra of native and
NO-treated XO reduced under conditions where the very rapid signal can
be obtained. As shown in Fig. 4 (trace a), native XO showed
a Mo(V) very rapid signal that has large g anisotropy when reduced with
6-methylpurine for a short time as reported previously (32). The
species that gives the very rapid signal has a very short lifetime if
xanthine is used as a substrate and corresponds to a transient
intermediate Mo(V)-substrate complex in enzyme turnover (36). Thus, the
observation of the very rapid signal provides evidence of reaction
between molybdopterin and the substrate. However, as shown in Fig.
4B, no EPR signal was observed for the NO-treated enzyme
under the same conditions, indicating that molybdopterin in NO-treated
XO has no ability to interact with purine derivatives and that NO
damages the molybdopterin.

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Fig. 4.
Mo(V) EPR spectra of XO reduced with
6-methylpurine. Native XO (trace a) and NO-treated XO
(trace b) (15 µM) in 20 mM Ches
buffer (pH 10.2) were anaerobically reduced with 6-methylpurine (5 mM) for 1 min, and 0.5 ml of the solution was frozen
20 s after reoxidation with bubbled O2. The EPR
spectra were recorded at 123 K. mT, milliteslas.
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Fig. 5 shows Mo(V) EPR spectra of native
XO (trace a), NO-treated XO (trace b), and
CN-treated desulfo-XO (trace c) (15 µM) in 20 mM frozen Bicine buffer (pH 8.2) reduced with dithionite for 1 min. The native enzyme exhibits a rapid type 1 signal (32), as
shown in Fig. 5 (trace a). On the other hand, NO-treated XO showed a different EPR spectrum (Fig. 5, trace b). The
spectrum showed no evidence of an extra hyperfine interaction of the
molybdenum center with NO nitrogen, which should split the signal into
three lines. Thus, the molybdopterin in NO-treated XO had not been
changed into its NO adduct. Furthermore, NO-treated XO showed a
spectrum substantially identical to that of the CN-treated
desulfoenzyme (Fig. 5, trace c), having similar apparent g
values to those reported previously (32). It is known that reduced
desulfo-type xanthine oxidase, which is inactive because an essential
sulfur atom coordinated to the molybdenum is replaced by an oxygen
atom, shows the slow signal due to Mo(V), which is stable under
anaerobic conditions (32). The Mo(V) in reduced NO-treated XO was also
stable under anaerobic conditions. Thus, it is suggested that nitric
oxide reacts with an essential sulfur in molybdopterin and removes
it.

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Fig. 5.
Mo(V) EPR spectra of XO reduced with
dithionite. Native XO (trace a), NO-treated XO
(trace b), and desulfo-XO (trace c) (15 µM) in 20 mM Bicine buffer (pH 8.2) were
anaerobically reduced with dithionite for 1 min, and the EPR spectra of
the frozen solutions (0.5 ml) were recorded at 123 K. mT,
milliteslas.
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Reactivation of CN-treated Desulfo-XO and NO-treated XO--
It
has been demonstrated that desulfo-XO undergoes reactivation by a
sulfide-generating system containing rhodanese, thiosulfate, and
sulfhydryl reagent up to half its maximal activity (34). To determine
whether NO-treated XO can recover its activity by incubation with a
sulfide-generating system, the change in the activity of NO-treated XO
was determined over 3 h, and the time course was compared with
that of the CN-treated desulfoenzyme (Fig.
6). In the presence of sulfide,
NO-treated XO was also reactivated, and its time course was almost the
same as that of CN-treated desulfo-XO. Thus, it was confirmed that
NO-treated XO has the same reactivity to sulfide as CN-treated
desulfo-XO. The recovery of NO-treated XO was slightly low compared
with that of the CN-treated desulfoenzyme, presumably because NO
treatment caused minor inactivation via a route other than conversion
to desulfo-XO.

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Fig. 6.
Reactivation of CN- and NO-treated XO.
Both CN-treated XO (initial AFR = 0; ) and NO-treated XO
(initial AFR = 4.6; ) (4.6 µM) were incubated at
37 °C in 0.1 M sodium pyrophosphate (pH 8.5), 0.2 mM EDTA, 58 mM dithiothreitol, 38 mM sodium thiosulfate, and 0.14 mg/ml rhodanese. The total
volume was 0.5 ml. Aliquots (10 µl) were analyzed for urate
production at the indicated times for the determination of
xanthine-O2 activity.
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Effects of Nitric Oxide on XDH--
Since XDH has the same
molybdopterin as XO, XDH is also expected to be inhibited by NO. As
shown in Fig. 7, XDH (10 µM) reduced with xanthine (0.15 mM) under
anaerobic conditions was inhibited by NO in the same manner as XO,
whereas the oxidized enzyme was not markedly inhibited. Without
xanthine, XDH was not inhibited by NO either. Moreover, XDH reduced
with excess NADH was also inhibited by NO. Since the NADH-DCPIP,
NADH-ferricyanide, and NADH-methylene blue activities of NO-treated XDH
were almost equivalent to those of the native enzyme, as also found for
CN-treated XDH (data not shown), it is concluded that FAD remained
intact. Thus, it is concluded that reduced XDH is inhibited by NO in
the same manner as XO.

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Fig. 7.
Inhibition of XDH by NO. XO (10 µM) was treated with nitric oxide (130 µM)
anaerobically at 25 °C. , no additional treatment; ,
pre-reduced with xanthine; , pre-reduced with NADH. The
vertical axis indicates xanthine-NAD+ reductase
activity measured in terms of uric acid generation rate monitored at
295 nm. The data are expressed as percent activity of native xanthine
dehydrogenase.
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In the presence of enough NADH, XDH is kept in the reduced form even
under aerobic conditions. So, it is possible that XDH is inhibited by
NO in air. In the presence of 1 mM NADH, XO (5 µM) or XDH (5 µM) was mixed with saturated
NO solution so that the initial NO concentration was 470 µM, and the catalytic activities were measured after 20 min. With NO treatment in air, XO activity was not markedly decreased
(93.8 ± 2.8% of untreated XO), whereas XDH activity was almost
completely inhibited (13 ± 0.8% of untreated XDH). Thus, as long
as enough NADH is present, XDH is susceptible to the attack of NO even
under aerobic conditions, where XO is not inhibited.
 |
DISCUSSION |
XO contains multiple redox centers that are essential for enzyme
activity and are plausible targets of NO attack. We examined the effect
of NO on each redox center using enzyme kinetics and spectroscopic
studies. Since NO forms an iron-nitrosyl adduct with iron-sulfur
complexes (12), as was demonstrated in aconitase (37), we first
examined the iron-sulfur centers by means of absorption spectroscopy
and EPR. In NO-treated XO, the iron-sulfur centers were redox-active,
and EPR proved that their environment remained unchanged; no
nitrosyl-iron-sulfur complex was observed. Since aconitase has an
Fe4S4-type cluster with at least one iron coordination with a solvent molecule, it may be more susceptible to NO
attack than the Fe2S2-type clusters with
complete cysteinyl ligation in XO. This is consistent with the fact
that most of the reports on the NO sensitivity of iron-sulfur proteins
refer to Fe4S4 clusters (12, 37), except for a
report on nitrosyl-iron complex formation from
Fe2S2 ferredoxins (38).
Since it seemed that FAD in NO-treated XO was intact, the molybdenum
center was considered the next most likely candidate for NO attack. The
lack of electron transfer activity from xanthine to DCPIP and the very
slow reduction of NO-treated XO by xanthine suggested that the
molybdenum center was damaged. The EPR measurements indicated that NO
reacts with an essential sulfur atom coordinated to the molybdenum
center to afford a desulfo-type enzyme. This was confirmed by the fact
that the NO-inactivated enzyme was reactivated by the
sulfide-generating system, like CN-treated desulfo-XO. Since NO can
react only with the reduced enzyme, i.e.
Mo(IV)-SH, not
with the oxidized form, Mo(VI)=S (1), it seems that sulfhydryl coordinated to molybdenum is more susceptible to NO attack than Mo=S.
Although the chemical mechanism of conversion of the sulfo to desulfo
form by NO is not clear, probably cationic NO might attack the reduced
sulfide to release SNO. It is interesting to note that the desulfo form
of XO exists in a significant amount in rat liver cells (35).
The fact that NO reacts only with reduced XO explains why XO is not
markedly inhibited by NO exposure under aerobic conditions. The
moderate inhibition during enzyme turnover may reflect a comparatively low steady-state level of the reduced enzyme. However, since XO operates under a much lower oxygen tension in vivo, close to
10 mm Hg (39), this restriction does not reduce the potential
importance of this NO inhibition. Furthermore, the reactivity of NO
toward XO in a reducing environment rather reinforces the importance of
this inhibition under hypoxic conditions, where more reducing substrates such as hypoxanthine, xanthine, and NADH might be
accumulated. Our finding that, under aerobic conditions, only XDH was
inactivated in the presence of a reducing substrate, NADH, suggests
that XDH, but not XO, may be inactivated by NO under conditions such as post-ischemic reperfusion, where oxygen is resupplied. Although XDH can
produce a large amount of O
2 without NAD+ during
xanthine-O2 turnover, this pathway should be inhibited since an excess amount of NAD+ is present in the normal
cell (40). Therefore, the conversion of XDH to XO seems to be necessary
to explain the increased production of O
2 in so-called
superoxide-induced injury during ischemia/reperfusion. However, it is
still controversial whether conversion from XDH to XO occurs under
post-ischemic conditions as postulated by McCord (6). The O
2
generation, however, was suggested to increase when the apparent ratio
of XDH to XO changes. That is, as XDH is inhibited by the accumulated
NADH during ischemia, accumulated hypoxanthine is utilized more by XO
than would be the case under normal conditions, and increased
superoxide production may occur upon reperfusion even though conversion
from XDH to XO does not take place (41, 42). The increased O
2
generation can be alternatively explained in terms of NADH oxidation by
XDH (42, 43). The present results suggest another mechanism for change
in the relative contributions of XO and XDH to O
2 generation
if selective inactivation of XDH by NO occurs under post-ischemic
conditions prior to the formation enough O
2. If once
O
2 is generated in a large amount, which can react with NO
with diffusion-limited rates to give peroxynitrite, decreased NO and
formed peroxynitrite may make the situation worse for the tissue. Thus,
although the generation of NO upon reperfusion was suggested to
down-regulate O
2 generation through inhibition of XO by NO
(21), it is clear from the present experiments that the idea that NO
serves as an antioxidant effector by suppressing XO activity in actual
cellular systems is an oversimplification. It should also be noted that
O
2 formation due to NADH oxidation, in which the molybdenum
center does not participate, cannot be inhibited by NO.