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J. Biol. Chem., Vol. 278, Issue 37, 35086-35092, September 12, 2003
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¶ || ¶ 
¶
From the
CNRS-UMR 7000, Faculté de
Médecine Pitié-Salpêtrière, 75013 Paris, France,
the ¶UFR de Biochimie, Université Denis
Diderot-Paris 7, 75005 Paris, France, and the
**Institute of Cancer Research, CRUK Centre for Cell
and Molecular Biology, Fulham Road, London SW3 6JB, United Kingdom
Received for publication, April 11, 2003 , and in revised form, June 26, 2003.
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONEXPERIMENTAL PROCEDURESRESULTSDISCUSSIONREFERENCES |
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONEXPERIMENTAL PROCEDURESRESULTSDISCUSSIONREFERENCES |
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Redox-dependent regulation of catalytic activities by reversible oxidation of an active-site cysteine residue has been reported for several enzymes, including protein phosphatases (18–21) and cysteine proteases (22–24). H2O2 is one of the oxidants that has been shown to regulate cell function, by oxidizing active cysteine residues in proteins to cysteine sulfenic acid or to disulfide (20, 25–27). In vivo, H2O2 formation is mainly due to enzymatic reactions such as the superoxide dismutase-dependent dismutation of superoxide (26, 28, 29). Although superoxide can oxidize proteins (30), H2O2, its main product, has appeared as a critical element involved in many cellular functions, including cell signaling (20, 27, 28, 31). In addition, substantial increases in the intracellular concentration of H2O2 are generally associated with deleterious conditions such as apoptosis, necrosis, inflammation, and cancer (20, 26, 27, 31). The induction of H2O2 production by xenobiotics has also been described, with potential effects on cysteine-containing proteins (32–35). Redox-dependent detoxification and/or activation of xenobiotics also suggests that cellular redox conditions may affect XME-dependent biotransformations (35–38).
We investigated here whether the catalytic activity of human NAT1 was regulated by H2O2. We found that NAT1 activity was reversibly inhibited by physiologically relevant concentrations of H2O2. We demonstrate that H2O2 oxidized rapidly NAT1, leading to oxidative inactivation of the enzyme. This H2O2-dependent inactivation of NAT1 probably results from oxidative modification of the essential catalytic cysteine residue to a stabilized cysteine sulfenic acid, as shown for other enzymes (39–41). The inactivated NAT1 was reactivated by physiological thiols, such as reduced glutathione. Our results suggest that in addition to the polymorphic-dependent variation of NAT1 activity, H2O2 and, more broadly, cellular redox status, could also regulate NAT1 activity, which may have important consequences with regard to drug toxicity and cancer risk.
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EXPERIMENTAL PROCEDURES |
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TOPABSTRACTINTRODUCTIONEXPERIMENTAL PROCEDURESRESULTSDISCUSSIONREFERENCES |
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-D-glucose were purchased from Sigma. PGEX-2T vector and
Escherichia coli BL21(DE3) cells were supplied by Amersham
Biosciences. The pET28 vector was obtained from Novagen.
Nickel-nitrilotriacetic acid superflow resin was purchased from Qiagen.
Anti-fluorescein Fab' fragments conjugated to peroxidase,
fluorescein-conjugated iodoacetamide, and complete protease inhibitor tablets
were obtained from Roche Applied Science. All other reagents were obtained
from Sigma or Eurobio (Les Ulis, France). The Bradford protein assay kit was
purchased from Bio-Rad.
Expression and Purification of Recombinant Human NAT1—The
human NAT1 cDNA, kindly provided by Dr. D. Grant (Toronto, Canada), was
subcloned into pGEX-2T and pET28 vectors. The resulting constructs encoded the
enzyme as a glutathione S-transferase fusion protein (GST-NAT1) and
as a polyhistidine-tagged fusion protein (His-NAT1), respectively. We
transformed BL21(DE3) bacteria with these constructs, induced expression of
the transgene with 0.1 mM
isopropyl-1-thio--D-galactopyranoside, and cultured the cells
for 4 h at 37 °C. Bacteria were pelleted by centrifugation (6,000 x
g, 30 min), washed with cold phosphate-buffered saline, and harvested
by centrifugation (6,000 x g, 30 min). Pellets were stored at
–80 °C until required.
For the purification of recombinant proteins, pellets (from 1-liter cultures) were resuspended in 40 ml of 50 mM Tris-HCl, pH 8, 150 mM NaCl (lysis buffer) containing lysozyme (1 mg/ml final concentration) and protease inhibitors. Following incubation (1 h at 4 °C), protease inhibitors, DNase I (20 µg/ml final concentration), and 0.2% Triton X-100 (final concentration) were added, and the suspension was incubated for an additional hour at 4 °C. Lysates were then subjected to sonication on ice (five pulses of 10 s each), centrifuged (12,000 x g, 30 min), and the supernatant collected.
For the purification of GST-NAT1, this supernatant was added to glutathione-agarose beads (40 mg/liter of original culture) for 2 h at 4 °C. Beads were then poured into a column and washed successively with 5 volumes of lysis buffer containing 0.2% Triton X-100 followed by 5 volumes of lysis buffer alone. GST-NAT1 was then eluted with 10 mM reduced glutathione (final) in 50 mM Tris-HCl, pH 8, 1 mM EDTA. The purified enzyme was reduced by incubation with 10 mM DTT (final concentration) for 10 min at 4 °C and was then dialyzed against 25 mM Tris-HCl, pH 7.5, 1 mM EDTA.
The His-NAT1 protein was prepared in a similar manner to GST-NAT1. Briefly, supernatants from bacterial lysates were incubated with 1.5 ml of nickel-nitrilotriacetic acid Superflow resin in the presence of 20 mM imidazole (final concentration) for2hat4 °C. The resin was then poured into a column and washed successively with lysis buffer containing 0.2% Triton X-100 and lysis buffer containing 50 mM imidazole (final concentration). His-NAT1 was eluted with 300 mM imidazole (final concentration) in lysis buffer. Purified His-NAT1 was reduced by incubation with 10 mM DTT (final concentration) for 10 min at 4 °C and was then dialyzed against 25 mM Tris-HCl, pH 7.5, 1 mM EDTA.
SDS-PAGE analysis was carried out at each stage of purification, and protein concentrations were determined using a standard Bradford assay.
Recombinant-tagged NAT1 enzyme was used in a previous study for functional characterization of the NAT1 enzyme (42). In this study, we used both GST-NAT1 and His-NAT1 as sources of purified recombinant NAT1 enzyme, and similar results were obtained with both proteins in all experiments.
Enzyme Assays—NAT1 enzyme activity was determined spectrophotometrically (410 nm), using PNPA as the acetyl donor and PAS as the NAT1-specific arylamine substrate, as described by Mushtaq et al. (43). Briefly, treated or non-treated samples (10–20 µl) containing NAT1 enzyme were assayed in a reaction mixture containing 500 µM PAS (final concentration) in 25 mM Tris-HCl, pH 7.5, 1 mM EDTA. Reactions were started by adding 125 µM PNPA (final concentration). In all reaction mixtures (total volume of 1 ml), the final concentration of NAT1 was 15 nM. The reaction mixture was incubated for 10 min incubation at 37 °C, and the reaction was then quenched by adding SDS (1% final concentration). p-Nitrophenol, generated by the NAT1-mediated hydrolysis of PNPA in the presence of PAS, was quantified by measuring absorbance at 410 nm with an enzyme-linked immunosorbent assay plate analyzer (Metertech). One unit of enzyme was defined as the amount of enzyme giving an A410 value of 0.5 per 10 min per ml. For the controls, we omitted the enzyme, PNPA, or PAS. All enzyme reactions were performed in quadruplicate, in conditions in which the initial rates were linear. Enzyme activities are shown as percentages of control NAT1 activity.
Reaction of NAT1 with H2O2 in the Presence or
Absence of Other Compounds—Concentrations of stock
H2O2 solutions were determined by measuring absorbance
at 240 nm (
240 = 44
M–1·cm–1). In all
subsequent experiments, the final concentration of enzyme during the oxidation
step with H2O2 was 1.5 µM, giving a final
concentration in the enzyme assay of 15 nM. The total volume of the
enzyme assay (1 ml) provided a great enough dilution (1:50 or 1:100) of the
various compounds used to prevent these compounds from interfering with NAT1
enzyme activity measurements. NAT1 activity in the absence of
H2O2 was taken as 100% activity (control).
We assessed the effect of bolus addition of H2O2 on NAT1 enzyme activity by incubating purified NAT1 samples (1.5 µM final concentration) with various concentrations of H2O2 in 25 mM Tris-HCl, pH 7.5, 1 mM EDTA (total volume of 10 µl) for 10 min at 37 °C. Mixtures were then assayed for NAT1 activity, as described above.
In reactivation experiments, NAT1 (1.5 µM final concentration) was first oxidized by H2O2 (200 µM final concentration), as described above, and incubated with catalase (300 unit/ml) for 1 min at 37 °C. The mixture was then incubated for 10 min at 37 °C with various concentrations of DTT or GSH in a total volume of 20 µl. A NAT1 enzyme assay was then carried out. Assays performed in this conditions (with catalase) but without H2O2 gave 100% NAT1 activity.
In substrate protection experiments, NAT1 (1.5 µM final concentration) was first incubated with various concentrations of AcCoA or CoA (final concentrations of 0.4–4 mM) in 25 mM Tris-HCl, pH 7.5, 1 mM EDTA (total volume of 10 µl) for 5 min at 37 °C. Samples were then incubated with H2O2 (200 µM final concentration) in a total volume of 20 µl, as described above, and assayed. Assays performed in these conditions with AcCoA or CoA alone gave 100% NAT1 activity.
In dimedone assays, NAT1 (1.5 µM final concentration) was first incubated with H2O2 (200 µM final concentration) in the presence of 10 mM dimedone (a specific reagent of sulfenic acid) in 25 mM Tris-HCl, pH 7.5, 1 mM EDTA (10 µl total volume) for 10 min at 37 °C. It was then incubated with DTT (5 mM final concentration, 20 µl total volume) for 10 min at 37 °C. Residual NAT1 activity was assayed as described above. Assays performed in these conditions, with dimedone alone (10 mM final concentration) gave 100% NAT1 activity.
For the kinetic analysis of H2O2-dependent NAT1 inactivation, NAT1 (1.5 µM final concentration) was incubated with H2O2 (final concentration of 100–400 µM)at37 °Cin25mM Tris-HCl, pH 7.5, 1 mM EDTA. At various time intervals, aliquots were removed and assayed for residual activity. The equation for the rate of inactivation of recombinant NAT1 by H2O2 can be represented as: –d[NAT1]/dt = kinact[NAT1] [H2O2], where [NAT1] is the concentration of active enzyme, and kinact is the second-order rate constant. Provided that H2O2 is present in substantial excess, the apparent first-order inactivation rate constants (kobs = kinact[H2O2]) can be calculated for each H2O2 concentration from the slope of the natural log (ln) of percent residual activity plotted against time. The second-order rate constant was determined from the slope of kobs plotted against H2O2 concentrations.
Reaction of NAT1 with H2O2 Generated by Glucose
Oxidase—The effect of continuous generation of
H2O2 by glucose oxidase on NAT1 was carried out as
described previously (44,
45). Quantification of
H2O2 generated by 0.15 unit/ml of glucose oxidase was
determined by measuring the absorbance at 240 nm, and the
peroxidase/o-phenylenediamine dihydrochloride assay as described
elsewhere (44,
46). We assessed the effect of
continuous generation of H2O2 by glucose oxidase by
incubating purified NAT1 (1.5 µM) with glucose oxidase (0.15
unit/ml) and glucose (5 mM) in 25 mM Tris-HCl, pH 7.5, 1
mM EDTA (total volume of 15 µl) at 37 °C. At various time
intervals, catalase (300 units/ml) was added prior to NAT1 residual activity
measurements. Controls where catalase was added to the glucose/glucose oxidase
system were carried out. In addition, controls without glucose oxidase,
without glucose, and without catalase were also done. Assays were all done in
quaduplicate. H2O2 constant production rate in these
experiments was determined to be 
6 µM
H2O2/min.
Fluorescein-conjugated Iodoacetamide Labeling of Proteins—Purified NAT1 (1.5 µM final concentration) was preincubated with or without (control) various concentrations of H2O2 (final concentration from 50 to 400 µM) in 25 mM Tris-HCl, pH 7.5, 1 mM EDTA for 10 min at 37 °C. Samples were then incubated with fluorescein-conjugated iodoacetamide (20 µM final concentration) for 10 min at 37 °C. Samples were then analyzed by SDS-PAGE under reducing conditions followed by Western blotting, using anti-fluorescein Fab' fragments conjugated to peroxidase.
Statistical Analysis—Results are expressed as means ± S.D. of three independent experiments performed in quadruplicate, unless otherwise stated. We used Student's t test, as appropriate, with levels of significance set at p < 0.05 (*) or p < 0.01 (**).
Protein Determination, SDS-PAGE, and Western Blotting—Protein concentrations were determined using a Bradford assay (Bio-Rad). Samples were combined with reducing 4 x SDS sample buffer and separated by SDS-PAGE. Gels were stained with Coomassie Brilliant Blue R-250. For Western blotting, following separation by SDS-PAGE, proteins were electrotransferred onto nitrocellulose membrane. The membrane was blocked by incubation with Tris-buffered saline/Tween 20 (TBS) containing 5% nonfat milk powder for 1 h. Anti-fluorescein Fab' fragments conjugated to horseradish peroxidase (1:100,000) was added and the membrane incubated for 1 h in TBS. The membrane was washed and incubated with Supersignal reagent (Pierce) for detection.
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RESULTS |
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TOPABSTRACTINTRODUCTIONEXPERIMENTAL PROCEDURESRESULTSDISCUSSIONREFERENCES |
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First, reduced NAT1 enzyme (1.5 µM final concentration) was
incubated with various concentrations of H2O2
(5–200 µM final concentration) and residual NAT1 activity
was measured. NAT1 activity was significantly inhibited by
H2O2, in a dose-dependent manner
(Fig. 1A). At a concentration
of 200 µM, H2O2 inhibited the enzyme by
over 90%. An IC50 of 45 µM was obtained for an enzyme
concentration of 1.5 µM. To make a more realistic and
physiologic assessment of the effect of H2O2 on NAT1
activity, we used another source of H2O2, the steady
conversion of -D-glucose to D-gluconolactone and
H2O2, which is catalyzed by glucose oxidase
(45). As shown in
Fig. 1B (filled
circles), NAT1 (1.5 µM final) was also inactivated by the
constant production of H2O2 through the glucose/glucose
oxidase system. In the conditions used, the constant production rate of
H2O2 was estimated to be 6 µM
H2O2/min, which is physiologically relevant
(45,
51). After a 15-min
incubation, residual NAT1 activity was close to 30% of the control. After a
30-min exposure to H2O2, the residual activity was less
than 10%. No inactivation of NAT1 by H2O2 generated by
the glucose/glucose oxidase system was observed in presence of catalase (300
units/ml) (Fig. 1B,
open circles). Thus, these experiments suggest that NAT1 enzyme is
inactivated by bolus addition and constant generation of physiologically
relevant levels of H2O2.
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Detection of H2O2-oxidized Cysteine Residues by Labeling with Fluorescein-conjugated Iodoacetamide—We investigated whether NAT1 contained reactive cysteine residues susceptible to oxidation by physiological concentrations of H2O2, using an approach based on the labeling of cysteine with 5-iodoacetamidofluorescein (25). Incubation of NAT1 with various concentrations of H2O2 resulted in the dose-dependent modification of cysteine residues, as indicated by the disappearance of fluorescein-conjugated iodoacetamide labeling (Fig. 2). Thus, NAT1 cysteine residues are modified by H2O2.
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Reactivation of H2O2-inactivated NAT1 by
Thiol-reducing Agents—We investigated whether the
H2O2-dependent inactivation of NAT1 could be reversed by
thiol-reducing agents, as reported for other enzymes
(18–20,
23). NAT1 (1.5
µM final concentration) was first inactivated by incubation with
H2O2 (200 µM final concentration), and
excess H2O2 was removed by catalase (300 units/ml).
Inactivated NAT1 was then incubated with DTT or GSH, at various concentrations
(1, 5, or 10 mM final concentration), and NAT1 activity was
determined (Fig. 3). Both DTT
and GSH reactivated H2O2-inactivated NAT1
(Fig. 3). At a concentration of
5 mM DTT, the H2O2-dependent inactivation of
NAT1 was completely reversed. In contrast, 5 mM GSH gave 60% of
the NAT1 control activity. A final concentration of 10 mM GSH was
able to recover 100% of the original NAT1 activity. Thus, the
H2O2-dependent inactivation of NAT1 is reversible. Our
results also suggest that the H2O2-inactivated NAT1
enzyme could be reactivated by GSH.
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Inhibition Kinetics and Stoichiometry—We carried out kinetic analysis of the H2O2-dependent inactivation of NAT1 in the presence or absence of various concentrations of H2O2. Semilogarithmic plots of percent residual activity versus time for various concentrations of H2O2 gave straight lines, indicating that inactivation obeyed apparent first-order reaction (Fig. 4A). Replotting the observed pseudo-first-order rate constants (kobs) against H2O2 concentrations gave a straight line that passed very close to the origin (Fig. 4B), consistent with a single-step reaction in which the reverse rate (obtained from the y intercept) was close to zero. The slope gave the apparent second-order rate constant (kinact) of H2O2-dependent NAT1 inactivation as 420 M–1·min–1. For each H2O2 concentration, the first-order rate constant can be expressed as kobs = kinact[H2O2]n, where n is the order of H2O2 in its reaction with NAT1. Thus, replotting ln of kobs versus ln of H2O2 concentration gave a straight line (Fig. 4C). The slope was n = 0.87, indicating that the inactivation of NAT1 by H2O2 involved 1:1 stoichiometry. Thus, the oxidative inactivation of NAT1 by H2O2 is a rapid bimolecular process in which one molecule of H2O2 modifies the catalytic cysteine residue, leading to inactivation of the enzyme.
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Identification of the Site Modified by H2O2 during NAT1 Inactivation—We then investigated whether the active-site cysteine of NAT1 was the target of H2O2-dependent oxidative inactivation. We included AcCoA (physiological acetyl-donor substrate of NAT1), which forms a covalent acetyl-enzyme intermediate (49), in the reaction to provide protection from inactivation by H2O2. CoA, a product of AcCoA hydrolysis that does not form an acetyl-enzyme intermediate, was used as a control. AcCoA conferred dose-dependent protection (up to 64) of NAT1 from H2O2-induced inactivation, whereas CoA did not (Table I). Of the five cysteine residues present in NAT1, only the active site cysteine has been shown to be conserved in all known NAT sequences and to be critical for enzyme function (5, 7, 48). Although we cannot rule out the possibility that H2O2 modifies other cysteine residues of NAT1, these results clearly demonstrate that the catalytic active site cysteine residue of NAT1 is a target of H2O2-dependent oxidative modification, leading to reversible inactivation of the enzyme.
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Determination of the Chemical Nature of the H2O2-oxidized Active-site Cysteine of NAT1—The results reported above led us to investigate the chemical nature of the H2O2-modified catalytic cysteine of the NAT1 enzyme. As H2O2-dependent oxidative inactivation of NAT1 was fully reversible, the oxidized active site cysteine was unlikely to be in the form of a sulfinic (–SO2H) or sulfonic acid (–SO3H), neither of which could be reduced by thiol-reducing agents (27, 40). There were two other possibilities. The active-site cysteine residue of the H2O2-inactivated enzyme may be involved in an inter- or intramolecular disulfide bond or form a stable cysteine sulfenic acid (–SOH), any of which could be reduced to give cysteine. Intermolecular and intramolecular disulfide bonds were ruled out on the basis of electrophoretic mobility shift assays (54). In addition, the catalytic cysteine residue of human NAT1 has been predicted to be at the base of the active-site pocket and inaccessible to other cysteine residues within the same molecule (15). Dimedone has been shown to react specifically with sulfenic acids to form a stable thioether product that cannot be reduced by thiol-reducing agents (22). This compound has been shown to identify cysteine sulfenic acids at the active sites of various enzymes (22, 55, 56). We investigated whether the H2O2-dependent inactivation of NAT1 resulted from the formation of a stable sulfenic acid at the active site cysteine by incubating dimedone (10 mM final concentration) with H2O2-oxidized NAT1 (Fig. 5). The control, in which NAT1 was incubated with 10 mM dimedone alone, showed no inhibition of activity (data not shown). The enzymatic activity of the NAT1 sample treated with H2O2 alone was fully restored by incubation with 5 mM DTT (final concentration). In contrast, NAT1 cotreated with H2O2 and dimedone was only partially reactivated (63% of control activity), consistent with the formation of a stable sulfenic acid at the active site cysteine. The partial, but significant, effect of dimedone on H2O2-treated NAT1 was probably due to the low reactivity of dimedone with sulfenic acids. This partial effect of dimedone has been reported elsewhere for incubations for short periods of time (<1 h) at a final concentration of 10 mM (22, 56).
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONEXPERIMENTAL PROCEDURESRESULTSDISCUSSIONREFERENCES |
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We provide here chemical and kinetic evidence to support that human NAT1
activity is regulated by H2O2. Human NAT1 activity was
significantly inactivated by both the bolus addition and the constant
generation of physiologically relevant levels of H2O2 as
reported, in similar conditions, for certain proteintyrosine phosphatases
(PTPs), in particular the isoform 1B (PTP1B)
(18,
20,
57). Indeed, in experimental
conditions similar to ours (similar enzyme and H2O2
concentrations, incubation times), IC50 values ranging from 60 to
100 µM were reported for inactivation of these PTPs by
H2O2
(18,
20,
57). These values are very
close to the IC50 determined for the inactivation of NAT1 by
H2O2 (IC50 = 45 µM). Moreover,
kinetic analysis showed the rapid oxidation of NAT1 by
H2O2, with a second-order rate constant
(kinact) for enzyme inactivation of 420
M–1·min–1
(Fig. 5). Interestingly, this
value is, again, very close to the kinact constants
(600
M–1·min–1)
reported for the PTPs mentioned above, the activities of which are regulated
in vivo by H2O2
(18,
19,
57). Similar
kinact values were also obtained for caspase 3, a
redox-regulated cysteine protease involved in apoptosis
(23,
24).
The inactivation of NAT1 by H2O2 was fully reversed by the non physiological thiol reductant DTT and by physiological concentrations of GSH, showing that H2O2-dependent inactivation of NAT1 is reversible. In contrast, physiological concentrations (up to 10 mM) of oxidized glutathione (GSSG) had no effect on NAT1 activity, suggesting that GSSG is unlikely to regulate NAT1 activity in vivo. GSH, a cellular reductant, is the major determinant of cellular redox potential, with a concentration of 1–10 mM (52, 53). Thus, our results suggest that GSH could reactivate H2O2-inactivated NAT1 in the cell. GSH levels can be decreased by oxidative stress, in particular by xeniobiotic-induced oxidative stress, with potential effects on the biotransformation activity of the phase II XME glutathione S-transferase (35).
Our findings are consistent with the specific oxidation of cysteine or methionine residues. Kinetic analysis (Fig. 5C) and protection experiments with AcCoA clearly showed that the essential active-site cysteine of NAT1 was the specific target of H2O2-dependent inactivation and that cysteine thiolate was the reactive species. Thiolates are much more susceptible to oxidation by H2O2 in cells than other protein cysteine residues as they are intrinsically stronger nucleophiles (19, 20, 58). Enzymes that contain an essential thiolate in their active site are widely accepted to be potential candidates for reversible oxidation by H2O2 generated within the cell (20, 25, 31). For instance, PTP1B, a phosphatase mentioned above, has been shown to be reversibly inactivated in vitro and in vivo by H2O2, via oxidation of its catalytic cysteine residue (18, 57).
H2O2 can oxidize cysteine residues in proteins to give cysteine sulfenic acid or disulfide, which can be reduced back to cysteine by cellular reductants, such as GSH (19, 20). We provide strong evidence that the H2O2-dependent inactivation of NAT1 involved the formation of a stable sulfenic acid at the active-site cysteine (Cys-SOH) of the enzyme. Interest in this oxidative modification of cysteine residues has increased, since it was shown to play an important role in redox-regulated processes (41). More specifically, the formation of stable sulfenic acid at the active-site cysteine of various enzymes, such as tyrosine phosphatase and glutathione reductases, has been suggested to play a key role in the reversible inhibition of these enzymes during oxidative or nitrosative stress (39, 40). The stability of Cys-SOH in proteins depends mainly on the presence of an apolar microenvironment around the Cys-SOH and the absence of proximal cysteine residues (39). Interestingly, the active-site pocket of NAT enzymes, and more specifically human NAT1 and NAT2, has been reported to be apolar with no other cysteine residue being proximal to the catalytic cysteine of these enzymes (2, 8, 15, 16, 59). Thus, the formation of a stable sulfenic acid at the active-site cysteine residue seems to be a plausible mechanism for the reversible inactivation of human NAT1 by H2O2. Conversely, formation of an inter- or intramolecular disulfide formation similar to that observed in some H2O2-regulated proteins (18, 60) is unlikely. First, in NAT family, only the catalytic cysteine is absolutely conserved. Second, the catalytic cysteine residue of human NAT1 is buried at the base of the active-site pocket with no other proximal cysteine residue (15). Third, no electrophoretic mobility shifts were observed between reduced and H2O2-oxidized NAT1 (data not shown). Finally, dimedone experiments demonstrated the presence of a stable sulfenic acid after H2O2 inactivation.
Our results suggest that human NAT1 could be reversibly inactivated in vivo by H2O2, as shown for other enzymes such as PTPs. Given the importance of oxidative stress in the biotransformation and/or toxicity of many xenobiotics, the inactivation of human NAT1 by H2O2 may be of physiological significance. Our data suggest that, in addition to polymorphic variation of the NAT1 gene (61), redox conditions could regulate NAT1 functional activity. This supports recent reports (62, 63) suggesting that non-genetic factors, such as substrate-dependent inhibition, may also contribute to overall NAT1 activity.
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FOOTNOTES |
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Holds a Ph.D. fellowship from le Ministère de la Jeunesse, de
l'Education Nationale et de la Recherche.
|| Holds a postdoctoral fellowship from Université Paris 7-Denis
Diderot.
The last two authors share senior authorship.
To whom correspondence should be addressed: CNRS-UMR 7000, Faculté de
Médecine Pitié-Salpêtrière, 75013 Paris, France.
Tel.: 33-1-53-60-08-03; Fax: 33-1-53-60-08-02; E-mail:
rlima{at}ext.jussieu.fr.
1 The abbreviations used are: XME, xenobiotic metabolizing enzyme; NAT1,
human arylamine N-acetyltransferase 1; PAS, p-aminosalicylic
acid; PNPA, p-nitrophenylacetate; AcCoA, acetyl-coenzyme A; DTT,
dithiothreitol; GST, glutathione S-transferase; PTP, protein-tyrosine
phosphatase.
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ACKNOWLEDGMENTS |
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REFERENCES |
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TOPABSTRACTINTRODUCTIONEXPERIMENTAL PROCEDURESRESULTSDISCUSSIONREFERENCES |
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, 100 µM; 
, 200
µM; 
, 400 µM. Error bars indicate
S.D. values. B, determination of second-order rate constant
(kinact) by plotting the apparent first-order inactivation
constant (kobs) values versus the
H2O2 concentrations (correlation factor
r2 = 0.99). Error bars indicate S.D. values.
C, determination of reaction order by replotting the ln of
kobs versus the ln of H2O2
concentration (correlation factor r2 = 0.99). Error
bars indicate S.D. values.