(Received for publication, October 2, 1996, and in revised form, December 17, 1996)
From the Division of Pulmonary/Critical Care Medicine, Department of Internal Medicine, University of California, Davis, California 95616 and the § Neurodegenerative Diseases Research Center, Pharmacology Group, King's College, University of London, Manresa Rd., London SW3 6LX, United Kingdom
Involvement of peroxynitrite
(ONOO) in inflammatory diseases has been implicated
by detection of 3-nitrotyrosine, an allegedly characteristic protein
oxidation product, in various inflamed tissues. We show here that
nitrite (NO2
), the primary
metabolic end product of nitric oxide (NO·), can be oxidized by
the heme peroxidases horseradish peroxidase, myeloperoxidase (MPO), and
lactoperoxidase (LPO), in the presence of hydrogen peroxide
(H2O2), to most likely form
NO·2, which can also contribute to
tyrosine nitration during inflammatory processes. Phenolic nitration by
MPO-catalyzed NO2
oxidation is
only partially inhibited by chloride (Cl
), the presumed
major physiological substrate for MPO. In fact, low concentrations of
NO2
(2-10 µM)
catalyze MPO-mediated oxidation of Cl
, indicated by
increased chlorination of monochlorodimedon or 4-hydroxyphenylacetic
acid, most likely via reduction of MPO compound II.
Peroxidase-catalyzed oxidation of
NO2
, as indicated by phenolic
nitration, was also observed in the presence of thiocyanate
(SCN
), an alternative physiological substrate for
mammalian peroxidases. Collectively, our results suggest that
NO2
, at physiological or
pathological levels, is a substrate for the mammalian peroxidases MPO
and lactoperoxidase and that formation of
NO2· via peroxidase-catalyzed
oxidation of NO2
may provide an
additional pathway contributing to cytotoxicity or host defense
associated with increased NO· production.
Nitric oxide (NO·)1 is produced
by a wide variety of cell types by both constitutive and inducible
nitric oxide synthases (1) and has many physiological functions ranging
from regulation of vascular tone to neurotransmission and modulation of
inflammatory processes (2). Induction of NO· synthesis during
inflammatory processes represents a defense mechanism against invading
microorganisms, but excessive formation of NO· has also been
implicated in host tissue injury (2-4). Although NO· is a free
radical, it has selective reactivity and reacts predominantly with
other paramagnetic species, including ferrous or ferric iron in heme
proteins or iron-sulfur centers, and other radical species such as
molecular oxygen (O2), superoxide anion (O2), and
lipid or protein radicals (5-9). Autoxidation of NO· by
reaction with O2 results in the formation of nitrite
(NO2
) as the primary end-product (10),
although at physiological concentrations of NO· and
O2, this reaction may be too slow to be of major importance in vivo (4, 6). In the vascular system, NO· is
rapidly oxidized by reaction with oxyhemoglobin (HbO2),
resulting in formation of methemoglobin (Hb3+) and nitrate
(NO3
) (11). NO· also reacts
with Hb3+ to form a complex (Hb-NO), which can hydrolyze to
Hb2+ and NO2
(reviewed in
Ref. 12). NO· also reacts at a near diffusion-limited rate with
O
2 to yield peroxynitrite (ONOO
), a powerful
oxidizing species, and many of the cytotoxic properties of NO·
have in fact been attributed to the formation of ONOO
(4). Reaction of ONOO
or its conjugate acid, ONOOH, with
a wide variety of biomolecules also results in concomitant production
of NO2
(7). Hence, irrespective of the
mechanisms involved, NO2
is a major
oxidation product derived from NO·, and increased
NO2
levels can often be detected in
situations where NO· production is elevated.
In healthy human subjects, NO2 can be
detected at levels of 0.5-3.6 µM in plasma (13, 14),
~15 µM in respiratory tract lining fluids (15), 30-210
µM in saliva, and 0.4-60 µM in gastric juice (16). Oral NO2
levels are
increased dramatically to near millimolar levels after ingestion of
nitrate (NO3
) because of
NO3
reduction by the oral microflora
(17). Extracellular NO2
levels are
also markedly increased during inflammatory processes, reflecting
increased NO· production. For instance, increased
NO2
levels have been detected in
synovial fluids of patients with rheumatoid arthritis (18), and serum
NO2
levels of 36 µM have
been reported in human immunodeficiency virus-infected patients with
interstitial pneumonia (19), dramatically higher than normal serum
NO2
levels. Moreover, increased
NO2
levels have been detected in
condensed exhalates from patients with asthma compared with those of
healthy subjects (20), which is in accordance with increases in expired
NO· by asthmatics compared with healthy control subjects
(21).
Although NO2 is a major end product of
NO· metabolism, it does not accumulate in vivo but is
rapidly oxidized to NO3
(22).
NO2
can be oxidized by
HbO2 or oxymyoglobin to form methemoglobin or metmyoglobin
and NO3
(23, 24), and catalase has
also been demonstrated to contribute to
NO2
oxidation (25, 26). However,
NO2
can also be oxidized by
inflammatory oxidants such as hypochlorous acid (HOCl) (27, 28), and we
have recently discovered that oxidation of
NO2
by HOCl results in the production
of reactive nitrogen intermediates (29). Furthermore,
NO2
can also be oxidized by heme
peroxidases in the presence of hydrogen peroxide
(H2O2) (30-32), and it has been suggested that
reactive nitrogen intermediates are produced during such processes
(32). Oxidation of NO2
by such
mechanisms could be of importance at loci of inflammatory immune
processes, when NO· and NO2
levels are enhanced and myeloperoxidase (MPO) and/or eosinophil peroxidase are secreted from activated granulocytes.
The purpose of the present study was to investigate
NO2 oxidation by heme peroxidases and
to assess its potential physiological importance. The results indicate
that MPO and other peroxidases can catalyze oxidation of
NO2
, to most likely form
NO2· as an intermediate product. At
physiological or pathological levels,
NO2
can act as a substrate for MPO and
lactoperoxidase (LPO), even in the presence of chloride
(Cl
) and thiocyanate (SCN
), the proposed
major physiological substrates for these peroxidases. Hence, formation
of reactive nitrogen intermediates via peroxidase-catalyzed oxidation
of NO2
could represent an important
contributing mechanism to NO·-mediated toxicity.
Sodium nitrite (NaNO2),
DL-tyrosine, 3-nitrotyrosine, 4-hydroxyphenylacetic acid
(HPA), 4-hydroxy-3-nitrophenylacetic acid (3-NO2-HPA),
4-hydroxy-3-chlorophenylacetic acid (3-Cl-HPA), 4-methoxybenzoic acid,
4-methoxy-3-nitrobenzoic acid, 5,5-dithiobis(2-nitrobenzoic acid)
(DTNB), monochlorodimedon, hydrogen peroxide (30%), bovine serum
albumin (essentially fatty acid-free), diethylenetriaminepentaacetic acid (DTPA), D-glucose, glucose oxidase (type V-S),
catalase (from bovine liver; 25,000 units/mg protein), horseradish
peroxidase, and lactoperoxidase (from bovine milk; 120 units/mg
protein) were purchased from Sigma. Myeloperoxidase (from human
leukocytes; 250-300 units/mg protein) was obtained from Alexis Corp.
(San Diego, CA). All peroxidases were used without further
purification. 3,3
-Dityrosine and 3,3
-bis(4-hydroxyphenylacetic acid)
(3, 3
-diHPA) were synthesized by reaction of DL-tyrosine
or HPA with horseradish peroxidase and hydrogen peroxide
(H2O2) (33). All other reagents were of the
highest purity commercially available.
Initial studies were performed with
horseradish peroxidase (HRP). DL-Tyrosine (1 mM) was dissolved in 50 mM sodium phosphate buffer (pH 7.4) containing 100 µM DTPA, 1 µM HRP, and various concentrations of NaNO2.
DTPA was included in the reaction mixture to avoid interfering
reactions with contaminating free metal ions. Reactions were initiated
by addition of 1 mM hydrogen peroxide (H2O2) and allowed to proceed at 37 °C for
various periods of time. Reactions were terminated at various time
points by addition of 10 nM catalase, and tyrosine
oxidation products were analyzed by HPLC. Similar experiments were
performed in which H2O2 was generated in
situ, using D-glucose and glucose oxidase, instead of
adding reagent H2O2. Hereto, 10-50 milliunits
of glucose oxidase was added to a solution containing 100 µM DL-tyrosine, 1 µM
horseradish peroxidase (HRP), 280 µM
D-glucose in the absence or presence of
NO2 in 50 mM phosphate
buffer (pH 7.4) containing 100 µM DTPA, and reaction
mixtures were incubated at 37 °C. Reactions were terminated by
centrifugation (15,000 rpm) on Microcon-3 concentrators (3,000 molecular weight cutoff) (Amicon Inc., Beverly, MA) to remove proteins.
Tyrosine and its oxidation products in the filtrates were analyzed by
HPLC using a 5-µm Spherisorb ODS-2 RP-18 column and 93% 50 mM potassium phosphate (pH 3.0), 7% methanol as mobile phase at 1 ml/min, and UV detection at 274 nm and fluorescence detection (excitation, 284 nm and emission, 410 nm). Identification and
quantitation of tyrosine oxidation products were performed using
external standards and by spectral matching using photodiode array
detection (34).
Similar experiments were performed using the mammalian peroxidases
myeloperoxidase (MPO) or lactoperoxidase (LPO), in the absence or
presence of various concentrations of chloride (Cl)
and/or thiocyanate (SCN
). In these experiments, HPA (1 mM) was used as a substrate instead of tyrosine, to avoid
interfering reactions of intermediate oxidants with the amino group in
tyrosine. Incubations were terminated by filtration to remove proteins,
and the filtrates were analyzed by HPLC. HPA and its oxidation products
were separated on a 5-µm Spherisorb ODS-2 RP-18 column using 65% 50 mM potassium phosphate (pH 3.0), 35% methanol as mobile
phase at a flow rate of 1 ml/min and analyzed by UV (274 nm) or
fluorescence (excitation, 284 nm and emission, 410 nm) detection.
To study the potential
formation of hydroxylating species during peroxidase-catalyzed
oxidation of NO2, phenylalanine (5 mM in 50 mM phosphate buffer (pH 7.4)) was used
as a trap for aromatic hydroxylation and incubated for 60 min with 10 nM MPO, 700 µM glucose, and 25 milliunits/ml
glucose oxidase (resulting in H2O2 production
at 10 µM/min) in the absence or presence of 1 mM NO2
. After filtration
to remove proteins, the filtrates were analyzed by HPLC for
hydroxylation products, using a Spherisorb ODS-2 RP-18 column, 0.1%
sodium chloride in 1% (v/v) acetonitrile in water adjusted to pH 3.0 with acetic acid as mobile phase at 1 ml/min, and fluorescence
detection (excitation, 275 nm and emission, 305 nm) (35). Using this
procedure, the detection limit for p-, m-, and
o-tyrosine was approximately 20 nM.
Bovine serum albumin (1 mg/ml) in 50 mM sodium phosphate buffer (pH 7.4) was
incubated with 10 nM MPO, 10 nM LPO, or 1 µM HRP in the presence of D-glucose/glucose
oxidase and varying concentrations of
NO2. After incubation, 100-µl
aliquots were mixed with 20 µl of sample loading buffer (20%
glycerol, 10%
-mercaptoethanol, 6% SDS in 125 mM
Tris-HCl (pH 6.8)), heated (5 min, 95 °C), and loaded on 10%
SDS-polyacrylamide gels for electrophoresis. After electrophoresis, proteins were transferred to polyvinylidene difluoride membranes (Sigma) and immunoblotted with a rabbit polyclonal antibody against 3-nitrotyrosine (Upstate Biotechnology, Lake Placid, NY). The antibody was detected using an anti-rabbit secondary antibody conjugated with HRP (Sigma) and stained using
H2O2 and diaminobenzidine (Vector
Laboratories, Burlingame, CA).
5-Thio-2-nitrobenzoic acid
(TNB) was prepared by reduction of 1 mM DTNB in 100 ml of
50 mM sodium phosphate buffer (pH 7.4) with 4 µl of
2-mercaptoethanol (36). MPO or LPO (10 nM) was added to 50 mM sodium phosphate buffer (pH 7.4) containing 40 µM TNB in the absence or presence of various
concentrations of NO2,
Cl
, or SCN
. Reactions were initiated by
addition of 30 µM H2O2 and
allowed to proceed at 20 °C. Oxidation of TNB to DTNB was followed
spectrophotometrically at 412 nm (
412 = 27,200 M
1 cm
1; Ref. 36).
Monochlorodimedon (MCD) is a substrate often
used to study peroxidase-catalyzed chlorination (37). Chlorination of
MCD to dichlorodimedon results in a decrease in absorbance at 290 nm. MCD (40 µM) was dissolved in 50 mM phosphate
buffer containing 150 mM Cl and mixed with
100 µM H2O2 in the absence or
presence of NO2
(2-100
µM). Reactions were initiated by addition of 10 nM MPO, and the decrease in A290 was
followed spectrophotometrically. Reactions were performed at pH ranging
from 6.0 to 7.5, at 20 °C. Control experiments were performed in the
absence of Cl
to study MCD oxidation in the presence of
NO2
alone.
Production of H2O2 by glucose/glucose oxidase was quantitated by oxidation of Fe(II) and formation of a Fe(III)-thiocyanate complex. Aliquots of 800 µl of the reaction mixture were mixed with 100 µl of 10 mg/ml bovine serum albumin, and proteins were precipitated by addition of 100 µl of 100% trichloroacetic acid, after which 800 µl of the supernatants were mixed with 200 µl of Fe(NH4)2(SO4)2 and 100 µl of 2.5 mM KSCN. Formation of the Fe(III)-thiocyanate complex was measured spectrophotometrically at 450 nm within 10 min. Linear standard curves were obtained with 1-50 µM H2O2 solutions treated in a similar manner.
MPO activity was measured spectrophotometrically at 470 nm, using guaiacol oxidation (38). One unit of MPO activity is defined as the amount of enzyme that utilizes 1.0 µmol of H2O2/min in the oxidation of guaiacol at 25 °C and pH 7.0.
NO2 was determined
spectrophotometrically at 543 nm, using Griess reagent (1%
sulfanylamide, 0.1% naphthylethylenediamine, and 2.5%
H3PO4) (28).
In
the presence of H2O2, peroxidases such as
horseradish peroxidase (HRP) are known to catalyze oxidation of the
amino acid tyrosine to form tyrosyl radicals, as indicated by the
production of the dimerization product 3,3-dityrosine (34). In the
presence of NO2
, 3-nitrotyrosine is
formed as an additional product, suggesting that
HRP/H2O2 can oxidize both tyrosine and
NO2
(32). The identity of
3-nitrotyrosine was confirmed by photodiode array detection and
spectral matching compared with authentic 3-nitrotyrosine. Although
dityrosine levels reached a maximum after 15-30 min incubation of 1 mM tyrosine with 1 µM HRP, 1 mM H2O2 and varying concentrations of
NO2
, formation of 3-nitrotyrosine was
found to increase linearly over time during the incubation (Fig.
1A), the rate of tyrosine nitration being
proportional to the concentration of
NO2
(Fig. 1B). No
detectable tyrosine oxidation or nitration was observed in the
absence of HRP or H2O2. Qualitatively
similar results were obtained using D-glucose/glucose
oxidase to continuously generate H2O2 in
situ. Tyrosine (100 µM), incubated with 1 µM HRP, 280 µM D-glucose, and
10 milliunits/ml glucose oxidase (generating H2O2 at 1.0 µM/min), was oxidized
at a rate comparable with the rate of H2O2
production (Fig. 2). Dityrosine accumulated and
eventually it became a substrate for HRP/H2O2
and was oxidized further to trityrosine or other polymeric products
(39, 40). In the presence of NO2
, the
yield of dityrosine was decreased, and 3-nitrotyrosine was found to
accumulate during the incubation (Fig. 2), suggesting that
NO2
competes with tyrosine for
oxidation by HRP/H2O2.
The mammalian peroxidases myeloperoxidase (MPO) and lactoperoxidase
(LPO) are also able to oxidize tyrosine in the presence of
H2O2, to form dityrosine, and are reported to
do so more efficiently than HRP (39). In the presence of
NO2, these peroxidases also generate
3-nitrotyrosine as an additional product. Using 1 mM
tyrosine, 100 µM NO2
,
and 4.0 µM/min H2O2, the yield of
both oxidation products increased with the concentration of MPO or LPO
and was maximal at enzyme concentrations between 10 and 20 nM. At higher enzyme concentrations the yield of dityrosine
did not increase further, and production of 3-nitrotyrosine was found
to decrease (not shown). For this reason, further studies were
performed using 10 nM of MPO or LPO. Both tyrosine
oxidation products accumulated rapidly during the first 30 min, and the
rate of product formation declined dramatically thereafter, possibly
because of enzyme inactivation or due to depletion of dissolved
molecular oxygen necessary for H2O2 generation. Hence, in further experiments using glucose/glucose oxidase, reaction mixtures were incubated for 30 min. As demonstrated in Table
I, the extent of tyrosine nitration by MPO or LPO was
dependent on the initial NO2
concentration, although the production of dityrosine was not dramatically affected. The yield of both products was markedly lower
when LPO was used instead of MPO, consistent with the notion that MPO
more efficiently catalyzes oxidation of tyrosine (39) and perhaps
NO2
. However, the results indicate
that both peroxidases are capable of oxidizing
NO2
in the presence of
H2O2 to form a reactive intermediate that is
capable of nitrating tyrosine.
|
In the presence of NO2,
MPO/H2O2 was also found capable of nitrating
tyrosine residues in proteins. As shown in Fig. 3,
reaction of 10 nM MPO with 1 mg/ml albumin in the presence
of 0.5 or 1.0 mM NO2
and
10 µM/min H2O2 resulted in
nitration of tyrosine residues, as detected by SDS-polyacrylamide gel
electrophoresis and Western blotting using a rabbit polyclonal antibody
against 3-nitrotyrosine. Protein nitration was also observed when LPO
or HRP were used instead of MPO (not shown).
Possible Oxidation Mechanisms
Peroxidases commonly oxidize
aromatic substrates by a one-electron oxidation mechanism, although
oxidation of halides by mammalian peroxidases has been generally
thought to occur via two-electron oxidation (39-42).
Peroxidase-catalyzed oxidation of NO2
therefore results in formation of either
NO2· or an
NO2+-like intermediate. Oxidation of
NO2
by
MPO/H2O2 did not result in aromatic
hydroxylation of the amino acid phenylalanine, indicating that a
hydroxylating species such as ONOOH (43) is not formed during
MPO-catalyzed NO2
oxidation. Addition
of 1 mM NO2+ (as the nitryl
salt NO2BF4) to a solution of 1 mM
tyrosine at pH 7.4 resulted in formation of 2 ± 1 µM 3-nitrotyrosine (mean ± S.D.; n = 3), which is markedly less than the extent of nitration by
MPO/H2O2 in the presence of 1 mM
NO2
(Table I). Although
NO2+ is capable of nitrating aromatic
rings by electrophilic aromatic substitution, up to 2 mM
NO2BF4 did not induce detectable nitration of
the aromatic substrate 4-methoxybenzoic acid (a substrate incapable of
forming phenolic radicals) in aqueous solution, possibly because of
rapid hydrolysis of NO2+. Hence, it is
unlikely that phenolic nitration via peroxidase-catalyzed oxidation of
NO2
is due to formation of
NO2+.
The presumed major physiological
substrate for MPO is chloride (Cl), although other anions
such as bromide (Br
) or thiocyanate (SCN
)
are sufficiently abundant in biological fluids to act as alternative physiological substrates (44, 45). Similarly, SCN
and
Br
have been proposed as a physiological substrates for
eosinophil peroxidase (EPO) or LPO (46-48). We therefore investigated
whether MPO or LPO could catalyze oxidization of
NO2
in the presence of either
Cl
or SCN
, using TNB as an oxidizable
substrate. In the absence of other substrates,
MPO/H2O2 slowly oxidizes TNB to form DTNB. The
rate of TNB oxidation was, however, enhanced dramatically when either Cl
, NO2
, or
SCN
was also present in the reaction mixture, suggesting
that these anions act as substrates for
MPO/H2O2 to form diffusible oxidation products
which in turn oxidize TNB. Rates of TNB oxidation by MPO/H2O2 in the presence of these anionic
substrates are shown in Fig. 4 and demonstrate that
SCN
is used more efficiently as a substrate than
NO2
, which is preferred as a substrate
over Cl
for MPO. Similarly,
NO2
is a substrate for LPO, but
LPO/H2O2 preferentially utilizes SCN
in oxidation of TNB (not shown). Only minimal
oxidation of TNB was observed by LPO/H2O2 in
the presence of 100 mM Cl
. As LPO is unable
to catalyze Cl
oxidation (see below), this is most likely
due to contaminants, such as Br
.
The rate of TNB oxidation by MPO/H2O2 and 100 mM Cl was found to be markedly inhibited in
the presence of NO2
. High
concentrations of NO2
were found to
inhibit TNB oxidation by
MPO/H2O2/Cl
to rates more similar
to those observed in the presence of
NO2
alone (Fig.
5A), and dramatic (>75%) inhibition of
Cl
-dependent TNB oxidation was observed in
the presence of NO2
at concentrations
as low as 10 µM. Nearly identical results were obtained
when 100 µM SCN
was used instead of 100 mM Cl
(not shown). As 10 µM
NO2
is unlikely to inhibit oxidation
of 40 µM TNB by the intermediately formed oxidants (HOCl
or HOSCN) to such an extent, the results suggest that
NO2
also competitively inhibits
Cl
or SCN
oxidation by
MPO/H2O2. In contrast to the results with MPO,
SCN
-catalyzed TNB oxidation by
LPO/H2O2 was found to be enhanced by 10 µM NO2
(Fig.
5B), suggesting that NO2
also can act as a substrate for LPO in the presence of physiological levels of SCN
.
Effect of NO2
Addition of MPO to a solution containing 40 µM
MCD, 100 µM H2O2, and 150 mM Cl was found to result in chlorination of
MCD, as demonstrated by a decrease in A290 (37,
49). The rate of MPO-catalyzed MCD chlorination increased with
decreasing pH (between pH 6.0 and 7.5), consistent with earlier reports
(50). Reaction of MPO with excess H2O2 results
in rapid formation of MPO compound I, which is then spontaneously
reduced to compound II (50, 51). As MPO compound II is unable to
oxidize Cl
(49), accumulation of compound II will result
in a decreased rate of MCD chlorination, and this was indeed observed
(Fig. 6). Addition of 2-10 µM
NO2
to the reaction mixture did not
significantly affect the initial rate of MCD oxidation but diminished
the decrease in chlorination rate over time, and at
NO2
concentrations of 5 µM or higher, the rate of MCD chlorination was constant
over the course of the experiment (Fig. 6). The rate of MCD
chlorination was not increased much further in the presence of higher
NO2
concentrations (Fig. 6), possibly
because high NO2
concentrations may
scavenge the chlorinating species, thereby partly inhibiting MCD
chlorination. The MCD concentration was only minimally affected by
MPO/H2O2 and 100 µM
NO2
in the absence of Cl
(minimal decrease in A290; Fig. 6), indicating
that oxidation products of NO2
do not
importantly contribute to the observed increased MCD
oxidation/chlorination in the presence of Cl
.
Peroxidase-catalyzed Phenolic Nitration in the Presence of Cl
Modification of phenolic
substrates by MPO-catalyzed oxidation of
NO2 was also studied in the presence
of alternative peroxidase substrates. MPO-catalyzed oxidation of
Cl
is known to induce chlorination of aromatic substrates
(37, 52, 53), which was confirmed in the present study using HPA. Incubation of 1 mM HPA with
MPO/H2O2 and Cl
resulted in
formation of 3-chloro-HPA, the extent of HPA chlorination being
proportional to the concentration of Cl
. In the absence
of Cl
, MPO/H2O2 catalyzes
oxidation of HPA to 3,3
-diHPA (the dimerization product of HPA), and
formation of 3,3
-diHPA was partly inhibited by Cl
(the
yield of 3,3
-diHPA was decreased up to 30% in the presence of 50 mM Cl
, but to a lesser extent in the presence
of higher Cl
concentrations). Thus, high concentrations
of Cl
appear to outcompete HPA for oxidation by
MPO/H2O2, but Cl
oxidation may
also contribute to HPA oxidation and dimerization.
MPO-catalyzed HPA chlorination in the presence of physiological levels
of Cl (150 mM) was found to be enhanced by
low concentrations of NO2
. As shown in
Fig. 7, HPA chlorination was increased most dramatically in the presence of 2-10 µM
NO2
. Increasing
NO2
concentrations did not
dramatically enhance further HPA chlorination and, in fact, HPA
chlorination was partly inhibited in the presence of concentrations of
NO2
above 200 µM (not
shown). Addition of 2-10 µM
NO2
also enhanced
MPO/H2O2/Cl
-induced formation of
3,3
-diHPA, and 3-NO2-HPA was formed as an additional
product (Fig. 7), and the extent of HPA nitration and dimerization
continued to increase in the presence of increasing concentrations of
NO2
. These observations are consistent
with our results with MCD and suggest that
NO2
increases MPO-catalyzed
Cl
oxidation by reducing MPO compound II to ferric MPO.
At high concentrations, NO2
may also
partly scavenge the intermediate chlorinating species (HOCl or
Cl2; Ref. 54), thereby inhibiting aromatic chlorination by
MPO/H2O2/Cl
.
Nitration of HPA by
MPO/H2O2/NO2
could be partially inhibited by a large excess of Cl
.
When the inhibition of HPA nitration was plotted against the negative
logarithm of the concentration of Cl
, a sigmoidal curve
was obtained (Fig. 8), suggesting that Cl
competitively inhibits NO2
oxidation
by MPO/H2O2, and the IC50 for
Cl
increased when higher concentrations of
NO2
were used. HPA nitration was
inhibited maximally by 30-45% (Fig. 8), and nitration of HPA was
still observed even when Cl
was present in
1,000-10,000-fold excess over
NO2
.
Extracellular fluids also contain significant levels of
SCN (20-120 µM in plasma but much higher
in secretions such as milk or saliva, for example), which has been
proposed as an alternative substrate for peroxidases. Both nitration
and chlorination of HPA, observed after incubation of HPA with
MPO/H2O2 in the presence of 150 mM
Cl
and 100 µM
NO2
, were found to be inhibited by
SCN
(Fig. 9). However, chlorination of HPA
was inhibited with an IC50
5 µM, and no
chlorination of HPA could be detected in the presence of >25
µM SCN
(<0.2 µM 3-Cl-HPA).
On the other hand, nitration of HPA was inhibited much less efficiently
by SCN
(IC50
60 µM), and
formation of 3-NO2-HPA was still detectable in the presence
of 500 µM SCN
(0.05 µM
3-NO2-HPA after 60 min incubation), indicating that NO2
can act as a substrate for
MPO/H2O2 even in the presence of physiological levels of Cl
and SCN
. Nitration of HPA by
LPO/H2O2/NO2
was not affected by 150 mM Cl
, consistent
with the notion that Cl
is not a substrate for LPO.
Moreover, chlorination of HPA was not observed by
LPO/H2O2/Cl
in the presence or
absence of NO2
. HPA nitration by
LPO/H2O2 and 100 µM
NO2
was inhibited by SCN
(IC50
10 µM), and no HPA nitration was
detected in the presence of >100 µM
SCN
.
The results presented herein demonstrate that the peroxidases HRP,
MPO, and LPO can oxidize NO2 in the
presence of H2O2 or
H2O2-generating systems to form reactive nitrogen intermediate(s) capable of nitrating phenolic compounds such
as tyrosine. At acidic pH, NO2
is
partly protonated and reacts with H2O2 directly
to form peroxynitrous acid (ONOOH), which causes aromatic nitration
(34). However, this did not occur to an appreciable extent at pH 7.4 under our reaction conditions. In addition to being oxidized to more
reactive nitrogen species, NO2
was
also found to catalyze MPO-mediated oxidation of TNB or chlorination of
MCD or HPA. These results confirm and extend a recent study by Shibata
et al. (32), who demonstrated that HRP and
H2O2 oxidize NO2
to form an intermediate that is
capable of causing chlorophyll degradation and tyrosine nitration. They
postulated that HRP oxidizes NO2
to
NO2· in the presence of
H2O2 according to the following reaction scheme (Reactions 1-3):
![]() |
![]() |
![]() |
![]() |
![]() |
![]() |
![]() |
![]() |
Taken together, our results suggest that in the presence of
H2O2, heme peroxidases can catalyze oxidation
of NO2 to predominantly
NO2·, and this could contribute to
aromatic nitration in vivo at sites where
NO2
levels are sufficiently high.
Although tyrosine nitration by NO2· is
rather inefficient because two NO2·
molecules are needed for nitration of one tyrosine residue, and intermediately formed tyrosyl radicals undergo rapid reactions including dimerization, tyrosyl radicals in proteins are often more
long-lived than free tyrosyl radicals, rendering tyrosyl radicals in
proteins more susceptible targets for nitration by NO2·, via rapid radical-radical
reaction (k = 3 × 109
M
1 s
1; Ref. 59). Moreover,
tyrosyl radicals have been detected in various enzymes such as
ribonucleotide reductase and prostaglandin H synthase (60), and
formation of NO2· could result in
nitration of these tyrosine residues and potentially affect enzyme
activity.
Although mammalian peroxidases such as MPO and LPO
can catalyze oxidation of NO2,
biological fluids contain alternative substrates that might be
preferentially oxidized. For instance, intracellular and extracellular fluids contain 100-150 mM Cl
, which is
commonly thought to be the physiological substrate for MPO (48).
Eosinophil peroxidase (EPO) can also oxidize Cl
, but
Br
and SCN
(plasma levels are 20-100 and
20-120 µM, respectively; Ref. 45) have been suggested as
more important physiological substrates for EPO, because these anions
are oxidized more easily. Similarly, SCN
has been
proposed as the physiological substrate for LPO in saliva or milk,
whereas SCN
levels are as high as 5 mM (47).
Reported oxidation potentials of SCN
, Br
,
Cl
and NO2
are
summarized below in Reactions 6-9 (61, 62):
![]() |
![]() |
![]() |
![]() |
![]() |
![]() |
Our studies with TNB have indicated that
NO2 acts as a competing substrate for
MPO/H2O2 in the presence of Cl
or
SCN
, as low concentrations of
NO2
dramatically inhibited TNB
oxidation by MPO/H2O2/Cl
or
MPO/H2O2/SCN
. However,
NO2
could also scavenge the oxidation
products of Cl
or SCN
, thereby inhibiting
TNB oxidation. In contrast, TNB oxidation by
LPO/H2O2/SCN
was enhanced by
NO2
, indicating that
NO2
can act as a substrate for
LPO/H2O2, even in the presence of excess
SCN
. The opposing results obtained with both peroxidases
may be due to different enzyme specificity or differences in oxidation
mechanisms. The notion that NO2
can
act as a substrate for MPO/H2O2 in the presence
of physiological levels of Cl
is further supported by our
studies with MCD chlorination or HPA oxidation, which demonstrated that
low concentrations of NO2
markedly
enhanced MPO-catalyzed chlorination of these substrates. Reducing
agents such as ascorbate, urate, or O
2 have been reported to
enhance the chlorinating activity of
MPO/H2O2/Cl
by reducing MPO
compound II to the native ferric enzyme (50, 64, 65). As MPO compound
II is unable to oxidize Cl
(49, 65), accumulation of MPO
compound II would reduce the rate of aromatic chlorination, which can
be prevented in the presence of reducing agents. We propose that
NO2
may act similarly as a reducing
substrate for MPO compound II, thereby recycling it to ferric MPO,
which would again participate in oxidation of Cl
. In this
process, NO2
is oxidized by a
one-electron mechanism to NO2·,
causing enhanced HPA dimerization and nitration by
MPO/H2O2/Cl
in the presence of
NO2
. In a recent study with
water-soluble metalloporphyrins, oxidation of a Fe(III) porphyrin with
m-chloroperoxybenzoate was found to produce the O=Fe(IV)
species (equivalent to compound II), which could rapidly be reduced
back to the Fe(III) porphyrin by addition of equivalent amounts of
NO2
(66), which is consistent with the
proposed reduction of MPO compound II by
NO2
. The observed partial inhibition
of
MPO/H2O2/NO2
-induced
HPA nitration by Cl
is also consistent with this
mechanism. Although Cl
is able to compete with
NO2
for oxidation by MPO compound I,
Cl
cannot reduce MPO compound II (65) and hence does not
completely inhibit
MPO/H2O2/NO2
-mediated
HPA nitration. Fig. 10 schematically summarizes the
proposed mechanisms by which MPO can catalyze oxidation of
NO2
.
In addition to catalyzing MPO-mediated chlorination by reduction of MPO
compound II, NO2 may also react with
the initially formed chlorinating species (HOCl or Cl2;
Ref. 54), to form a reactive intermediate similar to nitryl chloride
(NO2Cl), which is capable of inducing aromatic chlorination
and nitration (29). Hence, NO2Cl may contribute to the
observed MCD chlorination or HPA chlorination and nitration by
MPO/H2O2/Cl
in the presence of
NO2
. However, this mechanism is
unlikely to fully account for the observed increases in aromatic
chlorination and nitration because (i) low concentrations of
NO2
(2-10 µM) are
unlikely to efficiently compete with the much higher concentrations of
MCD or HPA for reaction with HOCl or Cl2, and (ii) the
produced intermediates (including NO2Cl) are unstable and
rapidly hydrolyze to NO2
and
Cl
. In fact, studies with reagent HOCl have indicated
that chlorination of HPA by HOCl was decreased in the presence of
NO2
, with a concomitant increase in
HPA nitration (29), which most likely explains why increasing
concentrations of NO2
failed to
dramatically further enhance aromatic chlorination by
MPO/H2O2/Cl
(Figs. 6 and 7).
Both nitration and chlorination of HPA by
MPO/H2O2 in the presence of Cl
and NO2
were inhibited by
SCN
, an alternative physiological substrate for mammalian
peroxidases. However, the fact that nitration of HPA was still
detectable in the presence of physiological SCN
levels,
whereas HPA chlorination was completely inhibited, further suggests
that nitration is due to direct oxidation of
NO2
by
MPO/H2O2, rather than via oxidation by
chlorinating intermediates such as HOCl or Cl2. Similarly,
LPO/H2O2/NO2
also caused detectable HPA nitration in the presence of
SCN
at concentrations similar to that of
NO2
. SCN
may
competitively inhibit oxidation of Cl
and
NO2
by these enzymes but could also
act by scavenging reactive intermediates formed by oxidation of
Cl
or NO2
.
One important
implication of the results presented herein is that oxidation of
NO2 by mammalian peroxidases in the
presence of H2O2 may represent an alternative
or additional mechanism of aromatic nitration in vivo. We
have recently demonstrated that oxidation of
NO2
by HOCl also results in formation
of reactive intermediates capable of inducing phenolic nitration (29),
and both mechanisms of NO2
oxidation
may contribute to tyrosine nitration observed in tissues undergoing
inflammatory responses and where high levels of MPO are present.
Interestingly, 3-nitrotyrosine and large amounts of active MPO have
both been detected in atherosclerotic lesions (67-69). Moreover, high
levels of MPO are also present in the rheumatoid joint of patients with
rheumatoid arthritis, and synovial fluids of rheumatoid arthritis
patients are also reported to contain increased levels of
NO2
and 3-nitrotyrosine (18, 70).
Furthermore, tyrosine nitration has also been detected in lung sections
of patients with acute pulmonary inflammation (71, 72), a condition
characterized by infiltration of neutrophils as well as increased
production of NO· and NO2
(e.g. Refs. 20, 21). Detection of 3-nitrotyrosine in
vivo, often regarded specific evidence for formation of
ONOO
, may therefore also be due to formation of reactive
nitrogen species by oxidation of NO2
by MPO or by HOCl. Hence, 3-nitrotyrosine is not a unique biomarker of
ONOO
production but merely indicates formation of
reactive nitrogen species derived from NO·, and formation of
3-nitrotyrosine in vivo may often largely depend on the
presence of MPO. Interestingly, it was recently reported that
NO· synthase is induced in cytokine-stimulated human neutrophils and found to be co-localized with MPO in primary granules (73). Induction of NO· synthase and activation of MPO are both likely
to be involved in the observed tyrosine nitration around ingested
bacteria (73).
We have performed preliminary studies with isolated human
polymorphonuclear neutrophils (PMN), which demonstrated that
stimulation of 2 × 106/ml PMN in PBS (150 mM Cl) with 100 ng/ml phorbol myristate
acetate in the presence of 10-100 µM
NO2
and 1 mM HPA results
in HPA nitration. Furthermore, HPA chlorination was enhanced in the
presence of NO2
, similar to our
studies with purified MPO. Stimulation of PMN with phorbol myristate
acetate results in complete degranulation and secretion of MPO into the
incubation medium. After 60-min incubations, the medium MPO activity
was 14 ± 5 milliunits/ml with 2 × 106/ml
resting PMNs and 175 ± 56 milliunits/ml (mean ± S.D.;
n = 4) after stimulation of PMN with phorbol myristate
acetate. The amount of MPO released from these PMNs was similar to that
used in the studies with purified MPO; 10 nM MPO
corresponds to an enzyme activity of 150 milliunits/ml. The same
number of activated PMN produce O
2 at a rate of about
5 µM/min (74), which would yield 2.5 µM
H2O2/min upon dismutation, similar to the rate
of H2O2 production used in our studies with
purified MPO and glucose/glucose oxidase. Therefore, the
NO2
oxidation mechanisms described in
this study are likely to be relevant during conditions of acute
inflammation, where PMNs are recruited and activated.
Peroxidase-catalyzed oxidation of NO2
or oxidation of NO2
by oxidants such
as HOCl may result in underestimation of NO· production under
inflammatory conditions, when NO2
is
measured to quantitate NO· production. These oxidation
mechanisms may significantly contribute to
NO2
oxidation under inflammatory
conditions when MPO and/or EPO are secreted. Interestingly, it has
recently been demonstrated that the
H2O2-scavenging activity of respiratory tract
mucus is predominantly due to the presence of a peroxidase similar to
LPO (75), which in the presence of sufficient concentrations of
NO2
could lead to production of
reactive nitrogen intermediates such as
NO2· in the respiratory tract.
Similarly, in situations when SCN
concentrations are
sufficiently low, NO2
could be an
important substrate for peroxidases in the oral cavity, especially when
salivary concentrations of NO2
are
elevated after ingestion of NO3
.
As NO2· is a powerfully oxidizing
species (27), peroxidase-catalyzed NO2
oxidation could induce biomolecular modifications in host tissues and
contribute to cell or tissue injury as a result of increased NO·
production. Moreover, NO2
could also
contribute to tissue injury by catalyzing MPO-mediated oxidation and
chlorination reactions. Alternatively,
NO2
has also been demonstrated to
increase the bactericidal activity of MPO (28), and
peroxidase-catalyzed NO2
oxidation
could therefore also represent an additional host defense mechanism.
In conclusion, we have demonstrated that
NO2 is a potential physiological
substrate for heme peroxidases such as MPO and LPO, even in the
presence of physiological concentrations of the alternative substrates
Cl
or SCN
, and is able to catalyze
peroxidase-mediated oxidation and chlorination of biological targets.
Moreover, peroxidase-catalyzed oxidation of
NO2
results in formation of
NO2· or a related species, which can
contribute to tyrosine nitration and could be involved in cell and
tissue injury during situations of increased NO· production.
This potential contributing role of
NO2
to NO· biochemistry has
been relatively overlooked, and further studies are needed to evaluate
its physiological and/or pathological importance.