1 Departments of Anesthesiology, 2 Physiology and Biophysics, and 3 Pediatrics, University of Alabama at Birmingham, Birmingham, Alabama 35233
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ABSTRACT |
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We examined the effect of peroxynitrite
(ONOO) on the cloned rat
epithelial Na+ channel
(
-rENaC) expressed in Xenopus
oocytes. 3-Morpholinosydnonimine (SIN-1) was used to concurrently
generate nitric oxide (· NO) and superoxide
(O
2 ·), which react to
form ONOO
, a species known
to promote protein nitration and oxidation. Under control conditions,
oocytes displayed an amiloride-sensitive whole cell conductance of 7.4 ± 2.8 (SE) µS. When incubated at 18°C with SIN-1 (1 mM) for 2 h (final ONOO
concentration = 10 µM), the amiloride-sensitive conductance was reduced to
0.8 ± 0.5 µS. To evaluate whether the observed inhibition was due to ONOO
, as opposed
to · NO, we also exposed oocytes to SIN-1 in the presence of
urate (500 µM), a scavenger of
ONOO
and superoxide
dismutase, which scavenges
O
2 ·, converting SIN-1
from an ONOO
to an
· NO donor. Under these conditions, conductance values remained at control levels following SIN-1 treatment.
Tetranitromethane, an agent that oxidizes sulfhydryl groups at pH
6, also inhibited the amiloride-sensitive conductance. These data
suggest that oxidation of critical sulfhydryl groups within rENaC by
ONOO
directly inhibits
channel activity.
nitric oxide; reactive species; sodium conductance; tetranitromethane; 3-morpholinosydnonimine; oxidation; nitration
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INTRODUCTION |
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AMILORIDE-SENSITIVE SODIUM CHANNELS have been identified in the apical membranes of a variety of epithelial tissues and are in an important class of proteins that regulate Na+ transport and fluid homeostasis. These channels are classified into various categories based on their biophysical properties and the extent of their inhibition by amiloride and its structural analogs. One group of these channels [referred to as type Na(5)] have a low single-channel conductance (4-5 pS), high selectivity for Na+ to K+ (PNa/PK > 10), and long mean open and closed times (0.5-5 s) (32). Additionally, these channels have a high affinity for amiloride (Ki < 0.5 µM at high external Na+ concentrations) and display the following structure/inhibition pattern relationship: phenamil, benzamil > amiloride >>> N-ethyl-N-isopropyl amiloride (35). This group includes the so-called classic Na+ channels present in the apical membranes of epithelia with high transepithelial electrical resistance, such as frog skin, toad urinary bladder, mammalian colon, bovine renal papilla, and amphibian A6 cells (32, 35).
A cDNA encoding an amiloride-sensitive
Na+ channel, known as -rENaC
(for the
-subunit of the rat epithelial
Na+ channel), was cloned from the
colon of salt-deprived rats using functional RNA expression (8, 10).
When expressed in Xenopus oocytes,
-rENaC displayed the characteristics of Na(5) channels. Subsequently, Canessa et al. (10) identified and cloned two additional
subunits of this channel, named
-rENaC and
-rENaC. Coexpression
of all three subunits in oocytes generated 100-fold higher
amiloride-sensitive currents than
-rENaC alone (10).
Because of their location, epithelial tissues are often exposed to
reactive oxygen and nitrogen species generated by a variety of
intracellular and extracellular sources. In addition, most mammalian
cells have the capacity to produce nitric oxide (· NO) from
the oxidative deamination of
L-arginine by either the
Ca2+-sensitive (type I or III) or
the Ca2+-insensitive (type II)
forms of · NO synthase (14). · NO is an important signal transduction molecule with diverse physiological functions, including vasoregulation, neurotransmission, and immune host
defense (29). For example, · NO may modulate the function of
various proteins by bonding to transition metal centers. In the case of
guanylate cyclase, perhaps the best understood example, this leads to
synthesis of cGMP. However, the biological effects of · NO
depend on its concentration, the biochemical composition of target
molecules, and the presence of other free radicals. Under some
circumstances, · NO is known to produce significant tissue
injury. This is thought to be due, at least in part, to the
rate-limited reaction of · NO with superoxide
(O2 ·) to form the
highly reactive product peroxynitrite
(ONOO
) (4). This species
nitrates phenolics and oxidizes methionine and cysteine residues by
one- or two-electron transfer. Both processes have been shown to result
in altered protein function (15, 28, 30, 38).
There is thus considerable interest in identifying the contribution of
· NO and its reactive intermediates to the initiation and
propagation of injury to epithelial cells, including the effects on ion
transport. Our previous results indicate that bolus addition of
ONOO decreases
amiloride-sensitive
22Na+
uptake across membrane vesicles of colonic epithelial cells and freshly
isolated alveolar type II cells (3, 21). However, the mechanisms by
which reactive nitrogen intermediates alter Na+ transport have not been
elucidated.
Herein we isolated oocytes from Xenopus
laevis, injected them with equivalent amounts (8.3 ng)
of -,
-, and
-rENaC cRNA, and recorded whole cell currents
36-48 h later, in the presence and absence of amiloride. To
quantify the effects of · NO and its by-products on the
amiloride-sensitive currents, an index of
Na+ channel activity, we incubated
oocytes with ONOO
and
· NO donors. In addition, these effects were compared with measurements obtained after exposing oocytes to tetranitromethane (TNM), which, like ONOO
,
acts as a nitrogen dioxide
(· NO2) donor and
oxidizes sulfhydryl groups and, depending on the pH value, may also
nitrate phenolics (36, 37). Our results indicate that
ONOO
, but not
· NO, decreased the amiloride-sensitive
Na+ current in oocytes expressing
-rENaC. Moreover,
ONOO
most likely does this
through oxidation of critical amino acid residues in the rENaC protein.
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MATERIALS AND METHODS |
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RNA synthesis. The pSport plasmid
(GIBCO BRL, Gaithersburg, MD), which contained either -,
-, or
-rENaC (a generous gift from Dr. B. Rossier, University of Lausanne,
Switzerland), was linearized by overnight incubation with
Not I (Promega, Madison, WI). Sense
RNA was in vitro transcribed from purified plasmid DNA using T7
polymerase according to the manufacturer's instructions (Ambion,
Austin, TX). The integrity of the cRNA was verified by denaturing gel
electrophoresis through 1% agarose-formaldehyde gels.
Oocyte expression. Female X. laevis toads (Xenopus Express, Beverly Hills, FL) were maintained in dechlorinated tap water at 18°C and fed beef liver twice weekly. A partial ovarectomy was performed under anesthesia (immersion in ice-cold solution of 0.1% ethyl-m-aminobenzoate methane sulfonate; Sigma, St. Louis, MO) through a small (~5 mm) ventral abdominal incision. The ovarian tissue was then placed in half-strength Lebowitz-15 (1/2L15; GIBCO BRL) medium buffered with 15 mM HEPES, pH 7.6. Oocytes at maturation stages V and VI were manually defolliculated and incubated overnight in fresh oocyte culture media [1/2L15 medium supplemented with penicillin (100 U/ml) and streptomycin (100 µg/ml) at 18°C]. A Nanoject (Drummond, Broomall, PA) microinjector was used to inject 25 ng of total cRNA (8.3 ng of each rENaC subunit) in a 50-nl volume into each oocyte. Control oocytes were injected with 50 nl of KCl (100 mM). Injected oocytes were incubated in oocyte culture media at 18°C until use.
Electrophysiological measurements.
Membrane currents were evaluated 36-48 h postinjection using a
two-electrode voltage clamp. Oocytes were bathed in modified ND96
solution at 18°C. This solution contained (in mM) 96 NaCl, 2 KCl, 1 MgCl2, 0.2 CaCl2, and 15 HEPES at pH 7.6. Cells were impaled with two 3 M KCl-filled electrodes having
resistances of 0.5-1.5 M. The electrodes were connected to a
Geneclamp 500 (Axon Instruments, Foster City, CA) current-voltage (I-V) clamp amplifier via Ag-AgCl
pellet electrodes and referenced to an Ag-AgCl pellet connected to the
bath via a 3 M KCl-agar bridge. After impalement, membrane potentials
were allowed to stabilize before current measurements (5-10 min).
The voltage clamp was controlled by a Macintosh IIci computer running
Axodata (Axon Instruments) acquisition software.
Oocytes were clamped at a holding potential of 0 mV. Currents were
recorded every 20 s by stepping from the holding potential to
100 mV for 400 ms, back to the holding potential for 50 ms, then
to +100 mV for 400 ms. I-V
relationships were determined before drug administration and at timed
intervals thereafter by stepping from the holding potential to
100 mV through +60 mV in 20-mV increments for 500 ms. Current
values sampled over the last 50 ms of each voltage step were averaged
and used to construct I-V curves.
Whole cell conductance values were calculated as the slope of the
I-V curve over the range from
100 to
60 mV. Amiloride-sensitive I-V relationships were calculated by
subtracting the I-V relationship after
amiloride application (10 µM) from that obtained before.
Generation of ONOO and
· NO.
Stock solutions of 100 mM 3-morpholinosydnonimine (SIN-1; Calbiochem,
La Jolla, CA) in 10 mM phosphate buffer (pH 5.5), 10 mM urate (Sigma,
St. Louis, MO) in 1/2L15 media, 78 mg/ml (5,200 U/mg) human
Cu/Zn superoxide dismutase (SOD; provided to us by Dr. J. Beckman,
Dept. of Anesthesiology, University of Alabama at Birmingham), 8.4 M
TNM (Aldrich, Milwakee, WI) in ethanol, and 100 mM PAPA-NONOate (Cayman
Chemical, Ann Arbor, MI) in 10 mM degassed phosphate buffer (pH 8.5)
were stored at
20°C. Working solutions were prepared from
stock solutions at the time of each experiment.
Quantitation of ONOO production.
ONOO
generation by SIN-1 (1 mM) was quantified by
measuring the rate of rhodamine formation from the oxidation of
dihydrorhodamine 123 (DHR) as previously described (16). All
measurements were conducted in 1/2L15 media at 25 or 18°C.
Rates were also measured at 18°C in the presence of SOD and urate.
Samples were taken at either 10- or 15-min (SOD and urate) intervals.
Rhodamine formation was monitored in a spectrophotometer at 500 nm for
2 min (
500nm = 78,000 M
1 · cm
1)
in cuvettes containing DHR (50 µM). Because
ONOO
oxidizes DHR with an
efficiency of 45%, the rate of
ONOO
formation was
calculated by dividing the rate of rhodamine formation by 0.45.
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RESULTS |
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SIN-1 forms ONOO in the oocyte
recording solution.
The time course of ONOO
formation resulting from the addition of SIN-1 (1 mM) to the
1/2L15 media (pH 7.6) is shown in Fig. 1. At 25°C,
ONOO
levels rose gradually
to >30 µM by 2 h. In contrast, only ~10 µM
ONOO
was
produced over the same time course at 18°C, the temperature at
which oocytes were exposed to SIN-1 and at which electrophysiological recordings were performed.
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The inhibitory effect of SIN-1 is mediated by
ONOO.
When SIN-1 was added to 1/2L15 culture media in the presence of
urate (500 µM) or SOD (3,000 U/ml), there was a >90% drop in detectable DHR oxidation, indicating a significant decrease in ONOO
production (Fig. 1).
As shown in Table 1, similar
values for the amiloride-sensitive conductance of
-rENaC-expressing oocytes were seen in controls (7.4 ± 2.8 µS; n = 6), those treated with SIN-1
in the presence of urate (7.7 ± 2.2 µS;
n = 7), and those treated with SIN-1
in the presence of SOD (6.0 ± 1.6 µS;
n = 7). However, SIN-1 alone
profoundly inhibited the amiloride-sensitive conductance (0.8 ± 0.5 µS; n = 7). We further examined the
effect of · NO on the
-rENaC-expressing oocytes by
treating them with 400 µM PAPA-NONOate, which generates >3 µM
· NO (18). However, even at this supraphysiological
concentration, · NO failed to decrease either the total or
the amiloride-sensitive currents (data not shown). These data indicate
that the inhibitory effect of SIN-1 on the amiloride-sensitive
conductance was due to production of
ONOO
and not · NO
or hydrogen peroxide.
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The inhibitory effects of SIN-1 are not mediated through a
Ca2+-dependent
pathway.
Oocytes expressing -rENaC and pretreated with BAPTA-AM (50 µM) for 3 h followed by exposure to SIN-1 (1 mM) and BAPTA-AM for 2 h
had amiloride-sensitive conductance values that were not different from
those of oocytes exposed to SIN-1 alone [17.7 ± 6.4 µS
(n = 4) vs. 21.4 ± 4.1 µS
(n = 6)]. In this set of
experiments, these values were ~55% lower than the
amiloride-sensitive conductance of control oocytes expressing rENaC
(47.4 ± 3.0 µS; n = 3;
P < 0.05). These data suggest that
SIN-1 did not inhibit the amiloride-sensitive conductance through a
Ca2+-dependent pathway. This is
further supported by the fact that SIN-1 did not activate the
endogenous Ca2+-activated
Cl
currents present in
KCl-injected oocytes (Fig. 4).
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DISCUSSION |
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In agreement with other studies (1, 9, 25), we show that coinjection of
the -,
,- and
-rENaC subunits in
Xenopus oocytes resulted in the
expression of a constitutively active amiloride-sensitive conductance.
We further show that incubation of
-rENaC-expressing oocytes
with SIN-1 for
2 h resulted in a marked decrease in this
amiloride-sensitive conductance. Importantly, the
I-V relationships of rENaC-expressing
oocytes treated with amiloride or exposed to SIN-1 were not different
from those of KCl-injected oocytes. The changes in
ERev, as well as
the decrease in the whole cell conductance, support the conclusion that
SIN-1 inhibited rENaC activity, rather than the amiloride sensitivity of the channel expressed in oocytes. This is an important distinction in light of recent work by Ismailov et al. (25), who demonstrated that
deletion of amino acid residues 278-283 in the extracellular domain of the
-rENaC subunit reduced the amiloride sensitivity of
the channel by two orders of magnitude without affecting the conductance of the channel. SIN-1-induced modifications of amino acid
residues in this region of the protein could have affected amiloride
binding and therefore the amiloride sensitivity. However, the whole
cell conductance would be unchanged, and this was not the case in our
study.
The decomposition of SIN-1 at physiological pH leads to the
simultaneous generation of
O2 · and · NO,
which react at diffusion-limited rates to form the highly reactive
species ONOO
(20, 22). The
following observations implicate
ONOO
as the toxic agent
involved in the decrease of the amiloride-sensitive Na+ current. First,
amiloride-sensitive currents were decreased in the presence of SIN-1,
which produces ONOO
, but
not by SIN-1 plus SOD or PAPA-NONOate, which produce · NO. Second, the inhibitory effect of SIN-1 was ameliorated by urate, a
nonspecific scavenger of
ONOO
. Our data further
confirm that, in the presence of SOD or urate, SIN-1 failed to oxidize
DHR to rhodamine (Fig. 1), an index of ONOO
formation (16). The
impermeable nature of SOD further suggests that
ONOO
works within the
extracellular medium to inhibit rENaC function. Although we
have not specifically examined whether
O
2 · is responsible for
the effect of SIN-1, the fact that · NO reacts with
O
2 · at a rate of 6.7 × 109
M
1 · s
1
(22) makes it very unlikely that any free
O
2 · will be generated
by the spontaneous decomposition of SIN-1.
At higher concentrations,
ONOO has been shown to
induce the release of intracellular
Ca2+ (31). This could result in an
increase in protein kinase C activity, which may inhibit rENaC activity
in Xenopus oocytes (2). However, we
did not observe a decrease in the inhibitory effect of SIN-1 in the
presence of BAPTA-AM. At concentrations of 1-10 µM, BAPTA-AM has
been shown to block the effects of intracellular Ca2+ on expressed channel activity
in a number of previous studies (34, 39, 40). It therefore seems
unlikely that, under our experimental conditions, the effects of
ONOO
on rENaC are
mediated through an increase in intracellular
Ca2+ concentration. As previously
mentioned, this conclusion is also supported by the fact that SIN-1 did
not activate the endogenous Ca2+-activated
Cl
currents present in
KCl-injected oocytes (Fig. 4).
Our laboratory has previously demonstrated that
ONOO directly inhibits
Na+ transport across alveolar type
II cells and membrane vesicles isolated from colonic enterocytes (3,
21). In the former study, the decrease in amiloride-sensitive
Na+ uptake coincided with a
decrease in cellular metabolism. It was determined, however, that the
inhibitory effect of ONOO
in alveolar type II cells was not due to these changes in metabolism, as cell viability and
Na+-K+-ATPase
activity were unaffected. In both of these former studies, the effects
were observed following bolus application of
ONOO
(0.1-1 mM) to the
experimental media. In the present study, we elected to use SIN-1 to
generate ONOO
over a period
2 h, which more closely resembles the profile of in vivo
ONOO
generation. A novel
and most interesting aspect of this study is that significant
inhibition of the amiloride-sensitive current was observed at very
small concentrations of
ONOO
, likely to be
encountered in vivo during inflammation. For example, Ischiropoulos et
al. (23) reported that 106 rat
alveolar macrophages stimulated with phorbol myristate acetate generated 0.1 nmol
ONOO
/min. Direct evidence
for the reactivity of ONOO
in vivo has been demonstrated by the presence of nitrotyrosine, the
stable by-product of the interaction of
ONOO
with tyrosine
residues, under a number of inflammatory conditions (5, 17, 26).
Nitration, nitrosylation, or oxidation of key amino acid residues may
account for the observed responses in the present study. For example,
the external loop of -rENaC contains 26 tyrosine, 6 tryptophan, and
14 cysteine residues. In addition, regions at the outer borders of the
two transmembrane (TM) domains contain tyrosines (Y) in
close proximity to one another (Y134 and Y137 in TM1 and Y482, Y484,
and Y485 in TM2), which may form a portion of the outer opening of the
channel. Oxidation or nitration of any or all of these amino acids by
reactive oxygen-nitrogen species may affect the function of this
channel.
We have attempted to determine the nature of
ONOO-induced inhibition by
comparing it with the effects of a second
· NO2 donor, TNM. This
compound has been shown to promote nitration at pH > 7.6 but not at
pH
6.0 (19) and to oxidize sulfhydryl groups over a broad pH range
(36, 37). In the present study, TNM significantly inhibited the
amiloride-sensitive Na+
conductance, but only at pH 6.0. This finding is concordant with the
conclusion that oxidative modification of specific sulfhydryl groups
within the rENaC protein account for a decreased activity. All three
subunits of the protein have highly conserved cysteine-rich regions
within the putative extracellular region of the protein (10) that
represent potential targets for oxidative modification. In previous
work, ONOO
has been shown
to oxidize methionine by a two-electron donation (33). Methionine
oxidation by ONOO
was shown
to account for the inactivation of
1-antitrypsin by ONOO
(30). It is therefore
highly possible that oxidative injury to the rENaC protein will account
for the decreased activity observed in the present study. Oxidation of
critical sulfhydryl groups in nonselective cation channels was found to
alter their biophysical properties through a process that was reversed
by dithiothreitol (27). However, the usefulness of dithiothreitol in
ameliorating the oxidation of rENaC is questionable. It was shown that,
at concentrations as low as 50 µM, dithiothreitol altered the gating of the rENaC channel, an effect attributed to the disruption of coordinated multichannel activity (24).
We believe our data strongly support the conclusion that
ONOO, acting on the
extracellular side of the protein, directly inhibits
-rENaC
activity. However, it should be emphasized that we have no direct
evidence for modification of rENaC by
ONOO
at this time. It is
possible that ONOO
modified
other proteins, such as actin, which in turn may have altered the
function of Na+ channels. Compeau
et al. (12) have shown that endotoxin treatment of distal airway cells
resulted in the reduction of Na+
channel density, corresponding to a reduction in F-actin. Moreover, the
results were dependent on
L-arginine, which suggested a
role for · NO in the process. Other studies have established
the role of the actin in normal
Na+ channel activity (6, 7, 11,
12). Although we cannot exclude the potential effects of
ONOO
on other proteins, the
impermeable nature of SOD and the fact that this enzyme ameliorated the
effect of SIN-1 strongly suggest that damage to intracellular
regulators of rENaC function is not responsible for the
ONOO
-induced responses in
this study.
In summary, our results indicate that, at concentrations as low as 10 µM, ONOO decreases
amiloride-sensitive currents across
Xenopus oocytes expressing
-rENaC. Moreover, this effect is likely due to the oxidation
of specific amino acid residues within the extracellular domain of the
channel proteins. Because
ONOO
is produced in vivo
during acute inflammation (5, 17), these findings point out that
reactive nitrogen species released by inflammatory cells may interfere
with Na+ and fluid homeostasis by
damaging epithelial Na+ channels.
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ACKNOWLEDGEMENTS |
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We acknowledge the assistance of Dr. Lan Chen and Carpantato Myles.
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FOOTNOTES |
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This project was supported by National Heart, Lung, and Blood Institute Grants HL-31197 and HL-51173 and by a grant from the Office of Naval Research (N00014-97-1-0309).
M. D. DuVall is a Parker B. Francis Fellow.
Address for reprint requests: S. Matalon, Dept. of Anesthesiology, Univ. of Alabama at Birmingham, 619 South 19th St., Birmingham, AL 35233-6810.
Received 22 October 1997; accepted in final form 12 February 1998.
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REFERENCES |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
1.
Awayda, M. S.,
I. I. Ismailov,
B. K. Berdiev,
and
D. J. Benos.
A cloned renal epithelial Na+ channel protein displays stretch activation in planar lipid bilayers.
Am. J. Physiol.
268 (Cell Physiol. 37):
C1450-C1459,
1995
2.
Awayda, M. S.,
I. I. Ismailov,
B. K. Berdiev,
C. M. Fuller,
and
D. J. Benos.
Protein kinase regulation of a cloned epithelial Na+ channel.
J. Gen. Physiol.
108:
49-65,
1996[Abstract].
3.
Bauer, M. L.,
J. S. Beckman,
R. J. Bridges,
C. M. Fuller,
and
S. Matalon.
Peroxynitrite inhibits sodium uptake in rat colonic membrane vesicles.
Biochim. Biophys. Acta
1104:
87-94,
1992[Medline].
4.
Beckman, J. S.,
T. W. Beckman,
J. Chen,
P. A. Marshall,
and
B. A. Freeman.
Apparent hydroxyl radical production by peroxynitrite: implications for endothelial injury from nitric oxide and superoxide.
Proc. Natl. Acad. Sci. USA
87:
1620-1624,
1990[Abstract].
5.
Beckman, J. S.,
Y. Z. Ye,
P. G. Anderson,
J. Chen,
M. A. Accavitti,
M. M. Tarpey,
and
C. R. White.
Extensive nitration of protein tyrosines in human atherosclerosis detected by immunohistochemistry.
Biol. Chem. Hoppe-Seyler
375:
81-88,
1994[Medline].
6.
Berdiev, B. K.,
A. G. Prat,
H. F. Cantiello,
D. A. Ausiello,
C. M. Fuller,
B. Jovov,
D. J. Benos,
and
I. I. Ismailov.
Regulation of epithelial sodium channels by short actin filaments.
J. Biol. Chem.
271:
17704-17710,
1996
7.
Berdiev, B. K.,
V. G. Shlyonsky,
O. Senyk,
D. Keeton,
Y. Guo,
S. Matalon,
H. F. Cantiello,
A. G. Prat,
D. A. Ausiello,
I. I. Ismailov,
and
D. J. Benos.
Protein kinase A phosphorylation and G protein regulation of type II pneumocyte Na+ channels in lipid bilayers.
Am. J. Physiol.
272 (Cell Physiol. 41):
C1262-C1270,
1997
8.
Canessa, C. M.,
J. D. Horisberger,
and
B. C. Rossier.
Epithelial sodium channel related to proteins involved in neurodegeneration.
Nature
361:
467-470,
1993[Medline].
9.
Canessa, C. M.,
J. D. Horisberger,
L. Schild,
and
B. C. Rossier.
Expression cloning of the epithelial sodium channel.
Kidney Int.
48:
950-955,
1995[Medline].
10.
Canessa, C. M.,
L. Schild,
G. Buell,
B. Thorens,
I. Gautschi,
J. D. Horisberger,
and
B. C. Rossier.
Amiloride-sensitive epithelial Na+ channel is made of three homologous subunits.
Nature
367:
463-467,
1994[Medline].
11.
Cantiello, H. F.
Role of the actin cytoskeleton on epithelial Na+ channel regulation.
Kidney Int.
48:
970-984,
1995[Medline].
12.
Compeau, C. G.,
O. D. Rotstein,
H. Tohda,
Y. Marunaka,
B. Rafii,
A. S. Slutsky,
and
H. O'Brodovich.
Endotoxin-stimulated alveolar macrophages impair lung epithelial Na+ transport by an L-Arg-dependent mechanism.
Am. J. Physiol.
266 (Cell Physiol. 35):
C1330-C41,
1994
13.
Darley-Usmar, V. M.,
N. Hogg,
V. J. O'Leary,
M. T. Wilson,
and
S. Moncada.
The simultaneous generation of superoxide and nitric oxide can initiate lipid peroxidation in human low density lipoprotein.
Free Radic. Res. Commun.
17:
9-20,
1992[Medline].
14.
Forstermann, U.,
E. I. Closs,
J. S. Pollock,
M. Nakane,
P. Schwarz,
I. Gath,
and
H. Kleinert.
Nitric oxide synthase isozymes. Characterization, purification, molecular cloning, and functions.
Hypertension
23:
1121-1131,
1994[Abstract].
15.
Greis, K. D.,
S. Zhu,
and
S. Matalon.
Identification of nitration sites on surfactant protein A by tandem electrospray mass spectrometry.
Arch. Biochem. Biophys.
335:
396-402,
1996[Medline].
16.
Haddad, I. Y.,
J. P. Crow,
P. Hu,
Y. Ye,
J. Beckman,
and
S. Matalon.
Concurrent generation of nitric oxide and superoxide damages surfactant protein A.
Am. J. Physiol.
267 (Lung Cell. Mol. Physiol. 11):
L242-L249,
1994
17.
Haddad, I. Y.,
G. Pataki,
P. Hu,
C. Galliani,
J. S. Beckman,
and
S. Matalon.
Quantitation of nitrotyrosine levels in lung sections of patients and animals with acute lung injury.
J. Clin. Invest.
94:
2407-2413,
1994[Medline].
18.
Haddad, I. Y.,
S. Zhu,
J. Crow,
E. Barefield,
T. Gadilhe,
and
S. Matalon.
Inhibition of alveolar type II cell ATP and surfactant synthesis by nitric oxide.
Am. J. Physiol.
270 (Lung Cell. Mol. Physiol. 14):
L898-L906,
1996
19.
Haddad, I. Y.,
S. Zhu,
H. Ischiropoulos,
and
S. Matalon.
Nitration of surfactant protein A results in decreased ability to aggregate lipids.
Am. J. Physiol.
270 (Lung Cell. Mol. Physiol. 14):
L281-L288,
1996
20.
Hogg, N.,
V. M. Darley-Usmar,
M. T. Wilson,
and
S. Moncada.
Production of hydroxyl radicals from the simultaneous generation of superoxide and nitric oxide.
Biochem. J.
281:
419-424,
1992[Medline].
21.
Hu, P.,
H. Ischiropoulos,
J. S. Beckman,
and
S. Matalon.
Peroxynitrite inhibition of oxygen consumption and sodium transport in alveolar type II cells.
Am. J. Physiol.
266 (Lung Cell. Mol. Physiol. 10):
L628-L634,
1994
22.
Huie, R. E.,
and
S. Padmaja.
The reaction of NO with superoxide.
Free Radic. Res. Commun.
18:
195-199,
1993[Medline].
23.
Ischiropoulos, H.,
L. Zhu,
and
J. S. Beckman.
Peroxynitrite formation from macrophage-derived nitric oxide.
Arch. Biochem. Biophys.
298:
446-451,
1992[Medline].
24.
Ismailov, I. I.,
M. S. Awayda,
B. K. Berdiev,
J. K. Bubien,
J. E. Lucas,
C. M. Fuller,
and
D. J. Benos.
Triple-barrel organization of ENaC, a cloned epithelial Na+ channel.
J. Biol. Chem.
271:
807-816,
1996
25.
Ismailov, I. I.,
T. Kieber-Emmons,
C. Lin,
B. K. Berdiev,
V. G. Shlyonsky,
H. K. Patton,
C. M. Fuller,
R. Worrell,
J. B. Zuckerman,
W. Sun,
D. C. Eaton,
D. J. Benos,
and
T. R. Kleyman.
Identification of an amiloride binding domain within the -subunit of the epithelial Na+ channel.
J. Biol. Chem.
272:
21075-21083,
1997
26.
Kaur, H.,
and
B. Halliwell.
Evidence for nitric oxide-mediated oxidative damage in chronic inflammation. Nitrotyrosine in serum and synovial fluid from rheumatoid patients.
FEBS Lett.
350:
9-12,
1994[Medline].
27.
Koivisto, A.,
D. Siemen,
and
J. Nedergaard.
Reversible blockade of the calcium-activated nonselective cation channel in brown fat cells by the sulfhydryl reagents mercury and thimerosal.
Pflügers Arch.
425:
549-551,
1993[Medline].
28.
MacMillan-Crow, L. A.,
J. P. Crow,
J. D. Kerby,
J. S. Beckman,
and
J. A. Thompson.
Nitration and inactivation of manganese superoxide dismutase in chronic rejection of human renal allografts.
Proc. Natl. Acad. Sci. USA
93:
11853-11858,
1996
29.
Moncada, S.,
and
A. Higgs.
The L-arginine-nitric oxide pathway.
N. Engl. J. Med.
329:
2002-2012,
1993
30.
Moreno, J. J.,
and
W. A. Pryor.
Inactivation of 1-proteinase inhibitor by peroxynitrite.
Chem. Res. Toxicol.
5:
425-431,
1992[Medline].
31.
Packer, M. A.,
and
M. P. Murphy.
Peroxynitrite causes calcium efflux from mitochondria which is prevented by cyclosporin A.
FEBS Lett.
345:
237-240,
1994[Medline].
32.
Palmer, L. G.
Epithelial Na channels: function and diversity.
Annu. Rev. Physiol.
54:
51-66,
1992[Medline].
33.
Pryor, W. A.,
X. Jin,
and
G. L. Squadrito.
One- and two-electron oxidations of methionine by peroxynitrite.
Proc. Natl. Acad. Sci. USA
91:
11173-11177,
1994
34.
Saugstad, J. A.,
T. P. Segerson,
and
G. L. Westbrook.
Metabotropic glutamate receptors activate G-protein-coupled inwardly rectifying potassium channels in Xenopus oocytes.
J. Neurosci.
16:
5979-5985,
1996
35.
Smith, P. R.,
and
D. J. Benos.
Epithelial Na+ channels.
Annu. Rev. Physiol.
53:
509-530,
1991[Medline].
36.
Sokolovsky, M.,
D. Harell,
and
J. F. Riordan.
Reaction of tetranitromethane with sulfhydryl groups in proteins.
Biochemistry
8:
4740-4745,
1969[Medline].
37.
Sokolovsky, M.,
J. F. Riordan,
and
B. L. Vallee.
Tetranitromethane. A reagent for the nitration of tyrosyl residues in proteins.
Biochemistry
5:
3582-3589,
1966[Medline].
38.
Van der Vliet, A.,
D. Smith,
C. A. O'Neill,
H. Kaur,
V. Darley-Usmar,
C. E. Cross,
and
B. Halliwell.
Interactions of peroxynitrite with human plasma and its constituents: oxidative damage and antioxidant depletion.
Biochem. J.
303:
295-301,
1994[Medline].
39.
Yoshida, S.,
and
S. Plant.
A potassium current evoked by growth hormone-releasing hormone in follicular oocytes of Xenopus laevis.
J. Physiol. (Lond.)
443:
651-667,
1991[Abstract].
40.
Yoshimura, M.,
S. Yoshida,
and
K. Taniyama.
Property of receptor for vasoactive intestinal contractor (VIC) expressed in Xenopus oocytes injected with mRNA from rat intestine.
Life Sci.
58:
1731-1736,
1996[Medline].