Departments of Medicine and Physiology, Richard L. Roudebush Veterans Affairs and Indiana University Medical Centers, Indianapolis, Indiana 46202
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ABSTRACT |
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Nitric oxide (·NO) attenuates hydrogen peroxide
(H2O2)-mediated barrier dysfunction in cultured
porcine pulmonary artery endothelial cells (PAEC) (Gupta MP, Ober MD,
Patterson C, Al-Hassani M, Natarajan V, and Hart, CM. Am J
Physiol Lung Cell Mol Physiol 280: L116-L126, 2001). However,
·NO rapidly combines with superoxide (O), which we
hypothesized would cause PAEC monolayer barrier dysfunction. To test
this hypothesis, we treated PAEC with ONOO
(500 µM) or
3-morpholinosydnonimine hydrochloride (SIN-1; 1-500 µM).
SIN-1-mediated ONOO
formation was confirmed by monitoring
the oxidation of dihydrorhodamine 123 to rhodamine. Both
ONOO
and SIN-1 increased albumin clearance
(P < 0.05) in the absence of cytotoxicity and altered
the architecture of the cytoskeletal proteins actin and
-catenin as
detected by immunofluorescent confocal imaging.
ONOO
-induced barrier dysfunction was partially reversible
and was attenuated by cysteine. Both ONOO
and SIN-1
nitrated tyrosine residues, including those on
-catenin and actin,
and oxidized proteins in PAEC. The introduction of actin treated with
ONOO
into PAEC monolayers via liposomes also
resulted in barrier dysfunction. These results indicate that
ONOO
directly alters endothelial cytoskeletal proteins,
leading to barrier dysfunction.
nitrotyrosine; actin; catenin
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INTRODUCTION |
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THE VASCULAR ENDOTHELIUM provides a barrier that prevents the unregulated passage of fluids and macromolecules across the vessel wall. Reactive oxygen species (ROS), as well as nonoxidant inflammatory mediators, disrupt the endothelial barrier by activating intracellular signaling pathways that stimulate altered endothelial cell (EC) cytoskeletal architecture and the formation of paracellular gaps in the monolayer (14). Endothelial barrier dysfunction in the lung is generally believed to contribute to noncardiogenic edema and to abnormalities in gas exchange and lung compliance in acute respiratory distress syndrome (ARDS). Inhaled nitric oxide (·NO) has been investigated as a therapeutic modality in ARDS because of its ability to promote the selective vasodilatation of pulmonary vessels in ventilated regions of the lung. To date, however, clinical studies have demonstrated that ·NO-induced improvements in gas exchange are short-lived (10, 38). Additional studies, however, indicate that ·NO attenuates oxidant-induced endothelial barrier dysfunction in diverse experimental models (13, 18, 20, 27, 36, 37, 41) and in patients with ARDS (4). Together, these studies suggest that ·NO, delivered at the appropriate time and concentration, exerts a barrier protective effect on vascular endothelium during oxidative stress.
On the other hand, inflammatory mediators that disrupt the endothelial
barrier stimulate the production of superoxide (O) (2). For example, both
bradykinin and the calcium ionophore A-23187 stimulated
ONOO
production via O
in the pathogenesis of acute lung injury and ARDS.
Nitrotyrosine residues were detected in lung tissue from patients with
ARDS (22, 29). Furthermore, bronchoalveolar lavage fluid
and plasma from ARDS patients contain elevated levels of nitrated and
oxidized proteins (15, 31). ONOO
infusion
into a rabbit lung increased hydrostatic pressure, capillary permeability, and lung weight (1) consistent with a
pathogenetic role for ONOO
in lung injury.
ONOO could contribute to endothelial barrier dysfunction
during lung injury by several potential mechanisms. The diffusion of
ONOO
across cell membranes (33) indicates
that ONOO
generated outside the EC compartment as well as
that generated within ECs could directly interact with intracellular
targets. ONOO
reacts with numerous targets including
protein tyrosine residues to form nitrotyrosine (25).
Nitration of target proteins could thereby modulate the structure or
function of molecules, leading to cell dysfunction.
ONOO
-induced oxidant stress could also disrupt actin
polymerization (8) or activate upstream signaling
cascades, e.g., tyrosine kinases (45), that result in the
modification of cytoskeletal proteins, leading to barrier dysfunction.
ONOO
-induced accumulation of cGMP in EC could regulate
vascular EC shape and function (35), whereas activation of
poly(ADP-ribose) synthetase (PARS) by ONOO
can suppress
mitochondrial respiration and cause endothelial dysfunction
(44). These reports emphasize that ONOO
can
potentially alter endothelial function through complex mechanisms. The
current study extends these previous reports by directly examining the
ability of ONOO
to modulate cytoskeletal architecture and
monolayer barrier function in cultured pulmonary artery endothelial
cells (PAEC).
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METHODS |
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PAEC isolation and culture.
A well-characterized model using PAEC monolayers was employed to
examine how ONOO alters endothelial barrier function
independent of its effects on vascular perfusion. EC were isolated from
the main pulmonary artery of pigs as previously reported
(23). Once confluent, PAEC were passaged by treatment with
trypsin to polycarbonate filters or 24-well tissue culture plates.
After cells reached secondary confluence, the concentration of fetal
bovine serum was decreased from 10% to 4% (maintenance medium). In
all experiments, PAEC were studied 3-6 days after confluence, with
control and experimental dishes matched according to cell line, number
of passages, and number of days postconfluence. Monolayers were
identified as ECs by phase-contrast microscopy and periodically by
immunofluorescent staining for factor VIII antigen (5).
ONOO treatment protocols.
PAEC were exposed to either ONOO
(Cayman Chemicals, Ann
Arbor, MI) or 3-morpholinosydnonimine hydrochloride (SIN-1; Molecular Probes, Eugene, OR), a compound that simultaneously generates ·NO and
O
(3). Stock solutions of
ONOO
(20 mM in 0.3 M NaOH) and SIN-1 [10 mM in Hanks'
balanced salt solution (HBSS; GIBCO)] were prepared immediately before
addition to the culture medium. PAEC monolayers were treated for 2 h with RPMI 1640 medium containing 4% fatty acid-free albumin (Sigma), 20 mM HEPES (Sigma), and ONOO
or SIN-1 (1-500 µM)
at 37°C. In selected experiments, because of the short half-time of
ONOO
in neutral solutions (30), PAEC
were also treated with repeated doses of ONOO
(150 µM)
every minute for 4-8 min. In other experiments, cysteine (5 mM for
1 h before treatment with SIN-1), a scavenger of
ONOO
(6), was included during treatment with
SIN-1. In all experiments, control monolayers were treated with culture
medium to which an equivalent volume of NaOH or HBSS vehicle had been added.
Measurement of PAEC cytotoxicity.
Each PAEC monolayer was treated with ONOO or SIN-1 for
2 h. To determine the effect of ONOO
or SIN-1 on
PAEC viability, we collected the medium from each monolayer. PAEC were
then washed thoroughly with HBSS and collected by scraping with a
rubber policeman into 10 mM Tris · HCl buffer, pH 7.4, containing 0.2% Triton X-100. PAEC injury was assessed by measuring
the release of intracellular lactate dehydrogenase (LDH) as described
previously (23). Duplicate aliquots (40 µl) of culture
medium and cell lysates were placed in 96-well microtiter plates, and
LDH activity was measured by monitoring the consumption of nicotinamide
adenine dinucleotide (Sigma) at 340 nm with a spectrophotometric plate
reader (17). Results are expressed as the percentage of
total LDH activity released to the culture medium: %LDH release = [LDH activity in medium/(LDH activity in cells + medium)] × 100.
Measurement of SIN-1-generated reactive species.
ONOO generation by SIN-1 was confirmed by
spectrophotometrically monitoring the oxidation of dihydrorhodamine 123 (DHR; Molecular Probes) to rhodamine at 500 nm (molar extinction
coefficient = 78,000 M
1 · cm
1)
(21). ONOO
, but neither
O
Measurement of PAEC barrier function.
The ability of monolayers to prevent the transendothelial passage of
albumin was measured as an index of PAEC function as previously
reported (23, 39). In brief, PAEC monolayers on polycarbonate filters attached to plastic ring wells were floated in 45 ml of RPMI 1640 medium, forming upper and lower chambers, respectively.
Evans blue dye bound to 4% bovine serum albumin in RPMI 1640 plus 20 mM HEPES buffer was gently placed inside the luminal (upper) well of
each chamber. The abluminal (lower) medium was continuously stirred and
was maintained at 37°C by a circulating water bath. Before
experimental interventions took place, samples (0.3 ml) were taken from
the abluminal medium at 10-min intervals for 1 h and placed into a
96-well microtiter plate. After baseline albumin clearance had been
measured for 1 h, ONOO or SIN-1 was added to the
luminal medium, and albumin clearance was monitored for an
additional 2 h. The absorbance of the samples was read at 620 nm
with a spectrophotometric plate reader. The raw absorbance data were
transferred to a spreadsheet program on an IBM personal computer for
analysis. The net abluminal sample absorbance was used to calculate the
equivalent theoretical volume of luminal medium cleared to the
abluminal space (39). The clearance of albumin (expressed
in µl/min) was calculated by linear regression analysis of changes in
absorbance over 1-h intervals. To determine the recovery of PAEC
barrier function, PAEC monolayers treated with 500 µM SIN-1 or HBSS
vehicle alone in RPMI medium for 2 h were washed and placed back
into maintenance medium for 24 h before albumin clearance was
determined for 1 h.
Detection of nitrotyrosine by immunoprecipitation.
PAEC in 100-mm dishes were treated for 2 h with 500 µM SIN-1,
500 µM ONOO, or RPMI 1640 medium with 4% albumin and
20 mM HEPES alone. The cells were then solubilized in lysis buffer [30
mM potassium phosphate, pH 7.4, 150 mM NaCl, 10% (vol/vol) glycerol, 1 µM phenylmethylsulfonyl fluoride,10 µg/ml aprotinin, and 1% Triton
X-100] as previously reported (34). Equal amounts of PAEC
lysates were precleared with Sepharose and exposed to monoclonal
anti-nitrotyrosine antibodies (Cayman Chemicals) overnight. The lysates
were then combined with Sepharose and centrifuged, and proteins were
eluted with a saturated nitrotyrosine solution. The proteins were then
resolved using SDS-PAGE (4-12%). Samples were run in parallel
with albumin (0.2% in phosphate buffer, pH 7.4), nitrated albumin
(0.2% albumin in phosphate buffer, pH 7.4, treated with 500 µM
ONOO
and vortexed immediately), and molecular weight
markers. The proteins were transferred to polyvinylidene difluoride
membranes (Millipore, Bedford, MA), blocked with 1% albumin and 1%
goat serum in phosphate-buffered saline with Tween 20, and probed with monoclonal anti-nitrotyrosine antibodies (1:1,000). The membrane was
then treated with horseradish peroxidase-conjugated goat anti-mouse antibody (1:5,000; Jackson ImmunoResearch, West Grove, PA), and bands
were detected using enhanced chemiluminescence (ECL; Pierce, Rockford,
IL). In selected experiments, nitrotyrosine immunoprecipitates were
subjected to SDS-PAGE, followed by slot blotting with antibodies to
actin (1:250; courtesy of Dr. Fred Pavalko) or
-catenin (1:250; Zymed, San Francisco, CA).
Detection of oxidized proteins.
After treatment with control, SIN-1, SIN-1 plus SOD (15 U/ml), or
ONOO, PAEC lysates were prepared as described in
Detection of nitrotyrosine by immunoprecipitation, and
oxidized proteins were detected with a commercially available kit
(Oxyblot; Oncor). Briefly, oxidant-induced carbonyl side chains
were derivatized to 2,4-dinitrophenylhydrazone (DNP-hydrazone) by
2,4-dinitrophenylhydrazine (DNPH). The samples were then resolved with
SDS-PAGE as described above. DNP residues were then detected with the
use of polyclonal rabbit anti-DNP antibodies, followed by treatment
with secondary antibodies and ECL as described above.
Effects of ONOO on PAEC cytoskeletal architecture.
PAEC were seeded onto gelatinized glass coverslips in 35-mm dishes and
grown to confluence. Coverslips were then incubated in RPMI 1640 medium
containing 4% bovine serum albumin and 20 mM HEPES with vehicle,
SIN-1, or ONOO
for 1-2 h at 37°C. After fixation
with 5% paraformaldehyde in wash buffer (150 mM NaCl, 0.1% Na-azide,
and 50 mM Tris · HCl, pH 7.6) for 10 min at room temperature,
the coverslips were rinsed and then permeabilized for 3.5 min with
0.2% Triton X-100 in wash buffer. After coverslips were rinsed and
blocked with 1% BSA in buffer for 1 h, they were treated with
monoclonal anti-
-catenin antibody (1:50 in 1% BSA/buffer;
Transduction Laboratories, Lexington, KY) overnight at 4°C. After
they were thoroughly washed, the coverslips were treated with
FITC-labeled donkey anti-mouse IgG (1:50; Jackson ImmunoResearch) and
rhodamine phalloidin (1:200; Molecular Probes) in 1% BSA/buffer. The
coverslips were then rinsed and mounted on slides with
SlowFade/glycerol (Molecular Probes). Fluorescence was observed by
confocal microscopy (1024 System; Bio-Rad, Hercules, CA) with a ×60
oil objective by using a krypton-argon laser at 10% laser power with
an iris aperture of 3 mm. Rhodamine and FITC fluorescence were each
recorded for 10-17 sections at 0.5-mm intervals by using identical
contrast and gain settings. The data were processed using MetaMorph
(Universal Imaging, West Chester, PA). A minimum of six images from
each treatment group was analyzed. Representative images were imported
to PowerPoint for printing by Kodak dye sublimation (Rochester, NY).
Effect of ONOO-treated actin on PAEC barrier
function.
In an attempt to determine the direct effects of ONOO
on
actin and PAEC barrier function, nonmuscle actin from human platelets (Cytoskeleton, Denver, CO) was treated with 500 µM ONOO
for 1 h. Treated actin was then dialyzed against four exchanges of
5 mM Tris · HCl, pH 8.0, 0.2 mM CaCl2, 0.2 mM ATP,
and 0.5 mM dithiothreitol. Dialyzed ONOO
-treated or
control actin (1 mg/ml) was incubated for 20 min with an equal volume
(5 µl) of Lipofectamine (GIBCO BRL, Gaithersburg, MD) and then added
to the luminal medium above PAEC monolayers, and albumin clearance was
measured as described in Measurement of PAEC barrier
function.
Statistical analysis. In all experiments data were analyzed with analysis of variance or repeated-measures analysis of variance to determine the significance of treatment effects, followed by Bonferroni or Student-Newman-Keuls analysis to examine differences between individual treatment groups. The level of statistical significance was taken as P < 0.05.
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RESULTS |
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SIN-1-induced PAEC cytotoxicity.
Treatment with 10-500 µM ONOO or 10-500 µM
SIN-1 for 2 h in RPMI 1640 plus 4% albumin and 20 mM HEPES did
not cause PAEC cytotoxicity assessed as %LDH release (Table
1). Phase-contrast microscopy confirmed
no cytolytic alterations in PAEC morphology in ONOO
- or
SIN-1-treated cells (not shown).
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Effect of ONOO and SIN-1 on PAEC barrier function.
After baseline albumin clearance was measured for 1 h, PAEC were
treated with 1-500 µM SIN-1 (Fig.
1A) or bolus addition of ONOO
(Fig. 1B). As previously reported
(23, 26), confluent PAEC present a substantial barrier to
the transmonolayer clearance of albumin that is stable during the 3-h
measurement period (Fig. 1B). SIN-1 failed to cause barrier
dysfunction initially but caused significant barrier dysfunction during
the second hour following its addition that demonstrated no significant
concentration dependence between 1 and 500 µM. Bolus addition of
ONOO
caused more extensive barrier dysfunction but, like
SIN-1, had minimal effect during the first hour of treatment (Fig.
1B). Repeated dosing with a lower ONOO
concentration (150 µM) each minute for 4-8 min also caused
significant barrier dysfunction after 2 h (Fig. 1B). In
monolayers treated with 500 µM ONOO
for 2 h,
followed by washing and incubation in maintenance medium for 24 h
(Fig. 2, recovery), barrier dysfunction
was only partially reversible. Although cysteine alone increased
baseline albumin clearance, it significantly inhibited SIN-1-mediated
barrier dysfunction (Fig. 2).
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Characterization of reactive species generated by SIN-1.
To examine the kinetics of ONOO generation,
SIN-1-mediated DHR oxidation was measured under conditions identical to
those employed in Fig. 1A. SIN-1-mediated DHR oxidation was
both dose (1-500 µM) and time (15-120 min) dependent and
was fully inhibited by the presence of 5 mM cysteine (not shown). As
shown in Fig. 3A, SIN-1 caused
little DHR oxidation during the first hour of treatment, similar to the
time course of SIN-1-induced barrier dysfunction (Fig. 1).
SIN-1-mediated DHR oxidation was partially inhibited by SOD (15 U/ml)
or c-PTIO (100 µM) and more fully inhibited by the combination of
c-PTIO and SOD. In contrast, catalase (50 U/ml) had no significant
effect on SIN-1-mediated DHR oxidation, indicating that
H2O2 was not the oxidizing agent (Fig.
3A). In the absence of SIN-1, PAEC caused low levels of DHR
oxidation that changed little during the 2-h treatment interval
studied. Figure 3B shows that under the conditions employed
for measurements of barrier function, in the absence of SOD,
SIN-1-mediated ·NO generation was nearly undetectable, consistent
with its rapid reaction with O
. In the presence of SOD, however, ·NO production
by SIN-1 became detectable, due to SOD-mediated conversion of
O
generation (Fig. 3A).
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Effect of c-PTIO and SOD on SIN-1-mediated PAEC barrier
dysfunction.
Because SOD and c-PTIO attenuated SIN-1-mediated ONOO
generation and DHR oxidation, the ability of these interventions to modulate SIN-1-mediated PAEC barrier dysfunction was examined. After
baseline albumin clearance was measured for 1 h, PAEC monolayers on polycarbonate filters were exposed to 500 µM SIN-1 for 2 h in
RPMI 1640 plus 4% albumin and 20 mM HEPES, in the presence or absence
of c-PTIO (100 µM), SOD (15 U/ml), or both c-PTIO and SOD. Treatment
with SIN-1 for 2 h caused significant barrier dysfunction (Fig.
4), consistent with the findings reported
in Fig. 1. SIN-1-mediated barrier dysfunction was not attenuated by
either c-PTIO or SOD individually and was significantly exacerbated by
the combination of c-PTIO and SOD (Fig. 4). We postulated that
exacerbation of SIN-1-mediated barrier dysfunction was caused by
SOD-induced generation of H2O2 combined with
c-PTIO-induced removal of barrier protective effects of ·NO
(20). Therefore, in separate studies, PAEC monolayers were
treated with SIN-1 alone, the combination of SIN-1, c-PTIO, and SOD (as
described above), or SIN-1, c-PTIO, and SOD plus catalase (50 U/ml). In
these studies (n = 3), barrier dysfunction 2 h
after the addition of SIN-1, c-PTIO, and SOD was 199.2% of the
baseline value, whereas the addition of catalase reduced barrier
dysfunction to 144.2% of the baseline value.
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Detection of ONOO-induced protein modifications.
PAEC were treated with 500 µM ONOO
or SIN-1 for 2 h and then collected as described in METHODS. Lysates were
immunoprecipitated with monoclonal anti-nitrotyrosine antibodies, and
equal amounts of protein were then resolved with Western blotting and
probed with the same anti-nitrotyrosine antibody. Compared with PAEC treated under control conditions, treatment with either SIN-1 or
ONOO
increased the number of proteins with nitrotyrosine
epitopes (molecular mass ranging from ~20 to 125 kDa) (Fig.
5A). In control cells,
nitrotyrosine epitopes were detected in proteins with molecular masses
of ~41 and 125 kDa with faint bands at ~36 and 200 kDa (Fig.
5A, arrowheads). In ONOO
- and SIN-1-treated
PAEC, roughly eight additional proteins displayed evidence of tyrosine
nitration (Fig. 5A, arrows). The anti-nitrotyrosine immunoprecipitates were also probed with antibodies to actin and
-catenin. These studies (Fig. 5B) show that proteins with
molecular masses of ~40 and 84 kDa react with both anti-nitrotyrosine
and actin or
-catenin antibodies, respectively. Whole cell lysates prepared from PAEC treated with control, SIN-1, SIN-1 plus SOD, or
ONOO
for 2 h were also examined for evidence of
protein oxidation. Compared with control conditions in which oxidized
proteins were detected with molecular masses of ~42, 84, and 209 kDa
(Fig. 6, arrowheads), treatment with
SIN-1, ONOO
, or SIN-1 plus SOD increased the oxidation of
PAEC proteins as evidenced by the detection of more DNPH epitopes in
proteins with molecular masses of ~21, 30, 33, 53, and 130 kDa, along
with increased intensity of staining in the 42- and 209-kDa bands (Fig.
6, arrows). The presence of SOD had little effect on SIN-1-induced PAEC
protein oxidation, suggesting that the observed oxidation was mediated by reactive species other than O
generation, the current studies cannot discriminate whether
SIN-1-mediated protein oxidation was caused by ONOO
,
H2O2, other reactive compounds, or combinations
of these species. The comparable pattern of protein oxidation detected
in SIN-1, ONOO
, and SIN-1 plus SOD suggests that
ONOO
may be responsible.
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Effect of ONOO on PAEC cytoskeletal architecture.
The cytoskeletal architecture of vascular ECs plays an important role
in the regulation of EC shape and barrier function. To examine the
effects of ONOO
on PAEC cytoskeletal organization, PAEC
were stained with rhodamine phalloidin to examine the structure of
filamentous (F)-actin and with
-catenin antibodies to examine the
distribution of this protein, which links the actin cytoskeleton to the
adherens junction complex (19). Identical, representative
fields obtained by confocal immunofluorescence microscopy are shown for
F-actin (Fig. 7) and
-catenin (Fig.
8). The F-actin staining of control
PAEC shows the typical, regular polygonal arrangement of cultured EC
with a fine weblike network of cellular actin (Fig. 7A). The
F-actin is most prominent in dense peripheral bands that define the
cell borders (Fig. 7A, arrow). Treatment with
ONOO
increased F-actin content and caused reorganization
into more parallel arrays of actin fibers, typical of a contractile
state (Fig. 7B, arrowhead). Similar F-actin reorganization
was observed 1 h after treatment with 100 µM SIN-1 (Fig.
7C). Two hours after the addition of 100 µM SIN-1, partial
dissolution of the thick parallel actin filaments and diminution of
total actin staining were observed, consistent with ongoing actin
reorganization (Fig. 7D). Comparable but more accelerated
alterations in actin fibers were seen after treatment with higher
concentrations of SIN-1. For instance, treatment with 500 µM SIN-1
caused total dissolution of the dense peripheral bands and apparent
reorganization of actin into parallel fibers within 1 h (Fig.
7E), comparable to the alterations observed after treatment
with 100 µM SIN-1 for 2 h. Compared with the 100 µM
concentration, 500 µM SIN-1 also caused a more rapid and extensive
progression of actin depolymerization, manifest as loss of rhodamine
staining intensity after treatment for 1 and 2 h (Fig.
7F). In addition, the higher dose of SIN-1 caused a dramatic
change in cell shape to a contractile, spindle shape (Fig.
7F, open arrow). Thus ONOO
caused activation
of PAEC to a contractile state, followed by dissolution of total
filamentous polymerized actin that was roughly proportional to the dose
and duration of ONOO
to which the cells were exposed.
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Effect of actin treated with ONOO on PAEC barrier
function.
To more directly address ONOO
effects on cytoskeletal
protein function, we treated nonmuscle actin with ONOO
in
vitro and then introduced it into PAEC monolayers with liposomes as
described in METHODS. These ONOO
treatment
conditions caused nitration of nonmuscle actin as confirmed by
immunoblotting (not shown). As shown in Fig.
9, compared with baseline, empty
liposomes increased albumin clearance slightly, and liposomes
containing native nonmuscle actin had little effect on PAEC barrier
function, whereas liposomes containing ONOO
-treated actin
caused significant increases in albumin clearance. None of these
treatment conditions caused cytotoxicity (LDH release or
phase-contrast microscopy, not shown).
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DISCUSSION |
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Evidence for protein nitration and oxidation in lung tissue,
plasma, and bronchoalveolar lavage fluid from patients with ARDS (15, 22, 29, 31) suggests that ONOO may
participate in the pathogenesis of lung injury. However, direct
evidence establishing ONOO
and nitrotyrosine formation as
mediators of vascular pathology as opposed to markers of inflammation
has been lacking. Therefore, the current study examined the effects of
ONOO
on endothelial barrier function as a relevant
physiological parameter of vascular endothelial function, impairment of
which contributes to vascular leakiness, interstitial edema, and organ
dysfunction. The direct extrapolation of these in vitro studies to
human pathophysiology remains limited by our lack of understanding of
relevant concentrations of ONOO
generated in vivo.
Therefore, the goal of the present investigation was to identify novel
targets of ONOO
reactivity that might allow more focused
investigations in future in vivo studies.
ONOO added as a bolus or generated continuously by SIN-1
caused significant PAEC monolayer barrier dysfunction that was not associated with PAEC cytotoxicity as determined by LDH release and
morphological examination. Furthermore, ONOO
-mediated
barrier dysfunction was partially reversible and attenuated by
cysteine, a known scavenger of ONOO
(6).
Although recent reports have suggested that SIN-1 can generate a
variety of reactive species (32), several lines of evidence in our study confirm that SIN-1 generated ONOO
under the conditions employed. First, SIN-1-induced DHR oxidation was
attenuated by SOD and c-PTIO, indicating that scavenging of either
O
formation.
Furthermore, catalase failed to attenuate SIN-1-induced DHR oxidation,
indicating that H2O2 did not account for SIN-1 reactivity. Second, addition of SOD to SIN-1 containing medium increased the amount of ·NO that was released into the culture medium
bathing PAEC during measurements of barrier function, indicating that
SIN-1 generates both O
. Together, these
results demonstrate that SIN-1 generated ONOO
under the
conditions employed in the measurement of PAEC barrier function.
Although scavenging either ·NO or O formation and DHR oxidation by SIN-1,
neither c-PTIO nor SOD attenuated SIN-1-mediated barrier dysfunction,
and the addition of both c-PTIO and SOD exacerbated SIN-1-induced PAEC
barrier dysfunction (Fig. 4). c-PTIO, a potent scavenger of ·NO, can, under certain conditions, modulate reactions of ONOO
and
·NO by other mechanisms that remain to be completely defined (40). Under conditions employed in the current study,
neither c-PTIO nor SOD completely prevented DHR oxidation (Fig. 3) or attenuated barrier dysfunction (Fig. 4) caused by SIN-1. These findings
suggest that SIN-1 produced ONOO
that was detectable only
after 1 h (Fig. 3), accounting for the delayed development of
barrier dysfunction. However, barrier dysfunction caused by bolus
addition of ONOO
was also delayed and developed only
after 1 h of treatment. Coupled with the poor correlation between
barrier dysfunction and SIN-1 concentration (Fig. 1A), these
findings suggest that ONOO
might alter endothelial
function through cumulative effects on cytoskeletal dynamics that are
more dependent on time than ONOO
concentration.
Alternatively, the inability of ·NO or O
and the continuous
generation of ONOO
with SIN-1 cause PAEC monolayer
barrier dysfunction that is partially reversible, unassociated with
cytotoxicity, and comparable in extent to barrier dysfunction induced
by ROS.
The exact mechanisms by which ONOO causes PAEC barrier
dysfunction remain to be defined. At physiological pH,
ONOO
nitrates tyrosine residues in actin, a cytoskeletal
protein critical for maintenance of EC shape and barrier function
(43). In addition to the actin cytoskeleton, adherens
junctions are a critical structure in cell-cell adhesion and the
formation of the endothelial barrier (9). Adherens
junctions are composed of complexes of membrane-spanning cadherin
proteins and intracellular catenins. The extracellular domain of
cadherin links to the cadherin expressed on adjacent cells via
Ca2+-dependent binding. The intracellular domain of
cadherin binds to
-catenin, which links to
-catenin to attach the
actin cytoskeleton to the adherens junction complex (42).
The current study provides several lines of evidence that
ONOO
alters these cytoskeletal proteins to disrupt
endothelial barrier function. Treatment with either SIN-1 or
ONOO
, under conditions identical to those causing barrier
dysfunction, resulted in PAEC protein nitration and oxidation.
Alterations in the structure of PAEC actin and
-catenin detected
with confocal immunofluorescent microscopy provide additional evidence
for these cytoskeletal proteins serving as protein targets or effectors in ONOO
-induced endothelial barrier dysfunction. The
barrier disruptive effects of cytoskeletal proteins nitrated with
tetranitromethane, a powerful oxidizing and nitrating agent, were
previously reported in cultured EC (12). The current study
demonstrated that actin treated with ONOO
, but not native
actin, caused barrier dysfunction when introduced into PAEC with
liposomes (Fig. 9). We speculate that nitrated actin is incorporated
into endogenous actin and alters the dynamics of actin polymerization
and depolymerization, thereby changing cell shape and causing barrier
dysfunction. The barrier disruptive effect of
ONOO
-induced nitration of actin and
-catenin is
further supported by results showing that proteins immunoprecipitated
from ONOO
-treated PAEC with a monoclonal
anti-nitrotyrosine antibody reacted with antibodies to actin and
-catenin (Fig. 5B). These results indicate that
ONOO
directly alters EC cytoskeletal proteins, leading to
EC barrier dysfunction. Although we have identified actin and
-catenin as relevant targets for the barrier disruptive effects of
ONOO
, other nitrated and oxidized proteins may well
contribute to ONOO
-induced EC barrier dysfunction. The
identification of these additional ONOO
-altered proteins
constitutes an area of active investigation in our laboratory.
In addition to directly oxidizing or nitrating PAEC cytoskeletal
proteins, ONOO could potentially activate intracellular
signaling pathways that regulate endothelial barrier function. For
instance, oxidants stimulate tyrosine phosphorylation in cultured EC
(45). Tyrosine phosphorylation of adherens junction
proteins disrupts the functional integrity of the adherens junction
complex (46). ONOO
can either activate
tyrosine kinases or inhibit tyrosine kinase-mediated signaling through
tyrosine nitration (16). However, in preliminary studies
we found that inhibiting tyrosine kinases had no effect on
SIN-1-mediated PAEC barrier dysfunction (data not shown), suggesting that ONOO
-mediated tyrosine kinase activation does not
constitute a major pathway mediating ONOO
-induced
endothelial barrier dysfunction. Although other cellular effects of
ONOO
could also contribute to PAEC barrier dysfunction,
including ONOO
-mediated activation of PARS and subsequent
NAD+ depletion and inhibition of mitochondrial respiration
as reported in human umbilical vein endothelial cells
(44), or to ONOO
-induced alterations in EC
calcium (11), our results suggest that the direct effects
of ONOO
on key cytoskeletal proteins causes barrier dysfunction.
In summary, ONOO, either as a bolus or generated
continuously by SIN-1, caused significant PAEC barrier dysfunction that
was not associated with cytotoxicity. This barrier dysfunction was partially inhibited by cysteine, suggesting that it is mediated by
ONOO
. Several proteins from ONOO
-treated
PAEC displayed evidence of nitration and oxidation. Particularly relevant to endothelial barrier function, ONOO
caused
nitration of actin and
-catenin, suggesting that these cytoskeletal
protein modifications contribute to barrier dysfunction. Actin treated
with ONOO
and introduced into PAEC also caused
significant barrier dysfunction. Together, these findings demonstrate
that ONOO
modifies cytoskeletal proteins, resulting in
changes in cell shape and subsequent barrier dysfunction. Additional
studies are required to clarify other proteins that are nitrated and/or
oxidized by ONOO
to define how these modifications relate
to protein structure and function and to determine the relative
contribution of oxidation and tyrosine nitration to alterations in
protein and cell function. The current study expands the list of
proteins that are modified by ONOO
and identifies novel
cytoskeletal targets in the lung parenchyma as candidates for
ONOO
reactivity.
![]() |
ACKNOWLEDGEMENTS |
---|
We gratefully acknowledge the expert technical assistance of Shehnaz Khan, Delbert Bauzon, and Dean Kleinhenz.
![]() |
FOOTNOTES |
---|
This work was supported by the Roudebush Veterans Affairs Medical Center Research Service, National Heart, Lung, and Blood Institute (NHLBI) Grant PO1-HL-58064, NHLBI Training Grant T32-HL-07774, and the American Diabetes Association.
Address for reprint requests and other correspondence: C. Michael Hart, Division of Pulmonary and Critical Care Medicine, Atlanta Veterans Affairs Medical Center (151-P), 1670 Clairmont Rd., Decatur, GA 30033 (E-mail: michael.hart.3{at}med.va.gov).
The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
Received 10 July 2000; accepted in final form 30 April 2001.
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