1 Division of Pulmonary and Critical Care Medicine and Division of Cardiology, Department of Medicine, Johns Hopkins University School of Medicine, Baltimore 21224; and 2 Division of Infectious Disease, University of Maryland School of Medicine, Baltimore, Maryland 21201
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ABSTRACT |
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Hyperoxia increases reactive oxygen species (ROS) production in vascular endothelium; however, the mechanisms involved in ROS generation are not well characterized. We determined the role and regulation of NAD(P)H oxidase in hyperoxia-induced ROS formation in human pulmonary artery endothelial cells (HPAECs). Exposure of HPAECs to hyperoxia for 1, 3, and 12 h increased the generation of superoxide anion, which was blocked by diphenyleneiodonium but not by rotenone or oxypurinol. Furthermore, hyperoxia enhanced NADPH- and NADH-dependent and superoxide dismutase- or diphenyleneiodonium-inhibitable ROS production in HPAECs. Immunohistocytochemistry and Western blotting revealed the presence of gp91, p67 phox, p22 phox, and p47 phox subcomponents of NADPH oxidase in HPAECs. Transfection of HPAECs with p22 phox antisense plasmid inhibited hyperoxia-induced ROS production. Exposure of HPAECs to hyperoxia activated p38 MAPK and ERK, and inhibition of p38 MAPK and MEK1/2 attenuated the hyperoxia-induced ROS generation. These results suggest a role for MAPK in regulating hyperoxia-induced NAD(P)H oxidase activation in HPAECs.
reactive oxygen species; lung vascular endothelial cells; mitogen-activated protein kinases
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INTRODUCTION |
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PATIENTS with acute respiratory distress syndrome (ARDS) or chronic obstructive pulmonary diseases (COPD) and subjects suffering from severe hypoxia due to drowning, life-threatening soft-tissue infections, and exposure to smoke and toxic gases are often rescued by oxygen therapy (13). Although oxygen therapy is given to the critically ill patients, prolonged exposure to high concentrations of oxygen results in ventilator-induced lung injury (24). Thus exposure to supraphysiological concentrations of oxygen (hyperoxia) results in sustained elevation of inspired oxygen, extensive damage to the alveolar-capillary barrier, increased permeability, and decreased pulmonary function (11). Hyperoxia-induced pathological changes in the lung tissue are often accompanied by injury and death of endothelial and epithelial cells (14). Apoptosis plays an important role in animal models of hyperoxic lung injury as well as in ARDS (28). Molecular mechanisms of hyperoxia-induced lung injury and cell death are complex and regulated by generation of overwhelming levels of reactive oxygen species (ROS), cytokine-mediated inflammation, loss of antioxidant defense mechanisms and modulation of signal transduction pathways that regulate the expression of stress responsive and apoptotic regulatory genes (4, 35).
Hyperoxia-induced formation of superoxide (O and their potential role in pulmonary oxygen
toxicity, as well as the efficacy of superoxide dismutase (SOD) for
pulmonary defense against oxygen toxicity, have been emphasized
(52). It was shown that exposure to hyperoxia induced both
NADH- and NADPH-dependent generation of O
Aerobes generate ROS by nonenzymatic and enzymatic mechanisms
(50). Molecular O2 is activated by enzymatic
electron transfer in subcellular organelles such as mitochondria,
endoplasmic reticulum, nuclear membranes, peroxisomes, plasma
membranes, and cytosol (25). Mammalian cells possess
several potential sources of generation of ROS that include but are not
limited to the mitochondrial electron transport system, arachidonic
acid metabolism by cyclooxygenase/lipoxygenase, cytochrome
P-450, xanthine oxidase, NAD(P)H oxidase, nitric oxide synthase, peroxidase, and other heme proteins (3).
However, the professional phagocytes of the immune system (neutrophils, eosinophils, monocytes, and macrophages) primarily generate
O
Despite the fact that a majority of the earlier studies reported the
formation/release of H2O2 independent from the
xanthine oxidase-mediated mechanism, other potential contributor(s)
responsible for hyperoxia-induced formation/release of
O
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MATERIALS AND METHODS |
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Materials.
HPAECs (passage 4) were obtained from Clonetics (San Diego,
CA). MCDB medium and phosphate-buffered saline (PBS) were
obtained from Biofluids (Rockville, MD). Nonessential amino acids,
trypsin, FBS, penicillin-streptomycin, Tris · HCl, EGTA,
MgCl2, glycerophosphate, Triton X-100, sodium
orthovanadate, aprotinin, leupeptin, pepstatin, Tween 20, ferricytochrome C, human erythrocyte SOD, bovine liver catalase,
H2O2 (30%), diphenyleneiodonium (DPI),
lucigenin, and DMSO were all obtained from Sigma (St. Louis, MO).
Amplex Red Hydrogen Peroxide Assay kit, hydroethidine, and
6-carboxy-2',7'-dichlorodihydroxyfluorescein diacetate dicarboyxy
methyl ester (DCFDA) were obtained from Molecular Probes (Eugene, OR).
5,5-Dimethyl-1-pyrroline-N-oxide (DMPO) spin trap
was obtained from Dojindo Laboratories (Gaithersburg, MD). [-32P]ATP (specific activity 5 Ci/mmol in 10 mM Tris
buffer) and 2-deoxy-D-[3H]glucose were
obtained from New England Nuclear (Wilmington, DE). SB-202190,
SB-203580, and PD-98059 were obtained from Calbiochem (San Diego, CA).
Endothelial cell growth factor (ECGF) and antibodies against
recombinant subcomponents of phagocytic NADPH oxidase (phox 22, phox
47, phox 67 and gp 91) were obtained from Upstate Biotechnology (Lake
Placid, NY). Phospho-specific p38 MAPK and ERK1/ERK2 antibodies were
purchased from Cell Signaling (Beverly, MA). Polyclonal antibodies to
ERK1, ERK2, p38 MAPK, and c-Jun NH2-terminal kinase (JNK)
were procured from Santa Cruz Biotechnology (Santa Cruz, CA). The
enhanced chemiluminescence kit for the detection of proteins by Western
blots was obtained from Amersham (Arlington Heights, IL).
Endothelial cell culture. HPAECs, at passages between 5 and 9 were grown to ~80% confluence in MCDB medium supplemented with 10% FBS, 100 U/ml penicillin and streptomycin, 5 µg/ml ECGF, 1% nonessential amino acids, and 1 µg/ml hydrocortisone at 37°C in a 5% CO2-95% air atmosphere as described earlier (40). Cells were grown in 35- or 100-mm dishes or T-75 flasks and were used in all of the experiments.
Assay of endothelial injury and cytotoxicity. The morphology of ECs and detachment of cells from the monolayers were assayed by phase-contrast microscopy. The uptake of trypan blue and 2-deoxy-D-[3H]glucose release by the cells were measured as indexes of hyperoxia-induced cytotoxicity (39).
Exposure of cells to hyperoxia. HPAECs in complete MCDB medium were placed in a dry and airtight modular incubator chamber (Billups-Rothenberg, Del Mar, CA), flushed continuously with 95% O2-5% CO2 for 30 min until the oxygen level inside the chamber reached ~95%, and placed in a cell culture incubator at 37°C for the desired lengths of time of incubation. The concentration of O2 inside the chamber was continuously monitored with a digital oxygen monitor. The buffering capacity of cell culture medium did not alter throughout the study period under hyperoxic exposure and maintained neutral pH (~7.4).
Determination of O
Determination of H2O2 by Amplex red assay. H2O2 formation in the medium was determined by fluorescence method using Amplex Red Hydrogen Peroxide Assay kit (Molecular Probes). HPAECs (~80% confluent in 60-mm dishes) were exposed to normoxia (95% air-5% CO2) and hyperoxia (95% O2-5% CO2) for required lengths of time in 1.0 ml of phenol red-free medium (GIBCO-BRL medium 199) in the absence and presence of pharmacological inhibitors for specified lengths of time. At the end of incubation, the medium was collected and centrifuged at 4,000 g for 5 min, and fluorescence of the medium was measured on an Aminco Bowman series 2 spectrofluorimeter with excitation and emission set at 560 and 590 nm, respectively, using appropriate blanks and expressed as percentage of normoxic controls.
ROS measurement by DCFDA fluorescence. Formation of ROS in HPAECs was determined by DCFDA fluorescence method (54). HPAECs (~80% confluent in 35-mm dishes) were loaded with 10 µM DCFDA for 30 min in complete MCDB medium at 37°C in a 95% air-5% CO2 environment. At the end of incubation, the medium containing DCFDA was aspirated, cells were washed once with complete MCDB medium, 1.0 ml of complete MCDB medium was added, and, if necessary, cells were preincubated with test compounds for indicated time followed by exposure to normoxia (95% air-5% CO2) and hyperoxia (95% O2-5% CO2) for desired lengths of time. At the end of exposure to normoxia and hyperoxia, the dishes containing cells were placed on ice, cells were scraped, and the medium containing cells was transferred to 1.5-ml microfuge tubes and centrifuged at 8,000 g for 10 min at 4°C. The medium was aspirated, and the cell pellet was washed twice with ice-cold PBS and sonicated on ice with a probe sonicator at a setting of 5 for 15 s in 500 µl of ice-cold PBS to prepare cell lysates. Fluorescence of oxidized DCFDA in cell lysates, an index of formation of ROS, was measured on an Aminco Bowman series 2 spectrofluorimeter with excitation and emission set at 490 and 530 nm, respectively, using appropriate blanks. The extent of ROS formation was expressed as a percentage of normoxic control.
ROS detection in cells by fluorescence microscopy. Hyperoxia-induced ROS formation in cells was detected by fluorescence microscopy. HPAECs (~80% confluent) in 35-mm dishes were loaded with DCFDA (10 µM) for 30 min in complete MCDB medium at 37°C in a 95% air-5% CO2 environment. At the end of incubation, the medium containing DCFDA was aspirated, cells were washed once with complete MCDB medium, 1.0 ml of complete MCDB medium was added, and if necessary, cells were preincubated with test compounds for the desired amount of time followed by exposure to normoxia (95% air-5% CO2) and hyperoxia (95% O2-5% CO2) for required lengths of time. At the end of incubation, cells were washed two times with PBS and examined under a Nikon Eclipse TE 300 microscope with excitation and emission set at 490 and 530 nm, respectively. Fluorescence of oxidized DCFDA in cells was captured with Sony digital DKC 5000 camera.
Electron paramagnetic resonance spectroscopy and spin trapping. HPAECs (~80% confluent) in 35-mm dishes were exposed, in the presence of 50 mM DMPO in complete MCDB medium, to normoxia (95% air-5% CO2) and hyperoxia (95% O2-5% CO2) at 37°C for required lengths of time. Wherever necessary, pharmacological agents such as DPI (100 µM) or SOD (100 µg) were included in the medium throughout the experimental exposure. Electron paramagnetic resonance (EPR) spectra were recorded in flat cells at room temperature with a Bruker ESP 300 spectrometer operating at X-band using the following parameters: microwave power, 20 mW; modulation amplitude, 0.5 f; modulation frequency, 100 kHz; receiver gain, 5 × 105; and the samples were loaded into the TM-110 cavity (60).
Localization of NADPH oxidase subcomponents by immunofluorescence microscopy. Immunohistochemical localization of NAD(P)H oxidase subcomponents (p47 phox and gp 91) in HPAECs was done using antibodies against p47 phox and gp 91 subunits for immunohistochemical staining, followed by fluorescence microscopic examination. HPAECs were grown on coverslips to ~80% confluence in an atmosphere of 95% air-5% CO2 and washed with 2 ml of prewarmed PBS (37°C). The coverslips were immediately treated with fixation-permeabilization solution (0.5% Triton X-100 and 3% paraformaldehyde) for 2 min immediately followed by treatment with 3% paraformaldehyde for 20 min at room temperature. The coverslips were washed three times with PBS for 5 min and incubated with blocking buffer [1% BSA in Tris-buffered saline containing 1% Tween (TBS-T)] for 30 min at room temperature. The coverslips were incubated with primary (p47 phox and gp 91) antibodies (1:400 dilution in blocking buffer) for 1 h at room temperature. The coverslips were gently washed three times with TBS-T for 5 min and incubated with AlexA Fluo 488 anti-goat antibody (1:200 dilution) for 1 h at room temperature followed by washing with TBS-T. Mounting medium (Kirkegaard & Perry Laboratories, Gaithersburg, MD) was applied, and coverslips were sealed. The cells were examined for fluorescence on Nikon inverted fluorescence microscope ECLIPSE TE300.
Determination of NADPH oxidase activity by chemiluminescence assay. NAD(P)H oxidase activity in intact cells was assayed by lucigenin chemiluminescence assay (18). HPAECs (~80% confluent) in 35-mm dishes, after exposure to normoxia (95% air-5% CO2) and hyperoxia (95% O2-5% CO2) for required lengths of time, were gently scraped and centrifuged at 400 g for 10 min at 4°C. The cell pellet was resuspended in a known volume (250 µl) of ice-cold phenol red-free medium (medium 199), and the cell suspension was kept on ice. To a final 1-ml volume of prewarmed (37°C) phenol red-free medium (medium 199) containing either NADPH (1 µM-100 µM) or lucigenin (20 µM), 50 µl of cell suspension (0.1 × 106 cells) were added to initiate the reaction followed by immediate measurement of chemiluminescence in a Packard scintillation counter in out-of-coincidence mode. Wherever necessary, pharmacological agents such as DPI (100 µM) or SOD (100 µg) were also included in the incubation mixture. Appropriate blanks and controls were established, and chemiluminescence was recorded. Neither NADPH nor NADH enhanced the background chemiluminescence of lucigenin alone (30-40 counts per min). Chemiluminescence was measured continuously for 12 min, and the activity of NAD(P)H oxidase was expressed as counts per million per cells.
Preparation of cell lysates and Western blotting. Preparation of cell lysates and Western blotting of proteins were performed as described earlier (41). After the HPAECs were exposed to normoxia (95% air-5% CO2) and hyperoxia (95% O2-5% CO2) in complete MCDB medium for required lengths of time, cells were rinsed twice with ice-cold PBS, scraped in 1.0 ml of lysis buffer containing 20 mM Tris · HCl (pH, 7.4), 150 mM NaCl, 2 mM EGTA, 5 mM glycerophosphate, 1 mM MgCl2, 1% Triton X-100, 1 mM sodium orthovanadate, 10 µg/ml protease inhibitors, 1 µg/ml aprotinin, 1 µg/ml leupeptin, and 1 µg/ml pepstatin, incubated at 4°C for 20 min, sonicated three times for 15 s each on ice with a probe sonicator, and cleared by centrifugation in a microfuge at 10,000 g for 5 min at 4°C. After determination of the total protein in the lysates by the bicinchoninic acid method, 6× Laemmli sample buffer was added to cell lysates to give a protein concentration of 1 µg/ml, and the lysates were boiled for 5 min. Proteins were separated on 10 or 12% gels by SDS-PAGE, transferred to polyvinylidene difluoride membranes, and subjected to immunoblotting with anti-p38 MAPK (1:1,000 dilution), anti-ERK1/ERK2 (1:1,000 dilution), anti-JNK1/JNK2 (1:1,000 dilution), anti-gp91 (1:1,000 dilution), anti-phox 67 (1:500 dilution), anti-phox 47 (1:500 dilution), and anti-phox 22 (1:500 dilution) overnight at 4°C. For detection of phosphorylated MAPKs, immunoblotting was performed with phospho-specific anti-p38 MAPK (1:1,000 dilution), anti-ERK1/ERK2 (1:1,000 dilution), and anti-JNK1/JNK2 (1:1,000 dilution). The membranes were washed three times with TBS-T and were incubated for 2-4 h at room temperature in horseradish peroxidase-conjugated goat anti-mouse or goat anti-rabbit secondary antibodies (1:2,000-1:5,000 dilution in TBS-T containing either 5% BSA or nonfat milk). The immunoblots were developed with enhanced chemiluminescence according to the manufacturer's recommendation. Intensities of the visualized protein bands were determined using a computerized densitometric method (Un-Scan-It gel, Automated Digitizing System, version 5.1).
Infection of HPAECs with p38 MAPK dominant-negative adenoviral constructs. HPAECs were cultured in 35-mm dishes until they reached ~80% confluence. Each dish was infected with either vector or p38 dominant-negative p38 MAPK adenovirus constructs [50 plaque-forming units (PFU)/cell; ~3 × 105 cells/dish]. After 24 and 48 h of infection, the medium was replaced with fresh complete MCDB medium, and the cells were exposed to hyperoxia and analyzed for expression of p38 MAPK by Western blotting and generation of ROS as described earlier. HPAECs infected with p38 MAPK dominant-negative or p38 wild-type or vector controls were exposed to normoxia and hyperoxia (3 h) followed by determination of intracellular ROS by the DCFDA oxidation method.
Statistical analysis. Statistical analysis was carried out by ANOVA using SigmaStat (Jandel). The level of statistical significance was taken as P < 0.05.
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RESULTS |
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Hyperoxia induces generation of
O
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Hyperoxia induces formation of H2O2 in
HPAECs.
HPAECs exposed to hyperoxia (3 h) compared with the cells exposed to
normoxia (3 h) showed significant accumulation of extracellular H2O2 (150% of normoxic control) in the medium
as measured by Amplex red fluorescence, which was completely attenuated
by treatment with DPI (100 µM) (Fig.
5). These results indicate that hyperoxia induced the formation of H2O2 in the medium
from dismutation of O
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DPI, but not rotenone or oxypurinol, inhibits hyperoxia-induced
formation of ROS in HPAECs.
Experiments were conducted utilizing the DCFDA fluorescence method to
identify the source of generation of ROS in HPAECs exposed to
hyperoxia. Metabolic blockers such as DPI, rotenone, and oxypurinol were used as inhibitors of flavin-dependent enzyme such as NAD(P)H oxidase, mitochondrial electron transport chain, and xanthine oxidase,
respectively. Hyperoxia (12 h)-induced formation of intracellular ROS
in HPAECs was enhanced (170% of normoxic control) and was significantly inhibited by DPI (100 µM) (125% of normoxic control), but not by rotenone (100 µM) or oxypurinol (100 µM) (Fig.
6). These results further demonstrate a
role of NAD(P)H oxidase, but not of either mitochondrial electron
transport chain or xanthine oxidase, in hyperoxia-induced formation of
intracellular ROS in HPAECs.
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Hyperoxia stimulates NADPH oxidase activity in HPAECs.
The NADPH- and NADH-dependent formation of O
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HPAECs contain phagocytic NADPH oxidase subcomponents.
The presence of phagocytic NADPH subcomponents in HPAECs was examined
by immunofluorescence microscopy and Western blotting. HPAECs exhibited
intense and moderate fluorescence due to reactivities to human
neutrophil p47 phox and gp 91 antibodies, indicating the presence of
p47 phox and gp 91 in HPAECs (Fig.
8A). SDS-PAGE and Western
blotting analysis of lysates of HPAECs revealed the presence of gp 91, p67 phox, p47 phox, and p22 phox subcomponents, similar to the NADPH
subcomponents of human neutrophils (Fig. 8B). The only
phagocytic subcomponent of NADPH oxidase not detectable by SDS-PAGE and
Western blotting analysis in HPAECs was p40 phox (data not shown).
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p22 phox antisense blocks hyperoxia-induced formation of ROS in
HPAECs.
We measured the role of NAD(P)H oxidase in hyperoxia-induced formation
of intracellular ROS by DCFDA fluorescence in HPAECs by exposing cells
to hyperoxia after overnight transfection with p22 phox antisense
plasmid. The intracellular generation of ROS in HPAECs transfected with
vector control was significantly higher in hyperoxia (3 h) (158% of
normoxic vector control), whereas in the cells transfected with p22
phox antisense, the generation of ROS was significantly lower after
both normoxia (3 h) and hyperoxia (3 h) (48% of normoxic vector
control and 60% of hyperoxic vector control) (Fig.
9A). Figure 9B
shows diminished expression of p22 phox subcomponent of NAD(P)H oxidase
in cells transfected with p22 phox antisense for 12 h as revealed
by Western blotting analysis. These results suggest a role for p22 phox
in activation of NAD(P)H oxidase and formation of intracellular ROS in
hyperoxia.
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Hyperoxia activates MAPKs in HPAECs.
The activation of MAPKs in HPAECs upon exposure to hyperoxia (3 h) was
studied by SDS-PAGE and Western blotting analysis. As shown in Fig.
10, the extent of the phosphorylation
of ERK and p38 MAPK in cells exposed to hyperoxia (3 h) was markedly
higher than that in the cells exposed to normoxia (3 h). No such
increase in the phosphorylation of JNK was noticed in the cells exposed to hyperoxia (Fig. 10). These results demonstrate that hyperoxia activates both ERK and p38 MAPK in HPAECs.
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MAPK inhibitors attenuate hyperoxia-induced ROS formation in
HPAECs.
We studied the role of MAPKs in hyperoxia-induced formation of
intracellular ROS by exposing HPAECs to hyperoxia (3 h) in the absence
or presence of various MAPK-specific inhibitors and measuring DCFDA
fluorescence. Hyperoxia induced a significant increase in the
intracellular generation of ROS (270% of the normoxic control, Fig.
11A), which was
significantly attenuated by the p38 MAPK inhibitors SB-202190 (20 µM)
and SB-203580 (20 µM). The MEK1/2 inhibitor PD-98059 (20 µM),
although less effective than the p38 MAPK-specific inhibitors, also
significantly attenuated the hyperoxia-induced intracellular generation
of ROS in HPAECs (190% of the normoxic control, Fig. 11B).
These results show a role for both ERK and p38 MAPK in
hyperoxia-induced ROS generation in HPAECs.
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Infection of HPAECs with p38 MAPK dominant-negative adenoviral
constructs attenuates hyperoxia-induced generation of ROS.
To further establish the role of p38 MAPK in hyperoxia-induced
generation of ROS, we exposed HPAECs infected with vector control and
dominant-negative p38 MAPK adenoviral constructs to normoxia (3 h) and
hyperoxia (3 h), and the extent of intracellular oxidation of DCFDA was
determined. As shown in Fig.
12A, hyperoxia-induced generation of ROS was significantly attenuated in cells infected with
dominant-negative p38 MAPK adenoviral constructs (125% of normoxic
vector controls) compared with vector control infected cells (275% of
normoxic vector controls). Correspondingly, the expression of p38 MAPK
protein in cells infected with dominant-negative constructs was
markedly enhanced, as revealed by Western blotting of cell lysates
(Fig. 12B). These results further support the role of p38
MAPK in hyperoxia-induced generation of ROS in HPAECs.
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DISCUSSION |
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In the present study, we employed various established techniques
(5) to determine the generation/release of
O
Several studies have revealed the activities of both NADH and NADPH
oxidase(s) in vascular ECs (37, 49). Our results also demonstrate that HPAECs contain an NADH- and NADPH-dependent oxidase system that activates molecular oxygen to O
Several nonphagocytic cells such as the vascular smooth muscle and ECs have been shown to contain many of the subcomponents of the phagocytic NADPH oxidase complex (3, 6, 23, 26). Although the exact role in EC physiology and function and the mechanism of activation of vascular EC NAD(P)H oxidase are yet to be understood thoroughly, it is clear from our results that HPAECs possess most of the subcomponents of human neutrophil NADPH oxidase. The only phagocytic subcomponent of NADPH oxidase not detectable by SDS-PAGE and Western blotting analysis in HPAECs was p40 phox. Our results revealed that transfection with p22 phox antisense attenuated hyperoxia-induced generation of ROS in HPAECs (Fig. 8), further supporting the role of p22 phox subcomponent in hyperoxia-induced activation of NAD(P)H oxidase. From studies on growth-factor responsive human airway smooth muscle cells, the regulation of NAD(P)H oxidase by the p22 phox subcomponent and its role in the generation of ROS and proliferation of cells have been established (7), which also support our results on the involvement of p22 phox in hyperoxia-induced generation of ROS in HPAECs.
MAPKs are stress-activated kinases that participate in various cellular
signaling events (2). MAPKs are divided into three broad
categories as 1) ERK, 2) JNK, and 3)
p38 MAPK (51). It has been shown that several stresses
such as chemical toxicity, hypoxia, and mechanical stretch activate
MAPKs and MAPK-dependent signaling events in many cell systems
(42). Our results indicate that hyperoxia (3 h)-induced
generation of intracellular ROS generation was significantly inhibited
by both the p38 MAPK- and ERK-specific inhibitors (Fig. 11,
A and B) and also suggest that the activation of
p38 MAPK and ERK is upstream of the generation of ROS under hyperoxic
exposure in HPAECs. In vascular smooth muscle cells, it was shown that
angiotensin II activated MAPKs (ERK, p38 MAPK, and JNK) in association
with NAD(P)H oxidase-mediated generation of ROS (53).
Studies on hepatitis C virus nonstructural protein 3-induced oxidative
burst in human monocytes show a role of p38 MAPK in NADPH oxidase
activation and generation of ROS (10). In bovine
polymorphonuclear leukocytes, p38 MAPK was identified to participate in
NADPH oxidase activation, production of O
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ACKNOWLEDGEMENTS |
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We acknowledge Donghong He, Jianbin Yang, and Tonya Watkins for excellent technical assistance. We are thankful to Dr. William Gerthoffer for providing the p38 MAPK dominant-negative adenoviral construct and Dr. Michael Crow of the National Institute of Aging for providing the p22 phox antisense plasmid.
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FOOTNOTES |
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This work was supported by National Heart, Lung, and Blood Institute Grant HL-69909 (to V. Natarajan).
Present address for N. L. Parinandi, J. L. Zweier, and A. J. Cardounel: Dorothy M. Davis Heart and Lung Research Institute, College of Medicine and Public Health, Ohio State Univ., 473 W. 12th Ave., Columbus, OH 43210.
Address for reprint requests and other correspondence: V. Natarajan, Div. of Pulmonary and Critical Care Medicine, Dept. of Medicine, Johns Hopkins Univ. School of Medicine, Asthma & Allergy Center, 5501 Hopkins Bayview Cir., Baltimore, MD 21224 (E-mail: vnataraj{at}jhmi.edu).
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.
July 26, 2002;10.1152/ajplung.00123.2002
Received 25 April 2002; accepted in final form 19 July 2002.
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