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VASCULAR BIOLOGY

Pollutant particles enhanced H₂O₂ production from NAD(P)H oxidase and mitochondria in human pulmonary artery endothelial cells

¹Center for Environmental Medicine, Asthma and Lung Biology, University of North Carolina, Chapel Hill; and ²National Health and Environmental Effects Research Laboratory, Office of Research and Development, Environmental Protection Agency, Research Triangle Park, North Carolina

Submitted 20 July 2005 ; accepted in final form 24 March 2006

	ABSTRACT

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Particulate matter (PM) induces oxidative stress and cardiovascular adverse health effects, but the mechanistic link between the two is unclear. We hypothesized that PM enhanced oxidative stress in vascular endothelial cells and investigated the enzymatic sources of reactive oxygen species and their effects on mitogen-activated protein kinase (MAPK) activation and vasoconstriction. We measured the production of extracellular H₂O₂, activation of extracellular signal-regulated kinases1/2 (ERK1/2) and p38 MAPKs in human pulmonary artery endothelial cells (HPAEC) treated with urban particles (UP; SRM1648), and assessed the effects of H₂O₂ on vasoconstriction in pulmonary artery ring and isolated perfused lung. Within minutes after UP treatment, HPAEC increased H₂O₂ production that could be inhibited by diphenyleneiodonium (DPI), apocynin (APO), and sodium azide (NaN₃). The water-soluble fraction of UP as well as its two transition metal components, Cu and V, also stimulated H₂O₂ production. NaN₃ inhibited H₂O₂ production stimulated by Cu and V, whereas DPI and APO inhibited only Cu-stimulated H₂O₂ production. Inhibitors of other H₂O₂-producing enzymes, including N-methyl-L-argnine, indomethacin, allopurinol, cimetidine, rotenone, and antimycin, had no effects. DPI but not NaN₃ attenuated UP-induced pulmonary vasoconstriction and phosphorylation of ERK1/2 and p38 MAPKs. Knockdown of p47phox gene expression by small interfering RNA attenuated UP-induced H₂O₂ production and phosphorylation of ERK1/2 and p38 MAPKs. Intravascular administration of H₂O₂ generated by glucose oxidase increased pulmonary artery pressure. We conclude that UP induce oxidative stress in vascular endothelial cells by activating NAD(P)H oxidase and the mitochondria. The endothelial oxidative stress may be an important mechanism for PM-induced acute cardiovascular health effects.

mitogen-activated protein kinase; extracellular signal-regulated kinase; p38; vasoconstriction

Recent in vivo and in vitro evidence indicates that PM may also cause endothelial dysfunction and vasoconstriction. Exposure to concentrated ambient particles (CAPs) (median, 182.75 µg/m³) for 5 h/day for 3 days decreased the lumen/wall area ratio of small pulmonary arteries (PA) in rats indicating increased pulmonary vascular resistance (4). PM from motorcycle exhaust enhanced constriction of rat aortic rings induced by phenylephrine (45). Exposure to PM for 4 wk increased atherosclerotic plaque formation in rabbits (42). Co-exposure to CAPs (150 µg/m³) and ozone (120 ppb) for 2 h causes acute constriction of brachial artery in healthy human adults (9). An air pollution episode in Germany was associated with increases in systemic blood pressure by as much as 8 mmHg (26). Various vasoconstrictor mechanisms are affected by PM, including the release of endothelins (8, 44), activation of the epithelial growth factor receptor (25), and inhibition of nitric oxide (NO) production (2, 8, 25, 27). More recently, we demonstrated that PM caused pulmonary vasoconstriction and activated mitogen-activated protein kinase (MAPK) via activation of angiotensin II receptor subtype 1 (AT1R) (32).

PM also is known to produce oxidative stress in lung cells (17). Chapel Hill particles increased the production of reactive oxygen species (ROS) in human alveolar macrophages and bronchial epithelial cells (6). Diesel particles induced oxidative stress in macrophages (23). PM-induced secretion of amphiregulin was blocked by the antioxidant N-acetylcysteine (7). Ottawa ambient particles (EHC-93) induced blood oxidative stress markers in rats (46). The oxidative stress in these lung cells have been linked mostly to PM-induced pulmonary inflammation. Because the cardiovascular adverse effects induced by PM may occur without lung inflammation, we reasoned that PM may induce oxidative stress directly in vascular endothelial cells. The induction of endothelial oxidative stress is an important mechanism linking many known cardiovascular risk factors, including hyperglycemia, dyslipidemia, hypertension, and cigarette smoking (11), with the development of vascular diseases. To test this hypothesis, we exposed PA endothelial cells to PM and measured extracellular H₂O₂ production. Our goals were to determine the enzymatic sources of ROS production in vascular endothelial cells and to investigate the effects of ROS from different sources on MAPK activation and vasoconstriction, two cellular events known to occur with PM exposure (32). The study was performed in human PA endothelial cells (HPAECs), isolated PA ring, and isolated buffer-perfused lungs.

	MATERIALS AND METHODS

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Reagents and chemicals. HPAECs were obtained from Cell Applications (San Diego, CA). Endothelial growth medium (EGM-2) and supplements were from Clonetics (BioWhittaker, Walkersville, MD). Diphenyleneiodonium (DPI), apocynin, indomethacin, cimetidine, rotenone, allopurinol, antimycin, sodium azide and N-methyl-L-arginine (L-NAME) were obtained from Sigma (St. Louis, MO). Monoclonal antibodies against phospho-p38, total p38, phospho-ERK1/2, and total ERK1/2 were purchased from Cell Signaling Technology (Beverly, MA). Horseradish peroxidase (HRP)-conjugated goat anti-rabbit and goat anti-mouse IgG were obtained from Santa Cruz Biotechnology (Santa Cruz, CA). Molecular mass standards, polyacrylamide, and buffers were from Bio-Rad (Richmond, CA). The ECL Western blot analysis detection reagents were purchased from Amersham Biosciences (Piscataway, NJ).

Source of particles. Urban particles (SRM 1648) were purchased from the National Institute of Standards and Technology (Gaithersburg, MD). They were prepared from urban PM collected in the St. Louis, MO, area in a baghouse over a period of >12 mo. The material was removed from the filter bags, combined in a single lot, screened through a fine mesh sieve to remove extraneous materials and thoroughly blended in a v-blender. The material was then packaged into sequentially numbered bottles. The major constituent elements are (mass fraction in %) the following: 3.4% Al, 3.9% Fe, 1.1% K⁺, and the minor constituent elements are 0.43% Na, 0.66% Pb, and 0.48% Zn. There are also metal constituents (in ng/mg), including 115 As, 76 Cd, 403 Cr, 609 Cu, 786 Mn, 82 Ni, 27 Se, 5.5 U, and 127 V. UP was suspended in distilled deionized water for the use in experiments.

Isolated rat PA ring model. Segments of the right and left main PAs of Sprague-Dawley rats (250–350 g) measuring 2–3 mm were removed and placed in the Krebs-Henseleit buffer, as described previously (31). The artery segments were then suspended in the Radnoti 4-unit tissue organ bath system (Glass Technology, Monrovia, CA). The reservoirs held 20 ml of Krebs-Henseleit buffer and were bubbled constantly with 21% O₂ + 5% CO₂ gas. After a 10- to 15-min stabilization period, the baseline tension of the rings was adjusted to 1 g before all experiments. The artery rings were then exposed to increasing doses of particles from 1 to 100 µg/ml. Inhibitors were given 15 min before particle exposure. The rings were washed with buffer between the two doses of particles. The maximum tension within 5 min after each dose of particles was recorded.

Cultured HPAECs. HPAECs were grown in EGM-2 supplemented with 2% fetal bovine serum, 0.04% hydrocortisone, 0.4% human fibroblast growth factor-B, 0.1% vascular endothelial growth factor, 0.1% R³-insulin growth factor-1, 0.1% ascorbic acid, 0.1% human epithelial growth factor, 0.1% GA-1000, and 0.1% heparin. Cells at passages 4–9 grown to 80% confluence were used for the experiments.

Isolated perfused rabbit lung. The isolated buffer-perfused lung model using male New Zealand White rabbits (May's Farm, Wilson, NC) weighing 2.5–3.0 kg has been described previously (25). Briefly, the PA and left atrium were cannulated for monitoring mean PA pressure and left atrial pressure, respectively. The left atrial cannula was connected by Tygon tubing to a reservoir suspended freely from a force displacement transducer (model FT100, Grass Instrument, Quincy, MA) for monitoring weight gain of the lung. The reservoir was placed at the lowest portion of the lung to maintain a left atrial pressure of zero. Tubing connecting the reservoir with the main PA cannula completed the perfusion circuit. The perfusion circuit also consisted of a roller pump (Sarns, Ann Arbor, MI), a bubble trap, and a heat exchanger. The volume of the system was 250 ml. Trachea was also cannulated for monitoring tracheal pressure. The lung was ventilated with 21% O₂ + 5% CO₂ through the tracheostomy using an animal respirator (Harvard Apparatus, S. Natick, MA) delivering 30 breaths per min at 2–3 cmH₂O positive end-expiratory pressure. The tidal volume was adjusted to achieve a peak tracheal pressure of 7–10 mmHg (20 ml). All pressures and weight gain were transmitted to a four-channel analog-to-digital converter and amplifier connected to a personal computer equipped with data-acquisition software (DATAQ Instruments, Akron, OH). The Krebs-Henseleit-3% albumin buffer contains 82.8 mM NaCl, 4.7 mM KCl, 2.4 mM monobasic potassium phosphate, 25 mM sodium bicarbonate, 1.2 mM magnesium sulfate, 2.7 mM CaCl, 11.1 mM dextrose, and 3% wt/vol bovine serum albumin, fraction V, at a pH of 7.3–7.4. After the pulmonary circulation was washed free of blood with the buffer, a recirculating perfused lung system with a flow of 100 ml/min was established. The lung was then allowed 10–15 min for stabilization. Lungs with visible leaks and/or high PA pressure (>20 mmHg) during this period were excluded.

Western blot analysis. After exposure, the cells were washed once with ice cold phosphate-buffered saline (PBS) and then lysed with RIPA buffer (1% Nonidet P-40, 0.5% sodium deoxycholate, and 0.1% SDS in PBS, pH 7.4) containing 0.1 mM vanadyl sulfate and protease inhibitors (0.5 mg/ml aprotinin, 0.5 mg/ml E-64, 0.5 mg/ml pepstatin, 0.5 mg/ml bestatin, 10 mg/ml chymostatin, and 0.1 ng/ml leupeptin). The cell lysates were then centrifuged at 3,000 g for 10 min at 4°C. Protein concentration of supernatant was measured with Bio-Rad protein assay reagent. Cellular proteins were separated by 10% SDS-PAGE and transferred to a polyvinylidene difluoride membrane. The blot was blocked with 5% milk in PBS with 0.05% Tween 20 for 1 h at room temperature, washed briefly, and then probed with goat antibodies against human phospho-p38 or phospho-ERK1/2 overnight at 4°C. This was followed by incubation with HRP-conjugated secondary antibodies. Bands were detected with the use of ECL and films. The blot was then stripped and reprobed with monoclonal antibodies against human p38 or ERK1/2 and appropriate HRP-conjugated secondary antibodies.

Measurements of H₂O₂. Cells were cultured in 24-well plates. Production of extracellular H₂O₂ was measured by the Amplex red reagent (10-acetyl-3,7-dihydroxyphenoxazine; Molecular Probes-Invitrogen, Carlsbad, CA). Amplex red reacts with H₂O₂ to produce highly fluorescent resorufin in the presence of HRP. The fluorescent signals were measured over 120 min with a Bioassay HTS7000 plate reader (Perkin Elmer, Wellesley, MA) with HTSoft version 1.0 software (PE Applied Biosystems, Weiterstadt, Germany). The excitation and emission wavelengths were 530 and 590 nm, respectively. Generation of H₂O₂ was calculated by subtracting the fluorescence signal at 0 min (baseline) from that at 120 min. Positive controls using H₂O₂ (0.5–2 µM) were included for all Amplex red assays. All inhibitors were added 20 min before Amplex red.

Transfection of HPAEC with p47^phox siRNA. Cells were grown to 60–70% confluency in 6-well plates. They were then transfected with Gene Silencer transfecting agent plus (Gene Therapy System, San Diego, CA) with p47^phox siRNA (100 nM; Dharmacon, Chicago, IL) in serum-free EGM-2 medium for 3 h according to the manufacturer's recommendation. Control cells were incubated with Gene Silencer without p47^phox siRNA. Fresh EGM-2 medium with 2% fetal bovine serum was then added, and cells were cultured for an additional 48 h. Cell lysates from some wells were collected for p47^phox protein expression and phospho-ERK1/2 and phospho-p38 MAPKs by Western blot analysis. Other wells were used for measuring H₂O₂ production.

Statistical analysis. Data shown in text and figures are means ± SE. Data from the artery ring experiments and perfused rabbit lungs were analyzed by the repeated-measures ANOVA. Data from the cell experiments were analyzed by ANOVA, followed by Scheffé's test for post hoc comparisons. The statistical analysis was performed using commercially available software (Statview version 5.0.1, SAS Institute, Cary, NC). A P value of <0.05 was taken as statistically significant.

	RESULTS

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Effects of UP on H₂O₂ release. We first measured extracellular release of H₂O₂ using the Amplex red reagent. UP at up to 100 µg/ml did not produce cytotoxicity in 20 min as evidenced by the lack of lactate dehydrogenase release. UP increased H₂O₂ production in a time- and dose-dependent manner (Fig. 1). The increase was statistically significant as early as 5 min (the first time point) after incubation. H₂O₂ released over 120 min by cells treated with 100 µg/ml of UP was 0.80 ± 0.02 µM or 230% that by the control cells. The enhanced Amplex red signal induced by UP was completely abolished by the addition of catalase to the medium (changes in the Amplex red signal = 7,319 ± 210).

View larger version (14K): [in this window] [in a new window]   Fig. 1. Production of H2O2 by pulmonary artery endothelial cells stimulated by urban particles (UP). A: the cells were incubated with UP (10 µg/ml) for up to 120 min. B: the cells were incubated with UP (1–100 µg/ml) for 120 min. 10-Acetyl-3,7-dihydroxyphenoxazine (Amplex red; 50 µM) was given 20 min before UP, and the fluorescence signals were read with a fluorescence plate reader. *P < 0.05 vs. 0 µg/ml UP; n = 4 independent experiments performed in triplicate.

View larger version (13K): [in this window] [in a new window]   Fig. 2. Inhibition of UP-stimulated H2O2 production by dophenyleneiodonium (DPI; A), apocynin (APO; B), and sodium azide (NaN3; C). Quiescent cells were pretreated with 4 µM DPI, 10 µg/ml APO, or 3% NaN3 for 20 min and incubated with 10 µg/ml UP for 120 min. H2O2 production was measured by Amplex red; n = 4 independent experiments performed in triplicate.

View larger version (18K): [in this window] [in a new window]   Fig. 3. Effects of Cu on H2O2 production. A: the cells were incubated with various concentrations of CuSO4 for 2 h. B: cells were incubated with CuSO4 (50 µM) for 2 h. Amplex red (50 µM) was given 20 min before CuSO4, and the fluorescence signals were read with a fluorescent plate reader. The CuSO4-induced H2O2 production was inhibited by 4 µM DPI (C), 10 µg/ml APO (D), and 0.3% NaN3 (E). *P < 0.05 vs. time 0; n = 6–10 independent experiments performed in triplicate.

View larger version (18K): [in this window] [in a new window]   Fig. 4. Effects of V on H2O2 production. A: cells were incubated with various concentrations of vanadyl sulfate (VOSO4) for 2 h. B: cells were incubated with 50 µM VOSO4 for 2 h. B: amplex red (50 µM) was given 20 min before VOSO4, and the fluorescence signals were read with a fluorescent plate reader. The VOSO4-induced H2O2 production was not inhibited by 4 µM DPI (C) or 10 µg/ml APO (D) but was inhibited by 0.3% NaN3 (E). *P < 0.05 vs. time 0; n = 6–10 independent experiments performed in triplicate.

View larger version (13K): [in this window] [in a new window]   Fig. 5. Effects of 4 µM DPI (A) and 0.3% NaN3 (B) on UP-induced pulmonary vasoconstriction. The pulmonary arterial ring was pretreated with DPI or NaN3 for 15 min before UP administration (1–100 µg/ml). The constriction was measured by ring tension and expressed as % maximum tension induced by 1 µM phenylephrine. *P < 0.05 vs. UP; n = 6 rings from different animals. C: changes in pulmonary artery pressure (PAP) induced by glucose/glucose oxidase, which generates H2O2. D-Glucose (1 mM) and glucose oxidase (10 U/ml) were given as a bolus into the perfusate of isolated lungs at time 0. PAP was then monitored for 25 min. *P < 0.05 vs. time 0; n = 6.

View larger version (21K): [in this window] [in a new window]   Fig. 6. Effects of DPI on UP-induced phosphorylation of ERK1/2 (A) and p38 (B). Quiescent cells were pretreated with DPI (4 µM) for 30 min and incubated with 10 µg/ml UP for 10 min. Phosphorylation of ERK1/2 and p38 in cell lysates was measured by Western blot analysis. The blots were then stripped and probed for total ERK1/2 and p38. A representative Western blot and the densitometry results for ERK1/2 and p38 are shown. Data are means ± SE; n = 4 independent experiments.

View larger version (22K): [in this window] [in a new window]   Fig. 7. Effects of APO on UP-induced phosphorylation of ERK1/2 (A) and p38 (B). Quiescent cells were pretreated with APO (10 µg/ml) for 30 min and incubated with 10 µg/ml UP for 10 min. Phosphorylation of ERK1/2 and p38 in cell lysates was measured by Western blot analysis. The blots were then stripped and probed for total ERK1/2 and p38. A representative Western blot and the densitometry results for ERK1/2 and p38 are shown. Data are means ± SE; n = 4 independent experiments.

View larger version (23K): [in this window] [in a new window]   Fig. 8. Effects of NaN3 on UP-induced phosphorylation of ERK1/2 (A) and p38 (B). Quiescent cells were pretreated with NaN3 (0.3%) for 30 min and then incubated with 10 µg/ml UP for 10 min. Phosphorylation of ERK1/2 and p38 in cell lysates was measured by Western blot analysis. The blots were then stripped and probed for total ERK1/2 and p38. A representative Western blot and the densitometry results for ERK1/2 and p38 are shown. Data are means ± SE; n = 4 independent experiments.

View larger version (43K): [in this window] [in a new window]   Fig. 9. Effects of p47phox small interfering RNA (siRNA) on protein expression of p47phox (A), UP-induced H2O2 production (B), UP-induced phosphorylation of ERK1/2 (C), and UP-induced phosphorylation of p38 (D). Cells were grown to 60–70% confluency in 6-well plates. They were then transfected with GeneSilencer transfecting agent plus with p47phox siRNA (100 nM) in serum-free EGM-2 medium for 3 h. Control cells were incubated with GeneSilencer without p47phox siRNA. Fresh EGM-2 medium with 2% fetal bovine serum was then added and cells were cultured for an additional 48 h. Some wells were treated with UP (100 µg/ml) for 20 min (for phospho-ERK1/2 and phospho-p38) or for 2 h (for H2O2 measurement by Amplex red). A representative Western blot for p47phox after transfection is shown in A. A representative Western blot and the densitometry results for ERK1/2 and p38 are shown in C and D, respectively; n = 4 independent experiments.

	DISCUSSION

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H₂O₂ released by the UP-stimulated endothelial cells comes from two major enzymatic sources, the NAD(P)H oxidase and the mitochondrial electron transport chain, because the release of H₂O₂ was inhibited by two structurally distinct NAD(P)H oxidase inhibitors (DPI and apocynin), p47^phox siRNA and a mitochondrial complex IV inhibitor (NaN₃). Other enzyme systems that are potential sources for H₂O₂, including xanthine oxidase, cytochrome P-450, NO synthase, and cyclooxygenase-1, were less likely sources because their respective inhibitors, (allupurinol, cimetidine, L-NAME, and indomethacin) did not inhibit UP-induced H₂O₂ release. The NAD(P)H oxidase activity in vascular endothelial cells can be stimulated by many plasma membrane-mediated events, including cytokines, hormones, growth factors, G protein receptor agonists, and shear forces, and activation of NAD(P)H oxidase is known to be involved in the pathogenesis of vascular disease (15, 20, 21, 24, 34). Electron leaks from the mitochondrial electron transport chain of vascular endothelial cells, on the other hand, have been observed with different stimuli, such as hypoxia-reoxygenation and glucocorticoid excess (28, 43). Our results show that UP is a common stimulus for both systems because inhibitors for each system almost completely inhibit the UP-stimulated production of H₂O₂. It is not possible, however, to assess the relative contribution of each system in part because of the nonspecific effects of these inhibitors. Because DPI and apocynin inhibit UP-induced H₂O₂ production earlier than NaN₃, UP may activate NAD(P)H oxidase first. This hypothesis would be consistent with the cell membrane localization of NAD(P)H oxidase that makes it more accessible to extracellularly administered UP.

The exact components of UP responsible for activating NAD(P)H oxidase and the mitochondrial electron transport chain are not clear. The water-soluble fraction of UP was sufficient to stimulate the release of H₂O₂. Its transition metals, Cu and V, also enhanced H₂O₂ release. The patterns of inhibition by DPI, apocynin, and NaN₃ on Cu- and V-induced H₂O₂ production were similar to those on UP-induced H₂O₂ production. These results indicate that some components in the water-soluble fraction are responsible for changes induced by UP. Whether Cu and V are among the active components will require further studies in the future.

H₂O₂ produced by NAD(P)H oxidase contributed to PM-induced vasoconstriction because DPI inhibited UP-induced constriction of PA ring and intravascular administration of extracellular H₂O₂ produced by glucose and glucose oxidase produced acute vasoconstriction in perfused lungs. We and others have previously identified several signaling pathways that mediate PM-induced vasoconstriction, including epidermal growth factor receptor, angiotensin II receptor subtype 1, endothelins, and NO (8, 25, 27, 31, 32, 44). Our results indicate that H₂O₂ produced by the membrane NAD(P)H oxidase should also be added to the list. The cell membrane localization of NAD(P)H oxidase would allow H₂O₂ produced by this enzyme to be released directly into the extracellular space and exert its effects on the adjacent smooth muscle cells. The mitochondrial source of H₂O₂, on the other hand, appears not involved in regulating vascular reactivity because inhibition of H₂O₂ production by NaN₃ did not affect UP-induced vasoconstriction despite increases in ERK1/2 and p38 MAPK phosphorylation. We speculate that a subfamily of mitochondrial MAPKs may be affected by NaN₃ (1, 3). These mitochondrial MAPKs show a different response to ROS than the cytosolic MAPKs because H₂O₂ inhibits rather than increases phosphorylation of mitochondrial ERK1/2 (1). The primary function of the mitochondrial MAPKs appears different from that of the cytosolic MAPKs. Activation of the mitochondrial p38 MAPK mediates ceramide-induced apoptosis (30). Activation of mitochondrial ERK MAPK induces phosphorylation of a pro-apoptotic protein Bad preventing cell death in murine heart (3). Activation of the mitochondrial MAPKs subfamily and the specific role of mitochondrial MAPKs in cell death signaling may explain why NaN₃ did not affect UP-induced vasoconstriction despite an increase in phosphorylated ERK1/2 and p38 MAPKs. More studies in the future will be needed to test this hypothesis.

Extrapolation of our in vitro results to human exposure to ambient PM should be made with caution. The doses of UP to which endothelial cells were exposed were much higher than the one-time exposure in most ambient settings. With repeated exposures during severe pollution episodes (38, 40) and under certain occupational conditions (22, 47), the amount of PM inhaled into the lung may increase several hundred fold. Although the whole inhaled particles are unlikely in direct contact with vascular endothelial cells, water-soluble components contained in ambient PM can more readily permeate the alveolar-capillary barrier and gain access to the endothelial cells. Our study shows that PM can directly induce the production of H₂O₂ from NAD(P)H oxidase and the mitochondrial electron transport chain in vascular endothelial cells and that H₂O₂ produced from NAD(P)H oxidase mediates the PM-induced vasoconstriction. These results provided further evidence that PM may be an important risk factor for the development of cardiovascular diseases.

	GRANTS

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The study was supported in part by the cooperative agreement R829522-01 funded by the United States Environmental Protection Agency.

	DISCLOSURES

 
The research described in this article has been reviewed by the Health Effects and Environmental Research Laboratory, United States Environmental Protection Agency, and has been approved for publication.

	ACKNOWLEDGMENTS

 
The authors thank Drs. Martha Sue Carraway and Richard Whorton of Duke University Medical Center for advising the siRNA experiments.

A portion of the study was presented in the Annual Meeting of the Society of Free Radical Biology Medicine in St. Thomas, Virgin Islands (Free Rad Biol Med 37, Suppl 1: S142, 2004).

	FOOTNOTES

 

Address for reprint requests and other correspondence: Y.-C. T. Huang, CB 7315, 104 Mason Farm Rd., Chapel Hill, NC 27599 (e-mail: huang002{at}mc.duke.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.

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