Oxidant stress regulates basal endothelin-1 production by cultured rat pulmonary endothelial cells

John R. Michael1, Boaz A. Markewitz2, and Donald E. Kohan3

Divisions of 1 Respiratory, Critical Care, and Occupational Pulmonary Medicine and 3 Nephrology and Hypertension, Department of Medicine, Veterans Affairs Medical Center, University of Utah School of Medicine, Salt Lake City, Utah 84132; and 2 Section of Pulmonary and Critical Care Medicine, Department of Medicine, Overton Brooks Veterans Affairs Medical Center, Louisiana State University School of Medicine, Shreveport, Louisiana 71130

    ABSTRACT
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Abstract
Introduction
Materials & Methods
Results
Discussion
References

Endothelin-1 (ET-1) is a pluripotent mediator that modulates vascular tone and influences the inflammatory response. Patients with inflammatory lung disorders frequently have elevated circulating ET-1 levels. Because these pathophysiological conditions generate reactive oxygen species that can regulate gene expression, we investigated whether the level of oxidant stress influences ET-1 production in cultured rat pulmonary arterial endothelial cells (RPAEC). Treatment with the antioxidant 1,3-dimethyl-2-thiourea (10 mM) or the iron chelator deferoxamine (1.8 µM) doubles basal ET-1 release. Conversely, exposing cells to H2O2 generated by glucose and glucose oxidase (0.1-10 mU/ml) for 4 h causes a concentration-dependent decrease in ET-1 release. This effect occurs at concentrations of glucose oxidase that do not affect [3H]leucine incorporation or specific 51Cr release from RPAEC. Catalase prevents the decrease in ET-1 synthesis caused by glucose and glucose oxidase. Glucose and glucose oxidase decrease not only ET-1 generation but also ET-1 mRNA as assessed by semiquantitative polymerase chain reaction. Our results indicate that changes in oxidative stress can either up- or downregulate basal ET-1 generation by cultured pulmonary endothelial cells.

hydrogen peroxide; messenger ribonucleic acid; glucose oxidase; 1,3-dimethyl-2-thiourea; deferoxamine

    INTRODUCTION
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Abstract
Introduction
Materials & Methods
Results
Discussion
References

ENDOTHELIN-1 (ET-1) is a cytokine that can potently modulate pulmonary vascular tone, lung smooth muscle growth, and inflammatory processes. Proinflammatory effects of ET-1 include priming neutrophils, activating mast cells, stimulating oxygen radical production by macrophages, releasing growth factors from smooth muscle cells, and increasing adhesion molecule expression on endothelial cells (19). ET-1 also stimulates monocytes to produce interleukin-6, interleukin-8, and prostaglandin E2, all important in modulating immune responses (17). Besides the proinflammatory effects of ET-1, its ability to regulate vascular and airway tone, augment airway and vascular smooth muscle cell growth, and stimulate fibroblast growth, migration, and collagen synthesis indicate its potential to participate in the tissue reaction to ongoing inflammation (19).

Patients with a variety of inflammatory lung diseases, including the acute respiratory distress syndrome, have evidence for increased ET-1 production and reactive oxygen species generation in their lungs (7, 13, 21, 32). Because reactive oxygen species can regulate gene expression, we investigated the effect of oxidant stress on ET-1 production by cultured pulmonary endothelial cells.

    MATERIALS AND METHODS
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Abstract
Introduction
Materials & Methods
Results
Discussion
References

Reagents

We purchased phenol red-free (PRF) Dulbecco's modified Eagle's medium (DMEM), medium 199, liquid 10× Hanks' balanced salt solution without phenol red (HBSS), Moloney murine leukemia virus reverse transcriptase (RT), 5× first strand buffer, dithiothreitol, Taq DNA polymerase, and random primers from GIBCO Laboratories Life Technologies (Grand Island, NY); fetal bovine and bovine calf sera were from HyClone Laboratories (Logan, UT); tissue culture plastic ware was from Costar (Cambridge, MA); [51Cr]sodium chromate (51Cr), [3H]leucine, and the guanosine 3',5'-cyclic monophosphate (cGMP) 125I assay system were from Amersham Life Science (Arlington Heights, IL); deferoxamine mesylate was from Ciba Pharmaceutical (Summit, NJ); 1,3-dimethyl-2-thiourea (DMTU) was from Aldrich Chemical (Milwaukee, WI); bovine liver thymol-free catalase and glucose oxidase type II-S derived from Aspergillus niger were from Sigma Chemical (St. Louis, MO); ET-1 radioimmunoassay (RIA) kits were from Peninsula Laboratories (Belmont, CA); bicinchoninic acid (BCA) protein assay reagents were from Pierce Chemical (Rockford, IL); dNTP set solution was from Pharmacia LKB Biotechnology (Alameda, CA); formamide and random pd(N)6 were from Boehringer Mannheim (Indianapolis, IN); deoxynucleotide triphosphates, Wizard polymerase chain reaction (PCR) preps DNA purification system, and RNasin ribonuclease (RNase) inhibitor were from Promega (Madison, WI); [alpha -32P]GTP was from ICN Biomedicals (Costa Mesa, CA); 2-mercaptoethanol was from Bio-Rad Laboratories (Richmond, CA); phenol saturated solution was from Amresco (Solon, OH); chloroform and calcium chloride were from Mallinckrodt (Paris, KY); ethyl alcohol was from Quantum Chemical (Tuscola, IL); and RNase-free presiliconized microcentrifuge tubes were from Intermountain Scientific (Bountiful, UT). All other reagents and chemicals were purchased from Sigma Chemical unless otherwise stated.

Cell Culture

Rat pulmonary artery endothelial cells (RPAEC) were initially isolated with microcarrier beads as previously described and were generously provided by Dr. Una Ryan (28). The isolated cells have a cobblestone morphology by light and electron microscopy and have been identified as endothelial cells by the presence of factor VIII antigen, by the expression of angiotensin-converting enzyme activity, and by the uptake of acetylated low-density lipoproteins. The cells were maintained in monolayer culture at 37°C and 5% CO2, using Ryan's red medium (medium 199, 6.7% bovine calf serum, 3.3% fetal calf serum, 10-5 M thymidine, 1.3 mM L-glutamine, 60 U/ml penicillin, 60 µg/ml streptomycin, and 20 µg/ml gentamicin). The cells were passed without the use of enzymes and were plated onto 6- or 24-well plates. All studies were performed at confluence.

Protocols for Measurement of ET-1

RPAEC were exposed to various treatments for 4 h in serum-free PRF-DMEM. To test the effect of antioxidants or an iron chelator on basal ET-1 release, cells were incubated with DMTU (10 mM), ascorbic acid (1 mM), deferoxamine, (1.8 µM), and urea (10 mM) for 4 h. To study the effect of H2O2 on ET-1 production, cells were exposed to varying concentrations of glucose oxidase (0, 0.1, 1, 2.5, 5, and 10 mU/ml) in the presence of 5.6 mM glucose. We also studied the effect of treatment with catalase (2,000 U/ml), DMTU (10 mM), or urea (10 mM) in the presence of glucose and glucose oxidase (10 mU/ml) for 4 h.

ET-1 Assay

After incubation, the supernatants were removed, and ET-1 was measured using a commercially available RIA kit as previously described (15). The lower limit of sensitivity for ET-1 detection was 2 pg. Intra-assay variation was <9%; interassay variation was <15%. After extraction in 1 ml of 0.1 N NaOH, the protein level in each well was measured from an aliquot of the solubilized cells using the BCA protein assay reagents. ET-1 measurements were expressed as picograms ET-1 per milligram total cell protein.

Protocol for Measuring Intracellular cGMP

We tested the effect of glucose and glucose oxidase exposure for 15 min and 4 h on intracellular cGMP levels. In the experiments with 15 min of exposure to glucose and glucose oxidase, confluent monolayers of pulmonary endothelial cells were equilibrated with Krebs buffer containing 0.1 mM 3-isobutyl-1-methylxanthine (IBMX) at 37°C for 30 min before adding the glucose oxidase. The Krebs buffer contained (in mM) 145 NaCl, 10 N-2-hydroxyethylpiperazine-N'-2-ethanesulfonic acid, 5 glucose, 5 KCl, 1 Na2HPO4, 2.5 CaCl2, and 1.8 MgSO4-7H2O at pH 7.3. The cells were then incubated in Krebs buffer with or without 10 mU/ml glucose oxidase for 15 min at 37°C. Additional studies were performed in which the cells were incubated in media without IBMX and were exposed to varying concentrations of glucose oxidase (0, 2.5, 5, and 10 mU/ml) for 4 h. At the end of the experiments, the buffer was removed, and 1 ml of 100% ethanol-HCl solution (50 ml of 100% ethanol and 2 drops of concentrated HCl) was left on the cells overnight at 4°C.

cGMP Levels

cGMP levels were measured using a commercially available RIA kit. The cell homogenate was dried in a Speed-Vac (Savant, Farmingdale, NY). The pellet was resuspended in 500 µl of RIA assay buffer, and a 50-µl aliquot was incubated with 50 µl of 125I-labeled cGMP and 50 µl of anti-cGMP antiserum overnight at 4°C. Amerlex-M donkey anti-rabbit serum (200 µl) was then added for 10 min at room temperature. The samples were centrifuged at 3,000 revolutions/min for 10 min at 4°C, and the counts per minute in the pellet were determined. Total protein measurements were performed as outlined above, and the results were expressed as cGMP in femtomoles per milligram cell protein.

Measurement of ET-1 and beta -Actin mRNA by RT-PCR Amplification of RNA

Isolation of RNA. Confluent cultures of RPAEC were incubated in growth media for 24 h, the supernatant was removed, and the cells were washed with medium 199 without serum. Cells were exposed for 4 h to serum-free PRF-DMEM alone or with PRF-DMEM containing glucose oxidase (10 mU/ml). The supernatant was removed, and the cells were overlaid with 4 M guanidinium isothiocyanate, 1% 2-mercaptoethanol, and 1% sarcosyl (pH 7.0). The cells were homogenized by several passages through a 25-gauge needle, and 1/10th volume of 2 M sodium acetate was added. The RNA was phenol/chloroform extracted, precipitated in isopropanol, washed in 70% ethanol, and suspended in tris(hydroxymethyl)aminomethane (Tris) · HCl-EDTA. Each sample was quantified spectrophotometrically.

RT-PCR Amplification of RNA

RT-PCR amplification of RNA was performed as previously described by this laboratory (16). Five micrograms of total RNA from each sample were reverse transcribed by incubation with 250 pmol random hexamers, 400 units Moloney murine leukemia virus RT, 80 units RNasin, 2 mM deoxynucleotide triphosphates, 0.5 mM dithiothreitol, 75 mM KCl, 3 mM MgCl2, and 50 mM Tris · HCl (pH 8.3, final volume 50 µl) for 1 h at 37°C. The RT was inactivated by heating for 10 min at 94°C. The cDNA was stored at 4°C.

The cDNA was amplified by PCR. Each sample was measured for ET-1 and beta -actin in separate tubes using specific primers. The upstream and downstream primers for ET-1 were 5'-GCCAAGCAGACAAAGAACTCCGAG-3' and 5'-GCTCTGTAGTCAATGTGCTCGGTT-3', respectively. These give a 247-bp fragment that is complementary to position 371-618 in rat ET-1 cDNA (29). PCR of rat genomic DNA yields a single 1,300-bp product, indicating that these primers span an intron. The upstream and downstream primers for beta -actin were 5'-TGGAGAAGAGCTATGAGCTGCCTG-3' and 5'-GTGCCACCAGACAGCACTGTGTTG-3', respectively, which produces a single band corresponding to a 201-bp cDNA fragment. PCR of rat genomic DNA with the beta -actin primers yields a 289-bp product that is complementary to position 2499-2788 in the beta -actin gene, confirming that this primer set spans an intron. Finally, the ET-1 and beta -actin product sequences were verified by Margaret Robinson in Dr. Ray White's laboratory at the University of Utah, using fluoresceinated primer ends and cycle sequencing.

PCR was performed by incubating 5 µl of sample cDNA with 50 mM KCl, 10 mM Tris · HCl, 1.5 mM MgCl2, 0.01% gelatin, 200 µM total dNTP, 2 units Taq DNA polymerase, 2.5% formamide, 0.15 µCi [32P]dCTP, and 100 pmol ET-1 or beta -actin primers in 50 µl final volume (final pH 8.3 at room temperature). PCR using beta -actin primers was carried out for 25 cycles (15 s at 94°C, 15 s at 65°C, 30 s at 72°C) after 1 min of early DNA denaturation at 94°C using a Perkin-Elmer Cetus 9600 GeneAMP PCR System. PCR using ET-1 was carried out for 30 cycles under identical conditions. Different primers were never combined in the same tube. Twenty microliters of the final PCR reaction were electrophoresed on a 7% nondenaturing polyacrylamide gel. Gels were stained with ethidium bromide, and the bands corresponding to the cDNA product were excised, mixed with scintillation cocktail, and the counts per minute were determined on a Beckman beta counter.

ET-1 and beta -actin cDNA obtained from PCR of reverse transcribed RNA were used to generate standard curves. The cDNA was amplified by PCR, and the resultant amplified product was divided into small fractions that were, in turn, reamplified. After removal of primers using Magic PCR Prep (Promega), the purity of the final product was confirmed by electrophoresis. At the end of the purification, the amount of standard cDNA was quantified spectrophotometrically. Standard curves for beta -actin or ET-1 were made by simultaneously amplifying sample cDNA and, in separate tubes, standard cDNA (10-1 to 10-8 ng/tube). Every PCR amplification included a standard curve. All PCR consisted of simultaneous amplification (in separate tubes) of cDNA for ET-1 and beta -actin. All results are expressed as femtograms ET-1 cDNA per picogram beta -actin cDNA to control for the amount of RNA initially reverse transcribed. The accuracy of this semiquantitative PCR technique has been described previously in detail (8, 12).

51Cr Release

When monolayers of RPAEC were 70-80% confluent in six-well culture plates, they were incubated in 1.5 ml/well Ryan's medium containing 1 µCi/ml 51Cr for 16-18 h. After labeling, the monolayers were washed three times with 1.5 ml/well PRF-DMEM, followed by incubation in 2 ml/well PRF-DMEM in the absence or presence of varying concentrations of glucose oxidase (0, 1, 2.5, 5, and 10 mU/ml). All of the experiments were performed at 37°C in an environment containing 5% CO2 for 4 h unless otherwise indicated. The supernate was then removed and centrifuged, and the cell-free supernate was used. The cells were then dissolved in 2 ml/well of 0.1 N NaOH. The samples were counted for 1 min in a Packard A5530 gamma counter (Packard Instrument, Downers Grove, IL). The percent 51Cr release was calculated as follows: {(S - BK)/[(S - BK) + (C - BK)]} × 100, where S represents the counts per minute in the cell-free supernatant, C represents the counts per minute remaining within the cells, and BK represents the background counts per minute. Specific 51Cr release represents the measured release minus baseline release.

Leucine Incorporation Assay

Protein synthetic rate was estimated by [3H]leucine incorporation into trichloroacetic acid (TCA)-precipitable protein. Confluent cultures of RPAEC in 24-well plates were exposed to PRF-DMEM alone or containing 1, 2.5, 5, or 10 mU/ml glucose oxidase for 4 h at 37°C in 5% CO2. [3H]leucine (1 µCi/ml) in PRF-DMEM was added for 10 min at 37°C, and the cells were rinsed five times with phosphate-buffered saline and were solubilized with 0.1% sodium dodecyl sulfate. Proteins were precipitated in 10% TCA at 4°C for 2 h in the presence of 2 mg bovine serum albumin. The precipitate was centrifuged and rinsed two times in 10% TCA, and the counts per minute were determined in a Beckman LS6000 beta -counter (Beckman Instruments, Fullerton, CA).

Measurement of H2O2 Produced by Glucose and Glucose Oxidase

The amount of H2O2 produced by glucose and glucose oxidase in PRF-HBSS was determined. Experiments were performed in six-well plates. A 0.2-ml aliquot of the sample was mixed with 0.8 ml of 100 mM potassium phosphate buffer (pH 7.0). The final concentrations of the reagents in the 1 ml reaction mixture were 20 U/ml horseradish peroxidase (type II, 200 purpurogallin U/mg), 1.5 mM 4-aminoantipyrine, and 0.11 M phenol. Absorbance was measured at 510 nm with a Hitachi U-3210 spectrophotometer (Hitachi, Tokyo, Japan). The concentration of H2O2 was calculated using a molar extinction coefficient of 6.58 mM-1 · cm-1.

Statistical Analysis

Data were analyzed by the Mann-Whitney test, unpaired Student's t-test, or analysis of variance. Statistical significance was taken as P < 0.05. Values are presented as means ± SE.

    RESULTS
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Abstract
Introduction
Materials & Methods
Results
Discussion
References

Effect of Antioxidants on Basal ET-1 Release

Treatment of RPAEC with the antioxidant DMTU or the iron chelator deferoxamine doubles ET-1 production (Fig. 1). In contrast, ascorbate does not affect ET-1 release (Fig. 1). Because DMTU is a urea derivative, we used urea as a control, and it did not alter ET-1 production (Fig. 1).


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Fig. 1.   Effect of an iron chelator and an antioxidant on basal release of endothelin-1 (ET-1) from rat pulmonary arterial endothelial cells. Cells were exposed to phenol red-free Dulbecco's modified Eagle's medium (DMEM) with or without deferoxamine (1.8 µM), 1,3-dimethyl-2-thiourea (DMTU, 10 mM), urea (10 mM), or ascorbate (1 mM) for 4 h. Values are expressed as %control (mean + SE). Control ET-1 release was 48 ± 5 pg/mg cell protein; n = 12 for control, deferoxamine, and ascorbate; n = 8 for DMTU and urea. * P < 0.001 vs. control and dagger  P < 0.01 vs. control.

Effect of Oxidant Stress on Basal ET-1 Release

Glucose and glucose oxidase selectively generate H2O2 in a dose-dependent fashion. Glucose oxidase (10 mU/ml) and glucose (5.6 mM) produce ~195 µM H2O2 over 4 h. Exposure of cells to glucose oxidase and glucose causes a concentration-dependent decrease in ET-1 production (Fig. 2). The decrease in ET-1 synthesis occurs at doses that do not affect [3H]leucine incorporation into protein or specific 51Cr release (Table 1).


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Fig. 2.   Glucose oxidase (GO) combined with glucose causes a concentration-dependent decrease in ET-1 release from rat pulmonary arterial endothelial cells. Cells were exposed to varying concentrations of GO (0.1-10 mU/ml) in phenol red-free DMEM for 4 h; n = 8-12 at each concentration. Values are %control (mean + SE). Control ET-1 release was 65 ± 2 pg/mg cell protein. § P < 0.005 vs. control and * P < 0.001 vs. control.

                              
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Table 1.   Concentration effect of glucose oxidase on [3H]leucine incorporation and 51Cr release from rat pulmonary arterial endothelial cells

Catalase completely prevents the decrease in ET-1 production caused by glucose oxidase (Fig. 3). Exposing cells to both DMTU and glucose oxidase results in ET-1 levels that are intermediate between the effects of DMTU alone and glucose plus glucose oxidase by themselves [DMTU 209 ± 28% (SE) control, glucose and glucose oxidase 58 ± 8% control, and DMTU + glucose oxidase 121 ± 6% control]. Urea does not prevent the inhibitory effect of glucose oxidase on ET-1 release (Fig. 3).


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Fig. 3.   Effect of catalase (Cat; 2,000 U/ml) or DMTU (10 mM) on the decrease in ET-1 production caused by exposure of rat pulmonary arterial endothelial cells to GO (10 mU/ml) for 4 h. Urea (10 mM) is ineffective; n > 8 in each group. Values are %control (mean + SE). Control ET-1 release was 39 ± 2 pg/mg cell protein. dagger  P < 0.001 vs. control without (-) GO; § P < 0.005 vs. GO alone (+); * P < 0.005 vs. control; ¶ P < 0.01, DMTU vs. DMTU + GO; and infinity  P < 0.02, DMTU + GO vs. GO.

Oxidant exposure not only decreases ET-1 release (Fig. 2) but also ET-1 mRNA as assessed by semiquantitative PCR (Fig. 4). We tested the possibility that the H2O2 generated by glucose and glucose oxidase might increase intracellular cGMP levels and thereby reduce ET-1 release. Exposure of RPAEC to glucose and glucose oxidase for 15 min or 4 h, however, does not affect intracellular cGMP levels (Fig. 5).


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Fig. 4.   GO decreases ET-1 mRNA in rat pulmonary arterial endothelial cells, as assessed by semiquantitative polymerase chain reaction and normalized for the concentration of beta -actin mRNA. Cells were exposed to GO (0, 2.5, 5, and 10 mU/ml) in phenol red-free DMEM for 4 h; n = 3 for each concentration. Values are %control (mean + SE). Control values were 304 ± 16 fg ET-1 cDNA/pg beta -actin cDNA. * P < 0.01 vs. control without GO.


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Fig. 5.   GO and glucose do not alter intracellular cGMP levels in rat pulmonary endothelial cells. Cells in the presence of glucose were exposed with or without GO (10 mU/ml) for 15 min (A) in the presence of 0.1 mM 3-isobutyl-1-methylxanthine (IBMX). Cells were pretreated with IBMX for 30 min at 37°C before adding GO. Concentration effect of GO exposure (0, 2.5, 5, and 10 mU/ml) for 4 h in the absence of IBMX was also determined (B). Values are means + SE; n = 4-6 at each time point.

    DISCUSSION
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Abstract
Introduction
Materials & Methods
Results
Discussion
References

The current study demonstrates that changes in oxidant stress regulate ET-1 generation. Treatment with the antioxidant DMTU or the iron chelator deferoxamine substantially increases baseline endothelial ET-1 release (Fig. 1), implying that basal oxidant stress tonically reduces ET-1 production. In contrast, increasing oxidant stress with glucose oxidase reduces in a dose-dependent manner ET-1 production by rat pulmonary arterial endothelial cells. H2O2 mediates this effect of glucose and glucose oxidase since catalase, which converts H2O2 to water and oxygen, completely abrogates this inhibitory effect on pulmonary endothelial ET-1 release. Previous work indicates that the superoxide anion or hydroxyl radical may reduce the level of immunoreactive ET-1 by inducing structural changes in the molecule (9, 34). We and others, however, have demonstrated that H2O2 by itself does not alter ET-1 immunoreactivity (9, 34). Although H2O2 may cause cytotoxicity, we found no evidence of generalized cell damage or dysfunction (as assessed by [3H]leucine incorporation into protein and 51Cr release) at concentrations of glucose oxidase that substantially reduce ET-1 production. Additionally, exposure to glucose and glucose oxidase causes a concomitant decrease in ET-1 mRNA as assessed by semiquantitative PCR. Our finding that both endogenous and exogenous oxidants reduce ET-1 synthesis without frank cell damage provides strong evidence that short-term oxidative stress can inhibit pulmonary endothelial ET-1 production.

Catalase, which eliminates H2O2, completely prevents the effect of glucose and glucose oxidase (Fig. 3). Unlike catalase, DMTU does not alter H2O2 levels but likely scavenges subsequent radicals, such as the hydroxyl radical. Combining DMTU, glucose, and glucose oxidase results in ET-1 levels that are intermediate between the effects of DMTU alone versus glucose plus glucose oxidase by themselves. The intermediate ET-1 level seen with DMTU and glucose oxidase may reflect their net effect on intracellular oxidant stress. The ability of catalase, but not DMTU, to completely block the glucose oxidase-mediated decrease in ET-1 suggests that H2O2 by itself can lead to a decrease in ET-1 synthesis.

Relatively little data exist on oxidant regulation of ET-1 production. H2O2 (0.1-20 mM) has been reported to reduce ET-1 release by human umbilical vein endothelial cells (22). This study, however, did not assess cytotoxicity. In contrast, another group detected no effect of exogenous superoxide anion or H2O2 on ET-1 release from bovine pulmonary artery endothelial cells (24). These investigators found, however, that exogenous hydroxyl radical production augmented ET-1 release, but at concentrations that elicited cell damage (24). This report did not measure ET-1 mRNA levels. Other studies suggest that H2O2 may increase ET-1 mRNA levels in bovine aortic endothelial cells (ET-1 release was not measured; see Ref. 25) and ET-1 release by, and mRNA levels in, human renal mesangial cells (9). The reasons for these disparate results are unknown but may relate to differences in cell type, duration, or concentration of oxidant exposure, technique (most studies have failed to closely assess cytotoxicity), or other factors. As an example of the potential for cell-specific differences, one mechanism by which H2O2 may regulate gene expression is by activating transcription factors, such as nuclear factor (NF)-kappa B (18, 30). The ability of H2O2 to activate NF-kappa B, however, depends on the cell type studied (18, 30). Additionally, the capacity of H2O2 to induce NF-kappa B in endothelial cells varies with the vascular bed studied (1, 2). H2O2, for example, activates NF-kappa B in porcine aortic endothelial cells (1) but not in human umbilical vein endothelial cells (2).

The observation that oxidants can modulate vasoactive factor production has precedence. Reactive oxygen species, for example, can activate phospholipase A2 and enhance arachidonic acid release and mediator production from pulmonary endothelium (5), stimulate thromboxane A2 production in alveolar macrophages (31), differentially alter the synthesis of prostacyclin and 15-hydroxyeicosatetraenoic acid in coronary artery endothelium (4), increase platelet-activating factor synthesis in pulmonary endothelial cells (14), and augment cytokine-induced nitric oxide synthesis (20). Additionally, superoxide anion can combine with nitric oxide, thereby inactivating nitric oxide.

The mechanisms by which oxidants regulate ET-1 production remain speculative. Relatively few factors decrease ET-1 synthesis, and, frequently, they appear to act by enhancing cGMP accumulation (11). This mechanism does not explain the findings in the current study since glucose oxidase had no effect on RPAEC cGMP levels. Reactive oxygen species can affect a multitude of signaling pathways involved in regulating gene expression, including factors interacting with promoter elements, such as the antioxidant response element (27), CC(A/T)6GG sequences (6), and consensus binding sites for transcription factors like heat shock factor (3), activator protein (AP)-1 (18), AP-1/E26 virus transformation-specific protein (26), and NF-kappa B (30). Which, if any, of these factors contribute to H2O2 regulation of ET-1 production by RPAEC remains to be determined.

Elevated circulating levels of ET-1 have been observed in patients with the acute respiratory distress syndrome (7, 13, 21), a condition associated with increased H2O2 production by the lung (32). Plasma ET-1 levels in these patients average five- to eightfold higher than in control subjects. Animal models of acute lung injury associated with increased oxidant production, such as ischemia-reperfusion (23, 33), also elevate ET-1 levels. Pulmonary ischemia followed by reperfusion increases circulating ET-1 (23), ET-1 release from the lung (33), and ET-1 mRNA in pulmonary tissue (23). Phosphoramidon, an inhibitor of endothelin-converting enzyme, prevents ET-1 release and markedly reduces the increase in pulmonary insufflation pressure in a model of lung ischemia-reperfusion (33). Additionally, in a rat model of unilateral warm ischemia and reperfusion lung injury, an endothelin subtype A (ETA) receptor blocker reduces the fall in arterial oxygenation, the gain in wet/dry lung weight, and the extent of neutrophil influx that develops 90 min after reperfusion (23). Blockade of ETA receptors also prevents the increase in pulmonary capillary permeability caused by ischemia-reperfusion in rat lungs (10). Thus injury to the pulmonary vascular bed in both animals and humans significantly elevates circulating ET-1 levels, which may contribute to the pathogenesis. The mechanisms underlying the increase in circulating (7, 13, 21, 23) and pulmonary ET-1 (23, 33) are unknown.

Although oxidant exposure appears to increase ET-1 synthesis in the lung, the oxidant species involved or what cells produce ET-1 is unknown. Our results indicate that the level of oxidant stress can regulate ET-1 production by cultured pulmonary endothelial cells. Treatment with an antioxidant or an iron chelator increases basal ET-1 synthesis. Conversely, short-term exposure to exogenous H2O2 decreases ET-1 synthesis. This observation has important implications regarding the mechanisms responsible for the enhanced ET-1 generation in oxidant lung injury. It implies that other oxidant species or additional factors, such as the presence of inflammatory mediators, are necessary to augment pulmonary endothelial ET-1 synthesis or that other lung cells distinct from endothelial cells serve as the source of the increased ET-1 production.

    ACKNOWLEDGEMENTS

This research was supported by Department of Veterans Affairs Medical Research Funds (to D. E. Kohan and J. R. Michael) and by National Institutes of Health Grants R29 DK-44440 (to D. E. Kohan) and the University of Utah Specialized Center of Research on Acute Lung Injury Grant IP 50 HL-50153 (to J. R. Michael). B. A. Markewitz was supported by an American Lung Association Research Training Fellowship Award, an Edward P. Stiles Trust Grant from the Louisiana State University Medical Center at Shreveport, and the Board of Regents of the State of Louisiana through the Louisiana Education Quality Support Fund (1996-99)-RD-A-20.

    FOOTNOTES

Address for reprint requests: J. R. Michael, Div. of Respiratory, Critical Care and Occupational Pulmonary Medicine, 725 Wintrobe Bldg, 50 North Medical Dr., University of Utah Medical Center, Salt Lake City, UT 84132.

Received 5 November 1996; accepted in final form 20 June 1997.

    REFERENCES
Top
Abstract
Introduction
Materials & Methods
Results
Discussion
References

1.   Barchowsky, A., S. R. Munro, S. J. Morana, M. P. Vincenti, and M. Treadwell. Oxidant-sensitive and phosphorylation-dependent activation of NF-kappa B and AP-1 in endothelial cells. Am. J. Physiol. 269 (Lung Cell. Mol. Physiol. 13): L829-L836, 1995[Abstract/Free Full Text].

2.   Bradley, J. R., D. R. Johnson, and J. S. Pober. Endothelial activation by hydrogen peroxide: selective increases of intracellular adhesion molecule-1 and major histocompatibility complex class I. Am. J. Pathol. 142: 1598-1609, 1993[Abstract].

3.   Bruce, J. L., B. D. Price, C. N. Coleman, and S. K. Calderwood. Oxidative injury rapidly activates the heat shock transcription factor but fails to increase levels of heat shock proteins. Cancer Res. 53: 12-15, 1993[Abstract].

4.   Callahan, K. S., and J. G. Garcia. Oxidant exposure stimulates cultured coronary artery endothelial cells to release 15-HETE: differential effects on PGI2 and 15-HETE synthesis. J. Lab. Clin. Med. 124: 569-578, 1994[Medline].

5.   Chakraborti, S., G. H. Gurtner, and J. R. Michael. Oxidant-mediated activation of phospholipase A2 in pulmonary endothelium. Am. J. Physiol. 257 (Lung Cell. Mol. Physiol. 1): L430-L437, 1989[Abstract/Free Full Text].

6.   Datta, R., N. Taneja, V. P. Sukhatme, S. A. Qureshi, R. Weichselbaum, and D. W. Kufe. Reactive oxygen intermediates target CC(A/T)6GG sequences to mediate activation of the early growth response 1 transcription factor gene by ionizing radiation. Proc. Natl. Acad. Sci. USA 90: 2419-2422, 1993[Abstract].

7.   Druml, W., H. Steltzer, W. Waldhèusl, K. Lenz, A. Hammerle, H. Vierhapper, S. Gasic, and O. F. Wagner. Endothelin-1 in adult respiratory distress syndrome. Am. Rev. Respir. Dis. 148: 1169-1173, 1993[Medline].

8.   Hughes, A. K., R. C. Cline, and D. E. Kohan. Alterations in renal endothelin production in the spontaneously hypertensive rat. Hypertension 20: 666-673, 1992[Abstract].

9.   Hughes, A. K., P. K. Stricklett, E. Padilla, and D. E. Kohan. Effect of reactive oxygen species on endothelin-1 production by human mesangial cells. Kidney Int. 49: 181-189, 1996[Medline].

10.   Khimenko, P. L., T. M. Moore, and A. E. Taylor. Blocked ETA receptors prevent ischemia and reperfusion injury in rat lungs. J. Appl. Physiol. 80: 203-207, 1996[Abstract/Free Full Text].

11.   Kohan, D. E. Endothelins in the kidney: physiology and pathophysiology. Am. J. Kidney Dis. 22: 493-510, 1993[Medline].

12.   Kohan, D. E., and E. Padilla. Osmolar regulation of endothelin-1 production by rat inner medullary collecting duct. J. Clin. Invest. 91: 1235-1240, 1993[Medline].

13.   Langleben, D., M. DeMarchie, D. Laporta, A. H. Spanier, R. D. Schlesinger, and D. J. Stewart. Endothelin-1 in acute lung injury and the adult respiratory distress syndrome. Am. Rev. Respir. Dis. 148: 1646-1650, 1993[Medline].

14.   Lewis, M. S., R. E. Whatley, P. Cain, T. M. McIntyre, S. M. Prescott, and G. A. Zimmerman. Hydrogen peroxide stimulates the synthesis of platelet-activating factor by endothelium and induces endothelial cell-dependent neutrophil adhesion. J. Clin. Invest. 82: 2045-2055, 1988[Medline].

15.   Markewitz, B. A., D. E. Kohan, and J. R. Michael. Endothelin-1 synthesis, receptors and signal transduction in a cloned rat alveolar epithelial cell line: evidence for an autocrine role. Am. J. Physiol. 268 (Lung Cell. Mol. Physiol. 12): L192-L200, 1995[Abstract/Free Full Text].

16.   Markewitz, B. A., J. R. Michael, and D. E. Kohan. Cytokine-induced expression of a nitric oxide synthase in rat renal tubule cells. J. Clin. Invest. 91: 2138-2143, 1993[Medline].

17.   McMillen, M. A., M. Huribal, R. Kumar, and B. E. Sumpio. Endothelin-stimulated human monocytes produce prostaglandin E2 but not leukotriene B4. J. Surg. Res. 54: 331-335, 1993[Medline].

18.   Meyer, M., R. Schreck, and P. A. Baeuerle. H2O2 and antioxidants have opposite effects on activation of NF-kB and AP-1 in intact cells: AP-1 as secondary antioxidant-responsive factor. EMBO J. 12: 2005-2015, 1993[Abstract].

19.   Michael, J. R., and B. A. Markewitz. Endothelins and the lung. Am. J. Respir. Crit. Care Med. 154: 555-581, 1996[Medline].

20.   Milligan, S. A., M. W. Owens, and M. B. Grisham. Augmentation of cytokine-induced nitric oxide synthesis by hydrogen peroxide. Am. J. Physiol. 271 (Lung Cell. Mol. Physiol. 15): L114-L120, 1996[Abstract/Free Full Text].

21.   Mitaka, C., Y. Hirata, T. Nagura, Y. Tsunoda, and K. Amaha. Circulating endothelin-1 concentrations in acute respiratory failure. Chest 104: 476-480, 1993[Abstract].

22.   Mitchell, M. D., D. W. Branch, S. Lamarche, and D. J. Dudley. The regulation of endothelin production in human umbilical vein endothelial cells: unique inhibitory action of calcium ionophores. J. Clin. Endocrinol. Metab. 75: 665-668, 1992[Abstract].

23.   Okada, M., C. Yamashita, M. Okada, and K. Okada. Contribution of endothelin-1 to warm ischemia/reperfusion injury of the rat lung. Am. J. Respir. Crit. Care Med. 152: 2105-2110, 1995[Abstract].

24.   Prasad, M. R., R. M. Jones, and D. L. Kreutzer. Release of endothelin from cultured bovine endothelial cells. J. Mol. Cell. Cardiol. 23: 655-658, 1991[Medline].

25.   Rodríguez-Puyol, D., S. López-Ongil, J. Lucio, S. Lamas, P. Ruiz, and M. Rodríguez-Puyol. Modulation of pre-pro-endothelin and constitutive nitric oxide synthase mRNA expression by reactive oxygen species in bovine aortic endothelial cells (Abstract). J. Am. Soc. Nephrol. 5: 590, 1994.

26.   Roebuck, K. A., A. Rahman, V. Lakshminaravanan, K. Janakidevi, and A. B. Malik. H2O2 and tumor necrosis factor-alpha activate intracellular adhesion molecule (ICAM-1) gene transcription through distinct cis-regulatory elements within the ICAM-1 promoter. J. Biol. Chem. 270: 18966-18974, 1995[Abstract/Free Full Text].

27.   Rushmore, T. H., M. R. Morton, and C. B. Pickett. The antioxidant responsive element: activation by oxidative stress and identification of the DNA consensus sequence required for functional activity. J. Biol. Chem. 266: 11632-11639, 1991[Abstract/Free Full Text].

28.   Ryan, U. S., and L. White. Microvascular endothelium isolation with microcarriers: arterial, venous. J. Tissue Cult. Methods 10: 9-13, 1986.

29.   Sakurai, T., M. Yanagisawa, A. Inoue, U. S. Ryan, S. Kimura, Y. Mitsui, K. Goto, and T. Masaki. cDNA cloning, sequence analysis and tissue distribution of rat preproendothelin-1 mRNA. Biochem. Biophys. Res. Commun. 175: 44-47, 1991[Medline].

30.   Schreck, R., P. Rieber, and P. A. Baeuerle. Reactive oxygen intermediates as apparently widely used messengers in the activation of the NF-kappa B transcription factor and HIV-1. EMBO J. 10: 2247-2258, 1991[Abstract].

31.   Sporn, P. H. S., M. Peters-Golden, and R. H. Simon. Hydrogen-peroxide-induced arachidonic acid metabolism in the rat alveolar macrophage. Am. Rev. Respir. Dis. 137: 49-56, 1988[Medline].

32.   Sznajder, J. I., A. Fraiman, J. B. Hall, W. Sanders, G. Schmidt, G. Crawford, A. Nahum, P. Factor, and L. D. Wood. Increased hydrogen peroxide in the expired breath of patients with acute hypoxemic respiratory failure. Chest 96: 606-612, 1989[Abstract].

33.   Vemulapalli, S., M. Rivelli, P. J. S. Chiu, M. delPrado, and J. A. Hey. Phosphoramidon abolishes the increases in endothelin-1 release induced by ischemia-hypoxia in isolated perfused guinea pig lungs. J. Pharmacol. Exp. Ther. 262: 1062-1069, 1992[Abstract].

34.   Yasuda, N., Y. Kasuya, G. Yamada, H. Hama, T. Masaki, and K. Goto. Loss of contractile activity of endothelin-1 induced by electrical field stimulation-generated free radicals. Br. J. Pharmacol. 113: 21-28, 1994[Abstract].


AJP Lung Cell Mol Physiol 273(4):L768-L774