NO regulates LPS-stimulated cyclooxygenase gene expression and activity in pulmonary artery endothelium

Jian-Xiong Chen1, Leonard C. Berry Jr.2, Brian W. Christman2, Miles Tanner2, Paul R. Myers2, and Barbara O. Meyrick1,2

Departments of 1 Pathology and 2 Medicine, Center for Lung Research, Vanderbilt University, Nashville, Tennessee 37232


    ABSTRACT
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

We examined whether nitric oxide (NO) inhibits prostanoid synthesis through actions on cyclooxygenase (COX) gene expression and activity. Bovine pulmonary artery endothelial cells were pretreated for 30 min with the NO donors 1 mM S-nitroso-N-acetylpenicillamine (SNAP), 0.5 mM sodium nitroprusside (SNP), or 0.2 µM spermine NONOate; controls included cells pretreated with either 1 mM N-acetyl-D-penicillamine or the NO synthase (NOS) inhibitor 1 mM NG-nitro-L-arginine methyl ester with and without addition of lipopolysaccharide (LPS; 0.1 µg/ml) for 8 h. COX-1 and COX-2 gene and protein expression were examined by RT-PCR and Western analysis, respectively; prostanoid measurements were made by gas chromatography-mass spectrometry, and COX activity was studied after a 30-min incubation with 30 µM arachidonic acid. LPS induced COX-2 gene and protein expression and caused an increase in COX activity and an eightfold increase in 6-keto-PGF1alpha release. LPS-stimulated COX-2 gene expression was decreased by ~50% by the NO donors. In contrast, LPS caused a significant reduction in COX-1 gene expression and treatment with NO donors had little effect. SNAP, SNP, and NONOate significantly suppressed LPS-stimulated COX activity and 6-keto-PGF1alpha release. Our data indicate that increased generation of NO attenuates LPS-stimulated COX-2 gene expression and activity, whereas inhibition of endogenous NOS has little effect.

prostaglandins; endotoxin; arachidonic acid; nitric oxide; lipopolysaccharide


    INTRODUCTION
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

DURING INFLAMMATION, endothelial cells synthesize growth factors, cytokines, and other autacoids involved in cell signaling. These various messengers modulate blood and vascular cell functions and are involved in both local and systemic inflammation, e.g., septic shock. Exposure of endothelial cells to proinflammatory bacterial lipopolysaccharide (LPS) induces the synthesis of cytokines and secondary mediators such as prostaglandins and nitric oxide (NO) (4, 16, 17, 22, 23). Prostaglandins, arachidonic acid (AA) metabolites of cyclooxygenases (COXs), and NO have major roles in regulating inflammation and blood vessel dilation. Both COX and NO synthase (NOS) exist in a constitutive form (COX-1 and ecNOS, respectively) in many cells and are responsible for modulation of a number of basal physiological functions. The inducible forms of both enzymes (COX-2 and iNOS, respectively) are the major isoforms expressed in inflammatory conditions and catalyze the production of high levels of prostaglandins and NO (7, 12).

Stimulation of a wide variety of cell types, including endothelial cells, with cytokines and other agents results in upregulation of COX-2, and this is often accompanied by an increase in NOS (1, 31). Whether the increase in NOS occurs before that of COX-2 is not certain, and it is possible that regulation of each enzyme may be cell specific. For example, in vitro, NO stimulates COX activity in both a LPS-activated mouse macrophage cell line RAW 264.7 and fibroblasts (24) but inhibits the production of PGE2 by rat Kupffer cells (27) and in cultured bovine endothelial cells (14). The mechanism for these contradictory findings is unknown. Studies in rat mesangial and human lung epithelial cells suggest that NO may directly interact with COX at the transcriptional level and that this effect is mediated through a cGMP-dependent mechanism (30, 32). Regulation of COX-2 gene expression by NO also has been suggested to occur posttranscriptionally in rat osteoblasts (10).

In previous studies, we showed that pretreatment of LPS-stimulated endothelial cells with antioxidants such as dimethyl sulfoxide (DMSO) and dimethyl thiourea strikingly attenuated LPS-stimulated prostaglandin release, suggesting that intracellular generation of reactive oxygen species may contribute to increased synthesis of prostaglandins (4, 17, 22). A role for reactive oxygen species was further delineated by our finding that DMSO attenuated LPS-stimulated mangano superoxide dismutase mRNA (17, 26). The mechanism by which reactive oxygen species regulate COX is incompletely understood. The present study examines the hypothesis that NO modulates expression of the COXs in normal and LPS-treated endothelial cells. Using bovine pulmonary artery endothelial cells (BPAEC), we examined the effect of several NO donors and an inhibitor of NOS on COX-1 and COX-2 gene expression and protein, COX activity, and prostaglandin release in LPS-stimulated and untreated BPAEC. Our data demonstrate that increased generation of NO regulates basal expression of COX-1 and LPS-stimulated COX-2 expression, whereas decreased intracellular levels of NO have little effect on prostaglandin synthesis and release.


    MATERIALS AND METHODS
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

Cell culture. Primary cultures of BPAEC were used in all experiments. Fresh pulmonary arteries were obtained from a local slaughterhouse, and endothelial cells were isolated by scraping the endothelial surfaces as previously described (16). The cells were cultured in medium containing 5% Nuserum (Collaborative Research, Bedford, MA), 3% fetal bovine serum (FBS; Atlanta Biological, Atlanta, GA) 2% bovine calf serum (HyClone, Logan UT), 50 U/ml penicillin, 50 µg/ml streptomycin, and 40 µg/ml gentamicin in medium 199 (GIBCO BRL, Grand Island, NY). Each of the endothelial lines used in these experiments had a typical cobblestone morphology, showed uptake of acetylated low-density lipoprotein, and exhibited factor VIII-related antigen (16). Confluent endothelial monolayers from at least three different cell lines, between passages 4 and 10, were used for each experimental protocol.

Experimental protocol. BPAEC were exposed to one of the following three NO donors: 1) S-nitroso-N-acetylpenicillamine (SNAP; 1 mM), 2) sodium nitroprusside (SNP; 0.5 mM), or 3) spermine NONOate (0.2 µM). Further sister cells were treated with either NG-nitro-L-arginine methyl ester (L-NAME; 1 mM) or an inactive form of SNAP, N-acetyl-D-penicillamine (NAP; 1 mM). All experiments and controls were carried out in medium 199 containing 2% FBS. All drugs were dissolved and diluted in medium 199 containing 2% FBS except for NAP, which was dissolved in DMSO. The final dilution of DMSO used in the experiment was 0.01%, a concentration that we showed earlier does not affect the LPS response in BPAEC (4). Untreated sister wells also were used as controls. After 30 min of pretreatment with each drug, LPS (0.1 µg/ml Escherichia coli endotoxin 055:B5, Difco Laboratories, Detroit, MI) was added to some of the wells and the cells were incubated for periods up to 24 h. Cell-free supernatants were collected at 8 h from 24-well plates for measurements of 6-keto-PGF1alpha and PGE2 release. Cell pellets were collected at 0, 2, 4, 6, 8, 18, and 24 h from P100 dishes for RT-PCR and Western blot analysis.

The dilutions for each of the NO donors were chosen on the basis of dose-response curves. For SNAP, the dilutions examined were from 0.3 to 3 mM; for SNP, the dilutions were from 0.01 to 10 mM; and for NONOate, the dilutions were from 0.04 to 4 µM. The dilution chosen for use in our experiments was the dilution that gave the best protection against LPS-stimulated 6-keto-PGF1alpha release after 8 h of incubation (data not shown).

Prostanoid measurements. Measurements of 6-keto-PGF1alpha , the stable metabolite of prostacyclin, and PGE2 were made by gas chromatography-mass spectrography (GC-MS). Briefly, after 8 h of incubation, 1 ng of tetradeuterated PGE2 was added to 1 ml of supernatant. The pH of the supernatant was adjusted to 3.0 with 1 N HCl and purified with a preconditioned Sep-Pak C18 followed by a silica Sep-Pak cartridge. The samples were methoximated, and dried under nitrogen, and the free carboxylic acid groups were converted to corresponding esters with 20% pentafluorobenzyl bromide in acetonitrile. The samples were then purified by TLC and prepared for GC-MS. Samples for analysis of 6-keto-PGF1alpha were similarly prepared but with slightly different TLC mobile phases, and sample analysis was carried out after methoximation, conversion to pentafluorobenzyl esters, and trimethysilylation. GC-MS was performed on a 7-m SPB-1 fused silica column in a Varian-vista instrument. Selected ion currents monitored included mass-to-charge ratios of 524 (PGE2), 614 (6-keto-PGF1alpha ), 528 (tetradeuterated PGE2 internal standard), and 618 (the tetradeuterated 6-keto-PGF1alpha internal standard). Quantification occurred by stable isotope dilution, comparing the relative peak areas of endogenously produced eicosanoid and deuterated internal standard.

Measurement of COX activity. BPAEC were treated with either intervention alone or intervention plus LPS. After 8 h of incubation, the supernatant was replaced with fresh medium containing AA (30 µM) and the cells were incubated for a further 30 min. At the end of this time, the supernatant was collected for the measurement of prostanoids by GC-MS as outlined above.

RT-PCR. Total RNA was isolated using RNA STAT-60 (Teltest B, Friendswood, TX) after the Teltest protocol. The RNA concentration and purity were estimated from the optical density at 260 and 280 nm. Total RNA (1 µg) was then subjected to RT-PCR analysis using an Access RT-PCR System Kit (Promega, Madison, WI) and specific primers for bovine COX-1 and COX-2, which were designed and synthesized according to their sequences (2): bovine COX-1 sense, 5'-TCC AAC CTT ATC CCC AGC C-3' and antisense, 5'-CAT GGC GAT GCG GTT GC-3' (777 bp); bovine COX-2 sense, 5'-TCC AGA TCA CAT TTG ATT GAC A-3' and antisense, 5'-TCT TTG ACT GTG GGA GGA TAC A-3' (449 bp). Levels of glyceraldehyde-3-phosphate dehydrogenase (GAPDH) gene expression were used as internal standards. This was achieved by the simultaneous addition of GAPDH primers (size of PCR product 356 bp; sense, 5'-GAG ATG ATG ACC CTT TTG GC-3' and antisense, 5'-GTG AAG GTC GGA GTC AAC G-3') to the RT-PCR amplification mixture. First-strand cDNA sythesis was performed following the manufacturer's procedures. The conditions for the PCR were as follows: after initial denaturation at 94°C for 2 min, the thermocycler (Hybaid OMN-E, Middlesex, UK) was programmed for 30 cycles: 1 min at 94°C, 1 min at 55°C, and 90 s at 72°C. The reaction was concluded with final extension step at 72°C for 10 min.

Semiquantitative analysis of PCR products. A 15-µl aliquot from each PCR reaction was electrophoresed in a 2% agarose gel containing ethidium bromide (0.5 µg/ml) and was photographed under ultraviolet transillumination. The PCR band on the photograph was scanned using a densitometer (Bio-Rad model GS-700) linked to a computer analysis system (Molecular Analysis Software, Bio-Rad, Hercules, CA). COX-1 and COX-2 signals were normalized relative to the corresponding GAPDH signal from the same sample.

Western blot analysis. After incubation with the intervention, BPAEC were washed twice in PBS buffer, lysed in 200 µl of 1% Triton X-100, and harvested with a rubber policeman. The protein content was determined using a bicinchoninic acid (BCA) protein assay kit (Pierce, Rockford, IL), using BSA as the standard. Proteins (25 µg) were separated by SDS-PAGE on 10% polyacrylamide gels and then transferred to nitrocellulose. Nonspecific binding was blocked with 5% nonfat milk. This was followed by incubation with either human polyclonal COX-2 antibody (1:1,000 dilution) or COX-1 antibody (1:1,000 dilution, Oxford Biochem Research, Oxford, MI) for 1 h at room temperature. The membrane was then washed and incubated with horseradish peroxidase-conjugated, goat anti-rabbit IgG (1:2,000 dilution; Dako). The membrane was developed using Western blot chemiluminescence reagent (NEN Life Science Products, Boston, MA), and densitometric analysis of the COX-1 and COX-2 bands was carried out using an imager and densitometer (Bio-Rad) and Molecular Analysis Software as previously described.

Measurement of cell viability. Cell respiration, an indicator of cell viability, was assessed by the mitochondrial-dependent reduction of 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT) to formazan (19). After 8 h of incubation, cells in 96-well plates were incubated with MTT (0.2 mg/ml) for 1 h at 37°C. The culture medium was removed, and cells were solubilized in DMSO (100 µl). Reduction of MTT to formazan within cells was quantitated by measurement of optical density at 540 nm by using a microplate reader.

Measurement of NO release. NO release was measured using a specific chemiluminescence method and a NO analyzer (Dasibi model 2108, Glendale, CA) (20, 21). Briefly, monolayers of BPAEC grown on microcarrier beads were incubated for 2 h in 2% FBS in medium 199 containing either SNAP, SNP, or NONOate. In experiments with LPS, this was added for the last 90 min. The cells on beads were transferred to a siliconized glass perfusion chamber fitted with a frit and continuously perfused with Krebs physiological saline (4 ml/min) at 37°C. The medium was previously bubbled with 5% CO2-21% oxygen in nitrogen. After 10 min, the effluent was assayed to establish basal NO content. Measurements were carried out by transferring the cell effluent to a reflux chamber containing 1% sodium iodide in glacial acetic acid. The mixture was refluxed at 55°C while being continuously flushed with nitrogen gas, which in turn was transferred to the NO analyzer where NO reacted with ozone to produce light. Light was measured by a photomultiplier and converted into voltage and recorded on a standard flat-bed recorder. The signal produced was used to construct a voltage-concentration standard curve using known concentrations of NO.

Measurement of cGMP in RFL-6 reporter cells. As an assessment of NO release by the NO donors per se over time, cGMP levels were measured in the reporter cell line RFL-6. This is a widely used and sensitive assay for indirect measurement of NO production (5, 11). RFL-6 cells contain high levels of soluble guanylyl cyclase, and NO diffuses easily across membranes to activate this enzyme to produce cGMP. Each of the NO donors, SNAP, NONOate, and SNP, was dissolved in 2% FBS in medium 199 and incubated at 37°C for periods up to 150 min; NAP and L-NAME were used as controls. RFL-6 cells, grown to confluence in 6-well plates (1 × 106 cells/well), were washed twice with HEPES buffer, pH 7.4, and incubated for 20 min at 37°C in "spiked HEPES" [10 mM HEPES, 138 mM NaCl, 5 mM KCl, 2 mM CaCl2, 1 mM MgCl2, 10 mM glucose, and 0.5 mM 3-isobutyl-1-methylxanthine (Sigma)] containing 100 U/ml superoxide dismutase, 1 mM EGTA, 100 µM NADPH, 5 µM indomethacin, and 100 µM L-arginine, pH 7.0. At the end of this procedure, a 50-µl aliquot from each of the donors was taken after 60, 120, and 150 min of incubation, was added to the RFL-6 cells, and was incubated at 37°C for 2 min. The supernatant was removed, and the fibroblasts were frozen in liquid nitrogen. The fibroblasts were thawed on ice, and 1 ml of ice-cold 70% ethanol was added. The cells were then scraped, collected, and centrifuged at 600 g for 10 min, and the supernatant was transferred to microcentrifuge tubes and dried for 2-3 h in a spin-vac. Measurements of cGMP were made by ELISA (Amersham, Arlington Heights, IL). Each assay was carried out in duplicate. The results are expressed in femtomoles per microgram of protein. Protein levels were measured using a BCA protein assay kit on each cell pellet.

Statistics. The results are expressed as means ± SE. Further analysis was performed when appropriate using either a one-way ANOVA followed by the Scheffé's multiple comparison test or a paired Student's t-test. P < 0.05 was considered significant.


    RESULTS
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

Temporal effect of LPS on COX-2 and COX-1 gene expression. COX-2 gene expression was undetectable by RT-PCR analysis in control cells. LPS-induced COX-2 gene expression by 2 h; the expression of this gene peaked at 6 h and then gradually declined over the remainder of the study (Fig. 1A). Western analysis revealed a similar time course of induction for the 70-kDa COX-2 protein in LPS-treated cells (Fig. 1B). RT-PCR demonstrated COX-1 gene expression in untreated cells.


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Fig. 1.   A: representative RT-PCR showing induction of cyclooxygenase (COX)-2 gene expression after treatment with lipopolysaccharide (LPS) for periods up to 24 h. Gene expression peaked at 6 h and then declined (n = 3); gene expression of glyceraldehyde-3-phosphate dehydrogenase (GAPDH) is shown as a loading control. B: representative Western analysis showing peak induction of the 70-kDa COX-2 protein at 6 h of exposure to LPS.

In contrast to the findings for COX-2, treatment with LPS caused a gradual decline in COX-1 gene expression; by 2 h, the level was ~50% and continued to decline below the baseline value over the 24 h of the study (Fig. 2A). Western blot analysis for COX-1 protein was consistent with the LPS-induced reduction in gene expression (Fig. 2B).


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Fig. 2.   Representative RT-PCR (A) and Western analysis (B) (n = 3) showing gradual suppression of COX-1 gene expression after exposure to LPS for periods up to 24 h.

Effect of NO donors on COX-2 and COX-1 gene expression. Treatment with the NO donors SNAP, SNP, and NONOate significantly inhibited the LPS-stimulated increase in COX-2 gene expression seen at 8 h by ~50% (Fig. 3, A and B). Treatment with NAP and L-NAME failed to cause any detectable change in LPS-stimulated COX-2 gene expression. Western blot analysis confirmed these findings, demonstrating that SNAP, SNP, and NONOate resulted in a 50% reduction in COX-2 protein (Fig. 3, C and D, n = 3, P < 0.05); L-NAME and NAP failed to decrease LPS-stimulated COX-2 protein. None of the interventions alone caused induction of COX-2 gene expression and protein.


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Fig. 3.   A: representative RT-PCR showing that COX-2 gene (lane 1) expression after 8 h of incubation with LPS was suppressed by S-nitroso-N-acetypenicillamine (SNAP; lane 1), sodium nitroprusside (SNP; lane 2), and NONOate (lane 3). N-acetyl-D-penicillamine (NAP; lane 4) and NG-nitro-L-arginine methyl ester (L-NAME; lane 5) had little effect on LPS-stimulated gene expression. Gene expression of GAPDH is shown as a loading control. S, standard; C, control. B: densitometric assessment (COX-2 gene expression related to GAPDH) showing a significant reduction in COX-2 gene expression by SNAP, SNP, and NONOate (data are shown as a percent of LPS and represent means ± SE; n = 3, *P < 0.05). C: representative Western analysis demonstrating that the nitric oxide (NO) donors SNAP (lane 1), SNP (lane 2), and NONOate (lane 3) inhibit COX-2 gene expression after 8 h of treatment with LPS, NAP (lane 4), and L-NAME (lane 5) had little effect. D: densitometric data from Western analyses demonstrating a similar reduction in LPS-stimulated COX-2 protein in the presence of NO donors (data are shown as a percent of LPS and represent means ± SE; n = 3, *P < 0.05).

Treatment of cells with SNAP, SNP, and NONOate caused little change in the LPS-stimulated downregulation of COX-1 gene expression (n = 3-4); NAP and L-NAME also had little effect (Fig. 4A). In control cells, basal COX-1 gene expression was significantly decreased by SNAP, SNP, and NONOate but was little changed by NAP or L-NAME (Fig. 4, B and C, P < 0.05, n = 5-6). Western analysis showed a reduction in COX-1 after 8-h treatment with SNAP, SNP, NONOate, and L-NAME, whereas NAP had no significant effect (Fig. 4D).


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Fig. 4.   A: representative RT-PCR analysis demonstrating that SNAP (lane 1), SNP (lane 2), NONOate (lane 3), NAP (lane 4), and L-NAME (lane 5) had little effect on COX-1 gene expression in cells treated with LPS for 8 h. S, standard. B: representative RT-PCR analysis demonstrating that SNAP (lane 1), SNP (lane 2) and NONOate (lane 3) decreased basal or control (C) levels of COX-1 gene expression; NAP (lane 4) and L-NAME (lane 5) had little effect. C: densitometric data (COX-1 gene expression related to GADPH) showing that SNAP, SNP, and NONOate significantly suppressed basal COX-1 gene expression; NAP and L-NAME had little effect. D: densitometric data from Western blots showing that SNAP, SNP, NONOate, and L-NAME, but not NAP, caused a significant reduction in basal COX-1 protein (data are shown as a percent of control and represent means ± SE; n = 5-6, *P < 0.05).

Effect of NO donors on LPS-stimulated prostanoid production. By 8 h, LPS resulted in a significant eightfold increase in release of 6-keto-PGF1alpha (baseline, 124 ± 206 pg/ml; 8 h, 1,104 ± 576 pg/ml, n = 5, P < 0.05); release of PGE2 was not significantly increased (PGE2 baseline, 23 ± 17 pg/ml; 8 h, 37 ± 21 pg/ml). Preincubation with SNAP resulted in marked (~80%) suppression of the LPS-stimulated 6-keto-PGF1alpha release (Fig. 5). SNP and NONOate also resulted in significant attenuation of the LPS-stimulated 6-keto-PGF1alpha release, although this effect was less pronounced than that for SNAP (~60% inhibition). L-NAME and NAP had little effect on LPS-stimulated 6-keto-PGF1alpha release. None of the interventions had any significant effect on PGE2 release (data not shown). The observed inhibitory effects were not associated with a reduction of cell viability at 8 h as reflected by measurement of the MTT reaction (data not shown). None of the interventions had an effect on basal production of either 6-keto-PGF1alpha or PGE2.


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Fig. 5.   A: effect of SNAP, SNAP, and NONOate of LPS-stimulated release of 6-keto-PGF1alpha from endothelial cells following 8 h of incubation (n = 5, *P < 0.05); NAP and L-NAME had no effect of LPS-stimulated 6-keto-PGF1alpha release. CON, control. Data are shown as means ± SE; n = 5, *P < 0.05.

Effect of NO donors on COX activity. In control cells, stimulation with AA caused a significant increase in 6-keto-PGF1alpha (4,431 ± 1,942 pg/ml) and PGE2 (234 ± 20 pg/ml) release compared with unstimulated cells (6-keto-PGF1alpha , 27 ± 23 pg/ml, P < 0.001; PGE2, 14 ± 5 pg/ml, n = 3, P < 0.05). Treatment with SNAP resulted in a 75% reduction in AA-stimulated 6-keto-PGF1alpha release and an approximate 40% reduction in PGE2 release. SNP caused ~50% inhibition of 6-keto-PGF1alpha and a 30% decrease of PGE2 release. NONOate, L-NAME, and NAP had little effect on AA-stimulated 6-keto-PGF1alpha and PGE2 release (Fig. 6A).


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Fig. 6.   Effect of SNAP, SNP, NONOate, NAP, and L-NAME on COX activity as measured by release of 6-keto-PGF1alpha (open bars) and PGE2 release (solid bars) without (A) and with (B) addition of LPS for 8 h. Data are shown as a percent of baseline in A and as a percent of LPS in B and represent means ± SE; n = 3, *P < 0.05.

LPS caused a significant increase in AA-stimulated release of both 6-keto-PGF1alpha (8-h LPS, 5,617 ± 1,956 pg/ml; control, 851 ± 439 pg/ml, P < 0.001) and PGE2 (LPS, 376 ± 93 pg/ml; control, 32 ± 18 pg/ml; n = 3, P < 0.001). Of the NO donors, SNAP significantly reduced the LPS/AA-stimulated increase in both 6-keto-PGF1alpha and PGE2 release (65 and 50%, respectively, n = 3), NONOate significantly reduced 6-keto-PGF1alpha by 35% (P < 0.05) and SNP significantly reduced PGE2 release by ~20% (P < 0.05, Fig. 6B). L-NAME and NAP did not exert any significant inhibitory effect on LPS/AA-stimulated prostanoid release.

Effect of NO donors on NO release from endothelial cells. After a 2-h treatment with SNAP and NONOate, NO release was significantly elevated compared with untreated controls (SNAP, 14.5 ± 3.2 µM; control, 7.8 ± 3.2; NONOate, 22.8 ± 3.5; control, 14.3 ± 2.4; n = 5, P < 0.05). Treatment with SNP did not cause an increase in NO release (SNP, 8.2 ± 2.5; control, 9.3 ± 2.8). LPS either alone or after preincubation with the NO donors caused no further increase in NO release.

Effect of NO donors on cGMP production in RFL-6 reporter cells. Treatment with each of the NO donors resulted in a significant increase in cGMP levels in RFL-6 cells (Fig. 7). For both SNAP and NONOate, the increase at 60, 120, and 150 min was similar. Treatment with SNP resulted in a significantly smaller increase at 60 min, reached a level similar to that of SNAP and NONOate at 120 min, and by 150 min had declined and was again significantly less than SNAP and NONOate (Fig. 7). Neither NAP nor L-NAME had any effect on cGMP levels in the RFL-6 cells, remaining at a basal value of 0.03 fmol/µg protein at each time studied.


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Fig. 7.   Temporal effect of SNAP (solid bars), NONOate (hatched bars), and SNP (crosshatched bars) on cGMP levels in RFL-6 reporter cells. Open bars, untreated RFL-6 cells. Each of the donors was incubated at 37°C, and at 60, 120, and 150 min, an aliquot was taken for assay. Data are shown as means ± SE; n = 4, *P < 0.05 compared with SNAP and NONOate. +P < 0.05 compared with each of the NO donors.


    DISCUSSION
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

The present study demonstrates that increased release of prostaglandins by BPAEC in response to LPS is accompanied by rapid induction of COX-2 gene expression and COX activity and a gradual reduction in expression of constitutive COX-1. Pretreatment of cells with the NO donors SNAP, SNP, and NONOate resulted in suppression of LPS-stimulated 6-keto-PGF1alpha release and blunted the induction of COX-2 gene expression and protein; SNAP was the most effective inhibitor. Incubation with either SNAP or NONOate also led to a significant increase in basal release of NO from BPAEC; this elevation was unaffected by LPS. Suppression of endogenous production of NO through inhibition of NOS had little effect on LPS-stimulated prostaglandin production. In control cells, exogenous generation of NO downregulated expression of COX-1 and its activity, but the donors had no detectable effect on expression of COX-2.

It is generally accepted that under normal conditions COX-1 regulates prostaglandin production and thereby affects basal vascular tone and normal cell activity. The increased release of NO after incubation with NO donors and the subsequent downregulation of COX-1 gene expression and protein suggest that NO also contributes to the maintenance of vascular tone and endothelial cell function. Because both NO and prostacyclin are vasodilators and inhibit platelet aggregation, it is possible that each is a "backup" mechanism for the other. Our finding that L-NAME downregulates basal COX-1 protein suggests that constitutive generation of NO may modulate basal prostaglandin production. COX-2, on the other hand, is induced by inflammatory stimuli and synthesizes prostaglandins, which modulate vascular tone and mediate the inflammatory process and/or tissue damage. LPS is known to stimulate prostanoid production in a number of cells through an effect exerted on COX-2 (1, 9, 13). Our data not only confirm that prostanoid production during inflammation is COX-2 dependent but also demonstrate that COX-1 and COX-2 are differentially and disparately regulated by LPS.

Our data demonstrate that exogenously generated NO suppresses the LPS-stimulated induction of 6-keto-PGF1alpha release, and the induction of COX-2 mRNA and protein expression. We were unable to detect any significant effect of the NO donors on PGE2 release, presumably because the level of this prostaglandin was low at this early time point of LPS stimulation; by 24 h of LPS, PGE2 release was significantly elevated (16). It has been shown recently that exogenously generated NO upregulates NOS mRNA in BPAEC and in neonatal sheep pulmonary artery endothelial cells (5, 15, 33). We now demonstrate increased release of NO from cells incubated with NO donors. These findings are consistent with the notion that extracellular generation of NO results in increased levels of intracellular NOS and NO and suggest that NO regulates prostanoid production through an effect on COXs that occur at the transcriptional level. Recent data also show that NO may modulate prostacyclin production through an effect on prostacyclin synthase (13). This latter finding may contribute to our observation that the NO donors have a greater effect on prostacyclin synthesis than on PGE2 formation.

Increasing evidence indicates that NO modulates prostanoid production by COX-2 both at the transcriptional level and through an effect on enzyme activity (6, 14, 25, 27, 30, 31). Our results demonstrate that COX activity is significantly inhibited by the NO donors SNAP, SNP, and NONOate, with SNAP being the most effective, regulating both 6-keto-PGF1alpha and PGE2 release. Because neither NAP (an inactive analog of SNAP) nor L-NAME affected COX activity, it is likely that increased intracellular levels of NO play a role in the loss of COX activity. Our findings indicate that suppression of prostacyclin production by SNAP also involves an effect of NO at the posttranscriptional level.

The higher level of released NO from the endothelial cells after treatment with SNAP and NONOate is consistent with the notion that the level of intracellular NO is important to the suppression of LPS-induced COX activity. Individual classes of NO donors have different chemical reactivities and kinetics for NO release (8). Although we were unable to detect any increase in NO release from the endothelium after 2 h following treatment with SNP, we have recently shown in a cell-free system that SNP gives a rise in concentration of NO by 10 min, which gradually declines over the 35 min of the study; this increase in NO is strikingly less than that from either SNAP or NONOate (5). The small increase in NO release by SNP in this cell-free system is likely explained by the fact that SNP requires tissue for NO release to occur (3). Our finding of an increase in cGMP in a reporter cell line following exposure to SNP confirms that notion. A recent study has demonstrated that release of NO from SNP may be dependent on membrane-bound NADH oxidoreductase (18). It is likely that intracellular generation of NO by SNP may explain, at least in part, the protective effects of SNP on the LPS-induced changes. The reason for our failure to detect an increase in NO release from the endothelial cells after exposure to SNP is not obvious but may reflect a difference in its site of generation. The intracellular generation of NO by SNP is likely to be governed by local enzymes such as NADH oxidoreductase, and its release is likely to be positioned at critically important loci for immediate utilization. Generation of extracellular NO, on the other hand, is likely to be in excess of local requirements, and subsequent stimulation of NOS may result in an overabundance of intracellular NO and its release. The lack of effect of NONOate on basal COX activity in normal cells is more difficult to explain. However, our finding that L-NAME had little effect on LPS-stimulated or basal gene expression of COXs and prostanoid release indicates that the normally low intracellular concentrations of NO contribute little to prostanoid synthesis; only when NO levels are increased are the effects of NO on COX-1 and COX-2 gene expression and protein apparent. Based on these data, it seems likely that an intracellular threshold of NO is needed before prostanoid synthesis is modulated.

Other studies have reported an inhibitory action of NO on prostanoid synthesis and COX activity. For example, NO has been shown to inhibit prostanoid production by macrophages, Kupffer cells, and chondrocytes activated by inflammatory stimuli to express COX-2 activity (27-29). However, in contrast, several studies in the literature indicate that NO stimulates the activity of COX in various cell types, including endothelial cells (6, 25, 30, 32). How can these apparently conflicting results be explained? One possibility is that different levels of NO cause different responses in the COX system. For example, in rabbit articular chondrocytes, NG-monomethyl-L-arginine was found to inhibit PGE2 release when stimulated with interleukin-1beta (IL-1beta ; low production of NO) but to increase PGE2 production when the cells were stimulated concurrently with IL-1beta , LPS and tumor necrosis factor-alpha (high production of NO) (28).

In summary, our data demonstrate that LPS results in induction of COX-2 gene and protein expression and an increase in COX-2 activity; this is accompanied by a gradual reduction in COX-1 expression. Exogenous generation of NO results in an increased release of NO from BPAEC and regulates constitutive COX-1 gene and protein expression. Exogenous generation of NO also suppresses LPS-stimulated prostacyclin release and attenuates COX-2 expression both at the level of gene transcription and by a posttranscriptional mechanism. Intracellular generation of NO by SNP also inhibits LPS-induced prostaglandin synthesis. However, inhibition of endogenous NOS has little effect on LPS-stimulated prostaglandin synthesis. We conclude that increased levels of intracellular NO regulates basal and LPS-stimulated prostaglandin production. We speculate that the vasodilator effects of inhaled NO may have limited beneficial effects when administered chronically to patients with acute respiratory distress syndrome and pulmonary hypertension since the net result may be to downregulate synthesis and release of the vasodilator prostacyclin. Furthermore, administration of NO to normal lung may affect regulation of normal pulmonary vascular tone by modulation of COX-1.


    ACKNOWLEDGEMENTS

This work was supported by the National Heart, Lung, and Blood Institute Grant HL-55649.


    FOOTNOTES

Address for reprint requests and other correspondence: B. Meyrick, Center for Lung Research, Vanderbilt Univ. Medical Center, MCN T-1217, Nashville, TN 37232-2650 (E-mail: barbara.meyrick{at}mcmail.vanderbilt.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.

Received 3 March 2000; accepted in final form 13 October 2000.


    REFERENCES
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

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