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GASTROINTESTINAL, HEPATIC, PULMONARY, AND RENAL
Department of Pharmacological and Physiological Science, Saint Louis University School of Medicine, St. Louis, Missouri
Received May 15, 2006; accepted January 30, 2007.
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Abstract |
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; Zhu et al., 2000). Of these regioisomers, 5,6-EET has been reported to be the most abundant metabolite synthesized in rabbit lung (Zeldin et al., 1995), and it is the most potent pulmonary vasoconstrictor (Zhu et al., 2000). Although EETs relax most blood vessels in the systemic circulation, they contract intralobar segments of pulmonary arteries (PA) (Schwartzman et al., 1985; Zhu et al., 2000; Stephenson et al., 2003; Losapio et al., 2005) and increase pulmonary vascular resistance in isolated perfused lungs (Stephenson et al., 2003). Recently, the importance of this finding was illustrated in a report by Pokreisz et al. (2006), who demonstrated that inhibition of epoxygenase activity significantly reduced acute hypoxic pulmonary vasoconstriction (HPV) and chronic hypoxia-induced pulmonary vascular remodeling in a mouse model of hypoxia-induced pulmonary hypertension. Their findings give EETs a physiological role as participants in HPV and may represent an important target for therapeutic intervention in pulmonary hypertension. Because synthesis of EETs in perfused human lung exceeds synthesis of pulmonary vasoconstrictor leukotrienes and hydroxyeicosatetraenoic acids (Kiss et al., 2000), information that improves our understanding of EET-mediated pulmonary vascular contraction may be valuable for developing a rational therapeutic approach to treat hypoxia-induced pulmonary hypertension.
Nonselective inhibition of cyclooxygenase (COX) activity inhibits 5,6-EET-induced pulmonary vascular contraction (Schwartzman et al., 1985; Zhu et al., 2000; Stephenson et al., 2003; Losapio et al., 2005), suggesting that either 5,6-EET can be metabolized by COX activity (Oliw, 1984) to a pulmonary vasoconstrictor or that 5,6-EET stimulates endogenous arachidonic acid release, which is then metabolized by COX to a pulmonary vasoconstrictor compound (Zhu et al., 2000; Stephenson et al., 2003). Of the COX isoforms identified, both COX-1 and COX-2 (Xie et al., 1991) were reported to be expressed constitutively in pulmonary vessels of the rat and mouse (Ermert et al., 1998; Baber et al., 2003, 2005). COX-1 is constitutively expressed in many tissues (Vane and Botting, 1998), whereas COX-2, often associated with inflammation, usually requires induction by cytokines (Hempel et al., 1994). Constitutively expressed COX-2 was reported to contribute to arachidonic acid-induced contraction of PA (Baber et al., 2003, 2005). However, it is not known which COX isoform contributes to EET-induced pulmonary vasoconstriction. If both COX isoforms are expressed in isolated rabbit pulmonary arteries, it is of considerable importance to determine whether COX-1, COX-2, or both isoforms are required for 5,6-EET-induced pulmonary vasoconstriction (Zhu et al., 2000; Stephenson et al., 2003) to fully understand the mechanism by which 5,6-EET contracts PA.
In the present study, we report that COX-1 and COX-2 isoforms are expressed in freshly isolated PA of healthy rabbits. To determine which isoform is required for 5,6-EET-induced contraction of rabbit PA, we examined the tissue distribution and activities of each COX isoform in second-order (first branches distal to the main PA) intralobar PA segments (1.98 ± 0.09 mm outside diameter) by selective pharmacological inhibition of their activities. In addition, we examined whether preferential metabolism of 5,6-EET by either COX isoform might explain differential effects of selective COX isoform inhibition on 5,6-EET-induced PA contraction.
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Materials and Methods |
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Isolated PA Segment Protocol. Intralobar PA were dissected free of extravascular tissue, cut into rings 3 to 4 mm in length, and suspended in water-jacketed tissue chambers containing physiological salt solution (PSS) composed of 118.3 mM NaCl, 4.7 mM KCl, 2.5 mM CaCl2, 1.2 mM MgSO4, 1.2 mM KH2PO4, 25 mM NaHCO3, 0.026 mM Na-EDTA, and 11.1 mM glucose. The PSS was gassed with 95% O2, 5%CO2, pH 7.4, and maintained at 37°C as described previously (Stephenson et al., 2003). Each ring was mounted between two stainless steel support wires. Ring tension was measured from one of the support wires attached to an isometric force transducer (FT03; Astro-Med, West Warwick, RI) and recorded continuously on a polygraph (Astro-Med). Each ring was placed under a basal tension (0.75–1.5 g) determined to result in a maximal contractile response to 60 mM KCl. Before contraction with 5,6-EET, 1 to 5 x 10–6 M prostaglandin (PG)F2
was added to achieve a contraction that was 50 to 80% of that produced with 60 mM KCl.
In some experiments, PA rings were preincubated with inhibitors of COX activity. The selective COX-2 inhibitors DUP-697 at 100 nM and NS-398 at 3 µM, the selective COX-1 inhibitor SC-560 at 300 nM, and the nonselective COX inhibitor indomethacin at 10 µM (Landreth et al., 1994; Smith et al., 1998) were used to inhibit COX activity. To ensure that inhibition of COX-2 activity with DUP-697 did not also inhibit COX-1 activity, we used rabbit platelet thromboxane synthesis as an index of COX-1 inhibition. Because platelets express only COX-1 activity (Funk et al., 1991; Patrignani et al., 1994), any decrease in platelet thromboxane synthesis with DUP-697 would indicate inhibition of COX-1 activity. We determined that in isolated rabbit platelets, the selective COX-1 inhibitor SC-560 at 300 nM decreased A23187
[GenBank]
-stimulated thromboxane synthesis >98%, whereas the selective COX-2 inhibitor DUP-697 at 100 nM did not reduce A23187
[GenBank]
-stimulated thromboxane synthesis. However, concentrations of DUP-697 higher than 300 nM (1 x 10–6–1 x 10–5 M) decreased platelet thromboxane synthesis, demonstrating loss of COX-2 selectivity at these higher concentrations.
5,6-EET was added cumulatively (1.4 x 10–8–1 x 10–5 M) to intralobar rings at basal tension 30 min after incubation with inhibitor or vehicle. EETs and COX isoform inhibitors were dissolved in either ethanol or dimethyl formamide. At the concentrations used (0.01%), neither drug vehicle altered the basal tension or contractile responses to 5,6-EET. Since the presence of intact endothelium has been reported to be necessary for EET-mediated contraction in smaller pressurized PA segments (243–510-µm internal diameter) (Zhu et al., 2000), in some PA rings, the endothelium was removed by rubbing the intimal surface with a cotton applicator tip to determine whether endothelium was necessary for 5,6-EET-induced contractions in PA rings of the size used in the present study. The presence or absence of endothelium in all intralobar PA used in the present study was confirmed by relaxation of the PGF2-contracted rings in response to the addition of acetylcholine (1 x 10–6 M).
Preparation of 5,6-EET and 14C-Labeled 5,6-EET. Unlabeled 5,6-EET for isolated vessel studies was prepared according to the method of Corey et al. (1979) as described previously (Stephenson et al., 1998). The same method was used for preparation of [14C]5,6-EET with modifications. In brief, 10 µCi of [14C]arachidonic acid (American Radiolabeled Chemicals, St. Louis, MO) was reacted with potassium triiodide (0.4 Eq) in the presence of 0.25 Eq of potassium bicarbonate in 100 µl of tetrahydrofuran/water (2:1) overnight at 4°C. After overnight incubation, excess iodine was removed by adding an aqueous solution of saturated sodium sulfite. The iodolactone formed was extracted three times with 1 ml of hexane, dried under N2 gas, and reacted with 0.2 N lithium hydroxide in 1.5 ml of tetrahydrofuran/water (2:1) for 3 h at room temperature. The [14C]5,6-EET formed was extracted three times with acidified ethyl acetate and purified by reverse-phase HPLC using a C18 column (5 µm, 4.6 x 250 mm, Nucleosil; Phenomenex, Torrance, CA) with a linear gradient from 50% water in acetonitrile/acetic acid (999:1) to 100% acetonitrile/acetic acid (999:1) over 40 min at 1 ml/min. Eluate containing 5,6-EET was collected, evaporated to dryness, and stored under N2 gas in hexane at –80°C. The identity of the [14C]5,6-EET synthesized was obtained by comparing the HPLC retention time of the radioactive peak (Radiomatic detector; PerkinElmer Life and Analytical Sciences, Boston, MA) with the retention time of an authentic 5,6-EET standard (Cayman Chemical, Ann Arbor, MI), determined by its ultraviolet absorbance at 204 nm.
Metabolism of [14C]5,6-EET by COX-1 and COX-2. Metabolism of [14C]5,6-EET by cyclooxygenase was examined by incubating 2.6 or 14.2 µg of purified COX-1 (from ram seminal vesicles; Cayman Chemical) or COX-2 (obtained from sheep placenta; Cayman Chemical) with 250 nmol of [14C]5,6-EET (
30 nCi) at 37°C in 100 mM Tris-HCl buffer solution, pH 8.0, containing 2 mM phenol and 1 µM hematin (Smith and Marnett, 1991; Johnson et al., 1995). In some incubations, selective inhibition of COX isoforms was examined by the addition of either SC-560 or DUP-697 to the reaction mixtures 30 min before the addition of [14C]5,6-EET. Reactions were terminated at 2 min by the addition of ice-cold acidified acetonitrile. The HPLC solvent protocol for identification of 5,6-EET metabolites began isocratically with acetonitrile/water/acetic acid (33:67:0.01) (solvent A) for the first 16 min followed by a linear gradient from 100% solvent A to 65% acetonitrile/water/acetic acid (90:10:0.01) (solvent B) over 10 min. This solvent mixture of 35% solvent A, 65% solvent B was maintained until the end of the 50-min HPLC run was completed (Sola et al., 1992). Metabolism of [14C]5,6-EET by COX-1 and COX-2 enzymes was analyzed by comparing the integrated areas under the radioactive metabolite peaks in the presence or absence of COX enzymes.
Western Blot Analysis. Intralobar PA vessels similar to those used for contraction studies were obtained as described above. In some cases, the endothelium was removed from vessels by rubbing the intimal surface with a cotton applicator tip. A portion of intact or endothelium-denuded vessel was silver stained as described by Pawlowski et al. (1988) to confirm that endothelium had been removed by rubbing. In brief, vessels were washed with a 5% glucose solution followed by incubation with 0.25% AgNO3 in 5% glucose solution for 5 min. After removing the AgNO3 and washing with 5% glucose, vessels were incubated with a solution containing 3% CoBr2 and 1% NH4Br for 2 min. After removal of the bromide solution, the vessels were again washed with a 5% glucose solution, and then they were examined by light microscopy for the presence of endothelium.
For immunoblots, vessels were homogenized with a polytron-type tissue homogenizer in ice-cold lysis buffer containing 25 mM HEPES, 300 mM NaCl, 10 mM EDTA, 1.5 mM MgCl2, 20 mM 
-glycerophosphate, and 0.1 mM sodium vanadate with a cocktail of protease inhibitors (Complete; Roche Diagnostics, Indianapolis, IN). After a 30-min incubation with lysis buffer (on ice), samples were centrifuged at 20,000g for 15 min at 4°C. Total protein content of each sample supernatant was determined by the bicinchoninic acid assay (Pierce Chemical, Rockford, IL). Supernatants were diluted in sample buffer containing 2% SDS, 15% glycerol, 100 mM dithiothreitol, 62.5 mM Tris, and 0.01% bromphenol blue at pH 6.8. The samples were boiled for 5 min and equal amounts (10 µg) of sample protein were loaded on to a 4 to 20% SDS polyacrylamide gel (Ready Gel; Bio-Rad, Hercules, CA), resolved by electrophoresis at 150 V for 1 h, and transferred to polyvinylidene fluoride (PVDF) membranes. Defined concentrations of COX-1 and COX-2 electrophoresis standards (Ovine; Cayman Chemical) were included on all sample gels. The PVDF membranes were blocked overnight at 4°C in a buffer solution (Tris-buffered saline/Tween 20) containing 50 mM tris(hydroxymethyl)aminomethane, 150 mM NaCl, 0.1% Tween 20, and 5% nonfat milk or StartingBlock (Pierce Chemical) at pH 7.4. After blocking, PVDF membranes were incubated for 4 h with either goat anti-COX-1 polyclonal antibody raised against a peptide sequence corresponding to the carboxyl terminus of human COX-1 protein (Santa Cruz Biotechnology, Santa Cruz, CA) or goat anti-COX-2 polyclonal antibody raised against a peptide sequence corresponding to the amino terminus of rat COX-2 protein (Santa Cruz Biotechnology, Santa Cruz, CA). Membranes were washed with Tris-buffered saline/Tween 20 and incubated with horseradish peroxidase-conjugated anti-goat secondary antibody (GE Healthcare, Piscataway, NJ) for 1 h at room temperature. Proteins were detected with enhanced chemiluminescence (GE Healthcare) on Iso-Max autoradiography/X-ray film (SciMart, St. Louis, MO). Relative protein concentrations were analyzed by densitometry of the film image using QuantiScan, version 2.1, software (Biosoft, Ferguson, MO). In studies examining the effect of endothelium removal on COX protein expression, membranes were stripped with a proprietary stripping solution (Restore; Pierce Chemical) and reprobed for COX-1 as described above. The order of membrane stripping and reprobing, i.e., COX-2 first or COX-1 first, was examined previously and did not influence the results obtained. After measuring COX isoform protein expression, the membranes were stripped again and reprobed with a primary mouse antibody to -actin (Sigma-Aldrich, St. Louis, MO). The -actin immunoblot served as a lane-loading control.
Immunohistochemistry. Intralobar PA vessels similar to those used for contraction studies were obtained as described above. Immediately after removal of extravascular tissue, vessels were submersed in 0.1 M sodium phosphate buffer (SPB), pH 7.4, containing 4% formaldehyde at 4°C for 24 h. The vessels then were transferred to SPB at 4°C for an additional 24 h. Twenty-four hours before sectioning, the vessels were transferred to SPB containing 25% sucrose at 4°C. Vessels were embedded in Tissue-Tek OCT (Miles Inc., Elkhart, IN) frozen, and cut into 25-µm-thick sections using a Leica CM1900 cryostat (Leica Microsystems, Deerfield, IL). Vessel sections were placed on glass slides, and the slides were air-dried. After drying, slides were rinsed briefly in SPB containing 0.1% Triton X-100 (SPB-Triton), and then they were incubated overnight with either COX-1 antibody (1:1000), COX-1 antibody (1:1000) preadsorbed for 1 h with 25-fold excess of the cognate peptide, COX-2 antibody (1:500), or COX-2 antibody (1:500) preadsorbed for 1 h with 25-fold excess of the cognate peptide. The sections were then rinsed in SPB-Triton and immersed for 1 h in the same carrier solution containing a 1:200 dilution of biotinylated anti-goat IgG (Jackson ImmunoResearch Laboratories Inc., West Grove, PA). Another series of rinses preceded immersion of the sections for 1 h in the same carrier solution containing a 1:200 dilution of avidin-biotin-peroxidase complex (ABC; Vector Laboratories, Burlingame, CA), after which the sections were again rinsed and then reacted for 10 min in a nickel-diaminobenzidine solution containing 0.015% diaminobenzidine, 0.4% nickel ammonium sulfate, and 0.006% H2O2 in 0.025 M Tris-HCl buffer, pH 8.0. Sections were counterstained with eosin, coverslipped using Permount (Fisher Scientific, St. Louis, MO), and visualized with a Nikon (Tokyo, Japan) Eclipse E600 microscope.
Enzyme Immunoassay. To examine the activity of selective COX inhibitors on calcium ionophore (A23187
[GenBank]
)-induced PGI2 synthesis in intralobar vascular rings, enzyme immunoassay was performed for quantitative identification of 6-keto-PGF1 (the stable degradation product of PGI2) in the supernatant of rings that were preincubated with either SC-560 or DUP-697 for 30 min before a 30-min incubation with A23187
[GenBank]
at 37°C in PSS as described previously (Pradelles et al., 1985; Stephenson et al., 1998). In brief, enzymatic tracers consisted of 6-keto-PGF1 covalently linked to purified acetylcholinesterase (Pradelles et al., 1985). The sample (50 µl) was combined with 50 µl of enzymatic tracer in a well of a 96-well microtiter plate (Nalge Nunc, Naperville, IL) that had been precoated with 2 µg/well goat anti-rabbit IgG antibody (Calbiochem, San Diego, CA). We then added 50 µl of the antiserum for 6-keto-PGF1 (Cayman Chemical). The plates were incubated for 18 to 20 h at room temperature and washed three times with 500 µl of 1 x 10–2 M potassium phosphate buffer, pH 7.4, containing 0.05% Tween 20. After washing, 200 µl of Ellman's reagent was added to each well. Ellman's reagent consisted of 2 µg/ml acetylthiocholine iodide and 2.15 µg/ml 5,5'-dithiobis(2-nitrobenzoic acid) in 1 x 10–2 M potassium phosphate buffer. The reaction product [reduced 5,5'-dithiobis(2-nitrobenzoic acid)] was monitored at 405 nm in a Bio-Tek model EL-309 enzyme immunoassay plate reader (Bio-Tek Instruments (Winooski, VT). All samples and standards were run in duplicate. Sample unknowns were determined by comparison with standards with log-logit data transformation.
Statistical Methods. All values are expressed as means ± S.E. Differences between experimental groups were determined by analysis of variance. If the F ratio indicated significant differences, a Fisher's least significant difference (protected t) test was performed to establish differences between individual sample means. Values of P < 0.05 were considered to be statistically significant.
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Effects of Cyclooxygenase Inhibition on 5,6-EET-Induced Contraction in PA Rings. At basal tension, 5,6-EET increased active tension in intralobar PA rings with intact endothelium in a concentration-dependent manner (Fig. 3). Intralobar PA in which the endothelium had been removed did not contract in response to 5,6-EET (Fig. 4), but it contracted normally to PGF2, a COX-independent vasoconstrictor. Pretreatment of endothelium-intact PA rings with the nonselective COX inhibitor indomethacin at 10 µM for 30 min also prevented contraction at all 5,6-EET concentrations examined (Fig. 3). These results confirm that 5,6-EET-induced contraction is dependent on intact endothelium and cyclooxygenase activity in second-order intralobar PA.
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; Carroll et al., 1993), metabolism of 5,6-EET by the COX-2 isoform has not been reported previously. If COX-2 does not metabolize 5,6-EET or if the metabolites produced are different from those produced by COX-1, selective inhibition of COX-2 may be less effective at reducing 5,6-EET-induced contraction of rabbit intralobar PA than selective inhibition of COX-1. Incubation of [14C]5,6-EET without addition of either COX-1 or COX-2 isoforms, under conditions identical to those in which the enzymes were included, resulted in nonenzymatic hydrolysis of [14C]5,6-EET to the diol, 5,6-dihydroxy-epoxyeicosatrienoic acid and its 
-lactone as described previously (Capdevila et al., 1981) (Fig. 8A). When incubated with either purified COX-1 (Fig. 8B) or COX-2 (Fig. 8D) enzymes, metabolites of [14C]5,6-EET eluted from HPLC with a pattern consistent with that described previously for COX-1 metabolism of 5,6-EET (Oliw, 1984). The least polar of the metabolites eluted at about 24 min, consistent with the elution of 5,6-epxoy-PGE1. The most polar metabolite eluted at about 9 min, consistent with nonenzymatic hydrolysis of the 5,6-epoxy-PGE1 to 5,6-dihydroxy-PGE1. A metabolite of intermediate polarity, eluting at about 21 min, is consistent with the elution of the -lactone. The elution times for 5,6-EET metabolites of purified COX-2 enzyme were not different from those of COX-1 (Fig. 8, B and D). Analysis of the HPLC peak areas revealed that during a 2-min incubation of 5,6-EET with COX-1, COX-1 metabolized 0.8 ± 0.2 µmol [14C]5,6-EET · µmol COX-1 protein–1, whereas COX-2 metabolized 0.5 ± 0.3 µmol [14C]5,6-EET · µmol COX-2 protein–1 (n = 3). After a 30-min preincubation with SC-560, COX-1 metabolism of 5,6-EET was greatly attenuated (Fig. 8C). Likewise, DUP-697 inhibited nearly all of the COX-2 metabolism of 5,6-EET (Fig. 8E), suggesting that at the concentrations used, these selective COX inhibitors effectively prevented COX-mediated metabolism of 5,6-EET.
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Discussion |
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TopAbstractMaterials and MethodsResultsDiscussionReferences |
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; Smith, 2001). However, COX-2 as well as COX-1 was readily detectable in Western immunoblots of intralobar PA in healthy rabbits in the present study. Similar findings were reported previously for rat and mouse pulmonary arteries (Ermert et al., 1998; Baber et al., 2005). A variety of other tissues have also been reported to express the COX-2 enzyme constitutively (O'Neill and Ford-Hutchinson, 1993; Watkins et al., 1999). Determination of the absolute amounts of COX-1 and COX-2 isoforms present in rabbit PA cannot be made directly from the Western immunoblot results in the present study because the epitope-binding capacities of the polyclonal primary antibodies are likely to be different for each isoform as well as different for rabbit samples and sheep standards.
Previously, we reported that 5,6-EET increased pulmonary vascular resistance in isolated perfused rabbit lungs (Stephenson et al., 2003). The 5,6-EET-induced increases in pulmonary vascular resistance were associated with increases in prostacyclin and thromboxane and were inhibited by the nonselective COX inhibitor indomethacin. Those results, along with the results of the present study in which indomethacin inhibited 5,6-EET-induced contraction in second order, intralobar PA (800- to 1500-µm outside diameter) support previous studies in smaller (300–500 µm) pressurized rabbit PA segments (Zhu et al., 2000) in which 5,6- and 14,15-EET were reported to contract PA in a cyclooxygenase- and endothelium-dependent manner. Because our studies of intralobar pulmonary arteries described responses identical to those of isolated resistance vessels and intact lung vasculature, isolated vessel rings obtained from rabbit intralobar PA vessels were used as the model for examining potential mechanisms of 5,6-EET-induced pulmonary vasoconstriction in the rabbit pulmonary circulation (Losapio et al., 2005) and in the present study.
In most vascular beds studied, EETs have been reported to be vasodilators (Harder et al., 1995; Roman, 2002). However, EETs have been reported to contract the vasculature of rat kidney (Takahashi et al., 1990; Fulton et al., 1996). In the rat kidney, the vasoconstrictor effect of EETs was also reported to be COX-dependent (Takahashi et al., 1990; Imig et al., 1996), suggesting that the dependence of EET-mediated vascular contraction on COX activity may be a common feature of all EET-mediated vascular contraction.
Because both COX isoforms were expressed in intralobar PA of healthy rabbits, we examined the contribution of each COX isoform to 5,6-EET-induced PA contraction using selective pharmacological inhibitors of the two isoforms. Treatment of intralobar PA rings with the selective COX-1 inhibitor SC-560 prevented 5,6-EET-induced contraction of intralobar PA rings, demonstrating that inhibition of COX-1 activity is sufficient to abolish the 5,6-EET-induced contraction. Inhibition of COX-2 activity with either DUP-697 or NS-398 was considerably less effective at decreasing 5,6-EET-induced PA contraction than selective inhibition of COX-1 activity. These data are supported by those in which SC-560 was capable of inhibiting PGI2 synthesis in isolated rabbit PA segments, whereas DUP-697 was not (Fig. 7). These findings differ from studies reported for the rat (Baber et al., 2003) and mouse (Baber et al., 2005) in which selective inhibition of either COX-1 or COX-2 inhibited arachidonic acid-induced increases in pulmonary artery pressure to a similar extent.
In an immunohistochemical study of rat pulmonary tissue, Ermert et al. (1998) reported that, in rats (Ermert et al., 1998), COX-2 was localized primarily in vascular smooth muscle cells of the small pulmonary arteries and veins, whereas COX-1 was identified primarily in the endothelium. Since in rabbits, 5,6-EET-induced PA contraction was endothelium-dependent, localization of COX-1 predominantly to the vascular endothelium could explain the endothelium-dependence of 5,6-EET-induced contraction and the ability of selective COX-1 inhibition to prevent 5,6-EET-induced contraction. In Western immunoblots, we found that greater than 50% of both COX-1 and COX-2 expression was lost in endothelium-denuded PA vessels. Moreover, both COX isoforms were predominantly localized to the endothelium when analyzed immunohistochemically, with minor immunostaining of the adventitia. Unlike the rat, in which COX-2 was identified in pulmonary vascular smooth muscle (Ermert et al., 1998), there was little or no immunostaining of either COX isoform in the vascular smooth muscle of rabbit PA. It is unlikely that COX expressed within the adventitia contributes to PA contraction. Therefore, localization of COX activity to the endothelium may explain why 5,6-EET-induced PA contraction is endothelium-dependent. The sparse, nonuniform COX-2 immunostaining that seemed to be absent from much of the endothelium and limited to the nuclear or perinuclear regions of the endothelial cells differed markedly from the dense uniform COX-1 immunostaining of the endothelium. A predominantly nuclear or perinuclear pattern of COX-2 immunostaining has been reported previously in vascular endothelial cells (Morita et al., 1995; Parfenova et al., 2001). Parfenova et al. (2001) proposed that nuclear/perinuclear localization of COX-2 in unstimulated endothelium may influence its function because they observed increased product formation with increased expression of COX-2 in the cytoplasm of cytokine-treated endothelial cells. In the present study, the sparse, nuclear or perinuclear localization of COX-2 may limit its activity such that the COX-2 identified may be insufficient to support 5,6-EET-induced contraction.
5,6-EET has been reported to be metabolized by COX-1 (Oliw, 1984), suggesting that a COX-dependent metabolite of 5,6-EET could be responsible for 5,6-EET-induced contraction of rabbit PA. If both COX isoforms identified in rabbit intralobar PA are active and present at concentrations sufficient to produce physiologically relevant 5,6-EET metabolite concentrations, then either COX isoform could metabolize 5,6-EET to a potential PA vasoconstrictor. Because metabolism of 5,6-EET by purified COX-2 had not been reported previously, we proposed that an alternate explanation for the failure of selective COX-2 inhibition to appreciably inhibit 5,6-EET-induced PA contraction would be that COX-2 is incapable of metabolizing 5,6-EET. Here, we identify that purified COX-2 metabolizes [14C]5,6-EET to products with identical HPLC retention times to those produced by metabolism of [14C]5,6-EET by COX-1. Selective inhibition of COX-1 and COX-2 with 300 nM SC-560 and 100 nM DUP-697, respectively, prevented 5,6-EET metabolism, indicating that these inhibitors were active at the concentrations used to inhibit PA contraction. Because COX-1 and COX-2 were observed to metabolize similar amounts of [14C]5,6-EET during a 2-min incubation, differential metabolism of 5,6-EET by COX isoforms does not explain the greater dependence of 5,6-EET-induced contraction on COX-1 activity. Dependence of 5,6-EET-induced PA contraction on its metabolism by COX has been considered previously, but it has not been firmly established. 5,6-EET can also stimulate metabolism of endogenous arachidonic acid by COX (Carroll et al., 1993; Sakairi et al., 1995). Therefore, in lieu of metabolism of 5,6-EET by a COX isoform, in rabbits, 5,6-EET may stimulate synthesis of an endogenous arachidonic acid-derived PA vasoconstrictor, such as the endoperoxide PGH2. The latter mechanism was favored by Zhu et al. (2000) when they were unable to identify any indomethacin-inhibitable products of EET metabolism in peripheral rabbit lung microsomes incubated with [14C]5,6-EET. However, since metabolism of 5,6-EET by both COX isoforms was observed in the present study, synthesis of a pulmonary vasoconstrictor metabolite of 5,6-EET by either COX isoform as a mechanism of 5,6-EET-induced PA contraction cannot be ruled out.
In summary, we have identified expression of COX-1 and COX-2 isoforms in intralobar PA of healthy rabbits. The expression of both isoforms is located mainly in the PA endothelium with little or none present in the smooth muscle. We demonstrated that the cyclooxygenase-dependent contraction of intralobar PA stimulated by 5,6-EET in the rabbit is primarily dependent on the activity of the COX-1 isoform and is also endothelium-dependent. Selective inhibition of COX-2 had only a small effect on 5,6-EET-induced contraction compared with that of COX-1 inhibition, consistent with the sparse and nonuniform distribution of COX-2 within the vascular endothelium. Our results suggest that, in the rabbit, the primary dependence of 5,6-EET-induced PA contraction on COX-1 activity cannot be explained by differential metabolism of 5,6-EET by COX-1 and COX-2 but that it may result from a denser, uniform expression of the COX-1 isoform in the rabbit PA endothelium. In the rabbit PA, COX-2 expression seemed to be absent from much of the endothelium, and where present, it seemed limited to the nuclear and perinuclear regions of the endothelial cells.
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Acknowledgements |
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Footnotes |
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Article, publication date, and citation information can be found at http://jpet.aspetjournals.org.
ABBREVIATIONS: EET, epoxyeicosatrienoic acid; PA, pulmonary artery; HPV, hypoxic pulmonary vasoconstriction; COX, cyclooxygenase; PSS, physiological salt solution; PG, prostaglandin; DUP-697, 5-bromo-2-(4-fluorophenyl)-3-(4-methylsulfonyl) thiophene; NS-398, N-(2-cyclohexyloxy-4-nitrophenyl) methanesulfonamide; SC-560, 5-(4-chlorophenyl)-1-(4-methoxyphenyl)-3-(trifluoromethyl)-1H-pyrazole; A23187 [GenBank] , calcimycin; HPLC, high-performance liquid chromatography; PVDF, polyvinylidene fluoride; SPB, sodium phosphate buffer.
Address correspondence to: Dr. Alan H. Stephenson, Department of Pharmacological and Physiological Science, 1402 S. Grand Blvd., St. Louis, MO 63104. E-mail: stephens{at}slu.edu
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