Divisions of Pulmonary and Critical Care Medicine and Cardiology, The Johns Hopkins University School of Medicine, Baltimore, Maryland 21224
Submitted 10 October 2002 ; accepted in final form 24 March 2003
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ABSTRACT |
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vascular smooth muscle; endothelium; electron paramagnetic resonance; dichlorofluorescein; lucigenin; superoxide dismutase; catalase
None of these proposals includes endothelial cells, which can be a potent source of ROS and other vasoactive factors (7, 29), and are thought to play an essential role in HPV (52). In small distal porcine pulmonary arteries, we found that HPV was 1) similar to HPV in isolated or intact lungs in terms of magnitude, O2 dependence, temporal characteristics, and inhibition by Ca2+-free conditions or nifedipine; 2) abolished by endothelial denudation or the endothelin-1 (ET-1) receptor antagonist BQ123; and 3) restored in endothelium-denuded arteries primed by exposure to a threshold concentration of ET-1 (28). Furthermore, in myocytes isolated from these vessels, ET-1 priming potentiated a small but significant hypoxic contraction nearly eightfold (44). On the basis of these results and observations that ET-1 receptor antagonists inhibited HPV in intact animals (38), we concluded that full expression of HPV required basal release of ET-1 from endothelium to facilitate mechanisms of hypoxic reactivity in pulmonary arterial smooth muscle; however, the role of ROS remained unclear. In this context, we found it interesting that ROS altered production of ET-1 in endothelial cells (31), whereas ET-1 altered production of ROS in vascular smooth muscle (55, 56).
The present study was performed to determine whether ROS are required for HPV, how ROS in pulmonary arteries are altered by hypoxia, and whether the cellular site of ROS production and action during HPV is pulmonary arterial smooth muscle or endothelium.
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METHODS |
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Male pigs (2025 kg) were anesthetized with ketamine (30 mg/kg im) followed by pentobarbital sodium (12.5 mg/kg iv). After exsanguinating the animal of the femoral arteries, we rapidly removed the lungs and placed them in ice-cold Krebs-Ringer bicarbonate (KRB) solution containing (in mM): 118.3 NaCl, 4.7 KCl, 1.2 MgSO4, 1.2 KH2PO4, 2.5 CaCl2, 25.0 NaHCO3, and 11.1 glucose. Under a dissecting microscope, 1-mm segments of pulmonary arteries [outer diameter (OD) 100150 µm] were isolated from seventh-generation branches in the top left lobe and carefully cleaned of adventitial tissue.
Measurement of vascular diameter. To measure vasomotor responses, the arterial segments were cannulated at one end with a glass micropipette mounted in a chamber filled with KRB solution equilibrated with 16% O2,5%CO2, and 79% N2 at 37°C. After securing the vessel to the pipette with a 120 nylon monofilament suture, we infused KRB solution slowly until the artery was filled. In some vessels, endothelial cells were disrupted by gently rubbing the intraluminal surface with a steel wire (OD 70 µm) followed by perfusion with 2 ml of air and 2 ml of KRB solution (perfusion pressure <5 mmHg). The other end of the vessel was cannulated with a second micropipette filled with KRB solution. Both cannulas were connected to a reservoir, which was adjusted to set transmural pressure (Ptm) at the desired value. Ptm was measured with a pressure transducer positioned at the level of the vessel lumen. The chamber was covered and the vessels superfused at 20 ml/min with recirculating KRB solution (total vol 50 ml) gassed with 16% O2, 5% CO2, and 79% N2 and maintained at 37°C. The same gas mixture flowed over the surface of the superfusate in the covered chamber. To measure vascular internal diameter (ID), the chamber was placed on the stage of an inverted microscope (Nikon TMS-F, Japan) connected to a video camera (Panasonic, CCTV camera, Japan). The vascular image was projected onto a video monitor, and ID was determined with a video dimension analyzer (Living Systems Instrumentation, Burlington, VT). ID and Ptm were recorded continuously (Gould, Cleveland, OH).
Vascular viability testing. Before experiments, arteries were allowed to equilibrate for 20 min at Ptm = 10 mmHg. Ptm was then increased to 20 mmHg and thereafter held constant. To test viability, the vessels were exposed to KCl (60 mM) and then to the thromboxane A2 agonist U-46619 (10-9 M), followed by acetylcholine (ACh, 10-6 M). After stabilization of responses (510 min), agonists were washed out by 1520 min of nonrecirculating superfusion with fresh KRB solution. Arteries in which ID decreased <30% after KCl and U-46619 were excluded. Endothelium-intact (E+) vessels were excluded if ACh reversed the U-46619 contraction by <50%. Endothelium-denuded (E-) vessels were excluded if ACh caused any vasodilatation. In some preparations, viability tests were repeated at the end of the experiment.
Exposure to hypoxia. Vasomotor responses to hypoxia were measured
in untreated control E+ pulmonary arteries (n = 15), E+ arteries
treated for 30 min with superoxide dismutase (SOD, 150 U/ml, n = 10)
or SOD + catalase (CAT, 200 U/ml, n = 6), untreated control
E-arteries (n = 15), and E-arteries treated for 30 min with ET-1
(10-10 M, n = 13) or ET-1 + SOD (n = 6), where
n is the number of animals as well as vessels. Hypoxia was produced
by decreasing O2 concentration from 16 to 4% in the gas mixture
bubbling the reservoir and flowing over the perfusate surface in the chamber.
This caused oxygen tension measured with a microelectrode (Microelectrodes,
Londonderry, NH) at the vessel to decrease rapidly from 114 ± 1 to 29
± 1 mmHg (28). Duration
of hypoxia was 30 min except in 8 of 15 untreated control E-arteries, where it
was 60 min. After hypoxia, oxygen concentration was returned to 16%. Responses
were quantified as changes in ID (ID) from values at the onset of
hypoxia. Data from control arteries and E-arteries primed with ET-1 have been
previously reported (28).
Measurement of ROS in pulmonary arteries. To assess production of ROS in E+ pulmonary arteries, we used lucigenin-derived chemiluminescence (LDCL) and electron paramagnetic resonance (EPR) spectroscopy. LDCL was measured in arterial segments isolated from five pigs as described above. Two segments, 1 mm in length, were placed into each of four stoppered test tubes containing 1 ml of KRB solution bubbled gently with 16% O2,5%CO2, and 79% N2 at 37°C. One pair of tubes served as an experimental group and the other as a control group. After 30 min, baseline production of ROS in the experimental group was assessed by adding lucigenin (bis-N-methyacidinium nitrate, 20 µM) to one of the tubes immediately before transfer to a luminometer (Lumat LB 9506; E. G. and G. Berthold, Bad Wildbad, Germany) for continuous recording of emitted light as relative light units per second. In the other test tube of the experimental pair, the gas mixture was then switched to 4% O2, 5% CO2, and 91% N2. After 30 min of exposure, LDCL in this tube was measured in the same manner. Baseline and exposure measurements were repeated in the control pair of test tubes, which were treated similarly but not exposed to hypoxia.
We performed EPR spectroscopy in E+ pulmonary artery segments isolated from seven pigs. Three arterial segments, 1 mm in length, were placed into each stoppered test tube, which contained 1 ml of KRB solution bubbled with 16% O2, 5% CO2, and 79% N2 at 37°C. After a 1-h equilibration period, the gas mixture was either switched to 4% O2, 5% CO2, and 91% N2 or not changed, and the spin trap, 5-(diethoxyphosphoryl)-5-methyl-1-pyrroline-N-oxide (DEPMPO, 40 mM) (12), was added to the tube. After a 1-h exposure period, 50-µl samples of the media bathing the vessels were taken into microcapillary tubes, which were placed in a 3-mm EPR quartz tube centered in the TM 110 cavity of a Bruker ER 300 EPR spectrometer operating at X-band. Serial acquisitions of spectra were performed at 25°C by accumulating 10 1-min scans per spectrum for up to 30 min. The following data acquisition parameters were used: microwave frequency, 9.7829 GHz; sweep width, 160 G; microwave power, 20 mol wt; modulation amplitude, 0.5 G; modulation frequency, 100 kHz; time constant 328 ms. The digital data were processed using custom-developed software (SPEX and EPRDAP) capable of component simulation, fitting, and quantification (21).
Pulmonary arterial smooth muscle cells and endothelial cells. We
used primary cultures of smooth muscle cells from porcine distal pulmonary
arteries to assess direct effects of hypoxia on production of ROS by smooth
muscle. As in our previous study
(44), we used the method of
Clapp and Gurney (6) to isolate
the cells. After removal of adventitial tissue from pulmonary artery segments
obtained as described above, we opened the vessels longitudinally and removed
endothelium by rubbing the intimal surface with a cotton swab. The cleaned
vessels were cut into small pieces and transferred to a vial filled with 10 ml
of dissociation solution containing (in mM): 0.16 CaCl2, 110 NaCl,
5 KCl, 2 MgCl2,10 HEPES, 10 NaHCO3, 0.5
KH2PO4, 0.5 NaH2PO4, 10 glucose,
0.04 phenol red, 0.49 EGTA, and 10 taurine as well as papain (3.2 U/ml),
bovine serum albumin (0.2 mg/ml), penicillin (100 U/ml), and streptomycin (0.1
mg/ml). pH was titrated to 7.4 with NaOH. We stored the vessel fragments in
this solution overnight at 4°C to allow papain to penetrate the tissue.
The next day, the tissue was transferred to fresh dissociation solution
containing dithiothreitol (0.1 mM) and equilibrated with 16% O2, 5%
CO2, and 79% N2 at 37°C (pH 6.6) to activate the
enzyme. After 20 min of digestion, the tissue was transferred to fresh
dissociation solution equilibrated with 16% O2, 5% CO2,
and 79% N2 at room temperature. Cells were dissociated by
trituration with a wide-bore pipette and plated on autoclaved glass coverslips
in petri dishes. After 3040 min to allow cell adherence, we added
growth media to the dishes and placed them in an incubator (37°C, 5%
CO2 in air). Growth medium was Clonetics SmGM-2 (BioWhittaker,
Walkersville, MD), composed of basal media (Clonetics SmBM) supplemented with
fetal bovine serum (5%), bovine insulin (5 µg/ml), human fibroblast growth
factor (2 ng/ml), human epidermal growth factor (0.5 ng/ml), gentamycin (50
µg/ml), amphotericin (50 ng/ml), streptomycin (50 µg/ml), and penicillin
(50 U/ml). When cells had grown to 50% confluence (13.5 ± 1.9
days), medium was changed to Clonetics SmBM containing 0.3% fetal bovine
serum, and the cells were maintained in this serum-starved state until study
(6.4 ± 0.4 days). Throughout the period of culture, medium was renewed
at 2- to 3-day intervals. Cells were identified as pulmonary arterial smooth
muscle cells (PASMCs) by their elongated spindle shape and positive
immunoperoxidase staining for
-actin
(15,
16). Staining was performed
with commercially available kits (Avidin/Biotin Blocking kit, Vectastain Elite
ABC reagent, and Vector SG Substrate kit; Vector Laboratories, Burlingame, CA)
using mouse anti-
-actin and biotinylated goat anti-mouse
IgG2a (Roche Molecular Biochemicals, Indianapolis, IN) as primary
and secondary antibodies, respectively.
In a few protocols (see below), we used rat PASMCs or bovine pulmonary arterial endothelial cells (PAECs). Rat PASMCs were isolated enzymatically from greater than fourth-generation pulmonary arteries obtained from 250- to 350-g male Wistar rats as previously described (46). Culture conditions were the same as for porcine PASMCs except that primary cultures were exposed to growth media for 46 days and serum starved for 12 days before study due to a faster rate of growth. Bovine PAECs frozen at passage 3 were obtained from Cell Systems (Kirkland, WA) and cultured in DMEM (Life Technologies, Rockville, MD) supplemented with 20% fetal bovine serum (Sigma, St. Louis, MO), 15 µg/ml of endothelial cell growth supplement (Upstate Biotechnology, Lake Placid, NY), 1% antibiotic-antimycotic solution (Life Technologies), and 0.1 mM nonessential amino acids (Life Technologies). When PAECs were 3040% confluent at passage 56, they were exposed to medium without serum, nonessential amino acids, and endothelial cell growth supplement for 12 days before study.
Measurement of dichlorofluorescein fluorescence. For cell perfusion and measurement of intracellular ROS, coverslips with porcine PASMCs were mounted in a closed polycarbonate chamber (RC-21BR; Warner Instrument, Hamden, CT) clamped in a heated aluminum platform (PH-2, Warner Instrument) on the stage of a laser-scanning confocal fluorescence microscope (LSM-510; Carl Zeiss, Thornwood, NY). The chamber was perfused at 0.5 ml/min with KRB solution containing 5 µM dihydro-2',7'-dichlorofluorescein diacetate (DCFH2-DA). The perfusate was equilibrated with 16 or 4% O2-5% CO2-balance N2 at 38°C in heated reservoirs and led via stainless steel tubing and a manifold to an inline heat exchanger (SF-28, Warner Instrument), which rewarmed the perfusate just before it entered the cell chamber. The temperature of the heat exchanger and chamber platform was controlled by a dual-channel heater controller (TC-344B, Warner Instrument). Perfusion of a cell chamber modified to allow continuous measurement of oxygen tension (Microelectrodes) and temperature at the coverslip confirmed maintenance of temperature at 37°C and oxygen tensions at 112 ± 2.0 and 34 ± 1.8 mmHg when reservoir O2 concentrations were 16 and 4%, respectively. After the reservoir O2 concentration was changed, PO2 in the chamber changed with a half-time of 2.6 ± 0.3 min.
Initially, porcine PASMCs were perfused under normoxic conditions for 13 h to allow DCFH2-DA uptake and stabilization of cell fluorescence. Upon cell entry, DCFH2-DA is cleaved by cellular esterases to DCFH2. DCFH2 remains inside the cell, where it can be oxidized by ROS to form the fluorescent product dichlorofluorescein (DCF). Thus an increase in DCF fluorescence suggests increased intracellular concentrations of ROS. In these experiments, two-dimensional images (230 x 230 µm; acquisition time 231 ms) of intracellular DCF fluorescence (>505 nm) were acquired at 2-min intervals using a x40 objective (Zeiss Plan-Neofluor, numerical aperture 1.3) and a helium-neon laser to deliver a scanning excitation line at 488 nm. After obtaining baseline images under normoxic conditions, we exposed cells to 4% O2 for 30 min followed by 16% O2 for 20 min. This protocol was performed in untreated cells (n = 10 experiments) and cells treated with SOD (150 U/ml, n = 8), CAT (200 U/ml, n = 6), or SOD + CAT (n = 4). Perfusion with antioxidant enzymes began 3040 min before the onset of hypoxia and continued throughout the experiment. To analyze images, we defined multiple regions of interest representing 519 cells and a single cell-free area. Fluorescence intensity measured in the cell-free region was taken as background and subtracted from fluorescence in other regions. These differences were calculated for each cell at each time, normalized by values at the onset of hypoxia, and averaged across cells. Averages for each experiment were then used to determine group means and perform statistical comparisons among groups.
To further evaluate the effects of CAT during normoxia, we also measured DCF fluorescence in normoxic rat PASMCs and bovine PAECs. After a 1- to 2-h stabilization period, fluorescent images were obtained at 2-min intervals as described above before and during a 40-min exposure to CAT (200 U/ml, n = 3 for each cell type) or heat-inactivated CAT (n = 3 for each cell type).
Imaging of fluorescein-conjugated SOD and CAT. To determine
whether perfusate SOD and CAT entered porcine PASMCs, we perfused cells for 1
h with normoxic KRB solution containing either SOD or CAT labeled with FITC
after a 1- to 2-h stabilization period during which the perfusate was normoxic
KRB solution without fluorescent indicators. SOD and CAT were labeled by
mixing with a 50-fold higher molar concentration of FITC for 1 h at room
temperature in phosphate-buffered saline alkalinized with NaHCO3
(final concentration 0.1 M). To remove unbound FITC, the reaction mixture
was passed through columns containing polyacrylamide gel (Bio-Spin P-6 and
P-30; Bio-Rad Laboratories, Hercules, CA) with molecular weight exclusion
limits of 6,000 (SOD) or 40,000 (CAT). If we assume complete recovery, the
final concentration of FITC-labeled proteins in perfusate was 1,000 (CAT) or
200 (SOD) U/ml. The higher concentration of FITC-labeled CAT was necessary to
permit detection by laser-scanning confocal microscopy. We obtained sequential
confocal images in the x-y plane (230 x 230, 115 x 115,
or 77 x 77 µm) at 0.3-µm intervals in the z plane
(Z-series) of cells fluorescing at >505 nm in response to excitation at 488
nm. Z-series images of the same cells were acquired at the same settings
immediately before and 1 h after exposure to labeled SOD or CAT followed by
15 min washout with normal KRB. Control cells were subjected to the same
protocol, except that SOD or CAT was not added to the labeling reaction
mixture.
Drugs and Reagents
ACh, SOD, and CAT were purchased from Sigma Chemical. For cell studies, CAT was also obtained from Calbiochem (San Diego, CA). ET-1 was purchased from American Peptide (Sunnyvale, CA) and U-46619 from Cayman Chemical (Ann Arbor, MI). Stock solutions of SOD and CAT were prepared in 0.05 M phosphate buffer and stored in aliquots at -20°C until use. ACh and ET-1 were prepared each day in deionized water and stored at 4°C until use. DCFH2-DA, lucigenin, and FITC were obtained from Molecular Probes (Eugene, OR). Stock solutions in DMSO (DCFH2-DA, FITC) or deionized water (lucigenin) were protected from light and stored in aliquots at -20°C until use. DEPMPO was a gift from Dr. Valerie Roubaud (Marseille, France) and was prepared as reported (12). Concentrations are expressed as final molar concentrations in perfusate.
Data Analysis
T-tests and analysis of variance were used for statistical
comparisons, taking repeated measures into account when appropriate. When
analysis of variance yielded a significant F ratio, least significant
differences were calculated for pairwise comparison among means. We assumed
that P values 0.05 indicated statistical significance. Values
given in the text are means ± SE.
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RESULTS |
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As shown in Table 1, KCl and
U-46619 caused vigorous constriction in both E+ and E-arteries. ACh reversed
constrictions induced by U-46619 in E+ arteries but had no effect in
E-arteries. Viability tests were repeated under normoxic conditions at the end
of the experiment in 14 untreated control E+ arteries, 5 E+ arteries treated
with SOD, and 6 E+ arteries treated with SOD + CAT
(Fig. 1). Vasoconstrictor
responses to KCl at the end of the experiment were decreased compared with
responses at the beginning of the experiment in untreated and SOD-treated
arteries (ID =-138 ± 10 vs. -30 ± 11 µm and -145
± 14 vs. -40 ± 18 µm, respectively) but not altered in
arteries treated with SOD + CAT (
ID =-175 ± 13 vs. -146 ±
19 µm). Vasoconstrictor responses to U-46619 were not changed in any group.
Vasodilator responses to ACh were decreased at the end of the experiment in
untreated and SOD-treated arteries (
ID = 114 ± 15 vs. 26
± 15 µm and 102 ± 29 vs. 21 ± 20 µm, respectively)
but not altered in arteries treated with SOD + CAT (
ID = 116 ±
14 vs. 99 ± 13 µm).
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Hypoxic Constriction in Pulmonary Arteries
In untreated E+ arteries, ID decreased from 198 ± 10 to 185 ±
9 µm in the 30- to 40-min normoxic period preceding the onset of hypoxia.
This spontaneous change in normoxic baseline ID was not different from that
observed over the same period of time after treatment with SOD or SOD + CAT
(195 ± 21 to 201 ± 19 µm and 196 ± 23 to 179 ±
23 µm, respectively). As shown in Fig.
2, decreasing O2 concentration from 16 to 4% for 30 min
in untreated control E+ arteries caused vigorous reversible constriction
(ID at 30 min = -57 ± 10 µm), which was blocked by SOD or SOD
+ CAT (
ID =-17 ± 7 and -5 ± 9 µm, respectively). The
effects of SOD + CAT on HPV were not significantly different from those of SOD
alone. Although endothelial denudation
(Fig. 3) abolished hypoxic
constriction (
ID =-8 ± 6 µm), the response could be restored
by pretreating ("priming") E-arteries with 10-10 M ET-1
(
ID =-39 ± 8 µm). This restored hypoxic constriction was also
blocked by SOD (
ID = 4 ± 3 µm).
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ROS in Pulmonary Arteries
LDCL in porcine distal pulmonary artery segments (Fig. 4) was maximum immediately on placing samples in the luminometer and declined progressively thereafter (Fig. 4A), perhaps due to cooling. Analysis of variance did not reveal significant differences in the time course of LDCL between baseline and exposure periods in either control or experimental groups. Similarly, maximum LDCL measured in the control group was not altered (Fig. 4B); however, maximum LDCL measured in the experimental group increased during hypoxia. This increase approached but did not achieve statistical significance (P < 0.07).
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In a further attempt to determine whether porcine distal pulmonary arteries produced radicals during hypoxia and to characterize the species generated, we performed EPR spectroscopy using DEPMPO as a spin-trapping molecule. Representative EPR spectra are shown in Fig. 5. No spin adduct signals were detected in four of four experiments in vessels subjected to normoxia (Fig. 5A); however, radicals were detected in three of six experiments in vessels subjected to hypoxia (Fig. 5B). These hypoxic spectra were composed of multiple signals, which were identified as hydroxyland carbon-centered alkyl radical adducts of DEPMPO (42) by simulating the signals independently (Fig. 5, C and D) and adding them together (Fig. 5E) to match the experimentally measured spectrum (Fig. 5B). The good correlation between experimental and composite-simulated spectra suggests that hydroxyl- and carbon-centered alkyl radicals were produced by pulmonary arteries during hypoxia.
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ROS in PASMCs
During hypoxia, DCF fluorescence increased in untreated PASMCs from distal porcine pulmonary arteries (Figs. 6 and 7A), reaching a plateau after 1015 min of hypoxia and returning nearly to baseline levels on resumption of normoxia. This response was blocked in PASMCs treated with SOD, SOD + CAT, or CAT. As shown in Fig. 7B, CAT or SOD + CAT (but not SOD) caused an increase in baseline DCF fluorescence during the normoxic period preceding hypoxia, which achieved a stable plateau after 3040 min. CAT also increased DCF fluorescence in rat PASMCs and bovine PAECs; however, the increase was smaller in PAECs than in PASMCs (Figs. 7B and 8). In contrast, DCF fluorescence fell slightly in rat PASMCs and bovine PAECs treated with heat-inactivated CAT (Fig. 8). In Figs. 7B and 8, fluorescence intensity (F) is expressed as differences from background in arbitrary fluorescence units. Initial values of F were lower in PASMCs treated with CAT or SOD + CAT because detector gain was reduced to avoid saturation of the photometer.
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Confocal images of PASMC fluorescence after exposure to FITC-labeled SOD or CAT in perfusate are shown in Fig. 9. Before exposure, myocytes exhibited low-intensity autofluorescence localized to spherical regions in cytoplasm, which were 12 µm in diameter and consistent with mitochondria. After exposure to labeled SOD or CAT, this fluorescence diminished, and new high-intensity fluorescence appeared on cell surfaces and small punctate and linear-reticular regions within the cells. In control cells, where FITC alone was added to the labeling reaction mixture, baseline autofluorescence was slightly diminished and no new fluorescence appeared.
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DISCUSSION |
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Several previous studies have examined the effects of antioxidants and inhibitors of antioxidant enzymes on HPV. Diethyldithiocarbamic acid (DETCA) and triethylenetetramine (TETA), inhibitors of SOD, potentiated hypoxic contractions in guinea pig pulmonary arteries (1), and CAT inhibited hypoxic contractions in sheep pulmonary veins (45); however, SOD did not alter hypoxic contractions in sheep pulmonary arteries (9). In isolated lungs, SOD potentiated or did not affect HPV (5, 58), whereas DETCA, TETA, and the antioxidants nitro blue tetrazolium, pyrrolidine dithiocarbamate, and ebselen inhibited HPV (53, 58, 59). Aminotriazole, an inhibitor of CAT, did not alter a small hypoxic pressor response in isolated rat lungs (36) but nonspecifically inhibited HPV in isolated rabbit lungs (58). The inconsistency among these results could be due to variability in type, concentration, source, and location of ROS produced; specificity and efficacy of the interventions employed; state of cellular antioxidant defenses; alteration of ROS-mediated physiological responses by ROS-mediated injury; and other factors. Although this inconsistency makes it difficult to distinguish which ROS might be involved in HPV, most studies suggest that ROS plays a role. Indeed, HPV may depend on changes in redox state inducible by a variety of pathways instead of a particular ROS. In our experiments, however, O2-· appeared to be essential.
KCl-induced constrictions and ACh-induced dilations were diminished after exposure to hypoxia and reoxygenation in control pulmonary arteries and arteries treated with SOD but not arteries treated with SOD + CAT (Fig. 1). In a previous report (28), we showed that vasomotor responses to KCl and ACh were more depressed at the end of the experiment in arteries exposed to hypoxia than in time-control vessels exposed only to normoxia. Together, these results suggest that ROS-induced injury occurred in our vessels; however, it is unlikely that SOD inhibited HPV by enhancing injury through increased production of H2O2 or its products because addition of CAT to SOD did not alter the inhibitory effects of SOD on HPV (Fig. 2). The cellular source of ROS responsible for injury was probably not endothelium because endothelial denudation did not prevent reduction of KCl responses after reoxygenation (data not shown). Furthermore, ROS production in porcine PAECs, unlike bovine cells, was not enhanced by anoxia reoxygenation (51, 61, 62).
Anoxia alone decreased ROS production in cultured PAECs (61, 62); however, this might not be true for less severe degrees of hypoxia in freshly isolated vessels. To determine whether endothelium-derived ROS contributed to HPV, we measured the effects of SOD on HPV in endothelium-denuded pulmonary arteries primed with the endothelium-derived contracting factor, ET-1. This approach was based on our previous observations that exposure to threshold concentrations (10-10 M) of ET-1 markedly enhanced hypoxic contractions in porcine PASMCs and prevented abolition of HPV by endothelial denudation in porcine pulmonary arteries (28, 44). As shown in Fig. 3, SOD blocked hypoxic constriction in these vessels. These results indicate that HPV depended on ROS produced by and acting on pulmonary arterial smooth muscle instead of endothelium.
Consistent with previous results (20, 53), hypoxia increased DCF fluorescence in PASMCs (Figs. 6 and 7). In theory, increased DCF fluorescence could result from enhanced entry of DCFH2-DA into cells; enhanced intracellular deacetylation of DCFH2-DA to DCFH2; enhanced intracellular oxidation of DCFH2 to DCF; diminished metabolism of DCF to a nonfluorescent product; or diminished leakage of DCF out of cells. With respect to the first two possibilities, we exposed PASMCs to DCFH2-DA throughout the experiment, including the 1- to 3-h period before hypoxia, to maximize intracellular DCFH2 concentration. The last two possibilities are suggested by the return of DCF fluorescence to baseline levels upon reoxygenation (Fig. 7). To our knowledge, DCF metabolism has not been studied in PASMCs. In cancer cells, leakage of calcein, a dye closely related to DCF, was due in part to ATP-dependent extrusion by plasma membrane transporters such as P-glycoprotein and multidrug resistance protein (11, 14). Whether these proteins play a role in PASMCs is unknown. Although the possibility that hypoxia inhibited DCF metabolism or leakage cannot be ruled out, our observation that SOD and/or CAT blocked the increase in fluorescence induced by hypoxia (Fig. 7A) suggests that hypoxia acted by enhancing intracellular oxidation of DCFH2.
In reaction solutions, H2O2 but not O2-· oxidized DCFH2 to DCF; however, in cells, DCFH2 oxidation could occur via a variety of pathways (24, 41). Thus an increase in DCF fluorescence may be more accurately interpreted to indicate an increase in oxidative stress rather than increased production of a particular ROS. Our observation that both SOD and CAT inhibited the increase in DCF fluorescence caused by hypoxia (Fig. 7A) is consistent with this notion. By converting O2-· to highly diffusible H2O2, SOD may have promoted net extrusion of ROS from cells. By converting H2O2 to H2O, CAT may have prevented oxidation of DCFH2 by intracellular peroxidases. By reducing intracellular concentrations of O2-· and H2O2, SOD plus CAT may have limited hydroxyl radical production via the Fenton reaction.
The inhibitory effect of CAT on DCF fluorescence during hypoxia
(Fig. 7A) confirms
previous observations in rat PASMCs
(20) and is consistent with
the recent demonstration that overexpression of CAT attenuated hypoxia-induced
increases in intracellular calcium concentration in PASMCs
(54). However, our results
should be interpreted with caution because CAT and SOD plus CAT also caused an
unexpected increase in normoxic baseline fluorescence
(Fig. 7B), indicating
that CAT increased DCF concentration. This increase did not result from
reaction of CAT with DCFH2-DA in perfusate, because fluorescence in
"background" regions of interest was not altered (data not shown)
and was in any case subtracted from fluorescence in cells. It is also unlikely
that the increase was caused by a constituent of the enzyme preparation other
than CAT, because different lots of CAT preparations from two companies
increased fluorescence while heat-inactivated CAT did not
(Fig. 8). Qualitatively, the
effect of CAT on DCF fluorescence was similar in PASMCs from pig and rat and
PAECs from cow, suggesting that it did not depend on species or cell type;
however, increases in fluorescence were larger in PASMCs than in PAECs
(Fig. 8), consistent with
previous observations that CAT entered vascular smooth muscle more readily
than endothelium (50).
Increased intracellular DCF concentrations could result from decreased
elimination of DCF from cells or increased oxidation of DCFH2. With
respect to the former, CAT may have decreased cell injury induced by
H2O2 or its products and thereby limited leakage of DCF
across cell membranes. With respect to the latter, it seems unlikely that CAT
and SOD plus CAT increased production of ROS, which then caused secondary
oxidation of DCFH2; however, CAT may have oxidized DCFH2
directly, as has been reported for other heme proteins
(37). CAT compound I, the
oxyferryl -cation porphyrin radical produced by reaction of ferric CAT
with H2O2, can regenerate ferric CAT by oxidizing a
variety of hydrogen donors, including ethanol, methanol, phenol, and ascorbate
(19,
23,
39,
49). Although steric hindrance
would be expected to exclude hydrogen donors as large as DCFH2 from
the heme-containing active site of the native enzyme
(23), partial denaturation of
lyophilized CAT might increase accessibility and permit oxidation of larger
molecules (48). Alternatively,
charge transfer from the active site of CAT compound I to its surrounding
protein might allow reactions with larger molecules, as proposed for oxidation
of NADPH by compound I (18,
40). Further investigation
will be necessary to determine which of these explanations is correct.
Because of their size, SOD (mol wt 31,200) and CAT (mol wt
250,000) are frequently assumed not to enter cells. If this assumption
were true, the inhibitory effects of SOD and CAT on HPV
(Fig. 2) and DCF fluorescence
during hypoxia (Fig. 7) would
have to be due to catabolism of ROS released from myocytes into the
extracellular space. To test this assumption, we used laser-scanning confocal
fluorescence microscopy and FITC protein labeling to determine whether SOD and
CAT entered PASMCs in our experiments. As shown in
Fig. 9, FITC-labeled SOD and
CAT were detected in PASMCs, whereas free FITC subjected to the same labeling
and separation protocol was not. These results are consistent with previous
reports that SOD entered lung cells after intratracheal administration
(8,
43) and that CAT entered
vascular smooth muscle cells after administration into culture media
(50). Thus the effects of SOD
and CAT (Figs. 2 and
7) could be due to catabolism
of ROS in the intracellular space (i.e., it is not necessary to conclude that
ROS released from myocytes mediated HPV by acting at the external sarcolemmal
surface or at intracellular loci after reentering cells).
The effects of hypoxia on DCF fluorescence (Figs. 6 and 7) confirm previous assessments of ROS production in PASMCs (20, 30, 53); however, studies in isolated lungs (4, 32), pulmonary arteries (32, 33), and pulmonary arterial homogenates (35) suggested that ROS concentrations decreased during hypoxia. These discrepancies could be due to production of ROS by cells other than PASMCs, limitation of ROS production by severe levels of hypoxia, destruction of a necessary cellular milieu, or use of proximal pulmonary arteries instead of small distal pulmonary arteries, which are thought to be the major locus of HPV (47). In our experiments, hypoxia increased LDCL in small distal porcine pulmonary arteries (Fig. 4), consistent with increased O2-· production; however, this increase did not achieve statistical significance (P < 0.07). Moreover, the lucigenin concentration needed to detect luminescence (20 µM) may not have been low enough to avoid artifacts due to lucigenin redox cycling (2527). We, therefore, repeated these experiments using EPR spectroscopy and the spin trap DEPMPO to assess ROS production. Hydroxyl and alkyl spin adducts, which could be produced secondary to O2-· generation, were detected in three of six preparations exposed to hypoxia and zero of four preparations exposed to normoxia (Fig. 5). Thus neither LDCL nor EPR spectroscopy provided decisive evidence that hypoxia increased ROS generation in distal pulmonary arteries. Possibly, this was due to an inadequate amount of tissue. Alternatively, the absence of distending pressure may have played a role, since stretch has been found to enhance ROS production in both endothelium and vascular smooth muscle (2, 17). In any case, LDCL and EPR spectroscopy in distal pulmonary arteries provided results consistent with the significant increases in DCF fluorescence caused by hypoxia in myocytes from these vessels (Figs. 6 and 7).
Our results add to a growing consensus that HPV depends on ROS produced by and acting on pulmonary arterial smooth muscle; however, more work will be necessary to clarify the intracellular sources of ROS required for HPV, the affects of hypoxia on these sources, and the transduction pathways involved.
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ACKNOWLEDGMENTS |
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Present address of J. Q. Liu: Div. of Pulmonary and Critical Care Medicine, Box 2620, Duke University Medical Center, Durham, NC 27710.
Present address of P. Kuppusamy: Dept. of Internal Medicine, Ohio State College of Medicine and Public Health, 114A Tzgournis, 420 W. 12th Ave., Columbus, OH 43210.
DISCLOSURES
This work was supported by National Heart, Lung, and Blood Institute Grant HL-51912.
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FOOTNOTES |
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The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
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REFERENCES |
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