1 Division of Pulmonary and Critical Care Medicine, Department of Internal Medicine, Asan Medical Center, College of Medicine Ulsan University, Songpa-gu, 138-736 Seoul; 2 Department of Clinical Pathology, Seoul National University Hospital, Chongro-gu, 110-744 Seoul; and 3 Asan Life Science Institute, 138-736 Seoul, Korea
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ABSTRACT |
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We examined the mechanism of
endothelin (ET)-1 regulation by cigarette smoke extract (CSE) and the
effect of platelets on CSE-induced stimulation of ET-1 gene expression
in human and bovine pulmonary artery endothelial cells (PAECs). Our
data show that CSE (1%) induces ET-1 gene expression (after 1 h)
and ET-1 peptide synthesis (after 4 h) in bovine PAECs. The
induction of preproET-1 mRNA level was due to de novo transcription,
and new protein synthesis was not required for this induction. The
protein kinase C inhibitors staurosporine (108 mol/l) and
calphostin C (10
7 mol/l) abolished the induction of ET-1
gene expression by CSE in bovine and human PAECs. Although a lower
concentration of platelets (106 cells/ml in bovine PAECs;
107 cells/ml in human PAECs) did not significantly alter
ET-1 gene expression in PAECs, incubation of platelets with CSE (1%)
and PAECs produced a significant increase in preproET-1 mRNA and ET-1 peptide compared with the values in the presence of CSE (1%) alone. CSE (1%) induced platelet aggregation and increased the expression of
platelet membrane glycoproteins ex vivo. Thus our data suggest that CSE
stimulates ET-1 gene expression via PKC in PAECs. CSE and platelets
showed a synergistic effect on ET-1 gene expression, possibly through
the activation of platelets by CSE.
endothelium
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INTRODUCTION |
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CIGARETTE SMOKE HAS BEEN IMPLICATED as a major risk factor in chronic obstructive pulmonary disease (COPD), chronic hypoxic cor pulmonale, atherosclerosis, and related cardiovascular dysfunction (18, 19, 38). The mechanism for the increased risk of vascular dysfunction is not well understood. It is presumed to be due to the absorption of tobacco smoke constituents that affect endothelial cell function (6, 23, 29, 33, 36), but the true mediator of these vascular diseases associated with smoking is not known. Endothelin (ET)-1 is a potent vasoconstrictor and mitogenic agent released by endothelial cells (41) and has been implicated in the pathogenesis of pulmonary hypertension (2, 14, 28). In humans, cigarette smoking results in a significant increase in plasma ET-1 levels (15, 16), and regular cigarette smoking is associated with functional and morphological changes in the pulmonary arteries (6, 7, 11, 17, 23). The ETA receptor antagonist BQ-610 blocks cigarette smoke-induced mitogenesis in rat airways and vessels (10). Pulmonary hypertension in patients with COPD is associated with the increased expression of ET-1 in vascular endothelial cells, suggesting that the local production of ET-1 may contribute to the vascular abnormalities associated with this disorder (14). Evidence obtained by a variety of approaches indicates that platelets are activated in the circulation of chronic smokers in vivo (13), and platelets stimulate expression of ET mRNA and ET biosynthesis in cultured endothelial cells (31). In the present study, we tested the hypothesis that cigarette smoke extract (CSE) stimulates ET-1 gene expression and that CSE and platelets show a synergistic effect on ET-1 gene expression, possibly through the activation of platelets by CSE, in human and bovine pulmonary artery endothelial cells (PAECs). Protein kinase C (PKC) has been implicated in the regulation of ET-1 production in various endothelial cells (12, 25, 40), and a recent study (21) has shown that cigarette smoke condensate induces PKC activity in endothelial cells. So the role of PKC in CSE-induced stimulation of ET-1 gene expression was also determined in this study.
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MATERIALS AND METHODS |
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Reagents. The Rp diastereomer of adenosine 3',5'-cyclic monophosphorothioate (Rp-cAMPS) was purchased from BIOMOL Research Laboratories (Plymouth, PA). Staurosporine, calphostin C, phorbol 12-myristate 13-acetate (PMA), actinomycin D, cycloheximide, prostacyclin, and ADP were purchased from Sigma (St. Louis, MO).
Cell culture and experimental design.
Bovine PAECs were obtained from American Type Culture Collection (CCL
209) and grown in DMEM supplemented with 20% fetal bovine serum and
1% penicillin-streptomycin for 4 or 5 days before they were
subcultured. Experiments were performed on confluent contact-inhibited cells that had been kept in serum-free DMEM for 24 h to induce quiescence. For experiments designed to measure ET-1 peptide in conditioned medium, the medium was replaced by serum-free DMEM immediately before the studies began. CSE was added to the cell suspension as indicated. Controls consisted of either untreated or
vehicle-treated cells. Human PAECs were obtained from Stratagene (La
Jolla, CA) and grown in endothelial cell growth medium (Clonetics). After achieving confluence, the cells were washed with MCDB-131 alone
and incubated with MCDB-131 containing human albumin (1 mg/ml) and test
reagents. Rp-cAMPS, staurosporine, calphostin C, PMA,
actinomycin D, and cycloheximide were added in concentrations and time
intervals indicated in RESULTS and Figs. 1-5. None of
the compounds used caused significant cytotoxic effects as judged by
trypan blue exclusion (cell viability in all experiments >95%). The
cells were harvested at the indicated time points and centrifuged, and
RNA was extracted as described in RNA isolation and Northern blot
analysis. Each experiment was performed at least three
times.
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Preparation of CSE solutions. An extract of cigarette smoke in phosphate-buffered saline (PBS) was prepared fresh for each day's experiment. Commercial cigarettes with filter (88GOLD, Korea Tobacco & Ginseng), which contain 10 mg tar and 1.0 mg nicotine/cigarette, were used. For the preparation of CSE solutions, we made an automatic smoking machine that simulates the act of smoking by humans. This peristaltic pump was equilibrated at a rate of 5 puffs/min, with a draw time of 2 s, producing 200 cm3 cigarette smoke/min and 1 cigarette/5 min. A cigarette was connected to an automatic peristaltic pump apparatus and lit, and the mainstream smoke was bubbled through 30 ml of PBS that were prewarmed to 37°C. Six cigarettes per thirty milliliters of volume were used. The cigarettes were 60 mm long, and ~40 mm were consumed. Assuming that cigarette smoke is extracted in the blood and equilibration occurs with the total blood volume, the 1% CSE solution used in our study approximately corresponded to exposures associated with smoking ~0.5 pack/day. Concentrations of nicotine in the 1% CSE solution used in the present studies ranged from 10 to 20 ng/ml, comparable to the plasma levels of nicotine found in smokers. The CSE solution was then sterile filtered and diluted into the medium as a percentage of the total volume. The CSE was diluted from 0.1 to 10% for use in these studies. No significant cell death was observed at <10% CSE. To generate a PBS solution of the particulate-phase extract (PPE) of cigarette smoke, smoke from three cigarettes was passed through a glass fiber Cambridge filter (Fisher, Orlando, FL) that retained 99% of all particulate matter in the smoke. The Cambridge filters with the absorbed particulate matter were then extracted for 15 min at 37°C in PBS. The smoke passing through the Cambridge filter was used to prepare a gas-phase extract (GPE) solution for the CSE as described earlier (36). Control solutions were prepared with the same protocols used to generate CSE, PPE, and GPE solutions except that the cigarettes were unlit. CSE was tested for the presence of endotoxin by the Limulus amebocyte lysate assay.
Isolation and preparation of washed human platelets. Platelet-rich plasma (PRP) was prepared from peripheral venous blood obtained from healthy nonsmokers by centrifuging the blood at 150 g for 20 min at 23°C. PRP was aspirated into another centrifuge tube, and prostacyclin (0.8 µM) was added to prevent platelet aggregation. PRP was then centrifuged for 10 min at 800 g, and the pellet was resuspended in 15 ml of Ca2+-free Tyrode buffer of the following composition (in mM): 137 NaCl, 2.7 KCl, 1.0 MgCl2, 0.35 NaH2PO4, 11.9 NaHCO3, and 5.5 glucose, pH 7.35, with 0.8 µM prostacyclin. The mixture was centrifuged for 10 min at 600 g, and the pellet was resuspended in 15 ml of Ca2+-free Tyrode solution with the addition of prostacyclin (0.8 µM final concentration). This mixture was then centrifuged for 10 min at 800 g, the pellet was resuspended in 2 ml of Ca2+-free Tyrode buffer, and CaCl2 was added for a final concentration of 1.8 mM (31). The pellet number was adjusted to 2.5 × 109 cells/ml with the use of a Coulter counter (Coulter, Hialeah, FL).
RNA isolation and Northern blot analysis.
RNA from PAECs was extracted by the method described by Chomczynski and
Sacchi (8). Total RNA (20 µg) was fractionated by
electrophoresis on a 1% agarose-6% formaldehyde denaturing gel and
transferred onto a positively charged nylon membrane (Sure Blot, Oncor,
Gaithersburg, MD). The RNA was cross-linked to the filter by
ultraviolet light. The membrane was prehybridized in Rapid-Hyb buffer
(Amersham, Arlington Heights, IL) for 20 min at 65°C. A plasmid
containing the ovine preproET-1 cDNA probe (550 bp) was kindly provided
by Dr. S. Abman (University of Colorado, Denver, CO). The probes were
labeled with [-32P]dCTP with a random-primer labeling
kit (Amersham) and added to the prehybridization solution at ~1 × 106 dpm/ml. After hybridization for 2 h at 65°C,
the filters were washed three times for 15 min at room temperature in
2× SSC-0.1% SDS and two times for 30 min at 65°C in 1× SSC-0.1%
SDS followed by autoradiography and scanning densitometry. A cDNA probe
for glyceraldehyde-3-phosphate dehydrogenase (GAPDH) was used as
control for RNA loading. The quantification of preproET-1 mRNA was
determined by scanning densitometry. Briefly, films containing the
hybridization signals were developed after 24 h and 2-3 days
of exposure. The 24-h exposure signal had a lighter signal, and it was
compared with a 2- to 3-day exposure to ensure that saturation of the
radiographic film had not occurred. The film with the best
signal-to-noise ratio was then scanned with a Molecular Dynamics series
400 phosphorimager and ImageQuant software (Sunnyvale, CA). The
preproET-1 mRNA signal was then normalized to the GAPDH signal.
Nuclear transcription run-off analysis.
Confluent cultures of bovine PAECs were incubated for 2 h in the
presence and absence of CSE (1%). At the end of the incubation, bovine
PAECs were lysed in ice-cold 10 mM Tris · HCl (pH 7.4), 10 mM
NaCl, 3 mM MgCl2, and 0.5% (vol/vol) Nonidet P-40, and the nuclei were isolated with a Dounce homogenizer as described by Kavanaugh et al. (22) Aliquots of nuclear suspension were
incubated with 0.5 mM each CTP, ATP, and GTP and 250 µCi of
[-32P]-labeled UTP (3,000 Ci/mmol; New England
Nuclear, Boston, MA). The samples were phenol-chloroform extracted,
precipitated, and resuspended at equal counts per minute (cpm) per
milliliter in hybridization buffer (10-20 × 106
cpm/ml). Hybridization to denatured probes (1 µg) dot blotted on
nitrocellulose filters was performed at 40°C for 3 days in the
presence of 50% formamide. cDNA probes for ovine preproET-1 gene were used.
Radioimmunoassay of ET-1. Levels for ET-1 peptide secretion in endothelial cell culture supernatants were quantified by radioimmunoassay (RIA; Peninsula Laboratories, Belmont, CA) in triplicate. Cell supernatants were collected, lyophilized, and suspended in RIA buffer consisting of 100 mmol/l of NaH2PO4, 0.05 mol/l of NaCl, 0.1% bovine serum albumin, 0.1% Triton X-100, and 0.01% NaN3. The rabbit anti-ET-1 that was used showed 100% specificity for ET-1, 17% cross-reactivity to Big ET-1, and 7% reactivity to ET-2 and ET-3. Antiserum (100 µl) was added in equal amounts to 100 µl of either ET standards or reconstituted cell supernatants and incubated for 24 h before 100 µl of 125I-ET-1 at a final concentration of 3,500 cpm were added for 24 h. ET bound to the antibody was immunoprecipitated with 100 µl of anti-rabbit serum before being separated from unbound antibody by centrifugation. Finally, the amount of radioactivity in the immunoprecipitants was determined by gamma counting. ET-1 standard curves revealed 50% displacement by 8-12 pg of ET-1. ET concentrations were calculated by computer-aided processing of the counting data using a logit-log transformation of the calibration curve and were corrected for protein concentration per dish with a Bradford protein assay (4).
Platelet aggregation tests. An 18-ml sample of blood obtained from a healthy nonsmoker was collected into a plastic syringe containing 2 ml of sodium citrate (129 mM adjusted to pH 7.4 with 10% citric acid). Aggregation was studied in PRP obtained by centrifugation (200 g for 15 min) and adjusted with platelet-poor plasma (PPP) to 0.3 × 109 platelets/ml. ADP (1.0 µM) was used as an inducer. Ex vivo PRP aggregometry was performed in a automated platelet aggregation analyzer (PA-3220 Aggregometer II, Daichi), according to the method of Born and Cross (3). Aggregation was quantified as the extent (intensity) of light transmittance in stimulated PRP calibrated as 100% light transmission for PPP and 0% for nonstimulated PRP.
Quantitative analysis of platelet membrane glycoprotein expression. Murine monoclonal antibodies to Gp IIb/IIIa (CD41a), Gp Ib (CD42b), Gp IIIa (CD61), and P-selectin (CD62P); isotypic control; fluorescein isothiocyanate (FITC)-labeled F(ab')2 fragments of human Ig-absorbed sheep anti-mouse IgG antibodies; and calibration beads with four different known amounts of antibody per bead were provided by Dr. M. Canton (Biocytex, Marseille, France).
For the ligand binding studies, 9 parts of blood from healthy nonsmokers who had not taken aspirin or any other antiplatelet agent in the previous 7 days were collected to 1 part of sodium citrate (3.8%). This was centrifuged at 150 g for 10 min, and the PRP was aspirated. The PRP was diluted to 1 × 107 platelets/ml with PBS and incubated with CSE (1%) or PBS for 10 min at room temperature. Aliquots of each dilution were incubated with monoclonal antibodies (10 µg/ml) at room temperature for 20 min. Antibody binding was determined with FITC-labeled F(ab')2 fragments of human Ig-absorbed sheep anti-mouse IgG (heavy plus light) antibodies. The samples were fixed with 1 ml of 1% formaldehyde after 10 min of incubation and analyzed by flow cytometry (FACScan, Becton Dickinson) at 488-nm excitation. Platelet populations were gated according to their forward and side light scatter. Histograms were generated with 10,000 counts, and geometric mean fluorescence was calculated with the CELLQUEST software of the FACScan system (Becton Dickinson). The binding of an isotypic control antibody was taken as nonspecific binding and was subtracted from the observed geometric mean fluorescence. Calibration beads consisting of a mixture of four different populations of 2-µm-diameter latex beads, each with a different defined amount of murine antibody per bead, were used to estimate the number of antibodies bound per platelet, similar to the method described by Poncelet et al. (32) The beads were analyzed in parallel with the samples, with the same FITC reagent and the same settings as the samples. The single bead populations were gated according to their forward and side scatter. Histograms of the geometric mean fluorescence intensity of 10,000 events were recorded and used to plot a log-log graph of the mean fluorescence intensity versus the number of antibodies attached to each bead. The number of platelet-bound monoclonal antibody molecules was estimated from this graph on the basis of the geometric mean fluorescence intensity of the sample.Data analysis. For Northern analysis, densitometry values for ET-1 were first divided by the values for the internal control to correct for any gel loading errors. ET-1 control values were then normalized to a value of 1; the multiple of induction of ET-1 levels from stimulated samples is displayed relative to these values. Statistical comparisons for each stimulus versus control were calculated with a paired two-tailed t-test. RIA results are reported as nanograms of ET-1 per milligram of total protein and were analyzed for significance with an unpaired t-test. Platelet aggregation results are reported as the intensity (in percent) of light transmittance and were analyzed for significance with an unpaired t-test. The value of platelet membrane glycoprotein expression is expressed as the number of platelet-bound molecules and was analyzed for significance with an unpaired t-test. All data are expressed as means ± SE, and significance was set at P < 0.05 or P < 0.01 as noted.
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RESULTS |
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Induction of ET-1 by CSE in bovine PAECs.
CSE caused an increase in preproET-1 mRNA in the tested range of
0.1-10%, with its maximal effect at 1% (Fig.
1A). CSE (1%) led to an
increase in preproET-1 mRNA that was time dependent. Quantitative
densitometric analysis of this experiment indicated a significant
difference in preproET-1 mRNA levels compared with the control level
after 60 min of exposure to CSE. The peak induction occurred after
2 h, and transcript levels remained significantly elevated through
4 h (Fig. 1B). The noted increase in preproET-1 mRNA in
response to CSE may depend on newly synthesized transcripts, the
heightened stability of preproET-1 transcripts, or a combination of
both. To identify which mechanisms may be responsible for the increase
in transcript level, an actinomycin D chase experiment and a nuclear
run-off analysis were done. Actinomycin D (10 µg/ml) was used to
arrest new RNA synthesis, thus allowing quantification of the rate of
disappearance of preproET-1 mRNA. PreproET-1 mRNA disappeared with a
half-life of <15 min in both vehicle- and CSE-treated cells (Fig.
2A). Nuclear run-off analyses
were carried out to determine the transcriptional rate of preproET-1
gene after 2 h of exposure of the cells to CSE (1%). Compared
with the transcriptional rate of preproET-1 in the control cells, there
was an average 3.4-fold increase in preproET-1 gene transcription in
cells treated with CSE (range 1.9- to 4.8-fold; Fig. 2B). To
elucidate whether new protein synthesis is required to elicit the
CSE-induced increase in preproET-1 mRNA, cells were treated with
cycloheximide in the absence and presence of CSE (1%), and preproET-1
mRNA levels were compared with those in control or CSE-treated cells
not exposed to cycloheximide (Fig. 2C). When the cells were
exposed to either CSE or cycloheximide for 2 h, an increase in
preproET-1 mRNA was observed. In cells pretreated with cycloheximide,
CSE induced a further increase in preproET-1 mRNA. To evaluate the time
course of ET-1 production by unstimulated and 1% CSE-stimulated bovine PAECs, cell culture supernatants were withdrawn at 1, 4, 8, and 24 h (Table 1). Extracellular ET-1
production increased from 0.49 ± 0.04 to 2.61 ± 0.10 ng/mg
protein in control cultures and from 0.51 ± 0.04 to 4.80 ± 0.21 in CSE-stimulated cultured cells. After 4 h of incubation,
ET-1 production in the group stimulated with CSE was significantly
higher than that of unstimulated cells.
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Effect of PPE, GPE, and nicotine on preproET-1 mRNA level in bovine PAECs. To investigate which component of cigarette smoke is responsible for the induction of preproET-1 mRNA, cigarette smoke was separated into PPE and GPE. Incubation of bovine PAECs with PPE (1%) or GPE (1%) resulted in an increase in preproET-1 mRNA similar to that observed with CSE (1%). In contrast, incubation of bovine PAECs with nicotine (0.1 and 1 µmol/l) did not change the level of preproET-1 mRNA (Fig. 3).
Role of PKC in the CSE-induced ET-1 gene expression in human and
bovine PAECs.
Activation of PKC by PMA, PKC depletion before agonist stimulation, and
inhibition with PKC antagonists was used to determine the role of PKC
in CSE-induced stimulation of ET-1 gene expression. The PKC inhibitors
staurosporine (at 108 mol/l) and calphostin C (at
10
7 mol/l) abolished the induction of preproET-1 mRNA and
ET-1 peptide by CSE (1%; Fig. 4,
A, B, and D). In bovine PAECs,
short-term activation of PKC with PMA (0.1 µmol/l) showed a modest
increase in preproET-1 mRNA at 30 min, which declined rapidly to below the basal level over 12 h of incubation with PMA. PKC depletion by
PMA pretreatment for 12 h also reduced the CSE-mediated preproET-1 mRNA induction (Fig. 4C). These findings suggest that PKC is
involved in CSE-induced ET-1 gene expression in PAECs. On the other
hand, Rp-cAMPS (at 5 × 10
4 mol/l), which
blocks the activation of PKA by cAMP, had no effect on CSE-induced ET-1
gene expression in bovine PAECs (Fig. 4, A and
D).
Effects of washed human platelets on CSE-induced ET-1 gene expression in human and bovine PAECs. The addition of washed human platelets (107 cells/ml) to bovine PAECs led to an increase in preproET-1 mRNA and ET-1 peptide, whereas a lower concentration of platelets (106 cells/ml) did not significantly alter preproET-1 mRNA and ET-1 peptide levels. Incubation of platelets (106 cells/ml) with CSE (1%) and bovine PAECs produced a significant increase in preproET-1 mRNA and ET-1 peptide compared with the values in the presence of CSE (1%) alone (Fig. 5, A and C). In human PAECs, platelets (107 cells/ml) did not significantly alter preproET-1 mRNA and ET-1 peptide levels, whereas incubation of platelets (107 cells/ml) with CSE (1%) and human PAECs produced a significant increase in preproET-1 mRNA and ET-1 peptide compared with the values in the presence of CSE (1%) alone (Fig. 5, B and C).
Effects of CSE on platelet aggregation and platelet membrane
glycoprotein expression.
To investigate whether the observed synergistic effect of CSE (1%) and
platelets on ET-1 gene expression in PAECs was due to activation of
platelets by CSE, we used a platelet aggregation test and a flow
cytometric method, which permit the detection of a spectrum of
activation-dependent modifications in the platelet surface membrane. In
the presence of platelet activation agonists, a quantitative variation
of the membrane glycoprotein expression is observed. In stimulated
platelets, surface expression of Gp IIb/IIIa, Gp IIIa, and P-selectin
increases, whereas expression of Gp Ib decreases (27). In
this study, we used a panel of monoclonal antibodies directed against
different platelet surface glycoproteins (Gp IIb/IIIa, Gp Ib, Gp IIIa,
and P-selectin). CSE induced platelet aggregation, which was
concentration dependent in the tested range of 0.1-10%, and
enhanced the aggregation induced by ADP (Table 2). CSE (1%) also increased the
expression of GP IIb/IIIa, Gp IIIa, and P-selectin and decreased the
expression of Gp Ib ex vivo (Table 3).
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DISCUSSION |
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In the studies reported here, CSE caused a concentration- and time-dependent increase in preproET-1 mRNA and an increase in ET-1 peptide in bovine PAECs. The induction of preproET-1 mRNA by CSE was a result of an increased transcriptional rate. The preproET-1 mRNA induction was not dependent on de novo protein synthesis as evidenced by the cycloheximide experiments. In fact, treatment with cycloheximide alone produced a dramatic increase in preproET-1 mRNA levels, and cotreatment with CSE and cycloheximide potentiated the induction over either treatment alone. The cycloheximide effect could result from stabilization of the transcript by interference with a translation-dependent ribonuclease or increased transcription by inhibition of a labile repressor protein. A number of AUUUA motifs known to be present in the 3'-untranslated region of several mRNA species with short half-lives (5) are found in the 3'-untranslated region of preproET-1 mRNA. Destabilization of these other AUUUA-containing mRNA species has been shown to be translationally dependent. This corroborates our cycloheximide evidence, suggesting that destabilization of bovine preproET-1 mRNA is translationally dependent. CSE (10%) caused a decrease in preproET-1 mRNA, which seems to be due to its toxic effect because CSE (10%) induced significant apoptosis of bovine PAECs in our experiment (data not shown).
Cigarette smoke is known to contain ~4,000 different constituents
distributed in gas and particulate phases (20). To
determine whether the component(s) responsible for CSE-induced
stimulation of ET-1 gene expression was confined to either the gas or
particulate phase, we separated CSE into a GPE and a PPE. Both GPE and
PPE solutions induced comparable increases in preproET-1 mRNA
expression, suggesting that the components in CSE responsible for
stimulation of ET-1 gene expression exist in both the gas and
particulate phases. Our results demonstrate that nicotine is unlikely
to be responsible for the stimulation of ET-1 gene expression by CSE (1%) because nicotine (107 to 10
6 mol/l)
in comparable concentrations to those in CSE (1%) did not affect ET-1
gene expression in our bovine PAECs. This is supported by the study
that showed that ET-1 is upregulated in the plasma of smokers but not
in the plasma of patients administered transdermal nicotine
(15). In vivo, pulmonary endothelial cells are not the
only source of plasma ET-1. ET-1 is also produced by airway epithelial
cells, neurons and astrocytes in the central nervous system,
endometrial cells, hepatocytes, kidney mesangial cells, Sertoli cells,
breast epithelial cells, neutrophils, and macrophages (24). Aside from the direct effects of the components of
cigarette smoke, several factors related to smoking could also
stimulate ET-1 production in vivo. For example, it is well established
that cigarette smoking acutely increases catecholamine levels in blood (9), which, in turn, could stimulate ET-1 production
(24). So the effect of cigarette smoking on ET-1
production in vivo is complicated, and this area needs further investigation.
In vascular endothelial cells, PKC seems to mediate ET-1 gene induction
caused by thrombin, angiotensin II, or hemodynamic shear stress
(37), and a recent study (21) has shown that cigarette smoke induces PKC activity in endothelial cells. In this
study, the PKC inhibitors staurosporine and calphostin C abolished the
induction of ET-1 gene expression by CSE in human and bovine PAECs.
Prolonged exposure of cells to activators of PKC can result in
downregulation of the enzyme (30). When cells are exposed
to PMA, activation of PKC in the membrane fraction continues for up to
8 h, after which downregulation is apparent for 16-24 h
(42). In our study in bovine PAECs, short-term activation of PKC with PMA (0.1 µmol/l) showed a modest increase in preproET-1 mRNA at 30 min, which declined rapidly to below basal level over 12 h of incubation with PMA. PKC depletion by PMA pretreatment for
12 h reduced the CSE-mediated preproET-1 mRNA induction in bovine
PAECs. From these data, it seems likely that the CSE effects on ET-1
gene expression in PAECs were mediated by PKC. The 5'-flanking region
of the human preproET-1 gene contains octanucleotide sequences for the
12-O-tetradecanoylphorbol 13-acetate-responsive element [TRE; activator protein-1 (AP-1)/c-Jun binding element]. The
synthesis and TRE binding activity of c-Jun protein, which is a major
component of the transcription factor AP-1, is increased by activation
of PKC (37). Cigarette smoke condensate exposure increased
DNA binding of AP-1 in human type II alveolar epithelial cells
(34), and ferrets given a -carotene supplement and
exposed to tobacco smoke had elevated expression of the
c-jun and c-fos genes (39).
Evidence obtained by a variety of approaches indicates that platelets are activated in the circulation of chronic smokers (13). Platelets stimulate expression of preproET mRNA and ET biosynthesis in cultured endothelial cells, and activation of platelets results in a further enhancement of ET release (31). In this study, we examined the effect of platelets on CSE-induced stimulation of ET-1 gene expression in human and bovine PAECs. The addition of platelets (107 cells/ml) to bovine PAECs led to an increase in preproET-1 mRNA and ET-1 peptide, whereas a lower concentration of platelets (106 cells/ml) did not significantly alter preproET-1 mRNA and ET-1 peptide levels. Incubation of platelets (106 cells/ml) with CSE (1%) and bovine PAECs produced a significant increase in preproET-1 mRNA and ET-1 peptide compared with the values in the presence of CSE (1%) alone. Similar results were obtained in experiments with human PAECs. We investigated whether the synergism of CSE and platelets was due to the activation of platelets by CSE. To evaluate the CSE-induced alterations in platelet function, we used a platelet aggregation test and a flow cytometric method, which permit the detection of a spectrum of activation-dependent modifications in the platelet surface membrane. In our study, CSE induced platelet aggregation and enhanced the aggregation induced by ADP. This result is in agreement with the study (1) that showed that platelet aggregation to thrombin and ADP increased 10 min after smoking compared with that before smoking. CSE also increased the expression of platelet membrane Gp IIb/IIIa, Gp IIIa, and P-selectin and decreased the expression of Gp Ib. These data suggest that CSE induces platelet activation ex vivo.
Regular cigarette smoking is associated with morphological changes in the muscular pulmonary arteries that evolve in parallel with small-airway disease and emphysema (17), and endothelium-dependent vasorelaxation is diminished in cigarette smokers and in the lungs of individuals with COPD and hypoxic cor pulmonale (6, 7, 11, 23). The cellular mechanism responsible for these pulmonary vascular alterations is not well understood. It is presumed to be due to the absorption of tobacco smoke constituents that affect endothelial cell function (6, 23, 29, 33, 36), but the true mediator of these vascular diseases associated with smoking is not known. ET-1 is a potent vasoconstrictor and mitogenic agent released by endothelial cells (41) and has been implicated in the pathogenesis of pulmonary hypertension (2, 14, 28). Cigarette smoking results in a significant increase in plasma ET-1 levels (15, 16), and the ETA-receptor antagonist BQ-610 blocks cigarette smoke-induced mitogenesis in rat airways and vessels (10). These findings suggest that ET-1 may contribute to the pulmonary vascular abnormalities associated with smoking and may be important in the eventual development of pulmonary hypertension and cor pulmonale. However, the functional significance of these vascular abnormalities associated with smoking is unknown because smokers without development of COPD (i.e., without alveolar hypoxia) rarely develop overt pulmonary hypertension. Pulmonary hypertension in patients with COPD is associated with the increased expression of ET-1 in vascular endothelial cells (14). BQ-123, an ETA-receptor antagonist, attenuates hypoxic pulmonary hypertension in rats (2), and it is generally accepted that hypoxia plays a major role, through the induction of ET-1, in the development of pulmonary hypertension in patients with COPD (26).
Nitric oxide (NO) is an endogenous vasodilator that contributes to the low vascular resistance in the pulmonary circulation (35). Exposure to CSE results in a decrease in endothelial NO synthase (eNOS) protein and eNOS mRNA contents as well as in eNOS activity in PAECs (36). A reduction in NO production by cigarette smoke is presumed to be responsible, at least in part, for the increased risk of systemic and pulmonary vascular disease and dysfunction in cigarette smokers (33). Taken together, CSE causes an upregulation of ET-1 gene expression and downregulation of eNOS expression in PAECs, and the local imbalance in the release of these mediators may play a part in the development or maintenance of pulmonary vascular disease and dysfunction associated with smoking.
In summary, our study demonstrates that CSE stimulates ET-1 gene expression via PKC in PAECs. Platelets and CSE showed synergism in the stimulation of ET-1 gene expression, possibly through the activation of platelets by CSE. Further studies are needed to find the components of CSE that are responsible for this stimulating effect and for the mechanisms of synergism between platelets and CSE in the stimulation of ET-1 gene expression.
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ACKNOWLEDGEMENTS |
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This work was supported by Korea Research Foundation Grant 1998-0342.
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FOOTNOTES |
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Address for reprint requests and other correspondence: S.-D. Lee, Division of Pulmonary and Critical Care Medicine, Dept. of Internal Medicine, Asan Medical Center, College of Medicine Ulsan Univ., 388-1 Poongnab-dong, Songpa-gu, 138-736 Seoul, Korea (E-mail: sdlee{at}www.amc.seoul.kr).
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 26 July 2000; accepted in final form 26 February 2001.
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REFERENCES |
---|
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---|
1.
Blache, D,
Bouthilliet D,
and
Davignon J.
Acute influence of smoking on platelet behaviour, endothelium and plasma lipids and normalization by aspirin.
Atherosclerosis
93:
179-188,
1992[ISI][Medline].
2.
Bonvallet, ST,
Zamora MR,
Hasunuma K,
Sato K,
Hanasato N,
Anderson D,
Sato K,
and
Stelzner TJ.
BQ123, an ETA-receptor antagonist, attenuates hypoxic pulmonary hypertension in rats.
Am J Physiol Heart Circ Physiol
266:
H1327-H1331,
1994
3.
Born, GVR,
and
Cross MJ.
The aggregation of blood platelets.
J Physiol (Lond)
168:
178-195,
1963[ISI][Medline].
4.
Bradford, M.
A rapid and selective method for the quantitation of microgram quantities of protein utilizing the principle of protein-dye interaction.
Anal Biochem
72:
248-254,
1976[ISI][Medline].
5.
Caput, D,
Beutler B,
Hartog K,
Thayer R,
Brown-Shimer S,
and
Cerami A.
Identification of a common nucleotide sequence in the 3'-untranslated region of mRNA molecules specifying inflammatory mediators.
Proc Natl Acad Sci USA
83:
1670-1674,
1986[Abstract].
6.
Celermajer, DS,
Adams MR,
Clarkson P,
Robinson J,
McCredie R,
Donald A,
and
Deanfield JE.
Passive smoking and impaired endothelium-dependent arterial dilatation in healthy young adults.
N Engl J Med
334:
150-154,
1996
7.
Celermajer, DS,
Sorensen KE,
Gergakopoulos D,
Bull C,
Thomas O,
Robinson J,
and
Deanfield JE.
Cigarette smoking is associated with dose-related and potentially reversible impairment of endothelium-dependent dilatation in healthy young adults.
Circulation
88:
2149-2155,
1993[Abstract].
8.
Chomczynski, P,
and
Sacchi N.
Single-step method for RNA isolation by acid guanidinium thiocyanate-phenol-chloroform extraction.
Anal Biochem
162:
156-159,
1987[ISI][Medline].
9.
Cryer, PE,
Haymond MW,
Santiago JW,
and
Shah SD.
Norepinephrine and epinephrine release and adrenergic medication of smoking-associated hemodynamic and metabolic events.
N Engl J Med
295:
573-577,
1976[Abstract].
10.
Dadmanesh, F,
and
Wright JL.
Endothelin-A receptor antagonist BQ-610 blocks cigarette smoke-induced mitogenesis in rat airways and vessels.
Am J Physiol Lung Cell Mol Physiol
272:
L614-L618,
1997
11.
Dinh-Xuan, AT,
Higenbottam TW,
Clelland CA,
Pepke-Zaba J,
Cremona G,
Butt AY,
Large SR,
Wells FC,
and
Wallwork J.
Impairment of endothelium-dependent pulmonary-artery relaxation in chronic obstructive lung disease.
N Engl J Med
324:
1539-1547,
1991[Abstract].
12.
Emori, T,
Hirata Y,
and
Ohta K.
Secretory mechanism of immunoreactive endothelin in cultured bovine endothelial cells.
Biochem Biophys Res Commun
160:
93-100,
1989[ISI][Medline].
13.
FitzGerald, GA,
Oates JA,
and
Nowak J.
Cigarette smoking and hemostatic function.
Am Heart J
115:
267-271,
1988[ISI][Medline].
14.
Giaid, A,
Yanagisawa M,
Langleben D,
Michel RP,
Levy R,
Shennib M,
Kimura S,
Masaki T,
Duguid W,
and
Stewart DJ.
Expression of endothelin in the lungs of patients with pulmonary hypertension.
N Engl J Med
328:
1732-1739,
1993
15.
Goerre, S,
Staehli C,
Shaw S,
and
Luscher TF.
Effect of cigarette smoking and nicotine on plasma endothelin-1 levels.
J Cardiovasc Pharmacol
26, Suppl3:
S236-S238,
1995[ISI][Medline].
16.
Haak, T,
Jungmann E,
Raab C,
and
Usadel KH.
Elevated endothelin-1 levels after cigarette smoking.
Metabolism
43:
267-269,
1994[ISI][Medline].
17.
Hale, KA,
Niewoehner DE,
and
Cosio MG.
Morphologic changes in the muscular pulmonary arteries: relationship to cigarette smoking, airway disease, and emphysema.
Am Rev Respir Dis
122:
273-278,
1980[ISI][Medline].
18.
Higgins, M.
Epidemiology of COPD. State of the art.
Chest
85, Suppl:
3S-8S,
1984[Medline].
19.
Holbrook, JH,
Grundy SM,
Hennekens CH,
Kannel WB,
and
Strong JP.
Cigarette smoking and cardiovascular disease. A statement for health professionals by a task force appointed by the steering committee of the American Heart Association.
Circulation
70:
1114A-1117A,
1984[Medline].
20.
Janoff, A,
Pryor WA,
and
Bengali ZH.
Effects of tobacco smoke components on cellular and biochemical processes in the lung.
Am Rev Respir Dis
136:
1058-1064,
1987[ISI][Medline].
21.
Kalra, V,
Ying Y,
Deemer K,
Natarajan R,
Nadler JL,
and
Coates TD.
Mechanism of cigarette smoke condensate induced adhesion of human monocytes to cultured endothelial cells.
J Cell Physiol
160:
154-162,
1994[ISI][Medline].
22.
Kavanaugh, WM,
Harsh GR,
Starksen NF,
Rocco CM,
and
Williams LT.
Transcriptional regulation of the A and B chain genes of platelet-derived growth factor in microvascular endothelial cells.
J Biol Chem
263:
8470-8472,
1988
23.
Kiowski, W,
Linder L,
Stoschitzky K,
Pfisterer M,
Burckhardt D,
Burkart F,
and
Buhler FR.
Diminished vascular response to inhibition of endothelium-derived nitric oxide and enhanced vasoconstriction to exogenously administered endothelin-1 in clinically healthy smokers.
Circulation
90:
27-34,
1994[Abstract].
24.
Levin, ER.
Endothelins.
N Engl J Med
333:
356-363,
1995
25.
Marsden, PA,
Dorfman DM,
and
Collins T.
Regulated expression of endothelin-1 in glomerular capillary endothelial cells.
Am J Physiol Renal Fluid Electrolyte Physiol
261:
F117-F125,
1991
26.
Michael, JR,
and
Markewitz BA.
Endothelins and the lung.
Am J Respir Crit Care Med
154:
555-581,
1996[ISI][Medline].
27.
Michelson, AD.
Flow cytometry: a clinical test of platelet function.
Blood
87:
4925-4936,
1996
28.
Miyauchi, T,
Yorikane R,
Sakai S,
Sakurai T,
Oxada M,
Nishikibe M,
Yano M,
Yano I,
Yamaguchi I,
Sugishita Y,
and
Goto K.
Contribution of endogenous endothelin-1 to the expression of cardiopulmonary alterations in rats with monocrotaline-induced pulmonary hypertension.
Circ Res
73:
887-897,
1993[Abstract].
29.
Nadler, JL,
Velasco JS,
and
Horton R.
Cigarette smoking inhibits prostacyclin formation.
Lancet
1:
1248-1250,
1983[ISI][Medline].
30.
Nishizuka, Y.
The molecular heterogeneity of protein kinase C and its implications for cellular regulation.
Nature
334:
661-665,
1988[ISI][Medline].
31.
Ohlstein, EH,
Storer BL,
Butcher JA,
Debouck C,
and
Feuerstein G.
Platelets stimulate expression of endothelin mRNA and endothelin biosynthesis in cultured endothelial cells.
Circ Res
69:
832-841,
1991[Abstract].
32.
Poncelet, P,
George F,
Papa S,
and
Lanza F.
Quantitation of hemopoietic cell antigens in flow cytometry.
Eur J Histochem
40, Suppl1:
15-32,
1996[ISI][Medline].
33.
Powell, JT,
and
Higman DJ.
Smoking, nitric oxide and the endothelium.
Br J Surg
81:
785-787,
1994[ISI][Medline].
34.
Rahman, I,
Smith CA,
Lawson MF,
Harrison DJ,
and
MacNee W.
Induction of gamma-glutamylcysteine synthetase by cigarette smoke is associated with AP-1 in human alveolar epithelial cells.
FEBS Lett
396:
21-25,
1996[ISI][Medline].
35.
Stamler, JS,
Loh E,
Raddy M,
Currie KE,
and
Creager MA.
Nitric oxide regulates basal systemic and pulmonary vascular resistance in normal human circulations.
Circulation
89:
2035-2040,
1994[Abstract].
36.
Su, Y,
Han W,
Giraldo C,
Li YD,
and
Block ER.
Effect of cigarette smoke extract on nitric oxide synthase in pulmonary artery endothelial cells.
Am J Respir Cell Mol Biol
19:
819-825,
1998
37.
Tasaka, K,
and
Kitazumi K.
The control of endothelin-1 secretion.
Gen Pharmacol
25:
1059-1069,
1994[Medline].
38.
US Public Health Services, Office of Smoking and Health, Surgeon General.
The Health Consequences of Smoking: Cardiovascular Disease. Washington, DC: US Department of Health and Human Services, 1982. [DHHS (PHS) Rep 84-50204]
39.
Wang, XD,
Liu C,
Bronson RT,
Smith DE,
Krinsky NI,
and
Russell M.
Retinoid signaling and activator protein-1 expression in ferrets given beta-carotene supplements and exposed to tobacco smoke.
J Natl Cancer Inst
91:
60-66,
1999
40.
Yamada, Y,
and
Mitsuhiro Y.
Effects of protein kinase C activation and inhibition on endothelin-1 release from human aortic and pulmonary artery endothelial cells.
Am J Hypertens
10:
32-42,
1997[ISI][Medline].
41.
Yanagisawa, M,
and
Masaki T.
Endothelin, a novel endothelium derived peptide. Pharmacological activities, regulation and possible roles in cardiovascular control.
Biochem Pharmacol
38:
1877-1883,
1989[ISI][Medline].
42.
Ziegler, A,
Knesel J,
Fabbro D,
and
Nagamine Y.
Protein kinase C down-regulation enhances cAMP-mediated induction of urokinase-type plasminogen activator mRNA in LLc-PK1 cells.
J Biol Chem
266:
21067-21074,
1994