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(The FASEB Journal. 2000;14:1518-1528.)
© 2000 FASEB

Oxidative stress and expression of p22phox are involved in the up-regulation of tissue factor in vascular smooth muscle cells in response to activated platelets

AGNES GÖRLACH*1, RALF P. BRANDES*, STEFFEN BASSUS§, NICOLA KRONEMANN*, CARL M. KIRCHMAIER§, RUDI BUSSE* and VALÉRIE B. SCHINI-KERTH*

* Institut für Kardiovaskuläre Physiologie, Klinikum der J. W. Goethe-Universität, 60590 Frankfurt/Main, Germany; and
§ Stiftung Deutsche Klinik für Diagnostik, Fachbereich Hämostaseologie, 65191 Wiesbaden, Germany

1Correspondence: Institut für Kardiovaskuläre Physiologie. Klinikum der J. W. Goethe-Universität, Theodor-Stern-Kai 7, D-60590 Frankfurt/Main, Germany. E-mail: A.Goerlach{at}em.uni-frankfurt.de


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Vascular injury after balloon angioplasty results in the rapid activation of platelets leading to the release of growth factors and vasoactive substances. In addition, up-regulation of tissue factor (TF) and an increased production of reactive oxygen species (ROS) have been detected at sites of vascular injury. We investigated whether platelet-derived products (PDP) released from activated human platelets increase ROS production, resulting in the induction of TF expression in vascular smooth muscle cells (SMC). PDP induced a time- and concentration-dependent increase in ROS generation in cultured SMC that was mediated mainly by PDGF-AB and TGF-ß1 and impaired by the flavin inhibitor diphenylene iodonium. Increased ROS formation was associated with enhanced mRNA levels of the small NAD(P)H oxidase subunit p22phox or its smooth muscle isoform. Transient transfection with a p22phox antisense vector decreased PDP-induced ROS generation. PDP up-regulated TF mRNA expression, which was redox sensitive and reduced by transfection of the p22phox antisense vector. In addition, PDP-stimulated reporter gene activity of two TF promoter constructs was decreased by coexpression of the p22phox antisense vector. These results indicate that activated platelets up-regulate TF expression and that this response involves ROS generation and a p22phox-containing NAD(P)H oxidase in SMC.—Görlach, A., Brandes, R. P., Bassus, S., Kronemann, N., Kirchmaier, C. M., Busse, R., Schini-Kerth, V. B. Oxidative stress and expression of p22phox are involved in the up-regulation of tissue factor in vascular smooth muscle cells in response to activated platelets.


Key Words: NAD(P)H oxidase • reactive oxygen species • platelet-derived products • vascular injury • antisense technique


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
BALLOON ANGIOPLASTY OF arteries causes endothelial denudation and, as a consequence of the exposure of the subendothelium to the blood, provokes an instantaneous activation of platelets and the coagulation cascade (1 , 2) . Activated platelets release mitogens and growth factors such as platelet-derived growth factor (PDGF), transforming growth factor-ß1 (TGF-ß1), insulin-like growth factor 1 (IGF-1), epidermal growth factor (EGF) and 5-hydroxytryptamine (5-HT) (3) . In addition, tissue factor (TF), a major activator of the coagulation cascade leading to the generation of thrombin, is rapidly induced in the arterial smooth muscle injured by balloon catheterization (4) ; thrombin activity and thrombin receptor expression have been detected at the luminal surface of such injured arteries for several weeks (5 , 6) . Besides its central role in hemostasis, thrombin is also a potent stimulator of vascular smooth muscle cell proliferation (7) . Both the hemostatic and platelet-derived factors have been implicated in the increased proliferation of vascular smooth muscle cells (SMC) and the enhanced synthesis of extracellular matrix proteins that is observed during the development of restenosis after balloon angioplasty (2 , 7) . This is in agreement with findings that both, antiplatelet agents (i.e., IIb/IIIa inhibitors) and direct inhibitors of thrombin (i.e., hirudin), retard the development of intimal thickening in animal models (8 , 9) .

An increase in the generation of reactive oxygen species (ROS) such as H2O2, O2-, or OH- not only by blood cells, but also by cells of the vascular wall, has been observed at sites of balloon injury (10) . In addition to the induction of proliferation (11) , oxidative stress might also contribute to the increased expression of proatherosclerotic genes at sites of vascular injury, including TF, monocyte chemoattractant protein 1 (MCP-1), E-selectin, and vascular cell adhesion molecule 1 (VCAM-1) (12 13 14) . However, the systems generating ROS in SMC and the subsequent activation of signaling cascades leading to redox-sensitive gene expression have not been fully elucidated. In phagocytes, the multicomponent NAD(P)H oxidase has been characterized as the prime enzyme to produce ROS (15) . On activation, assembly of the membrane-bound cytochrome b558 consisting of a 91 kDa and a 22 kDa protein (gp91phox and p22phox, respectively) with the cytosolic factors p47phox, p67phox, and p40phox as well as the small GTP binding protein rac2 allows the generation of large amounts of O2- in the respiratory burst (15) . There is now increasing evidence that nonphagocytic cells, including hepatoma cells, mesangial cells, fibroblasts, and vascular cells, may also express one or more proteins identical or similar to the subunits of the phagocyte NAD(P)H oxidase (16 17 18 19) . Recently, the expression of p22phox was demonstrated in rat SMC (20) and has been shown to be functionally involved in angiotensin II-mediated generation of ROS (21) . A polymorphism in the p22phox gene has been discussed to be associated with the incidence of coronary heart disease (22 , 23) . Moreover, a recently discovered isoform of gp91phox, mitogenic oxidase 1, is expressed in smooth muscle cells (24) .

The aim of the present study was to determine whether 1) activated platelets increase ROS generation in SMC and, if so, 2) to characterize the ROS-generating system(s) and 3) to determine whether this response leads to the induction of TF expression.

We show here that platelet-derived products (PDP) isolated from activated platelets, in particular PDGF-AB and TGF-ß1, induce oxidative stress in SMC in a rapid and sustained manner involving the p22phox subunit of NAD(P)H oxidase, which in turn causes an up-regulation of TF expression.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 REFERENCES
 
Reagents
Diphenylene iodonium was purchased from Alexis Corp. (San Diego, Calif.). Dichlorodihydrofluorescein diacetate (DCFH-DA) was supplied by Molecular Probes (Eugene, Oreg.). Recombinant human PDGF-AB and TGF-ß1 as well as a neutralizing antibody against TGF-ß, rabbit IgG, and goat IgG were provided by R&D Systems (Wiesbaden, Germany). PDGF neutralizing antibody was purchased from Upstate Biotechnology Inc. (Lake Placid, NY.). Recombinant human IGF-1 and EGF were obtained from Prepro Tech EC Ltd. (Rocky Hill, N.J.). Minimal essential medium (MEM) containing Earle’s salts, penicillin and streptomycin were from Life Technologies Inc. (Gaithersburg, Md.); fetal calf serum was from Biochrom KG (Berlin, Germany). Deoxycytidine 5'-{alpha}32P-triphosphate (3000 Ci/mmol) was obtained from Hartmann Analytic (Braunschweig, Germany). Male Wistar rats were purchased from Charles River Wiga GmbH (Sulzfeld, Germany). All other chemicals were obtained from Sigma Chemical Co. (Deisenhofen, Germany).

Preparation and treatment of platelet-derived products
Human PDP were prepared and activated as described previously (25) . Briefly, washed platelets in suspension (3.5x109 platelets/ml) were stimulated with {alpha}-thrombin (1 U/ml) for 2 min, followed by addition of hirudin (10 thrombin-inactivating U/ml). Platelet suspensions were centrifuged, and supernatants containing PDP were collected and stored at -80°C until use. The protein concentrations of the 10 different platelet preparations were in the range of 190 to 500 µg protein/ml. Platelet buffer [0.02 mM tris(hydroxymethyl)aminomethane-HCl (pH 7.4), 0.14 mM NaCl, 5 mM glucose, 1 mM CaCl2] treated as described above served as blank control. In some experiments, PDP were incubated with neutralizing antibodies against PDGF or TGF-ß (0.2 mg/ml) or nonimmune rabbit or goat IgG (0.2 mg/ml) for 1 h at room temperature.

Cell culture
SMC were isolated by explant technique from thoracic aortas from male Wistar rats or from human mesenterial arteries. Human aortic SMC were also obtained from Clonetics (Walkersville, Md.). SMC were cultivated in MEM containing 2 mM L-glutamine, 5 mM TES, 5 mM HEPES (both pH 7.3), 100 U/ml penicillin, 50 µg/ml streptomycin, and either 10% (rat) or 20% (human) fetal calf serum. All experiments were performed with SMC from passages 5 to 18. Prior to stimulation, confluent cells were incubated with MEM without serum in the presence of 0.1% fatty acid-free bovine serum albumin and nonessential amino acids (serum-free medium) for 48 h.

DCF measurements
Acute formation of intracellular ROS was measured in confluent SMC grown in 24-well plates exposed to serum-free medium for 48 h. Cells were washed with Hank’s balanced salt solution (HBSS, Life Technologies GmbH, Karlsruhe, Germany) and loaded with 20 µM DCFH-DA dissolved in HBSS containing 300 µM N-{omega}-nitro-L-arginine for 30 min at 37°C. The dye was removed and 300 µl HBSS containing N-{omega}-nitro-L-arginine and 20 µg protein/ml PDP or equal amounts of blank control was added. DCF fluorescence was measured immediately in a Wallac Victor 1420 fluorescence plate reader (EG&G Wallac, Freiburg, Germany) at 37°C at an excitation wavelength of 488 nm and an emission wavelength of 535 nm for 60 min every 10 min.

Determination of long-term ROS formation was performed in SMC cultivated in serum-free medium for 48 h and subsequently stimulated with PDP or growth factors as indicated. In some experiments, SMC were incubated with various inhibitors for 1 h prior to the addition of either 20 µg protein/ml PDP or blank control. SMC were then washed with HBSS and loaded as described above. After removal of the dye, DCF fluorescence was determined in the absence of PDP in 300 µl HBSS containing 300 µM N-{omega}-nitro-L-arginine after 10 min. DCF fluorescence was corrected for the amount of viable cells at the end of the experiment using the MTT (thiazolyl blue) test according to the manufacturer’s instructions (Sigma). Results are given in arbitrary units (a.u.).

Cytochrome c measurements
The release of superoxide anions was determined by measuring superoxide dismutase (SOD) -inhibitable reduction of ferricytochrome c in SMC grown in 12-well plates to confluence. After 48 h incubation in serum-free medium, cells were washed three times with HEPES-modified Tyrode’s solution containing 1.8 mM CaCl2, 2.6 mM KCl, 0.49 mM MgCl2, 137 mM NaCl, 0.36 mM NaH2PO4, 5.6 mM glucose at pH 7.4 and incubated in 300 µl of the same buffer with and without 150 U/ml SOD for 10 min at 37°C in humidified air. Subsequently, 100 µl ferricytochrome c to obtain a final concentration of 80 µM was added to the reaction buffer solution, followed by addition of 20 µg protein/ml PDP or blank control. After 1 h, the buffer was removed and absorbances at 550 nm and 525 nm were measured immediately. Reduction of ferricytochrome c was calculated from the differences in absorbance at 550 nm normalized to the absorbance at the isosbestic point at 525 nm. Superoxide-specific cytochrome c reduction was calculated between cells incubated with and without SOD by use of an extinction coefficient of 21100 l · mol-1 · cm-1. The rates of O2- generation were determined as mol · min-1 · well-1.

Plasmids and oligonucleotides
A p22phox antisense vector (21 , kindly provided by K. Griendling, Atlanta, Ga.) containing a 465 bp rat p22phox cDNA fragment was digested with PmeI and the resulting fragment was cloned into the PmeI site of pcdna3- (Invitrogen, Groningen, The Netherlands) in sense or antisense direction yielding pcp22s and pcp22as, respectively. Alternatively, phosphorothioate-modified oligonucleotides derived from the rat p22phox sequence (26) were used: Antisense p22phox: 5'GATCTGCCCCATGGTGAGGACC3'; Sense p22phox: 5'GGTCCTCACCATGGGGCAGATC3'; Scrambled p22phox: 5'TAGCATAGCCCTCCGCTGGGGA3'.

Human TF promoter firefly luciferase constructs (kindly provided by A. Bierhaus and P. Nawroth, Tübingen, Germany) contained sequences from -278 bp to +112 bp (pgl2TF1) or -111 bp to +112 bp (pgl2TF6) cloned into the promoterless vector pgl2basic (Promega, Mannheim, Germany) as described previously (27) . The pRL-TK Renilla Luciferase vector (Promega) was used as control for transfection efficiency.

Transient transfections
Transient transfections of SMC were performed by liposomes using the Superfect reagent (Qiagen, Hilden, Germany) according to the manufacturer’s instructions. Briefly, cells were seeded in 24-well plates (for DCF measurements) or 6-well plates (for reporter gene assays and RNA isolation) and grown to 60% confluency. After 24 h cells were washed with serum-free medium.

For DCF measurements, 2 µg of p22phox plasmid DNA or 3 µg of p22phox oligonucleotides were incubated with 100 µl MEM and 15 µl Superfect reagent for 10 min, followed by addition of 2300 µl complete growth medium. Cells were incubated with 200 µl of this mixture per well for 3 h, followed by incubation for 12 h with complete growth medium. Cells were then incubated for another 16 h in serum-free medium and treated with 20 µg protein/ml PDP or blank control before DCF measurements.

For reporter gene assays, 1.5 µg of each reporter gene construct, 0.5 µg of the pRL-TK Renilla luciferase vector, 80 µl MEM, and 15 µl Superfect reagent per well were incubated for 10 min; 600 µl complete medium was added and cells were incubated for 3 h. Then, the mixture was replaced by complete growth medium (1.5 ml) and subsequently treated as described above. For cotransfection experiments, 1.5 µg pcp22s, pcp22as, or pcdna3 control vector was incubated with 1.5 µg of each reporter gene construct and 0.5 µg Renilla luciferase vector. Transfection and stimulation were performed as described above. Cells were washed twice with phosphate-buffered saline (PBS) on ice and lysed in 200 µl reporter lysis buffer delivered with the Dual luciferase kit (Promega). Firefly luciferase and Renilla luciferase activities were measured in a Biolumat 9505 bioluminescence reader (Berthold, Wildbad, Germany) using the reagents provided with the Dual luciferase kit according to the manufacturer’s recommendations. Differences in transfection efficiency and extract preparation were corrected by normalization to the corresponding Renilla luciferase activities.

For RNA determination, SMC were transfected with 2 µg of p22phox plasmid DNA and 0.5 µg of the pRL-TK Renilla luciferase vector to account for transfection efficiency, as described above.

Northern blot analysis
Total cellular RNA from SMC cells was prepared according to standard protocols (28) . Total RNA (25 µg) was separated by electrophoresis through a 1.2% agarose gel containing 6% formaldehyde dissolved in 0.04 M morpholinopropanesulfonic acid, 0.01 M sodium acetate, 1 mM EDTA, pH 7.0, visualized by ethidium bromide staining, transferred to a nylon membrane (Porablot NY amp, Macherey-Nagel, Düren, Germany), and UV cross-linked. A PmeI fragment derived from pcp22s containing 465 bp of rat p22phox cDNA or a 1026 bp human TF cDNA fragment derived by HindIII/XbaI digestion of pN3-hTF-S (kindly provided by A. Bierhaus and P. Nawroth) was labeled with 32P-{alpha}-dCTP using the Ready to Go DNA labeling kit from Amersham Pharmacia Biotech Inc. (Freiburg, Germany) and used at 2 x 106 cpm/ml. Hybridization was performed at 42°C for 16 h. Subsequently, the blots were washed twice with 6x SSPE, 0.1% sodium dodecyl sulfate (SDS) at room temperature and at 42°C and twice with 2x SSPE, 0.1% SDS at 42°C and 54°C for 30 min each time. The blots were exposed to a PhosphorImager screen (Fujifilm Bioimaging Analyzer BAS-1500, Kyoto, Japan) prior to mRNA quantification. The images were displayed using a linear relationship between signal and image intensity. Equal loading was confirmed by staining 18S and 28S ribosomal RNA with ethidium bromide or by hybridization with an 18S ribosomal RNA probe.

Tissue factor determination
For tissue factor antigen measurements, human SMC were seeded in 3.5 cm wells and grown until confluence. After 48 h exposure to serum-free medium, cells were stimulated with 20 µg protein/ml PDP or blank controls. Cells were washed once with PBS and lysed on ice with 400 µl lysis buffer containing 0.5% Triton X and 50 mM TEA in PBS. Cells were frozen, thawed three times, and then sonicated. Cell lysates were stored at -80°C until assayed. TF antigen expression was determined using the Imubind Tissue factor ELISA kit according to the manufacturer’s instructions (American Diagnostica, Greenwich, Conn.).

Statistical analysis
Values presented are means ± SE. Results were compared by ANOVA for repeated measurements, followed by the Newman-Keuls test. A probability level P<0.05 was accepted as significant. The blots are representative of data obtained in two to three additional experiments. Each DCF experiment consists of at least six data points.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 REFERENCES
 
PDP induce extra- and intracellular ROS production via PDGF-AB and TGF-ß1
Exposure of SMC to PDP (20 µg protein/ml) for 1 h induced a significant increase in the extracellular release of superoxide anions from 1.96 ± 0.94 to 7.84 ± 0.39 pmol O2– · well-1 · min-1 as determined by SOD-inhibitable cytochrome c reduction (n=3, P<0.05) as well as an increase in the intracellular ROS production, as measured by DCF fluorescence, from 387 ± 19.6 to 678.2 ± 37.6 a.u. (n=3, P<0.05).

To examine whether PDP up-regulate ROS generating systems, SMC were incubated for various time periods with 20 µg protein/ml PDP or blank controls and DCF fluorescence was measured subsequently in the absence of PDP. PDP-induced DCF fluorescence was significantly elevated after 4 h of incubation, and maximal responses were observed within 24 h (Fig. 1A ). Moreover, the response to PDP was concentration dependent, with a maximal effect obtained in the presence of 10 or 20 µg protein/ml PDP (Fig. 1B ).



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  		Figure 1. PDP induce ROS production in smooth muscle cells. A) Smooth muscle cells were incubated for various time periods with 20 µg protein/ml PDP or B) with different PDP concentrations for 8 h prior to measurement of ROS production by DCF fluorescence. Data are shown as mean ± SE. n = 4, *P<0.05 vs. 0.

To determine which platelet-associated growth factors might contribute to PDP-induced oxidative stress, SMC were exposed to a panel of growth factors including PDGF-AB, TGF-ß1, EGF, IGF-1, and 5-HT for 8 h. PDGF-AB and TGF-ß1 stimulated DCF fluorescence significantly by 67 ± 12% and 63 ± 16%, respectively, and IGF-1 increased DCF fluorescence by 24 ± 4% (Fig. 2A ). EGF (30 ng/ml) and 5-HT (1 µM) had no significant effects (data not shown). Accordingly, preincubation of PDP with neutralizing antibodies against PDGF or TGF-ß, but not with control IgGs, significantly reduced PDP-induced DCF fluorescence by 73 ± 6% and 70 ± 7%, respectively (Fig. 2B ).



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  		Figure 2. PDGF-AB and TGF-ß1 stimulate ROS production. A) Smooth muscle cells were incubated with the platelet associated growth factors PDGF-AB (30 ng/ml), TGF-ß1 (10 ng/ml) and IGF-1 (30 ng/ml) for 8 h prior to measuring DCF fluorescence. Data are shown as mean ± SE. n = 4, *P<0.05 vs. ctr. B) PDP (20 µg protein/ml) were preincubated for 1 h with either 0.2 mg/ml neutralizing antibody against PDGF (PDGF-Ab) or TGF-ß (TGFß-Ab) or with 0.2 mg/ml rabbit (IgG rabbit) or goat IgG (IgG goat). SMC were incubated for 8 h with these mixtures and DCF fluorescence was measured subsequently. Data are shown as mean ± SE. n = 3, *P<0.05 vs. ctr.

PDP-induced ROS production involves the NAD(P)H oxidase subunit p22phox
PDP-stimulated but not basal DCF fluorescence was inhibited by more than 70% by the antioxidants N-acetylcysteine (NAC, 10 mM) and pyrrolidinedithiocarbamate (PDTC, 100 µM) or the iron chelator o-phenanthroline (10 µM), indicating the generation of ROS (Table 1 ). Exogenously added SOD (0.2 mg/ml) or catalase (100 U/ml) did not significantly affect the PDP-induced intracellular ROS generation whereas the cell-permeable catalase inhibitor aminotriazole (10 mM) increased basal and PDP-induced DCF fluorescence by 26 ± 9% and 48 ± 10%, respectively. The CuZnSOD inhibitor diethylthiocarbamate (DETC, 100 µM) abolished PDP-stimulated DCF fluorescence completely and reduced the basal signal by 20 ± 8% (Table 1) . PDP-stimulated DCF fluorescence was not significantly affected by the cyclooxygenase inhibitor diclofenac (100 µM), the xanthine oxidase inhibitor oxypurinol (100 µM), or the respiratory chain inhibitor sodium cyanide (10 µM) (Table 1) . However, PDP-induced DCF fluorescence was completely abrogated by diphenylene iodonium (DPI, 10 µM), an inhibitor of flavin-containing enzymes such as NAD(P)H oxidase, whereas basal DCF fluorescence was minimally affected (Table 1) .


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  		Table 1. ROS generation in vascular smooth muscle cellsa

Since no specific inhibitor is available for NAD(P)H oxidase, an antisense approach was used to further determine the role of NAD(P)H oxidase in PDP-induced ROS formation. PDP-stimulated ROS generation as measured by DCF fluorescence in SMC transiently transfected with a p22phox antisense vector (pcp22as) was significantly reduced to 40 ± 5% of the ROS production in cells transfected with control vector pcdna3 (Fig. 3A ). In additional experiments, similar results were obtained with p22phox antisense oligonucleotides (data not shown). Transfection of SMC with a p22phox sense vector (pcp22s) enhanced PDP-stimulated ROS generation by 94 ± 36% compared to SMC transfected with pcdna3. Basal DCF fluorescence was not affected by p22phox sense or antisense treatment (data not shown).



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  		Figure 3. PDP-induced ROS production is impaired by p22phox antisense cDNA. A) Smooth muscle cells were transfected with a p22phox sense vector (pcp22s), a p22phox antisense vector (pcp22as), or empty control vector (pcdna3) and DCF fluorescence was measured after 8 h incubation with 20 µg protein/ml PDP or blank control. PDP-induced increases in DCF fluorescence were determined for pcp22s- and pcp22as-transfected cells and related to the PDP-stimulated increase in DCF fluorescence in pcdna3-transfected cells. Data are shown as mean ± SE. n = 4, *P<0.05 vs. pcdna3 or pcp22s. B) RNA was isolated from SMC transfected with p22phox sense (pcp22s), p22phox antisense (pcp22as), and control vector (pcdna3); Northern blot analysis was performed using a rat p22phox cDNA probe.

Northern blot analysis demonstrated that p22phox mRNA expression was substantially reduced in SMC transfected with pcp22as compared to cells transfected with pcdna3 (Fig. 3B ); in pcp22s-transfected cells, enhanced p22phox mRNA levels were observed. Transfection efficiencies were comparable, as deduced from luciferase activity of the concomitantly transfected Renilla luciferase vector.

PDP up-regulate p22phox expression via PDGF-AB and TGF-ß1
On exposure to PDP, a time- and concentration-dependent increase in p22phox mRNA expression was observed. After 2 h of incubation with 20 µg protein/ml PDP, p22phox mRNA levels were significantly up-regulated, with a peak value obtained within 8 h (Fig. 4A ). Expression of p22phox mRNA was significantly increased at concentrations greater than 5 µg protein/ml PDP (Fig. 4B ).



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  		Figure 4. PDP induce a time- and concentration-dependent increase in p22phox mRNA levels. Smooth muscle cells were incubated A) for different time periods with 20 µg protein/ml PDP or B) with different PDP concentrations for 4 h; Northern blot analysis was performed using a rat p22phox cDNA probe. Data are shown as mean ± SE. n = 4, *P<0.05 vs. 0.

When SMC cells were exposed to a set of platelet-associated growth factors, p22phox mRNA levels were increased after incubation with PDGF-AB, TGF-ß1, and to a lesser degree with IGF-1 (Fig. 5A ), whereas EGF or 5-HT had no effects. Furthermore, preincubation of PDP with neutralizing antibodies against PDGF or TGF-ß decreased p22phox mRNA up-regulation (Fig. 5B ).



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  		Figure 5. PDGF-AB and TGF-ß1 increase p22phox mRNA levels. A) Smooth muscle cells were incubated with 10 ng/ml TGF-ß1, 30 ng/ml PDGF-AB, 30 ng/ml IGF-1, 30 ng/ml EGF, or 1 µM 5-hydroxytryptamine (5-HT) for 4 h. Northern blot analysis was performed using a rat p22phox probe. Data are shown as mean ± SE. n = 4, *P<0.05 vs. ctr. B) PDP (20 µg protein/ml) were preincubated for 1 h with either 0.2 mg/ml neutralizing antibody against PDGF-AB (PDGFAB-Ab) or TGF-ß (TGFß-Ab). SMC were then incubated for 4 h with these mixtures prior to RNA isolation and subsequent Northern blot analysis using a rat p22phox cDNA probe. The blot is representative of three experiments.

PDP up-regulate tissue factor expression via PDGF-AB and TGF-ß1
Next we investigated whether PDP modulate TF expression. Incubation of SMC with PDP for 6 h enhanced TF mRNA levels significantly at concentrations greater than 5 µg protein/ml (Fig. 6A ). A significant increase in TF protein levels was observed after incubation of SMC with more than 10 µg protein/ml PDP for 8 h (Fig. 6B ). Neither EGF, 5-HT, nor IGF-1 alone affected TF mRNA levels, whereas PDGF-AB or TGF-ß1 markedly enhanced TF expression (Fig. 7A ). Moreover, in the presence of neutralizing antibodies against PDGF or TGF-ß, PDP-induced TF mRNA up-regulation was substantially decreased (Fig. 7B ).



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  		Figure 6. PDP induce tissue factor expression. A) Smooth muscle cells were incubated with increasing concentrations of PDP for 6 h and Northern blot analysis was performed using a human TF cDNA probe. n=4, *P<0.05 vs. 0. B) Smooth muscle cells were incubated for 8 h with 10, 20, or 30 µg protein/ml PDP and TF antigen was measured in cell lysates by ELISA. Data are shown as mean ± SE. n = 4, *P<0.05 vs. 0.



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  		Figure 7. PDGF-AB and TGF-ß1 induce tissue factor mRNA levels. A) Smooth muscle cells were incubated with 10 ng/ml TGF-ß1, 30 ng/ml PDGF-AB, 30 ng/ml IGF-1, 30 ng/ml EGF, or 1 µM 5-hydroxytryptamine (5-HT) for 6 h. Northern blot analysis was performed using a human TF probe. Blot is representative of two additional independent experiments. B) PDP (20 µg protein/ml) were preincubated for 1 h with neutralizing antibodies (0.2 mg/ml) against PDGF or TGF-ß. SMC were exposed for 6 h to these mixtures. Blot is representative of two additional independent experiments.

PDP induce tissue factor expression via a redox-sensitive pathway involving p22phox
PDP-stimulated TF mRNA expression was markedly decreased by the antioxidants PDTC and NAC and abolished by DPI (Fig. 8A ). Moreover, externally applied H2O2 stimulated TF expression in a concentration-dependent manner (data not shown). To obtain evidence for a possible role of NAD(P)H oxidase as a source for ROS-mediated regulation of TF expression, TF mRNA levels were determined in SMC transiently transfected with pcp22s, pcp22as, or the control vector pcdna3. A marked decrease in PDP-induced TF mRNA levels was observed in cells transfected with pcp22as compared to cells transfected with pcdna3. In pcp22s-transfected cells, PDP-induced TF expression was not significantly different from pcdna3-transfected cells (Fig. 8B ).



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  		Figure 8. PDP-induced tissue factor expression is redox-sensitive. A) Smooth muscle cells were incubated for 6 h with 20 µg protein/ml PDP alone or in the presence of the antioxidants pyrrolidinedithiocarbamate (PDTC, 100 µM) and N-acetylcysteine (NAC, 10 mM) or the flavin inhibitor diphenylene iodonium (DPI, 10 µM). TF expression was determined by Northern blot analysis. Blot is representative of two additional independent experiments. B) Smooth muscle cells were transiently transfected with p22phox sense (pcp22s), p22phox antisense (pcp22as) or control vector (pcdna3). After 6 h incubation with 20 µg protein/ml PDP, RNA was isolated and Northern blot analysis was performed using a human TF cDNA probe. The blot is representative of three independent experiments.

To further support the role of PDP and p22phox in the regulation of TF expression, reporter gene experiments were performed using two TF promoter constructs (27) . The plasmid pgl2TF6 harbored the minimal promoter, including a serum-response element with three egr-1 sites (-111 to +14 bp), whereas the plasmid pgl2TF1 contained in addition an LPS-response element (-227 to -172 bp) with an NF-{kappa}b and two AP-1 consensus sequences. Although pgl2TF1 and pgl2TF6 differed in their basal reporter gene activity, showing a respective 5.3 ± 1.7-fold and 2.0 ± 0.8-fold increase in luciferase activity compared to the control vector pgl2basic, both constructs were stimulated approximately twofold (1.9±0.2 and 2.3±0.3, respectively) compared to basal levels by 20 µg protein/ml PDP (Fig. 9A ). PDTC and DPI attenuated PDP-induced luciferase activity (data not shown), suggesting the involvement of a redox-mediated pathway in the PDP-induced TF reporter gene activity. Furthermore, in the presence of pcp22as, PDP-induced luciferase activities of pgl2TF1 and pgl2TF6 were significantly reduced to 50 ± 4% and 45 ± 12% of the corresponding luciferase activities in the presence of pcdna3 (Fig. 9B ). Cotransfection of pcp22s significantly increased PDP-induced luciferase activities of pgl2TF1 and pgl2TF6 by 2.7 ± 0.5 and 3.1 ± 0.6-fold, respectively, compared to the respective values in pcdna3-transfected cells.



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  		Figure 9. p22phox is involved in PDP-induced tissue factor reporter gene activity. Smooth muscle cells were transiently transfected with a promoterless luciferase reporter gene (pgl2basic) or two luciferase reporter genes containing different human TF promoter sequences. The plasmid pgl2TF1 contains TF sequence from -278 to +112 bp harboring a distal enhancer (-227 to -172 bp) with two AP-1 sites and a NF-{kappa}B site and a proximal enhancer (-111 to +14 bp) with three egr-1 consensus sequences. The plasmid pgl2TF6 contains TF sequence from -111 to +112 bp, including the proximal enhancer. Firefly luciferase activity was normalized to Renilla luciferase activity. A) Transiently transfected SMC were exposed to 20 µg protein/ml PDP for 12 h and luciferase activity was determined in relation to the control vector pgl2basic. The ratio of PDP-stimulated to unstimulated luciferase activity is given for each plasmid. Data are shown as mean ± SE. n = 4, *P<0.05 vs. pgl2basic. B) SMC were cotransfected with TF promoter luciferase constructs and a p22phox sense vector (pcp22s), a p22phox antisense vector (pcp22as) or the control vector pcdna3. Luciferase activity was measured in unstimulated cells and in cells exposed to 20 µg protein/ml PDP or blank control for 12 h. The PDP-induced increase in luciferase activity for each TF plasmid in the presence of pcp22s or pcp22as was related to the respective PDP-induced luciferase activity in the presence of pcdna3. Data are shown as mean ± SE. n = 4, *P<0.05 vs. pgl2basic, +P<0.05 vs. pgl2basic.


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
REFERENCES
 
The present study shows that PDP isolated from activated, washed human platelets induce oxidative stress in vascular SMC. The enhanced ROS production was most likely derived from an NAD(P)H oxidase since 1) the flavin inhibitor DPI impaired ROS production, 2) PDP up-regulated p22phox mRNA in a time- and concentration-dependent manner closely resembling the kinetics of PDP-induced DCF fluorescence, and 3) transfection of an p22phox antisense vector or p22phox antisense oligonucleotides markedly impaired PDP-induced ROS production.

Oxidative stress has been implicated in the pathogenesis of several cardiovascular diseases including atherosclerosis, hypertension, and diabetes mellitus (29) . Moreover, increased ROS generation has also been observed at sites of vascular injury and has been related to the development of restenosis after balloon angioplasty (30) , since in vitro experiments have shown that ROS can stimulate migration and proliferation of SMC and fibroblasts (11 , 31) . In addition to activated phagocytes, endothelial cells and vascular smooth muscle cells may also contribute to the increased ROS generation at sites of vascular injury (32 , 33) . However, in contrast to macrophages, the rate of ROS production in most cases is substantially lower in vascular cells, suggesting that this ‘low-output’ production of ROS might not only be targeted to damage cells, but might fulfill a more subtle task in acting as signaling molecules. Indeed, there is increasing evidence that ROS might play an important role in inducing gene expression (34 35 36) . The present findings demonstrate a rapid increase of intra- and extracellular ROS generation in SMC by PDP that requires the continuous presence of PDP. These findings suggest that PDP cause direct activation of a radical generating system. In addition, long-term exposure to PDP also increased ROS production, which was inhibited by p22phox antisense treatment and elevated in the presence of p22phox sense vector. In parallel, PDP-induced up-regulation of p22phox mRNA expression was decreased by p22phox antisense treatment and increased by p22phox sense treatment. Since transfection efficiencies were comparable, the p22phox cDNA fragment, although it does not contain full-length p22phox cDNA, may be sufficient to enhance the level of p22phox, for example, by enhancing p22phox mRNA stabilization. Thus, up-regulation of NAD(P)H oxidase by activated platelets in smooth muscle may provide an explanation for sustained formation of ROS after vascular injury.

Although PDP-stimulated ROS generation was inhibited by the antioxidants PDTC and NAC, neither extracellularly applied catalase nor SOD had any effect. The failure to reduce ROS production by exogenously applied catalase or SOD to intact cells has been attributed to the low cell permeability of these enzymes (37) . However, inhibition of catalase by aminotriazole and of CuZnSOD by DETC increased and decreased, respectively, basal and stimulated DCF fluorescence, indicating the PDP-induced generation of H2O2 by SMC. Since the iron chelator o-phenanthroline also inhibited PDP-stimulated DCF fluorescence, one might speculate that a Fenton-type reaction is involved in the activation process of DCFH-DA, as has been proposed for activation of the ROS-sensitive dye dihydrorhodamine (38) .

Based on the addition of single growth factors and on the use of neutralizing antibodies, PDGF-AB and TGF-ß1 were identified as the major platelet-associated growth factors to stimulate ROS generation and induce p22phox mRNA expression, suggesting that these growth factors may be potent activators of NAD(P)H oxidase in SMC. Previously, PDGF-stimulated ROS formation was found to be sensitive to the flavin inhibitor DPI in SMC (39) and to be dependent on the small GTPase rac1 known to be required for activation of the phagocyte NAD(P)H oxidase (34) .

ROS have not only been suggested to be involved in the proliferative and chemotactic response of vascular cells, but they may also contribute to the procoagulant state observed at sites of vascular injury (40) . TF expression has been shown to be redox sensitive in endothelial cells stimulated with TNF-{alpha} (27) ; it was found to be induced in the arterial wall by balloon injury and to accumulate in atherosclerotic plaques (4 , 41) . In addition to its important role in coagulation, TF has also been involved in mitogenic and chemotactic responses in vascular SMC (42) . The present findings indicate that PDP up-regulate TF expression in SMC and identify PDGF-AB and TGF-ß1 as the platelet-derived growth factors largely contributing to this response. Consistent with these findings, PDGF-AA and PDGF-BB have been shown to enhance TF mRNA in rat and human SMC (43 , 44) . Furthermore, reporter gene experiments using two TF promoter constructs containing either a proximal serum response element with three egr-1 sites or, in addition, a distal LPS-responsive element with two AP-1 and one NF-{kappa}B consensus sites (27 , 45) , show that PDP stimulated TF reporter gene activity of both plasmids to a similar extent. This suggests that the proximal serum response element containing egr-1 binding sites is sufficient for transcriptional activation of the TF promoter by PDP. Recently, vascular endothelial growth factor (VEGF) has been shown to induce TF expression via the redox-sensitive transcription factor egr-1 in endothelial cells (46) . Moreover, a marked increase in the expression of egr-1 has been observed at sites of mechanical injury of the arterial wall (47) . Thus, activation of egr-1 might be an important step in PDP-stimulated TF expression in SMC.

Furthermore, PDP-induced TF expression was impaired in the presence of the antioxidants PDTC and NAC, whereas exogenously added H2O2 increased TF mRNA levels, indicating an important role of ROS in mediating TF up-regulation in SMC. This redox-sensitive response most likely involves a p22phox-containing NAD(P)H oxidase since 1) the flavin inhibitor DPI reduced PDP-stimulated TF mRNA levels, 2) transfection of p22phox antisense vector reduced PDP-induced TF mRNA levels, and 3) TF promoter reporter gene activity stimulated by PDP was significantly decreased in the presence of p22phox antisense vector. Transfection with p22phox sense vector enhanced PDP-induced TF promoter reporter gene activity but did not significantly affect PDP-stimulated TF mRNA expression. Since SMC were only transiently transfected, the higher sensitivity of the reporter gene assays compared to Northern blot analysis might account for this difference. Thus, ROS generated by a p22phox-containing NAD(P)H oxidase appear to be critically involved in the regulation of TF expression by PDP in SMC.

Taken together, our findings suggest that platelets activated at sites of vascular injury release growth factors such as PDGF-AB and TGF-ß1, which induce NAD(P)H oxidase, leading to the rapid and sustained generation of ROS in SMC. Increased oxidative stress, in turn, induces redox-sensitive transcription factors, possibly egr-1, resulting in the up-regulation of TF. The platelet-dependent expression of TF may lead to the generation of thrombin that catalyzes the formation of fibrin, but may also contribute to the chemotactic and mitogenic responses of the injured arterial wall. In addition, thrombin-induced activation of platelets may further enhance the expression of TF as described in the present study. Such a positive feedback loop may account for the substantial accumulation of TF observed in mechanically injured arteries. Thus, generation of ROS by a p22phox-containing NAD(P)H oxidase is likely to play a pivotal role in the redox-sensitive regulation of proatherosclerotic genes. Furthermore, these findings may provide the basis for the development of new therapeutic strategies to improve the outcome of vascular injury.


   ACKNOWLEDGMENTS
 
R.P.B. is the recipient of a young investigator award from the Klinikum der J. W. G.-Universität, Frankfurt/Main. This work was supported in part by Deutsche Forschungsgemeinschaft (DFG), Bonn, Germany, and by grants from the Heinrich und Fritz Riese-Stiftung, Germany, and the Paul- und Ursula Klein-Stiftung, Germany. The authors thank Mrs. I. Winter, Mrs. M. Piepenbrook, and Mrs. G. Römer for expert technical assistance.

Received for publication September 24, 1999. Revision received January 25, 2000.
   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 

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