Oxidized LDL inhibit hepatocyte growth factor synthesis in coronary smooth muscle cells

International Journal of Cardiology 103 (2005) 298 – 306 www.elsevier.com/locate/ijcard

Oxidized LDL inhibit hepatocyte growth factor synthesis in coronary smooth muscle cellsB Cornelia HaugT, Christina Lenz, Max Georg Bachem Central Department of Clinical Chemistry, University Hospital Ulm, Robert-Koch-Strasse 8, D-89070 Ulm, Germany Received 12 May 2004; accepted 13 August 2004 Available online 19 February 2005

Abstract Hepatocyte growth factor (HGF) is a potent regeneration factor for endothelial and epithelial cells, and has also been shown to modulate extracellular matrix synthesis and matrix metalloproteinase activity in renal epithelial cells and tumor cells. Controversial results have been published concerning the possible role of HGF in the pathogenesis of coronary atherosclerosis. In this study, we have investigated the effect of oxidized low density lipoproteins (LDL) and elevated glucose concentrations on HGF synthesis in cultured human coronary artery smooth muscle cells. In addition, we have studied whether HGF modulates the release of extracellular matrix, extracellular matrix metalloproteinase inducer (EMMPRIN) and matrix metalloproteinases (MMP) by coronary artery smooth muscle cells. Oxidized LDL (1–10 Ag/ml) induced a significant dose-dependent decrease of HGF release and a concomitant decrease of HGF mRNA expression, whereas native LDL and elevated glucose concentrations induced no significant changes of HGF synthesis. Incubation of cultured human coronary smooth muscle cells with human HGF (1–100 ng/ml) did not significantly alter cell migration and collagen I, fibronectin, EMMPRIN, MMP-1, MMP-2 and MMP-9 release. In summary, our results provide evidence that HGF does not promote coronary plaque growth or plaque destabilization. Regarding the fact that HGF is a potent endothelial cell regeneration factor, the observed downregulation of HGF synthesis by oxidized LDL supports the concept that HGF might be a protective factor in coronary atherosclerosis and that a decrease rather than an increase of HGF synthesis might promote coronary atherosclerosis. D 2005 Elsevier Ireland Ltd. All rights reserved. Keywords: Atherosclerosis; Hepatocyte growth factor; Low density lipoproteins; Extracellular matrix; Matrix metalloproteinases

1. Introduction Hepatocyte growth factor (HGF), a heparin-binding glycoprotein originally isolated from the sera of partially hepatectomized rats [1,2], enhances proliferation, motility and differentiation of various cell types and also acts as an anti-apoptotic factor. HGF is composed of a large a-subunit and a small h-subunit and exerts its mitogenic and motogenic effects on epithelial and endothelial cells by stimulating the tyrosine kinase activity of its specific B

This work was supported by a grant from Deutsche Forschungsgemeinschaft (SFB 451, Teilprojekt B3 to Cornelia Haug and Max Georg Bachem). T Corresponding author. Tel.: +49 731 50024586; fax: +49 731 50024584. E-mail address: [email protected] (C. Haug). 0167-5273/$ - see front matter D 2005 Elsevier Ireland Ltd. All rights reserved. doi:10.1016/j.ijcard.2004.08.059

receptor c-met [3–6]. The HGF receptor is a heterodimeric protein consisting of an a- and a h-subunit and is encoded by the c-met proto-oncogene [5,7,8]. Due to its mitogenic, motogenic and anti-apoptotic action on endothelial cells, HGF seems to play an important role in endothelial cell regeneration and therefore might have a protective function in the complex process of atherogenesis. In animal studies, HGF gene transfer induced therapeutic angiogenesis [9], and after balloon injury HGF inhibited neointima formation and increased re-endothelialization [10]. However, in contrast, recently published studies have suggested that HGF might also promote smooth muscle cell migration and proliferation [11–13]. In addition, HGF has also been shown to induce collagen and fibronectin expression in mesangial cells and renal epithelial cells [14,15] and to induce matrix metalloproteinases (MMP) in endothelial cells, renal epithelial cells and various tumor cells [16–19]. The presently

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available, partly controversial data suggest an atheroprotective effect of HGF due to enhanced endothelial cell regeneration, but might also suggest that HGF promotes the development of atherosclerotic lesions and plaque destabilization by stimulating smooth muscle cell migration and extracellular matrix synthesis and by enhancing matrix degradation. To further elucidate the role of HGF in coronary atherosclerosis we have investigated the effect of native and oxidatively modified low density lipoproteins (LDL) and elevated glucose concentrations on HGF synthesis in cultured human coronary artery smooth muscle cells and whether HGF modulates cell migration, extracellular matrix, extracellular matrix metalloproteinase inducer (EMMPRIN/CD147) and matrix metalloproteinase (MMP) release.

2. Methods 2.1. Culture of human coronary artery smooth muscle cells Human coronary artery smooth muscle cells (HCASMC; passage 3, Clonetics, USA) were subcultured in a 1:1 (v:v) mixture of Dulbecco’s modification of Eagle’s medium (DMEM) with Ham’s F12 medium containing 2 mM lglutamine and 10% fetal calf serum (FCS). Cells from passages 5–8 were seeded into 6- or 24-well plates, and after 48 h, experiments were performed with subconfluent monolayers in culture medium containing 0.1%, 1% and 5% fetal calf serum (FCS).

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2.3. Measurement of HGF in cell culture supernatants HCA-SMC, seeded in 6-well plates were incubated for 48 h with native or oxidatively modified LDL at final concentrations of 1 and 10 Ag/ml, respectively. Cell culture supernatants were concentrated with centricon 30000 (Millipore, Germany; 1 h), and HGF was measured by an enzyme-linked immunosorbent assay (ELISA) kit (R&D Systems, USA). Intra-assay coefficients of variation (n=20) were 7.1% (425 pg/ml) and 4.4% (991 pg/ml), inter-assay coefficients of variation (n=40) were 7.2% (409 pg/ml) and 7.1% (1058 pg/ml). Standards and samples were incubated for 2 h at room temperature in coated microtiter plates. After washing, conjugated antibody was added, followed by incubation for 1.75 h at room temperature and addition of substrate solution. After 30 min, stop solution was added and optical density was read at 450 nm with correction at 570 nm. No immunoreactive HGF was detected in cell culture medium which had not been incubated with the cells. HGF concentrations in the cell culture supernatants were referred to the DNA content in the corresponding culture wells. 2.4. Measurement of DNA DNA content of cells was measured as described previously [21] by fluorescent DNA staining with bisbenzimide (Sigma, Germany) using calf thymus DNA as a standard. Fluorescence (excitation 350 nm, emission 450 nm) was measured with a Victor 1420 Multilabel Counter (Wallac, Finland).

2.2. Isolation and modification of LDL LDL were isolated from blood samples of healthy volunteers by sequential ultracentrifugation with density adjustment by potassium bromide. For preparation of native LDL (nLDL) plasma samples (4 Amol ETDA/ml blood) were supplemented with butylated hydroxytoluene (BHT, 20 AM) in order to prevent oxidation. LDL, isolated from serum samples of the same donors were oxidized by exposure to 5 AM CuSO4 and oxygen for 24 h at 37 8C, respectively. Lipopolysaccharide content of isolated LDL, assessed by the Limulus amebocyte lysate assay (BioWhittaker, Belgium), was b0.05 EU/ml. Lipoprotein concentrations are expressed in terms of their protein content determined by a modified Lowry protein assay (Bio-Rad DC Protein Assay, Bio-Rad, Germany). The degree of oxidation was quantified by i) measurement of absorption at a wavelength of 234 nm, indicating conjugated diene formation of fatty acids and ii) fluorescence measurement at 430 nm (excitation at 360 nm), attributed to the derivatization of apo B-100 lysine residues by reactive aldehydes [20]. Oxidative modification resulted in a 3.4F0.36-fold increase of absorption at 234 nm and a 21.7F4.2-fold increase of fluorescence emission at 430 nm. LDL preparations were stored at 4 8C and used within 1–2 days.

2.5. RNA isolation and reverse transcription polymerase chain reaction (RT-PCR) of HGF and c-met mRNA For detection of HGF mRNA, HCA-SMC were incubated for 3 and 5 h with oxLDL (1 and 10 Ag/ml) and nLDL (10 Ag/ml) in cell culture medium containing 0.1% FCS (control). For detection of c-met mRNA cells were incubated for 3, 5 and 24 h in cell culture medium containing 0.1% FCS. Total RNA was extracted with a High Pure RNA Isolation Kit which includes DNA digestion (Roche Diagnostics, Germany), the concentration and purity of RNA was determined by measuring the absorbance at 260 and 280 nm, and 1 Ag of total RNA was reversely transcribed into cDNA with 0.2 U/ml reverse transcriptase (Superscript; Gibco, Germany) at 42 8C for 50 min. For RT-PCR the following oligonucleotide primers (0.5 AM) were used: human HGF [2] from nucleotide +1405 to +1831 (a 427 bp fragment, Acc M060718), sense primer 5VATGATGATGCTCATGGACCCT-3V, antisense primer 5VCTGGCAAGCTTCATTAAAACC-3V (exon11–exon15); cmet [5] from nucleotide +1281 to +1955 (a 675 bp fragment, Acc J02958), sense primer 5V-AATGGATCGATCTGCCATGT-3V, antisense primer 5V-TCCGAAATCCAAAGTCCCA-3V; h-actin [22] from nucleotide +144 to

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+683 (a 540 bp fragment, Acc M10277), sense primer 5VGTGGGGCGCCCCAGGCACCA-3V, antisense primer 5VCTCCTTAATGTCACGCACGATTTC-3V (exon2–exon4). DNA amplification was performed with 40 ng cDNA using the LightCycler technology (Idaho Technology, USA; HGF and c-met: LightCycler — FastStart DNA Master SYBR Green I (FastStart DNA Taq Polymerase was activated by pre-incubation at 95 8C for 10 min); h-actin: LightCycler — DNA Master SYBR Green I; Roche Diagnostics). Reactions were cycled 34–40 times (95 8C denaturation: HGF and cmet 10 s, h-actin 1 s; annealing: HGF and c-met 56 8C, 10 s, h-actin 58 8C, 5 s; extension (72 8C): HGF and c-met: 25 s, h-actin 22 s; slopes were 20 8C/s). Fluorescence was measured at the end of the extension phase. To confirm the specificity of the amplified products, melting curves were performed at the end of the amplification by cooling the sample at 20 8C/s to 56 8C (HGF and c-met) or 68 8C (hactin) and then increasing the temperature to 95 8C at 0.1 8C/s with fluorescence measurement every 0.1 8C. In addition, PCR products were purified with the QIAquick Gel Extraction Kit (Qiagen, Germany) and sequenced (Sequiserve, Germany). Quantification of the PCR products was performed with the LightCycler Software using purified, sequenced PCR product as standard in a serial dilution. HGF mRNA was referred to h-actin mRNA in the corresponding samples. No detectable PCR products were present in water controls and in controls, amplified without prior reverse transcription. For visualization, PCR products were applied to 1% agarose gel in 0.5 Tris-borate and stained with GelStar (Biozym, Germany). 2.6. Quantitative determination of extracellular matrix synthesis For quantitative measurement of fibronectin and collagen I in cell culture supernatants cells, seeded in 24-well plates, were incubated with human HGF (10, 35 and 80 ng/ml; R&D Systems) in the presence of 0.1% or 1% FCS, ascorbic acid (100 Ag/ml) and h-aminopropionitrile (100 Ag/ml) for 48 h. Cellular fibronectin was measured by time-resolved fluorescence immunoassay as described previously [23]. For measurement of collagen type I 96well microtiter plates were coated overnight with 50 Al/ well rabbit-anti-mouse-IgG (6 Ag/ml; Dako, Germany) and for 4 h at room temperature with monoclonal mouse-antihuman collagen type I (15 Ag/ml; Sigma) in coating buffer (50 mM NaHCO3, pH 9.1). After blocking unspecific binding with assay buffer (0.5% milk powder, 50 mM Tris, 150 mM NaCl, 0.5% sodium azide, pH 7.7) for 2 h at room temperature, samples (100 Al/well) and standards (7.8–1000 ng/ml human placental collagen type I; Biozol, Germany) were added and incubated overnight at 4 8C. After 3 washing steps with wash buffer (150 mM NaCl, 50 mM Tris, 0.05% Tween 40, pH 7.4) the biotinylated second antibody (goat anti-human collagen type I, 1:300; Biozol) was added (4 h). After washing and incubation for

2 h at room temperature with streptavidin–Europium (1:500; DelfiaR Wallac), another washing step was performed, 100 Al enhancement solution (DelfiaR Wallac) was added and after 45 min time-resolved fluorescence of the Europium-chelate was measured with a Victor multiwell-counter (Wallac). All measurements were done in duplicate. Coefficients of variation of duplicate measurements were between 0.6% and 9.8%. Fibronectin and collagen I concentrations in the cell culture supernatants were referred to the DNA content in the corresponding culture wells. 2.7. Measurement of EMMPRIN release Microtiter plates were coated overnight at 4 8C with rabbit-anti-mouse IgG (Dako, 6 Ag/ml, 50 Al/well), after washing (150 mM NaCl, 50 mM Tris, 0.05% Tween 40, pH 7.4) wells were coated with monoclonal anti-CD147 antibody (0.25 Ag/ml, 100 Al/well; R&D Systems). After blocking of unspecific binding (0.5% bovine serum albumin, 50 mM Tris, 150 mM NaCl, 0.5% sodium azide, pH 7.7; 2 h at room temperature), 100 Al standard (R&D Systems) or sample were added to the wells and incubated overnight at 4 8C. After washing, 100 Al biotinylated polyclonal anti-CD147 (0.05 Ag/ml; R&D Systems) were added and the wells were incubated for 4 h at room temperature. After another washing step, wells were incubated for 1 h at room temperature with 100 Al streptavidin–Europium (1:500; DelfiaR Wallac), and after addition of 100 Al enhancer solution (DelfiaR Wallac, 45 min at room temperature), time-resolved fluorescence of the Europium-chelate was measured with a Victor multiwellcounter (Wallac). All measurements were done in duplicate (intra-assay coefficient of variation: 8.7%, n=30). EMMPRIN concentrations in the cell culture supernatants were referred to the DNA content in the corresponding culture wells. 2.8. Measurement of MMP release HCA-SMC, seeded in 6-well plates were incubated for 48 h with human HGF (R&D Systems) at final concentrations of 1, 10, 35, 80 and 100 ng/ml, respectively. Total MMP-1 was measured with a commercially available ELISA kit (Amersham Biosciences, Germany; the assay recognizes free MMP-1 and MMP-1 complexed with inhibitors such as TIMP-1, it does not recognize MMP1 bound to a2-macroglobulin). Intra-assay coefficients of variation (n=10) were 5.5% (16.9 ng/ml) and 7.9% (35.5 ng/ ml), inter-assay coefficients of variation (n=12) were 11.6% (23.2 ng/ml) and 12.0% (55.3 pg/ml). Standards and samples were incubated for 2 h at room temperature in coated microtiter plates, and after washing, the microtiter plates were incubated for 2 h at room temperature with the second anti-MMP-1 antibody. After washing, wells were incubated for 1 h with conjugated anti-rabbit-antibody and

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for 30 min with substrate solution. Stop solution was added and optical density was read at 450 nm. For measurement of MMP-2, microtiter plates were coated overnight at 4 8C with rabbit-anti-mouse IgG (6 Ag/ml, 50 Al/well; Dako). After washing (150 mM NaCl, 50 mM Tris, 0.05% Tween 40, pH 7.4), wells were coated (4 h at room temperature) with 100 Al monoclonal anti-MMP-2 antibody (1 Ag/ml; R&D Systems, antibody recognizes proMMP-2 and active MMP-2), washed and unspecific binding was blocked for 2 h at room temperature (0.5% bovine serum albumin, 50 mM Tris, 150 mM NaCl, 0.5% sodium azide, pH 7.7). After another washing step wells were incubated with 100 Al standard (purified human MMP2, Chemicon, Canada) or sample, followed by washing and 4 h incubation at room temperature with a polyclonal biotinylated anti-MMP-2 antibody (0.1 Ag/ml, 100 Al/well; R&D Systems). Thereafter, wells were washed, incubated for 1 h at room temperature with 100 Al streptavidin– Europium (1:500; DelfiaR Wallac), and for 45 min with 100 Al enhancer solution at room temperature (DelfiaR Wallac). Time-resolved fluorescence of the Europium-chelate was measured with a Victor multiwell-counter (Wallac). All measurements were done in duplicate (intra-assay coefficient of variation: 7.9%, n=30). Total MMP-9 was measured with a commercially available ELISA kit (Oncogene, Germany; the assay detects free MMP-9 and MMP-9 bound to TIMP-1; intra-assay coefficients of variation (n=8): 3–18%, inter-assay coefficients of variation (n=8): 5–20%). Standards and samples (50 Al) were incubated for 2 h at room temperature in coated microtiter plates, and after washing, the microtiter plates were incubated for 2 h at room temperature with conjugated antibody. After washing, substrate solution was added. After 30 min, stop solution was added and optical density was read at 450 nm. MMP concentrations in the cell culture supernatants were referred to the DNA content in the corresponding culture wells. 2.9. Zymography For detection of MMP activity cells were incubated for 48 h with HGF (1–100 ng/ml) in culture medium containing 0.1% FCS. Supernatants were concentrated and adjusted according to their protein content (Ultrafree 0.5 Centrifugal Filters; Millipore). Non-reducing sodium dodecyl sulfate polyacrylamide gel electrophoresis was performed in 7.5% polyacrylamide gels containing 0.2% gelatin (0.2%, w/v). After electrophoresis, gels were washed at room temperature in 2.5% Triton X-100 (215 min) to remove SDS and incubated overnight at 37 8C in 50 mM Tris/HCl, pH 7.5, containing 200 mM NaCl, 5 mM CaCl2 and 0.02% Brij 35. After staining with 0.34% Coomassie Blue R-250 (30 min) gels were destained with 15% (v/v) acetic acid and 40% (v/v) methanol until gelatinolytic activity was seen as a clear band against the blue background of stained gelatin.

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2.10. Detection of cell migration—Wound healing assay Cell migration was assessed by the ability of the cells to migrate into a cell-free area. Cells were seeded in 6-well culture plates and grown for 24 h to reach confluence. After starvation with medium containing 0.1% FCS overnight, the monolayers were wounded by scrapping off a strip of cells in a cross pattern with a 100 Al pipette tip. The medium and detached cells were removed, the remaining cells were washed twice and HGF (80 ng/ml) was added in fresh medium containing 0.1% FCS or 5% FCS, respectively. Cells were then incubated in a humidified incubator (5% CO2), coupled to a microscope (Olympus IX81 Biosystems, Germany) and the migration of the cells into the open gap was documented by a series of photomicrographs (every 10 min for 18 h). 2.11. Statistical analysis Results are expressed as meansFS.E.M. and were evaluated by one way analysis of variance, followed by the Newman–Keuls test.

3. Results 3.1. Effect of LDL and elevated glucose concentrations on HGF synthesis in HCA-SMC Oxidized LDL induced a significant, concentrationdependent reduction of HGF release. The inhibitory effect of oxLDL on HGF release was less pronounced with increasing FCS concentrations in the cell culture medium (Fig. 1A). In contrast to oxLDL, nLDL induced no significant changes of HGF release (Fig. 1B). Investigation of HGF mRNA expression demonstrated that the oxLDL-induced inhibition of HGF release was accompanied by a corresponding decrease of HGF mRNA expression (Fig. 2). Increasing glucose concentrations in the cell culture medium induced no significant changes of HGF release (Fig. 3). 3.2. Effect of HGF on fibronectin, collagen I, EMMPRIN, MMP-1, -2 and -9 release HGF did not significantly alter cellular fibronectin and collagen I release in coronary smooth muscle cells (Fig. 4) and induced only slight, not significant changes of EMMPRIN, MMP-1, MMP-2 and MMP-9 release (Fig. 5A). Zymography also showed no significant changes of MMP activity after incubation with increasing HGF concentrations (Fig. 5B). The expression of the HGF receptor c-met on human coronary artery smooth muscle cells was confirmed by RT-PCR.

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125

HGF (% of control)

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Fig. 3. Effect of different glucose concentrations on HGF release by cultured human coronary artery smooth muscle cells (HCA-SMC). HCA-SMC were incubated for 48 h in culture medium containing increasing glucose concentrations and different FCS concentrations (0.1%, 1% and 5% FCS). HGF release was quantified by an ELISA, and was referred to the DNA content in the corresponding culture wells. Results represent meansFS.E.M. (n=3) and are expressed as relative HGF release compared to controls.

3.3. Effect of HGF on smooth muscle cell migration

60 40

HCA-SMC migration, investigated by the wound healing assay, was not accelerated by incubation with HGF (80 ng/ ml, Fig. 6).

20 0

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nLDL 1 µg/ml

0.1% FCS

1% FCS

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Fig. 1. Effect of (A) oxidized LDL (oxLDL) and (B) native LDL (nLDL) on HGF release by cultured human coronary artery smooth muscle cells (HCA-SMC). HCA-SMC were incubated for 48 h with oxidized and native LDL in culture medium containing different FCS concentrations (0.1%, 1% and 5% FCS), and HGF concentrations were determined by an ELISA. HGF release was referred to the DNA content in the corresponding culture wells. Results represent meansFS.E.M. (n=5) and are expressed as relative HGF release compared to controls; **Pb 0.01, ***Pb 0.001 versus control.

Fibronectin (% of control)

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Fig. 2. Effect of oxidized LDL (oxLDL) and native LDL (nLDL) on HGF mRNA expression in human coronary artery smooth muscle cells (HCASMC). Cells were incubated with oxLDL and nLDL for 3 and 5 h in cell culture medium containing 0.1% FCS. HGF mRNA expression was quantified by RT-PCR using the LightCycler technology, and is expressed as relative mRNA expression compared to the corresponding control (meansFS.E.M. of 3 and 5 h incubation, n=8).

0.1% FCS

1% FCS

Fig. 4. Influence of HGF on cellular fibronectin and collagen I release by cultured human coronary artery smooth muscle cells (HCA-SMC). HCASMC were incubated for 48 h with HGF in culture medium containing 0.1% FCS and 1% FCS, respectively. Fibronectin and collagen I were measured by a time-resolved fluorescence immunoassay, and were referred to the DNA content in the corresponding culture wells. Results represent meansFS.E.M. (n=6) and are expressed as relative release compared to controls.

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EMMPRIN/MMP release (% of control)

A

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4. Discussion The pathogenesis of atherosclerosis is a complex process including endothelial cell injury, deposition of LDL in the vessel wall, migration and proliferation of vascular smooth muscle cells as well as extracellular matrix synthesis. In advanced atherosclerotic lesions extracellular matrix degradation seems to be involved in induction of plaque instability. Until now, little is known about a possible role of HGF in the development or progression of atherosclerotic lesions. In the present study, we have demonstrated a significant concentration-dependent inhibitory effect of oxidatively modified LDL on HGF synthesis in HCA-SMC. Native LDL induced no significant changes of HGF release, and the inhibitory effect of oxLDL on HGF release was attenuated by increasing FCS concentrations in the cell culture medium. These findings suggest that oxidative stress might at least in part be responsible for the inhibitory effect of oxLDL on HGF release by HCA-SMC. In a previous study, HGF release by human coronary artery endothelial cells was significantly increased after incubation with native LDL and oxLDL, and only the highest oxLDL concentration (200 Ag/ml) induced a reduction of HGF release [24]. This discrepancy might be due to cell type-specific susceptibility to nLDL- and oxLDL-induced actions or to cell type-dependent differences in regulation of HGF release. In addition, this discrepancy might also be connected to different biological effects of HGF on these cell types. Several studies have shown that HGF is a potent endothelial cell regeneration factor inducing migration, proliferation, angiogenesis and exerting anti-apoptotic actions [3,25–28]. In contrast, controversial results have been reported concerning the effect of HGF on smooth muscle cells. Some groups observed no effect of HGF on

100 75 50 25 0 Control

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MMP-1

MMP-2

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B kDa 118 83 MMP-2

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Fig. 5. (A) Effect of HGF on EMMPRIN, MMP-1, MMP-2 and MMP-9 release by cultured human coronary artery smooth muscle cells (HCASMC). HCA-SMC were incubated for 48 h with HGF in culture medium containing 0.1% FCS. EMMPRIN and MMP-2 release were quantified by time-resolved fluorescence immunoassay, MMP-1 and MMP-9 release were measured by ELISA. EMMPRIN and MMP release was referred to the DNA content in the corresponding culture wells. Results represent meansFS.E.M. (n=8) and are expressed as relative release compared to controls. (B) Gelatin zymography of HCA-SMC supernatants after 48 h incubation with HGF (photograph shows one of three experiments).

HGF

0h

9h

12 h

15 h

18 h

Control 0 h

9 h

12 h

15 h

18 h

Fig. 6. Influence of HGF (80 ng/ml) on human coronary artery smooth muscle cell migration, investigated by the wound healing assay. Cells, seeded in 6-well plates were wounded by scrapping off a strip of cells in a cross pattern with a pipette tip. Cells were then incubated (0.1% FCS) in a humidified incubator coupled to a microscope and the migration of the cells into the open gap was documented by a series of photomicrographs (photographs show one of three experiments).

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smooth muscle cell growth [10,28,29], whereas others observed a stimulatory effect on smooth muscle cell proliferation and migration [11–13]. In the context of these data, our results, demonstrating oxLDL-induced compensatory upregulation of HGF in endothelial cells and oxLDLinduced downregulation in smooth muscle cells might support the concept of a protective role of HGF in coronary atherosclerosis. In our study, elevated glucose concentrations in the cell culture medium did not significantly alter HGF release by HCA-SMC. In previous studies with endothelial cells, a significant concentration-dependent inhibitory effect of elevated glucose concentrations on HGF release was observed, whereas in smooth muscle cells only the highest glucose concentration (4.5 g/l) exerted an inhibitory effect on HGF release [26,30], again suggesting a different regulation of HGF release in endothelial cells and smooth muscle cells. To further elucidate the possible pathophysiological role of HGF in coronary atherosclerosis, we have investigated the effect of exogenously added HGF on extracellular matrix synthesis, EMMPRIN release, MMP release and migration of coronary artery smooth muscle cells. An enhancement of extracellular matrix synthesis plays an important role in plaque growth. In contrast to previous reports on mesangial cells and renal epithelial cells [14,15], we observed no significant HGF-induced changes of fibronectin and collagen I release in human coronary artery smooth muscle cells. Accumulating evidence suggests that MMP play an important role in the progression of atherosclerotic lesions. An enhanced MMP expression has been observed in plaque tissue [31–33], and it has been shown that cytokines and oxidized LDL induce an upregulation of MMP [34–37]. MMP seem to increase smooth muscle cell migration and proliferation [38–40] and also might promote plaque instability by inducing extracellular matrix degradation [41]. Several studies have demonstrated that HGF induces matrix metalloproteinases (MMP) in endothelial cells, renal epithelial cells and various tumor cells [15–17,19,42]. EMMPRIN is a cell surface glycoprotein which has been shown to induce the production of MMP in fibroblasts and tumor cells [43,44]. It also has been reported that EMMPRIN is expressed in human atheroma [45,46] and own studies have demonstrated that soluble EMMPRIN is upregulated by cytokines (unpublished observations) and modified LDL [47]. In the present study, we found no significant changes of EMMPRIN, MMP-1, MMP-2 and MMP-9 release by coronary artery smooth muscle cells after incubation with HGF. These findings again provide evidence that HGF has different effects in different cell types. Additionally, in the wound healing assay we observed no relevant acceleration of coronary smooth muscle cell migration after incubation with HGF. In summary, we have demonstrated that oxidized LDL induce a dose-dependent reduction of HGF synthesis in human coronary artery smooth muscle cells. HGF did not

significantly alter extracellular matrix synthesis, EMMPRIN release, MMP-1, MMP-2 and MMP-9 release by coronary artery smooth muscle cells and did not accelerate smooth muscle cell migration in the wound assay. These findings suggest that HGF does not promote coronary plaque growth or destabilization. With respect to the fact that HGF acts as a potent endothelial cell regeneration factor, our results provide further evidence that HGF might have a protective function in coronary atherosclerosis and that a decrease rather than an increase of HGF synthesis might promote coronary atherosclerosis.

Acknowledgements We thank Martina de Groot, Martina Adam-J7ger and Gisela Sailer for expert technical assistance. This work was supported by a grant from Deutsche Forschungsgemeinschaft (SFB 451, Teilprojekt B3 to C.H. and M.G.B.).

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