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Howard Florey Institute of Experimental Physiology and Medicine (C.S.S., I.M., R.A.D.B., S.L.L., G.W.T.), University of Melbourne, Victoria 3010, Australia; Connetics Corporation (E.N.U., J.M.), Palo Alto, California 94303; and Baker Heart Research Institute (X.-J.D.), Melbourne, Victoria 8008, Australia
Address all correspondence and requests for reprints to: Chrishan S. Samuel, Ph.D., Howard Florey Institute, Gate 11, University of Melbourne, Parkville, Victoria 3010, Australia. E-mail: c.samuel{at}hfi.unimelb edu.au.
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-smooth muscle actin expression, collagen synthesis, and collagen deposition (by up to 95% with TGF-ß and 40% with Ang II; all P < 0.05). Fibroblast proliferation was significantly increased by 10–8 M and 10–7 M Ang II (63–75%; P < 0.01) or 0.1–1 µg/ml IGF-I (27–40%; P < 0.05). Relaxin alone had no marked effect on these parameters, but it significantly inhibited Ang II- and IGF-I-mediated fibroblast proliferation (by 15–50%) and Ang II- and TGF-ß-mediated fibroblast differentiation, as detected by decreased expression of -smooth muscle actin (by 65–88%) and collagen (by 60–80%). Relaxin also increased matrix metalloproteinase-2 expression in the presence of TGF-ß (P < 0.01) and Ang II (P < 0.05). Furthermore, relaxin decreased collagen overexpression when administered to two models of established fibrotic cardiomyopathy, one due to relaxin deficiency (by 40%; P < 0.05) and the other to cardiac-restricted overexpression of ß2-adrenergic receptors (by 58%; P < 0.01). These coherent findings indicate that relaxin regulates fibroblast proliferation, differentiation, and collagen deposition and may have therapeutic potential in diseased states characterized by cardiac fibrosis. |
Introduction |
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Cardiac fibrosis is a hallmark of heart disease and is the result of a variety of structural changes that occur after pathological stimuli to the cardiovascular system (8). The fibrosis is characterized by a disproportionate accumulation of fibrillar collagen that occurs after myocyte death, inflammation, enhanced workload, hypertrophy, and stimulation by a number of hormones, cytokines, and growth factors (9, 10, 11, 12). The proximal effector cells in this process are fibroblasts, which are activated to become myofibroblasts, which produce an excessive amount of collagen in response to inflammatory mediators, such as TGF-ß (11) and angiotensin II (Ang II) (12). It has also been demonstrated that Ang II induces TGF-ß release from cardiac fibroblasts, and together, they act in synergistic pathways to interfere with normal structure and function of the surrounding myocardium (13, 14, 15). Despite the pathophysiological significance of fibrosis, no effective treatment strategies exist.
The role of relaxin in the heart has been demonstrated by our recent studies showing elevated collagen content, leading to increased left ventricular end-diastolic pressure and chamber stiffness in relaxin-deficient mice (16). The relevance of relaxin in human heart disease has also been indicated by a recent study showing substantial changes in myocardial and circulating levels of relaxin in patients with heart failure (17). Furthermore, relaxin was shown to modulate the effects of Ang II (17), identifying it as a potential therapeutic agent against cardiovascular disease. However, direct effects of relaxin administration on cardiac fibrosis and the potential mechanisms involved have never been demonstrated.
Based on these previous observations (2, 3, 4, 5, 6, 16, 17), we propose that relaxin acts through its G protein-coupled receptor (LGR7) (18) as a collagen-repressing hormone against cardiac fibrosis. In this study, we sought to investigate relaxin’s effect on cardiac fibroblast proliferation and differentiation/activation and on the synthesis and deposition of collagen stimulated by profibrotic factors, which are known to be altered in diseased states. Additionally, we evaluated the antifibrotic properties of relaxin in two independent murine models of fibrosis, the relaxin-deficient (relaxin knockout) mouse, which develops an age-related progression of cardiac fibrosis (16), and the ß2-adrenergic receptor (ß2-AR) transgenic mouse, a more severe model of fibrotic cardiomyopathy (19).
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Animals
Neonate (1 d old) Sprague Dawley rats were used for tissue collection and subsequent cardiac cell isolation and preparation. All male relaxin wild-type and relaxin knockout mice used in this study were generated from RLX heterozygous (C57Blk6Jx129SV) parents (16). The male wild-type and heterozygous ß2-AR transgenic mice used were generated from heterozygous (C57Blk6JxSJL) parents (19). The animals were housed in a controlled environment and maintained on a 14-h light, 10-h dark schedule with access to rodent lab chow (Barastock Stockfeeds, Pakenham, Victoria, Australia) and water. These experiments were approved by the Howard Florey Institute’s and Baker Heart Research Institute’s Animal Experimental Ethics Committees, which adhere to the Australian code of practice for the care and use of laboratory animals for scientific purposes.
Cell culture
To determine the distribution of relaxin and LGR7 expression in cardiac cells, atrial myocytes and fibroblasts, ventricular myocytes and fibroblasts, and vascular smooth muscle cells (VSMC) were obtained from neonate rats using standard collagenase digestion methods. The cells were maintained in DMEM supplemented with 10% fetal calf serum, penicillin (50 U/ml), and streptomycin (50 µg/ml) (DMEM-FBS). Cardiac fibroblast isolation from myocytes and subsequent preparation were performed as described previously (20). These preparations contained more than 95% cardiac fibroblasts as determined by morphological appearance and immunocytochemical staining. Fibroblasts were used between passages 2 and 4 for all studies.
Determination of collagen synthesis and deposition from cardiac fibroblasts
To study the effects of relaxin on collagen expression, atrial and ventricular fibroblasts were plated at an equal density of 1–2 x 105 cells/cm2 in DMEM-FBS. To investigate the effects of relaxin on TGF-ß-stimulated collagen expression, cells were then treated for 72 h with rhRLX (100 ng/ml), TGF-ß (1 or 2 ng/ml), or a combination of rhRLX (100 ng/ml) and TGF-ß (2 ng/ml) in serum-free DMEM supplemented with lactalbumin hydrolysate (2). To investigate the effects of relaxin on Ang II-stimulated collagen expression, fibroblasts were treated with Ang II (1 x 10–7 M and 5 x 10–7 M) or a combination of rhRLX (100 ng/ml) and Ang II (1 or 5 x 10–7 M). To study the effects of relaxin alone on low-density cells, which rapidly differentiate to myofibroblasts (21), fibroblasts were also plated at 5/mm2 and grown for 7 d in the absence or presence of rhRLX (100 ng/ml). The rhRLX dose used in these experiments was based on previous studies on fibroblast cultures, whereby at this concentration, rhRLX demonstrated maximal effects on collagen expression (2, 3). Furthermore, the dose of rhRLX used was well within physiological levels of circulating relaxin in pregnant rats, which attain 50–100 ng/ml relaxin from d 14–20 of gestation and up to 180 ng/ml during parturition (22). For measurement of collagen synthesis, collagen was biosynthetically labeled with [3H]-proline in the presence of ascorbate and ß-aminopropionitrile, as previously described (2). For measurement of collagen deposition into the cell matrix, cell layers (per well) were isolated and hydrolyzed with 6 M hydrochloric acid for measurement of hydroxyproline content, as described previously (23). Hydroxyproline values were then converted to total collagen content by multiplying by a factor of 6.94 (based on hydroxyproline representing approximately 14.4% of the amino acid composition of collagen in most mammalian tissues) (24).
Determination of MMP activity from cardiac fibroblasts
To determine the effects of rhRLX on MMP expression, gelatin zymography of conditioned media was performed, as previously described (25). Briefly, the cells (1 x 105/cm2) were treated with either TGF-ß (2 ng/ml) or Ang II (10–7 M) in the absence or presence of rhRLX (100 ng/ml) for 72 h; the final 24 h of treatment were under serum-free conditions. Equal aliquots of the collected media were then analyzed on zymogram gels consisting of 7.5% acrylamide and 1 mg/ml gelatin, and the gels were subsequently treated as previously detailed (25). Clear bands indicated gelatinolytic activity.
Determination of cardiac fibroblast proliferation
In separate experiments, the effects of relaxin on IGF-I- or Ang II-stimulated atrial and ventricular fibroblast proliferation was investigated based on their reported abilities to stimulate fibrosis (12, 13) and potently increase cardiac fibroblast number (26, 27). Cardiac fibroblasts were plated at an equal density of 104 cells /96-well plate in DMEM-FBS. Twenty-four hours later, media were changed to DMEM supplemented with lactalbumin hydrolysate, and cells were labeled with [3H]-thymidine (3 µCi/ml) in the presence of rhRLX (100 ng/ml) alone or in combination with IGF-I (0.1 µg/ml, 0.5 µg/ml, and 1 µg/ml) or Ang II (10–8 M and 10–7 M) for 72 h. [3H]-thymidine incorporation and cell counts, using a hemocytometer, were evaluated.
Western blot analysis of -smooth muscle actin (-SMA)
Fibroblast differentiation into myofibroblasts was determined by -SMA expression. Ventricular fibroblasts were plated at a density of 2 x 105 cells/12-well plate and incubated with media (0.7 ml) alone, media containing TGF-ß (2 ng/ml) or TGF-ß (2 ng/ml), and rhRLX (100 ng/ml) for 72 h of culture. Collected media were stored at –80 C. Cell layers were trypsinized, and the protein was extracted using Trizol reagent (according to the manufacturer’s instructions; Life Technologies, Gaithersburg, MD) and analyzed by the Bio-Rad dye-binding protein assay (Bio-Rad, Richmond, CA).
Protein extracts (in 1% sodium dodecyl sulfate; 10 µg total protein/lane) were electrophoresed under nonreducing conditions on 12.5% acrylamide gels, as previously described (28). Western blot analysis was performed with a monoclonal antibody to -SMA (M0851, 1:1000 dilution; Dako Corp., Carpinteria, CA) overnight and a goat antimouse IgG secondary antibody (1:2000 dilution; Dako) for 2 h. Densitometry of the -SMA bands was performed using a Bio-Rad GS710 Calibrated Imaging Densitometer and Quantity-One software (Bio-Rad).
RT-PCR analysis of relaxin and relaxin receptor expression in cardiac cells
Freshly isolated cells or cells plated in six-well plates and grown to confluency in DMEM-FBS were used for RNA extraction and analysis. RNA extraction and RT-PCR of relaxin-1 (homologous to human relaxin-2) and LGR7 gene expression in atrial myocytes and fibroblasts, ventricular myocytes and fibroblasts, and VSMCs of rats (n = 3–4 samples per cell type) were performed as previously described (29, 30). Rat cardiac cells were screened for relaxin-1 using primer sequences detailed previously (29). Relaxin-3 expression was also determined using specific forward (5'-TGCGGAGGCTCACGATGGCGC-3') and reverse (5'-ATAGCTGACAGCAGGTTGGAC-3') primers. LGR7 expression was determined using the following primers: forward, 5'-GTGTATCCTTTTCGGTGTTTAAGG-3'; and reverse, 5'-GAATAAGAATTGAGTCTAGATGAC-3'. For relaxin-1 gene expression, an annealing temperature of 55 C (40 cycles) was used; for relaxin-3 expression, an annealing temperature of 58 C (40 cycles) was used; and for LGR7 expression, the following (touch-down) annealing temperatures were used: 54 C (two cycles), 52 C (two cycles), 50 C (two cycles), and 48 C (34 cycles). Glyceraldehyde-3-phosphate dehydrogenase was used in separate PCR reactions to control for quality and equivalent loading of the cDNA. Aliquots of the PCR products were electrophoresed on 2% (wt/vol) agarose gels stained with ethidium bromide and photographed.
cAMP bioassay
The ability of rhRLX to induce cAMP production by rat ventricular fibroblasts was also determined using Ang II-stimulated and unstimulated ventricular fibroblasts. Fibroblasts were plated at a density of 4 x 104 cells/cm2 in DMEM-FBS and then incubated with or without Ang II (10–7 M) for 72 h. Before cAMP measurement, cells were preincubated for 15 min with isobutylmethylxanthine (50 µM). rhRLX (10 nM) alone, forskolin (1 µM) alone, or a combination of rhRLX and forskolin were then added to cells, before intracellular cAMP was assayed as previously described (30) following cell lysis at 2, 5, 10, 15, and 30 min after incubation.
Relaxin treatment in vivo by osmotic mini-pumps
The ability of rhRLX to reduce myocardial collagen accumulation in vivo was investigated in two models of cardiac fibrosis, one due to relaxin disruption (16) and the other to cardiac-restricted overexpression of ß2-AR (19). The age at which cardiac fibrosis was already established in these animals (12 months of age in male relaxin-deficient mice and 5 months of age in male ß2-AR transgenic mice) was chosen for treatment studies based on our previous observations (16, 19). For each model, wild-type littermates were used as controls. Osmotic mini-pumps (model 2002; Alza Corp., Cupertino, CA), loaded with either vehicle (20 mM sodium acetate buffer, pH 5.0) or rhRLX (500 µg/kg·d) were implanted sc for 14 d, which produced circulating relaxin concentrations of 20–40 ng/ml in these animals (3, 6), which is within physiological levels of serum relaxin in pregnant rodents (1, 22). The dose of rhRLX used in these studies was identical to that used to treat pulmonary fibrosis in relaxin-deficient (6) and bleomycin-treated (3) mice. Male relaxin-null mice (n = 4 treated with vehicle, n = 4 treated with rhRLX), ß2-AR transgenic mice (n = 8 treated with vehicle, n = 8 treated with rhRLX), and their respective wild-type mice (n = 8 for each group) were used for analysis. After 14 d, ventricular tissues were collected and used for hydroxyproline analysis, as previously described (23).
Statistical analysis
The results were analyzed using one-way ANOVA and the Newman-Keuls test for multiple comparisons between groups. All data in this article are presented as the mean ± SEM, with P < 0.05 described as statistically significant.
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), which was consistent with our previous findings (29). Relaxin-3 mRNA expression, however, was clearly identified in all atrial and ventricular cells studied (Fig. 1A), suggesting that it represents the predominant form of relaxin in the rat heart. LGR7 gene expression was detected in freshly isolated atrial and ventricular myocytes but was either not detected (2-d cultured atrial myocytes) or decreased (2-d cultured ventricular fibroblasts) in cultured myocytes (Fig. 1B). LGR7 was expressed in atrial and ventricular fibroblasts, and this expression persisted during culture (from passages 1–4). LGR7 transcripts were not detected in 2-d cultured VSMC (Fig. 1B). These findings demonstrated that neonatal cardiac fibroblasts expressed LGR7 and should, therefore, respond to relaxin. A comparison of the amino acid sequence of rat relaxin-1, rat relaxin-3, and rhRLX demonstrates a conserved receptor binding domain (of relaxin) that interacts with the LGR7 receptor (Fig. 1C).
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). Similarly, Ang II (5 x 10–7 M, 1 x 10–7 M) alone increased collagen secretion by 15–20% and 50–60% (P < 0.05), respectively (Fig. 2A). rhRLX (100 ng/ml) alone had no effect on collagen expression in fibroblasts, as previously described (2, 3), but significantly inhibited the TGF-ß- (by 60–65%, P < 0.05; Fig. 2A) and Ang II- (by 70–80%, P < 0.05, Fig. 2A) induced effects on collagen secretion over a 72-h period.
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). However, rhRLX (100 ng/ml) treatment of cardiac fibroblasts in the presence of TGF-ß (2 ng/ml) significantly increased both the latent (by 55–60%, P < 0.01) and active (by 20–25%, P < 0.01) forms of MMP-2, while having no marked effects on MMP-9 expression over 72-h of culture (Fig. 2B). rhRLX also increased MMP-2 expression in the presence of Ang II (by 10–30% of that from untreated cultures, P < 0.05) over 3 d in culture (data not shown).
Collagen deposition into the matrix of fibroblasts was also up-regulated by TGF-ß (65–85%, P < 0.05; Fig. 2C) and Ang II (20–50%, P < 0.05; Fig. 2C). As with its effects of collagen secretion into the media, rhRLX (100 ng/ml) alone had no effect on collagen deposition into the matrix when applied to higher-density (atrial and ventricular) cells, but rhRLX significantly decreased collagen deposition stimulated with TGF-ß (P < 0.05) or Ang II (P < 0.05). rhRLX (100 ng/ml) treatment of low-density fibroblast cultures (which accelerate the differentiation of fibroblasts to myofibroblasts) also significantly (P < 0.05) inhibited the collagen content of the ECM by 38% compared with the collagen content from untreated cultures (Fig. 2C).
Relaxin inhibits Ang II- and IGF-I-stimulated fibroblast proliferation
The ability of rhRLX to modulate cardiac fibroblast proliferation was investigated in the absence or presence of Ang II and IGF-I, which both potently stimulate fibroblast proliferation. Ang II significantly increased atrial and ventricular fibroblast proliferation, as determined by cell counts and [3H]-thymidine incorporation, when administered at concentrations of 10–8 M (63–75%, P < 0.01) and 10–7 M (64–73%, P < 0.01) over a 72-h culture period (Fig. 3A). IGF-I also significantly augmented fibroblast proliferation at doses of 0.1 µg/ml (27–30%, P < 0.05), 0.5 µg/ml (36–40%, P < 0.05), and 1.0 µg/ml (34–40%, P < 0.05) over 3 d of culture (Fig. 3B). rhRLX (100 ng/ml) alone had no significant effects on basal fibroblast proliferation but significantly inhibited the Ang II- (by 15–30%, P < 0.05; Fig. 3A) and IGF-I- (by 25–50%, P < 0.05, Fig. 3B) mediated effects on cell proliferation.
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-SMA expression by ventricular fibroblasts. In all experiments, an equal amount of total protein (10 µg/sample) was analyzed. Consistent with our earlier observations, TGF-ß (2 ng/ml) and Ang II (10–7 M) significantly increased fibroblast differentiation, as detected by Western blotting of -SMA protein. -SMA expression increased by 85–95% with TGF-ß (P < 0.01) and 25–35% with Ang II (P < 0.05) over 72 h of culture (Fig. 4). rhRLX (100 ng/ml) significantly inhibited the TGF-ß-induced effects on -SMA expression (by 65%, P < 0.05). Similarly, rhRLX significantly inhibited the Ang II-mediated increase in -SMA (by 88%, P < 0.05; Fig. 4).
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Relaxin decreases collagen accumulation in two rodent models of cardiac fibrosis
To extend the in vitro findings of inhibited cardiac collagen synthesis and deposition by rhRLX, the effects of rhRLX were determined on two models of cardiac fibrosis in vivo. Cardiac fibrosis was established in relaxin-deficient mice by 12-months of age, resulting in a 30% (P < 0.05) increase in collagen concentration (Fig. 5A). Myocardial fibrosis was more rapidly established in ß2-AR transgenic mice at 5 months of age, as evidenced by a 60% increase (P < 0.001) in collagen concentration (Fig. 5B). This increment in myocardial collagen concentration over respective wild-type control values was significantly reduced by rhRLX (500 µg/kg·d) treatment of animals over 14 d by 40% (P < 0.05) in relaxin-deficient mice (Fig. 5A) and by 58% (P < 0.01) in ß2-AR transgenic mice (Fig. 5B).
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Discussion |
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In the normal heart, cardiac fibroblasts are relatively quiescent, but upon pathological stimuli to the heart (8, 9, 10, 11, 12), they proliferate and differentiate into activated myofibroblasts. These myofibroblasts, the hallmarks of which are -SMA expression and production of prodigious amounts of collagen, are important contributors to fibrosis and the pathology of the heart (31, 32). In the diseased heart, a number of autocrine, paracrine, and neuronal hormone systems have been found to be activated, and several of these systems, including TGF-ß, Ang II, catecholamines, and endothelin-1, are known to directly stimulate fibroblast activation and collagen production (31, 32, 33, 34). Thus, inhibitors that are able to block such profibrotic signaling are expected to have therapeutic potential against cardiac fibrosis. Interestingly, in this study, we have shown that relaxin inhibits fibroblast activation and collagen synthesis that were stimulated by two of these factors (TGF-ß and Ang II) in vitro, and such action was consistent with observations made in vivo. These findings are consistent with relaxin’s ability to inhibit TGF-ß- and IL-1ß-induced collagen production by human dermal and lung fibroblasts (2, 3) and with relaxin’s ability to antagonize endothelin-1 activity in a number of human-derived cells (35).
The collagen-lowering effects of rhRLX observed in this study in vivo and in vitro are similar to relaxin’s ability to decrease collagen expression by human dermal (2) and lung (3) fibroblasts and in other organs from animal models of fibrosis (3, 4, 5, 6). In those studies, as well as in the current investigation, relaxin’s ability to decrease fibrosis was the result of its ability to directly inhibit collagen secretion, as well as its potential ability to induce collagen degradation by increasing the release of MMPs (2, 3, 4, 5). MMP-2 has been shown to cleave type I collagen (36), as well as fibronectin and nonfibrillar collagens. Therefore, it is likely that the deposition of a number of interstitial collagens and ECM molecules associated with fibrosis could be influenced by relaxin via its ability to induce MMP-2 activation.
Previous work has demonstrated that IGF-I, as well as Ang II, can stimulate cardiac fibroblast proliferation and may contribute to fibrosis (26, 27). Our data suggest that relaxin’s ability to inhibit organ fibrosis may also be attributable to its antiproliferative effects on fibroblasts, thus keeping in check the number of those cells that synthesize and secrete collagen. Therefore, our findings now demonstrate that relaxin may be able to modulate organ fibrosis in the following ways: 1) by inhibiting the activation of fibroblasts, as assessed by expression of -SMA (a marker of myofibroblasts); 2) by inhibiting the proliferation of activated fibroblasts; 3) by antagonizing collagen deposition by activated fibroblasts; and 4) by increasing collagen degradation via activation of MMPs. It is of interest to note that relaxin did not influence cardiac collagen expression under basal conditions when applied over a short-term period in vitro. Thus, relaxin only reduced cardiac collagen synthesis and accumulation in activated cells (myofibroblasts) when stimulated by a number of factors.
Relaxin’s ability to stimulate sustained increases in cAMP when applied to other cells types has been well documented (30, 37, 38), and the relaxin receptor (LGR7) has been clearly demonstrated to couple to Gs proteins and cAMP accumulation (18). However, relaxin was previously shown not to influence cAMP when applied to human uterine fibroblasts over a 15-min period (37). We investigated whether relaxin stimulation of cardiac fibroblasts was coupled to sustained increases in cAMP. Our data demonstrated that relaxin was only able to induce a weak transient rise in cAMP over a 2- to 5-min period, which returned to basal levels after 10 min, but this occurred in only two of four experiments. Therefore, relaxin stimulation is not coupled to sustained increases in cAMP in fibroblasts, which is consistent with the study on human uterine fibroblasts (37). Furthermore, the weak transient rise in cAMP in two experiments was also seen in normal fibroblasts not stimulated with Ang II; hence, the mechanism by which relaxin reverses myofibroblast phenotype is likely to be independent of cAMP signaling. Therefore, it is likely that relaxin induces its effects via other mechanisms. Indeed, relaxin has been shown to modulate a number of its effects in other target cells via the MAPK pathway (37, 38, 39) and induce MAPK phosphorylation in human uterine fibroblasts (37). The potential interactions of relaxin signaling with that of Gq-mediated activation of Ras/Raf/MAPK/ERK by Ang II (40) and activation of MAPK and ERK by TGF-ß (41) are the subject of future investigation.
In this study, we used two murine models of fibrotic cardiomyopathy to test whether relaxin treatment could reverse already established fibrosis in the heart. Our data indicated that relaxin treatment was associated with a similar and significant reduction in collagen content in both models within only a 2-wk period of treatment with relaxin. It might be argued that treatment of the relaxin-null mice with exogenous relaxin would simply be replacing the hormonal deficit and that the results on cardiac fibrosis are not unexpected. However, the selection of animals at 12 months of age was made to ensure that cardiac fibrosis was already well established in these animals (16). Furthermore, this efficacy of reversing fibrosis was found using another strain of mouse in which the etiology of cardiac fibrosis is independent of relaxin (i.e. due to cardiac overexpression of ß2-ARs). Although treatment with drugs like angiotensin converting enzyme inhibitors have been reported to reduce the amount of collagens in the heart (42), such efficacy usually requires much longer periods of treatment. Thus, the significant reduction in collagen expression by short-term relaxin treatment is encouraging for this peptide as a therapeutic agent, specifically for the reversal of cardiac fibrosis.
In conclusion, we have demonstrated that relaxin inhibits cardiac collagen secretion and deposition by directly inhibiting cardiac fibroblast proliferation, differentiation, and activation and increasing MMP activity. Relaxin also markedly reversed collagen overexpression in two models of cardiac fibrosis in vivo. Of great interest was the ability of relaxin to induce such potent and rapid antifibrotic effects in the heart over a 2-wk period. Thus, relaxin provides an important means to regulate excessive collagen deposition in heart diseases characterized by fibrosis.
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Acknowledgments |
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Footnotes |
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Present address for E.N.U.: BAS Medical Inc., San Mateo, California 94402.
Abbreviations: ß2-AR, ß2-Adrenergic receptor; Ang II, angiotensin II; DMEM-FBS, DMEM supplemented with 10% fetal calf serum, penicillin, and streptomycin; ECM, extracellular matrix; MMP, matrix metalloproteinase; rhRLX, recombinant human gene-2 relaxin; -SMA, -smooth muscle actin; VSMC, vascular smooth muscle cells.
Received February 18, 2004.
Accepted for publication May 10, 2004.
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, P < 0.05 compared with values from TGF-ß- or Ang II-treated cells. Additionally, rhRLX (100 ng/ml) treatment of low-density cells (5/mm2) over 7 d caused an inhibition of collagen deposition. Results are presented as the mean ± SE of relative collagen content from three separate experiments (six assays per group from each experiment). 
