1Department of Nephrology, Royal Melbourne Hospital, 2Department of Medicine, University of Melbourne and 3Howard Florey Institute, Melbourne and 4Microvascular Biology and Wound Healing Group, RMIT University, Bundoora, Victoria, Australia
Correspondence and offprint requests to: Rosemary Masterson, Department of Nephrology, Royal Melbourne Hospital, Grattan Street, Victoria 3050, Australia. Email: Rosemary.Masterson{at}mh.org.au
![]() |
Abstract |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Methods. Rat cortical fibroblasts were obtained from outgrowth culture of renal tissue isolated from kidneys 3 days post-unilateral ureteric obstruction and constituted 100% of cells studied. A relaxin radio-receptor assay was used to establish binding of relaxin to renal fibroblasts in vitro. Functional studies then examined the effects of H2 relaxin (0, 1, 10 and 100 ng/ml) on fibroblast kinetics, expression of alpha-smooth muscle actin (-SMA), total collagen synthesis, collagenase production and collagen-I lattice contraction. CTGF mRNA expression was also measured by northern analysis.
Results. H2 relaxin bound with high affinity to rat renal fibroblasts, but receptor numbers were low. Consistent with its previously reported bimodal effect, transforming growth factor (TGF-ß1) reduced fibroblast proliferation, an effect abrogated by H2 relaxin. Fibroblasts exposed to H2 relaxin (100 ng/ml) for 24 h demonstrated decreased immunostaining for -SMA and reduced
-SMA protein expression compared with controls. There was a trend for a relaxin-mediated reduction in total collagen synthesis and
1(I) mRNA expression with large dose-related increases in collagenase protein expression being observed. TGF-ß1-stimulated collagen-I lattice contraction was significantly inhibited following co-incubation with 100 ng/ml relaxin. Incremental doses of H2 relaxin had no significant effect on CTGF mRNA expression.
Conclusions. The findings of this study suggest that the antifibrotic effects of relaxin involve down-regulation of fibroblast activity, increase in collagenase synthesis and restructuring of collagen-I lattices, which are consistent with its known physiological role of matrix remodelling. Although there appears to be an interaction between TGF-ß1 and H2 relaxin, this does not appear to involve a reduction in CTGF mRNA expression.
Keywords: contraction; fibroblast; matrix remodelling; relaxin; renal; transforming growth factor-ß1
![]() |
Introduction |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
An overwhelming body of evidence has shown that interstitial fibroblasts are important effector cells in renal fibrosis [1]. Although phenotypic markers for fibroblasts are poorly characterized, several groups have recognized activated fibroblasts, so-called myofibroblasts, by their de novo expression of the smooth muscle-associated protein, alpha-smooth muscle actin (-SMA) [2,3]. While widely acknowledged for their matrix-synthesizing and collagenase-producing properties, the less well reported contractile properties of these cells are being increasingly recognized as integral to the overall scarring process. Based on this knowledge, potential antifibrotic strategies have focused on blocking naturally occurring activators of these cells or directly down-regulating their function.
Relaxin, a peptide hormone and member of the insulin-like growth factor family, has traditionally been associated with growth and remodelling of the female reproductive tract during pregnancy [4]. Following the availability of recombinant human gene (H2) relaxin, there has been increasing interest in the potential therapeutic application of this hormone as an antifibrotic agent. In vitro studies of stimulated dermal [5], lung [6] and hepatic [7] fibroblasts have demonstrated that relaxin can reduce type-I and -III collagen production, increase procollagenase synthesis and reduce TIMP-1 expression. In vivo, recombinant human relaxin has been shown to reduce the extent and severity of scarring in a number of experimental models of non-renal [6] and renal [8,9] fibrosis. The administration of relaxin in a rodent model of Bromoethylene acetate (BEA)-induced renal fibrosis was associated with a 75% decrease in collagen deposition at the corticomedullary junction, with resultant preservation of renal function compared with controls [8].
The objective of this study was to further elucidate the in vitro effects of relaxin on renal fibroblasts. Based on the observation that relaxin can reduce established fibrosis following injury, we particularly focused on its potential matrix-degrading and remodelling effects. Previous studies have reported the effect of relaxin to be amplified by co-stimulation of dermal fibroblasts with transforming growth factor (TGF)-ß1 [5]. To further establish the relationship between these two cytokines, we looked at the effects of relaxin on the synthesis of CTGF, a putative downstream mediator of TGF-ß1.
![]() |
Subjects and methods |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Cultures were subsequently exposed to recombinant H2 relaxin (Connetics Corp., Palo Alto, CA, USA), with functional parameters being measured as below.
Experimental unilateral ureteric obstruction
Under inhalational general anaesthesia, the left ureter of male SpragueDawley rats (250300 g) was ligated in two places using 3/0 gauge surgical silk (Davis-Geck, Melbourne, Vic., Australia) and then cut between the ligatures. The contralateral kidney was left intact. Animals were sacrificed 3 days post-procedure, with removal of renal tissue for explant culture.
Cell isolation and characterization
Renal interstitial fibroblasts were propagated using explanting methods described previously [10]. Renal tissue was collected in ice-cold Hank's salt solution treated with gentamicin (ICN Pharmaceuticals, Costa Mesa, CA, USA). Cultures were established by cutting tissue into gelatin (Sigma, St Louis, MO, USA)-coated Petri dishes and covering with DMEM media (CSL, Parkville, Vic., Australia) supplemented with 20% fetal calf serum (FCS; CSL) and penicillin/streptomycin (ICN). Tissue was maintained at 37°C in 5% CO2/95% air, with media being changed on day 3. Once confluent, primary outgrowth cells were trypsinized and subcultured for subsequent experiments.
Cell outgrowths were characterized as myofibroblasts based on their immunohistochemistry and growth characteristics. For immunohistochemistry, cells were grown on coverslips (Nunc, Roskilde, Denmark) and fixed in methanol. Cells were consecutively immersed in 3% H2O2 in methanol to quench endogenous peroxidase activity, incubated with primary antisera against vimentin, cytokeratin, SMA and desmin (Dako, Carpenteria, CA, USA), rinsed in phosphate-buffered saline (PBS), incubated with appropriate species-specific biotinylated secondary antisera (Vector Laboratories, Burlingame, CA, USA), washed in PBS and incubated with the avidinbiotin complex (Vector) and diaminobenzidine (DAB; Dako). DAB-enhancing solution (Vector) was used to enhance the reaction product. Subcultured cells were universally stellate in appearance and positive for the mesenchymal marker vimentin, but negative for endothelial (RECA), epithelial (cytokeratin) and macrophage markers (ED-1). Fibroblasts constituted 100% of cells, of which 60% were positive for the myofibroblast marker SMA, with only occasional cells expressing desmin (myofibroblast and mesangial cells).
Cell viability assays
Confluent fibroblasts in 96-well plates (Nunc) were incubated for 24 h in media supplemented with 0, 1, 10 and 100 ng/ml of recombinant H2 relaxin (Connetics Corp., Palo Alto, CA, USA), as appropriate. After incubation with 50 µl of 5 mg/ml stock 3-(4,5-dimethylthiazole-2-yl)-2,5-diphenyl tetrazolium bromide (MTT; Sigma) for 4 h, cells were lysed by the addition of 200 µl dimethyl sulfoxide and 25 µl Sorensen's glycine buffer (0.1 M glycine, 0.1 M NaCl, pH 10.5). The purple formazan reaction product was measured at 570 nm with an enzyme-linked immunosorbent assay plate reader (Tecan, Salzberg, Austria).
Relaxin receptor identification
Receptor binding assays. The ability of recombinant human relaxin to bind to rat renal fibroblasts was tested in a relaxin radio-receptor assay. H2 relaxin was labelled with [33P] using the catalytic subunit of cyclic AMP-dependent protein kinase. Medium was removed from the cells followed by washing with PBS before pre-incubation in 300 µl binding buffer [20 mM HEPES, 50 mM NaCl, 1.5 mM CaCl2, 1% bovine serum albumin (BSA), 0.1 mg/ml lysine, 0.01% NaN4, pH 7.5]. Binding studies were performed with 100 µl 33P-labelled H2 relaxin (100 pM) and 100 µl of increasing concentrations of competitor or blank in binding buffer at 25°C for 60 min. Non-specific binding was determined by an excess of H2 relaxin (1 µM). After incubation, the cells were washed with PBS followed by their recovery from the plates using 500 µl of 1 M NaOH, which was transferred to scintillation vials. Liquid scintillation cocktail (Ultima Gold, Packard, Meriden, USA) was added and the vials were counted in a liquid scintillation analyser (Packard 1900 TR). The assay was performed in quadruplicate and is represented as mean±SEM of the pooled data and plotted using one-site competition function of the PRISM programme (Graphpad Inc., San Diego, USA). LIGAND analysis was performed on one representative experiment.
RTPCR analysis of LGR7 and LGR8 expression
Total RNA extraction from cells and tissues, reverse transcription of RNA, reverse transcriptionpolymerase chain reaction (RTPCR) methodology and subsequent isolation of products and sequencing on both strands has been described [11]. The specific primers for RTPCR were LGR7 (fwd: 5'-GTGTATCCTTTTCGGTGTTTAAGG-3'; rev: 5'-TTCCACCCAGATGAATGATGG-3'), LGR8 (fwd: 5'-GATGGGGGTGTACCTGTTCTC-3'; rev: 5'-AGTAGACAGGTGTGTTACCTT-3') and GAPDH (fwd: 5'-tgatgacatcaagaaggtgg-3'; rev: 5'-GTTTCTTACTCCTTGGAGGCC-3'), which generated 627, 803 and 247 bp products, respectively.
Cell growth
The effect of relaxin and TGF-ß1 on serum-stimulated cell growth was evaluated by measuring DNA synthesis (proliferation) and total cell counts (cell kinetics).
DNA synthesis
[3H]Thymidine incorporation was used to determine the effects of relaxin, TGF-ß1 and a combination of H2 relaxin plus TGF-ß1 on DNA synthesis. Equal numbers of cells were seeded in 36-mm2 multiwell plates (Nunc) (1 x 105/well) and were cultured for 24 h in DMEM + 10% FCS before arrest of cell growth with exposure to DMEM + 5% FCS for a further 24 h. Cells were subsequently incubated for 24 h in DMEM + 10% FCS containing 0, 1, 10 or 100 ng/ml relaxin, 2.5 ng/ml TGF-ß1 and a combination of 100 ng/ml relaxin and 2.5 ng/ml TGF-ß1. A 0.25 µCi aliquot of [3H]thymidine (Amersham, Little Chalfont, Bucks, UK) was added to each well for 24 h. Cell membranes were lysed using 0.25 M sodium hydroxide and removed from plates using a cell scraper (Nunc). Scintillation counts were read on a LS3801 ß-counter (Beckman, Palo Alto, CA, USA). Experiments were performed in triplicate and data are expressed as counts per minute.
Total cell counts
Cell kinetics was determined by measuring total cell number over a 5 day growth period. Cells were trypsinized and seeded into 6-well plates (Costar) at a density of 1 x 105 cells per well in DMEM + 10% FCS (n = 5 per dose tested). Following cell adhesion, fresh medium containing relaxin in incremental doses (1, 10 and 100 ng/ml) was added. This medium was changed after 3 days. After 5 days, cell number was estimated by measuring the ability of each well to reduce MTT, as above.
Collagen synthesis
The effect of H2 relaxin, TGF-ß1 and a combination of H2 relaxin plus TGF-ß1 on collagen synthesis was quantified by measuring [3H]proline incorporation in collagenous proteins [10]. For each experimental group, wells of 36-mm2 multiwell plates (Costar) were seeded with 1 x 105 and grown to confluency in DMEM + 10% FCS at 37°C with 95% air/5% CO2. At confluence, cells were incubated for 72 h in DMEM + 10% FCS, 0.25 mM ascorbic acid (Sigma) and H2 relaxin (100 ng/ml), TGF-ß1 (2.5 ng/ml) or H2 relaxin (100 ng/ml) plus TGF-ß1 (2.5 ng/ml), as appropriate. At the end of this period, fresh media containing H2 relaxin, TGF-ß1 or H2 relaxin plus TGF-ß1 in the above doses were combined with 20 µCi [3H]proline (Amersham), added to the cells and incubated for a further 24 h. Protein from the medium fraction was precipitated overnight at 4°C by adding trichloroacetic acid (TCA; final concentration 10% w/v) in the presence of 0.04% proline and 0.1% BSA carrier with samples centrifuged to pellet protein. Protein pellets were washed once with 10% TCA/1 mM proline and then twice more with 5% TCA/1 mM proline. Pellets were then dissolved in 0.2 M NaOH and duplicate aliquots were transferred to microfuge tubes and neutralized by adding HEPES buffer and sufficient HCl to adjust to pH 7.4.
Reagent solutions were prepared with (+) or without () 10 µl of (50 units/ml) purified collagenase (Worthington CSPA, Lakewood, NJ, USA) in 10 µl 25 mM CaCl2 and 20 µl 62.5 mM N-ethylmaleimide. Reagent volume was made up to 40 µl with 0.05 M Tris (pH 7.6). Reagent solutions were added to duplicate samples and digestion allowed to proceed for 90 min at 37°C. Protein was then precipitated with 20% TCA/0.5% tannic acid, the vials centrifuged and supernatants transferred to scintillation vials together with a wash of 0.5 ml 5% TCA. Ten millilitres of scintillation fluid (Starscint; Packard, Meriden, CT, USA) was added to each sample and radioactivity determined in a ß-counter (LS3890; Beckman). The difference in counts between samples prepared with and without collagenase provided a measure of the amount of radioactivity incorporated into newly synthesized collagen. Results are expressed as disintegrations per minute.
Western blotting of -SMA and collagenase
Confluent fibroblasts in 36-mm diameter multiwell plates (Costar) were cultured overnight in DMEM + 10% FCS supplemented with or without incremental doses of H2 relaxin. The presence of cellular -SMA and interstitial collagenase were quantitated using western blotting and appropriate murine monoclonal antisera. The
-SMA antiserum (Dako) recognizes a single band of
43 kDa. Anti-human matrix metalloproteinase (MMP)-1 antiserum (Calbiochem, San Diego, CA, USA), which recognizes a 57 kDa band corresponding to active MMP-1, was used to quantitate expression of MMP-13, the rat orthologue of MMP-1 [12]. Cross reactivity was confirmed by western blotting of rat fibroblast-derived protein and a MMP-13-positive control (Calbiochem), with subsequent immunostaining with anti-MMP-1 antiserum (Calbiochem) demonstrating a single 57 kDa band in both samples (data not shown).
Cell-associated protein was isolated from the organic fraction of Trizol (Gibco Life Technologies, Rockville, MD, USA). Protein concentration was estimated by protein assay (Bio-rad, Hercules, CA, USA) and equal quantities of protein were made up to uniform volume before being electrophoresed on a 4.4%/12.5% sodium dodecyl sulphatepolyacrylamide gel and electroblotted on Hybond nitrocellulose membrane (Amersham). The membrane was blocked in 0.01 M PBS containing 5% skimmed milk powder (Bonlac, Melbourne, Vic., Australia), 0.01% Tween-20 (Bio-rad) and incubated for 1 h with 1:1000 dilution anti--SMA (Dako) or 1:1000 anti-MMP-1 (Calbiochem), as appropriate. After washing, the membrane was incubated with a 1:2000 dilution of horseradish peroxidase-conjugated secondary antiserum (Dako) in PBSTween. Blots were developed using the ECL detection kit (Amersham) to produce a chemiluminescent signal, which was captured on X-ray film (Kodak, Rochester, NY, USA), according to manufacturer's instructions. Results were quantitated using computerized densitometry (NIH Image; Scion Corp., Fredrick, MA, USA) and are represented as the mean of triplicate assays.
Collagen lattice contraction assays
Collagen lattices were prepared by mixing cells with 1.8 ml acid-solubilized collagen-I (final collagen concentration: 2.1 mg/ml) (ICN Biochemicals) in 24-mm diameter cluster dishes (ICN Biochemicals), as described previously [13]. Lattices were solidified at 37°C with 95% air/5% CO2 before being dislodged from the sides of the dish with a sterile scalpel blade. Five control lattices and five lattices for each experimental group were poured. Lattices contained 1 x 106 cells in either DMEM + 10% FCS (control), DMEM + 10% FCS containing relaxin (100 ng/ml), DMEM + 10% FCS containing TGF-ß1 (2.5 ng/ml) or DMEM + 10% FCS with relaxin (100 ng/ml) and TGF-ß1 (2.5 ng/ml).
Cell-mediated contraction of collagen-I substrate, as determined by a decrease in lattice diameter, was measured at 24 and 48 h after lattice dislodgment. Contraction, determined by a decrease in lattice diameter, was measured using a metric ruler, as described previously [13]. We have previously validated diameter as a legitimate estimation of contraction. Lattice diameter was shown to be directly proportional to volume (unpublished data).
Northern hybridization for CTGF and procollagen-1(I)
Northern blot analysis was used to determine expression of CTGF and procollagen-1(I) mRNA. Total cellular RNA was isolated from confluent cell monolayers using Trizol reagent (Gibco Life Sciences Inc., Grand Island, NY, USA) in accordance with the manufacturer's instructions.
CTGF. A cDNA probe for CTGF was generated using RTPCR. Primers for CTGF were designed from the rat CTGF sequence (GenBank accession no. AF079531) and produced a single 318 bp PCR product, as predicted. RTPCR was performed on RNA extracted from a fibrotic rat kidney 7 days post-UUO. The 318 bp fragment was confirmed by electrophoresis before subcloning.
Procollagen-1(I). A 0.6 kb EcoRI/HindIII fragment of rat
1(I): cDNA from plasmid Pgem-3Z [10].
For northern hybridization, isolated RNA was denatured and electrophoresed through a 1% agarose gel containing formaldehyde (2.2 mol/l). Signals were normalized to ethidium bromide-stained gels to control for variation in loading. RNA for experimentation was then transferred and UV-fixed onto Hybond nylon membranes (Amersham). Before hybridization with [32P]-labelled cDNA probe prepared by multiprime® labelling (Amersham), RNA expression was quantified using densitometry with signals normalized to ethidium bromide-staining of the 28S band. Data are represented as the CTGF:28S ribosomal RNA ratio and the procollagen-1(I):28S ribosomal RNA ratio for each sample. Both experiments were performed in triplicate.
Statistical analysis
Data are represented as means±SD. Treatment groups were compared using KruskalWallis analysis of variance with Dunnett's correction for multiple comparisons (Sigma Stat; SPSS, Chicago, IL, USA).
![]() |
Results |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
|
Relaxin does not affect renal fibroblast mitogenesis
Dose response studies indicated that 1, 10 and 100 ng/ml H2 relaxin had no significant effect on [3H]thymidine incorporation and, therefore, on the proliferation of rat renal fibroblasts (P = NS, all doses vs control). TGF-ß1 (2.5 ng/ml) reduced fibroblast proliferation (P<0.05 vs control; n = 3), an effect abrogated by H2 relaxin (Figure 2A). There was no significant difference in cell population growth over 5 days in H2 relaxin-treated groups vs control (P = NS, all doses vs control; Figure 2B).
|
|
Effect of H2 relaxin and TGF-ß1 on total collagen synthesis and procollagen-1(I) mRNA expression
In renal fibroblasts cultured in DMEM + 10% FCS, basal levels of collagen synthesis, as assessed by [3H]proline incorporation, were not significantly altered by the addition of H2 relaxin (100 ng/ml). The addition of exogenous TGF-ß1 (2.5 ng/ml) resulted in a significant increase in collagen synthesis (P<0.05 vs control). Co-stimulation of cells with TGF-ß1 (2.5 ng/ml) and H2 relaxin (100 ng/ml) to a large extent prevented this effect (P = NS vs control; P = 0.06 vs TGF-ß1).
Cellular mRNA expression of procollagen-1(I) was measured using northern hybridization and a 32P-labelled cDNA sequence complementary to rat procollagen-
1(I). A representative northern blot is illustrated in Figure 4B. When the 4.7 kb mRNA transcript was normalized for 28S ribosomal RNA (Figure 4C), densitometry indicated that the TGF-ß1-induced increase in procollagen-
1(I) mRNA expression (P<0.05 vs control) was reduced by co-stimulation of H2 relaxin with TGF-ß1 (P = NS vs control; P = 0.07 vs TGF-ß1; Figure 4B).
|
|
|
|
![]() |
Discussion |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
The first step in further elucidating the effects of relaxin on rat renal fibroblasts was to establish that these cells could bind H2 relaxin. Recently, two orphan leucine-rich repeats containing G-protein-coupled receptors (LGRs) were shown to bind relaxin [14]. It was clearly demonstrated that the LGR7 receptor was a relaxin receptor, however, more recent data have shown that the LGR8 receptor is the receptor for the related peptide hormone INSL3 [15]. We investigated the expression of both of these receptors in rat renal fibroblasts in comparison with known LGR7- (cerebral cortex) and LGR8- (gubernaculum) expressing tissues. As expected, the LGR7, but not the LGR8, receptor mRNA is expressed in renal fibroblasts. Further evidence for the presence of functional relaxin receptors in renal fibroblast cells is the presence of high-affinity relaxin-binding sites, as determined by radio-receptor assays. The receptor numbers measured on renal fibroblasts were low (1100 receptors/cell), but are consistent with numbers seen in THP1 cells (250 receptors/cell) and human lower uterine segment fibroblasts (3220 receptors/cell) [16]. All fibroblasts used in this study were derived from renal tissue explanted from a male rat. It is possible that the number of relaxin receptors or receptor affinity may be different in fibroblasts derived from female animals or that receptor activity may be influenced by the presence or absence of oestrogen. These issues were not addressed in this study.
Myofibroblasts are well characterized in most, if not all forms of progressive renal disease where their presence correlates with disease progression. These cells are recognized by the presence of -SMA, which has long been thought of as a surrogate marker of activated fibroblasts [1]. By inference, the results of western blotting demonstrating a decrease in total
-SMA expression, suggest that relaxin can down-regulate the activity of these matrix-producing cells. Immunohistochemistry was used to determine if the reduced
-SMA expression was reflected by a change of cell phenotype. We confirmed that following 24 h of exposure to relaxin, 24% of fibroblasts originally immunostaining for
-SMA had lost this phenotypic marker. These findings are consistent with a relaxin-mediated reversal of myofibroblast differentiation.
Relaxin has previously been found to abrogate the effects of TGF-ß1 [5,8], recognized as a pivotal driver of glomerulosclerosis and tubulointerstitial fibrosis in renal disease. In the case of fibroblasts, the profibrotic effects of TGF-ß1 are mediated through its ability to reduce matrix degradation while simultaneously stimulating matrix protein production [17] and increasing the contraction of the surrounding extracellular matrix [13].
Large increases in interstitial collagenase synthesis were seen following exposure to H2 relaxin, consistent with both its physiological role in pregnancy and its ability to abrogate scarring by reducing the net accumulation of collagen in response to injury.
There was a trend for relaxin to prevent the TGF-ß1-mediated increase in collagen synthesis. Given the demonstrated ability of H2 relaxin to ameliorate other effects of TGF-ß1 and in the light of evidence from previous in vitro studies, which clearly show that H2 relaxin does reduce type-I and -III collagen synthesis [57], it is likely that similar effects on renal fibroblast-mediated total collagen production occur.
Although matrix synthesis is important in the early stages of chronic inflammation, the contraction of the surrounding matrix and the subsequent loss of parenchymal volume play a dominant role in the later stages of the scarring process. We have previously demonstrated a temporal relationship between renal myofibroblasts and reducing scar size, suggesting that fibroblast-mediated contraction of interstitial collagens contributes to scar formation by increasing the relative density of the extracellular matrix [13]. The contraction of scar tissue relies on the ability of fibroblasts to organize and contract collagen fibres by tractional structuring, a process mediated by 2ß1 integrins [18]. The demonstrated ability of H2 relaxin to reduce interstitial matrix contraction is of relevance particularly in the setting of established fibrosis. The prevention of contraction-mediated parenchymal collapse might allow preservation of the remaining functional tissue.
It is well established that TGF-ß1 is a bimodal regulator of cell growth, with a number of studies showing it might have both stimulatory and inhibitory effects on fibroblast mitogenesis [19,20]. Recently, it has been reported that low levels of TGF-ß receptor activation are sufficient to stimulate fibroblast proliferation, whereas higher degrees of receptor activation are associated with growth inhibition [19]. In this study, we have demonstrated a significant inhibitory effect of TGF-ß1 on fibroblast proliferation, an effect abolished by co-stimulation with H2 relaxin. This result further supports the widely reported interaction between these cytokines.
To further elucidate the relationship between TGF-ß1 and H2 relaxin, we studied the effects of incremental doses of relaxin on CTGF, a putative downstream mediator of TGF-ß1 activity. No effect of incremental doses of H2 relaxin on CTGF mRNA expression was found (Figure 7).
In summary, in renal fibroblasts, the most striking in vitro effects of H2 relaxin are an increase in collagenase activity, the inhibition of collagen-I lattice contraction and the down-regulation of fibroblast activity as determined by -SMA expression. These effects, which are consistent with the known physiological role of relaxin in matrix remodelling, might have therapeutic application not only in the prevention of fibrosis following injury, but also in repair and tissue remodelling following established fibro-contractive disease.
Conflict of interest statement. None declared.
![]() |
References |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|