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Endocrinology, doi:10.1210/en.2006-1324
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Endocrinology Vol. 148, No. 4 1582-1589
Copyright © 2007 by The Endocrine Society

Relaxin Antagonizes Hypertrophy and Apoptosis in Neonatal Rat Cardiomyocytes

Xiao-lei Moore, Su-ling Tan, Chen-yi Lo, Lu Fang, Yi-Dan Su, Xiao-Ming Gao, Elizabeth A. Woodcock, Roger J. Summers, Geoffrey W. Tregear, Ross A. D. Bathgate and Xiao-Jun Du

Baker Heart Research Institute (X.L.M., S.L.T., C.Y.L., L.F., Y.D.S., X.M.G., E.A.W., X.J.D.), Melbourne, Victoria 8008, Australia; Howard Florey Institute of Experimental Physiology and Medicine (G.W.T., R.A.D.B.), University of Melbourne, Melbourne, Victoria 3010, Australia; and Department of Pharmacology (R.J.S.), Monash University, Clayton, Victoria 3800, Australia

Address all correspondence and requests for reprints to: X. L. Moore, Baker Heart Research Institute, P.O. Box 6492, St. Kilda Road Central, Melbourne, Victoria 8008, Australia. E-mail: shirley.moore{at}baker.edu.au.


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
The pregnancy hormone relaxin has recently been shown to be cardio-protective. Despite its well-established antifibrotic actions in the heart, the effects of relaxin on cardiomyocytes (CM) remain to be determined. We investigated effects of isoform 2 of the human relaxin (H2-relaxin) on CM hypertrophy and apoptosis. In cultured neonatal rat CM, phenylephrine (50 µM) and cardiac fibroblast-conditioned medium were used respectively to induce CM hypertrophy. The degree of hypertrophy was indicated by increased cell size, protein synthesis and gene expression of atrial natriuretic peptide. Although H2-relaxin (16.7 nM) alone failed to suppress hypertrophy induced by phenylephrine, it repressed the cardiac fibroblast-conditioned medium-induced increase in protein synthesis by 24% (P < 0.05) and reversed the increase in cell size (P < 0.001) and atrial natriuretic peptide expression (P<0.01). We further studied the effect of H2-relaxin on CM apoptosis induced by H2O2 (200 µM). Studies of DNA laddering and nuclear staining demonstrated that H2-relaxin treatment reduced H2O2-induced DNA fragmentation. Real-time PCR and Western blot analysis revealed a significant increase in the Bcl2/Bax ratio in H2-relaxin-treated CM. Further analysis showed that activation of Akt (1.8-fold, P< 0.001) and ERK (2.0-fold, P<0.01) were involved in the antiapoptotic action of H2-relaxin in CM, and that Gi/o coupling of relaxin receptors was associated with the H2-relaxin-induced Akt activation in CM. In conclusion, these results extend our current knowledge of the cardiac actions of relaxin by demonstrating that H2-relaxin indirectly inhibits CM hypertrophy and directly protects CM from apoptosis.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
RELAXIN, A 6-kDa PEPTIDE hormone belonging to the IGF family, has long been regarded as a pregnancy hormone since its discovery in 1926 (1). It is primarily secreted by the corpus luteum in females and reaches the highest plasma levels during pregnancy to prepare and facilitate the processes of parturition and lactation (2). Recent research has revealed that relaxin is also produced by nonreproductive organs and acts via endocrine, paracrine, and autocrine mechanisms on a range of target tissues including the heart (2, 3, 4). Relaxin suppresses cardiac fibrosis as evidenced by the fibrotic phenotype of the heart in relaxin deficient mice (5) and by reversal of established cardiac fibrosis following relaxin treatment (6, 7). In animal models of ischemia and reperfusion, relaxin reduced myocardial injury and preserved ventricular function (8, 9). Although conflicting results exist (10, 11, 12), in patients with congestive heart failure or renal failure associated with increased cardiac events, the circulating and/or cardiac contents of relaxin have been reported to be increased and related to the severity of the syndrome or a higher risk of cardiovascular deaths (13, 14).

Myocardial hypertrophy and apoptosis are documented events that play pivotal roles in the development and progression of heart disease (15, 16, 17). In this regard, potential effects of relaxin on cardiomyocytes (CM) are largely unexplored. The antiapoptotic action of relaxin was firstly reported in rat reproductive tissues during pregnancy (18) and then in swine hearts following ischemia reperfusion injury (9). However, direct evidence supporting the antiapoptotic effects of relaxin in the heart is still lacking. Similarly, the effect of relaxin on cardiac hypertrophy is ambiguous (6, 19). The evaluation of the effects of relaxin directly on CM is therefore a critical step in exploration of the possibility that relaxin is a potential therapeutic agent in the setting of cardiac diseases. Here we have investigated the action of isoform 2 of the human relaxin (H2-relaxin) on cultured neonatal rat CM, focusing on CM hypertrophy and apoptosis. In addition, the relevant signaling pathways activated by relaxin in CM were also explored.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Cell cultures
CM and cardiac fibroblasts (CF) were prepared from ventricles of 1- to 3-d-old Sprague Dawley rats by multiple rounds of digestion using 0.4% collagenase II (Worthington, Lakewood, NJ) and 0.05% trypsin (Invitrogen, Carlsbad, CA) as previously described (20). CM were enriched by 2 h of differential plating (20) and viable cells counted after trypan blue staining. The cells were then seeded at 500 or 1000 cells/mm2 in DMEM with 10% newborn calf serum, 100 U/ml antibiotics-antimycotic agent, 0.1 mM bromodeoxyuridine, 2 mM L-glutamine, 0.1 mM MEM-nonessential amino acids, 0.05 mM MEM-essential amino acids and 1 mM sodium pyruvate [all from Invitrogen, except bromodeoxyuridine from Sigma (St. Louis, MO)]. One day after plating, the media were replaced with fresh serum-free medium (SFM) containing the components specified above, with newborn calf serum omitted but with addition of 0.01% vitamins, 2 µg/ml insulin and 10 µg/ml apo-transferrin (Invitrogen). Cells were maintained in this SFM for 2 d before treatment. This protocol yields more than 98% CM as defined by sarcomeric actin (Sigma) and Hoechst dye 33342 (Sigma) staining.

CF were retained from differential plating during the preparation of CM. They were cultured in DMEM containing 10% fetal bovine serum. To reduce possible contamination by CM or other cells, CF were subcultured twice and, at third passage, were seeded into six-well plates (for CF experiments) or 10-cm plates (for preparation of CF-conditioned medium, FCM) at a density of 500 cells/mm2. The purity of CF was approximately 100% as examined by vimentin (Santa Cruz Biotechnology, Inc., Santa Cruz, CA) and Hoechst dye staining. One day after seeding, CF were rinsed with PBS, and the media changed to SFM. After 2 d of serum starvation, cardiac fibroblast-conditioned medium was prepared with SFM in the presence or absence of H2-relaxin (16.7 nM; BAS Medical, San Mateo, CA) for 48 h and then these media, CF conditioned with H2-relaxin (FCM-relaxin) and CF conditioned without H2-relaxin (FCM), were transferred onto serum-starved CM. The concentration of H2-relaxin was selected on the basis of its physiological level and previous studies (7, 19, 21).

Presence of the relaxin receptor RXFP1 (LGR7) in CM and CF was confirmed by RT-PCR as previously described (7).

Experimental protocols
Before the investigation of the indirect effect of H2-relaxin on CM hypertrophy, experiments to confirm CF inhibition by H2-relaxin were carried out. Serum-starved CF were treated with SFM, TGFß (2 ng/ml; R&D Systems, Minneapolis, MN) and TGFß with H2-relaxin (16.7 nM) for 48 h to determine protein expression of {alpha}-smooth muscle actin ({alpha}-SMA), a marker for myofibroblasts (7, 22).

Experiments studying the direct effect of H2-relaxin on CM hypertrophy were carried out using serum-starved CM treated with phenylephrine (PE) (50 µM; Sigma) in the presence or absence of H2-relaxin (16.7 nM; 48 h). To examine whether H2-relaxin could indirectly inhibit CM hypertrophy, serum-starved CM were treated with SFM, FCM, or FCM-relaxin for 48 h. After these treatments, cell size, protein synthesis, and gene expression of atrial natriuretic peptide (ANP) were determined.

To examine the antiapoptotic effect of relaxin, serum-starved CM were challenged with H2O2 (200 µM; Merck, Whitehouse Station, NJ) with and without H2-relaxin (16.7 nM), using CM cultured in SFM as a control. CM were harvested at different times, based on our preliminary experiments, for determination of DNA fragmentation, Akt and ERK activation and Bcl2/Bax ratio at both mRNA and protein levels.

To explore whether the pertussis toxin (PTX)-sensitive G-proteins, Gi/o, are involved in H2-relaxin-induced Akt activation in CM, serum-starved CM were pretreated with PTX (100 ng/ml; Sigma) overnight, and then subjected to H2O2 (200 µM) challenge in the presence or absence of H2-relaxin (16.7 nM). After 15 min, cells were harvested for determination of phosphorylated Akt (pAkt) expression by Western blotting.

All culture medium contained 0.2% lactalbumin hydrolysate (Becton Dickinson, Franklin Lakes, NJ) as a protein carrier to ensure the stability of H2-relaxin for at least 72 h.

Measurement of cell size
Morphological changes in CM were observed by phase-contrast light microscopy (Olympus IX70). Live cell images, at x200 magnification, were acquired by a digital camera attached to the microscope. Three images were taken for each well and three or four wells were set up per group. For cell size measurement, one of three images was randomly selected and all cells in that image (at least 30/image) were measured using software Optimas (version 6.5; Media Cybernetics, Silver Spring, MD). Results from three to four wells per group (about 100 cells) were expressed as average cell size per group.

Measurement of protein synthesis
Protein synthesis was determined by incorporation of [3H]-leucine. [3H]-leucine (1 µCi/ml; Amersham, Arlington Heights, IL) was coincubated with the cells for the last 24 h of treatment. Cells were washed with PBS and treated with 10% trichloroacetic acid at 4 C for 30 min to precipitate proteins, which were then dissolved in NaOH (0.25 M). Aliquots of quadruplicates per group were counted with a scintillation counter (TRI-CARB Liquid Scintillation Analyzer 1900CA; Packard Instruments, Meriden, CT).

Real-time PCR
Total RNA was extracted from cells using TRIzol reagent (Invitrogen). After deoxyribonuclease I (Promega, Madison, WI) treatment, 1 µg of total RNA was converted to cDNA using Superscript III (Invitrogen). Real-time PCR was performed on 5 ng cDNA using SYBR Green PCR Master Mix (Applied Biosystems, Foster City, CA) on an ABI Prism 7500 Sequence Detection System to determine mRNA levels of ANP, Bcl2, and Bax. Results were normalized to glyceraldehyde-3-phosphate dehydrogenase (GAPDH). Primers were designed using Vector NTI (Invitrogen) and purchased from Sigma.

Western blot analysis
Cells were lysed in buffer [50 mM Tris (pH 7.5), 100 mM NaCl, 50 mM NaF, 1 mM EDTA, 0.5% deoxycholic acid, 0.1% sodium dodecyl sulfate, 1% Triton X-100 and 1x protease inhibitor cocktail from Roche (Indianapolis, IN)] with fresh addition of 0.2 mM Na3VO4, 0.4 mM phenylmethylsulfonyl fluoride and 1x phosphatase inhibitor cocktail I and II (Sigma). Protein was determined using the Bradford assay (Bio-Rad, Hercules, CA). Equal amounts of protein from each treatment were separated on 10% SDS-PAGE and transferred to polyvinylidene difluoride membranes which were blocked with 5% skim milk and incubated in a primary antibody solution overnight at 4 C. All of the primary antibodies were diluted in Tris-buffered saline/Tween 20 solution containing 5% skim milk and 0.1% sodium azide [1:1000 for p-ERK1/2, 1:1000 for p-Akt; Cell Signaling Technology (Beverly, MA); 1:1000 for {alpha}-SMA (Dako, Carpinteria, CA); 1:1000 for GAPDH, 1:200 for Bcl2, 1:200 for Bax, Santa Cruz and 1:8000 for {alpha}-tubulin; Sigma]. After the primary antibody incubation, membranes were washed four times in Tris-buffered saline/Tween 20 and incubated in a corresponding secondary antibody conjugated with horseradish peroxidase (Santa Cruz) for 3 h at room temperature. Membranes were washed four times before exposure using enhanced chemiluminescence reagent (Pierce, Rockford, IL).

To quantify band intensity, the images of Western blot films were scanned. Band intensity was quantitated using the software Quantity One (version 4.5.2; Bio-Rad) and results normalized by respective level of either GAPDH or {alpha}-tubulin.

Quantitative analysis of apoptotic nuclei
Hoechst 33342, a fluorescent DNA binding dye that allows clear distinction between apoptotic and normal cells on the basis of nuclear morphology, was used to quantify apoptotic myocytes (23). Dye was added to the culture medium to a final concentration of 10 µM for 1 h before termination of the experiments. The cells were incubated at 37 C for 1 h and evaluated under fluorescence microscopy at x400 magnification. Cells were scored as apoptotic if they exhibited unequivocal nuclear chromatin condensation and/or fragmentation (23). For each experimental condition, three independent CM populations were prepared and triplicates set for each condition every time. To quantify apoptosis, at least 300 nuclei were analyzed from 10 randomly selected fields per well and the results from three wells per group (about 900 nuclei) were quantified for each condition in one independent experiment. The percentage of apoptotic cells was calculated. These independent quantification procedures were performed blind.

Analysis of DNA fragmentation
DNA was extracted from cells using ApopLadder Ex kit (Takara, Shiga, Japan) following the manufacturer’s instructions. The amount of DNA isolated was determined by spectrophotometry at 260 nm. DNA samples (10 µg) were subjected to electrophoresis in 3% NuSieve 3:1 agarose gels and imaged by ethidium bromide staining.

Statistical analysis
Either triplicates or quadruplicates were used for each experimental group in each experiment. All data, from at least three independent experiments, are presented as mean ± SEM, with P < 0.05 being considered as statistically significant. An unpaired t test was used to compare results between two groups, and one-way ANOVA, with Newman-Keuls’s test if P < 0.05, used for analysis of results from ≥ two groups. Software used was GraphPad Prism (version 4.01; GraphPad Software Inc., San Diego, CA).


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
Relaxin did not alter PE-induced cardiomyocyte hypertrophy
Treatment with PE (50 µM) alone caused significant hypertrophy of CM, as measured by a 25% increase in cell size (P < 0.01), 2.0-fold increase in [3H]-leucine uptake (P < 0.001) and a 6.8-fold increase in ANP gene expression (P < 0.001), compared with untreated control cells (Fig. 1Go). Cotreatment with PE and H2-relaxin (16.7 nM) failed to repress hypertrophy. Although H2-relaxin treatment alone caused a 4-fold increase in ANP expression (SFM, 1.1 ± 0.2; relaxin, 3.9 ± 0.7; P < 0.01) there was no significant change in cell size or protein synthesis (Fig. 1Go). Pretreatment with H2-relaxin for 24 h produced no effect on CM hypertrophy induced by PE (data not shown). Collectively, these results indicate that H2-relaxin has no significant effect on PE-induced hypertrophy.


Figure 1
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  		FIG. 1. H2-relaxin (Rln) did not alter PE-induced cardiomyocyte hypertrophy. Mean cell size (top) was determined from 100 cells/group/cell preparation. Mean protein synthesis (middle) was estimated by [3H]-leucine (1 µCi/ml) incorporation. Mean mRNA level of ANP (bottom) was measured by real-time PCR. ANP expression levels were normalized by GAPDH. All data above were obtained from either triplicates or quadruplicates per experimental group for each CM preparation and three or four independent preparations of CM were conducted in total.

 
Relaxin suppressed cardiomyocyte hypertrophy induced by cardiac fibroblast-conditioned medium
Compared with untreated cells, treatment with TGFß alone increased {alpha}-SMA expression in CF by 1.6-fold (P < 0.01). H2-relaxin abolished this increase in {alpha}-SMA expression (P < 0.001). Moreover, H2-relaxin further reduced the {alpha}-SMA expression to only 50% of the control level (P < 0.05; Fig. 2AGo). We then tested whether H2-relaxin could influence, via its inhibitory action on CF, CM hypertrophy induced by FCM. FCM caused a significant CM hypertrophy, as measured by increased CM size (SFM, 100 ± 6%; FCM, 125 ± 5%; P < 0.01), enhanced [3H]-leucine incorporation (SFM, 100 ± 16%; FCM, 320 ± 29%; P < 0.001), and elevated ANP gene expression (SFM, 1.0 ± 0.1; FCM, 1.3 ± 0.04; P < 0.05; Fig. 2BGo). All of the three measurements were significantly reduced in CM treated with FCM-relaxin, indicating inhibition of hypertrophy by H2-relaxin. Cell size and ANP gene expression were reduced to equivalent to or below the basal level, while [3H]-leucine incorporation was suppressed by about 24% (FCM, 320 ± 29%; FCM-relaxin, 243 ± 19%; P < 0.05; Fig. 2BGo). This set of data suggests that H2-relaxin inhibits CM hypertrophy via its actions on CF.


Figure 2
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  		FIG. 2. H2-relaxin (Rln) suppressed cardiomyocyte hypertrophy induced by FCM. A, Relaxin inhibited CF activation. Western blot analysis of {alpha}-SMA was used as a marker of activated CF (myofibroblasts). CF were either untreated (SFM) or treated with TGFß (2 ng/ml) alone or TGFß and H2-relaxin (16.7 nM) (TGFß + Rln) for 48 h. Images shown are representatives of Western blots (top) and corresponding quantification by densitometry (bottom) of three independent experiments using CF. The {alpha}-SMA expression was normalized against {alpha}-tublin for protein loading. B, Measurement of degree of CM hypertrophy. Mean cell size (top) was determined from 100 cells/group/cell preparation. Protein synthesis (middle) was estimated by [3H]-leucine (1 µCi/ml) incorporation. mRNA level of ANP (bottom) was measured by real-time PCR. ANP expression levels were normalized by GAPDH. All data above were obtained from either triplicates or quadruplicates per experimental group for each CM preparation and three or four independent preparations of CM were conducted in total.

 
Relaxin protects cardiomyocytes against apoptosis under oxidative stress
A well-characterized apoptotic inducer, H2O2 (200 µM) (24), was used to induce apoptosis in CM with and without H2-relaxin to determine whether the peptide affects CM apoptosis under oxidative stress. In agreement with previous reports (24), CM apoptosis was morphologically evident after 24 h treatment with H2O2. Hoechst dye 33342 binds to DNA allowing for visualization of chromatin condensation as small, dense and brighter nuclei in H2O2-treated cells (Fig. 3AGo). Image quantification (Fig. 3BGo) revealed that the percentage of apoptotic cells identified in the H2O2-treated group was approximately 7-fold higher than that in the control group (SFM, 5.3 ± 2.4%; H2O2, 37.4 ± 3.7%; P < 0.001). This increased apoptotic rate induced by H2O2 was halved by H2-relaxin (16.7 nM) treatment (H2O2, 37.4 ± 3.7%; H2O2 + relaxin: 17.6 ± 1.5%; P < 0.001), demonstrating that H2-relaxin antagonizes oxidative stress-induced apoptosis in CM. This was further supported by DNA electrophoresis studies, where an intense apoptotic laddering pattern was evident after H2O2 treatment, that was reduced in cells cotreated with H2O2 and H2-relaxin (Fig. 3CGo).


Figure 3
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  		FIG. 3. H2-relaxin (Rln) reduced DNA fragmentation in oxidatively stressed cardiomyocyte. A, Representative images of Hoechst dye 33342-stained CM at 24 h either untreated (SFM) or treated with H2O2 (200 µM) alone (H2O2), H2-relaxin (16.7 nM) alone (Rln), or H2O2 and H2-relaxin (H + R). Cells were considered as apoptotic if they exhibited unequivocal nuclear chromatin condensation and/or fragmentation (indicated by red circles and arrows). B, Quantitative results of antiapoptotic action of relaxin in oxidatively stressed CM. Each column represents mean of percentages of apoptotic cell numbers calculated from 900 nuclei/group/cell preparation and three independent cell preparations in total. C, Relaxin attenuated DNA fragmentation shown by DNA laddering. Fragmented genomic DNA was isolated from CM. DNA was separated by electrophoresis in 3% NuSieve 3:1 Agarose gel and imaged by ethidium bromide staining. M, Denotes 1-kb Plus DNA Ladder used for size indication. The image is representative of four independent experiments.

 
We also examined the Bcl2/Bax ratio at both the mRNA and protein levels to better understand the antiapoptotic action of H2-relaxin on CM. H2-Relaxin treatment alone did not influence the ratio of Bcl2/Bax at mRNA level, whereas H2O2 induced a significant and sustained fall (~70%) in the ratio from control (P < 0.05; Fig. 4AGo). Importantly, cotreatment with H2O2 and H2-relaxin preserved the ratio, and, in fact, increased it by nearly 7-fold compared with that of H2O2 treatment group (P < 0.001). Although the cotreated cells failed to sustain the elevation following prolonged exposure of H2O2, the Bcl2/Bax ratio in 2 h after H2O2 challenge was maintained at the level comparable with that of control cells and was still significantly higher than that of H2O2-treated cells. In agreement with the gene expression data, protein expression studies showed that H2-relaxin increased the Bcl2/Bax ratio of CM 6 h after H2O2 challenge (Fig. 4BGo), with a 5-fold increase (P < 0.01), suggesting that relaxin exerts its antiapoptotic effect by maintaining the Bcl2/Bax ratio in oxidatively stressed CM.


Figure 4
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  		FIG. 4. H2-relaxin (Rln) augmented the Bcl2/Bax ratio in cardiomyocyte under oxidative stress. A, H2-relaxin (16.7 nM) (H+R) increased Bcl2/Bax mRNA ratio in oxidatively stressed CM (H2O2) determined by real-time PCR. Each column represents triplicates/group/cell preparation and three independent preparations of CM in total. mRNA expression levels of Bcl2 and Bax were normalized against GAPDH. B, H2-relaxin (16.7 nM) (H + R) increased Bcl2/Bax protein ratio in CM subjected to oxidative stress (H2O2). Western blot images (top) and quantitative analysis using densitometry (bottom) are representative of duplicates/group/cell preparation and four independent preparation of CM in total. Protein expression levels were normalized against GAPDH.

 
Relevant antiapoptotic signaling pathways activated by relaxin in oxidatively stressed cardiomyocytes
To further confirm that relaxin possesses antiapoptotic actions in CM under oxidative stress and to explore its signaling pathways, phosphorylation of Akt and ERK were examined by Western blotting. Comparable but weak phosphorylated ERK (pERK) signals were detected in both untreated control and H2-relaxin treated cells (Fig. 5AGo), indicating that H2-relaxin alone had no significant effect on ERK phosphorylation in control CM. After stimulation with H2O2, however, there was a significant increase in pERK, to some 14-fold of that in the basal group (P < 0.05). Cotreatment with H2-relaxin induced a further increase in phosphorylated ERK after H2O2 challenge (H2O2 + relaxin: 2.9 ± 0.4; H2O2: 1.4 ± 0.2; P < 0.01). Similar observations were made for Akt activation. pAkt was detected to a similar extent in untreated control, H2-relaxin treated or H2O2-treated cells (Fig. 5BGo). Addition of H2-relaxin to H2O2-treated CM increased pAkt expression by up to 1.8-fold, compared with that in untreated cells (P < 0.01) or cells treated with H2O2 alone (P < 0.001).


Figure 5
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  		FIG. 5. Relevant antiapoptotic signaling pathways activated by relaxin in cardiomyocytes. A, H2-relaxin treatment (16.7 nM) (H + R) increased levels of pERK in H2O2 (200 µM) treated CM (H2O2), determined by Western blot analysis. Protein expression levels of pERK were normalized against {alpha}-tublin. B, H2-relaxin treatment (16.7 nM) (H + R) increased levels of pAkt in H2O2 (200 µM) (H2O2) treated CM and pretreatment of PTX (100 ng/ml) (H + R + P) blunted this up-regulation of pAkt by H2-relaxin. Western images (top) and their corresponding quantitative analysis using densitometry (bottom) are representative of duplicates/group/cell preparation and three independent preparations of CM in total for both determinations.

 
To determine whether Gi/o proteins are involved in phosphorylation of Akt in CM treated with H2-relaxin (16.7 nM), Western blot analysis was employed to compare phosphorylation of Akt induced by H2-relaxin with or without pretreatment by PTX, a specific inhibitor of Gi/o. PTX (100 ng/ml) was applied overnight followed by H2O2 stimulation with and without H2-relaxin. Treatment with H2-relaxin, PTX, or H2O2 alone had no significant effect on Akt phosphorylation compared with untreated control cells (Fig. 5BGo). However, under oxidative stress induced by H2O2, H2-relaxin significantly increased pAkt level up to 1.8-fold (P < 0.01), and this was significantly repressed by pretreatment with PTX (H2O2 + relaxin, 1.6 ± 0.1; H2O2 + relaxin + PTX, 1.4 ± 0.02; P < 0.05). Thus, Gi/o coupling is involved in relaxin-induced Akt activation in CM.


   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
The present study demonstrates, for the first time, an indirect antihypertrophic and direct antiapoptotic effects of relaxin in cultured neonatal rat CM. Our data suggest that relaxin inhibits the ability of FCM to induce hypertrophy in CM but does not have a direct antihypertrophic effect. We also demonstrated that relaxin directly attenuates apoptosis in CM induced by oxidative stress. Furthermore, we showed that the two important antiapoptotic signaling pathways involving Akt and ERK were activated by relaxin in oxidatively stressed CM.

Relaxin has been shown to induce cardiac expression of ANP (25), which mediates antihypertrophic signals (26). We thus hypothesized that relaxin could directly influence CM hypertrophy via induction of ANP. In contrast to the findings by Dschietzig et al. (19) of the inhibitory effect of relaxin on gene expression of ANP and ß-myosin heavy chain in PE-treated CM, we showed that CM hypertrophy induced by PE, as measured by cell size, protein synthesis, and ANP gene expression, was not influenced by relaxin. We observed that treatment with relaxin increased ANP expression by almost 4-fold compared with untreated cells. However, pretreatment of CM with relaxin failed to suppress the PE-induced hypertrophy confirming that relaxin did not direct inhibit CM hypertrophy in this model.

The lack of agreement between this and an earlier study (19) implying a direct inhibition of CM hypertrophy by relaxin could result from difference in the cell types used. We used neonatal Sprague Dawley rat CM, whereas the previous study used CM from adult spontaneously hypertensive rats. Based on the lack of cardiac hypertrophy in aged relaxin-deficient mice (5), and the current data showing indirect inhibition of CM hypertrophy by relaxin, we speculate that the reported inhibitory action of relaxin on CM hypertrophy is related largely to its antifibrotic property.

It is well documented that pathological CM hypertrophy and progressive matrix remodeling, including CF activation and fibrosis, coexist in the diseased heart in vivo. In the diseased heart, CF are activated and differentiate into myofibroblasts (22). These activated myofibroblasts secret a series of cytokines, growth factors, and inflammatory mediators, such as IL-6, basic fibroblast growth factor, TGFß, endothelin-1, and TNF-{alpha}, that induce and promote CM hypertrophy in a paracrine fashion (22, 27, 28, 29, 30). Because relaxin possesses antifibrotic properties (6, 7), we speculated that this inhibition of CF might in turn suppress CM hypertrophy. Our results demonstrated that relaxin did not only reverse the elevated {alpha}-SMA expression in CF following TGFß stimulation, but also halved the level in myofibroblasts relative to the untreated control CF. Cultured myofibroblasts proliferate faster than CF (22, 30). Thus, H2-relaxin is also capable of inhibiting proliferation in activated CF, confirming our previous findings (7). This inhibition would result in reduced levels of paracrine factors secreted by activated CF and, eventually cause CM hypertrophy in response to these factors. Significant hypertrophy was revealed in CM treated with FCM compared with untreated CM. Interestingly, all three parameters for hypertrophy, cell size, 3[H]-leucine incorporation and ANP gene expression, were reduced in CM treated with FCM-relaxin compared with those treated with FCM. This provided support for the notion that relaxin would indirectly inhibit CM hypertrophy.

The antiapoptotic action of relaxin was first described in reproductive tissues (2, 18, 31). There are reports of a protective action of relaxin against ischemia-reperfusion injury (8, 9). It is well established that apoptotic cell death resulting from oxidative stress is a major pathology under ischemia-reperfusion injury (32). Perna et al. (9) showed in pigs subjected to ischemia-reperfusion that relaxin treatment significantly reduced apoptotic cell number associated with a marked attenuation of caspase-3 activity, indicating antiapoptotic property of relaxin. However, the percentage of apoptotic cells (over 70%) was significantly higher than that usually seen in hearts following ischemia-reperfusion in vivo (5–15%) (33, 34, 35, 36). In addition, relaxin is known to inhibit the activation and endothelium-adhesion of neutrophils (8, 37, 38), the migration of eosinophils (39), histamine release from mast cells (40), and platelet aggregation (41), effects potentially contributing to cardiac protection by relaxin. Hence, it was necessary to investigate the antiapoptotic effects of relaxin and the mechanisms involved in a pure CM model. Using oxidatively stressed CM, we explored whether H2-relaxin directly protects CM from apoptotic cell death, and determined the relevant key molecules and pathways involved.

H2O2 is a well-defined inducer of apoptosis with its level increased in the setting of ischemia-reperfusion injury (42). H2O2 stimulation induces reactive oxygen species but inhibits nitric oxide synthase and both effects could trigger apoptosis in CM (43, 44). In our study, H2O2-treated cells exhibited the typical changes of apoptosis such as distinct DNA laddering and smaller and/or brighter nuclei staining with Hoechst dye. Treatment with relaxin halved the percentage of apoptotic cells and increased Bcl-2 but decreased Bax expression, resulting in an elevated Bcl2/Bax ratio, indicative of an antiapoptotic effect. These results demonstrate that relaxin directly protects CM from apoptotic death under oxidative stress, which very likely underpins one of the major mechanisms involved in relaxin’s cardio-protective effects in ischemia-reperfusion injury (8, 9).

We further explored the signaling pathways involved in this antiapoptotic action of relaxin in CM, which constitutively express RXFP1 (7). Relaxin signaling is complex, target cell-specific and only partially understood (2, 4, 45). When relaxin binds to its cognate G protein-coupled receptor, RXFP1, a Gs protein signaling pathway is stimulated leading to activation of adenylate cyclase and intracellular accumulation of cAMP (46). Binding of relaxin to RXFP1 may also activate pAkt-NOS, via Gß{gamma}-subunits from Gi/o (45); or stimulate a tyrosine kinase-ERK pathway (45). Because the signaling pathways used by relaxin in CM are poorly explored, we examined in CM the levels of activation of Akt and ERK, two well-documented antiapoptotic signal molecules (47, 48) and found that both were significantly up-regulated by relaxin under conditions of oxidative stress. Akt activated in response to relaxin phosphorylates and inactivates Bad, leading to disintegration of the apoptosome and then enhanced cell survival (49). In addition, increased NO produced by relaxin via the pAkt-NOS pathway could also potentially suppress apoptosis (50, 51, 52). ERK may be phosphorylated as a result of a relaxin-induced increases in cAMP (53) and/or relaxin stimulated activation of tyrosine kinase, leading to increased CM survival (45, 48). Thus, it is likely that relaxin exerts its antiapoptotic action in CM via both Akt and ERK pathways.

RXFP1-Gs coupling was first demonstrated in 1999 (54). Recently, it has been reported that relaxin’s signaling is partially mediated by Gi coupling (21). Therefore, Akt could be activated in CM by relaxin as a result of augmentation of the PI3K signaling via the Gß{gamma} subunits of Gi/o (21). One of our earlier studies had displayed a reduced inotropic response to relaxin by treatment of PTX, an inhibitor of Gi/o, in a rat model of myocardial infarction (55). The present study further demonstrated, for the first time at cellular level, that relaxin-induced activation of Akt in CM is partially suppressed by pretreatment by PTX, thus suggesting that Gi/o coupling of RXFP1 plays a part in relaxin’s signaling in CM.

In summary, the data from the present study reveal novel information on the antihypertrophic and antiapoptotic actions of relaxin in CM and the relevant signaling pathways. These findings extend our current knowledge of the action of relaxin on the heart and provide direct evidence at the cellular and molecular levels for the notion that relaxin is a candidate for the treatment of heart disease, targeting at myocardial hypertrophy, apoptosis, and fibrosis.


   Footnotes
 
This work was supported by a grant from the Australian Research Council (LP0560620) and the Australian National Health and Medical Research Council (Grant 436713 to R.J.S. and R.A.D.B.) and Fellowships (to E.A.W., G.W.T., and X.J.D).

Author Disclosure: X.L.M., S.L.T., C.Y.L., L.F., Y.D.S., X.M.G., E.A.W., X.J.D., G.W.T., R.A.D.B, R.J.S. have nothing to declare.

First Published Online January 4, 2007

Abbreviations: ANP, Atrial natriuretic peptide; CF, cardiac fibroblasts; CM, cardiomyoctyes; FCM, cardiac fibroblast-conditioned medium; GAPDH, glyceraldehyde-3-phosphate dehydrogenase; H2-relaxin, isoform 2 of the human relaxin; pAkt, phosphorylated Akt; PE, phenylephrine; pERK, phosphorylated ERK; PTX, pertussis toxin; SFM, serum-free medium; {alpha}-SMA, {alpha}-smooth muscle actin.

Received September 27, 2006.

Accepted for publication December 27, 2006.


   References
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 

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