Abnormal stiffness and altered cardiac function arising from abnormal collagen deposition occur in hypertrophy and heart failure. ANG II has been shown to play a role in this process. To evaluate the mechanism, we developed an in vitro model by subjecting fibroblasts to ANG II treatment in the presence or absence of myocytes in coculture (25). Employing this model, we demonstrated that ANG II-induced collagen gene transcription in cardiac fibroblasts was potentiated by myocyte-derived factors. In attempting to identify mechanisms of collagen upregulation and to define the role of myocytes, we found that interleukin (IL)-6, tumor necrosis factor (TNF)-α, and the transforming growth factor (TGF)-β superfamily were also involved in collagen upregulation. Collagen transcripts were increased after fibroblasts were treated with IL-6 (20–50 ng/ml) and TNF-α (0.1–0.5 ng/ml). In this study, we show that cardiomyocytes induce secretion of active TGF-β in the presence of ANG II and that a paracrine action of TGF-β subsequently induces different cytokines (IL-6) in fibroblasts, thereby promoting collagen synthesis. The cross-talk between myocytes and fibroblasts and involvement of these cytokines in the upregulation of collagen transcript levels are novel findings that may explain their possible roles in the upregulation of collagen.
- fibroblast-myocyte cross-talk
- angiotensin II
cells respond in various ways to mechanical stimuli. Such stimuli often trigger signals that lead to increased gene expression, protein synthesis, or mitogenesis (29, 35). In other situations, cells respond by differentiating (28), rearranging their cytoskeleton or focal contacts (9), or altering the composition of their extracellular matrix. Excessive deposition of collagen may be responsible for the abnormal tissue stiffness and altered cardiac function that develop during hypertrophy and heart failure (7). Our laboratory has shown that as hypertrophy develops, collagen phenotypes become altered in humans and rats. Although collagen is universally regarded as a major player in hypertensive hypertrophy, leading to stiffness of the heart and eventual heart failure, the exact mechanism of collagen gene upregulation in the heart in the diseased state is not known. Evidence suggests that cardiac hypertrophy is induced by mechanical load (22), humoral factors such as ANG II (31), endothelin (37), α1- and β1-adrenergic agonists (36), transforming growth factor (TGF)-β (34), and insulin-like growth factors (13). Also, ANG II has been shown to affect gene expression in both fibroblasts and myocytes. ANG II acts as a growth-promoting factor on myocytes in an autocrine or paracrine manner and activates a variety of signaling molecules to induce various genes that promote cardiac hypertrophy. Evidence further suggests that ANG II has a defining role in stimulating collagen gene expression and protein turnover in cultured cardiac fibroblasts (30, 41). ANG II induces collagen mRNA expression in vivo via angiotensin type 1 (AT1) receptors (15, 18), whereas the deposition of type I collagen in vitro occurs only when the medium is supplemented with ascorbate, independent of the age of the animal (40).
The mechanism by which ANG II stimulates abnormal collagen production remains unclear. Different signaling pathways of AT1 receptors may promote collagen synthesis (12). Irrespective of signaling mechanisms, the end result is activation of transcription factors that bind to various “cis-acting” elements in the regulatory sequences of α1(1), α2(1), and α1(III) genes (14). We have shown previously that cellular cross-talk between myocyte and fibroblast is necessary for upregulation of collagen in the heart (25), providing new insight into the mechanism of collagen gene activation in the diseased heart. The present study is a continuation of our previous observation that myocytes potentiate ANG II-mediated collagen production by fibroblasts. The present study was undertaken to determine the role of myocytes in this process. This study showed that ANG II exerts two independent effects: 1) it induces active TGF-β production from myocytes, and 2) it modulates the cytokine response in fibroblasts. Our data demonstrate how autocrine/paracrine mechanisms regulate the cross-talk between various cell types (fibroblasts and myocytes) and may provide novel insights into molecular mechanisms of collagen synthesis during myocardial hypertrophy.
MATERIALS AND METHODS
The Wistar-Kyoto (WKY) rats used in this study were obtained from Taconic Farms (Germantown, NY). This investigation conformed to the National Institutes of Health Guide for the Care and Use of Laboratory Animals (NIH Pub. No. 85-23, Revised 1996) and guidelines of the Institutional Animal Care and Use Committee of the Cleveland Clinic Foundation.
Preparation of Cardiac Cells and the Beating, Nonworking Rat Heart
Rat cardiac fibroblasts from normal 28-wk-old WKY rats were isolated as described previously (33). Flasks containing pure fibroblasts were then kept at 37°C until cells were confluent; these were subsequently passaged. The cells were used at 70–75% confluency and kept in serum-free medium for 24 h before each experiment.
Neonatal myocytes from WKY rat pups were isolated and cultured on laminin-coated cover glasses placed in standard six-well plates following the procedure described previously (33). Adult myocytes were prepared on a laminin-coated coverglass or coculture inserts, as described previously (32). In brief, rats were heparinized intraperitoneally (100 U/100 g body wt), and the hearts were taken out aseptically and washed with ice-cold Joklik's medium containing Joklik's minimal essential medium, 25 mmol/l glutamic acid, 30 mmol/l taurine, and 1 mmol/l adenosine. A cannula was introduced into the lumen of the aorta, and the heart was retrogradely perfused at 37°C on a modified Langendorff apparatus for 10 min. The perfusion was continued in the same medium containing collagenase type II (100 U/ml, Worthington) for 30 min at 37°C. After perfusion, the atria and vessels were removed, and the ventricles were cut into small pieces. These were incubated for an additional 5 min in collagenase media with occasional shaking. The tissue was disaggregated by trituration with a sterile transfer pipette, and the released cells were removed by filtration through a piece of sterile nylon net in 5% FBS-containing media. α-Actinin antibody staining showed that the isolated myocytes were ∼90% pure.
The beating, nonworking rat heart model was prepared as described by Pathak et al. (25). For the beating, nonworking heart preparation, normal 28-wk-old WKY rats were injected with 500 units of heparin and anesthetized with 20 mg/kg of pentobarbital intraperitoneally. Each heart was taken out and washed with modified Krebs-Henseleit buffer (pH 7.35). The buffer was bubbled with 95% O2-5% CO2 for 30 min. The heart was then cannulated through the aorta and was then perfused with buffer at a control pressure of 75 mmHg for 3 h. [Sar1]ANG II (10−8M, Sigma) was perfused by a small peristaltic pump along with the buffer into the heart. Hearts perfused with only Krebs-Henseleit buffer for 3 h were used as controls.
Coculture Experiments with Rat Myocytes and Fibroblasts with ANG II
Adult fibroblast cells (2 × 106 cells/well of a standard 6-well plate) from 28-wk-old WKY rats were incubated for 24 h at 37°C in the absence (control) or presence (treated) of 10−8 mol/l [Sar1]ANG II in standard six-well plates with [fibroblasts + myocytes + ANG II (F+M+A)] or without myocytes [fibroblasts + ANG II (F+A)] on coverslips or coculture inserts, as described previously (25). The coculture of fibroblasts and myocytes was incubated for 4 h alone and then with 10−8 mol/l [Sar1]ANG II for 20 h at 37°C. The same concentration of ANG II was replenished every 6 h during the experiment. Both of these cell types (myocytes and fibroblasts) were under the influence of ANG II throughout the incubation period.
Extraction of RNA and Northern Hybridization
Total RNA from rat hearts and fibroblasts was isolated using the RNeasy minikit (QIAGEN) following the manufacturer's protocol. Then, 10 μg RNA from each sample was transblotted on a GeneScreen membrane and hybridized with three different radiolabeled probes: 1) collagen type I rat cDNA probe (kind gift from Dr. David Rowe, Univeristy of Connecticut Health Center); 2) collagen type III human skin fibroblast cDNA probe (American Type Culture Collection); and 3) 18S rRNA oligoprobe (Oncogene), respectively. The cDNA probes were labeled using a random primed DNA labeling kit (Roche Diagnostics). 18S oligoprobes were labeled by [γ-32P]dATP using T4 kinase (Roche) and were used as internal controls for all the blots used in this study. The cDNA-mRNA hybrids were visualized by autoradiography on Kodak Biomax-MR films exposed for 24 h with intensifying screens. For quantification, the intensity of each band was determined by videodensitometry and normalized with the internal control.
Quantification of Total Collagen
Total collagen was quantified from the 28-wk-old WKY rat fibroblast supernatant by measuring hydroxyproline with a modified Stegemann procedure (25). DMEM-F12 medium was used as a reference. The amount of hydroxyproline in unknown samples was calculated with a standard curve. Collagen content was estimated by multiplying the hydoxyproline content by a factor of 8.2 and is expressed as micrograms of collagen per milliliter of cell supernatant.
RNase Protection Assay and RT-PCR
RNase protection assays were performed according to the manufacturer's protocol (RiboQuant, Pharmingen; cytokine template sets mCK-3b) with 15 μg of total RNA from fibroblasts. After RNase digestion, protected probes were resolved on denaturing polyacrylamide gels and quantified by a PhosphoImager (ImageQuant software, Molecular Dynamics). The value of each hybridized probe was normalized to that of GAPDH, included in each template set as an internal control.
RT-PCRs were performed according to the protocols of Ambion. Rat oligonucleotide primers for TGF-β, tumor necrosis factor (TNF)-α, and interleukin (IL)-6 were provided by the company. PCR was performed for 30 cycles (94°C for 30 s, 57°C for 30 s, and 72°C for 30 s), and the products were visualized on 2% agarose gels by ethidium bromide staining.
Treatment of Fibroblasts
Fibroblasts from 28-wk-old WKY rats were treated with TGF-β2 (2–8 ng/ml), TNF-α (0.1–0.5 ng/ml), and IL-6 (20–50 ng/ml) (Life Technologies) for 12–16 h in six-well plates either alone or with fibroblasts cocultured with myocytes. Anti-IL-6-neutralizing antibody (1 μg/ml, R&D Systems; Minneapolis, MN) was added to the F+M+A model for 4–6 h after the coculture of fibroblasts and myocytes were incubated with 10−8 mol/l [Sar1]ANG II for 20 h.
In a separate experiment, fibroblasts were incubated with anti-TGF-β receptor RII antibody (5 μg/ml, Santa Cruz Biotechnology) in culture for 6 h before myocytes and ANG II were added to it. In another set of experiments, neutralizing anti-TGF-β antibody (10 μg/ml, R&D Systems) was added to growing myocytes in culture before the supernatant was supplemented to the fibroblasts in the F+M+A model.
With the use of lipofectamine (Life Technologies), the 28-wk-old WKY rat fibroblasts were transiently transfected with dominant negative constructs of Smad 2 and 3 [Smad 2DN and Smad 3DN (8, 26)] using the manufacturer's protocol. An appropriate empty vector was also transfected. The transfection efficiency (Smad 2DN expression) was estimated separately by immunoblotting cell lysates with rabbit anti-phospho-Smad 2 from Upstate Biotechnology (Lake Placid, NY) and anti-Smad2 and anti-Smad 3 from Zymed Laboratories (San Francisco, CA) after the fibroblasts were treated with TGF-β. Expression of Smad 2DN and Smad 3DN was sometimes checked by Northern blots using a collagen type I probe to these transfected cells after TGF-β treatment.
Quantitation of Active TGF-β in Conditioned Media
Serum-free conditioned media from the cardiac cells (fibroblast, myocytes, and combination) grown in the presence or absence of ANG II were collected after 24 h. Extensive washing was done to eliminate carryover of serum. The media were briefly centrifuged, and protease inhibitor cocktail set III (Calbiochem; La Jolla, CA) was added. Media were extensively dialyzed against water, lyophilized, and resuspended in 1× PBS. Next, 10–20 μg/ml of resuspended media were added to CCL-64 mink lung epithelial cells, maintained in high-glucose formulation of 1× DMEM (Life Technologies) with 10% FBS (11). Then, 1–2.5 μg/ml of TGF-β2 were added to these cells as a positive control. The cells were treated with the media for 17 h and pulsed with 0.5 μCi (12.6 mCi/mmol) [3H]thymidine (Amersham) for 3 h. The cells were lysed with 0.3 N NaOH-0.1% SDS, and the lysate was transferred to Whatman microfiber filters. The filters were washed, and the radioactivity was counted using a β-scintillation counter. The values are expressed as the percent decrease over control.
Active TGF-β1 and TGF-β2 from the cell supernatant were measured by the Emax ImmunoAssay system (Promega; Madison, WI), following the manufacturer's protocol. The wells were precoated overnight with TGF-β coat mAb and then blocked with 1× blocking buffer (supplied) at 37°C for 35 min. Standard curves for TGF-β1 and TGF-β2 were prepared using six 1:2 serial dilutions with the supplied acidified standards. One hundred microliters of each sample were incubated at room temperature for 2 h. After being washed, the sandwich was incubated with anti-TGF-β1 and anti-TGF-β2 pAb for 2 h, followed by incubation with TGF-β horseradish peroxide (HRP) conjugate for 2 h at room temperature. The color was generated by TMB One solution for 15 min in the dark, and the reaction was stopped by 1 N hydrochloric acid. The plate was immediately read at 450-nm absorbance using an ELISA plate reader (Vector II, Perkin-Elmer).
Estimation of Secreted Cytokines by ELISA and Dot Blots
Estimation of secreted cytokines, e.g., TNF-α and IL-6, from 28-wk-old WKY rat fibroblast-conditioned media was performed using kits from Pierce Endogen (Rockford, IL). Standard curves were obtained using eight data points with serial dilutions using supplied standards. Samples as well as standards were applied on the antibody-precoated ELISA plates and were incubated at room temperature for 2 h. Biotinylated antibody was then added and incubated for 1 h at room temperature. After 30 min of incubation with streptavidin-HRP solutions, the samples were allowed to undergo enzymatic reactions with TMB substrate solutions in the dark for an additional 30 min. The reaction was stopped using stop solution (supplied) and was measured for absorbance on an ELISA plate reader set at 450 nm. The amount of secreted cytokines (in pg/ml) was calculated from the absorbance values for each sample. The data represent the means of three independent experiments.
For dot blots, 10–30 μg of supernatant proteins from fibroblasts and myocyte culture were spotted on a polyvinylidene difluoride (PVDF) membrane and probed with antibodies of TGF-β, TNF-α, and IL-6 (R&D Systems), followed by incubation with HRP-conjugated secondary antibodies (Pierce). Immunoreactive bands were visualized by using enhanced chemiluminescence (NEN).
Results are expressed as means ± SE. Data were analyzed by two-way ANOVA, and the differences between groups were determined by the least-squares means test (SUPERNOVA). Differences were considered statistically significant at a value of P < 0.05.
Role of Myocytes in Collagen Upregulation During Hypertensive Hypertrophy: In Vitro Vs. Ex Vivo Experiments
When 10−8 M ANG II was perfused ex vivo through beating, nonworking rat hearts, the transcript levels of both type I and type III collagen increased considerably (2.5-fold compared with control; Fig. 1A, lanes 1 and 2). In contrast, the transcript levels did not change significantly when ANG II was added to the fibroblasts alone in vitro at various concentrations (83.2 ± 6.3 density units/10 μg RNA in F vs. 92.3 ± 10.2 in F+A for collagen type I; 80.1 ± 7.9 density units/10 μg RNA in F vs. 79.4 ± 5.8 in F+A for collagen type III, n = 5, P < 0.05; Fig. 1B). Collagen mRNA expression was significantly increased when 10−8 M ANG II was added and fibroblasts were cocultured with cardiac myocytes [∼2-fold compared with either fibroblast alone or fibroblast treated with ANG II (P < 0.01, n = 5; Fig. 1A, lanes 3–6)]. No change in collagen transcripts was observed with coculture of fibroblasts and myocytes alone (data not shown).
In parallel, a significant increase in collagen protein production was observed in each sample in vitro (5.925 ± 0.4 μg/ml in F vs. 19.09 ± 1.2 μg/ml in F+M+A, P < 0.01; Fig. 1C). The data showed a parallel increase in both collagen transcripts and protein levels.
ANG II is Necessary for Myocyte-Mediated Collagen Upregulation in Fibroblasts
To ascertain the role of ANG II in the process of collagen upregulation in vitro, fibroblasts were pretreated with the receptor blockers for AT1 receptors (losartan) and AT2 receptors (PD-123319) singly (10−6 M for 28 h) and in combination before coculture with cardiac myocytes. Pretreatment with losartan or PD-123319 alone resulted in downregulation of collagen I and collagen III transcripts (Fig. 2). However, the inhibition of collagen I transcripts was more significant when both losartan and PD-123319 were added (1.8-fold compared with F+M+A). The additive effect of the receptor blockers was more pronounced in the case of collagen I than collagen III transcript levels. This result suggests that the upregulation of collagen transcripts is mediated through both AT1 and AT2 receptors.
Role of Cytokines and Growth Factors in Collagen Upregulation by Fibroblast-Myocyte Interaction
Cellular cross-talk induces cytokine levels with ANG II.
To evaluate the participation of different factors in the upregulation of collagen, RNase protection assays were performed with fibroblast RNA using mouse cytokine templates as probes. As shown in Fig. 3A, different cytokine transcripts were upregulated in fibroblasts and myocytes with ANG II (lane 3) compared with either fibroblasts alone in culture (lane 2) or without myocytes (lane 1). The TGF-β isoforms (TGF-β1, -β2, and -β3), TNF-α, IL-6, interferon (IFN)-γ, and macrophage migration-inhibitory factor transcripts were found to be significantly upregulated in fibroblasts treated with myocytes and ANG II (lane 3). A considerable increase in TGF-β isoforms was also observed in lane 1 when fibroblasts were treated only with ANG II compared with control (lane 2). However, interactions between myocytes and fibroblasts (lane 3) induced TGF-β transcripts 1.2-fold greater than the level observed in the absence of myocytes (lane 1). No significant induction in the remaining cytokine transcripts tested was observed when fibroblasts were treated with only ANG II. The increase in TNF-α, IL-6, and TGF-β transcripts in fibroblasts and myocytes with ANG II by RNase protection assay was cross-checked by RT-PCR (data not shown).
Levels of IL-6 secreted by fibroblasts and myocytes cocultured in the presence of ANG II were 4.5-fold greater than levels of either fibroblasts or fibroblasts with ANG II (F+M+A = 521 ± 0.5 pg/ml; F = 114 ± 1.4 pg/ml; F+A = 132 ± 1.23 pg/ml; n = 4, P < 0.05; Fig. 3B); however, some induction in secreted IL-6 levels was observed in fibroblasts treated with myocytes without ANG II (F+M = 225 ± 2.1 pg/ml). Secreted TNF-α protein was detected in much less quantity; however, the level was fourfold higher in fibroblasts and myocytes with ANG II compared with fibroblasts alone in culture (36.6 ± 0.23 pg/ml in F+M+A vs. 9.3 ± 0.6 pg/ml in F; Fig. 3B). Fibroblasts with ANG II showed some induction (15 ± 0.36 pg/ml), but no change was found in fibroblasts and myocytes without ANG II. Because fibroblasts and myocytes with ANG II showed maximal upregulation of collagen transcripts as well as transcripts of different cytokines in vitro, we further confirmed the involvement of these factors in the induction of collagen transcript levels, as described below.
Effect of TGF-β, TNF-α, and IL-6 on collagen transcripts in vitro.
The cytokines TGF-β, IL-6, and TNF-α, when added separately to either fibroblasts alone or along with ANG II in the absence of myocytes, resulted in a significant upregulation of collagen transcripts. On the other hand, IFN-γ downregulated collagen transcripts to some extent when added to the fibroblasts in culture (data not shown).
The effect of different TGF-β2 concentrations (2, 4, and 8 ng/ml) on fibroblasts alone is shown in Fig. 4A. Maximum stimulation of collagen I transcripts (∼2.8 fold) was observed when 4 ng/ml TGF-β was added to fibroblasts alone (lane 4) for 12 h. Collagen transcripts were also upregulated appreciably in fibroblasts treated with 2 or 8 ng/ml TGF-β (lanes 3 and 5). The induction of the collagen I transcript level in fibroblasts by TGF-β was also comparable to induction by fibroblasts and myocytes with ANG II (data not shown).
Treatment of fibroblasts with TNF-α (0.1 and 0.5 ng/ml) resulted in upregulation of collagen I transcripts (Fig. 4B) compared with fibroblasts alone in culture or fibroblasts with ANG II. A significant increase (∼2-fold) in collagen I transcripts was observed when ANG II was added to fibroblasts treated with 0.5 ng/ml TNF-α (lane 5).
A similar increase in collagen I transcripts (2.3-fold) was also observed when different doses (20–50 ng/ml) of IL-6 were added to the fibroblasts in culture alone or with ANG II (Fig. 4C, lanes 3–5). Pretreatment of fibroblasts with anti-IL-6-neutralizing antibody (R&D Systems) for 4–6 h in the F+M+A model (lane 7) resulted in a significant downregulation of collagen I transcripts (2-fold) compared with the untreated fibroblasts and myocytes with ANG II (lane 6), whereas pretreatment of fibroblasts with only rat IgG had no effect on collagen I transcripts (Fig. 4C, lane 8).
When fibroblasts were pretreated with the angiotensin receptor blockers losartan and PD-123319 and incubated with TNF-α (0.5 ng/ml), upregulation of collagen I transcripts was inhibited (almost 31%; Fig. 4D, lane 6). The AT1 and AT2 receptor blockers, however, did not affect the induction of collagen I transcripts by IL-6 (2.3%; Fig. 4D, lane 4). These results suggest that cytokines play an important role in the process of collagen upregulation in the heart and that the effect of IL-6 on collagen gene activation is independent of AT1 and AT2 receptors.
Factor from Cardiac Myocytes is Secretary in Nature
To determine if the factor(s) from myocytes is secretary in nature, serum-free supernatant media from neonatal myocytes were added to the fibroblast culture alone (lane 3) and in presence of 10−8 M ANG II for 24 h (lane 4). Fibroblasts treated with the myocyte supernatant and ANG II showed significant stimulation of both collagen I and collagen III transcript levels (n = 3, P < 0.01; Fig. 5, A and B, lane 4).
Serum-free myocyte supernatant was examined for the presence of IL-6, TNF-α, and TGF-β, which can independently stimulate collagen synthesis in fibroblasts. After lyophilization and extensive dialysis, cell supernatants were blotted onto PVDF membranes and probed with antibodies to IL-6, TNF-α and TGF-β. TGF-β was detected in the myocyte supernatant, but IL-6 and TNF-α were not detected (Fig. 5C).
Estimation of Active TGF-β Production by Cardiac Cells
Because cardiac fibroblasts are also known to produce TGF-β de novo, the amount of active TGF-β produced by each cell type, i.e., myocyte and fibroblast, was estimated. TGF-β2 (5 ng/ml, GIBCO-BRL) inhibited the growth of mink lung epithelial CCL-64 cells by almost 90% (Fig. 6). Fibroblast supernatants were prepared as discussed in materials and methods and, when added at 20 μg/ml, inhibited the growth of CCL-64 cells by 32%. ANG II-supplemented myocyte supernatant (20 μg/ml) or F+M+A supernatant also inhibited the growth of mink lung epithelial cells by 77.25%. F+A and F+M supernatants also showed inhibition in growth, but the effect was much less (45%) compared with myocytes treated with ANG II. These data clearly suggest that cardiac myocytes are a more potent source of active TGF-β than cardiac fibroblasts, even in the presence of ANG II.
Active TGF-β (TGF-β1 and TGF-β2) was estimated from the cellular supernatant of different combinations with and without 10−8 mol/l ANG II by the sandwich ELISA technique as described in materials and methods. One hundred microliters of each of the cellular supernatant were used to measure biologically active TGF-β isoforms produced by cells alone or in combination. The activity was plotted against individual standards, and the result of this experiment is shown in Fig. 6B (n = 5). Maximum activity of TGF-β1 and -β2 was observed in F+M+A supernatant, which was 85% more compared with fibroblasts alone. The addition of ANG II to fibroblasts alone would induce the active TGF-β production by 27%. Isolated myocyte supernatant (with or without ANG II) has been shown to produce significantly more active TGF-β (both TGF-β1 and -β2) than the fibroblast supernatant (with or without ANG II). The additive effect of myocytes and ANG II to fibroblasts in culture induces the activity of TGF-β production to the maximum level.
Collagen Upregulation is Mediated Through Paracrine Action of TGF-β from Myocytes
To prove that TGF-β from cardiac myocytes plays a key role in the upregulation of collagen in fibroblasts, three experiments were performed. In the first experiment (Fig. 7A, lane 4), fibroblasts were pretreated with anti-TGF-β receptor RII antibody (5 μg/ml) with a subsequent addition of myocytes and ANG II [(F+anti-TGF-RII)+M+A]. In the second experiment, myocytes were pretreated with anti-TGF-β antibody (10 μg/ml) and the supernatant was added to fibroblasts that had been treated with ANG II [F+(M+anti-TGF-β)+A; lane 5]. Fibroblasts, fibroblasts with ANG II, and fibroblasts and myocytes with ANG II were used as controls (Fig. 7A, lanes 1–3). Pretreatment of fibroblasts with anti-TGF-β receptor RII antibody (Fig. 7A, lane 4) or pretreatment of myocytes with anti-TGF-β antibody (Fig. 7A, lane 5) resulted in a significant downregulation of collagen I transcripts (47% for 4.5 kb and 32% for 4.8 kb in lane 4, 49.5% for 4.5 kb and 48% for 4.8 kb in lane 5) compared with controls (lane 3) as well as myocytes treated with only rat IgG (lane 6).
In the third experiment, transient transfection of fibroblasts using Smad DN2 and Smad DN3 constructs (which blocked the TGF-β downstream signaling pathway in these cells) followed by cellular exposure to myocytes and ANG II (Smad2DN/F+M+A and Smad3DN/F+M+A; Fig. 7B, lanes 4 and 5) significantly downregulated collagen I gene transcripts (almost 42% reduction) compared with either fibroblasts and myocytes with ANG II or empty vector transfected fibroblasts (Fig. 7B, lanes 3 and 6). These results suggest that TGF-β from myocytes has a paracrine role in the process of collagen upregulation in vitro.
IL-6 transcripts were downregulated appreciably in fibroblast RNA samples that had been treated with anti-TGF-β receptor RII antibody before the addition of myocytes and ANG II (Fig. 7C, lane 3). Compared with fibroblasts and myocytes with ANG II (lane 2), IL-6 transcripts were also downregulated in fibroblasts, when myocytes pretreated with anti-TGF-β antibody were added to fibroblasts and ANG II (lane 4). Collagen transcripts were also downregulated in these samples, as shown in Fig. 7A.
The molecular mechanism for increased collagen formation (fibrosis) during the chronic phase of hypertrophy and heart failure is not yet clearly understood. Previously, Kim et al. (17) reported that ANG II modulates collagen production. Also, others have shown that TGF-β plays a role in collagen upregulation (19, 21). Our report is the first, to the best of our knowledge, to demonstrate that in addition to autocrine action of fibroblasts, the paracrine action of active TGF-β from myocytes on fibroblasts also plays a causal role in the upregulation of collagen.
This study has presented several new and intriguing observations. Specifically, this study is the first to demonstrate 1) a paracrine role of cardiomyocytes by their secreting active TGF-β in ANG II-mediated upregulation of collagen by fibroblasts (Fig. 7), and 2) participation of cytokines such as TNF-α and IL-6 in collagen upregulation by fibroblasts (Fig. 4). Our data also showed that ANG II-induced collagen upregulation mediated through both AT1 and AT2 receptors (Fig. 2).
We have shown previously that there is a major difference between results from in vitro and in vivo studies with respect to collagen upregulation by fibroblasts in response to treatment with ANG II (25). This paradox led us to believe that other signals or factors may be necessary for collagen upregulation and that other cardiac cell types may play a direct or indirect role in activation of this gene. We have already shown that fibroblast-myocyte cross-talk is essential for ANG II upregulation of collagen expression in vitro (25). Experiments in the present study were designed to address the question of how myocytes participate in stimulating ANG II-mediated collagen synthesis by fibroblasts.
The role of ANG II in collagen upregulation is well documented (17, 25). However, the underlying mechanism, especially the role of ANG II receptors, in collagen production is not well understood. Our data showed that treatment with AT1 and AT2 receptor blockers significantly downregulated collagen gene expression (Fig. 2). Importantly, when a combination of AT1 and AT2 receptor blockers was administered, an almost 90% downregulation of collagen gene expression was observed, significantly more compared with what was blocked by losartan or PD-123319 alone. These data suggest that ANG II-induced collagen upregulation is mediated not only through AT1 receptors but also through AT2 receptors (Fig. 2). In support of this observation, previously, in an in vivo study, when spontaneously hypertensive rats (SHRs) were treated with AT1 and AT2 receptor blockers separately and in combination, a significant reduction in collagen content was observed with either the AT1 blocker (losartan) or the AT2 blocker (PD-123319) separately (29% and 39%, respectively). When a combination was given, an 80% inhibition of collagen content was observed (unpublished observations), suggesting that collagen upregulation is mediated via both AT1 and AT2 receptors.
Until now, the role of cardiomyocytes in collagen gene upregulation was not known. Previous data from our studies convincingly suggest that rat cardiac myocytes (both neonatal and adult) play an important role in stimulating collagen gene expression in vitro in response to ANG II (25). The fact that serum-free myocyte supernatant also upregulates collagen transcripts in response to ANG II, confirming the role of cardiac myocytes in triggering collagen production and demonstrates that the inducing factor is secretory in nature. Furthermore, in this study, we have demonstrated convincingly that myocytes in addition to fibroblasts and ANG II not only stimulate TGF-β production but also maximally convert inactive TGF-β to the active form. This has been shown by both indirect and direct assay, as summarized in Fig. 6, A and B. This is a new observation and could open new doors to study the mechanism and define the condition of such a conversion that stimulates collagen production and thus promotes fibrosis.
A number of growth factors have been shown to regulate collagen production both in vitro and in vivo (6). The ability of cells to produce or release growth factors in response to load appears to be a key modulator of collagen production. However, the responses, in terms of gene expression and cell replication, appear to be cell type specific (3). Interactions between growth factors and mechanical load may regulate fibroblast activation, and collagen deposition is a likely outcome of these combined effects. One of the important growth factors that induce collagen deposition appears to be TGF-β. In the rat collagen α1(1) gene promoter, the TGF-β1-responsive element contains a potential binding site for the transcription factor nuclear factor 1, through which TGF-β1 activates transcription of the collagen gene (20). TGF-β1, -β2, and -β3 transcripts were upregulated in fibroblasts cocultured with myocytes and ANG II (the F+M+A model), resulting in induced collagen gene expression (both type I and III). This finding is of pathophysiological significance, as we have shown previously that during the chronic phase of hypertrophy in SHRs, a significant increase in type III collagen occurs (39). These findings, as well as those from several other studies (19, 21, 25), have suggested that TGF-β plays an important role in the regulation of collagen synthesis in the myocardium, but the underlying mechanism is still not clear. Kupfahl et al. (19) also showed that ANG II treatment in isolated cardiac fibroblasts significantly increased TGF-β1 mRNA, whereas ANG II had no effect on collagen type I or collagen type III mRNA; this corroborates our observation on the effect of ANG II in vitro. The recent identification of Smads as effector proteins (23) provided a major advance in understanding TGF-β1 postreceptor signaling pathways. Many workers have shown that transient transfection of Smad 2 and Smad 3 dominant negative mutants blocked the TGF-β downstream signaling pathways in isolated cell systems (26). Activation of TGF-β1 and increased expression of novel downstream Smad signaling proteins were observed in infarct scars during chronic myocardial infarction in rat hearts (16). Although it was postulated that ANG II potentiates or possibly regulates TGF-β production in fibroblasts (27), in both fibroblasts and myofibroblasts (10), our data suggest that ANG II-mediated collagen upregulation is a result of cross-talk between cardiomyocytes and fibroblasts via the paracrine action of TGF-β from myocytes (Fig. 7).
We have demonstrated that, in addition to TGF-β, TNF-α and IL-6 could independently induce collagen upregulation by fibroblasts. Interestingly, our data showed that AT2 and AT1 receptor blockers inhibited TNF-α-induced upregulation of collagen but not IL-6. Our data suggest that TNF-α-mediated upregulation of collagen requires ANG II, whereas IL-6-mediated upregulation is independent of ANG II. Further studies are necessary to elucidate these processes. Downregulation of collagen transcripts by preincubation of fibroblasts with IL-6-neutralizing antibodies confirms the role of IL-6 in this process. Results from RNase protection assay (Fig. 3A) suggest that myocytes induce the production of cytokines in fibroblast cultures only under the influence of ANG II. However, whether these cytokines were produced directly or indirectly during the cross-talk between myocytes and fibroblasts needs to be determined.
Once the secretory nature of the myocyte factor(s) that upregulates collagen transcription in fibroblasts was established, our next aim was to determine whether these cytokines were actually released by myocytes. We detected TGF-β but not IL-6 or TNF-α in the myocyte supernatant. Azuma et al. (1) also reported activation of TGF-β1 transcript by ANG II in cardiomyocytes. This finding led us to hypothesize that TGF-β from cardiomyocytes, but not from fibroblasts, can induce cytokine production in ANG II-treated fibroblasts. To prove this hypothesis, we designed an experiment to block the action of TGF-β released by cardiac myocytes. In one set, fibroblasts were pretreated with TGF-β receptor RII neutralizing antibody, followed by the addition of myocytes and ANG II. In another set, cardiac myocytes were preincubated with TGF-β-neutralizing antibody, and the conditioned medium was used to supplement fibroblasts, which were then treated with ANG II. Both treatments were designed to prevent secretion of any active TGF-β from myocytes to act on fibroblasts. In both cases, collagen type I transcripts were found to be downregulated compared with either the F+M+A model or fibroblasts treated with myocyte supernatant and ANG II (Fig. 7A), suggesting that TGF-β produced by myocytes plays an important role in collagen upregulation. IL-6 was also found to be downregulated in these samples compared with the F+M+A model (Fig. 7C), suggesting that IL-6 produced by fibroblasts also plays a role in collagen upregulation. Transient transfection of fibroblasts with Smad DN2 and Smad DN3 mutants blocked the TGF-β downstream signaling pathway in these cells. In this setting, TGF-β from any outside source could not potentiate the downstream signaling cascades in fibroblasts, and thus collagen upregulation would be prevented. Fibroblasts transfected with the Smad DN2 and Smad DN3 mutants showed significant downregulation of collagen type I transcripts compared with the F+M+A model (Fig. 7B), confirming the paracrine role of myocytes in collagen production. Thus we have convincingly shown that myocytes must synergize with fibroblasts in controlling ANG II-mediated collagen upregulation and that active TGF-β from myocytes appears to be instrumental in this complex process.
Cardiovascular cells in vivo normally reside in an environment in which they are constantly exposed to mechanical load, and changes in those mechanical forces or additional stimuli might be a causal factor for heart failure. Although the data are currently limited, many known signaling pathways appear to be activated by load (5, 24, 38). Our study suggests that ANG II and TGF-β (from myocytes and from fibroblasts) activate cytokines such as IL-6 from fibroblasts to upregulate collagen production during stress (ANG II treatment). However, any relation between the activated postreceptor mechanisms for ANG II and TGF-β in heart failure is still not known. Our main focus in this study is to define the role of myocytes in collagen upregulation in the presence of ANG II. We have clearly demonstrated that even though a slight increase (≈10%) was observed in fibroblasts alone treated with ANG II, the addition of myocytes in the presence of ANG II always resulted in significantly more upregulation in collagen transcripts (80%). Our data also suggest that TGF-β plays a role in this process to upregulate collagen under the influence of cardiac myocytes. One important criterion to be considered is the nature of TGF-β secreted by each cell type (active or inactive form). In this study, we have shown that the TGF-β coming from fibroblasts perhaps is not fully active. Whenever myocytes were added, an increase in collagen production was found, which was not the case with fibroblasts alone with ANG II (Fig. 1). One possibility is that myocytes, when added to fibroblasts, convert inactive TGF-β to the active form. Now, we have convincingly demonstrated by quantifying active TGF-β in the presence and absence of myocytes that myocytes produce active TGF-β that stimulates collagen production by fibroblasts in the presence of ANG II (Fig. 6, A and B). ANG II regulates this process because the upregulation can be inhibited by angiotensin receptor blockers. This study was designed primarily to understand the mechanism of collagen upregulation in the heart during hypertrophy. However, whether upregulation of collagen means increased transcription or increased stability or both for collagen type I and collegen type III has not been addressed in this study. Our data suggest that the mechanism of collagen upregulation is a very complicated phenomenon and may include cellular cross-talk, synergistic effects of different growth factors, or multiple signaling pathways. We have shown that cardiomyocytes play an important role in stimulating collagen gene expression in fibroblasts. We believe that myocytes release factors such as TGF-β, along with an additive effect with TGF-β from fibroblasts, which in turn stimulates secretion of other cytokines such as IL-6 or TNF-α from fibroblasts. IL-6 and/or TNF-α and then trigger collagen gene upregulation by fibroblasts under the influence of ANG II (Fig. 8). In addition to the cytokines and growth factors investigated in this study, there are other potent inducers of collagen production, such as endothelin I (4) and norepinephrine (2), which we have not examined. Once the role of all these factors is identified, we might be able to outline specific pathways for excessive collagen accumulation. A greater understanding of the processes involved in mechanosignaling and how growth factors interact with these pathways might lead to the development of more effective drug regimens to combat the deadly disease of heart failure.
This study was funded by National Heart, Lung, and Blood Institute Grants HL-27838 and HL-47794 (to S. Sen).
We sincerely thank Dr. P. Howe, Dept. of Cell Biology, Cleveland Clinic Foundation, for the kind gift of the Smad 2 DN and Smad 3 DN constructs and thank Drs. G. Wildey and S. Karnik for critical review of the manuscript. We also acknowledge skilled secretarial help from Joanne Holl and editorial assistance by Christine Kassuba.
The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked “advertisement” in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
- Copyright © 2004 by the American Physiological Society