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Cardiovascular Research 2007 74(2):323-333; doi:10.1016/j.cardiores.2006.12.010
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Copyright © 2006, European Society of Cardiology

Induction of pulmonary connective tissue growth factor in heart failure is associated with pulmonary parenchymal and vascular remodeling

Mohammad Shakil Ahmeda,c, Erik Øiea,b, Leif Erik Vingea,c, Thomas G. von Luedera,c, Toril Attramadala and Håvard Attramadala,b,c,*

aInstitute for Surgical Research, University of Oslo, N-0027 Oslo, Norway
bDepartment of Cardiology, Rikshospitalet University Hospital, Sognsvannsveien 20, N-0027 Oslo, Norway
cFaculty Division Rikshospitalet, University of Oslo, N-0027 Oslo, Norway

* Corresponding author. Institute for Surgical Research, A3.1013, Rikshospitalet University Hospital, N-0027 Oslo, Norway. Tel.: +47 23 073520; fax: +47 23 07353. Email address: havard.attramadal{at}medisin.uio.no

Received 24 May 2006; revised 7 December 2006; accepted 11 December 2006


   Abstract
Top
 Abstract
1. Introduction
 2. Methods
 3. Results
 4. Discussion
 Acknowledgments
 References
 
Objectives: Pulmonary remodeling is a well recognized consequence of heart failure (HF). However, the cellular and molecular mechanisms orchestrating the structural alterations of the lungs in HF are poorly understood. We have previously reported induction of the profibrotic peptide connective tissue growth factor (CTGF) in myocardial tissue of rats with HF, suggesting a role of CTGF during myocardial remodeling. The aim of the present study was to explore the potential role of CTGF in pulmonary remodeling in HF.

Methods: Pulmonary tissue samples were obtained from rats with myocardial infarction (MI) subsequent to ligation of the left coronary artery. Real-time quantitative RT-PCR was employed to investigate mRNA levels. The cellular distribution of CTGF was analysed by immunohistochemistry.

Results: Seven days after induction of myocardial infarction (MI) and HF in rats we found 2.3-fold and 1.9-fold increase of pulmonary transforming growth factor-β1 and procollagen {alpha}1(I) mRNA levels, respectively, and typical morphological characteristics of pulmonary remodeling including interstitial fibrosis and medial thickening of pulmonary arteries. Pulmonary CTGF mRNA levels were substantially elevated in HF rats compared to sham-operated rats (4-fold; P<0.05) and corresponded with similar increase (3-fold; P<0.05) of pulmonary CTGF protein contents. Immunohistochemical analysis revealed increased pulmonary anti-CTGF immunoreactivity in HF, with immunostaining predominantly localized to alveolar macrophages and interstitial fibroblasts. Isolated alveolar macrophages from HF rats demonstrated substantial induction of CTGF mRNA expression (16-fold; p<0.05). Interestingly, platelets caused robust induction of CTGF mRNA expression in alveolar macrophages upon co-culture in vitro.

Conclusion: Pulmonary CTGF was substantially increased in parallel with pulmonary remodeling in rats with HF. Our data indicate that alveolar macrophages are a major source of increased pulmonary CTGF in HF and that CTGF may be a player in the profibrotic mechanisms associated with HF.

KEYWORDS Heart failure; Growth factor; Remodeling; Macrophages; Pulmonary circulation


   1. Introduction
Top
 Abstract
 1. Introduction
2. Methods
 3. Results
 4. Discussion
 Acknowledgments
 References
 
Heart failure (HF), the terminal end-point of cardiovascular disease of diverse etiologies, carries high mortality despite implementation of several new treatment modalities during recent years. The pathophysiologic interactions between the heart and the lungs in HF are well recognized. Indeed, HF is associated with pulmonary arterial hypertension and structural alterations of the lungs [1,2]. The most distinctive features of these structural alterations include thickening of interstitial and alveolar septa with increased deposition of collagen and reticulin, proliferation of myofibroblasts (pericytes) and medial thickening of pulmonary arteries [1]. However, the cellular and molecular mechanisms orchestrating these structural alterations of pulmonary tissue during evolving HF are poorly understood. Previous studies have demonstrated that not only hemodynamic alterations, but also increased circulating levels of neurohormones and cytokines known to be involved in myocardial remodeling may directly affect the lungs during evolving HF [3–6].

Connective tissue growth factor (CTGF) is a member of the CCN (acronym of Cyr61/CEF-10, CTGF/Fisp-12, and Nov) family of growth factors [7–9]. These growth factors are secreted, extracellular matrix (ECM)-associated proteins involved in multiple cellular events including synthesis of ECM and proliferation of fibroblasts [10,11]. Indeed, CTGF is associated with several fibrotic disorders [12–14], and it has been reported that CTGF stimulates both proliferation of fibroblasts as well as deposition of ECM through increased secretion of collagen and fibronectin. In this respect, CTGF has been shown to mediate at least part of the actions of transforming growth factor-β (TGF-β), which is an important stimulator of fibrosis [15]. Furthermore, tissue expression of CTGF has been shown to correlate with development of pulmonary fibrosis of non-cardiac etiology [16,12]. We therefore hypothesized that CTGF is a mediator of pulmonary remodeling during evolving HF. To further explore the potential role of CTGF in pulmonary remodeling associated with HF, we employed the post-myocardial infarction HF model in rats and investigated the cellular distribution and regulation of pulmonary CTGF in HF.


   2. Methods
Top
 Abstract
 1. Introduction
 2. Methods
3. Results
 4. Discussion
 Acknowledgments
 References
 
2.1 Animal preparation and study protocol
The left coronary artery-ligated rat model of HF was employed as previously described [17]. Briefly, male Wistar rats (~ 250 g) were anesthetized with 1% isoflurane in 1/3 O2 and 2/3 N2O and subjected to left thoracotomy with subsequent ligation of the proximal portion of the left coronary artery. Except for ligation of the artery, sham-operated rats underwent the same procedure. In rats subjected to ligation of the left coronary artery, mortality was 37% at 24 h after the surgical procedure. In the time span from 24 h to 7 days post-MI, mortality was 6%. Rats subjected to MI that met the following criteria at the indicated time points of the study (7, 15, and 56 days post-MI) were included as HF rats in the study: Left ventricular (LV) end-diastolic pressure (LVEDP) > 15 mm Hg and MI size >40% of the ventricular circumference. In the first series of experiments HF rats (n=8) and sham-operated rats (n=9) were euthanized 7 days after induction of MI, and pulmonary tissue samples were obtained for subsequent analysis of gene expression, tissue contents, and cellular distribution of CTGF.

The purpose of the second series of experiments was to investigate the mRNA levels of CTGF and TGF-β1 in isolated alveolar macrophages of HF rats (n=8) and sham-operated rats (n=6) 15 days after surgery. In the third series of experiments HF rats (n=7) and sham-operated rats (n=7) were euthanized 8 weeks after induction of MI, and pulmonary tissue samples were obtained for gene expression and immunohistochemical analyses.

The animal experiments and housing were in accordance with institutional guidelines and national legislation conforming to the Guide for the Care and Use of Laboratory Animals published by the US National Institutes of Health (NIH Publication No. 85-23, revised 1996).

2.2 Hemodynamic measurements and tissue sampling
On the day of the experiments, the rats were anesthetized with 1% isoflurane in 1/3 O2 and 2/3 N2O. Aortic pressure, LV end-systolic pressure (LVSP), and LVEDP were recorded by a 2F micromanometer-tipped catheter (model SPR-407, Millar Instruments, Houston, USA) inserted through the right carotid artery. After completion of the hemodynamic recordings, the rats were sacrificed by excision of the heart.

Heart and lung weights were determined, and pulmonary tissue samples from the rats of the first series of experiments were snap-frozen in liquid nitrogen and stored at –70 °C until analysis. The lungs from the rats of the second series of experiments were dried in order to determine pulmonary dry weight. Pulmonary tissue samples for immunohistochemistry were fixed in Bouin's solution [2% paraformaldehyde and 0.2% picric acid in phosphate-buffered saline (PBS)], embedded in paraffin wax, and stored at 4 °C.

2.3 Bronchoalveolar lavage and isolation of alveolar macrophages
Alveolar macrophages were obtained by bronchoalveolar lavage of rats as described previously [18]. Phosphate-buffered saline (PBS) containing heparin (10 U/ml) was instilled in the trachea using a volume equivalent to total lung capacity. The lavage fluid was aspirated and the cells in the aspirate were recovered by centrifugation (800 xg at 4 °C for 5 min). The procedure was repeated three times and the collected cells were either 1) snap-frozen in liquid nitrogen and kept at –70 °C for subsequent analysis of gene expression, 2) cultured in Dulbecco's modified Eagle's medium supplemented with 10% fetal calf serum (Bio Whitaker) and 40 µg/ml of garamycin in a humidified atmosphere containing 5% CO2 at 37 °C and employed for analysis of ligand-stimulated induction of CTGF mRNA expression, or 3) dispersed onto glass slides and fixed in 50% methanol (v/v in acetone) and immunostained with monoclonal mouse anti-rat CD68 IgG1 (Serotec, Oxford, UK) in order to assess the homogeneity of macrophages aspirated by bronchoalveolar lavage.

2.4 Real-time quantitative reverse-transcription polymerase chain reaction
Total RNA was isolated from pulmonary tissue samples or alveolar macrophages using RNeasy (Qiagen, Germany) and subsequently treated with RNase-free DNase I (RQ1 DNase I, Promega). Reverse transcription (RT) and PCR of each sample were run in triplicates using the TaqMan PCR Core Reagent Kit and the ABI Prism 7900 Sequence Detection System and software (Applied Biosystems, Foster City, CA, USA) according to the manufacturer's instructions. Sequence specific PCR primers and TaqMan probes were designed using the Primer Express software version 1.5 (Applied Biosystems); see Table 1 for details. Each pair of oligonucleotide primers was designed to span an intron to avoid amplification of genomic DNA. SyBr Green assays (Table 1) were performed using 2x SyBr Green Universal Master Mix (Applied Biosystems) and 300 nM sense and anti-sense oligonucleotide primers. The specificity of the SyBr Green assays was assessed by melting point analysis. Prepro ET-1 mRNA levels were analysed using rat ET-1 pre-developed assay reagents (Applied Biosystems). A standard curve was generated by amplifications of cDNA obtained from serial dilutions of lung total RNA. For all specific mRNA amplified linear inverse correlations were observed between amount of RNA and CT value (number of cycles at threshold lines). Gene expression was presented relative to the levels of 18S ribosomal RNA as the housekeeping gene.


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  		Table 1 Characteristics of the real-time PCR assays used in the study

 
2.5 Western blot analysis
Pulmonary tissue samples were homogenized and solubilised in buffer containing 3% Triton X-100, 300 mM NaCl, 100 mM Tris; pH 7.3, 1 mM Na3VO4, 20 mM EDTA, 1% NP-40, 2 mM PMSF, and protease inhibitors (1 µg/ml each of aprotinin, pepstatin A and leupeptin). After solubilisation, the extracts were centrifuged (8000 xg) at 4 °C for 15 min, denatured in Laemmli buffer, and subsequently separated on 12% SDS-PAGE, transferred by electroblotting to a PVDF membrane, and finally subjected to immunoblot analysis as previously described [19]. Generation of rabbit anti-CTGF antiserum and purification of anti-CTGF-specific IgG have previously been described [19].

2.6 Immunocytochemistry/histochemistry and histological analysis
Pulmonary tissue for histochemical and immunohistochemical analysis was from HF rats (n=2, each time point) and sham-operated rats (n=2, each time point) euthanized 7 and 56 days after induction of MI. Immunohistochemical analysis of pulmonary tissue sections was performed as described previously [20] using the purified rabbit anti-CTGF IgG described above. In addition, monoclonal mouse anti-rat CD68 IgG1 (Serotec, Oxford, UK) was used for detection of alveolar macrophages in pulmonary tissue sections. The avidin–biotin–peroxidase system (Vectastain Elite kit, Vector Laboratories, CA, USA) was used for signal amplification. Purified rabbit or mouse IgG from non-immune serum or omission of primary antibody was used as negative controls. Other pulmonary tissue sections of HF rats (n=2, each time point) and sham-operated rats (n=2, each time point) were stained with van Gieson's solution and hematoxylin and at least two sections (4–6 fields/section) per animal were observed under light microscopy at 400x magnification to assess the contents of extracellular collagen.

2.7 Co-incubation of alveolar macrophages and platelets
Platelets were isolated as described previously [21]. Briefly, venous blood was drawn from rats and collected into acid-citrate-dextrose buffer and centrifuged at room temperature for 10 min at 100 xg to obtain platelet-enriched plasma. The platelet-enriched plasma was centrifuged at 800 xg for 15 min. The pellet of platelets was washed once and resuspended in PBS. Rat alveolar macrophages were seeded at a density of 2x105 cells/60 mm dish in serum-free DMEM and co-cultured with platelets (1x107 platelets/60 mm dish). All co-cultures were performed in triplicates. The macrophages were analyzed following 20 h co-incubation with platelets.

2.8 Statistical analysis
All values are expressed as mean±SEM. Statistical analysis was assessed by the Mann–Whitney nonparametric test and linear regression was used to test the relation between the variables when relevant. P values <0.05 were considered to be statistically significant.


   3. Results
Top
 Abstract
 1. Introduction
 2. Methods
 3. Results
4. Discussion
 Acknowledgments
 References
 
3.1 Hemodynamic measurements
Table 2 shows mean arterial pressure (MAP), LVEDP and LVSP of sham-operated rats and HF rats 7, 15 and 56 days after the surgical procedure. The hemodynamic measurements confirmed LV dysfunction in the HF rats with MAP and LVSP significantly decreased and LVEDP significantly increased compared to the sham-operated group.


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  		Table 2 Hemodynamic measurements in sham-operated rats and HF rats 7, 15 and 56 days after MI or sham operation

 
3.2 Heart and lung weights
The heart weight-to-body weight ratio was increased in the HF rats compared to the sham rats (Table 2; P<0.05) demonstrating robust cardiac hypertrophy in the HF rats. Wet lung weight-to-body weight ratio displayed a significant increase in the HF group compared with the sham-operated group (P<0.05) consistent with pulmonary congestion. Dry lung weight-to-body weight ratio was also increased in the HF group (2.6-fold relative to sham-operated rats, Table 2; P<0.05) providing evidence of pulmonary remodeling.

3.3 Pulmonary gene expression
Pulmonary CTGF mRNA levels were significantly increased in the HF rats compared to sham-operated rats (4-fold; P<0.01; Fig. 1A). In addition, pulmonary TGF-β1 and procollagen {alpha}1(I) mRNA levels were also elevated in the HF rats compared to sham rats (2.3-fold and 1.9-fold, respectively; P<0.01; Fig. 1B and C), whereas procollagen {alpha}1 (III) mRNA did not display significant alterations (Fig. 1D). We also investigated the expression of pulmonary CTGF and procollagen {alpha}1 (I) mRNA levels at a time point of chronic HF. Real-time quantitative PCR analysis revealed significant increase of CTGF and procollagen {alpha}1 (I) mRNA levels in HF rats of 8 weeks (56 days) after MI compared to their corresponding sham-operated rats (2-fold and 1.4-fold, respectively; P<0.01; Fig. 5E and F).


Figure 1
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  		Fig. 1 Pulmonary mRNA levels of CTGF (A), TGF-β1 (B), procollagen {alpha}1 (I) (C), and procollagen {alpha}1 (III) (D) in rats 7 days after induction of MI or sham-operation. Total RNA of pulmonary tissue samples was extracted and mRNA levels were analyzed by quantitative real-time RT-PCR. Each RNA sample was analyzed in triplicates. The data in the histograms are ratios of indicated mRNA relative to 18S rRNA levels. The values are mean±SEM of each group for sham-operated rats (n=9) and HF rats (n=8). *P<0.05 vs. sham-operated group.

 

Figure 5
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  		Fig. 5 Representative photomicrographs of rat pulmonary tissue sections 8 weeks after sham-operation or induction of MI stained with van Gieson's solution and hematoxylin (A and B). Section from pulmonary tissue of sham-operated rat demonstrating thin alveolar septa and scarce amounts of extracellular collagen (A). Pulmonary section from HF rat demonstrating increased thickness of the alveolar septa and substantial interstitial collagen accumulation (arrowheads) (B). Representative photomicrographs demonstrating CTGF immunostaining of pulmonary tissue sections of sham-operated rat and HF rat 8 weeks after induction of MI (C and D). Low levels of CTGF immunoreactivity could be discerned in alveolar epithelial cells (arrows) and subepithelial interstitial tissue (arrowheads) in sections from sham-operated rat (C). Immunoreactive macrophages could not be detected in pulmonary sections from sham rats. Numerous alveolar macrophages demonstrating strong anti-CTGF immunoreactivity (arrowheads) can be discerned in the pulmonary sections of HF rat in addition to CTGF immunostaining in alveolar interstitial tissue (D). Magnification A–B: x200 (bars=100 µm). Magnification C–D: x400 (bars=50 µm). Pulmonary mRNA levels of CTGF (E) and procollagen {alpha}1 (I) (F) in rats 8 weeks after induction of MI or sham-operation. The data in the histograms are ratios of indicated mRNA relative to 18S rRNA levels. The values are mean±SEM of each group for sham-operated rats (n=7) and HF rats (n=7). *P<0.01 vs. sham-operated group.

 
3.4 Pulmonary CTGF levels
Western blot analysis of pulmonary tissue extracts revealed immunoreactivity corresponding to the expected molecular mass of CTGF (38 kDa). The immunoreactive band corresponded with the immunoreactive band in extracts from COS-7 cells transfected with expression vector encoding rat CTGF cDNA (data not shown). As shown in Fig. 2, 3-fold increase of CTGF levels was found in pulmonary tissue of HF rats compared with that of sham-operated rats (P<0.05).


Figure 2
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  		Fig. 2 Western blot analysis of pulmonary CTGF 7 days after sham-operation or induction of MI. Representative photograph of Western blot showing specific immunoreactive band at 38 kD representing rat CTGF (A). Seventy µg of protein was loaded in each lane of the SDS-polyacrylamide gel. Histograms of pulmonary tissue contents of CTGF determined by densitometric analysis of immunoreactive bands (B). The data are mean±SEM (arbitrary units) of each group (sham-operated rats, n=4; HF rats, n=6). *P<0.05 vs. sham-operated rats.

 
3.5 Histochemical analysis of pulmonary tissue sections
Fig. 3 shows photomicrographs of pulmonary sections from sham-operated rats and HF rats (7 days after MI) stained with van Gieson's solution and hematoxylin. Thin alveolar walls with few or no alveolar macrophages were characteristic of pulmonary sections of sham-operated rats (A). Conversely, in the pulmonary sections of HF rats structural alterations characterized by increase of alveolar thickness with increased interstitial and subepithelial deposition of collagen were found (B). Furthermore, substantially increased deposition of collagen in the media of small arteries was observed in pulmonary tissue of HF rats (D and F) vs. that of sham-operated rats (C and E) as visualized in van Gieson's solution/hematoxylin-stained sections. Fig. 5 shows photomicrographs of pulmonary sections from sham-operated rats and HF rats 8 weeks (56 days) after MI stained with van Gieson's solution and hematoxylin (A and B). The characteristic features of pulmonary tissue of HF rats 8 weeks after MI were similar to those observed in pulmonary tissue sections from HF rats 7 days after MI.


Figure 3
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  		Fig. 3 Representative photomicrographs of rat pulmonary tissue sections stained with van Gieson's solution and hematoxylin. Section from pulmonary tissue of sham-operated rat demonstrating thin alveolar septa and scarce amounts of extracellular collagen (A). Pulmonary section from HF rat demonstrating increased thickness of the alveolar septa and substantial interstitial collagen accumulation (arrowheads) 7 days after induction of MI (B). Pulmonary artery of sham-operated rat (C and E) and HF rat (D and F) with extensive circumferential medial thickening and fibrosis (arrowheads) could be discerned only in the HF rat. Magnification A–D: x200 (scale bars=100 µm). Magnification E–F: x400 (scale bars=50 µm).

 
3.6 Immunohistochemical analysis of pulmonary distribution of CTGF
Representative photomicrographs of immunohistochemical staining of CTGF in pulmonary tissue sections 7 days after MI are shown in Fig. 4. Immunohistochemical analysis of pulmonary tissue sections of sham-operated rats revealed weak anti-CTGF immunostaining of alveolar epithelial cells and interstitial macrophages (A). In HF rats, on the other hand, CTGF immunoreactivity of alveolar macrophages (C), which were predominantly anti-CD68-positive macrophages (D), was substantially increased. Enhanced CTGF immunoreactivities of spindle-shaped fibroblast-like cells of the alveolar septal walls could also be discerned, whereas immunostaining of alveolar epithelial cells were weak (C). No immunostaining was seen in pulmonary tissue sections after omission of primary antibody or after incubation with non-immune rabbit serum, demonstrating specificity of the CTGF immunostaining (B). Interestingly, the number of anti-CD68 positive alveolar macrophages was substantially increased in pulmonary sections of HF rats (E) compared to sham-operated rats (F). Another consistent finding demonstrated in Fig. 4G was anti-CTGF immunostaining of spindle-shaped cells of the medial layer of pulmonary arteries of HF rats. On the contrary, immunostaining of the medial layer of pulmonary arteries of sham-operated rats could not be discerned (H). Furthermore, in order to investigate the expression of pulmonary CTGF at a time point of chronic HF, immunohistochemical analysis of pulmonary tissue sections from rats 8 weeks post-MI was performed. As demonstrated in Fig. 5 (C and D), similar patterns of anti-CTGF immunostaining were observed in pulmonary tissue sections of rats 8 weeks post-MI as in those from rats 7 days post-MI. Robust CTGF immunostaining could be discerned in tissue sections of HF rats, predominantly in alveolar macrophages.


Figure 4
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  		Fig. 4 Representative photomicrographs demonstrating immunostaining of pulmonary tissue sections of sham-operated rat and HF rat. Low levels of CTGF immunoreactivity in alveolar epithelial cells (arrows) and subepithelial interstitial tissue (arrowheads) in sections from sham-operated rat (A). Pulmonary tissue sections incubated with non-immune rabbit serum as control do not exhibit immunostaining of any of the cellular elements demonstrating specificity of the purified anti-CTGF IgG (B). Strong anti-CTGF immunoreactivity in alveolar macrophages (arrowheads) and in alveolar interstitial tissue (arrows) in pulmonary section of HF rat (C). Pulmonary section (serial section of tissue in C) immunostained with mouse anti-rat CD68 IgG1 as marker of alveolar macrophages (arrowheads) (D). Note: Abundant anti-CD68-positive alveolar macrophages in HF rat (E) compared to sham-operated rat (F). CTGF immunoreactivity in pulmonary artery of HF rat (G) and sham-operated rat (H) with CTGF immunoreactivity only discernable in the media of the HF rat. Magnification A–D and G–H: x400 (scale bars=50 µm). Magnification E–F: x200 (scale bars=100 µm).

 
3.7 CTGF, prepro-endothelin-1 (prepro-ET-1) and TGF-β1 mRNA levels in isolated alveolar macrophages
As shown in Fig. 6A and B, CTGF and prepro-ET-1 mRNA levels were substantially elevated in alveolar macrophages obtained by bronchoalveolar lavage of HF rats as compared to those obtained from sham-operated rats (16-fold and 2.5-fold respectively; P<0.05), whereas TGF-β1 mRNA levels were not significantly altered in alveolar macrophages among the groups (C). The homogeneity of isolated macrophages was assessed by immunocytochemistry using monoclonal anti-CD68 IgG1. More than 95% of the cells obtained by bronchoalveolar lavage were anti-CD68 positive macrophages (D and E).


Figure 6
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  		Fig. 6 CTGF (A), ET-1 (B) and TGF-β1 (C) mRNA levels in alveolar macrophages from rats 15 days after induction of MI or sham-operation. mRNA levels were analyzed by quantitative real-time RT-PCR. Each RNA sample was analyzed in triplicates. The data are presented as ratios of indicated mRNA relative to the levels of 18S rRNA and are mean±SEM of each group (sham-operated rats, n=8; HF rats, n=6). *P<0.05 vs. sham-operated group. Immunocytochemical analysis of cells aspirated by bronchoalveolar lavage. The cells were immunostained with anti-CD68 IgG1 (D) or non-immune mouse serum (E) as negative control. Magnification D–E: x200 (bars=100 µm).

 
3.8 Effects of angiotensin II (Ang II) and TGF-β1 on expression of CTGF mRNA in alveolar macrophages
Expression of CTGF mRNA in cultured alveolar macrophages was analyzed by real-time quantitative RT-PCR. As shown in Fig. 7 (A and B), neither Ang II (100 nM) nor TGF-β1 (5 ng/ml) induced expression of CTGF mRNA in alveolar macrophages after 24 h of stimulation. We also investigated whether induction of CTGF mRNA in vitro was dependent on pre-activation of alveolar macrophages by lipopolysaccharide (LPS). For this purpose, alveolar macrophages were pre-incubated with LPS (1 µg/ml) for 16 h and subsequently stimulated with Ang II (100 nM) or TGF-β1 (5 ng/ml) for another 24 h. Stimulation of LPS-activated macrophages with Ang II or TGF-β1 did not affect CTGF mRNA expression levels (data not shown).


Figure 7
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  		Fig. 7 Quantitative real-time RT-PCR analysis of CTGF mRNA expression in rat alveolar macrophages. The cells were adapted to serum-free medium and subsequently stimulated for 24 h in the absence or presence of Ang II (100 nmol/l) (A), TGF-β1 (5 ng/ml) (B), or co-culture with platelets (1x107 platelets/60 mm dish) (C). Data are presented as ratios of CTGF mRNA levels relative to the levels of 18S rRNA. Values are mean ± SEM of n=4 independent experiments. *P<0.005 vs. control.

 
3.9 Effect of co-incubation of platelets and alveolar macrophages on CTGF mRNA expression in alveolar macrophages
To provide evidence of platelet-stimulated induction of CTGF mRNA in rat alveolar macrophages, alveolar macrophages obtained by bronchoalveolar lavage were co-cultured with platelets as described in Methods. As shown in Fig. 7C, CTGF mRNA levels were significantly elevated upon co-culture of alveolar macrophages with platelets (4-fold increase compared to control, i.e. alveolar macrophages alone; P<0.005). No detectable CTGF mRNA has been observed in platelet extracts [21].


   4. Discussion
Top
 Abstract
 1. Introduction
 2. Methods
 3. Results
 4. Discussion
Acknowledgments
 References
 
Pulmonary arterial remodeling with medial hypertrophy and fibrosis is a well recognized patho-anatomic finding in pulmonary arterial hypertension of various etiologies, including HF [22,23]. However, pulmonary parenchymal remodeling in HF has also been observed, including thickening of alveolar septa with proliferation of myofibroblasts and increased deposition of extracellular matrix proteins. The current study has uncovered the novel findings that pulmonary fibrosis and medial hypertrophy of pulmonary arteries in rats with experimentally induced HF are associated with increased pulmonary levels of CTGF. Increased levels of CTGF were detected in alveolar macrophages at sites of alveolar septal thickening and deposition of collagen. Furthermore, increased levels of CTGF immunoreactivity were detected in fibroblasts/myofibroblasts in arteries displaying medial hypertrophy.

In several disease states in which fibrosis is considered to be a key pathological process, CTGF plasma or tissue levels appear to correlate with the severity of disease [24]. Previous investigations have demonstrated increased levels of pulmonary CTGF in several models of fibrotic lung disease. Specifically, increased expression of pulmonary CTGF has been reported in bleomycin-induced pulmonary fibrosis, idiopathic pulmonary fibrosis, and pulmonary sarcoidosis [14,25,12]. Thus, increased pulmonary CTGF in HF, as observed in the present study from acute to chronic HF 8 weeks post-MI, strongly suggests that CTGF is involved in the structural alterations of pulmonary tissue secondary to HF. Evidence of alveolar macrophages as a major source of pulmonary CTGF further corroborates this notion. Alveolar macrophages have previously been demonstrated to mediate profibrotic stimuli in various pulmonary disorders. For example, in bleomycin-induced pulmonary fibrosis, the alveolar macrophages were found to be an important source of profibrotic cytokines [26]. Furthermore, alveolar macrophages obtained by bronchoalveolar lavage after bleomycin treatment spontaneously release fibroblast mitogens [27]. In addition, alveolar macrophages from patients with idiopathic pulmonary fibrosis (IPF) were shown to secrete fibroblast regulatory peptides [28]. Indeed, CTGF may represent such a fibroblast mitogen. CTGF has been shown to stimulate fibroblast proliferation and synthesis and secretion of collagen type I and fibronectin in vitro [13,29]. Recent evidence from adenovirus-mediated overexpression of CTGF in rat lungs in vivo also indicates that CTGF may elicit pulmonary fibrosis [30]. Congruently, accumulation of pulmonary macrophages demonstrating robust immunoreactivity to CTGF in chronic HF, as demonstrated in the present study, suggests that influx of pulmonary macrophages may be a key factor of increased CTGF production and pulmonary fibrosis in chronic HF. The key role of pulmonary macrophages is also supported by induction of ET-1 mRNA, an important proinflammatory cytokine. Another finding of the current report also points to the profibrotic role of CTGF during pulmonary remodeling in HF. CTGF immunoreactivity in the media of pulmonary arteries in HF rats correlated with increased levels of medial collagen. Thus, increased levels of CTGF appear to be a consistent finding at sites of fibrosis. Yet, the strong association of pulmonary arterial and parenchymal CTGF expression and fibrosis demonstrated in the present study, does not prove that CTGF causes pulmonary remodeling in HF. Causative role for CTGF in pulmonary fibrosis in HF will best be addressed in models that abrogate induction of CTGF in HF.

The mechanisms of induction of pulmonary CTGF in HF are unknown. Neurohormones, autocrine/paracrine factors, or shear stress due to pulmonary hypertension may all be envisaged as potential inducers of pulmonary CTGF expression in HF. Several reports have shown a strong correlation between TGF-β1, a key cytokine involved in fibrosis, and CTGF in various fibrotic disorders [31,32,7]. Although, pulmonary TGF-β1 mRNA levels were increased in the current study, TGF-β1 mRNA levels in alveolar macrophages were not found to be altered in HF rats as compared with sham rats. Thus, to the extent that pulmonary TGF-β1 is involved in synthesis and secretion of CTGF from alveolar macrophages, a paracrine mechanism would have to be implicated. Nevertheless, TGF-β1 did not elicit increased CTGF mRNA levels in primary cultures of alveolar macrophages obtained by bronchoalveolar lavage. Furthermore, as plasma Ang II levels are increased in the current model of HF [33,34], circulating Ang II may be another factor to be considered in induction of pulmonary CTGF. In recent studies, it has been demonstrated that infusion of Ang II in rats induces expression of CTGF in aorta and kidney [35,36]. Furthermore, in a previous study from our laboratory both Ang II and TGF-β1 were found to elicit induction of CTGF in primary isolates of adult rat cardiac fibroblasts [19]. On the contrary, neither Ang II nor TGF-β1 caused induction of CTGF mRNA expression in isolated alveolar macrophages from rats as reported in the current study. Thus, alveolar macrophages from rats do not appear to be target cells of Ang II or TGF-β1 under physiological conditions. However, alveolar macrophages from rats with chronic HF may become activated or altered in ways that may confer sensitivity to Ang II or TGF-β1. Although the latter scenario remains to be investigated, activation of alveolar macrophages by pre-incubation with LPS in vitro did not confer sensitivity to Ang II or TGF-β. Conceivably, other neurohormones or cytokines may stimulate pulmonary expression and release of CTGF in HF. In this respect, Rishikof and colleagues recently reported interleukin-4-mediated induction of CTGF mRNA expression human lung fibroblasts [37]. The current study points to another paracrine mechanism, i.e. induction of CTGF mRNA in alveolar macrophages by a platelet-derived factor. Although, this study does not resolve which platelet factor or factors mediate induction of CTGF mRNA expression in alveolar macrophages, recent reports have shown that platelets contain large amounts of CTGF [38,39]. The current study indicates that CTGF expression by alveolar macrophages is induced by direct contact with platelets. Conceivably, platelet-derived CTGF may be such a factor mediating induction of CTGF expression and other proinflammatory players in alveolar macrophages. Although the relevance of these findings to human HF still remains to be shown, thickening of alveolar septa with proliferation of myofibroblasts has been reported in autopsy findings of patients with pulmonary arterial hypertension and HF, indicating that pulmonary parenchymal remodeling also occurs in human HF [40]. In conclusion, the present study demonstrates induction of CTGF at sites of pulmonary fibrosis and remodeling of the pulmonary vasculature in chronic HF rats. The present results indicate that activated alveolar macrophages are a major source of increased levels of pulmonary CTGF in HF. In addition, increased levels of CTGF in the medial layer of arteries displaying medial fibrosis suggest that CTGF may be involved in remodeling of pulmonary arteries in HF. Further studies are needed to elucidate the prevailing mechanism of induction of pulmonary CTGF in HF and whether inhibition of CTGF signalling may offer a therapeutic strategy to halt pulmonary remodeling during HF.


   Acknowledgments
Top
 Abstract
 1. Introduction
 2. Methods
 3. Results
 4. Discussion
 Acknowledgments
References
 
This study was supported by grants from the Norwegian Research Council, The Rikshospitalet-Radiumhospitalet Research Fund, and the Norwegian Council on Cardiovascular Research.


   Notes
 
Time for primary review 28 days


   References
Top
 Abstract
 1. Introduction
 2. Methods
 3. Results
 4. Discussion
 Acknowledgments
 References
 

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