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Am J Physiol Lung Cell Mol Physiol 290: L153-L161, 2006. First published August 12, 2005; doi:10.1152/ajplung.00287.2005
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Connective tissue growth factor and vascular endothelial growth factor from airway smooth muscle interact with the extracellular matrix

Janette K. Burgess,1,2 Qi Ge,1 Maree H. Poniris,1 Sarah Boustany,2 Stephen M. Twigg,3,4 Judith L. Black,1,2 and Peter R. A. Johnson1,2

1Respiratory Research Group, Department of Pharmacology, University of Sydney; 2Woolcock Institute of Medical Research, 3Discipline of Medicine, University of Sydney; and 4Department of Endocrinology, Royal Prince Alfred Hospital, Sydney, Australia

Submitted 4 July 2005 ; accepted in final form 4 August 2005


   ABSTRACT
TOP
 ABSTRACT
METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Airway remodeling describes the structural changes that occur in the asthmatic airway that include airway smooth muscle hyperplasia, increases in vascularity due to angiogenesis, and thickening of the basement membrane. Our aim in this study was to examine the effect of transforming growth factor-{beta} on the release of connective tissue growth factor and vascular endothelial growth factor from human airway smooth muscle cells derived from asthmatic and nonasthmatic patients. In addition we studied the immunohistochemical localization of these cytokines in the extracellular matrix after stimulating bronchial rings with transforming growth factor-{beta}. Connective tissue growth factor and vascular endothelial growth factor were released from both cell types and colocalized in the surrounding extracellular matrix. Prostaglandin E2 inhibited the increase in connective tissue growth factor mRNA but augmented the release of vascular endothelial growth factor. Matrix metalloproteinase-2 decreased the amount of connective tissue growth factor and vascular endothelial growth factor, but not fibronectin deposited in the extracellular matrix. This report provides the first evidence that connective tissue growth factor may anchor vascular endothelial growth factor to the extracellular matrix and that this deposition is decreased by matrix metalloproteinase-2 and prostaglandin E2. This relationship has the potential to contribute to the changes that constitute airway remodeling, therefore providing a novel focus for therapeutic intervention in asthma.

transforming growth factor-{beta}; airway remodeling; matrix metalloproteinase; prostaglandin E2

ASTHMA IS A COMPLEX DISEASE characterized by airway inflammation and airway hyperresponsiveness. Chronic inflammation leads to structural changes within the airways including an increase in the mass of smooth muscle, an increase in the number of blood vessels (angiogenesis) (23), and increased deposition of extracellular matrix (ECM) proteins (collectively referred to as airway remodeling). The airway smooth muscle (ASM) cell was originally thought to have a role only in the contractile response of the airways but is now recognized to actively contribute to airway remodeling. The mechanisms underlying airway remodeling are poorly understood; however, recent studies suggest a role for cytokines such as transforming growth factor (TGF)-{beta}, connective tissue growth factor (CTGF), and VEGF (11–13, 24).

Concentrations of TGF-{beta} are higher in the bronchial lavage fluid of asthmatic subjects (29), TGF-{beta} gene expression is increased in bronchial tissue (26, 27, 40), and increased immunoreactivity for TGF-{beta} has been reported in bronchial biopsies and submucosal eosinophils from asthmatic subjects (26, 40). TGF-{beta} also stimulates release of CTGF from ASM cells (5, 44), and the amount released from cells derived from asthmatic individuals is significantly greater (5). CTGF has intrinsic angiogenic and fibrogenic properties (33, 34) and can also bind to angiogenic factors such as VEGF and thereby modulate their activity (10, 15).

VEGF is a key regulator of endothelial cell growth and may thus play an important role in the angiogenic component of airway remodeling. Levels of VEGF in lavage fluid and induced sputum are higher in asthmatics (1, 22), and VEGF gene expression is increased in bronchial tissue (12). Blood vessels are more numerous and comprise a greater area of bronchial biopsies taken from asthmatic subjects. As is the case for CTGF and TGF-{beta}, VEGF is released from a variety of airway cells including ASM cells (19, 20, 42).

An intimate relationship exists between the ASM and the ECM. We have previously reported that abnormalities exist in the in vitro properties of the asthma-derived ASM cells in terms of proliferation rate (18), release of endogenous mediators (5, 7), and the profile of matrix proteins released (16, 17). Whether differences exist in the levels of VEGF released by the muscle is not known.

Little is known of the modulation of airway remodeling either by therapeutic intervention, such as the use of long-term corticosteroids, or by endogenous factors. However, it is known that PGE2 regulates TGF-{beta}-induced CTGF and collagen I in fibroblasts (30) and that matrix metalloproteinases (MMPs) can cleave CTGF and activate the angiogenic activity of VEGF (10). Human ASM cells express MMP2 and release pro-MMP2 (9). The role of PGE2 and MMPs in the modulation of fibrogenic and angiogenic factors such as CTGF and VEGF and their relationship to the ASM cell and its surrounding ECM are yet to be examined.

In this study we examined the interaction of released CTGF and VEGF with the ECM from ASM cells derived from both asthmatic and nonasthmatic subjects. The induction of VEGF following stimulation with TGF-{beta} and recombinant CTGF was studied, and the role of PGE2 and MMP2 in the regulation of these events was also elucidated. We hypothesized that in ASM an intimate relationship exists between CTGF and VEGF that is modulated by MMP2 and PGE2 and that has the potential to contribute to the changes which constitute airway remodeling.


   METHODS
TOP
 ABSTRACT
 METHODS
RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Chemicals. DMEM, insulin transferrin selenium (ITS), Dulbecco’s PBS, penicillin, streptomycin, amphotericin B, trypan blue (Life Technologies, Heidelberg, Australia); FBS (Commonwealth Serum Laboratories, Melbourne, Australia), recombinant human TGF-{beta}, PGE2, EDTA, benzamidine, PMSF, and aprotinin (Sigma, St. Louis, MO) were obtained from the sources given in parentheses. Recombinant human CTGF was produced in and purified from an adenoviral expression system using 911 cells, as previously described (25). Recombinant human VEGF was obtained from R&D Systems (Minneapolis, MN).

ASM. ASM was obtained from human bronchial airways by methods previously described (5, 18, 31). The details of the individuals from whom airways were obtained are shown in Table 1. Approval for all experiments with human lung was provided by the Human Ethics Committee of the University of Sydney and the Central Sydney Area Health Service, and all patients provided written informed consent. ASM cell characteristics were confirmed by immunofluorescence and light microscopy (8).


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  		Table 1. Patient details

 
Stimulation with growth factors. ASM cells were seeded into six-well plates at 1 x 104 cells/cm2 in 5% FBS DMEM for 24 h before being synchronized for 24 h in 0.1% ITS in DMEM. Cells were stimulated with TGF-{beta} (1 ng/ml) in the presence or absence of PGE2 (10 µM), recombinant human CTGF (250 ng/ml), or VEGF (50 ng/ml) for 8 or 24 h. Control cells were maintained in 0.1% ITS in DMEM. Cells were lysed and then stored at –20°C until extraction. Total RNA was extracted from ASM cells using the NucleoSpin RNA II (Macherey-Nagel, Duren, Germany) according to the manufacturer’s instructions. After extraction, samples were eluted in 30 µl of RNase-free water and stored at –20°C until use.

ELISA. For measurement of release of VEGF165 [a splice variant of VEGF-A, the most potent angiogenic factor in its family, and the best-characterized isoform in ASM cells (3, 19, 20, 35, 42)] from the cells, supernatants were collected from the six-well plates described above. The levels of VEGF165 were measured with commercial ELISA kits according to the manufacturer’s instructions (R&D Systems, Minneapolis MN).

Immunoprecipitation. ASM cells were seeded into 175-cm2 flasks at 1 x 104 cells/cm2 in 5% FBS DMEM for 24 h before being synchronized for 24 h in 0.1% ITS in DMEM and then stimulated with TGF-{beta} (1 ng/ml) for 24 h. The supernatants were collected and stored at –20°C until analysis.

The supernatants were defrosted and spun at 500 g at 4°C for 5 min before the addition of protease inhibitors (5 mM EDTA, 1 mM benzamidine, 0.5 mM PMSF, and 10 µm aprotinin). Biotin 3-sulfo-N-hydroxysuccinimide ester sodium salt (0.4 mM, Sigma) was incubated with the supernatants for 30 min at room temperature before the addition of 0.8 mM glycine. The samples were precleared with protein G agarose (Roche Diagnostics, Basel, Switzerland) for 30 min at room temperature before centrifugation at 13,000 g at 4°C for 3 min. Anti-CTGF (2.5 µg/ml CTGF-ab6992; Abcam, Cambridge, UK) or anti-VEGF (2 µg/ml goat anti-human VEGF, R&D Systems) were added to the samples and incubated for 1 h at 4°C. Fresh protein G agarose was then added and incubated for 1.5 h at 4°C before centrifugation at 13,000 g at 4°C for 2 min. The precipitated agarose was washed three times with cold immunoprecipitation buffer (20 µM Tris, 5 mM EDTA, 1 mM benzamidine, 0.5 mM PMSF, and 10 µm aprotinin) before resuspension in 20 µl of SDS loading buffer [0.0625 M Tris·HCl, pH 6.8, 2% (wt/vol) SDS, 0.1 M DTT, 10% (vol/vol) glycerol, and 0.01% (wt/vol) bromphenol blue]. Cellular proteins were size fractionated on 10% PAGE, transferred to polyvinylidene difluoride membranes, and blocked overnight in 5% (wt/vol) skim milk solution as described previously (6). The membranes were incubated with horseradish peroxidase-conjugated streptavidin (1 in 1,000 in skim milk solution; Amersham Biosciences, Buckinghamshire, UK) for 1 h before being washed. Immunoblot detection was performed with a Supersignal West Dura extended-duration substrate kit (Pierce, Rockford, IL), and bands were analyzed with a 440F Kodak imaging system and software.

Real-time reverse transcription PCR. Real-time reverse transcription PCR for CTGF and real-time PCR for collagen I and fibronectin were performed as described previously (5). Data from the reactions were collected and analyzed by the complementary computer software.

Immunohistochemistry. Human ASM cells were grown for 24 h in 5% FBS DMEM on glass coverslips. Medium was then changed to 0.1% ITS DMEM for 24 h before stimulation with or without TGF-{beta} (1 ng/ml) for a further 24 h. Human lung tissue was obtained from nonasthmatic lung specimens resected for carcinoma or transplantation. Bronchial rings (2–5 mm diameter and 3 mm in length) were dissected free from surrounding parenchymal tissue. Corresponding rings from asthmatic tissue were not available for comparison in these experiments. The bronchial rings from the same patient were incubated for 24 h in a 5% CO2 incubator at 37°C in 0.1% ITS DMEM with TGF-{beta} (1 ng/ml) in the presence or absence of PGE2 (10 µM). Control rings were maintained in 0.1% ITS in DMEM. Tissues were frozen in optimal cutting temperature compound embedding medium (Fronine Laboratory Supplies, Riverstone, Australia) and cut with a cryostat at 7-µm thickness. Sequential sections were rehydrated in water for 5 min and washed with PBS for 2 min before staining. Primary antibodies for CTGF (4 µg/ml rabbit anti-human CTGF, Torrey Pines Biolabs) and VEGF (10 µg/ml, mouse anti-human VEGF, R&D Systems) were added to the cells or sections simultaneously, and those for fibronectin (1 µg/ml mouse antifibronectin, Chemicon) or collagen I (1 µg/ml mouse anti-collagen I, Chemicon) were added to sequential sections and incubated for 1 h at room temperature. Cells or sections were washed twice with PBS and secondary antibodies added for 30 min at room temperature (3 µg/ml donkey anti-rabbit TRITC, Jackson Immunoresearch; and 1 µg/ml goat anti-mouse FITC, BD Biosciences). After two further washes with PBS, cells or sections were mounted with Vectashield mounting medium (Vector Laboratories). Hematoxylin and eosin-stained sequential sections were used to identify airway morphology.

Release of CTGF and VEGF from the ECM by MMP2. ASM cells were grown in eight-well chamber slides (Lab-Tek, Nalge Nunc International) for 24 h and then quiesced in 0.1% ITS for a further 24 h. The cells were then stimulated with either TGF-{beta} (1 ng/ml), TGF-{beta} (1 ng/ml) + active recombinant MMP2 (100 ng/ml), or MMP2 (100 ng/ml) alone for 24 h before fixation in 4% paraformaldehyde.

Cells were stained for CTGF, VEGF, or fibronectin (as above). The Envision staining system (Dakocytomation), coupled with chromogen 3,3'-diaminobenzidine, was used to visualize positive staining. Cells were counterstained with hematoxylin to allow visualization of the nuclei. Rabbit IgG (4 µg/ml, Jackson Immunoresearch) and mouse IgG1 and IgG2B (5 µg/ml, R&D Systems) isotype controls were used.

Image analysis was carried out with QWIN image analysis software (Leica Microsystems, Wetzlar, Germany). Images were transformed to gray scale, and the gray intensity levels were measured per cell. Twenty cells were measured per treatment per patient, and the mean and SE of the gray level for each treatment were calculated.

Statistical analysis. In real-time PCR experiments the number of cycles needed to attain a threshold fluorescence set to lie on the exponential part of the amplification plot was determined. Results for CTGF, collagen, or fibronectin were normalized against those obtained for 18S and expressed as a fold increase. Results from triplicate wells from each individual patient were meaned, and an overall mean and SE were calculated for cycle number. ANOVA using repeated measures and the Fisher paired least significant difference posttest was performed on the results for real-time RT-PCR and mean densitometric values for immunoprecipitation blots. Factorial ANOVA and paired Student’s t-test, where appropriate, were used to compare VEGF, CTGF, collagen, and fibronectin mRNA or protein production from asthmatic and nonasthmatic cells. In all cases a P value of < 0.05 was considered significant.


   RESULTS
TOP
 ABSTRACT
 METHODS
 RESULTS
DISCUSSION
 GRANTS
 REFERENCES
 
Growth factor induction of VEGF. ASM cells, stimulated with TGF-{beta}, released significant levels of soluble VEGF165 after 8 h in asthmatic cells (data not shown) and 24 h in both cell types (P < 0.004 compared with 0.1% ITS, asthmatic n = 8, nonasthmatic n = 9) (Fig. 1A), in agreement with previous reports (19, 42). Stimulation of the ASM cells with recombinant human CTGF also induced significant release of VEGF165, after 24 h (P < 0.0004, asthmatic n = 5, nonasthmatic n = 5) (Fig. 1A). There was no difference in the level of VEGF165 released by asthmatic and nonasthmatic cells with either stimulus.



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  		Fig. 1. Release of VEGF165 from airway smooth muscle (ASM) cells. Asthmatic and nonasthmatic ASM cells were stimulated with either transforming growth factor (TGF)-{beta} (1 ng/ml) or recombinant human connective tissue growth factor (CTGF, 250 ng/ml; A) or TGF-{beta} (1 ng/ml), TGF-{beta} + PGE2 (10, µM) or PGE2 alone (B) for 24 h, and the supernatants were collected. Control cells were maintained in 0.1% insulin transferrin selenium (ITS) in DMEM. The release of VEGF165 was measured by ELISA. Expression significantly different to 0.1% ITS, *P < 0.02. Expression significantly different to TGF-{beta}, #P < 0.003. Asthmatic cells stimulated with TGF-{beta} n = 8 and CTGF n = 5. Nonasthmatic cells stimulated with TGF-{beta} n = 9 and CTGF n = 5. Asthmatic and nonasthmatic cells stimulated with TGF-{beta} + PGE2 n = 7 and PGE2 n = 3.

 
PGE2 augmented TGF-{beta}-induced VEGF release after 24 h in asthmatic and nonasthmatic ASM cells (P < 0.003, asthmatic n = 7, nonasthmatic n = 7) (Fig. 1B). Once again there was no difference in the level of release between the two cell types.

CTGF and VEGF coimmunoprecipitate. Immunoprecipitation of lysates from cells that had been stimulated with TGF-{beta} resulted in the precipitation of both CTGF (Fig. 2A) and VEGF (Fig. 2B) proteins when antibodies specific for CTGF or VEGF165 were used as the capture antibody. Both antibodies precipitated significantly greater amounts of specific proteins than the Sepharose beads alone.



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  		Fig. 2. Coimmunoprecipitation of CTGF and VEGF. Proteins released from ASM cells following TGF-{beta} stimulation were labeled with biotin before immunoprecipitation with either anti-CTGF or anti-VEGF165 antibodies and protein G agarose. The precipitated proteins were size fractionated on SDS-PAGE, and the bands were detected by chemiluminescence. The precipitation of CTGF protein (A) and VEGF protein (B) with each of the antibodies was measured in densitometric units. The immunoprecipitation blots are representative of the results seen with 4 asthmatic cell lines, and the graphs illustrate means ± SE for n = 4 asthmatics. Immunoprecipitation significantly greater than beads alone *P < 0.04.

 
CTGF and VEGF colocalize. Using immunohistochemistry, in ASM cells grown in culture (Fig. 3A) and tissue sections from bronchial rings (Fig. 3B) that had both been stimulated with TGF-{beta}, we showed that CTGF and VEGF165 were upregulated. Dual staining of the same cells demonstrated that the CTGF and the VEGF colocalized around a subpopulation of the ASM cells. The CTGF and VEGF appear to be localized in the ECM surrounding the cells and the ASM bundles in the tissue sections, although the resolution of our images was not of a high enough magnitude to elucidate the exact location of these proteins. PGE2 decreased the TGF-{beta}-induced CTGF and VEGF165 expression in bronchial segments (Fig. 3C).



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  		Fig. 3. Colocalization of CTGF and VEGF and modulation by TGF-{beta} and PGE2 in ASM cells and tissue sections. ASM cells (A) were incubated in 0.1% ITS DMEM or 0.1% ITS DMEM + TGF-{beta} (1 ng/ml) or human bronchial rings from the same patient or were incubated in 0.1% ITS DMEM or 0.1% ITS DMEM + TGF-{beta} (1 ng/ml) (B) or 0.1% ITS DMEM, 0.1% ITS DMEM + TGF-{beta} (1 ng/ml), 0.1% ITS DMEM + TGF-{beta} + PGE2 (10 µM) or 0.1% ITS DMEM + PGE2 (C) for 24 h. Cells and tissue sections were simultaneously stained with rabbit anti-CTGF coupled with donkey anti-rabbit TRITC (red staining) and mouse anti-human VEGF coupled with goat anti-mouse FITC (green staining). The images were merged using imaging software, and regions of colocalization were identified (yellow staining). Controls are representative of the staining seen with both isotype control antibodies. The hematoxylin and eosin-stained sequential sections were used to identify airway morphology. The results are representative of tissue from 3 nonasthmatic patients.

 
VEGF does not induce CTGF, collagen, or fibronectin. Stimulation of ASM cells with VEGF165 for 24 h did not induce any CTGF, collagen I, or fibronectin mRNA expression; it did, however, induce proliferation of human umbilical vein endothelial cells (data not shown). In addition, TGF-{beta} or VEGF did not increase the release of soluble fibronectin above the level released in the presence of 0.1% ITS alone (data not shown).

Effect of PGE2 on TGF-{beta}-mediated CTGF, collagen, and fibronectin release. PGE2 significantly attenuated the TGF-{beta}-induced CTGF mRNA (P ≤ 0.003) after 8 h in the asthmatic ASM (n = 5) (Fig. 4A) and after 24 h in the nonasthmatic ASM (n = 6) (Fig. 4B). Similarly, PGE2 significantly attenuated the TGF-{beta} induction of collagen I mRNA (P ≤ 0.002, asthmatic n = 5, nonasthmatic n = 6) and fibronectin mRNA (P ≤ 0.001, asthmatic n = 5, nonasthmatic n = 6) after 24 h (data not shown). PGE2 also significantly inhibited TGF-{beta}-induced collagen I and fibronectin protein deposition by asthmatic ASM cells (P ≤ 0.02, A n = 5) (Fig. 4, C and D).



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  		Fig. 4. PGE2 attenuates TGF-{beta}-induced CTGF, collagen, and fibronectin. ASM cells were stimulated with TGF-{beta} (1 ng/ml) in the presence or absence of PGE2 (10 µM) for up to 24 h. Control cells were maintained in 0.1% ITS in DMEM. The results are expressed as a percentage of the expression seen in the presence of TGF-{beta} alone in asthmatic (n = 4) and nonasthmatic (n = 4) cells. CTGF expression was assessed at 8 (A) and 24 h (B) by real-time PCR, and the cycle thresholds were normalized to the expression of the control (18S rRNA) gene in each sample. Collagen I (C) and fibronectin (D) protein expression was assessed at 24 h by extracellular matrix (ECM) ELISAs. Expression significantly different from TGF-{beta} #P < 0.05.

 
Moreover PGE2 decreased TGF-{beta}-induced collagen I and fibronectin (Fig. 5) in tissue sections from bronchial rings.



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  		Fig. 5. PGE2 attenuates TGF-{beta}-induced collagen and fibronectin expression in tissue sections. Human bronchial rings from the same patient were incubated in 0.1% ITS DMEM, 0.1% ITS DMEM + TGF-{beta} (1 ng/ml), 0.1% ITS DMEM + TGF-{beta} + PGE2 (10 µM), or 0.1% ITS DMEM + PGE2 for 24 h. Sequential tissue sections were stained with mouse anti-collagen I or mouse antifibronectin primary antibodies and goat anti-mouse FITC (green staining). The hematoxylin and eosin-stained sequential sections were used to identify airway morphology. The results are representative of tissue from 3 nonasthmatic patients.

 
Effect of MMP2 on TGF-{beta}-mediated CTGF and VEGF binding to cells. The addition of active recombinant MMP2 (100 ng/ml) to TGF-{beta}-stimulated ASM cells in culture significantly attenuated the binding of both CTGF and VEGF165 to the cell layer in asthmatic cells and CTGF in nonasthmatic cells (Fig. 6, Table 2). In contrast, the increased deposition of fibronectin around the ASM cells, induced by TGF-{beta}, was not altered in either cell type (Fig. 6, Table 2). Active MMP2 alone had no effect on the amount of CTGF or VEGF165 bound to the cells in the presence of 0.1% ITS alone.



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  		Fig. 6. Matrix metalloproteinase (MMP)-2 dissociates CTGF-VEGF complex from ECM. ASM cells were incubated in 0.1% ITS DMEM, 0.1% ITS DMEM + TGF-{beta} (1 ng/ml), 0.1% ITS DMEM + TGF-{beta} + active recombinant MMP2 (100 ng/ml), or 0.1% ITS DMEM + active recombinant MMP2 for 24 h. Cells were stained for CTGF, VEGF, or fibronectin using the Envision staining system coupled with chromogen 3,3'-diaminobenzidine (brown staining). Cells were counterstained with hematoxylin to allow visualization of the nuclei. The images are representative of the results seen with n = 3 asthmatic and n = 3 nonasthmatic cell lines.

 

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  		Table 2. Effect of MMP2 on TGF-{beta}-mediated CTGF, VEGF, and fibronectin binding to cells

 

   DISCUSSION
TOP
 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
GRANTS
 REFERENCES
 
In this study we have shown for the first time that CTGF and VEGF released from ASM cells bind and colocalize in the ECM. The presence of CTGF is required for the VEGF to be localized to the ECM. TGF-{beta} increases the expression of both growth factors, but PGE2 has divergent effects, inhibiting the TGF-{beta}-induced CTGF and enhancing the TGF-{beta}-induced VEGF. Our results were obtained from not only cultured ASM cells but also sections of bronchial tissue, and this was important for two reasons. First, it established that what we observed were not merely culture artifacts, and second, it enabled us to observe events occurring beyond the confines of a single cell.

CTGF and VEGF are released from ASM cells following TGF-{beta} stimulation. This is the first study, to our knowledge, to examine the interaction of CTGF and VEGF released from primary cells in culture and to demonstrate the colocalization of CTGF and VEGF in airway sections following TGF-{beta} stimulation. It was not possible to determine the exact cellular or subcellular localization of these growth factors from our study. Our results are consistent with those of Inoki and colleagues (15), who previously demonstrated that recombinant CTGF and VEGF proteins can bind. We also found that CTGF was able to induce the release of VEGF from ASM cells, indicating that CTGF may play an autocrine role in the regulation of VEGF release.

Our finding that TGF-{beta} induced the release of VEGF from asthmatic and nonasthmatic ASM cells is consistent with previous studies examining release of VEGF from nonasthmatic ASM cells (19, 42). Although we did not find a difference in the level of VEGF released from asthmatic and nonasthmatic ASM cells, since Hoshino et al. (12) have shown that there is increased immunoreactivity to VEGF in sections/biopsies of asthmatic airways, this is likely to be the result of the increased TGF-{beta} present in the airways of asthmatic patients. In this study our nonasthmatic ASM cells and tissue were derived from patients with a variety of disease states, but no clear relationship emerged between the disease states of these patients and the results observed.

Stimulation of ASM cells with VEGF did not induce CTGF, collagen I, or fibronectin message or the release of soluble fibronectin in our study. This finding is in contrast to that by Kazi and colleagues (19), who reported that VEGF induced the release of soluble fibronectin from ASM cells. The difference between the findings of the two studies may be related to the composition of the cell culture medium. In our study we used 0.1% ITS rather than 0.1% BSA. Insulin is known to induce collagen and fibronectin in smooth muscle cells (14, 32, 38). The levels of soluble fibronectin released from ASM cells in 0.1% ITS in our study were similar to the levels induced by VEGF in the previous study. It is possible that we did not see any further induction of soluble fibronectin as the levels induced by insulin alone were maximal. Suzuma and colleagues (37) reported that VEGF induced CTGF mRNA in bovine retinal endothelial cells. In contrast, however, and consistent with our data, Wunderlich et al. (43) could not show CTGF induction by VEGF. Cell type and species differences may account for these disparate findings.

PGE2 inhibited TGF-{beta}-induced CTGF, collagen I, and fibronectin mRNA and protein in the ASM cells in culture and in the airway tissue sections in our study. In agreement with this finding, PGE2 attenuated TGF-{beta}-induced CTGF mRNA transcription in a human lung fibroblast cell line (30) and iloprost (which also acts by elevation of cAMP) blocked TGF-{beta}-induced CTGF and collagen in human skin fibroblasts (36). However, the effect of PGE2 on TGF-{beta}-induced VEGF from the ASM cells in our experiments was different. The VEGF was increased in the cell supernatants but decreased in the ECM surrounding the ASM cells in culture and in the airway tissue sections. As we have previously reported that the PGE2 released by asthmatic ASM cells is decreased (7), we may have expected the release of VEGF from asthmatic ASM cells to be reduced compared with nonasthmatic cells. Bradbury et al. (3) recently reported that PGE2-induced release of VEGF from ASM cells is mediated via the E-prostanoid (EP) 2 and EP4 receptors. The level of VEGF released from the asthmatic ASM cells may be regulated by the increased number of EP2 receptors on the asthmatic ASM cells (4) that may compensate for the decreased PGE2 release. This would explain the fact that no differences were observed between the asthmatic and nonasthmatic cells. Others have reported that PGE2 can increase VEGF release. Stocks and colleagues (35) recently observed that the induction of VEGF165 from ASM cells by TGF-{beta} and PGE2 is mediated via separate pathways. Kazi and colleagues (19) found that PDGF-induced VEGF release in cell supernatants was increased by PGE2 in ASM cells, which raises the possibility of stimulus specific effects.

We found that PGE2 increased TGF-{beta}-induced soluble VEGF in cells in culture but reduced matrix-associated VEGF protein following treatment of bronchial rings. Since PGE2 downregulates CTGF expression, then if CTGF was required to colocalize VEGF in cell-associated sites and in tissue ECM this could explain the PGE2-induced changes in VEGF. Our findings indicate that VEGF depends on the presence of CTGF to be anchored to the ECM. This supports the proposal that ctgf/cyr61/nov family proteins such as CTGF act primarily as matrix-associated proteins that provide a bridge between structural proteins in the ECM and effector molecules such as growth factors recruited to the matrix (2, 21). We have previously reported that asthmatic ASM cells release significantly more CTGF in response to TGF-{beta} than nonasthmatic cells (5). Asthmatic ASM cells release less PGE2 (7), which may result in less downregulation of the CTGF induced by TGF-{beta} in these cells. Increased local concentrations of CTGF would lead to greater amounts of VEGF being anchored in the ECM in the vicinity of the ASM in the asthmatic airway. The presence of more ECM associated VEGF could increase the angiogenic potential in this region of the airway.

CTGF may also play a role in regulating VEGF activity (10, 15) and has intrinsic angiogenic activity (33, 34). In nonpulmonary tissue in areas of strong CTGF staining, CD31-positive staining (indicative of new blood vessel formation) was observed (39). Inoki and colleagues (15) demonstrated that when CTGF interacts with VEGF165 it inhibits VEGF binding to endothelial cells and immobilized recombinant VEGF receptor (KDR/IgG Fc). This interaction resulted in the inhibition of the VEGF angiogenic activity, suggesting that this activity is negatively regulated by CTGF.

MMP2 is known to proteolyse CTGF, and we determined whether its application would also affect VEGF presence in the ECM after the induction of both VEGF and CTGF by TGF-{beta}. In our study active recombinant MMP2 decreased the binding of TGF-{beta}-induced CTGF and VEGF to the ECM surrounding the ASM cells in culture. This occurred in both asthmatic and nonasthmatic cells, and no significant differences between the cell types were observed. This degradative effect was a specific event, as the deposition of TGF-{beta}-induced fibronectin was not altered. MMP2 therefore has the capacity to regulate the release of VEGF from the ECM. Treatment of a recombinant VEGF/CTGF complex with a variety of recombinant MMPs (including MMP2) resulted in degradation of the CTGF and release of the VEGF in an active form (10). The source of endogenous MMP2 is not known, although we do know that ASM cells release pro-MMP2, which could be activated in the vicinity of the ECM. VEGF upregulates MMP expression in vascular smooth muscle cells (41), but whether this occurs in ASM cells is not known. This has the potential to provide an additional control for the angiogenic processes that could be occurring around the ASM bundles in the airways of asthmatic individuals.

CTGF has the potential also to play a role in regulating events leading to remodeling in other tissues or other disease states. As CTGF is released from other cell types [for example fibroblasts (30) and endothelial cells (43)], it has the potential also to anchor VEGF in the ECM surrounding these cells. In this way CTGF may be an important regulatory factor for the processes involved in remodeling in many tissues. Thus the implications of our findings are not restricted to airway remodeling but may also be relevant to many other circumstances where angiogenesis plays a role.

It is now becoming clear that it is of importance to think of asthma in terms of dysfunctional injury and wound healing and that there is a need to identify markers to distinguish asthmatics who will develop chronic airway remodeling. Understanding the role of angiogenesis in the pathophysiology of asthma will be a vital part of this process. In this study we have shown that TGF-{beta} induces the release of CTGF and VEGF from ASM cells and that these factors bind and colocalize in the ECM surrounding the ASM cells. The presence of CTGF is required for the VEGF to be localized to the ECM. PGE2 inhibits the TGF-{beta}-induced CTGF but enhances the TGF-{beta}-induced VEGF. MMP2 also has a role in the regulation of the formation of the CTGF/VEGF complex. It is possible that CTGF, via its interaction with VEGF, may have an important role in regulating the new blood vessel formation around the ASM bundles in the asthmatic airway. CTGF may be a novel marker of airway remodeling, and increased understanding of its function may lead to the development of new avenues for therapeutic interventions.


   GRANTS
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 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
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This work was supported by the NH&MRC, Australia. J. K. Burgess is supported by National Health and Medical Research Council (NH&MRC) Peter Doherty Fellowship 165722.


   ACKNOWLEDGMENTS
 
The authors acknowledge the collaborative effort of the cardiopulmonary transplant team and the pathologists at St. Vincent’s Hospital, Sydney, and the thoracic physicians and pathologists at Royal Prince Alfred Hospital, Concord Repatriation Hospital, Strathfield Private Hospital, and Rhodes Pathology, Sydney. We also acknowledge the contribution of Drs. Gregory King and Melissa Baraket at the Woolcock Institute of Medical Research for supplying the asthmatic biopsies and Joanne Thompson and Pablo Britos for excellent technical assistance.


   FOOTNOTES
 

Address for reprint requests and other correspondence: J. Burgess, Respiratory Research Group, Dept. of Pharmacology, Bosch Bldg., D05, Univ. of Sydney, Sydney, NSW, Australia 2006 (e-mail: janette{at}pharmacol.usyd.edu.au)

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.


   REFERENCES
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 RESULTS
 DISCUSSION
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