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Published online February 15, 2007
Diabetes 56:1421-1428, 2007
DOI: 10.2337/db06-1644
© 2007 by the American Diabetes Association
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Functional Characteristics of Connective Tissue Growth Factor on Vitreoretinal Cells

Takeshi Kita, Yasuaki Hata, Muneki Miura, Shuhei Kawahara, Shintaro Nakao, and Tatsuro Ishibashi

From the Department of Ophthalmology, Graduate School of Medical Sciences, Kyushu University, Fukuoka, Japan

Address correspondence and reprint requests to Yasuaki Hata, MD, PhD, Department of Ophthalmology, Graduate School of Medical Sciences, Kyushu University, 3-1-1 Maidashi, Higashi-Ku, Fukuoka 812-8582, Japan. E-mail: hatachan{at}med.kyushu-u.ac.jp

Abbreviations: BAEC, bovine aortic endothelial cell; BREC, bovine retinal capillary endothelial cell; BRPE, bovine retinal pigment epithelial cell; CTGF, connective tissue growth factor; DMEM, Dulbecco's modified Eagle's medium; FBS, fetal bovine serum; HUVEC, human umbilical vein endothelial cell; MAPK, mitogen-activated protein kinase; PDR, proliferative diabetic retinopathy; PVR, proliferative vitreoretinopathy; REC, retinal capillary endothelial cell; RPE, retinal pigment epithelial cell; RRD, rhegmatogenous retinal detachment; TGF-ß, transforming growth factor-ß; VEGF, vascular endothelial growth factor


   ABSTRACT
TOP
 ABSTRACT
RESEARCH DESIGN AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Connective tissue growth factor (CTGF) level is elevated in eyes with proliferative vitreoretinal diseases, such as proliferative diabetic retinopathy and proliferative vitreoretinopathy (PVR), as we previously reported, but its functional characteristics on vitreoretinal cells are yet to be clarified. In this study, we demonstrated a growth-promoting effect of CTGF on cultured hyalocytes and bovine retinal pigment epithelial cells (BRPEs) with the induction of p44/p42 mitogen-activated protein kinase phosphorylation and [3H]thymidine incorporation. CTGF also stimulated the synthesis of fibronectin by hyalocytes and BRPEs without significant effect on collagen gel contraction by these cells. On the other hand, CTGF had no direct effects on the proliferation, migration, or in vitro tube formation by vascular endothelial cells. Nevertheless, CTGF promoted vascular endothelial growth factor (VEGF) gene expression by hyalocytes and BRPEs. Although the concentrations of both CTGF and VEGF in the human vitreous samples with proliferative vitreoretinal diseases were elevated, there was no significant correlation between these concentrations. These findings indicate that CTGF appears to be involved in the formation of proliferative membranes without direct regulation of their cicatricial contraction in the pathogenesis of proliferative vitreoretinal diseases. Whereas CTGF might have no direct effects or minimal effects, if any, on retinal neovascularization, it is possible that CTGF has indirect effects by modulating the expression of VEGF.

Proliferative vitreoretinal diseases, such as proliferative vitreoretinopathy (PVR) and proliferative diabetic retinopathy (PDR), still remain common causes of severe vision loss or blindness despite the dramatic development of vitreoretinal surgery (1,2). Better understanding of the pathogenesis of proliferative vitreoretinal diseases is thus needed for improved management of the diseases.

PVR and PDR cause tractional retinal detachment due to the formation of contractile preretinal fibrous membranes as a result of excessive wound healing in the pathological conditions. The membrane formation is characterized by the proliferation and migration of the cells and the excessive synthesis and deposition of extracellular matrix proteins (3). Previous studies including our own revealed that various types of ocular cells, such as pigment epithelial cells, hyalocytes, glial cells, vascular endothelial cells, and fibroblast-like cells, appear to play a crucial role in the formation of preretinal fibrous membrane (3–5). Another important pathological complication of PDR is retinal neovascularization, which often leads to vitreous hemorrhage and causes blindness. Retinal neovascularization due to PDR is known to be associated with retinal capillary nonperfusion and to be characterized by the proliferation and migration of retinal vascular endothelial cells and the development of new vessels, frequently followed by the formation of fibrovascular membrane together with the contractile proliferative membrane (6).

The tissue repair process is regulated by a number of polypeptides including cytokines and growth factors. Connective tissue growth factor (CTGF) is a 38-kDa cysteine-rich polypeptide that was originally identified from conditioned medium of human umbilical vein endothelial cells (HUVECs) (7). CTGF, considered to be a downstream mediator of transforming growth factor-ß (TGF-ß) (8,9), is indicated to induce the production of extracellular matrix, such as collagen and fibronectin, and to cause fibrosis (10). Recent studies have shown that CTGF is overexpressed in the membranes of eyes with PVR (11) or PDR (12), suggesting that CTGF might be involved in the pathogenesis of PVR and PDR. In addition, our previous study revealed that CTGF is overexpressed also in the vitreous with PVR and PDR and additionally demonstrated that various types of vitreoretinal cells could be the sources of CTGF (13). However, little is known about the cellular effects of CTGF in the pathogenesis of proliferative vitreoretinal diseases. We hypothesized that CTGF would have some effects on vitreoretinal cells in an autocrine and/or a paracrine manner and play a role in the progression of proliferative vitreoretinal diseases, such as PVR and PDR.

Furthermore, CTGF has been recently indicated to be one of the regulators of angiogenesis. In vitro, CTGF has been demonstrated to have proangiogenic effects on HUVECs (14) and bovine aortic endothelial cells (BAECs) (15), and in vivo, CTGF has been indicated to induce angiogenesis in rat corneal pocket implants (16) and to be involved in tumor angiogenesis (17) and choroidal neovascularization (18,19). However, to the best of our knowledge, the effects of CTGF on retinal neovascularization and retinal vascular endothelial cell functions have yet to be investigated.

In the present study, we investigated cellular effects of CTGF using hyalocytes and RPEs to address the effect of CTGF on the formation of contractile proliferative membrane and also using retinal vascular endothelial cells to address the effect of CTGF on retinal neovascularization. Furthermore, we addressed the expressional interaction between CTGF and vascular endothelial growth factor (VEGF), a potent angiogenic molecule in vitro and in vivo.


   RESEARCH DESIGN AND METHODS
TOP
 ABSTRACT
 RESEARCH DESIGN AND METHODS
RESULTS
 DISCUSSION
 REFERENCES
 
Reagents.
Recombinant human CTGF was purchased from PeproTech (London), and recombinant human TGF-ß2 and VEGF were from Sigma-Aldrich (Tokyo). Rabbit polyclonal antibodies against phospho-p44/p42 mitogen-activated protein kinase (MAPK) and p44/p42 MAPK were obtained from Cell Signaling Technology (Beverly, MA), and mouse monoclonal antibody against fibronectin was from Santa Cruz Biotechnology (Santa Cruz, CA). In addition, horseradish peroxidase–conjugated secondary antibody was obtained from Bio-Rad (Hercules, CA).

Cell culture.
Bovine eyes were obtained from a local abattoir. The posterior part of the vitreous body was extracted and washed once in Dulbecco's modified Eagle's medium (DMEM; Sigma). The vitreous was chopped into several pieces and incubated in type I collagen–coated dishes filled with DMEM containing 10% fetal bovine serum (FBS; Invitrogen-Gibco, San Diego, CA) for 1 week. The cells that proliferated on the dishes were then subcultured in type I collagen–coated dishes in DMEM supplemented with 10% FBS as well. Cultured hyalocytes obtained between passages 4 to 7, which had shown no obvious morphological changes, were used in the following experiment. Isolated cells were immnocytochemically identified as hyalocytes expressing S-100 protein, but expressed neither glial fibrillary acidic protein nor cytokeratin, as previously described (20). Bovine retinal pigment epithelial cells (BRPEs) and bovine retinal capillary endothelial cells (BRECs) were also isolated from bovine eyes and cultured as previously reported (21,22).

Phosphorylation of p44/p42 MAPK.
Hyalocytes, BRPEs, and BRECs were starved with 1% calf serum and untreated or treated with CTGF (100 ng/ml) for the indicated time (2, 5, 15, 30, and 60 min) with the same media, and the cells were lysed with 1x Laemmli buffer (50 mmol/l Tris [pH 6.8], 2% SDS, and 10% glycerol) containing protease inhibitor and phosphatase inhibitors (1 mmol/l phenylmethylsulfonyl fluoride, 1 mmol/l NaF, and 0.5 mmol/l Na3VO4). The lysed protein samples were separated by SDS-PAGE, followed by electrophoretic transfer to nitrocellulose membranes (New England Biolabs, Beverly, MA). The blots were blocked with skim milk and incubated overnight at 4°C with antibody against phospho-p44/p42 MAPK (1:1,000). Washed with Tris-buffered saline with Tween (20 mmol/l Tris [pH 7.5], 500 mmol/l NaCl, and 0.1% Tween-20) three times for 10 min each, the membranes had been incubated with horseradish peroxidase–labeled goat anti-rabbit IgG (Bio-Rad) (1:4,000) for 30 min at room temperature. Visualization was performed with an enhanced chemiluminescence detection system (Amersham, Arlington Heights, IL) according to the manufacturer's protocol. Lane-loading differences were normalized by reblotting the membranes with antibodies against p44/p42 MAPK (1:1,000).

[3H]thymidine incorporation.
The hyalocytes, BRPEs, and BRECs were respectively seeded into 24-well plates at a density of 1 x 104 cells/well. The media were replaced by DMEM with 1% calf serum the next day. After 24 h, the cells were unstimulated or stimulated with CTGF (1, 10, and 100 ng/ml) for 18 h with the same media. [3H]thymidine was then added (0.25 µCi/well) for an additional 6 h, and after that, the cells were washed, fixed, and lysed. Incorporated [3H]thymidine was determined by scintillation counting, as previously described (23).

Collagen gel contraction assay.
The contraction assay was performed as we previously described (24). Hyalocytes and BRPEs were collected by the treatment of cultures with trypsin-EDTA for 3 min, washed with DMEM, and resuspended in DMEM at a density of 2.2 x 106 cells/ml. Type I collagen (Koken, Tokyo, Japan), two kinds of reconstitution buffer, hyalocytes or BRPEs suspension, and distilled water were mixed on ice at a ratio of 7:1:1:1:1 (final concentration of type I collagen gel 1.9 mg/ml; final cell density 2 x 105 cells/ml). The resultant mixture (0.5 ml) was added to a 24-multiwell plate (Nunc, Roskilde, Denmark), and the formation of collagen gel was induced by incubation at 37°C for 60 min. After gels formed, 0.5 ml DMEM containing 10% FBS was added to each well. After 48 h of incubation, the cells were starved with DMEM containing 1% calf serum. The gels were freed from the walls of the culture wells with a microspatula and used for the experiments. The diameter of the collagen gel was measured with a ruler at indicated time points after stimulation. For quantitative purposes, contraction data are presented as the reduction in diameter of the collagen gels.

Northern blot analysis.
Northern blot analysis was performed as previously described (21). Subconfluent hyalocytes, BRPEs, and BRECs under normal growth medium conditions were collected, and total RNA was extracted by the acid guanidine thiocyanate-phenol-chloroform extraction method. Radioactive fibronectin or VEGF cDNA probe was generated using a labeling kit (Multiprime; Amersham) and [32P]dCTP (NEN, Boston, MA). Quantitation of Northern blot analysis was performed using a computing phosphorescence imager (PhosphorImager with ImageQuant software analysis; Molecular Dynamics, Sunnyvale, CA). Lane-loading differences were normalized by rehybridization with radiolabeled 36B4 cDNA probe as an internal control gene.

Scratch wound assays.
The migration of BRECs was assessed in scratch wound assays. After the cells in six-well plates were confluent and starved with DMEM containing 1% calf serum, monolayers in each well were scratched using a micropipette tip (1-ml blue tip) and rinsed gently with PBS before stimulating the cells with or without VEGF (25 ng/ml) and CTGF (100 or 300 ng/ml). Wound closure was monitored for 48 h and photographed using a digital camera, and the number of cells that migrated from initial borders per same unit area was counted. These experiments were repeated three times with similar results.

In vitro tube formation assay.
The effect of CTGF on capillary formation was evaluated using an angiogenesis assay kit (KZ-1000; Kurabo, Tokyo, Japan) according to the provided instructions. Briefly, at day 1, HUVECs cocultured with human fibroblasts were incubated in optimized medium that was contained in the kit (KZ-1500; Kurabo), and the cells were stimulated with or without CTGF (100 or 300 ng/ml). Recombinant human VEGF (25 ng/ml) was added to the medium as a positive control. The media were replaced at days 4, 7, and 9. At day 11, the HUVECs were fixed and stained by an anti-human CD31 antibody (Kurabo). Total areas of tube-like structures were quantified by image analysis using angiogenesis quantitation software (KSW-5000U; Kurabo) and statistically analyzed.

Enzyme-linked immunosorbent assay.
This study was carried out with approval from the institutional review board and performed in accordance with the ethical standards of the 1989 Declaration of Helsinki. We obtained written informed consent to participate from the patients. Vitreous samples were collected from patients who underwent pars plana vitrectomy because of macular hole, rhegmatogenous retinal detachment (RRD), PDR, and PVR, which are the same samples in which we investigated the concentrations of TGF-ß2 and CTGF in our previous study (13). Concentrations of VEGF in the vitreous were measured by human VEGF immunoassay kit (R&D Systems) according to the manufacturer's protocol. Undetectable concentrations of VEGF were determined <31.2 pg/ml. The concentrations of CTGF had been analyzed already in our previous study (13). We investigated whether there is a correlation between the concentrations of VEGF and CTGF in the vitreous with PDR and PVR.

Statistical analysis.
The experimental data are expressed as means ± SD. Statistical significance was assumed when P < 0.05 using the Student's t test in normally distributed populations. The correlation between the differences in CTGF and the differences in VEGF was statistically analyzed using StatView version 5.0 (SAS Institute, Cary, NC), and statistical significance was assumed at P < 0.05 as well.


   RESULTS
TOP
 ABSTRACT
 RESEARCH DESIGN AND METHODS
 RESULTS
DISCUSSION
 REFERENCES
 
Growth-promoting effect of CTGF on hyalocytes and BRPEs.
We addressed the phosphorylation of p44/p42 MAPK and [3H]thymidine incorporation in the presence of CTGF to investigate whether CTGF stimulates cell growth activity. As shown in Fig. 1A and C, whereas quiescent hyalocytes and BRPEs both showed a slight phosphorylation state of p44/p42 MAPK, CTGF stimulated the phosphorylation state of p44/p42 MAPK within 5 min, and maximum activation was achieved after 15 min and maintained at least up to 60 min by both types of cells. Furthermore, CTGF significantly promoted [3H]thymidine incorporation by hyalocytes and BRPEs in a dose-dependent manner (0–100 ng/ml) (Fig. 1B and D). It suggests that CTGF might have growth-promoting effect on hyalocytes and BRPEs.


Figure 1
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  		FIG. 1. Growth-promoting effect of CTGF on hyalocytes and BRPEs. Phosphorylation states of p44/p42 MAPK by hyalocytes (A) and BRPEs (C) are shown. The cells were starved and unstimulated or stimulated with CTGF (100 ng/ml) for the indicated periods of time. Total cell lysates were subjected to Western blot analysis with an antibody against phospho-p44/p42 MAPK (pp44/42 MAPK). Lane-loading differences were normalized by reblotting the membrane with an antibody against p44/p42 MAPK. [3H]thymidine incorporation in the presence of CTGF by hyalocytes (B) and BRPEs (D). After starvation, the cells were untreated or treated with CTGF (1, 10, and 100 ng/ml) for 18 h, and [3H]thymidine was then added (0.25 µCi/well) for an additional 6 h. Incorporated [3H]thymidine was determined by scintillation counting and expressed as a percentage by the induction ratio under the condition of CTGF(–). *P < 0.05 and **P < 0.01 compared with CTGF(–) (n = 4, respectively).

 
Collagen gel contraction by TGF-ß2 and CTGF.
Next, to examine the contractile property of hyalocytes and BRPEs in the presence of CTGF, hyalocytes, and BRPEs, respectively, embedded in type I collagen gels were starved and untreated or treated with TGF-ß2 (0.3 nmol/l) or CTGF (100 ng/ml) for 5 days. Whereas TGF-ß2 significantly decreased the diameter of the collagen gel (hyalocytes, 34.9%, and BRPEs, 36.3% reduction in diameter compared with control, P < 0.01), the gels treated with CTGF showed no significant contraction up to 5 days (Fig. 2A and B). In addition, protein expressions of fibronectin, one of the extracellular matrix proteins, of those kinds of cells embedded in collagen gels were both enhanced in the presence of CTGF compared with control, but the expression was less promoted than that in the presence of TGF-ß2 (Fig. 2C).


Figure 2
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  		FIG. 2. Collagen gel contraction and fibronectin protein expression by hyalocytes and BRPEs in the presence of CTGF. A: Hyalocytes and BRPEs were embedded in type I collagen gels (n = 4). Five days after stimulation in the presence or absence of TGF-ß2 (0.3 nmol/l) and CTGF (100 ng/ml), the gels were photographed. B: The diameter of the collagen gels was measured and expressed as a percentage of the average diameter of control group. **P < 0.01 compared with control. C: The gels were dissolved by collagenase, and the cells were isolated. Total cell lysate was then extracted, and the same amount of protein from the extractions was subjected to Western blot analysis for the detection of fibronectin protein. Signal intensities were quantified and are expressed as a percentage by control. FN, fibronectin. *P < 0.05 and **P < 0.01 compared with control.

 
CTGF-induced expression of fibronectin mRNA by hyalocytes and BRPEs.
To confirm the effect of CTGF on extracellular matrix synthesis, we investigated changes in mRNA expression of fibronectin by hyalocytes and BRPEs in the presence of CTGF. The cells were starved and left unstimulated or stimulated with CTGF (100 ng/ml) for 1, 4, 10, and 24 h, and then total RNA was extracted for Northern blot analysis. As shown in Fig. 3A and B, fibronectin mRNA expression by both hyalocytes and BRPEs was significantly promoted by CTGF in a time-dependent manner up to 24 h (2.07-fold by hyalocytes and 1.75-fold by BRPEs, respectively, compared with control, P < 0.01), demonstrating that CTGF was associated with the synthesis of the extracellular matrix by hyalocytes and BRPEs.


Figure 3
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  		FIG. 3. Fibronectin mRNA expression by hyalocytes and BRPEs in the presence of CTGF. Hyalocytes (A) and BRPEs (B) were untreated or treated with CTGF (100 ng/ml) for the indicated time. CTGF mRNA expression in the presence of CTGF was analyzed by Northern blot analysis. Lane-loading differences were normalized by rehybridization with radiolabeled 36B4 cDNA probe as an internal control gene. Representative blots from three independent experiments are demonstrated respectively. Signal intensities were quantified and are expressed as a percentage by the ratio of CTGF/36B4 at time 0. *P < 0.05 and **P < 0.01 compared with time 0.

 
CTGF does not induce proliferation and migration of BRECs.
We also investigated the proliferative effect of CTGF on BRECs by the examination of phosphorylation of p44/p42 MAPK and [3H]thymidine incorporation. Whereas phosphorylation state of p44/p42 MAPK and thymidine incorporation by hyalocytes and BRPEs in the presence of CTGF were stimulated, neither of them by BRECs were stimulated in the presence of CTGF (100 ng/ml) (Fig. 4A and B). As shown in Fig. 5A and B, compared with the positive control VEGF (2.37-fold compared with control, P < 0.01), CTGF at concentration of 100 or 300 ng/ml did not significantly promote migration of BRECs in scratch wound assays.


Figure 4
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  		FIG. 4. Proliferative effect of CTGF on BRECs. A: Phosphorylation states of p44/p42 MAPK. The cells were starved and unstimulated or stimulated with CTGF (100 ng/ml) for the indicated periods of time. Total cell lysates were subjected to Western blot analysis with an antibody against phosphorylated p44/p42 MAPK (pp44/42 MAPK). Lane-loading differences were normalized by reblotting the membrane with an antibody against p44/p42 MAPK. B: [3H]thymidine incorporation in the presence of CTGF. After starvation, the cells were untreated or treated with CTGF (1, 10, and 100 ng/ml) for 18 h, and [3H]thymidine was then added for an additional 6 h. Incorporated [3H]thymidine was determined by scintillation counting and expressed as a percentage by the induction ratio under the condition of CTGF(–). n = 4.

 

Figure 5
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  		FIG. 5. Migration of BRECs in the presence of CTGF. Scratch wound assays were performed to evaluate the migration of BRECs. Confluent cells in six-well plates were starved, and monolayers in each well were scratched using a micropipette tip. The cells were unstimulated or stimulated with VEGF (25 ng/ml) or CTGF (100 or 300 ng/ml). A: Wound closure was monitored for 48 h and photographed. B: The number of cells that migrated from wounded edge was counted and expressed as a number of cells per same unit area. *P < 0.01 compared with control; N.S., not significant compared with control.

 
CTGF did not promote in vitro tube formation.
In the presence of VEGF, HUVECs generated tube-like structures, and the area of the structures was significantly greater than control (1.95-fold compared with control, P < 0.01), whereas CTGF did not promote formation of tube-like structures at the concentration of either 100 or 300 ng/ml compared with control (Fig. 6A and B).


Figure 6
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  		FIG. 6. In vitro tube formation. In vitro tube formation in the presence of CTGF was analyzed using an angiogenesis assay kit (Kurabo) in co-cultures of HUVECs and fibroblast of dermal origin. A: After 11 days of treatment with or without VEGF (25 ng/ml) and CTGF (100 or 300 ng/ml), the HUVECs were fixed and stained by CD31 antibody. B: Total areas of tube-like structures were quantified by image analysis using an angiogenesis assay program (Kurabo) and expressed as a percentage of the area under the condition of control. *P < 0.01 compared with control; N.S., not significant compared with control.

 
CTGF induced VEGF mRNA expression by hyalocytes and BRPEs.
To investigate whether CTGF enhances VEGF expression, hyalocytes, BRPEs, and BRECs were starved and unstimulated or stimulated with CTGF (100 ng/ml) for 4 h. Total RNA was extracted, and Northern blot analysis was performed for the detection of VEGF mRNA. As shown in Fig. 7, VEGF mRNA was enhanced by hyalocytes and BRPEs in the presence of CTGF, whereas CTGF did not promote VEGF mRNA expression by BRECs.


Figure 7
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  		FIG. 7. CTGF-induced VEGF gene expression by vitreoretinal cells. Hyalocytes, BRPEs, and BRECs were starved and untreated or treated with CTGF (100 ng/ml) for 4 h. Total RNA was subjected to Northern blot analysis for VEGF mRNA. Lane-loading differences were normalized by rehybridization with radiolabeled 36B4 cDNA probe as an internal control gene. Representative blots from three independent experiments are demonstrated, respectively. Signal intensities were quantified and are expressed as a percentage by the ratio of VEGF/36B4 at time 0. *P < 0.01 compared with time 0, respectively.

 
Correlation between CTGF and VEGF in the vitreous in eyes with PDR and PVR.
Average concentrations of CTGF protein in the vitreous were 38.90 ng/ml (macular hole; n = 15), 39.87 ng/ml (RRD; n = 18), 56.63 ng/ml (PDR; n = 41), and 69.47 ng/ml (PVR; n = 10). Concentrations of CTGF in eyes with PDR or PVR were significantly elevated compared with those with macular hole and RRD, respectively (P < 0.01) (13). Average concentrations of VEGF protein in the vitreous were 11.17 pg/ml (macular hole; n = 15), 17.49 pg/ml (RRD; n = 18), 939.2 pg/ml (PDR; n = 41), and 173.3 pg/ml (PVR; n = 10). Concentrations of VEGF in eyes with PDR or PVR were also significantly elevated compared with those with macular hole and RRD, respectively (PDR vs. macular hole and RRD, P < 0.01 and PVR vs. macular hole and RRD, P < 0.05) (data not shown). However, there was no significant correlation between the concentrations of CTGF and VEGF in the vitreous with PDR and PVR (r = 0.195, P = 0.171, Fig. 8).


Figure 8
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  		FIG. 8. Correlation between the concentrations of CTGF and VEGF in the vitreous with PDR and PVR. Vitreous samples were collected from patients with proliferative vitreoretinal diseases (PDR and PVR), and then concentrations of CTGF and VEGF in the vitreous were measured by enzyme-linked immunosorbent assay. ({blacksquare}, PDR [n = 41]; {square}, PVR [n = 10]). Statistical analysis for the correlation between CTGF and VEGF was performed (r = 0.195, P = 0.171, not significant). Values below 0 on the y-axis denote undetectable concentrations of VEGF <31.2 pg/ml.

 

   DISCUSSION
TOP
 ABSTRACT
 RESEARCH DESIGN AND METHODS
 RESULTS
 DISCUSSION
REFERENCES
 
Recent studies, including our previous one, have demonstrated that CTGF levels were elevated in the vitreous and proliferative preretinal membranes in eyes with PDR and PVR (11–13) and thus suggested the involvement of CTGF in the pathogenesis of proliferative vitreoretinal diseases. In the present study, we demonstrated the functional characteristics of CTGF on vitreoretinal cells, such as hyalocytes, BRPEs, and BRECs, for the first time and clarified its functional mechanisms, at least in part. Furthermore, we addressed the expressional interaction between CTGF and VEGF in vitro and in vivo.

CTGF enhanced the phosphorylation state of p44/p42 MAPK and [3H]thymidine incorporation, increased the number of cells significantly (at the concentration of 100 ng/ml; data not shown), and also promoted mRNA and protein expression of fibronectin by both hyalocytes and BRPEs, suggesting that CTGF might stimulate cell growth and extracellular matrix synthesis by hyalocytes and BRPEs. In addition, our previous study revealed that hyalocytes and BRPEs could also be the sources of CTGF (13), and fibronectin is indicated to be a major component of the extracellular matrix proteins observed in epiretinal membranes of PDR and PVR (25). Taken together, CTGF secreted by hyalocytes and RPEs might modulate the formation of proliferative fibrous membrane in an autocrine and paracrine manner in the pathogenesis of proliferative vitreoretinal diseases, such as PDR and PVR. On the other hand, CTGF could not significantly induce the contraction of type I collagen gel embedding hyalocytes and BRPEs in this study, whereas TGF-ß2 remarkably induced its contraction (26,27). CTGF is indicated to share many biological functions with TGF-ß in causing fibrosis, and thus it is considered to be one of the downstream mediators of TGF-ß (8,9). CTGF might not share the contractile property with TGF-ß, despite enhanced expression of fibronectin by both types of cells. Therefore, it is considered that CTGF might not be involved or have minimal involvement, if any, in the cicatricial contraction of proliferative membranes in eyes with proliferative vitreoretinal diseases.

Whereas CTGF might have a growth-promoting effect on hyalocytes and BRPEs, CTGF enhanced neither phosphorylation state of p44/p42 MAPK nor [3H]thymidine incorporation by BRECs, suggesting that CTGF might not have a direct effect on the proliferation of BRECs. These results indicated that the effect of CTGF on cellular proliferation might be cell-type dependent. In addition, CTGF did not promote the migration by BRECs or in vitro tube-like formation by HUVECs co-incubated with dermal fibroblasts, suggesting that CTGF might not be directly associated with neovascularization. However, those results are more controversial, because this result does not support some of previous findings that examined cellular functions of CTGF on various types of endothelial cells. Regarding cell proliferation, CTGF has been indicated to promote cell proliferation by BAECs (15) but not to directly mediate cell proliferation by HUVECs (14) or choroidal endothelial cells (18). In addition, CTGF has been demonstrated to promote the migration of endothelial cells by Boyden chamber assay and to promote in vitro tube formation by HUVECs, BAECs, and choroidal endothelial cells (14,15,18). This difference from our results may be due to endothelial cell heterogeneity, which has been well documented.

Another reason is presumed that CTGF has been indicated to interact with other angiogenic proteins at many different levels and to use indirect mechanisms to regulate endothelial cell function (28). It has been reported that CTGF modulates the activity of basic fibroblast growth factor, one of the potent angiogenic molecules, by HUVECs (14), and it has recently been revealed that CTGF transcriptionally regulates VEGF by airway smooth muscle cells (29). We also demonstrated that CTGF stimulates VEGF mRNA expression by hyalocytes and BRPEs but not by BRECs. VEGF is widely known as a mitogen of endothelial cells and as a vasopermeability factor (30). Therefore, even though CTGF had no direct effects on the proliferation or migration of BRECs or on tube formation by HUVECs, it is possible that CTGF-induced VEGF secreted by hyalocytes and RPEs might have such effects on retinal vascular endothelial cells in a paracrine manner in vivo. Besides, conversely, VEGF has been reported to enhance CTGF expression by retinal capillary endothelial cells (RECs) (31). It is also possible that VEGF-induced CTGF secreted by RECs might mediate the formation of proliferative fibrous membrane in proliferative vitreoretinal diseases through the proliferation and extracellular matrix synthesis by hyalocytes and RPEs in a paracrine manner.

According to those in vitro analyses, there might be an expressional correlation between CTGF and VEGF in the progression of proliferative vitreoretinal diseases, such as PDR and PVR. In addition, CTGF knockout mice exhibit reduced VEGF expression and vascular defects during embryogenesis and fetal development, despite the absence of the examination of retinal capillary vessels (32). However, we demonstrated that there is no significant correlation between the concentrations of CTGF and VEGF in the vitreous with PDR and PVR, even though concentrations of CTGF and VEGF were both significantly higher in the vitreous with PDR and PVR than those in the vitreous with nonproliferative vitreoretinal diseases (macular hole and RRD), respectively (data not shown), similar to the previous reports (13,33). CTGF expression is known to be strongly enhanced by TGF-ß and also promoted by hypoxia (34) and high glucose (35), and VEGF expression is also known to be potently enhanced by hypoxia and further promoted by high glucose and many other cytokines and growth factors, such as TGF-ß and tumor necrosis factor-{alpha}. Therefore, VEGF-induced CTGF and CTGF-induced VEGF are respectively presumed to be only minor components in entire expressions of CTGF and VEGF in the vitreous with proliferative vitreoretinal diseases. A fairly recent study sustains our results at least partially, which demonstrated that the CTGF level in the vitreous significantly correlates with the severity of fibrosis but not neovascularization activity by multifactorial ANOVA in consideration of sex, age, diabetes, degree of neovascularization, degree of hemorrhage, and fibrosis score (36).

In this study, we demonstrated functional characteristics of CTGF on vitreoretinal cells and indicated that CTGF might play an important role in the formation of proliferative fibrous membranes in proliferative vitreoretinal diseases, such as PDR and PVR, but CTGF might not directly associate with the cicatricial contraction of the membranes. Furthermore, our results also suggested that CTGF might have no direct effects or minimal effects, if any, on retinal neovascularization, however, we demonstrated the possibility that CTGF might have indirect effects by modulating the expression of VEGF by neighboring cells. Further investigations in vitro and in vivo are still necessary to clarify the interaction among CTGF, VEGF, and other cytokines or growth factors and their direct or indirect effects on the pathogenesis of proliferative vitreoretinal diseases.


   ACKNOWLEDGMENTS
 
This study has received Grant-in-Aid for Scientific Research 17591839 from the Ministry of Education, Science, Sports and Culture, Japan.


   FOOTNOTES
 
Published ahead of print at http://diabetes.diabetesjournals.org on 15 February 2007. DOI: 10.2337/db06-1644.

The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

Received for publication November 23, 2006 and accepted in revised form February 5, 2007


   REFERENCES
TOP
 ABSTRACT
 RESEARCH DESIGN AND METHODS
 RESULTS
 DISCUSSION
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