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Endocrinology Vol. 145, No. 8 3739-3745
Copyright © 2004 by The Endocrine Society

Crucial Role of Activin A in Tubulogenesis of Endothelial Cells Induced by Vascular Endothelial Growth Factor

Kyoko Maeshima, Akito Maeshima, Yoshiro Hayashi, Shoji Kishi and Itaru Kojima

Institute for Molecular & Cellular Regulation (K.M., A.M., Y.H., I.K.), Gunma University; and Department of Ophthalmology (K.M., S.K.); and Third Department of Internal Medicine (A.M.), Gunma University School of Medicine, Maebashi 371-8512, Japan

Address all correspondence and requests for reprints to: Itaru Kojima, M.D., Institute for Molecular & Cellular Regulation, Gunma University, Maebashi 371-8512, Japan. E-mail: ikojima{at}showa.gunma-u.ac.jp.


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
The present study was conducted to elucidate the role of activin A in tubulogenesis of vascular endothelial cells. Activin A was produced in bovine aortic endothelial cells (BAEC). These cells also expressed the type I and type II activin receptors. When added to BAEC cultured in a collagen gel, activin A induced capillary formation. Activin A was as potent as vascular endothelial growth factor (VEGF) and markedly enhanced VEGF-induced tubulogenesis. To examine the role of endogenous activin A, we added follistatin, an inhibitor of activin A. Follistatin nearly completely blocked the VEGF-induced tubulogenesis, and the effect of follistatin was reproduced by transfection of the dominant-negative type II activin receptor gene. In BAEC, activin A increased the expression of VEGF and the VEGF receptors, Flt-1 and Flk-1. On the other hand, VEGF increased the production of activin A. Finally, addition of follistatin, which blocks the action of endogenous activin A, reduced the expression of Flt-1 and Flk-1. These results indicate that an autocrine factor activin A amplifies the effect of VEGF by up-regulating VEGF and its receptors. This effect of activin A is critical in the VEGF-induced tubulogenic morphogenesis in BAEC.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
ANGIOGENESIS, THE FORMATION of new blood vessels by sprouting from existing vessels, plays a critical role in a variety of physiological events including organogenesis during development, tissue repair, and maintenance of reproductive functions (1). Angiogenesis is also involved in the pathophysiology of various diseases: for example, tumor growth and metastasis, rheumatoid arthritis, and proliferative retinopathies (2). Vascular endothelial growth factor (VEGF), originally identified as a vascular-permeability factor (3), is one of the most important factors regulating physiological as well as pathological angiogenesis (4). It is essential for the formation of blood vessels during development, and disruption of the VEGF gene is lethal in mice (5). Because angiogenesis is involved in the pathogenesis of various diseases, manipulation of the VEGF action can provide therapeutic approaches for various vascular disorders. Thus, administration of VEGF (6) or introduction of the VEGF gene (7) may be beneficial for the treatment of ischemic vascular diseases by promoting angiogenesis. Conversely, blocking the effect of VEGF may be effective for the treatment of cancers and diabetic retinopathy (8). There are three tyrosine kinase VEGF receptors (VEGFRs), Flt-1 (VEGFR-1), Flk-1/KDR (VEGFR-2) and Flt-4 (VEGFR-3). Flt-1 and Flk-1 are expressed in vascular endothelial cells and are thought to regulate angiogenesis (9). Although the signal transduction pathways activated by VEGF are complex, VEGF induces angiogenesis by stimulating growth and migration of endothelial cells and increasing protease synthesis. Although angiogenesis is a complex phenomenon comprised of many biological reactions, the angiogenic activity of VEGF can be assessed in an in vitro culture system by measuring tubulogenic morphogenesis. Thus, VEGF induces the formation of capillary structures when added to endothelial cells cultured in a collagen gel (9).

Activin A is a member of the TGF-ß superfamily and elicits diverse effects in various biological systems (10). These include morphogenesis and organogenesis during development, modulation of growth and differentiation of various types of cells, and regulation of reproduction (11). The action of activin A is modified by follistatin, an activin-binding protein that blocks its activity (12). Thus, the activin-follistatin system provides a complex regulation of growth and differentiation in various tissues. Recent studies have shown that activin A acts as an autocrine or paracrine factor to regulate branching tubulogenesis in the pancreas, kidneys, and lung (13). Both activin and follistatin are expressed in vascular tissues and are up-regulated in pathological conditions (14, 15). They are thought to be involved in vascular remodeling (16). Because tubulogenic morphogenesis is a critical step in the formation of blood vessels, it is of interest to determine whether or not activin A regulates tubulogenesis of vascular endothelial cells. In the present study, we investigated the role of activin A in VEGF-induced tubulogenesis using bovine aortic endothelial cells (BAEC). The results indicate that autocrine factor activin A is critical in tubulogenesis of BAEC induced by VEGF.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Materials
Recombinant human activin A and follistatin were provided by Dr. Y. Eto of the Central Research Laboratory, Ajinomoto Inc. (Kawasaki, Japan). Recombinant human VEGF121 was purchased from R&D Systems (Minneapolis, MN). Recombinant human TGF-ß1 was obtained from Sigma (St. Louis, MO). Polyclonal antihuman VEGF, polyclonal antihuman Flt-1, polyclonal antihuman Flk-1, and polyclonal antibody against the type II activin receptor were purchased from Santa Cruz Biotechnology (Santa Cruz, CA).

Cell culture and measurement of cell proliferation
BAEC were purchased from Dainippon Pharmaceutical Co. Ltd. (Osaka, Japan). They were cultured in DMEM (Invitrogen Life Technologies, Grand Island, NY), with 10% fetal bovine serum (Invitrogen Life Technologies), penicillin, and streptomycin in an atmosphere of 5% CO2-95% air at 37 C. BAEC were used from passages 8–20. Morphologically, these cells had cobble stone-like appearance and more than 95% of them expressed factor VIII, a marker of endothelial cells.

The number of living cells was assessed by using MTT [3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide] (17). After incubation for an indicated period, cells were treated with MTT for 3 h at a final concentration of 1 mg/ml in the culture medium, MTT extracted by adding 2-propanol containing 0.04 M HCl, and OD was measured at 590 nm.

Measurement of tubulogenesis
Cells were suspended at 1 x 105 cells/ml in a neutralized collagen solution (Koken, Tokyo, Japan), dispensed into 24-well plates and incubated at 37 C. After the collagen solution had gelled, complete culture medium with the indicated agent was added, which was renewed every 2–3 d. Photographs of the cells were taken at the indicated time using a Nikon Diaphot TMD inverted microscope (Tokyo, Japan).

To semiquantify tubulogenic activity, morphological change after 5 d of culture with indicated factors was observed. Fifty colonies per experimental condition were randomly selected. Then, the percentage of cell clusters containing capillary structures was determined, and the tube length (long axis) was also measured by using image analysis software (Image program, National Institutes of Health, Bethesda, MD).

For histological analysis, collagen gels were fixed with 4% formaldehyde in PBS, removed from the cultured dishes, dehydrated in graded concentrations of ethanol and embedded in paraffin blocks according to standard procedures. Four-micrometer sections were cut and stained with hematoxylin and eosin.

RT-PCR
Total RNA was isolated with the Trizol Reagent (Invitrogen Life Technologies) from BAEC. First-strand cDNA was made from total RNA using the superscript Preamplification System (Invitrogen Life Technologies) as described previously (18). Contaminated genomic DNA was removed with ribonuclease-free deoxyribonuclease. Five micrograms of deoxyribonuclease-treated RNA were incubated with 1 µl oligo(deoxythymidine) at 70 C for 10 min. Two microliters of 10x PCR buffer, 1 µl dithiothreitol (0.1 M), 2 µl deoxynucleotide triphosphate mix (10 mM), and 2 µl MgCl2 (25 mM) were added to each reaction. After incubation for 5 min at 42 C, 1 µl reverse transcriptase was added. Samples were incubated at 42 C for 50 min, then at 70 C for 15 min. Ribonuclease H (1 µl) was added to each reaction and samples were incubated at 37 C for 20 min. PCR was performed as indicated by the manufacturer (PerkinElmer) with the primers described in Table 1Go. Reactions included 5 µl of 10x PCR buffer, 2 µl MgCl2 (50 mM), 1 µl deoxynucleotide triphosphate mix, 1 µl 3'-primer, 1 µl 5'-primer, 0.5 µl Taq polymerase, and 1 µl cDNA. Samples were incubated at 94 C for 5 min, followed by the indicated cycles of 30 sec at 94 C, 30 sec at 53 or 54 or 56 or 63 C, 90 sec at 72 C, and final extension at 72 C for 10 min in a PerkinElmer (Foster City, CA) DNA Thermal Cycler.


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  		TABLE 1. Primers used in this study

 
Immunoblot analysis
Cells were washed three times with PBS, suspended in Laemmli buffer, and heated to 100 C for 10 min. After centrifugation, the supernatant was collected, and the protein concentration was determined by using a protein assay kit (Bio-Rad Laboratories, Hercules, CA). Twenty micrograms of protein from each sample were separated by SDS-PAGE under reducing conditions and transferred to a polyvinlidene difluoride membrane (Nihon Millipore, Yonezawa, Japan) by electroblotting. To reduce nonspecific antibody binding, the membrane was blocked with 5% BSA, and 0.1% NaN3 dissolved in Tris-saline for 1 h at 37 C, then incubated overnight with primary antibody and washed with Tris-PBS. After incubation with peroxidase-labeled secondary antibody for 1 h at room temperature, the membrane was washed with Tris-PBS and analyzed by exposure to x-ray film using ECL Western blotting detection reagent (Amersham Life Science, Buckinghamshire, UK).

Construction and transfection of replication-deficient recombinant adenovirus vector
The recombinant adenovirus AdextARII, carrying truncated type II activin receptor (tARII) cDNA was generated as described previously (19). The recombinant adenovirus AdexLacZ, carrying the LacZ gene, which encodes the Escherichia coli ß-galactosidase, was provided by Dr. T. Takeuchi (Gunma University) and was used as a control for confirming successful transfection.

Serum-starved BAEC in a monolayer culture were infected with AdexLacZ or AdextARII at a titer of indicated multiplicity of infection (MOI) for 2 h at 37 C and then cultured in serum-free medium for 24 h. Cells were then thoroughly washed with PBS, trypsinized, and used for collagen gel culture.

ß-Galactosidase expression was detected as the development of blue pigmentation due to the enzymatic cleavage of X-gal (19).

Statistical analysis
The significance of differences between means was determined by Student’s t test.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
Expression of the ßA-subunit of activin and activin receptors in BAEC
To investigate the role of activin A in endothelial cells, we first examined the expression of the ßA-subunit of activin and activin receptors in BAEC by RT-PCR. As shown in Fig. 1AGo, the expression of mRNA for these proteins was observed in BAEC. The ßA-subunit of activin and the type II activin receptor were also detected in BAEC by Western blotting (Fig. 1BGo). These results suggest that activin acts as an autocrine factor in BAEC.



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  		FIG. 1. Expression of activin A and activin receptors in BAEC. A, mRNA was obtained from BAEC and the expression of ßA-subunit of activin (lane 1), type II (lane 2), and type I (lane 3) activin receptors was measured by RT-PCR. M, Markers. Results are representative of three independent experiments. B, Expression of ßA-subunit and type II activin receptor was measured by Western blotting. Results are representative of three independent experiments.

 
Effect of activin A and VEGF on tubulogenesis of BAEC
As shown in Fig. 2AGo-a, BAEC cultured in complete medium did not form tube structure. We then examined the effect of activin A on the morphology of BAEC. When cultured with activin A, BAEC formed a capillary network (Fig. 2AGo, b and c). Note that we observed formation of a lumen in tubular structure induced by activin A (Fig. 2AGo-d). TGF-ß (50 pM) did not induce tube formation in BAEC cultured in collagen gel.



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  		FIG. 2. Effects of activin A and VEGF on tubulogenesis of BAEC. A, BAEC were cultured in a collagen gel for 7 d in the absence (a) or presence (b and c) of 4 nM activin A. d, HE staining of the section. a and b, x100; c, x400; d, x1000. Results are representative of five independent experiments. B, BAEC were cultured for 7 d in a collagen gel in the absence (a) or presence of 100 ng/ml VEGF (b), 4 nM activin A (c), or 100 ng/ml VEGF plus 4 nM activin A (d). a–d, x100. Results are representative of five independent experiments. C, BAEC were cultured for 7 d in a collagen gel in the absence (none) or presence of 4 nM activin A (act), 100 ng/ml VEGF or 4 nM activin A plus 100 ng/ml VEGF (act + VEGF). The numbers of tube structure (a) and tube length (b) were measured as described in Materials and Methods. Values are the mean ± SE for six experiments. *, P < 0.05 vs. none.

 
We next examined the effect of VEGF, an important regulator of angiogenesis, on the morphology of BAEC. As shown in Fig. 2BGo-b, BAEC formed a capillary network when cultured with VEGF. We observed a lumen in tubular structure induced by VEGF (data not shown). We then examined the effect of activin A on VEGF-induced capillary formation in BAEC cultured in collagen gel. Indeed, when cultured with a combination of VEGF and activin A, BAEC formed considerably more elongated capillary networks (Fig. 3BGo-d). Note that these concentrations of VEGF and activin A exerted the maximal effect when added one at a time. Morphometrical analysis was carried out by measuring two parameters, the number of tube structure, and the tube length. In terms of the number of tube structures formed, 4 nM activin A was much more effective than 100 ng/ml VEGF. There was no additivity between the actions of activin A and VEGF (Fig. 2CGo-a). In terms of tube length, 4 nM activin A was slightly more effective than VEGF, and the effects of the two factors are additive (Fig. 2CGo-b).



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  		FIG. 3. Effect of follistatin on VEGF-induced capillary formation. A, BAEC were incubated for 7 d in collagen gel in the absence (a) or presence of 100 ng/ml VEGF (b), 10 nM follistatin (c), or 100 ng/ml VEGF plus 10 nM follistatin (d). a–d, x100. Results are representative of five independent experiments. B, BAEC were incubated for 7 d in a collagen gel in the presence of 100 ng/ml VEGF and various concentrations of follistatin. The numbers of the tube structure (a) and tube length (b) were measured. Values are the mean ± SE for six experiments. *, P < 0.05 vs. without follistatin.

 
Effect of follistatin on VEGF-induced capillary formation
As mentioned above, activin A is produced in BAEC. To assess the role of endogenous activin A in tubulogenesis of BAEC, we administered follistatin, an antagonist of activin A. Follistatin by itself had no significant effect on angiogenesis, suggesting that endogenous activin A does not have significant effect under basal condition. Surprisingly, when cultured with both VEGF and follistatin, VEGF-mediated capillary formation was markedly inhibited (Fig. 3AGo). Morphometrical analysis showed that follistatin inhibited the number of VEGF-induced tube structures in a dose-dependent manner, and at a concentration of 1 nM, follistatin nearly completely blocked the effect of VEGF (Fig. 3BGo). In terms of tube length, follistatin nearly completely inhibited the VEGF-induced tube structure at a concentration of 1 nM and above (Fig. 3BGo).

Gene transfer of truncated type II activin receptor using adenovirus vector
To further examine the role of activin A in branching morphogenesis of BAEC, we constructed a cDNA encoding truncated type II activin receptor, which lacks intracellular kinase domain (19), and transfected BAECs with this cDNA using adenovirus vector (19). To determine the efficacy of transfection using the adenovirus vector, we infected BAEC with AdexLacZ at various titers of MOI and examined the expression of ß-galactosidase. The efficacy of transfection by using adenoviral vector was dependent on the titer of the viral solution. We used an adenoviral vector at a titer of 10 MOI for the next study using AdextARII because no cytotoxity was observed at this concentration and almost all BAEC exhibited ß-galactosidase activity. When we infected BAEC with AdextARII, we were able to detect the expression of this mutant receptor by Western blotting (Fig. 4AGo). Two bands (62 and 75 kDa) indicate endogenous type II activin receptor, and the band around 45 kDa represents an overexpressed mutant receptor. As predicted, we could not observe an activin-induced capillary network in BAEC infected with AdextARII (Fig. 4BGo-g).



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  		FIG. 4. Infection of adenovirus vector encoding truncated type II activin receptor. A, BAEC were infected with adenovirus vector containing tARII or LacZ and the expression of the type II receptor was determined by Western blotting. B, BAEC infected with adenovirus vector containing LacZ (a–d) or AdextARII (e–h) were incubated in a collagen gel for 7 d in the absence (a and c) or presence of 100 ng/ml VEGF (b and f), 4 nM activin A (c and g), or 100 ng/ml VEGF plus 4 nM activin A (d and h). Results are representative of five independent experiments.

 
To further investigate the role of endogenous activin A in VEGF-induced capillary formation, we examined the changes in morphology of BAEC infected with AdextARII. As shown in Fig. 4BGo, a and e, BAEC infected with AdextARII or AdexLacZ did not form capillary structures under basal condition. When cultured with VEGF, BAEC infected with AdexLacZ formed capillary network, which was similar in noninfected BAEC. In contrast, BAEC infected with AdextARII did not form capillary structures even in the presence of VEGF (Fig. 4BGo-f). Furthermore, combination of VEGF and activin A, which induced marked tubulogenesis in normal BAEC, did not induce the formation of a capillary network in BAEC expressing tARII (Fig. 4BGo-h).

Effect of activin A on the expression of VEGF, Flt-1, and Flk-1
To clarify the mechanism of activin action, we studied the effect of activin A on the expression of VEGF in BAEC by RT-PCR. As shown in Fig. 5Go, activin A increased the expression of VEGF in BAEC. Western blot analysis also showed increase in the production of VEGF protein in BAEC treated with activin A (Fig. 5BGo). Activin A also increased the expression of VEGF receptors, Flt-1 and Flk-1 at mRNA and protein levels (Fig. 5Go, A and C). We next addressed whether or not VEGF modified the production of activin A in BAEC. We determined the effect of VEGF on the production of activin A by Western blotting. As shown in Fig. 5DGo, VEGF induced the production of activin A in BAEC. Above results indicated that activin A increases production of VEGF. To address whether or not activin-induced tubulogenesis was dependent on VEGF produced in endothelial cells, we measured the effect of activin A in the presence of excess amount of neutralizing anti-VEGF antibody. The activin-induced tubulogenesis was only slightly inhibited by anti-VEGF antibody. Thus, the number of tube formation was approximately 90% in the presence of neutralizing excess amount of anti-VEGF antibody. We also examined the effect of activin A on the VEGF receptors. Anti-VEGF antibody did not affect activin-induced up-regulation of the VEGF receptors (data not shown). We also examined whether TGF-ß increased the expression of VEGF. As depicted in Fig. 5EGo, 50 pM TGF-ß did not affect the production of VEGF.



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  		FIG. 5. Interaction of the activin and VEGF signaling system. A, Effect of activin A on the mRNA expression of VEGF and VEGF receptors. BAEC were cultured in monolayer and incubated for indicated time with 4 nM activin A and changes in the mRNA expression of VEGF, Flt-1 and Flk-1 were measured by RT-PCR. The results are representative of three experiments. B, BAEC were cultured in monolayer and incubated for 24 h with 4 nM activin A. Cells were then lysed and the expression of VEGF was determined by Western blotting. Results of the densitometric analyses are shown in the lower panel. Values are the mean ± SE for three experiments. *, P < 0.001. C, BAEC were incubated for 24 h with 4 nM activin A, and the expression of Flt-1 and Flk-1 was determined by Western blotting. Values are the mean ± SE for three experiments. *, P < 0.01. D, BAEC were incubated for 24 h with 100 ng/ml VEGF and the expression of ßA-subunit of activin was measured by Western blotting. Values are the mean ± SE for three experiments. *, P < 0.01. E, BAEC were incubated for 24 h with 50 pM TGF-ß, and the expression of VEGF was measured by Western blotting. Values are the mean ± SE for three experiments.

 
Effect of follistatin on the expression of VEGF, Flt-1, and Flk-1
The above results indicate that endogenous activin A is critical in VEGF-induced tubulogenesis. To further confirm the role of endogenous activin A, we addressed whether or not endogenous activin A functions to maintain the expression of VEGF and its receptors. To this end, we incubated BAEC with follistatin to block the endogenous activin action and measured the expression of VEGF, Fllt-1, and Flk-1 in BAEC. As shown in Fig. 6Go, follistatin reduced the expression of VEGF and its receptors in BAEC.



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  		FIG. 6. Effect of follistatin on the expression of VEGF and the VEGF receptor. BAEC were incubated for 24 h with 10 nM follistatin, and the expression of VEGF, Flt-1, and Flk-1 was measured by Western blotting. Results of the densitometoric analyses are shown in the lower panel. Values are the mean ± SE for three experiments. *, P < 0.01; **, P < 0.001.

 

   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
Results obtained in the present study clearly show that activin A is an autocrine factor produced in BAEC and promotes the formation of capillary structure in collagen gel. To our knowledge, this is the first report describing tubulogenic action of activin A in vascular endothelial cells, and at least in BAEC, activin A is more potent than VEGF in inducing tubulogenic morphogenesis. The present results also show that activin A exerts diverse effects on the VEGF system in BAEC, including up-regulation of the ligand, VEGF, and the VEGF receptors, Flt-1 and Flk-1.

Of particular importance is the interaction of the activin A and the VEGF signaling systems. Exogenously added VEGF up-regulates the production of activin A, a ligand capable of inducing branching morphogenesis by itself. Not only is it tubulogenic by itself, activin A also up-regulates the expression of VEGF receptors, Flt-1 and Flk-1, which may lead to amplification of the VEGF signaling. Activin A also stimulates the production of VEGF in BAEC and feeds forward the two systems. These events may explain the synergism between the actions of VEGF and activin A. The most important observation is that follistatin, an antagonist of activin A, nearly completely blocks the tubulogenesis of endothelial cells induced by VEGF. In addition, the effect of follistatin is reproduced by transfection of the dominantly negative type II activin receptor. Hence, the amplification mechanism involving autocrine activin A is functionally important in the VEGF-induced capillary formation. Without this amplification mechanism, the VEGF action is abolished. Autocrine factor activin A is therefore critical in the tubulogenic action of VEGF. The role of autocrine factor activin A under the basal condition is best demonstrated by the finding that addition of follistatin significantly reduced the expression of the VEGF receptors, Flt-1 and Flk-1. The signal transduction pathway of VEGF is thus maintained by endogenously produced activin A in BAEC. Without the activin action, VEGF is no longer effective in promoting tubulogenesis.

It is well known that TGF-ß induces angiogenesis both in vitro and in vivo. Although this factor inhibits growth of endothelial cells (22), it induces tubular formation in three-dimensional collagen gel (23, 24). TGF-ß promotes the formation of new capillary when injected to mice (25, 26). The role of TGF-ß in the formation of blood vessels was also demonstrated in mutant mice lacking genes for TGF-ß1, or the type I and type II TGF-ß receptors (27, 28, 29). Deletion of these genes results in defective vasculogenesis, defective endothelial cell differentiation, and derangements in capillary formation (27, 28, 29). Taken together, members of the TGF-ß superfamily play critical roles in the formation and remodeling of blood vessels.

Some reports have shown that activin A inhibits angiogenesis in vivo: Kozian et al. (30) showed that follistatin induces angiogenesis in rabbit cornea; and activin A has also been identified as an antiangiogenic factor produced in cancer cells (31). These reports are apparently contradictory to the present results. Although the exact reason for the discrepancy is not certain, some explanations are possible. For instance, we measured the capillary formation of endothelial cells in a collagen gel. This assay is a simple system to monitor tubulogenesis in vitro but does not fully represent angiogenesis in vivo. Actually, angiogenesis in vivo is a complicated event involving a variety of reactions, and it is possible that activin A inhibits angiogenesis in vivo despite the tubulogenic actions. Another possibility is that the action of activin A differs depending upon the types of endothelial cells. Alternately, the effect of activin A is modified by other factors, and the expression of the putative factor differs depending upon types of tissues. Further studies are required to clarify these issues.

The present results provide some new insights into the therapeutic approaches to diseases involving angiogenesis. Because VEGF plays a pivotal role in the pathophysiology of such diseases, manipulation of the VEGF action would be useful for the treatment of pathological angiogenesis. For example, tumor growth is dependent on angiogenesis, and blockade of angiogenic action of VEGF would suppress tumor growth (8). Similarly, VEGF is an important factor in the pathophysiology of diabetic retinopathy. On the other hand, promotion of angiogenesis is beneficial in various types of ischemic vascular diseases. Administration of VEGF (6) and gene therapy using the VEGF gene (7) have been applied for such disorders. The present results raise a possibility that, instead of VEGF agonist or VEGF antagonist, administration of follistatin or activin A would be effective to modify the angiogenic action of VEGF. Modification of the activin-follistatin system in blood vessels may have a potential for the treatment of various disorders involving angiogenesis.


   Acknowledgments
 
The authors are grateful to Mayumi Odagiri for secretarial and technical assistance.


   Footnotes
 
Abbreviations: BAEC, Bovine aortic endothelial cells; MOI, multiplicity of infection; MTT, 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyl-tetrazolium bromide; tARII, truncated type II activin receptor; VEGF, vascular endothelial growth factor; VEGFR, VEGF receptor.

Received February 18, 2004.

Accepted for publication April 21, 2004.


   References
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 

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  20. Deleted in proof
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