- Research article
- Open Access
The response of VEGF-stimulated endothelial cells to angiostatic molecules is substrate-dependent
© Addison et al; licensee BioMed Central Ltd. 2005
- Received: 30 December 2004
- Accepted: 31 October 2005
- Published: 31 October 2005
The microenvironment surrounding cells can exert multiple effects on their biological responses. In particular the extracellular matrix surrounding cells can profoundly influence their behavior. It has been shown that the extracellular matrix composition in tumors is vastly different than that found in normal tissue with increased amounts of certain matrices such as collagen I. It has been previously demonstrated that VEGF stimulation of endothelial cells growing on type I collagen results in the induction of bcl-2 expression and enhanced endothelial cell survival. We sought to investigate whether this increased endothelial cell survival resulted in the failure of angiostatic molecules to inhibit angiogenesis.
We now demonstrate that VEGF-induced survival on collagen I impairs the ability of three known angiostatic molecules, TSP-1, IP-10 and endostatin to inhibit endothelial cell proliferation. Apoptosis of endothelial cells, growing on collagen I, induced by TSP-1 and IP-10 was also inhibited following VEGF stimulation. In contrast, endostatin induced apoptosis in these same cells. Further analysis determined that endostatin did not decrease the expression of bcl-2 nor did it increase activation of caspase-3 in the presence of VEGF. Alternatively, it appeared that in the presence of VEGF, endostatin induced the activation of caspase-8 in endothelial cells grown on collagen I. Furthermore, only endostatin had the ability to inhibit VEGF-induced sprout formation in collagen I gels.
These data suggest that TSP-1, IP-10 and endostatin inhibit endothelial cells via different mechanisms and that only endostatin is effective in inhibiting angiogenic activities in the presence of collagen I. Our results suggest that the efficacy of angiostatic treatments may be impaired depending on the context of the extracellular matrix within the tumor environment and thus could impede the efficacy of angiostatic therapies.
- Vascular Endothelial Growth Factor
- Endothelial Cell Proliferation
- Endothelial Cell Apoptosis
- Hand Panel
- Human Dermal Microvascular Endothelial Cell
The growth of tumors beyond 1 mm3 is dependent on the induction of angiogenesis, defined as the growth of new blood vessels from preexisting vasculature . The process of angiogenesis is regulated by a number of promoters (angiogenic) and inhibitors (angiostatic), and it is the balance of expression of these opposing molecules that ultimately dictates whether or not angiogenesis proceeds. There are a number of endogenous angiostatic inhibitors that have been actively investigated including thrombospondin, interferon-inducible protein 10 (IP-10) and endostatin. Thrombospondin (TSP) was initially identified as a human platelet derived protein  that played a key role in platelet aggregation [3, 4]. It was subsequently shown to modulate the biological responses of endothelial cells [5, 6], induce apoptosis of endothelial cells via a caspase-3 dependent mechanism [7–9] and inhibit pathological angiogenesis [10–13]. IP-10 is a CXC chemokine that is secreted by a variety of different cell types in response to interferon stimulation . In addition to being a T-cell chemoattractant [15, 16], IP-10 has been shown to be an inhibitor of angiogenesis [17–19] and tumor growth in vivo [19–21]. Endostatin is the 20 kDa C-terminal cleavage product of collagen XVIII that has been shown to have a number of anti-angiogenic properties including inhibition of endothelial cell proliferation  and migration [23, 24], induction of endothelial cell apoptosis , and inhibition of tumor growth in vivo [22, 26–28]. Although these angiostatic molecules have been shown to be efficacious in pre-clinical models, their success in clinical trials has been more limited. Part of this may be due to the fact that very little is understood about the mechanisms by which these molecules exert their biological effects and how their efficacy might be altered by different tumor microenvironments.
The extracellular matrix (ECM) composition in tumors is vastly different than that found in its normal tissue counterparts. Certain ECM proteins such as collagen I, fibronectin and tenascin C are increased, while the basement membrane proteins collagen IV and laminin are decreased in neoplastic as compared to normal breast tissue [29, 30]. Previous data suggested that stimulation of human dermal microvascular endothelial cells (HDMEC) grown on collagen I with vascular endothelial growth factor (VEGF) resulted in the increased survival of these cells by a mechanism involving the upregulation of bcl-2 . It has also been shown that increased endothelial cell survival following overexpression of bcl-2 was associated with enhanced tumorigenesis in a xenograft model of human tumorigenesis . These observations suggested that increased endothelial cell survival might be modulated by extracellular matrix components such as collagen I, and may contribute to the progression of tumor growth and metastasis as a result of enhanced angiogenic potential within these tumors. Furthermore, this increased survival may render the endothelial cells more resistant to the inhibitory effects of certain angiostatic molecules, thus limiting their efficacy in certain tumor microenvironments. As collagen I is found to be overexpressed in tumor versus normal tissue of the breast [29, 30], and is itself a substrate for attachment of integrins that can profoundly influence endothelial function [33–37], we therefore examined the ability of three known angiostatic molecules, TSP-1, IP-10 and endostatin to inhibit endothelial cell proliferation, induce endothelial cell apoptosis on both plastic and collagen I coated surfaces in vitro and to inhibit VEGF-induced endothelial cell tube formation in collagen I gels in vitro. We found that although TSP-1 and IP-10 could inhibit endothelial cell proliferation and induce apoptosis when HDMEC were cultured on plastic tissue culture dishes, their ability to inhibit these processes was impaired following culture of HDMEC on collagen I. We observed similar results when cells were exposed to endostatin in that it inhibited proliferation of HDMEC cultured on plastic, but failed to inhibit proliferation of HDMEC grown on collagen I. In contrast to the results observed with TSP-1 and IP-10, we found that endostatin retained the ability to induce apoptosis of HDMEC even in the presence of collagen I. Furthermore, only endostatin could inhibit the formation of VEGF-induced sprouts on collagen I gels in vitro, while TSP-1 and IP-10 remained ineffective in this model system. These data suggest that the context of the tumor matrix microenvironment may modulate the inhibitory activity of angiostatic molecules and this may have significant impact on the ability of these molecules to function as anti-angiogenic therapeutics clinically in certain tumor types.
Growth on collagen I impairs the ability of TSP-1, IP-10 and endostatin to inhibit endothelial cell proliferation
Collagen I impairs endothelial cell apoptosis induced by TSP-1 and IP-10, but not by endostatin
The pro-apoptotic activity of endostatin is independent of bcl-2 expression
Endostatin-induced endothelial cell apoptosis is independent of caspase-3 activation
Endostatin induces the activation of caspase-8 in HDMEC grown on collagen I
Endostatin but not TSP-1 or IP-10 inhibits VEGF-induced HDMEC sprout-formation on collagen I gels
We had previously observed that VEGF enhanced survival of HDMEC grown on collagen I , suggesting that factors such as elevated collagen I levels within tumor microenvironments could increase endothelial cell survival and thus could potentially induce resistance to angiostatic molecules in vivo. We decided to formally examine this hypothesis and determined that growth on the tumor-associated matrices collagen I and tenascin C impaired the ability of TSP-1 and IP-10 to inhibit endothelial cell proliferation and induce endothelial cell apoptosis (in the case of collagen I). The ability of TSP-1 to inhibit endothelial cell proliferation has previously been suggested to be dependent on its ability to bind to heparin and compete with heparin-binding growth factors for binding sites on cells . As TSP-1 has been previously shown to be able to bind collagen I, it is quite possible that TSP-1 is sequestered from cells by binding collagen I, thus allowing the heparin-binding growth factors to stimulate the proliferation of HDMEC on collagen I . Thrombospondin has also been previously shown to induce apoptosis in endothelial cells via activation of caspase-3 [8, 9]. Surprisingly, although we did see a dose-dependent decrease in the levels of procaspase-3 following treatment of HDMEC on collagen I with TSP-1 in combination with VEGF, indicating activation of caspase-3, this was not associated with the induction of significant apoptosis as compared to stimulation with VEGF alone. Unlike other studies suggesting that TSP-1 induced endothelial cell apoptosis in a caspase-3 dependent manner [8, 9], our studies were performed in the presence of exogenous VEGF, thus although some processing of caspase-3 occurs in response to TSP-1 in our experimental conditions, it is not enough to overcome the survival effects induced by simultaneous exogenous VEGF stimulation. Other studies have implicated caspase-8, fas and fasL as the main mediators of TSP-1-induced apoptosis . We attempted to determine if caspase-8 was playing a role in TSP-1-induced apoptosis in our model system, however were unable to detect active caspase-8 using western blot assays. There are a number of differences between our study and those previously reported, including different endothelial cell types that are known to behave differently in experimental conditions [44–46] and different TSP-1 concentrations than those used in our studies. In fact, it was reported that when TSP-1 was delivered in combination with 50 ng/ml of VEGF, the cells became resistant to TSP-1 , thus confirming our observations.
We also observed that growth on collagen I prevented the angiostatic chemokine IP-10 from inhibiting endothelial cell proliferation and inducing endothelial cell apoptosis. The mechanisms of endothelial cell inhibition by IP-10 have not been extensively studied. Previous data has suggested that IP-10 can inhibit endothelial cell proliferation , however other reports suggest that IP-10 had no effect on endothelial cell growth, attachment or migration, but did however inhibit bFGF-induced tube formation in vitro and in vivo . More recently, using IP-10 mutants that have impaired binding domains, the angiostatic activity of IP-10 was shown to be dependent on its binding to the g-protein coupled receptor CXCR3 . It has been previously demonstrated that cell migration in response to SDF-1 via activation of CXCR4 is enhanced in the presence of fibronectin, suggesting that signaling through integrins can affect the downstream signal transduction pathway activated by the CXC receptors . To date, there are no studies implicating integrins in CXCR3 signaling, however the CXCR family of receptors are very homologous so it is highly possible that ECM binding through the integrins can modulate the response following CXCR3 activation. As very little is known about the mechanism by which IP-10 inhibits angiogenesis, further investigation into the mechanism of inhibition of endothelial cell proliferation and induction of apoptosis by IP-10, and the effects of tumor-associated extracellular matrices is warranted.
We also examined the ability of endostatin to inhibit endothelial cell proliferation following growth on plastic, collagen I, laminin or tenascin C. As seen with the other inhibitors, endostatin was not able to inhibit endothelial cell proliferation when HDMEC were cultured on collagen I or tenascin C as a substrate. Previous data suggested that inhibition of proliferation of endothelial cells is one of the primary mechanisms of its anti-angiogenic capabilities [22, 24]. Subsequently, endostatin has been shown to affect multiple facets of the angiogenic process including cell migration [23, 24], survival , protease activity [51, 52], and vessel stabilization [50, 53]. Discrepancies in the results obtained using endostatin can be observed depending on the source of endostatin used in the experiments, which can be either bacterially derived , purified from murine hemangioendothelioma cells , or derived from Pichia pastoris. We have used the latter source of endostatin for our studies, and it has previously been shown to inhibit endothelial cell proliferation and migration and to induce a G1 arrest in cells . Our observations support previous observations that endostatin can inhibit VEGF-induced endothelial cell proliferation to some extent; however, we have extended these findings to demonstrate that in the presence of the "tumor-associated" ECM proteins collagen I and tenascin C, this inhibition of proliferation by endostatin is dramatically impaired. The receptor that binds endostatin to inhibit angiogenic activities remains unclear despite numerous studies indicating its ability to bind cell surface molecules such as glypicans , heparin , tropomyosin-3 , VEGFR1 and VEGFR2 , and integrins αv and α5 . It has been previously shown that α5β1 integrins cluster following endostatin binding , and since collagen I and tenascin C can both bind β1 integrins (in association with various α integrin chains) it is highly possible that binding of these matrices by integrins could modulate either the affinity for endostatin binding or integrin signal transduction cascades through an inside-out signal . Furthermore, a peptide derived from endostatin has recently been shown to be a potent inhibitor of angiogenesis in a β1 integrin and heparin-dependent manner , suggesting that perhaps collagen I and tenascin C compete with endostatin for integrin-binding on cells and supporting our observations that endostatin was ineffective at inhibiting endothelial cell proliferation following growth on collagen I or tenascin C.
In contrast to our results with endothelial cell proliferation, we found that endostatin could induce apoptosis of endothelial cells regardless of whether they were cultured on plastic or collagen I substrates. In fact, we observed endothelial cell apoptosis at doses 100-fold lower than has previously been described . The previous studies indicated that endostatin's ability to induce apoptosis was due to its ability to decrease bcl-2 expression and induce activation of caspase-3 . In contrast to these results, we did not observe either a decrease in bcl-2 expression nor activation of caspase-3 in HDMEC following endostatin stimulation in either the presence or absence of VEGF. In these studies, bovine pulmonary artery endothelial cells were used, thus differences observed between the doses of endostatin required to induce apoptosis could again be a reflection of differences in the behavior of different endothelial cell types.
As endostatin has been previously shown to bind to a number of different integrin molecules , and recently it has been demonstrated that the inappropriate ligation of integrins can lead to what has been referred to as integrin-mediated death , it is possible that endostatin may induce apoptosis via alternative mechanisms such as integrin-mediated death. This form of apoptosis was shown to be dependent on caspase-8, and our observation of activation of caspase-8 observed at the higher doses of endostatin supports the notion that the apoptosis observed in our system following treatment with endostatin may be related to integrin-mediated death. Recently, it has also been shown that endostatin inhibited the migration on and attachment of endothelial cells to collagen I, but did not affect the proliferation of endothelial cells cultured on this matrix , which is in complete agreement with our results. Other evidence would also suggest that the main mechanism of inhibition of angiogenesis by endostatin is via its ability to inhibit cell migration through α5β1 integrin, heparan sulfate, and lipid raft-mediated interactions , supporting our observations regarding endostatin's ability to inhibit vessel formation in our tube formation assays, where endothelial cell migration is a prerequisite for structure formation, and our results would suggest that this effect is independent of signals from collagen I.
In conclusion, we have demonstrated that TSP, IP-10 and endostatin mediate inhibition of angiogenesis via different mechanisms that are affected to different degrees by growth of the endothelial cells on substrates such as collagen I. Our data suggests that the biological effects that various angiostatic molecules have on endothelial cells may be affected by the type of extracellular matrix upon which the cells are in contact. Indeed, the importance of cues from the extracellular matrix in vessel survival are also evident from transgenic animal systems where targeted deletion of the collagen I gene in mice led to embryonic death, due to rupture of blood vessels . The effects of ECM on vessel survival and angiogenesis is extremely important in the context of inhibition of angiogenesis as an anti-tumor therapy as tumors usually have extensively remodeled matrix with differences in the composition of this matrix as compared to that found in normal tissues. Given the differences in the human tumor matrix microenvironment, and our results that "tumor-associated" matrices such as collagen I and tenascin C may enhance the resistance of endothelial cells to certain anti-angiogenic agents, it is imperative that we gain a clearer understanding of the mechanism of inhibition of angiogenesis by these molecules and how this may be affected by differences in the tumor microenvironment. These insights will potentially help to predict patient response to these inhibitors and elucidate targets of intervention that will be unaffected by differences in the tumor matrix microenvironment.
Endothelial cells and antibody reagents
Human dermal microvascular endothelial cells (HDMEC) derived from neonatal tissue were obtained from Cambrex Corporation (Walkersville, MD) and were propagated in EGM-2MV media (Cambrex Corp., Walkersville, MD). All experiments were performed using cells between passages 4 and 10. Rabbit anti-Bax and goat anti-Caspase-3 antibodies were from R&D Systems (Minneapolis, MN). Hamster anti-Bcl-2 antibody was purchased from BD Pharmingen (Mississauga, ON) and rabbit anti-active caspase-3 antibody was from Biovision (Palo Alto, CA). Mouse anti-caspase-8 antibody was purchased Cell Signaling Technologies (Beverly, MA).
Coating tissue culture dishes with collagen
Vitrogen 100 bovine dermal collagen (Cohesion Technologies, Palo Alto, CA) was used to coat tissue culture vessels in all experiments, and is a mixture of ~97% collagen I and ~3% collagen III matrices. The acidified collagen solution was kept on ice, diluted to a concentration of 1.5 mg/ml, and neutralized following addition of 10× PBS and 0.1 N NaOH to a pH of approximately 7.4. The appropriate volume of collagen solution was added to coat each vessel and the plates were rocked to ensure even distribution of collagen across the surface. Plates were then incubated at 37°C for 4 h to overnight to allow gelation to occur. The collagen surfaces were washed with Hanks Buffered Salt Solution (HBSS, Invitrogen, Carlsbad, CA), and incubated in EGM-2MV for a minimum of 2 h to equilibrate the collagen prior to addition of endothelial cells. For coating with laminin, human placenta laminin (Sigma, Oakville, ON) was diluted in PBS and added to dishes at a concentration of 1 μg/cm2. The dishes were allowed to dry overnight, uncovered, in a laminar-flow hood and cells were seeded on the surfaces the next day. For tenascin C, dishes were coated with tenascin C purified from a human tumor cell line (Chemicon International, Temecula, CA). For coating, tenascin-C was diluted to 0.1 μg/ml in PBS, and added to dishes which were then incubated overnight at 4°C. The next day, the solution was aspirated, and the dishes were blocked with 0.1% casein solution for at least 1 h. The dishes were then washed three times with 1× PBS, and seeded with cells.
HDMEC were seeded at a density of 15,000 cells per well in EGM-2MV into untreated or matrix coated 12-well plates as described above and allowed to incubate overnight. The next day, media was removed, cells were washed twice with HBSS and then starved overnight in MCDB 131 (Invitrogen, Carlsbad, CA) containing 1% fetal bovine serum (FBS). The following day cells were stimulated with varying concentrations of either TSP-1 (Calbiochem, San Diego, CA), IP-10 (Intergen, Purchase, NY), or endostatin (Calbiochem, San Diego, CA) together with 50 ng/ml VEGF (kind gift from the Biological Resources Branch of the National Cancer Institute) in MCDB 131 containing 5% FBS. Stimulation with 50 ng/ml VEGF in MCDB 131 containing 5% FBS was used as a positive control for proliferation, and stimulation with MCDB 131 containing only 1% FBS was used as a negative control for proliferation. Following 72 h of incubation, cells were collected by trypsinization or by collagenase treatment in the case of cells seeded on collagen substrate, and the number of cells per unit volume in each well was determined following counting in a Coulter counter. Each treatment was performed in triplicate and the entire experiment was performed a minimum of three independent times.
BrdU Cell Proliferation ELISA
HDMEC were seeded at a density of 2,500 cells per well in EGM-2MV into untreated or collagen I coated 96-well plates as described above and allowed to incubate overnight. The next day, media was removed, cells were washed twice with HBSS and then starved overnight in MCDB 131 containing 1% FBS. The following day cells were stimulated with varying concentrations of either TSP-1, IP-10, or endostatin together with 50 ng/ml VEGF in MCDB 131 containing 5% FBS. BrdU labeling solution was subsequently added 48 h post-stimulation with growth factors and inhibitors, and cells were incubated overnight. Incorporated BrdU was subsequently detected using the BrdU Cell Proliferation ELISA, kit according to the manufacturers directions (Roche, Laval, PQ), using a Fluoroskan Ascent Plate reader (Thermolabs, Franklin, MA) for luminescent detection.
Determination of apoptosis by FACS
HDMEC were seeded into untreated or collagen I coated dishes and allowed to adhere overnight. The following day, monolayers were washed with HBSS to remove non-adherent cells, and adherent cells were stimulated with MCDB 131 media containing 5% FBS (unstimulated), or MCDB 131 media with 5% FBS supplemented with 50 ng/ml VEGF alone, 10, 50 or 250 ng/ml TSP-1, 16, 80 or 400 ng/ml IP-10, or 100, 500 or 2500 ng/ml endostatin alone or in combination with 50 ng/ml VEGF. Cells were incubated for 60 h, and then harvested by collecting the non-adherent and adherent cell populations either by trypsinization or collagenase treatment for plastic and collagen I coated dishes respectively. Collected cells were pelleted by centrifugation at 300 × g, washed twice with PBS, and then resuspended in ice-cold 70% ethanol. Cell suspensions were incubated in 70% ethanol at -20°C for a minimum of 24 h. Following permeabilization, cells were washed two times with PBS, and then resuspended in 500 ul of propidium iodide solution (48 ug/ml propidium iodide, 40 ug/ml Rnase A in PBS). The percentage of apoptotic cells was determined following identification of the sub-G1 population of cells by flow cytometric analysis.
Detection of caspase-3 or caspase-8 activity
HDMEC were seeded into untreated or collagen I coated dishes and allowed to adhere overnight. The following day, monolayers were washed with HBSS to remove non-adherent cells, and cells were stimulated with EGM-2MV media alone, 50 ng/ml VEGF alone, or 50 ng/ml TSP-1, 80 ng/ml IP-10, or 500 ng/ml endostatin alone or in combination with 50 ng/ml VEGF in EGM-2MV. Cells were incubated for 48 h, and then harvested by collecting the non-adherent and adherent cell population by either trypsinization or collagenase treatment for plastic and collagen I coated dishes respectively. Collected cells were pelleted by centrifugation at 300 × g, washed with PBS, counted and then stored at -80°C. At time of assay, cells were lysed in the appropriate volume of lysis buffer (10 mM HEPES, 1 mM EDTA, 100 mM NaCl, 5 mM MgCl, 142.5 mM KCl, and 1 mM DTT) so that equal cell number/unit volume was obtained for each sample. Samples were incubated on ice for 45 minutes followed by a centrifugation at 13,000 rpm at 4°C for 30 minutes in a microfuge. Supernatants were removed and 20 ul of each sample was added to assay buffer (50 mM HEPES, 1 mM EDTA, 100 mM NaCl, 10% glycerol, 0.1% CHAPS, 10 mM DTT, pH 7.4) in a 96-well microtiter plate. Plates were allowed to equilibrate at 37°C for 10 minutes, and then 10 ul of a 1 mM stock of the fluorogenic caspase-3 substrate Ac-DEVD-AMC (Alexis Biochemicals, San Diego, CA) was added to each well. Plates were incubated at 37°C to allow the enzymatic reaction to proceed, and the fluorescence was measured at various times post-addition of substrate using a fluorescent plate reader set at 460 nm emission/360 nm excitation wavelengths. Purified caspase-3 was used as a positive control in all assays, and all samples were assayed in triplicate. Experiments were performed a minimum of three independent times.
For detection of caspase-8 activity, cells were seeded into dishes and stimulated as described above. Following incubation for 48 h, cells were isolated following trypsinization, washed once with PBS, and cell pellets were stored at -80°C. Subsequently, the pellets were lysed in provided caspase activity lysis buffer according to the manufacturer's directions (Sigma, Oakville, ON), the protein concentration for each lysate was determined, and caspase-8 activity was determined according to the manufacturer, using a fluorogenic caspase-8 activity kit (Sigma, Oakville, ON) that used Ac-IETD-AMC as a substrate and fluorometric detection in a fluorescent plate reader set at 460 nm emission/360 nm excitation wavelengths.
Western blot analysis
HDMEC were seeded on plastic or collagen I coated dishes as described above and allowed to adhere overnight. The next day, monolayers were washed twice with HBSS and then stimulated with varying concentrations of TSP-1, IP-10 or endostatin alone or in combination with 50 ng/ml VEGF. Total protein lysates were generated in boiling lysis buffer (1% SDS, 1.0 mM sodium ortho-vanadate, 10 mM Tris pH 7.4, 0.2 mM PMSF, 2 μg/ml aprotinin) following recovery of cells from dishes by trypsinization or collagenase treatment for growth on plastic or collagen I respectively. Aliquots containing 5–40 μg of total protein were subjected to SDS-PAGE electrophoresis followed by transfer to nitrocellulose membranes. Specific proteins were detected following incubation with primary and horse-radish peroxidase-conjugated secondary antibodies and visualization using chemiluminescent detection (Supersignal, Pierce, Rockford, MD).
Endothelial cell sprouting assays
Collagen I gels were prepared as described above. 2 × 105 HDMEC were seeded onto each 60 mm dish of collagen I and allowed to adhere overnight. The following day the dishes were washed twice with HBSS and then stimulated with EGM-2MV alone or supplemented with 50 ng/ml VEGF alone or in combination with 50 ng/ml TSP, 80 ng/ml IP-10 or 500 ng/ml endostatin. Cells were counted on day 0 prior to stimulation to ensure similar numbers of cells had been seeded on each dish, and were then counted daily for 12 days. Media containing supplements was replaced every 48 hours.
We would like to thank Diana Malenica for technical assistance. This work has been supported in part by the Canadian Institute of Health Research (MOP-53076, C.L.A.) and by the National Institutes of Health/National Institute of Dental and Craniofacial Research (DE13161, P.J.P.). C.L.A. is the recipient of a CIHR New Investigator Award.
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