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CELLULAR AND MOLECULAR
Laboratoire d'Organogénèse Expérimentale, Hôpital du Saint-Sacrement, Département de Chirurgie, Faculté de Médecine, Université Laval, Québec, Canada
Received for publication
May 16, 2005
Accepted
July 26, 2005.
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Abstract |
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). Angiogenesis leads to activation of endothelial cells, which degrade their basement membrane and surrounding extracellular matrix to migrate through the tissue and proliferate (Wilting and Christ, 1996; Risau, 1997). In adults, angiogenesis is mainly induced after a trauma, such as a wound or an ischemia, or during pathological processes such as retinopathy, leukemia, rheumatoid arthritis, diabetic retinopathy, fibrosis, psoriasis, ulcer, and tumor growth (Carmeliet and Jain, 2000; Folkman, 2001). A new therapeutic approach was developed to inhibit the development of solid tumors by blocking their vascularization with angiostatic drugs (Carmeliet, 2003). The angiostatic strategy is now the most promising way to inhibit angiogenesis-dependent diseases, although only few drugs have yet been approved for clinical application [Avastin, an anti-VEGF monoclonal antibody, was approved in 2004 (Kabbinavar et al., 2005)]. Several angiogenesis models have been developed to investigate the mechanisms of angiogenesis and to allow a better targeting of angiostatic drugs in a preclinical step (Cockerill et al., 1995). Various animal models were produced, such as the chick chorioallantoic membrane, the rabbit cornea assay, the sponge implant models, the Matrigel plug, and conventional tumor models (Auerbach et al., 2003). Nevertheless, these models remain extremely complex, relatively expensive, and difficult to interpret for routine screening. Thus, in recent years, a number of in vitro models have been developed. The most common in vitro models are the three-dimensional gel assays made of various extracellular matrix components such as type I collagen, fibrin, fibronectin, basement membrane extracts, or Matrigel, in which endothelial cells are embedded (Auerbach et al., 2003; Ley et al., 2004; Robinet et al., 2005). These in vitro models reconstitute an environment made of only one type of extracellular matrix molecule (except for Matrigel) with one type of cell, the endothelial cells, in contrast with the physiological environment, which features several different molecules and cells that are known to be essential to modulate neovascularization in vivo and in vitro (Vernon and Sage, 1999). In addition, most of these models are produced with animal endothelial cells (Vernon and Sage, 1999; Auerbach et al., 2003), which have a much lower growth factor requirement to survive, proliferate, and form tubular structures (Cavallaro et al., 2001). Moreover, many in vitro angiogenesis models achieve the formation of cord-like structures instead of capillary-like ones (Zimrin et al., 1995) and can present a basement membrane on the lumen side (Sage and Vernon, 1994).
We developed a model of endothelialized reconstructed dermis that promotes the spontaneous formation of a capillary-like network by coculture of human umbilical vein endothelial cells (HUVEC) with dermal fibroblasts in a collagen biopolymer (Black et al., 1998; Hudon et al., 2003). In contrast to most in vitro angiogenesis models produced with human endothelial cells (Cockerill et al., 1995), this angiogenesis process takes place without the addition of any exogenous modulator-like growth factors (except during the first 10 days of culture) or tumor-promoting agents. This phenomenon has been attributed to the synthesis of angiogenic molecules by endothelial cells and fibroblasts and the presence of a complex extracellular matrix synthesized by the fibroblasts (Berthod et al., 1996). We previously demonstrated that capillary-like structures (CLS) were only composed of endothelial cells that deposited basement membrane proteins (laminin and type IV collagen) on their basal side. These tubular structures featured a lumen the entire length of the CLS and were organized in a network of branching tubes that reached the bottom-most portion of the dermis (Hudon et al., 2003). In this study, we demonstrated that the ERCT, made of HUVEC or human microvascular endothelial cells (HMVEC), was a powerful model to finely analyze, quantitatively and qualitatively, the influence of angiostatic drugs on angiogenesis.
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Materials and Methods |
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HUVEC were obtained from healthy newborns by enzymatic digestion with 0.250 µg/ml thermolysin (Sigma-Aldrich Canada). Veins were cannulated at both ends and washed with calcium-free HEPES solution (Valeant Canada Limited, St-Laurent, QC, Canada). A thermolysin solution was then injected to rinse and fill the vein, and the cord was placed in calcium-free HEPES solution at 37°C. After 30-min incubation, the veins were perfused with M199 medium (Sigma-Aldrich Canada) containing 10% FetalClone II serum (HyClone) and antibiotics (100 U/ml penicillin G and 25 µg/ml gentamicin) (Black et al., 1998).
HUVEC were centrifuged and resuspended in M199 medium supplemented with 20% FetalClone II serum (HyClone), 2.28 mM glutamine, 40 IU/ml heparin (Leo Laboratories, Pickering, ON, Canada), 20 µg/ml endothelial cell growth supplement (Sigma-Aldrich Canada), and antibiotics (100 U/ml penicillin G and 25 µg/ml gentamicin). HUVEC were plated on gelatin-coated tissue culture flasks, and characterization was assessed as previously described (Black et al., 1998).
HMVEC were purified from the foreskin of a healthy baby using Dynabeads CD31 (Dynal Biotech, Lake Success, NY) (Richard et al., 1998). Briefly, pieces of skin tissue were incubated overnight in a Hanks' balanced salt solution (Sigma-Aldrich Canada) containing 2% fetal calf serum (HyClone) and 1% dispase II (Roche Diagnostics Canada). The epidermis was removed, and endothelial cells were extruded by pressing the dermal layer. Mixed cells were centrifuged, resuspended in EGM-2 medium (Cambrex Bio Science Baltimore, Inc., Baltimore, MD), and then plated on gelatin-coated tissue culture flasks for 1 week. Cells were then removed from the culture flask by trypsin/EDTA (Valeant Canada Limited) and incubated with Dynabeads CD31 for 30 min. HMVEC characterization was assessed as previously described (L'Heureux et al., 1993).
Collagen-Glycosaminoglycan-Chitosan Biopolymer Preparation. Collagen-glycosaminoglycan-chitosan biopolymers were prepared using a technique previously described (Berthod et al., 1993). These sponges were prepared with types I and III bovine collagen (Laboratoire Perouse Implant, Chaponost, France), chitosan (95% deacetylated; SADUC, Lyon, France), and chondroitin 4 to 6 sulfates (SADUC), which were dissolved in 0.1% acetic acid. After mixing, 1 ml/well (3.8 cm2) of the final solution was poured into 12-well plates (Becton Dickinson, Toronto, ON, Canada) and frozen overnight at -80°C. The frozen plates were then lyophilized in a Genesis 12EL vacuum freeze dryer (Virtis, Gardiner, NY).
Endothelialized Reconstructed Dermis Preparation. The ERCT was prepared with a suspension of 1:1 ratio of human fibroblasts and HUVEC or HMVEC. A volume of 3 ml was seeded on the top of a chitosan-glycosaminoglycan-collagen sponge at a final concentration of 2.1 x 105 cells/cm2 (Black et al., 1998). The cells were cultured under submerged conditions and fed with 3 ml of EGM-2 medium (Cambrex Bio Science Baltimore, Inc.) supplemented with 50 µg/ml ascorbic acid (Sigma-Aldrich Canada). After 10 days, ERCT were cultured in a Dulbecco-Vogt Eagle's modified medium-Ham's F-12 medium (Invitrogen) supplemented with 10% fetal bovine serum, 0.4 µg/ml hydrocortisone (Calbiochem, La Jolla, CA), 5 µg/ml bovine insulin (Sigma-Aldrich Canada), 100 µg/ml ascorbic acid, and antibiotics. Seven days later, the ERCT were elevated at the air-liquid interface and cultured in the same medium for 2 additional weeks.
ERCT Treatment with Drugs. The angiogenesis-inducing agent vascular endothelial growth factor (VEGF; Sigma-Aldrich Canada) or inhibiting agents tamoxifen (Sigma-Aldrich Canada), ilomastat (Chemicon, Temecula, CA), and echistatin (Sigma-Aldrich Canada) were added three times in the medium for a total period of 7 days. In the first experiment, drugs were added between days 17 and 24 (three successive doses on days 17, 19, and 22) at the following concentrations: VEGF (15, 30, 45, and 60 ng/ml), tamoxifen (10, 20, 30, and 40 µM), ilomastat (12.5, 25, 50, and 75 µM), and echistatin (0.03, 0.06, 0.09, and 0.12 µM) to assess their dose-response effect. In a second experiment, a unique concentration of VEGF (15 ng/ml), tamoxifen (40 µM), ilomastat (50 µM), and echistatin (0.06 µM) was added between days 10 and 17 to analyze the long-term effect of each drug on HUVEC-ERCT, compared with HMVEC-ERCT. The analyses were performed on biopsies taken after 10, 17, 24, and 31 days of treatment. Results obtained with ilomastat were compared with a control treated with 30 µl/ml DMSO.
ERCT Treated under a Condition of Hypoxia. HUVEC-ERCT were cultured for 24 days and then incubated 16 h in an anaerobic jar (VWR, Montreal, PQ, Canada) containing a disposable carbon dioxide/hydrogen gas-generating system (Anaerocult A; VWR), which induced a deprivation of oxygen by the supplementation of atmosphere with CO2. The total deprivation of oxygen was assessed with an Anaerotest test strip (VWR). The histological analyses were performed on ERCT biopsies taken 1, 4, 7, and 10 days after the 16-h hypoxia treatment.
Immunohistochemistry. Frozen 4-µm sections were incubated in a blocking solution containing phosphate-buffered saline and 1% (w/v) bovine serum albumin (Sigma-Aldrich Canada). Fluorescent staining was performed by subsequent incubations with a primary mouse monoclonal anti-human platelet-endothelial cellular adhesion molecule-1 (PECAM-1) antibody (Chemicon) and a rhodamine-conjugated goat anti-mouse IgG-IgM (Chemicon) secondary antibody, mixed with Hoechst (1/100) (Sigma-Aldrich Canada) to stain the cell nuclei. Sections were examined using a Nikon Eclipse E600 fluorescence microscope (Nikon, Melville, NY).
Quantification and Evaluation of Areas Containing Capillary-like Structures. Samples were fixed with Histochoice's solution (Armesco, Solon, OH), dehydrated, and embedded in paraffin. Six microns sections were cut, stained with Masson's trichrome, and visualized on a Nikon Eclipse E600 fluorescence microscope. The area of each CLS was calculated on digital microphotographies using Metaview software (Molecular Devices, Sunnyvale, CA). The number of CLS was measured with the NIH image version 1.62 software (National Institutes of Health, Bethesda, MD). Four samples (with the mean of two slides per sample) were analyzed for each group for both the number and surface area of CLS. The data obtained were analyzed by the two-way analysis of variance test, and p < 0.05 was considered as significant.
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Results |
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When the ERCT was treated with VEGF (15 to 60 ng/ml) (Fig. 3), a significant increase of the number of CLS was observed (*p < 0.05, **p < 0.005; n = 4), compared with the control, after drug removal at 45 ng/ml. Seven days after drug removal, this effect was also observed at 30 ng/ml.
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When echistatin (0.03 to 0.12 µM) (Fig. 3) was added to the culture medium, a diminution of the number of CLS at 0.06, 0.09, and 0.12 µM was observed, although this effect was only maintained at the 0.12 µM concentration during the 7 days following the removal of the drug. Thus, echistatin has a moderate angiostatic effect early after its addition, but this effect was rapidly reversible after drug removal.
The addition of ilomastat (12.5 to 75 µM) (Fig. 3) induced a significant decrease in the number of CLS at 50 (p < 0.05) and 75 µM (p < 0.005) after drug removal. This angiostatic effect was observed at all concentrations 7 days after drug removal (p < 0.005). At 75 µM, a 4.5-times decrease in the number of CLS was observed compared with the control. This inhibition of angiogenesis was abolished after drug removal, since at day 31, more CLS were observed at all concentrations compared with day 24. Ilomastat induced a very robust and very precocious inhibitory effect on CLS formation, since at 75 µM nearly no additional CLS was formed following the addition of the drug (the number of CLS at day 17 and day 24 was similar at this concentration; data not shown).
The ERCT allowed us to analyze the modulation of hypoxia on CLS formation. The in vitro model was placed for 16 h in an anaerobic jar, and histological analyses were performed on biopsies taken 1, 4, 7, and 10 days after the treatment. One week after hypoxia (Fig. 4, b–d), no significant difference in the number of CLS (Fig. 4c) was detected compared with control (Fig. 4c), but the number of very large CLS (over 300 µm2) was elevated 10 times compared with the control (*p < 0.05; n = 3) (Fig. 4d).
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Discussion |
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VEGF, which is known to have a strong angiogenic effect (Ferrara, 2004), induced an augmentation in the number of CLS in the ERCT at 45 ng/ml after the end of a 7-day treatment. Seven days after the removal of VEGF, a significant increase was observed at both 30 (p < 0.05) and 45 ng/ml (0.005). The 30-ng/ml concentration needed a 1-week time period after treatment to induce an increase in the CLS number. This showed that the induction of the VEGF-mediated angiogenesis was a slow process at lower concentrations. However, at 60 ng/ml, the angiogenic drug did not show a significant augmentation. This could be explained by an increased competition of the molecule with its receptor and by the fact that VEGF is also known to affect the intussusception (nonsprouting angiogenesis) (Ferrara, 2001).
Tamoxifen was shown to have concurrent pro- and antiangiogenic actions when it interacts with estrogen receptors (Lee et al., 2004). However, this mediation through the estrogen receptor can not account for all of the tamoxifen effects (Reddel et al., 1985). Tamoxifen is also known to inhibit angiogenesis in vitro and in vivo by the inhibition of endothelial cell proliferation (Gagliardi et al., 1996). This effect could be induced by blocking ornithine decarboxylase, cholesterol synthesis, microsomal lipid peroxidation, or cyclins, but predominantly by blocking the calcium-dependent protein kinase C (Gagliardi et al., 1996). In contrast, tamoxifen demonstrates an angiogenic effect by stimulating the expression of VEGF (Hyder et al., 1997), transforming growth factor-
(Grainger et al., 1993), and by down-regulating CD36 expression (Silva et al., 1997).
Our model depicted these complex processes very well. In the ERCT, an angiogenic effect of tamoxifen was observed at 10 µM, which could be explained at least in part by the stimulation of VEGF expression by fibroblasts. A concentration-dependent antiangiogenic effect was detected starting at 20 µM. This effect became statistically significant 1 week after the drug removal at 30 µM and was clearly cytotoxic at 40 µM. The lag time needed before seeing a decrease in the CLS number could be explained by the delay necessary for tamoxifen metabolites to reach the critical concentration for inducing an irreversible modification of protein kinase C (McNamara et al., 2001). However, this decrease should not be specific to an angiogenesis inhibition since the total number of cells seemed to be notably decreased (see number of blue nuclei in Fig. 1i compared with Fig. 1a).
Echistatin is well known to inhibit adhesion, migration, and survival of human umbilical cord vein endothelial cells (Juliano et al., 1996). This effect is induced by its binding to 
v3 and 51, two integrins that participate together in different angiogenic processes like proliferation, migration, and survival (Alimenti et al., 2004), on endothelial cells. Echistatin induces a considerable dose-dependent decrease in the number of CLS, as observed in the literature with other antagonists of integrin v3 (Brooks et al., 1994). This inhibition effect was still detectable at the highest dose 7 days after drug removal but was abolished at 0.06 and at 0.09 µM. This prolonged effect at 0.12 µM could be explained by the noncovalent irreversible binding of echistatin to v3, which ultimately induces apoptosis in endothelial cells (Alimenti et al., 2004). At lower doses, only a transient effect was observed and is most likely due to internalization of blocked integrins, which never reached the critical level that activate apoptosis.
Ilomastat is known to inhibit angiogenesis by blocking some matrix metalloproteinases (MMP) such as the gelatinases MMP-2 and MMP-9 (Galardy et al., 1994). Both are specialized in the degradation of different basement membrane components and seem to modulate endothelial cell attachment, proliferation, and migration during angiogenesis (Stetler-Stevenson, 1999). The angiostatic effect obtained with ilomastat hinged highly upon the time it was added to the ERCT culture. If added between days 10 and 17, ilomastat had no clear inhibitory effect on CLS formation (Fig. 2). In contrast, ilomastat added to the ERCT culture between days 17 and 24 induced a dose-response diminution in the number of CLS at all concentrations 7 days after drug removal, as observed in other angiogenesis models (Galardy et al., 1994). This inhibition of CLS formation was drastic (3 times less capillaries compared with control at 75 µM) and took effect rapidly after drug addition in the culture medium, since the effect was already achieved at the end of the drug incubation period. However, 1 week after the drug removal, a 2-fold increase of the number of CLS was observed at 75 µM, indicating that the inhibitory effect of the drug was abolished immediately after drug removal, as compared with the 2-fold increase in CLS number in controls during the same period. Indeed, ilomastat has a half-life of 13 min and necessitates a continuous treatment to maintain its inhibitory effect on the MMP synthesis or secretion (Galardy et al., 1994). Since the addition of ilomastat between days 10 and 17, which is a period of low CLS formation, did not induced a significant decrease in CLS formation compared with its addition between days 17 and 24, this result highlights the critical importance on the kinetic timing of drug addition during the CLS formation. Finally, ilomastat appeared to be the most powerful angiostatic drug since it rapidly and completely inhibits CLS formation at 75 µM, while at the same time it did not seem to induce a toxic effect on cells.
Since the local oxygen tension is well known to have a profound effect on the vasculature by compensating the vascular insufficiency through the induction of angiogenesis (Knighton et al., 1983), we investigated the modulation of hypoxia on the CLS in the ERCT. We demonstrated that hypoxia can modulate the shape of CLS formed in the ERCT. We observed a 10-fold increase in the formation of CLS larger than 300 µm in diameter, although the total number of CLS remained unchanged. This effect could be explain by a hypoxia-mediated VEGF expression, which might induce nonsprouting angiogenesis (Risau, 1997). An increase in the capillary diameter of cardiac tissue has also been described under conditions of oxygen deprivation in fetal sheep (Martin et al., 1998). Thus, the ERCT allowed us to detect an unexpected modulation of the capillary-like network by hypoxia, which gives evidence of the powerfulness of this model in discriminating subtle modulations of different parameters in the angiogenesis process.
Most in vitro studies performed on endothelial cell activation use cells isolated from human umbilical cord veins. However, it is well known in the literature that endothelial cells are greatly heterogeneous, depending on the tissue from which they originated (Garlanda and Dejana, 1997). To assess whether similar results could be obtained with microvascular endothelial cells cultured in our model, we have compared the effect of the drugs on CLS produced with endothelial cells isolated from large vessel (HUVEC) versus CLS made of endothelial cells from microvascular endothelium (HMVEC). The two endothelial cell types showed similar results except in the case of ilomastat, where the HMVEC-ERCT featured 2 times less CLS number compared with the HUVEC-ERCT (Figs. 1 and 2).
By using three angiostatic compounds which inhibited three major components of the angiogenic process, i.e., endothelial cell proliferation (tamoxifen), endothelial cell migration and apoptosis control via integrins (echistatin), and MMP function (ilomastat), we demonstrated the strength of our model to accurately discriminate the angiostatic potential of drugs. One of the major advantages of the ERCT is that it can be maintained in culture for a long period (over 30 days), in contrast with other models in which CLS are produced in a few hours in response to a boost of growth factors and are degenerated in less than 72 h (Troyanovsky et al., 2001). With the ERCT, the moment and the duration of the treatment can be chosen over at least a 31-day period to observe an inhibition in the CLS development (earlier treatment) or a regression of pre-existing vessels (later treatment). In addition, this culture period gives the possibility to analyze the long-term effect of the drugs on angiogenesis, as we have demonstrated that some molecules (such as echistatin and ilomastat) can induce a rapid but transient decrease on CLS formation, whereas others have a delayed inhibitory effect on angiogenesis, as is the case with tamoxifen. A toxic effect of the molecule can also be suspected by observation of histological tissue morphology and, in turn, quantified by cell extraction and counting or by the use of viability tests such as a 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium test. Finally, the analysis of CLS morphology also allows the control of the CLS size.
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Footnotes |
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Article, publication date, and citation information can be found at http://jpet.aspetjournals.org.
ABBREVIATIONS: VEGF, vascular endothelial growth factor; HUVEC, human umbilical vein endothelial cell(s); CLS, capillary-like structure(s); ERCT, endothelialized reconstructed connective tissue; HMVEC, human microvascular endothelial cell(s); PECAM-1, platelet-endothelial cellular adhesion molecule-1; MMP, matrix metallo proteinase.
Address correspondence to: Dr. François A. Auger, Laboratoire d'Organogénèse Expérimentale, Hôpital du Saint-Sacrement, 1050 chemin Sainte-Foy, Québec, QC Canada G1S 4L8. E-mail: francois.auger{at}chg.ulaval.ca
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