Integrin signaling is critical for pathological angiogenesis

Previous studies using cell lines and platelets demonstrated that phosphorylation of β₃ integrin cytoplasmic tyrosines is involved in outside-in integrin signaling. Therefore, we assessed whether EC adhesion to extracellular matrix proteins could induce tyrosine phosphorylation of β₃ integrin. We isolated ECs from the lungs of WT and DiYF mice, plated them on various substrates, and assessed the phosphorylation of β₃ integrin on Tyr747 and Tyr459 (Fig. 1, A and B). Very low levels of phosphorylation of WT β₃ integrin were observed when cells were kept in suspension or plated on poly-l-lysine. Extracellular matrix proteins that serve as ligands for α_vβ₃ such as vitronectin, fibronectin, fibrinogen, and gelatin strongly stimulated Tyr759 and Tyr747 phosphorylation of WT ECs. Laminin and collagen, which are recognized primarily by integrins other than α_vβ₃, induced lower levels of tyrosine phosphorylation. Sodium pervanadate, known to block phosphatase activity, was used as a positive control in this experiment. As anticipated, no tyrosine phosphorylation of β₃ integrin was observed in DiYF EC under any conditions.

The role of the β₃ subunit phosphorylation in extracellular matrix recognition was previously assessed in CHO cells and K562 cells. These systems do not provide insight into the roles of β₃ integrin in angiogenesis and in the regulation of VEGF or FGF-induced EC responses. These processes can be only assessed in ECs expressing appropriate receptors and signaling intermediates. We sought to determine whether VEGF treatment affected the phosphorylation status of β₃ integrin. Treatment of ECs with VEGF-A165 induced both Tyr747 and Tyr759 phosphorylation in a time-dependent manner (Fig. 1 C). Both time curves of β₃ tyrosine phosphorylation followed a bell-shaped pattern, which is typical for growth factor–induced responses. Thus, not only integrin engagement, but also VEGF treatment stimulated tyrosine phosphorylation of β₃ subunit in ECs, suggesting a possible regulatory role for this process in VEGF-induced angiogenesis.

We next evaluated whether β₃ integrin cytoplasmic tyrosine motifs are crucial for a complete angiogenic response to VEGF in vivo. We implanted VEGF-A–containing Matrigel subcutaneously into WT and DiYF mice and assessed the angiogenic response based on the amount of hemoglobin extracted from Matrigel. As shown in Fig. 2 A, the hemoglobin concentration was at least fivefold lower in Matrigel plugs isolated from DiYF mice compared with WT counterparts. The vascular density in Matrigel implants was assessed by von Willebrand factor (vWF) staining. Only 50% of Matrigel plugs from DiYF mice had a distinguishable vasculature and exhibited staining for vWF (Fig. 2 B, top). The vascular density was 4.2-fold lower in DiYF mice than in WT controls (P = 0.01) (Fig. 2 B, bottom). Thus, VEGF-induced angiogenesis was significantly impaired in DiYF mice.

Further, we assessed tumor-induced pathological angiogenesis in DiYF and WT mice. To this end, mouse melanoma cells (B16F10) were implanted subcutaneously into mice, and after 10 d tumors were excised. The average weight of tumors formed in DiYF mice was at least twofold lower than average tumor weight in WT mice (0.14 g vs. 0.29 g) (Fig. 2 C). Tissue section analysis revealed the presence of well-developed blood vessels that stained positively for CD31, and laminin in tumors from WT mice. In contrast, in tumors formed in DiYF mice, blood vessels were sparse and thin walled based on laminin staining (Fig. 2 E, top). The vascular density in tumors grown in DiYF mice was sixfold lower than that in WT mice (P = 0.009) (Fig. 2 E, bottom). Similar defects in tumor growth and neovascularization were observed using alternative model of tumor growth in vivo. Mouse prostate cancer cells (RM-1) were implanted into WT and DiYF mice, and tumor was excised after 10 d. Tumors formed in DiYF mice were smaller than in WT mice (1.17 g vs. 0.8 g) (Fig. 2 D). Tissue section analysis indicated the presence of substantially larger number of vWF-positive blood vessels in tumor developed in WT mice compared with DiYF mice (Fig. 2 F). Collectively, our in vivo studies demonstrate that tumor angiogenesis is impaired in DiYF mice, resulting in reduced tumor growth. Thus, it appears that β₃ integrin cytoplasmic tyrosine motifs play crucial role in the regulation of pathological angiogenesis.

Knowing that monocytes/macrophages use β₃ integrin for their arrest and extravasation at sites of inflammation, we assessed whether DiYF mutation affects macrophage recruitment into tumors using anti-F4/80 antibody staining. As shown in Fig. 2 G, considerable amounts of macrophages were found to be present in tumors implanted in both WT and DiYF mice. No substantial reductions in macrophage recruitment were found in DiYF mice. Similar results were observed in Matrigel model of VEGF-induced angiogenesis (Fig. 2 H). Thus, DiYF mutations within β₃ integrin do not impair the process of macrophage recruitment in tumors.

Homozygous WT and DiYF mice were bred to obtain homozygous littermates. The average number of embryos per litter was the same. Size and weight of pups were not substantially different between WT and DiYF mice. In contrast to β₃ integrin knockout mice, no embryonic mortality caused by placental defects was observed in DiYF mice. DiYF mutation did not affected survival of embryos at early stages of development. DiYF embryos did not exhibit any broad vascular defects (Fig. 3 A). Prenatal DiYF mutant mothers did not suffer from intrauterine hemorrhage and placental defects (Fig. 3 B) as observed in β₃ integrin knockout mice. To examine the survival of DiYF mutant mice after birth, the numbers of pups per litter from homozygous crosses were monitored. Even after 7 d of postpartum there was no significant difference in mortality of pups between these two groups. The DiYF mutant embryos (P = 12) or pups did not show any vasculature defects and hemorrhage in skin or in gastrointestinal tract as observed in β₃ integrin knockout mice (Fig. 3 C). Development of large blood vessels exemplified by aorta also appeared to be normal because no significant differences (P = 0.445) in thickness and general histological structures of aorta were observed in DiYF mice compared with WT counterparts (Fig. 3 D).

To further investigate the effect of DiYF mutation on adult vasculature, tissue sections of liver (Fig. 3 E) and kidney (Fig. 3 F) were stained for vWF and laminin. No substantial differences in blood vessel density of vascular maturation were observed in DiYF mice. Vascular density was assessed in skin sections from WT and DiYF mice using staining for vWF, and there were no significant differences between these two groups (P = 0.725, n = 16; Fig. 3 G). Since vascular maturation and pericyte recruitment are known to be essential for pathological angiogenesis (21), staining for smooth muscle actin was performed to reveal mature blood vessels (Fig. 3 H). Quantitative results demonstrate the similar density of smooth muscle actin–positive blood vessels in skins of WT and DiYF mice (P = 0.127; n = 11). Since β₃ integrin is known to be expressed on circulating white blood cells and involved in immune response, we demonstrated that the numbers of circulating immune cells do not differ among DiYF and WT mice. The numbers of circulating lymphocytes were 86.5 ± 0.93 and 80.1 ± 2.7 × 10³ cells/μl for WT and DiYF mice, respectively. Monocyte counts were also similar: 2.34 ± 0.5 and 2.32 ± 0.6 × 10³ cells/μl for WT and DiYF mice, respectively. Collectively, these data clearly indicate that DiYF mutation did not affect the embryogenesis, organogenesis, postpartum developments, normal vascular development and maturation, and levels of circulating white blood cells.

Previous studies demonstrated that α_vβ₃ integrin controls migration, invasive potential, and angiogenic phenotype of ECs (20). Therefore, we assessed whether impaired β₃ integrin tyrosine phosphorylation affected EC capillary and tube formation ex vivo. WT but not DiYF ECs were able to form well-assembled and complete capillary cord-like structures in the presence of VEGF-A (Fig. 4 A). In contrast, DiYF ECs remained randomly scattered without any signs of organization (Fig. 4 A). The number of cords formed by WT EC was 5.4-fold higher compared with DiYF cells (Fig. 4 B).

A major phenotypic characteristic of ECs is the ability to assemble into the interconnected network of tube-like structures when grown in a three-dimensional matrix (22). WT but not DiYF ECs formed clearly defined and well-connected networks of EC tubes (Fig. 4 C). These structures were relatively stable and remained well organized for at least 18 h. As evident from Fig. 4 C, DiYF ECs were not able to complete tube formation, and no obvious patterns were formed even in the presence of VEGF. The extent of tube formation was quantified by measuring the length of tubes; the mean values of three independent experiments are represented graphically in Fig. 4 D. VEGF treatment induced approximately threefold increase in the length of the tubes formed by WT ECs but had a little effect on DiYF ECs.

We next assessed whether DiYF mutations affected an outgrowth of vascular sprouts from aortic segments isolated from mice. First, an ex vivo angiogenic assay was performed in Matrigel enriched with growth factors. The aortic rings from WT mice produced an extensive network of vascular sprouts, whereas DiYF aortic rings failed to do so (Fig. 4 E). To elucidate the role of β₃ integrin in VEGF-induced responses, the aortic ring assay was performed in the presence or absence of VEGF using growth factor–reduced Matrigel. Under these conditions, aortic rings from WT mice produced a significantly higher number of vascular sprouts both in the absence and presence of VEGF than did aortic rings from DiYF mice (Fig. 4 F). Quantification of the number of aortic ring sprouts indicated that DiYF cells formed fourfold fewer vascular sprouts ex vivo, regardless of stimulation, than did WT aortic rings (Fig. 4 G). VEGF produced a small increase in capillary formation of DiYF rings; however, the number of sprouts was only 20–25% of that observed in WT aortic rings (Fig. 4 G).

To further analyze the capillary growth from aortic rings, a detailed kinetic study was undertaken. The time curves of vascular growth are presented in Fig. 4 H. In the absence of stimulation, very few microvessels were detected in either WT or DiYF implants even after a prolonged incubation, whereas serum-induced neovascularization was considerably higher in WT implants than in DiYF implants (Fig. 4 H). The peak values of capillary growth were observed 8 d after implantation and were 10 and 45 microvessels per ring for DiYF and WT, respectively. VEGF with endothelial growth supplement was a stronger stimulus than endothelial growth supplement alone, but both produced extensive formation of capillaries in WT but not in DiYF aortic rings (Fig. 4 H). Collectively, these results indicate that the impaired pathological angiogenesis in DiYF mice was caused by the defective functional responses of endothelial cells.

To further define the nature of the angiogenic defect observed in DiYF mice, we compared angiogenesis-relevant functions of ECs isolated from WT and DiYF mice. We first assessed whether the mutation in β₃ integrin cytoplasmic domain had any effect on EC adhesion and subsequent cell spreading on extracellular matrix substrates. WT and DiYF ECs were plated on various integrin ligands and numbers of attached and spread cells per field were counted. WT and DiYF ECs adhered and spread equally well on fibronectin, laminin-1, and collagen-coated plates (Fig. 5, A and B). In contrast, a significant difference (P < 0.001) in the behavior of WT and DiYF ECs was found using the α_vβ₃ ligands vitronectin and entactin. On these substrates, DiYF ECs showed a twofold reduction in adhesion and a fourfold decrease in the number of spread cells on vitronectin and threefolds on entactin (Fig. 5, A and B). To further examine the role of β₃ integrin cytoplasmic tyrosine motifs in regulation of receptor ligand-binding strength, we examined cell adhesion under conditions of shear stress. WT and excessive DiYF endothelial cells were plated on vitronectin-coated glass coverslips to achieve equal number of adherent cells. Then, adherent cells were subjected to perfusion with shear rates of 5–30 dyn/cm² for 1 min. As shown in Fig. 5 C, shear stress revealed a dramatic difference in strength of adhesion between WT and DiYF endothelial cells. Shear stress of 10 to 15 dyne/cm² almost completely abolishes DiYF endothelial cell adhesion to vitronectin, whereas >60% of WT endothelial cells remained tightly adhered to the matrix. Shear rate sufficient to wash away 50% of adherent cells was 23 and 8 dyne/cm² for WT and DiYF endothelial cells, respectively (Fig. 5 C). This finding demonstrates that β₃ integrin cytoplasmic tyrosine motifs regulates ligand-binding strength.

When the migratory activity of WT and DiYF ECs toward various extracellular matrix proteins known to be recognized by integrins was compared, results were similar to those of the adhesion assays: WT and DiYF ECs migrated equally well toward fibronectin, laminin, and collagen, but not toward vitronectin and entactin, where a 2.5-fold reduction in migration of DiYF ECs compared with WT was observed (Fig. 5 D).

We then compared the VEGF-induced migratory activity of WT and DiYF ECs. Stimulation of WT ECs with VEGF at 5, 10, and 20 ng/ml induced 1.5-, 2.5-, and 2.9-fold increases of migration compared with untreated ECs (Fig. 5 E). DiYF ECs also responded to VEGF stimulation; however, the rate of migration was substantially reduced when compared with WT cells (Fig. 5 E). Thus, tyrosine phosphorylation of α_vβ₃ integrin appears to play an important role in VEGF-induced EC migration to extracellular matrix. To further confirm these results, we used an alternative and more physiologically relevant method to assess EC migration. WT and DiYF ECs were plated on various integrin ligands and were allowed to form a confluent monolayer. Then, a wound in the monolayer was created and the healing process was monitored at different time points. The quantitative aspects of wound recovery and representative images of ECs are presented in Fig. 5, F and G, respectively. Whereas WT and DiYF EC migrated equally well on fibronectin, laminin, and collagen, a threefold reduction in migration on vitronectin was observed in DiYF ECs compared with WT (Fig. 5 F). The DiYF mutation impairs α_vβ₃ integrin-dependent and VEGF-stimulated responses of ECs, underscoring the role of phosphorylation of tyrosines in the cytoplasmic domain in the regulation of a cross-talk between α_vβ₃ and VEGFRs.

Next, we sought to identify a molecular mechanism responsible for abnormalities observed in DiYF ECs. Previous studies using β₃-null mice demonstrated that the absence of β₃ leads to up-regulation of VEGFR-2 and consequently to augmentation of angiogenic responses of ECs (8). However, no differences in VEGFR-2 levels were observed between DiYF and WT ECs from lung or aortic origin (unpublished data). It was previously shown that integrin β₃ forms a complex with VEGFR-2 immediately upon stimulation with VEGF, and this association was proposed to be necessary for the activation of angiogenic program in ECs (23). Therefore, we sought to determine whether the DiYF mutations impaired the interaction of β₃ integrin with VEGFR-2. Low levels of β₃–VEGFR-2 interaction were observed in unstimulated WT ECs in suspension or upon adhesion to extracellular matrix. VEGF stimulated a dramatic increase in formation of the complex between β₃ and VEGFR-2 in WT ECs plated on vitronectin, but not in suspension or on laminin, demonstrating the ligand specificity of this phenomenon (Fig. 6 A). In contrast, no interaction between β₃ and VEGR-2 was observed in DiYF ECs under any conditions (Fig. 6 B). Thus, β₃ tyrosine phosphorylation was essential for an interaction between VEGFR-2 and α_vβ₃ integrin.

To investigate the mechanism of β₃ integrin-dependent VEGF signaling, we analyzed the VEGFR-2 phosphorylation status in WT and DiYF ECs after VEGF stimulation. The time courses of VEGR-2 phosphorylation are shown in Fig. 6 C. In WT ECs, VEGF induced a bell- shaped response with a maximum sixfold increase in VEGFR-2 phosphorylation relative to unstimulated cells. After 45 min, VEGFR-2 phosphorylation returned to the control level. In contrast, the maximum VEGFR-2 phosphorylation in DiYF ECs upon VEGF stimulation was only 2.5-fold over control. Importantly, VEGFR-2 remained phosphorylated three times longer in WT ECs than in DiYF ECs (Fig. 6 C). Thus, the lack of integrin phosphorylation in DiYF ECs resulted in reduced phosphorylation, and therefore activation, of VEGFR-2 in response to VEGF, which in turn affected all the signaling events downstream of VEGFR-2. To further examine this phenomenon WT and DiYF cells were induced with VEGF for various time periods and cell lysates were analyzed for phosphorylation of extracellular signal–regulated kinase (ERK)-1/2 as downstream target molecule of VEGFR-2. In WT ECs, VEGF induced strong bell-shaped response with a 3.5-fold increase in ERK-1/2 phosphorylation relative to unstimulated cells within 2.5 to 15 min. In contrast, the maximum ERK-1/2 phosphorylation in DiYF ECs upon VEGF stimulation was only 2.5-fold over control and appeared only after 15 min of VEGF stimulation. Importantly, VEGFR-2–mediated ERK-1/2 phosphorylation was immediate and longer in WT ECs than in DiYF ECs (Fig. 6, D and E).

An intrinsic property of integrin is an increase in soluble ligand binding in response to stimulation, a process referred to as integrin activation. We and others previously reported that VEGF activates α_vβ₃ integrin on ECs in response to VEGF (24). We sought to determine whether impaired activation of VEGFR-2 in DiYF ECs resulted in defective α_vβ₃ activation by VEGF. VEGF induced a sixfold increase of fibrinogen binding to WT ECs, but only a threefold increase in binding to DiYF ECs (Fig. 7 A). MnCl₂, an agonist known to activate integrins and stimulate β₃ integrin tyrosine phosphorylation (25), produced at least a 12-fold increase in fibrinogen binding to WT ECs compared with a 5-fold increase observed in DiYF ECs (Fig. 7 B). The specificity of ligand binding was confirmed by addition of 10-fold excess of unlabeled fibrinogen. Similar results were observed when integrin activation was monitored using a monovalent activation-dependent ligand WOW-1 Fab. VEGF and MnCl₂ stimulated 9- and 11-fold increases, respectively, in WOW-1 binding to WT ECs and 3.5- and 4-fold increases, respectively, to DiYF ECs (Fig. 7 C and Fig. 6 D). Thus, it is apparent that the mutations within the cytoplasmic domain of β₃ integrin in the DiYF mice significantly impair the process of integrin activation, resulting in defective cell adhesion and migration.

To investigate the role played by β₃ integrin in pathological angiogenesis, we replaced the WT β₃ integrin with a mutated form unable to undergo phosphorylation, creating the knock-in mouse DiYF. The mutant β₃ integrin is expressed and is present on cell surfaces at the normal levels, but is defective in signaling. The major advantage of this model is that it allows monitoring of α_Vβ₃ function in primary endothelial cells as opposed to model cell lines lacking appropriate receptors and signaling intermediates. The ex vivo and in vitro results from this model provided novel information that is different from the previously published results from in vitro studies and β₃ knockout mice experiments (8). A unique aspect of our experimental system is that compensatory or over-compensatory responses were not observed; these responses often occur in knockout models when a protein of interest is absent during development (26). Using DiYF knock-in mice, we assessed pathological neovascularization in vivo and performed an extensive analysis of the underlying mechanisms of angiogenesis using ex vivo models. The major findings of this paper are the following. (a) Phosphorylation of β₃ integrin in WT ECs occurred in response to integrin ligation and VEGF stimulation. (b) Lack of β₃ integrin phosphorylation in DiYF knock-in mice resulted in impaired angiogenic response in three distinct in vivo models. As a result of reduced vascularization, tumor growth was significantly inhibited in DiYF mice compared with WT. (c) At the same time, DiYF mutation did not affect embryogenesis, organogenesis, and overall vascular development. In contrast to the phenotype of β₃ integrin knockout mice, DiYF pups were broadly normal and did not show any hemorrhage in any organ. Vascular density and maturation in adult DiYF animal was also normal, indicating that integrin phosphorylation is crucial for pathological but not normal vascularization. (d) In ECs from DiYF mice, VEGF-induced functional responses (cell adhesion, spreading, migration, and capillary tube formation) were defective compared with WT. (e) Lack of β₃ integrin phosphorylation in DiYF ECs lead to disruption of the VEGFR-2–β₃ integrin complex and lack of VEGFR-2 phosphorylation in response to VEGF. (f) VEGF-induced integrin activation (inside-out signaling) was suppressed in DiYF ECs compared with WT ECs.

Our data show that, in WT ECs, phosphorylation of β₃ integrin occurs in response to integrin ligation and VEGF stimulation. The finding that β₃ phosphorylation occurs not only in response to adhesion to extracellular matrix (i.e., as a result of outside-in integrin signaling), but also upon treatment with VEGF, indicates that β₃ might play an important regulatory role in outside-in integrin signaling, also known as integrin activation. In DiYF ECs, β₃ cannot be phosphorylated, and integrin activation, as measured by soluble ligand binding to α_Vβ₃, was considerably reduced compared with WT. Previous studies demonstrated that ligation or the treatment with the agonist Mn²⁺ induces β₃ integrin phosphorylation in cells other than ECs (25). At the molecular level, it was also shown that β₃ integrin phosphorylation controls the strength of receptor–ligand interaction (27), and might contribute to the overall functional activity of the integrin.

In three distinct models, one that measured capillary formation in Matrigel and two others for vascularity of implanted tumors, we observed that pathological angiogenesis was severely impaired in DiYF mice. Our findings are consistent with the results of studies using α_Vβ₃ blocking antibodies and demonstrate a key role for α_Vβ₃ integrin in angiogenesis, suggesting that β₃ integrin is a positive regulator of angiogenesis and its phosphorylation is a critical step in EC responses during VEGF-induced neovascularization. Further, our findings provide an additional argument that increased angiogenesis observed in β₃ knockout mice (8) is a result of molecular compensation via VEGR-2; this emphasizes the physiological importance of VEGR-2–α_Vβ₃ cross-regulation in ECs, but does not reflect the true function of α_Vβ₃ integrin in angiogenesis in an organism expressing normal amounts of this receptor. Since our experimental approach does not involve a complete deletion of β₃ integrin from the cell surface, it does not trigger compensatory responses (i.e., up-regulation of VEGR-2). The phenomenon of molecular compensation—in many cases, over-compensation—in knockout mice has been described in numerous reports and represents a major drawback of the knockout approach. Frequently, the loss of a particular protein results in increased expression of other members of the same family, revealing the functional redundancy (28). It has been shown, for example, that α_Vβ₅ integrin can compensate for the loss of α₅β₁ integrin (26). Analysis of the substitute molecule, which is structurally and functionally distinct from the targeted protein (29, 30), might reveal new connections within the molecular network. It is intriguing that VEGR-2, a tyrosine kinase receptor, can compensate for the loss of a member of integrin family of cell adhesion molecules, as VEGR-2 is distinct in regard to ligand repertoire and other functional aspects.

In the present study, impaired angiogenesis in DiYF knock-in mice resulted from the decreased adhesion, spreading, and migration of ECs in response to VEGF. Interestingly, impaired cell adhesion and migration was observed only on vitronectin as a substrate, but not on fibronectin or collagen. Nevertheless, ex vivo angiogenesis in Matrigel, and in vivo angiogenesis, was defective, emphasizing a role for α_Vβ₃ in this process. At the molecular level, this study demonstrated that impaired tyrosine phosphorylation of β₃ integrin in DiYF ECs abrogated α_Vβ₃–VEGFR-2 complex formation and resulted in dramatically reduced phosphorylation of VEGFR-2 upon VEGF stimulation. Since VEGF (via VEGFR-2 as its major functional receptor on ECs) induces phosphorylation of α_Vβ₃ and phosphorylation of α_Vβ₃, in turn, is required for complete and sustained phosphorylation of VEGFR-2, it can be concluded that these two receptors are able to cross-activate each other in ECs, therefore forming a functional partnership that is essential for successful angiogenesis.

Collectively, our findings demonstrate that α_Vβ₃ and its inside-out and outside-in signaling is essential for pathological angiogenesis and undoubtedly represents a promising target for pharmaceutical approaches. However, when developing new inhibitors for angiogenesis, one should take into consideration the complexity of α_Vβ₃ regulation, its interactions with other receptors, and possible compensatory changes resulting from its suppression.

DiYF mice were generated in the laboratory of Dr. David R. Phillips and maintained on a C57/Bl6 background (seven generations of backcrossing). 6–8-wk-old WT and DiYF mice were used in our study. We performed all procedures according to protocols approved by Cleveland Clinic Foundation Institutional Animal Care and Use Committee.

WT and DiYF animals received an abdominal subcutaneous injection of 500 μl Matrigel mixed with VEGF (60 ng/ml) and heparin (60 units/ml). After 7 d, the animals were killed and dissected. Matrigel plugs were removed and digested using 5 ml Drabkin reagent, and quantification of neovascularization was assessed using a hemoglobin assay as per the manufacturer's protocol.

WT and DiYF mice were subcutaneously injected with freshly harvested 10⁶ B16F10 mouse melanoma or RM-1 mouse prostate cancer cells. Tumors were collected 10 d after injection and tumor weights were measured. Tumors were also photographed and processed for immunohistochemical staining.

Immunohistochemistry and image analysis was performed as described previously (21). Matrigel plugs or tumor tissues sections were stained with polyclonal rabbit anti-vWF antibody (Abcam, Inc.) or rat anti-F4/80 (Serotec, Inc). Sections were counterstained with hematoxylin. WT and DiYF skin, liver, and kidney tissue samples were processed and stained for vWF or laminin. B16F10 tumor sections were stained with polyclonal anti-CD31 (Fitzgerald Industries Intl.) and anti-laminin (Sigma-Aldrich) antibody.

The mouse aortic ring assay was performed as described previously (21). WT and DiYF mouse lungs were excised, minced, and digested using collagenase-dispase reagent (3 mg/ml). Digests were strained and the resulting cell suspension was plated on flasks coated with 1 mg/ml fibronectin. Endothelial cells were isolated and characterized as described previously (31).

The cell adhesion assay was performed as described previously (32). Mouse lung endothelial cell suspensions were added to ligand-coated wells and placed in humidified incubator for 45 min. The wells were gently washed three times with DMEM and photographs were taken. The numbers of attached and spread cells per field were counted. Adhesion assay under shear stress conditions was performed as described previously using flow chamber (33). Prior to the beginning of the experiment WT and 2.25-fold excess DiYF endothelial cells were added on vitronectin-coated coverslips to achieve approximately equal number of adherent cell population and allowed to interact with the matrix for 15 min. These coverslips were gently washed and introduced into the bottom of the flow chamber. Shear stress of 5–30 dyne/cm² was applied using syringe pump for 1 min. Number of WT endothelial cells adhered to vitronectin under static condition per field was assigned the value of 100% and relative adhesion was calculated.

Endothelial cell migration assay was performed as described previously (24). WT and DiYF mouse lung endothelial cells were grown to confluence in 12-well plates precoated with various integrin ligands. Cells were serum starved and then a wound was created. Images were recorded immediately after wounding (time zero) and 12 h later. Cell migration was quantified using image analysis of five randomly selected fields of denuded area. The mean wound area is expressed as percent of recovery (% R) from three identically treated plates using the equation % R = [1− (T_t/T₀)] × 100, where T₀ is the wounded area at 0 h and T_t is the wounded area after 12 h.

WOW-1 Fab binding assay was performed as described previously (34). To assess fibrinogen binding, WT and DiYF mouse lung endothelial cells were serum starved for 4 h and further induced with 20 ng/ml VEGF-165 or 1 mm MnCl₂. FITC-labeled fibrinogen was added at a final concentration of 200 nM for 45 min. Cells were fixed and washed twice with ice-cold PBS. FACS was performed using a FACS Calibur (Becton Dickinson), and data were analyzed using CellQuest software program.

The formation of vascular tube-like structures by WT and DiYF mouse lung endothelial cells were assessed on the basement membrane matrix preparation. WT and DiYF mouse lung endothelial cells were seeded on Matrigel-coated plates. Medium with or without 20 ng/ml VEGF was added, and cells were further incubated at 37°C for 8 h. The tube formation was observed using an inverted phase–contrast microscope, and photographs were taken from each well. Using ImagePro software, the degree of tube formation was quantified by measuring the length of tubes in random fields. The precapillary cord formation assay was performed as described previously (22).

Immunoprecipitation was performed as described previously (23). To analyze the β₃ integrin tyrosine phosphorylation of WT and DiYF mouse lung microvascular endothelial cells, ECs were added to various integrin ligand-coated plates and incubated at 37°C for 60 min. Cell lysates containing equal amount of proteins were subjected to Western blot analysis using rabbit anti–integrin β₃ [pY⁷⁴⁷] and [pY⁷⁵⁹] antibody (Biosource International, Inc). Cell lysates were also analyzed for β₃ integrin expression as a loading control. Serum-starved WT and DiYF ECs were also treated with 20 ng/ml VEGF for 0–60 min. Cell lysates were analyzed by Western blot using anti–integrin β₃ [pY⁷⁴⁷], anti–integrin β₃ [pY⁷⁵⁹], anti–ERK-1/2 and anti–p-ERK-1/2 (Cell Signaling Technology) antibodies.

Values were expressed as mean ± SDs. p values were based on the paired t test. Results were considered statically significant with P < 0.05.

We acknowledge financial support from the National Institutes of Health (HL071625 and HL073311 to T.V. Byzovat) and the American Heart Association (0625271B to G.H. Mahabeleshwar).

The authors have no conflicting financial interests.