Endothelial HIF-2α regulates murine pathological angiogenesis and revascularization processes

Localized tissue hypoxia is a consequence of vascular compromise or rapid cellular proliferation and is a potent inducer of compensatory angiogenesis. The oxygen-responsive transcriptional regulator hypoxia-inducible factor 2α (HIF-2α) is highly expressed in vascular ECs and, along with HIF-1α, activates expression of target genes whose products modulate vascular functions and angiogenesis. However, the mechanisms by which HIF-2α regulates EC function and tissue perfusion under physiological and pathological conditions are poorly understood. Using mice in which Hif2a was specifically deleted in ECs, we demonstrate here that HIF-2α expression is required for angiogenic responses during hindlimb ischemia and for the growth of autochthonous skin tumors. EC-specific Hif2a deletion resulted in increased vessel formation in both models; however, these vessels failed to undergo proper arteriogenesis, resulting in poor perfusion. Analysis of cultured HIF-2α–deficient ECs revealed cell-autonomous increases in migration, invasion, and morphogenetic activity, which correlated with HIF-2α–dependent expression of specific angiogenic factors, including delta-like ligand 4 (Dll4), a Notch ligand, and angiopoietin 2. By stimulating Dll4 signaling in cultured ECs or restoring Dll4 expression in ischemic muscle tissue, we rescued most of the HIF-2α–dependent EC phenotypes in vitro and in vivo, emphasizing the critical role of Dll4/Notch signaling as a downstream target of HIF-2α in ECs. These results indicate that HIF-1α and HIF-2α fulfill complementary, but largely nonoverlapping, essential functions in pathophysiological angiogenesis.

The therapeutic manipulation of angiogenesis holds great promise for treating diverse pathological conditions, including cancer, macular degeneration, atherosclerosis, and peripheral arterial disease (PAD) (1, 2). However, the ability to reliably inhibit (or promote) angiogenesis in clinical settings requires a thorough appreciation of the complex molecular events that coordinate vessel sprouting and remodeling (3, 4). These processes are clearly influenced by reductions in local O₂ concentration (hypoxia), which can arise as a consequence of ischemic insult or rapid cell division during solid tumor growth (5, 6). Elucidating the mechanisms by which hypoxic stress modulates the activity of vascular ECs is therefore critical to our understanding of disease-associated angiogenesis.

Cells respond to hypoxic stress through multiple mechanisms, including the stabilization of hypoxia-inducible factors (HIFs), which directly regulate the expression of more than 150 target genes that modulate angiogenesis, cell metabolism, proliferation, survival, and migration (reviewed in ref. 6). Mammalian HIFs function as heterodimers composed of either HIF-1α (7) or HIF-2α (8) bound to HIF-1β (also known as aryl hydrocarbon nuclear translocator). HIF-1α appears to be ubiquitously expressed, whereas HIF-2α is detected in a more restricted set of cell types, including vascular ECs, hepatocytes, type II pneumocytes, and macrophages. A third mammalian HIF-α subunit, HIF-3α (9), has also been described, although its role in hypoxic responses is less well understood.

HIF activity is controlled by hypoxia-induced posttranslational stabilization of HIF-α subunits, which are otherwise hydroxylated, ubiquitylated, and degraded under normoxic conditions through the activity of HIF-specific prolyl hydroxylases (PHD1–PHD3) and other enzymes (6, 10). Additionally, oncogenic signaling pathways have been reported to influence HIF-α levels, in part by regulating the rate of HIF-α protein translation. Although HIF-1α and HIF-2α share highly conserved structural features, oxygen-mediated stabilization, and numerous target genes, mounting evidence indicates that each protein has remarkably distinct functions in specific cellular contexts. These differences are mediated in part by regulation of unique target genes, but also by opposing influences on other critical factors, including c-Myc (11), p53 (12), and nitric oxide (13).

Emerging evidence suggests that HIF-1α and HIF-2α have important and independent effects on pathological angiogenesis. For example, femoral artery ligation (FAL) experiments revealed decreased limb perfusion and increased spontaneous amputation in heterozygous Hif1a^+/– mice, or in wild-type mice treated with the HIF-1α inhibitor 2-methoxyestradiol (14). Furthermore, delivery of adenovirus encoding constitutively active HIF-1α stimulated reperfusion following FAL and hindlimb ischemia in older wild-type mice or in diabetic mice (15); in each case, the effects of HIF-1α were attributed to mobilization of circulating angiogenic cells. EC-specific HIF-1α expression is also required for robust tumor angiogenesis, tumor growth, and wound healing by promoting EC proliferation and survival (16). These effects were correlated with HIF-1α–dependent expression of VEGF and other angiogenic factors.

The specific contribution of EC HIF-2α to ischemia-induced angiogenesis has not been investigated directly, despite the fact that HIF-2α is highly expressed in these cells. An indirect role for HIF-2α in ischemic responses has been reported for PHD1-deficient mice subjected to FAL; specifically, muscles in Phd1^–/– mice are protected from ischemic stress by alterations in glucose metabolism and reactive oxygen species production, phenotypes largely reversed by HIF-2α haplodeficiency (17). In contrast, we previously demonstrated that EC-specific deletion of HIF-2α caused defects in EC adhesion and vessel integrity (18). Moreover, HIF-2α ablation in ECs inhibited subcutaneous xenograft tumor growth and angiogenesis (18, 19), which were associated with greater intratumoral hypoxia and fewer functional vessels relative to tumors grown in control mice. HIF-2α was also implicated as a critical downstream effector modulating normalization of tumor vessels in Phd2^+/– mice, which correlates with improved tumor oxygenation and decreased metastasis (20).

In the current study, we use critical limb ischemia and autochthonous skin tumor models to directly assess the cellular and molecular mechanisms by which HIF-2α regulates EC function and pathological angiogenesis. Mice with EC-specific HIF-2α deletion produced increased numbers of newly formed vessels in ischemic muscles and autochthonous tumors; however, these vessels failed to remodel effectively, thereby reducing overall vascular area, blood flow, and tissue oxygenation. Purified HIF-2α–deficient ECs displayed increased migration, invasion, and tube formation in vitro as well as reduced hypoxic expression of important angiogenic molecules, including angiopoietin 2 (Ang2) and adrenomedullin (ADM1). Interestingly, similar effects on tumor angiogenesis were observed upon inhibition of the delta-like ligand 4/Notch (Dll4/Notch) pathway (21), and we further demonstrated that HIF-2α specifically regulated hypoxic expression of the Notch ligand Dll4 as well as downstream effectors of the Dll4/Notch pathway, both in vivo and in vitro. Finally, we showed that exogenously supplied Dll4 complemented the phenotypes of HIF-2α–deficient ECs both in vitro and in vivo. Collectively, these data identified EC-specific functions for HIF-2α that we believe to be novel, both in restricting angiogenic sprouting and promoting vascular remodeling, and demonstrated that HIF-1α and HIF-2α have fundamentally distinct, but apparently complementary, EC-specific functions in pathophysiological angiogenesis. Whereas previous reports demonstrated that HIF-1α promotes EC proliferation and VEGF expression, thereby driving new vessel formation (16), HIF-2α was essential for remodeling new vessels into a functional vasculature, partly through Notch pathway regulation. These data support the hypothesis that by promoting apparently antagonistic molecular responses, HIF-1α and HIF-2α expression in ECs modulate angiogenic responses in the context of pathophysiological hypoxia to produce a functional vasculature.

Endothelial HIF-2α deletion disrupts blood flow recovery after FAL. Our previous studies suggested that HIF-2α would be a critical regulator of vascular regeneration and angiogenesis in the context of peripheral vascular disease. To test this hypothesis, we challenged control mice or mice lacking EC-specific HIF-2α expression (referred to herein as KO mice) with FAL and investigated subsequent blood flow, vessel regeneration, and tissue damage. Blood flow in the adductor muscles and feet of control and KO mice was assessed before and after surgery using Speckle imaging techniques, which showed markedly decreased blood flow in the ligated limb (Figure 1A). Perfusion in ligated and nonligated limbs was then measured 7, 14, 21, and 28 days after surgery using laser Doppler flowmetry (LD), which detects blood flow in superficial tissues. In addition, diffuse correlation spectroscopy (DCS) was used to measure oxygenation and metabolic processes of the microvasculature in deeper tissues (22). Both LD and DCS analyses revealed significantly reduced flow characteristics in ligated limbs of control and KO animals after surgery compared with nonligated contralateral limbs (Figure 1, B and C), indicative of comparable levels of postprocedure ischemia in both strains. As expected, a steady recovery of blood flow in the ligated limb was evident beginning 7 days postsurgery; however, blood flow recovery was significantly attenuated in KO versus control mice over the entire observation period (Figure 1, B and C). Moreover, KO mice never fully recovered normal blood flow, even after 28 days, in contrast to control mice.

Figure 1

Endothelial HIF-2α deletion affects blood flow, foot movement recovery, and necrosis after femoral artery ligation (FAL). Blood flow, foot movement score, and necrosis were assessed for control and KO mice. Blood flow was quantified by laser Doppler (LD) and diffuse correlation spectroscopy (DCS) before (Pre), immediately after (Post), and on the indicated days following FAL. (A) Representative images obtained by Speckle imaging technique showing the efficiency of the surgery distally to the occlusion side. (B and C) Quantitative LD and DCS analyses showing the left limb/right limb ratio after occlusion of the femoral artery. KO mice displayed delayed restoration in perfusion compared with control mice. (D) Oxygen saturation in ligated and nonligated limb was determined for control and KO mice using diffuse reflectance spectroscopy at several time points. (E) Foot movement was determined and scored between 0 and 4 as a functional readout parameter to assess flow deficits after ischemia. Active foot movement was significantly impaired in KO mice. (F) Percent mice presenting necrosis or amputation. More KO mice had to be euthanized due to amputation or necrosis of the limb. n = 20 (control); 22 (KO). *P < 0.05; **P < 0.01.

Similarly, ligated limbs in both KO and control strains displayed equivalent reductions in oxygen saturation immediately postsurgery. Oxygen saturation returned to nearly normal levels in control mice by 7 days postsurgery, whereas blood oxygen saturation remained significantly reduced in KO mice over the 28-day postsurgery observation period (Figure 1D), which indicates that ligated limbs in KO mice remain substantially more ischemic than those in control mice.

The functional consequences of impaired revascularization in KO mice were investigated using a scoring system based on active foot movement (see Methods and ref. 14). Foot movement in the ligated limb was significantly reduced at days 7, 14, 21, and 28 in KO mice compared with control animals (Figure 1E). Finally, KO mice exhibited increased incidence of tissue necrosis and spontaneous amputation compared with control mice (Figure 1F), further suggesting that HIF-2α activity in ECs is required for proper vascular recovery after FAL.

Impaired postnatal arteriogenesis and angiogenesis in mice with EC-specific HIF-2α deletion. To investigate the cellular nature of the perfusion defects observed in ischemic KO muscles, we quantified collateral arterial growth and capillary formation in affected tissues. Prior to surgery, histomorphometry revealed no difference in the size of collateral arteries in contralateral limbs of control and KO mice. By 28 days postsurgery, control mice displayed increased collateral arterial diameter and wall area in affected tissues, consistent with active arterial remodeling and growth; however, this enhancement was significantly diminished in KO mice (Figure 2, A–C). Furthermore, smooth muscle cell recruitment to collateral arteries was also reduced in KO mice compared with controls (Supplemental Figure 1; supplemental material available online with this article; doi: 10.1172/JCI57322DS1), revealing a role for EC-specific HIF-2α expression in injury-induced arteriogenesis.

Figure 2

Impaired postnatal arteriogenesis and angiogenesis and increased necrosis in KO mice. (A) Representative H&E-stained sections of adductor muscle from ligated and nonligated limb in control and KO mice. Arrows indicate collateral wall. Arrowheads indicate collateral lumen. (B and C) Histomorphometric analysis of collateral lumen and wall area showing deficient collateral artery growth in KO mice. Arrows indicate collateral wall, arrowheads indicate collateral lumen. (D–F) Rhodamine-lectin staining (D) showed increased vessel density (E) but decreased vascular area (F) in adductor muscle of KO mice. Arrowheads indicate capillaries. (G) H&E-stained sections of adductor muscle for ligated and nonligated limb in control and KO mice. Boxed regions are shown at higher magnification below. Dashed outlines denote damaged or injured areas within muscle. (H) Percent injured area per section determined for ligated limb. (I) Levels of creatine kinase activity assessed in serum after ligation. Scale bars: 20 μm (A and C); 80 μm (G, top); 40 μm (G, bottom). Original magnification, ×200 (A and G, bottom); ×400 (D); ×100 (G, top). n = 8 (control); 9 (KO). *P < 0.05; **P < 0.01.

Capillary density was enhanced in ischemic muscles from all mice after 28 days, although ischemic KO muscles displayed a surprising increase in vessel density relative to controls (Figure 2, D and E). Nevertheless, the corresponding total vascular area (rhodamine-lectin–labeled vessels plus lumens) was actually reduced in ischemic KO muscles compared with control muscles (Figure 2F). These data, as well as the relative paucity of capillaries with large lumens in ischemic KO muscles (Figure 2, D and F), suggest that KO vessels have an impaired ability to remodel into fully functional vessels in ischemic tissue. This hypothesis is supported by the observation that muscles from ligated limbs in KO mice exhibited more extensive areas of injury and necrosis than those in control mice (Figure 2, G and H). Serum creatine kinase activity, a marker of muscle damage, was elevated in all mice after FAL; however, KO mice retained persistently higher creatine kinase activity than control mice (Figure 2I), reflecting greater muscle damage in these mice. Collectively, these results strongly suggest that recovery from FAL-induced ischemia requires EC HIF-2α activity; otherwise, incomplete or aberrant arteriogenesis and angiogenesis occur, with consequent increases in tissue damage.

EC HIF-2α deletion affects tumor growth and vasculature in a skin carcinogenesis model. To determine whether EC-specific HIF-2α expression was required for vessel sprouting and remodeling in a distinct model of neovascularization, we investigated the growth and angiogenesis of carcinogen-induced skin epithelial tumors in KO and control mice. Previous experiments using subcutaneous xenograft tumors indicated that EC-specific HIF-2α deletion inhibited tumor growth (18); however, rapid growth of Lewis lung carcinoma and B16 melanoma xenografts do not always reflect the physiology of bona fide autochthonous tumors. To address this point, skin papillomas were induced in control and KO mice using a standard DMBA/TPA protocol (see Methods) for 25 weeks (Figure 3A). After 5 weeks, epidermal tumors were apparent in all treated mice (Figure 3B), and their number and size increased with time (Figure 3C). Interestingly, no difference in the percentage of mice with small (0–3 mm) and medium (3–6 mm) skin tumors was observed between KO and control mice; however, the formation of large (>6 mm) skin tumors was delayed in KO mice compared with controls (Figure 3C). Large tumors appeared approximately 5 weeks earlier in control mice than in KO mice, and after 20 weeks, control mice were more likely to develop large papillomas than KO mice (4 of 8 control, versus 1 of 8 KO). A similar pattern was observed after 25 weeks: 6 of 8 control mice, compared with 3 of 8 KO mice, presented with large tumors (Figure 3C). Finally, control mice formed 10 times as many large papillomas at 25 weeks (control, 2.23 ± 1.17 papillomas/mouse; KO, 0.23 ± 0.60 papillomas/mouse; P < 0.05; Figure 3, D and E).

Figure 3

Endothelial HIF-2α deletion affects tumor formation and angiogenesis in a DMBA/TPA-induced tumorigenesis model. (A) Skin carcinogenesis was induced using the DMBA/TPA protocol for 25 weeks. Lesions with diameters of 0–3 mm, 3–6 mm, and >6 mm were recorded as tumors. (B) Representative images of tumor formation on the dorsal skin of control and KO mice after 25 weeks. Tumors of different sizes are shown by arrowheads. (C) Incidence of tumor formation, shown as percent mice with 0–3 mm, 3–6 mm, and >6 mm tumors during the 25 weeks of treatment. (D) Number of tumors per genotype. (E) Number of tumors per mouse. (F) Representative H&E-stained sections of skin tumors. Arrowheads denote vessels within tumors. (G) Vessel density, determined for vessels <30 μm and >30 μm in diameter. (H–J) Rhodamine-lectin staining (H) showed decreased lumen area (I), but increased capillary density (J), in skin tumors of KO mice. Arrows indicate capillaries; arrowheads indicate vessel lumens. (K) Desmin (pericyte marker) and CD31 (EC marker) costaining in skin tumors from control and KO mice. (L) Pericyte recruitment, expressed as the pericyte/EC ratio, was diminished for tumors grown in KO mice. Scale bars: 800 μm (F, left); 25 μm (F, right); 40 μm (H); 20 μm (K). Original magnification, ×50 (F, left); ×200 (F, right); ×400 (K). n = 10 (control); 11 (KO). *P < 0.05.

Immunohistochemical and histological analyses of skin papillomas from KO mice revealed impaired angiogenesis compared with control mice (Figure 3, F–L). Specifically, skin tumors in KO mice were characterized by an increased number of small vessels and a reduced number of large vessels compared with control mice (Figure 3G). Rhodamine-lectin staining confirmed increased functional capillary density and decreased lumen area in KO skin tumors (Figure 3, H–J), similar to results in ischemic muscle. Moreover, immunohistochemical staining for desmin and α–smooth muscle actin (Figure 3K and Supplemental Figure 1) revealed a significant decrease in the degree of mural cell coverage of tumor vessels in KO mice (Figure 3L).

Collectively, our data demonstrated that HIF-2α–deficient ECs produced elevated numbers of capillaries compared with controls in 2 distinct pathophysiological angiogenesis models (ischemic muscle tissue and autochthonous skin tumors). In each case, however, these HIF-2α–deficient capillaries failed to remodel into mature, functional blood vessels, resulting in decreased perfusion.

Cell-autonomous effects of HIF-2α or HIF-1α deletion in ECs. To identify possible cell-autonomous functions of HIF-2α that underlie the angiogenic phenotypes observed in vivo, we generated primary EC cultures from mice carrying a conditional floxed Hifa allele and the Ubc-CreER transgene (see Methods and refs. 23, 24). Independent cultures of immortalized lung ECs that expressed characteristic cell surface markers (CD31, ICAM2, and vWF) and took up the EC-diagnostic compound DiI-Ac-LDL (Supplemental Figure 2) were isolated. Subsequent deletion of the conditional Hif2a allele was induced by tamoxifen treatment, as previously described (18). PCR analysis revealed efficient recombination of the Hif2a floxed allele, and Western blot analysis confirmed loss of HIF-2α protein expression in these cells (Supplemental Figure 2). Similar analyses revealed efficient recombination of the conditional Hif2a allele in purified EC populations from muscle and skin of KO mice (Supplemental Figure 3 and ref. 25).

Tamoxifen-induced HIF-2α deletion had no statistically significant effect on the proliferation or viability of ECs cultured under normoxic (21% O₂) or hypoxic (0.5% O₂) conditions, compared with matched HIF-2α–expressing EC controls (Supplemental Figure 4, A–F). Similarly, no significant difference in EC proliferation was observed between control and KO ischemic muscle (Supplemental Figure 4, G and H). We next investigated the effects of HIF-2α deletion on EC migration, invasion, and adhesion (Figure 4 and Supplemental Figure 5), which are essential for physiological and pathophysiological angiogenesis. Scratch assays revealed no obvious difference in migration under normoxic conditions for control or KO ECs (Figure 4A); however, EC migration was markedly increased under hypoxic conditions and further elevated by HIF-2α deletion (Figure 4, A and B). Related experiments using Boyden chambers showed that control and KO ECs exhibited enhanced invasive behavior in hypoxia, which was also significantly increased by HIF-2α deletion (Figure 4, C and D). Hypoxic KO ECs also exhibited reduced adhesion to dishes coated with collagen (but not plastic), which correlated with elevated numbers of visible filopodial extensions (Supplemental Figure 5), consistent with increased mobility of these cells.

Figure 4

HIF-2α deletion affects EC migration and network formation. (A) Migration of control and KO lung ECs was assessed under normoxic (21% O₂) or hypoxic (0.5% O₂) conditions at several time points using a scratch wound assay. Representative images were taken 24 hours after scratch. Ctrl H2.1 and KO H2.1, control and HIF-2α KO ECs. Ctrl H2.2 and KO H2.2, control and KO ECs with an acute deletion of HIF-2α (Tamoxifen induced). (B) Percent wound closure, determined at consistent locations, was significantly increased for KO ECs 16 and 24 hours after scratch and under hypoxic conditions. (C) EC invasion was further assessed using a Boyden chamber assay. Images represent control and KO EC invasion after 8 hours under hypoxic conditions. (D) Number of invaded cells was quantified after 8 hours under normoxic or hypoxic conditions. Increased invasion was observed for KO ECs compared with control under hypoxic conditions. (E) EC network/capillary formation was assessed using a Matrigel assay. Images represent control and KO EC capillary formation after 8 hours under normoxic or hypoxic conditions. (F–H) Quantifications of EC network formation revealed significant increase in total cellular cord length (F), number of lumens (G), and branch points (H) for KO versus control ECs under hypoxic conditions. Original magnification, ×100. Scale bars: 30 μm. Experiments were performed in triplicate. *P < 0.05; **P < 0.01.

Finally, we measured the ability of HIF-2α–deficient ECs to form cytoplasmic extensions (also referred to as cords) when cultured in Matrigel, an established in vitro angiogenesis assay. Under normoxic conditions, matched control and KO ECs formed cords within 8 hours of plating (Figure 4, E–H). The number and length of EC cords, as well as the number of branch points and internal lumens formed, was enhanced by hypoxic conditions, but again significantly elevated by HIF-2α deletion (Figure 4, F–H). These results surprisingly indicated that HIF-2α normally restrains specific EC processes associated with angiogenesis, including EC migration, invasion, and cord formation in vitro. Our data suggest that the aberrant vascular remodeling observed in ischemic muscles and autochthonous tumors from KO mice may be caused by inappropriate activation of these processes in hypoxic HIF-2α–deficient ECs.

To confirm that our results were specific to HIF-2α deletion, we also generated HIF-1α–deficient immortalized EC cultures (Supplemental Figure 2). In contrast to HIF-2α, and consistent with a previous report (16), deletion of HIF-1α reduced EC viability and proliferation (Supplemental Figure 6), as well as migration and invasion under hypoxic conditions (Supplemental Figure 7). Moreover, the formation of cords, branch points, and lumens in Matrigel culture was strongly inhibited in HIF-1α–deficient ECs under hypoxia (Supplemental Figure 7). Collectively, these data suggest that HIF-1α and HIF-2α exhibit unique and apparently antagonistic effects on vascular remodeling and angiogenesis.

EC HIF-2α deficiency impairs hypoxic induction of genes regulating angiogenesis and arteriogenesis. Previous data indicate that HIF-2α regulates multiple genes encoding angiogenic factors such as Dll4, ADM1, and Ang2 (18, 26). We hypothesized that altered expression of these genes could account, at least in part, for the vascular phenotypes we observed in KO ECs, both in vitro and in vivo. For example, inhibition of Dll4 signaling has been shown to result in proliferation of disorganized capillaries, leading to decreased tumor perfusion and growth (21). ADM1 also modulates arterial differentiation and vascular development, in part by regulating the Dll4/Notch pathway (27–29). Finally, Ang2 promotes arteriogenesis and tissue reperfusion after artery occlusion (30).

Quantitative real-time RT-PCR (qRT-PCR) analyses confirmed that HIF-2α deletion greatly inhibited the hypoxic induction of Ang2, Dll4, and Adm1 mRNA expression in ECs (Figure 5A), although the absolute level of mRNA expression varied among independent EC lines. The hypoxic induction of multiple Notch target genes, including Hey1, Hey2, and Hes1, was also significantly inhibited in HIF-2α–deficient ECs. This response was selective, in that other genes encoding related angiogenic regulators (Vegf, Tie2, and Ang1) were unaffected by HIF-2α deletion. Of note, HIF-1α deletion in ECs had relatively minor effects on hypoxic Dll4, Adm1, Ang2, and Notch target gene induction, but significantly reduced the hypoxic induction of Vegf and Ang1 transcript levels (Figure 5B), in contrast to HIF-2α deletion (18). These data are consistent with the notion that HIF-2α and HIF-1α regulate the expression of distinct angiogenic factors in ECs. Interestingly, deletion of either HIF-1α or HIF-2α reduced the expression of the arterial marker Ephb2 in ECs, but had no effect on the expression of the venous marker Ephb4 (Figure 5, A and B).

Figure 5

Endothelial HIF-2α deficiency impairs hypoxic induction of angiogenic genes. (A and B) Hypoxic induction of HIF and Dll4/Notch pathway target gene expression was assessed by qRT-PCR in control, HIF-2α (A), and HIF-1α (B) KO lung ECs. The relative ratio of hypoxic to normoxic gene expression (fold hypoxic induction) is shown for control and KO ECs (n = 6). Note that Hes1 charts for HIF-2α KO ECs were separated for the 2 replicates. (C) Dll4 staining in adductor muscle for ligated and nonligated limb in control and KO mice at day 7 after surgery. (D) Quantification of fluorescence intensity revealed decreased Dll4 expression in the ligated limb of KO mice after ligation. (E) Levels of Ang2 were assessed in control (n = 8) and KO (n = 9) serum after ligation. (F) Ang2 staining in adductor muscle for ligated and nonligated limb in control and KO mice at day 7 after surgery. Arrowheads denote collaterals. (G) Quantification of fluorescence intensity revealed decreased Ang2 expression in the ligated limb of KO mice. (H) Dll4-immunofluorostained sections of skin tumors from control and KO mice after 25 weeks of DMBA/TPA treatment. (I) Quantification of fluorescence intensity revealed decreased Dll4 expression in skin tumors of KO mice (n = 10 [control]; 11 [KO]). Scale bars: 40 μm (C and H); 20 μm (F). Original magnification, ×200. *P < 0.05; **P < 0.01.

The differential expression of Dll4 and Ang2 were of particular interest, as the primary angiogenic phenotypes in KO mice (specifically, increased nonproductive capillary formation and reduced arteriogenesis) were similar to angiogenic phenotypes resulting from decreased Dll4 and Ang2 expression, respectively (30, 31). To determine whether changes in Dll4 and Ang2 transcript levels in cultured HIF-2α KO ECs correlated with altered EC protein expression in our in vivo models, we performed immunofluorescence on the ischemic muscle tissues and skin tumors described in Figures 1–4. Previous work demonstrated that Dll4 transcription is induced in adductor muscles after FAL (31), and whole-mount immunofluorescent analysis revealed similar elevation of Dll4 protein expression in vessels of ischemic muscles in control mice 7 days after FAL (Figure 5C). In contrast, Dll4 expression in KO ischemic muscle was reduced by 30% (Figure 5D).

Ang2 protein expression is also induced in ischemic adductor muscles in response to FAL, where it likely contributes to vessel enlargement and remodeling (30). We assessed circulating Ang2 protein levels in the serum of control and KO mice at multiple time points after FAL. In control mice, serum Ang2 levels increased 3-fold (to 120 ± 16 pg/ml) 7 days postsurgery, and gradually returned to starting levels (39 ± 6 pg/ml) 28 days postsurgery. This increase in serum Ang2 levels was dramatically inhibited in KO mice, reaching a peak of only 63 ± 7 pg/ml 7 days postsurgery and returning to starting levels 14 days postsurgery (Figure 5E). We also determined that Ang2 staining was increased, and localized primarily to larger collateral vessels, in vessels of ischemic muscles from control mice, whereas Ang2 staining intensity was reduced by 56% in ischemic muscles from KO mice (Figure 5, F and G).

Finally, we performed immunofluorescence analysis of Dll4 expression in skin tumors from control and KO mice. Vessels from control tumors exhibited robust Dll4 expression, which was reduced by 40% in tumors from KO mice (Figure 5, H and I). Collectively, these data indicate that Dll4 and Ang2 expression in ECs is regulated by HIF-2α both in vitro and in vivo and that reduced Dll4 and Ang2 expression in KO mice correlates with impaired tissue recovery from ischemia and tumor angiogenesis.

Dll4/Notch pathway inhibition phenocopies HIF-2α deficiency in EC network formation, migration, and adhesion. To investigate the specific role of Dll4/Notch signaling in the HIF-2α–dependent EC phenotypes, we attempted to manipulate the expression of specific Dll4/Notch pathway components in cultured ECs using gene transfer methods; however, these approaches were hampered by low transfection efficiencies (data not shown). We therefore tested whether the γ-secretase inhibitor N-[N-(3,5-difluorophenacetyl-l-alanyl)]-S-phenylglycine t-butyl ester (DAPT), which blocks Notch cleavage and activation (32), could phenocopy HIF-2α deficiency in control ECs. In initial experiments, we verified the efficacy of DAPT treatment by monitoring the accumulation of Notch intracellular domain (NICD) protein, as well as Notch target gene expression (Supplemental Figure 8). Robust induction of NICD protein accumulation in hypoxic control ECs was effectively blocked by DAPT treatment, which also inhibited the more meager NICD induction observed in KO ECs (Supplemental Figure 8A). Corresponding hypoxic induction of the Dll4/Notch pathway target genes Hes1 and Hey2 was also significantly impaired in ECs treated with DAPT (Supplemental Figure 8B).

We then investigated the effects of DAPT treatment (and Notch inhibition) on EC migration, invasion, and tube formation assays using control and KO cells grown under hypoxic conditions (Figure 6). Interestingly, control ECs treated with DAPT exhibited increased cell network/capillary formation (Figure 6, A–D) as well as increased migratory and invasive phenotypes (Figure 6, E–G), achieving levels similar to those observed for untreated KO ECs. Consistent with these results, DAPT treatment reduced the adhesion of control HIF-2α–replete ECs to Matrigel or fibronectin-coated plates to levels indistinguishable from those of untreated KO ECs (Supplemental Figure 8, D and E).

Figure 6

Notch pathway inhibition phenocopies HIF-2α deficiency in EC network formation and migration. (A) EC network/capillary formation was assessed using a Matrigel assay. Images show control and KO ECs after 8 hours under hypoxic conditions and treated with DAPT. (B–D) Quantification of EC network formation revealed no significant difference in total cellular cord length (B) and number of branch points (C) and lumens (D) between untreated KO ECs and DAPT-treated control cells under hypoxic conditions. (E) Migration of control and KO lung ECs treated with DAPT was assessed under hypoxic conditions at several time points using a scratch wound assay. Representative images were taken 16 hours after scratch. (F) Percent wound closure showed no significant difference between untreated KO ECs and DAPT-treated control cells at 16 hours after scratch and under hypoxic conditions. (G) EC invasion was further assessed using a Boyden chamber assay after 8 hours under hypoxic conditions and treatment or not with DAPT. The number of invaded cells showed no significant invasion difference between untreated KO ECs and DAPT-treated control cells under hypoxic conditions. Original magnification, ×100. Scale bars: 30 μm. Experiments were performed in triplicate. *P < 0.05; **P < 0.01.

In a complementary gain-of-function experiment, control and KO ECs were plated on tissue culture dishes coated with Fc-Dll4 fusion protein (Figure 7). In contrast to soluble Fc-Dll4 protein, which acts as a Notch inhibitor, immobilized Fc-Dll4 protein has been shown to activate Notch signaling (33, 34). The elevated migratory behavior of hypoxic KO ECs was diminished by exposure to Fc-Dll4 ligands (Figure 7, A and B). Control ECs exposed to Fc-Dll4 similarly displayed a slight decrease in migration, also to levels comparable to KO ECs (Figure 7B). Western blot and qRT-PCR analysis confirmed that the Fc-Dll4 ligand activated Notch signaling, as levels of NICD protein and Hes1 and Hey2 mRNA were strongly induced in hypoxic control and KO ECs (Figure 7, C and D). These results implicate the Dll4/Notch pathway as a principal downstream target of HIF-2α function in ECs in pathophysiological angiogenesis and arteriogenesis.

Figure 7

HIF-2α deletion affects EC migration through the Dll4/Notch pathway. (A) Migration of control and KO lung ECs was assessed on control- or Dll4-coated plates under hypoxic conditions at several time points using a scratch wound assay. Representative images were taken 24 hours after scratch. Original magnification, ×100. Scale bars: 100 μm. (B) Percent wound closure showed no significant difference between KO and control ECs on Dll4-coated plates at 24 hours after scratch and under hypoxic conditions. (C) Western blot analysis of NICD protein expression in control and KO ECs seeded on control- or Dll4-coated plates under hypoxic conditions. (D) Induction of Dll4/Notch pathway target genes expression Hes1 and Hey2 was assessed by qRT-PCR in control and KO lung ECs seeded on control- or Dll4-coated plates under hypoxic conditions. Experiments were performed in triplicate. *P < 0.05; **P < 0.01.

Exogenously supplied Dll4 and Ang2 rescue angiogenic phenotypes of HIF-2α–deficient ECs in vivo. To confirm Dll4/Notch signaling as a physiologically relevant downstream effector of HIF-2α function in ECs, we repeated the FAL procedure, but injected surgically exposed KO adductor muscle with adenovirus-associated virus (AAV) engineered to express Dll4 protein. Experiments using control AAV vector encoding GFP revealed extensive expression in muscle fibers and ECs (Supplemental Figure 9A). Whereas injection of control AAV vector had no effect on recovery of limb perfusion in control or KO mice, injection of virus expressing Dll4 protein (Supplemental Figure 9B) largely restored blood flow in ischemic KO muscle to control levels by 7–14 days postsurgery (Figure 8 and Supplemental Figure 10, A and B). Additional postoperative intravenous injection of soluble recombinant Ang2 protein augmented the effects of Dll4 expression, fully restoring blood flow in ischemic KO limbs to control levels (Figure 8B). Concomitant changes in collateral artery diameter (Figure 8, C and D), relative tissue injury (Figure 8, E and F), capillary density and vascular area (Figure 8, G–I), and foot movement and necrosis (Supplemental Figure 10, C and D) were also observed in KO mice treated with Dll4 and Ang2. These data strongly suggest that Dll4 and Ang2 are critical HIF-2α effectors required to regulate effective compensatory angiogenesis in ischemic tissue.

Figure 8

Dll4, alone or in combination with Ang2, is sufficient to reverse endothelial HIF-2α deletion phenotypes in a hindlimb ischemia model. Blood flow, arteriogenesis, and angiogenesis were assessed for control and KO mice after FAL and intramuscular injection of an empty viral vector or of viral vectors expressing Dll4 or Dll4 vector in combination with intravenous Ang2 (Dll4/Ang2). (A and B) Representative images obtained by LD at day 14 (A) showed full flow restoration for KO mice injected with viral vector expressing Dll4 or Dll4 plus Ang2. (B) Quantitative LD analysis showing the left limb/right limb ratio after occlusion. (C) Representative H&E-stained sections of adductor muscle from ligated limb in control and KO mice. (D) Histomorphometric analysis of collateral lumen showing rescue in collateral diameter by Dll4 expression and the combination of Dll4 and Ang2. (E) H&E-stained sections of adductor muscle for ligated limb in control and KO mice injected as indicated. Dashed outlines denote damaged or injured areas within muscle. (F) Percent injured area per section for ligated limb in control and KO mice. (G–I) Rhodamine-lectin staining (G) showed decreased vessel density (H) and increased vascular area (I) in adductor muscle of KO mice injected with Dll4 alone or in combination with Ang2. Arrows indicate capillaries. Scale bars: 20 μm (C and G); 80 μm (E). Original magnification, x200 (C); x100 (E); ×400 (G). n = 20 per group. *P < 0.05; **P < 0.01.

Angiogenesis and vascular remodeling are highly complex, dynamic processes strongly influenced by hypoxia and other microenvironmental stimuli (1). Multiple studies have revealed that the oxygen-responsive HIF transcriptional factors HIF-1α and HIF-2α are important regulators of pathological angiogenesis (10), although the precise mechanisms by which they coordinate vessel sprouting and remodeling remain unclear. In the current work, we identified unique cell-autonomous functions for HIF-2α in regulating EC behavior and demonstrated that EC-specific HIF-2α deletion in mice impaired angiogenesis in hindlimb ischemia and autochthonous solid tumor models. It is interesting to note that, although reduced HIF-1α expression also disrupted angiogenesis in these disease models, there were striking phenotypic differences between HIF-1α– and HIF-2α–deficient mice.

HIF-1α and HIF-2α share many target genes, but mounting evidence indicates that each protein fulfills distinct, and sometimes opposing, physiological functions in specific cell types, including ECs (35). For example, hypoxic VEGF expression is driven primarily by HIF-1α in ECs, whereas hypoxic Dll4 and Ang2 expression is regulated primarily by HIF-2α, suggesting an important division of transcriptional labor that underlies complementary effects on EC behavior. HIF-1α–null ECs cultured under hypoxic conditions exhibit reduced proliferation, VEGF expression, migration, and cytoplasmic cord formation (16). In contrast, the HIF-2α–deficient ECs of our KO mice displayed no significant differences in proliferation rates or VEGF expression, but instead exhibited significantly enhanced branching, migration, and invasion. Moreover, pathophysiological angiogenesis in KO mice was characterized by increased vessel density, but poor arteriogenesis and reduced perfusion, resulting in increased ischemic damage or reduced autochthonous tumor growth. Using both in vitro and in vivo techniques, we identified Dll4 and Ang2 as critical downstream effectors of HIF-2α function in ECs and further demonstrated that exogenous Dll4 and Ang2 expression was sufficient to fully restore perfusion in ischemic muscle with HIF-2α–deficient ECs.

Taken as a whole, the available data are consistent with a model in which HIF-1α promotes EC growth, proliferation, and morphogenesis in response to local hypoxic stress; however, these activities are restrained and fine-tuned by HIF-2α to promote vessel remodeling and produce a mature, functional vascular network (Figure 9). In previous reports, the effects of HIF-1α on angiogenesis in the FAL model were attributed primarily to the mobilization of circulating angiogenic cells (14, 36), and Takeda et al. demonstrated that Phd2^+/– mice were protected against hindlimb ischemia through modulation of macrophage populations, which regulate arteriogenesis (37). We are currently investigating whether HIF-2α regulates the recruitment of these cells to sites of pathological angiogenesis in our models; however, the EC-intrinsic effects on Dll4 and Ang2 appear to account for the majority of the phenotypes observed. Nevertheless, additional HIF-2α target genes almost certainly play important roles in pathophysiological angiogenesis. It would be of considerable interest to subject mice with EC-specific loss of both HIF-1α and HIF-2α to similar ischemia and tumor growth experiments, although an inducible deletion strategy would likely be required, as EC-specific deletion of the murine gene encoding HIF-1β produces embryonic and perinatal lethality associated with altered liver function (38).

Figure 9

Unique cell-autonomous functions for HIF-2α in physiological and pathophysiological angiogenic processes. (A) Inhibiting either HIF-1α or HIF-2α in ECs leads to an inadequate vasculature by different mechanisms, but results in the same consequences on tumor growth and vascularization processes. (B) Model of HIF-1α and HIF-2α functions in murine ECs in response to hypoxia/ischemia, reflecting unique transcriptional targets of HIF-1α and HIF-2α. HIF-1α and HIF-2α fulfill complementary, and largely nonoverlapping, essential functions in physiological and pathophysiological angiogenesis.

Our data are consistent with other reports demonstrating that HIF-1α and HIF-2α can promote opposing cellular responses in specific settings. Examples include inhibition or augmentation of c-Myc transcriptional activity in renal cell carcinomas (by HIF-1α or HIF-2α, respectively; ref. 11) and elevated or diminished nitric oxide production in macrophages (by HIF-1α or HIF-2α, respectively; refs. 13, 39, 40). These and other data (41) make it increasingly clear that HIF-1α and HIF-2α mediate hypoxic responses by acting either in concert or in opposition, depending on the precise physiological setting. This model also suggests that therapeutic targeting of the HIF pathway may require independent regulation of each isoform, depending on the specific pathological context (42).

Our data also revealed HIF-2α as the primary activator of hypoxic Dll4/Notch signaling in ECs, adding an interesting facet to the complex relationship between these important regulatory pathways. The Dll4/Notch pathway is required for angiogenesis in peripheral ischemia models; for example, blood flow recovery and postnatal neovascularization in response to hindlimb ischemia are impaired in Notch1^+/– or Dll1^+/– mice (43). Additionally, Dll4 inhibition impairs reparative angiogenesis after ischemia by the formation of a disorganized, low-perfused capillary network in ischemic muscles (31) and causes proliferation of nonfunctional vessels in xenograft tumors, resulting in poor perfusion and reduced tumor growth (21). These phenotypes are remarkably similar to those observed in our FAL and papilloma experiments, and we further demonstrated that enforced Dll4/Notch signaling largely reversed the impaired recovery from FAL in our KO mice.

Previous analysis demonstrated that Dll4 inhibition promoted the formation of vascular tip cells at the expense of proliferative stalk cells, resulting in vascular discontinuity and leakage (44, 45). Initial attempts to investigate the relative presence of tip and stalk ECs in ischemic muscles and papillomas of our control and KO animals were inconclusive, as were attempts to quantify these cells using a more tractable oxygen-induced retinopathy (OIR) model (ref. 46 and data not shown). The equivocal OIR results may reflect the mixed genetic background of the mice (47); consequently, we intend to repeat these experiments using mice that have been repeatedly backcrossed onto the C57BL/6 genetic background. It is also interesting to note that HIF-2α–deficient ECs have reduced expression of VEGFR2 (18), which collaborates with Dll4/Notch signaling to regulate tip and stalk cell fate (48). Whether HIF-2α, Dll4/Notch signaling, or both regulate VEGFR2 expression in tip and stalk cells is in interesting question for future study.

Hypoxia has previously been reported to regulate Notch pathway activity through direct and indirect mechanisms in other settings. For example, HIF-1α binds NICD directly in stem and progenitor cells, promoting NICD stabilization and increased target gene expression (49). Reduced HIF-1α expression has been shown to correlate with decreased NICD levels and activity in thymic lymphomas from Hif1a^+/–p53^R270/R270 mutant mice (50), and mRNA and protein levels of Notch1 are induced by hypoxia in a HIF-1α–dependent manner in melanoma cells (51). Moreover, ectopically expressed HIF-1α and HIF-2α activate the expression of genes encoding Dll4, Hey1, and Hey2 in endothelioma and endothelial precursor cells, although this regulation may be independent of direct HIF DNA binding activity (26). Interestingly, ADM1 is an established HIF-2α target and has also been shown to activate the Notch pathway, providing another potential link between hypoxia and Notch signaling in ECs (28). It is also possible that HIF and Notch pathways collaborate in stromal cells, such as pericytes and smooth muscle cells, to regulate angiogenesis and vascular function.

Determining the complex effects of local hypoxia on pathological angiogenesis is likely to have important therapeutic consequences (52). The ability to induce and regulate angiogenesis and vascular remodeling in a directed manner would represent a major advance in the treatment of ischemic vascular diseases, including myocardial infarction, atherosclerosis, and PAD, which constitute a major cause of death and morbidity in industrialized societies (53). Although angioplasty, bypass grafting, and other therapies are partly successful in restoring blood flow to affected tissues, there are currently no effective strategies to selectively generate new functional vessels to relieve ischemic stress, which can otherwise produce severe tissue necrosis and gangrene and necessitate amputation. In contrast, inhibiting angiogenesis with targeted therapies has shown some efficacy in treating solid tumors and macular degeneration, maladies typically associated with unregulated or excessive angiogenic activity (2). For example, the humanized VEGF antibody bevacizumab is an FDA-approved treatment for macular degeneration, in addition to being a first-line therapy for colon and non–small cell lung cancer (54). Similarly, the small-molecule VEGFR inhibitors sunitinib and sorafenib are approved for the treatment of renal carcinomas, as well as gastrointestinal stromal tumors and hepatocellular carcinomas, respectively (55). By elucidating the molecular mechanisms controlling endogenous responses to the hypoxic stress associated with ischemic attack and tumor growth, we hope to reveal novel targets with which to regulate angiogenesis therapeutically.

View Supplemental data

We thank Hongwei Yu for histological preparations; the M.C. Simon laboratory for helpful discussions and comments; Tom Kadesch for providing the Fc-Dll4 plasmid; and Regina Choe and the Joel Greenberg laboratory for their collaboration, particularly Wes Baker for his technical assistance. This work was funded by the Howard Hughes Medical Institute and the NIH (grants R01-NS060653, HL66130, and RR02305). M.C. Simon is an investigator of the Howard Hughes Medical Institute.

Address correspondence to: M. Celeste Simon, Howard Hughes Medical Institute, Abramson Family Cancer Research Institute, Department of Cell and Developmental Biology, Perelman School of Medicine at the University of Pennsylvania, 456 BRB II/III, 421 Curie Boulevard, Philadelphia, Pennsylvania 19104-6160, USA. Phone: 215.746.5532; Fax: 215.746.5511; E-mail: celeste2@mail.med.upenn.edu.

Conflict of interest: The authors have declared that no conflict of interest exists.

Reference information: J Clin Invest. 2012;122(4):1427–1443. doi:10.1172/JCI57322.

Anja Runge’s present address is: Division of Vascular Oncology and Metastasis, German Cancer Research Center, Heidelberg, Germany.

Liping Liu’s present address is: Merck Research Laboratories, West Point, Pennsylvania, USA.

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