Glucose-6-phosphate Dehydrogenase Modulates Vascular Endothelial Growth Factor-mediated Angiogenesis

Glucose-6-phosphate dehydrogenase (G6PD), the first enzyme of the pentose phosphate pathway, is the principal intracellular source of NADPH. NADPH is utilized as a cofactor by vascular endothelial cell nitric-oxide synthase (eNOS) to generate nitric oxide (NO•). To determine whether G6PD modulates NO•-mediated angiogenesis, we decreased G6PD expression in bovine aortic endothelial cells using an antisense oligodeoxynucleotide to G6PD or increased G6PD expression by adenoviral gene transfer, and we examined vascular endothelial growth factor (VEGF)-stimulated endothelial cell proliferation, migration, and capillary-like tube formation. Deficient G6PD activity was associated with a significant decrease in endothelial cell proliferation, migration, and tube formation, whereas increased G6PD activity promoted these processes. VEGF-stimulated eNOS activity and NO• production were decreased significantly in endothelial cells with deficient G6PD activity and enhanced in G6PD-overexpressing cells. In addition, G6PD-deficient cells demonstrated decreased tyrosine phosphorylation of the VEGF receptor Flk-1/KDR, Akt, and eNOS compared with cells with normal G6PD activity, whereas overexpression of G6PD enhanced phosphorylation of Flk-1/KDR, Akt, and eNOS. In the Pretsch mouse, a murine model of G6PD deficiency, vessel outgrowth from thoracic aorta segments was impaired compared with C3H wild-type mice. In an in vivo Matrigel angiogenesis assay, cell migration into the plugs was inhibited significantly in G6PD-deficient mice compared with wild-type mice, and gene transfer of G6PD restored the wild-type phenotype in G6PD-deficient mice. These findings demonstrate that G6PD modulates angiogenesis and may represent a novel angiogenic regulator. Glucose-6-phosphate dehydrogenase (G6PD), the first enzyme of the pentose phosphate pathway, is the principal intracellular source of NADPH. NADPH is utilized as a cofactor by vascular endothelial cell nitric-oxide synthase (eNOS) to generate nitric oxide (NO•). To determine whether G6PD modulates NO•-mediated angiogenesis, we decreased G6PD expression in bovine aortic endothelial cells using an antisense oligodeoxynucleotide to G6PD or increased G6PD expression by adenoviral gene transfer, and we examined vascular endothelial growth factor (VEGF)-stimulated endothelial cell proliferation, migration, and capillary-like tube formation. Deficient G6PD activity was associated with a significant decrease in endothelial cell proliferation, migration, and tube formation, whereas increased G6PD activity promoted these processes. VEGF-stimulated eNOS activity and NO• production were decreased significantly in endothelial cells with deficient G6PD activity and enhanced in G6PD-overexpressing cells. In addition, G6PD-deficient cells demonstrated decreased tyrosine phosphorylation of the VEGF receptor Flk-1/KDR, Akt, and eNOS compared with cells with normal G6PD activity, whereas overexpression of G6PD enhanced phosphorylation of Flk-1/KDR, Akt, and eNOS. In the Pretsch mouse, a murine model of G6PD deficiency, vessel outgrowth from thoracic aorta segments was impaired compared with C3H wild-type mice. In an in vivo Matrigel angiogenesis assay, cell migration into the plugs was inhibited significantly in G6PD-deficient mice compared with wild-type mice, and gene transfer of G6PD restored the wild-type phenotype in G6PD-deficient mice. These findings demonstrate that G6PD modulates angiogenesis and may represent a novel angiogenic regulator. Angiogenesis, the formation of new blood vessels in response to tissue ischemia or injury, is dependent upon a coordinated sequence of events involving vascular endothelial cell migration, proliferation, and tube formation (1Carmeliet P. Nat. Med. 2000; 6: 389-395Crossref PubMed Scopus (3490) Google Scholar, 2Vailhe B. Vittet D. Feige J.J. Lab. Invest. 2001; 81: 439-452Crossref PubMed Scopus (296) Google Scholar). Initially, vascular endothelial cells must acquire an angiogenic phenotype to migrate toward an angiogenic stimulus, proliferate behind the front of migration, and differentiate to form endothelial tubes and capillary-like structures. Nitric oxide (NO•) 1The abbreviations used are: NO•, nitric oxide; G6PD, glucose-6-phosphate dehydrogenase; eNOS, endothelial cell nitric-oxide synthase; VEGF, vascular endothelial growth factor; BAEC, Bovine aortic endothelial cells; AS, antisense; SS, scrambled sequence; PBS, phosphate-buffered saline; l-NAME, l-N G-nitroarginine methyl ester; l-NMMA, l-N G-monomethyl arginine citrate; hpf, high powered field; ROS, reactive oxygen species; WT, wild type; HEMI, hemizygous. has been shown to modulate angiogenesis by mediating growth factor-stimulated endothelial cell migration and proliferation. Nitric oxide is permissive for endothelial cell migration and enhances directional migration by inducing a switch from a stationary to a mobile phenotype (3Ziche M. Morbidelli L. Masini E. Amerini S. Granger H.J. Maggi C.A. Geppetti P. Ledda F. J. Clin. Invest. 1994; 94: 2036-2044Crossref PubMed Scopus (774) Google Scholar, 4Noiri E. Peresleni T. Srivastava N. Weber P. Bahou W.F. Peunova N. Goligorsky M.S. Am. J. Physiol. 1996; 270: C794-C802Crossref PubMed Google Scholar). In addition NO• promotes endothelial cell proliferation, and proliferating endothelial cells demonstrate increased expression of the endothelial isoform of nitric-oxide synthase (eNOS) compared with quiescent cells (5Zollner S. Aberle S. Harvey S.E. Polokoff M.A. Rubanyi G.M. Endothelium. 2000; 7: 169-184Crossref PubMed Scopus (18) Google Scholar). In vivo studies utilizing eNOS–/– mice have demonstrated the absolute requirement for endothelium-derived NO• for effective angiogenesis. In this murine model, compared with mice with normal eNOS activity, vascular endothelial growth factor (VEGF)-stimulated angiogenesis and vascular permeability are significantly attenuated (6Fukumura D. Gohongi T. Kadambi A. Izumi Y. Ang J. Yun C.O. Buerk D.G. Huang P.L. Jain R.K. Proc. Natl. Acad. Sci. U. S. A. 2001; 98: 2604-2609Crossref PubMed Scopus (813) Google Scholar). Glucose-6-phosphate dehydrogenase (G6PD), the first and rate-limiting enzyme in the pentose phosphate pathway, is the principal intracellular source of NADPH. NADPH, in turn, is utilized directly as a cofactor for eNOS and, indirectly, to maintain levels of another important cofactor, tetrahydrobiopterin, via de novo synthesis and the dihydrofolate reductase salvage pathway. In this manner, G6PD regulates eNOS activity and NO• levels. In this study, we demonstrate that G6PD activity modulates endothelial cell migration, proliferation, and tube formation by mediating NO• levels. G6PD may, therefore, serve as a novel regulatory determinant of the angiogenic phenotype. Antisense Transfection and Recombinant Adenovirus Gene Transfer—Bovine aortic endothelial cells (BAEC) (Cell Systems Co, Kirkland, WA) were grown to confluence in Dulbecco's modified Eagle's medium supplemented with 10% fetal bovine serum, 100 units/ml penicillin, and 100 μg/ml streptomycin. To decrease G6PD expression, an antisense phosphorothioate oligodeoxynucleotide to G6PD mRNA (AS) or a scrambled sequence control (SS) was utilized as described previously (7Leopold J.A. Cap A. Scribner A.W. Stanton R.C. Loscalzo J. FASEB J. 2001; 15: 1771-1773Crossref PubMed Scopus (125) Google Scholar). To increase G6PD expression, a recombinant adenovirus encoding rat G6PD cDNA under control of the cytomegalovirus promoter, AdG6PD, or an empty viral vector, Ad, were utilized as described previously (8Zhang Y.Y. Walker J.L. Huang A. Keaney J.F. Clish C.B. Serhan C.N. Loscalzo J. Biochem. J. 2002; 361: 267-276Crossref PubMed Scopus (48) Google Scholar). G6PD protein expression was determined by Western blotting (Fig. 1A), and corresponding NADPH levels were measured as described previously (7Leopold J.A. Cap A. Scribner A.W. Stanton R.C. Loscalzo J. FASEB J. 2001; 15: 1771-1773Crossref PubMed Scopus (125) Google Scholar, 8Zhang Y.Y. Walker J.L. Huang A. Keaney J.F. Clish C.B. Serhan C.N. Loscalzo J. Biochem. J. 2002; 361: 267-276Crossref PubMed Scopus (48) Google Scholar) (Fig. 1B). Experiments were conducted on cells from passages 4–10. Thymidine Incorporation Assay—BAEC (1 × 104) were seeded in triplicate 24-well plates, placed in reduced serum (1.0%) media for 24 h, and stimulated with vascular endothelial growth factor (VEGF) (100 ng/ml) in 1.0% serum for 12 h in the presence of 1 μCi/ml [methyl-3H]thymidine. After 12 h, cells were washed with PBS and fixed in 10% trichloroacetic acid at 4 °C for 16 h, washed with 100% ethanol, incubated with 1 n NaOH for 30 min at 37 °C, and neutralized with 10 n HCl. Thymidine incorporation was measured by liquid scintillation counting. Cell Migration—Migration was assessed by a cell wounding assay in BAEC grown to confluence in a P100 dish and synchronized in 1% serum for 24 h. A longitudinal incision was made in the midline of the plate with a sterile scalpel, and cells were scraped from one-half of the plate and stimulated with VEGF (100 ng/ml) for 12 h. After this time, cells that migrated across the midline were visualized using a Nikon TE 300 microscope, and 5 images per high powered field (hpf) per plate were captured digitally. Cells that crossed the midline were counted and averaged per plate. Data are presented as migrated cells/hpf. In addition, cell migration was assayed using a modified Boyden chamber (ChemoTx® plate, Neuro Probe, Inc., Gaithersburg, MD) according to the manufacturer's instructions. Tube Formation—In vitro formation of capillary-like tube structures was examined using Matrigel. Matrigel (0.5 ml) was polymerized on dual-chamber microscope slides. Cells were then plated on Matrigel in full-growth media for 1 h. Once the cells were seeded, the media were replaced with media containing 1% serum or VEGF (100 ng/ml). Tube formation was visualized using an inverted microscope (Nikon TE 300) equipped with digital imaging. For each treatment, 10 high power field images were captured, and area of tubes/networks formed was quantified using Scion Corp. (NIH Image) area analysis with background subtraction and averaged. Data are presented as density units. eNOS Activity—eNOS activity was measured in intact cells without the addition of exogenous cofactors as described previously (9Uittenbogaard A. Shaul P.W. Yuhanna I.S. Blair A. Smart E.J. J. Biol. Chem. 2000; 275: 11278-11283Abstract Full Text Full Text PDF PubMed Scopus (280) Google Scholar). Western Blotting—Western blot analyses were performed as described previously (9Uittenbogaard A. Shaul P.W. Yuhanna I.S. Blair A. Smart E.J. J. Biol. Chem. 2000; 275: 11278-11283Abstract Full Text Full Text PDF PubMed Scopus (280) Google Scholar). For analysis, equal amounts of cellular proteins (50 μg/lane) were resolved by SDS-PAGE and transferred to polyvinylidene difluoride membrane. After blocking in 5% milk solution, membranes were probed with anti-phosphotyrosine (clone 4G10) (Upstate Biotechnology, Inc.), anti-Flk-1/KDR (Santa Cruz Biotechnology), anti-phospho-Akt (Ser-473), anti-Akt, anti-phospho-eNOS (Ser-1177), or anti-eNOS antibodies (Cell Signaling Technologies) overnight at 4 °C. Membrane-bound antibodies were visualized by probing with horseradish peroxidase-conjugated secondary antibodies and visualized using the ECL detection system (Amersham Biosciences). Immunoprecipitation—Cells were incubated in lysis buffer for 1 h at 4 °C and centrifuged to remove insoluble material. The lysates were incubated overnight at 4 °C with protein-Sepharose A that had been incubated for 24 h with an antibody to eNOS (Transduction Laboratories) or G6PD (Sigma). After boiling, 100 μg of cell protein was loaded per lane, and samples were resolved by SDS-PAGE and transferred to polyvinylidene difluoride membrane. Blots were blocked with 5% skim milk and developed with antibodies for eNOS (1:500) or G6PD (1:1000) and visualized using the ECL detection system as described previously (7Leopold J.A. Cap A. Scribner A.W. Stanton R.C. Loscalzo J. FASEB J. 2001; 15: 1771-1773Crossref PubMed Scopus (125) Google Scholar). Real Time NO • Measurements—BAEC were grown to confluence in P100 dishes, washed twice with Dulbecco's phosphate-buffered saline, and placed in a balanced salt solution (130 mmol/liter NaCl, 5 mmol/liter KCl, 1 mmol/liter MgCl2, 1.5 mmol/liter CaCl2, 35 mmol/liter phosphoric acid, and 20 mmol/liter HEPES, pH 7.4) stimulated with VEGF (100 ng/ml). Real time NO• measurements were obtained using the inNO II Nitric Oxide Measuring System (Innovative Instruments) and the amino-FLAT probe. Data were recorded in real time; peak NO• levels were measured, and the area under the curve was integrated. 2′,7′-Dichlorodihydrofluorescein Diacetate Fluorescence—ROS accumulation was measured using 20 μm 6-carboxy-2′,7′-dichlorodihydrofluorescein diacetate di(acetoxymethyl) ester (Molecular Probes) as described previously (10Leopold J.A. Zhang Y.Y. Scribner A.W. Stanton R.C. Loscalzo J. Arterioscler. Thromb. Vasc. Biol. 2003; 23: 411-417Crossref PubMed Scopus (161) Google Scholar). Murine Aortic Ring Model of Angiogenesis—Murine thoracic aortas were harvested and placed in ice-cold Dulbecco's phosphate-buffered saline (PBS). The aorta was flushed with ice-cold PBS until free of blood. The adventitia was dissected free, and the aorta was cut into 1-mm rings. The rings were embedded in type I collagen gel such that the lumen was oriented horizontally and placed in MCDB 131 media and maintained at 37 °C. To assess the angiogenic response, the rings were visualized using a Nikon TE300 microscope, digital images captured, and image analysis performed using Scion Corp. (NIH Image) area analysis with background subtraction. Data are presented as density units (11Burbridge M.F. West D.C. Murray J.C. in Angiogenesis Protocols. Humana Press, Totowa, NJ2001: 185-204Google Scholar). G6PD-deficient Mouse Model—The Pretsch mouse, a murine model of G6PD deficiency on a C3H murine background (12Pretsch W. Charles D.J. Merkle S. Biochem. Genet. 1988; 26: 89-103Crossref PubMed Scopus (56) Google Scholar), was bred at our institution from frozen embryos obtained from Medical Research Council (Harwell, UK). Hemizygous (XbY) G6PD-deficient male mice (HEMI) and wild-type (XY) C3H control mice (WT) age 12–16 weeks were studied. Animals were genotyped as described previously (13Sanders S. Smith D.P. Thomas G.A. Williams E.D. Mutat. Res. 1997; 374: 79-87Crossref PubMed Scopus (39) Google Scholar). The animals were fed standard chow and handled following NIH guidelines. All procedures were approved by the Institutional Animal Care and Use Committee at Boston University Medical Center. The G6PD-deficient phenotype was confirmed by measuring hepatic G6PD activity and NADPH levels as described previously (7Leopold J.A. Cap A. Scribner A.W. Stanton R.C. Loscalzo J. FASEB J. 2001; 15: 1771-1773Crossref PubMed Scopus (125) Google Scholar). Liver was harvested from mice at the time of sacrifice, snap-frozen, and stored at –80 °C. G6PD and NADPH levels were measured in liver homogenates. In Vivo Matrigel Plug Assay and Cell Recovery—Matrigel (0.5 ml) was injected subcutaneously in the ventral groin area. One side was injected with Matrigel alone and the other with Matrigel mixed with VEGF (100 ng/ml). In this manner, each animal serves as its own control. After 14 days, the mice were euthanized by CO2 inhalation. Matrigel plugs were excised and fixed in 10% formalin, subjected to an ethanol dehydration series, and embedded in paraffin. Serial sections (10 μm) were cut using a cryotome and applied to glass slides. Slides were deparaffinized and stained with hematoxylin and eosin. To recover cells, the excised Matrigel plugs were minced with a sterile scalpel, passed 10 times through a 14-gauge needle, treated with BD™ Cell Recovery Solution for 1 h at 4 °C, centrifuged, and subjected to immunoprecipitation as described above to assay for proteins of interest. Immunohistochemistry—Immunohistochemistry on deparaffinized slides was performed using a rabbit polyclonal anti-von Willebrand factor antibody (Santa Cruz Biotechnology, Inc.) at 1:50, as described previously (14Eberhardt R.T. Forgione M.A. Cap A. Leopold J.A. Rudd M.A. Trolliet M. Heydrick S. Stark R. Klings E.S. Moldovan N.I. Yaghoubi M. Goldschmidt-Clermont P.J. Farber H.W. Cohen R. Loscalzo J. J. Clin. Invest. 2000; 106: 483-491Crossref PubMed Scopus (376) Google Scholar). An angiogenic response was quantified by cell counts from 10 high power fields per section and image analysis using Scion Corp. (NIH Image) with background subtraction to determine the area occupied by endothelial cells. Statistical Analysis—Continuous data were expressed as mean ± S.E. Comparison between groups was performed by Student's paired two-tailed t test. Two-way analysis of variance was used to examine differences in response to treatments between groups, with post hoc analysis performed by the method of Student-Newman-Keuls. A p value of <0.05 was considered significant. G6PD and Endothelial Cell Proliferation—To examine the role of G6PD in VEGF-mediated endothelial cell proliferation, BAEC were transfected with an antisense oligodeoxynucleotide to G6PD mRNA with a 36% transfection efficiency to decrease G6PD expression and activity by 76% (AS-EC) or a scrambled control sequence (SS-EC), and cells were stimulated with VEGF (100 ng/ml) for 12 h. Compared with SS-EC, AS-EC demonstrated a significant decrease in basal and VEGF-stimulated [3H]thymidine incorporation (Fig. 2A). The observed decrease in cell proliferation was not the result of increased cell death as determined by lactate dehydrogenase activity in the media (data not shown). To determine the contribution of NO• to VEGF-stimulated endothelial cell proliferation, SS-EC and AS-EC were pretreated for 1 h with l-NAME (1 mmol/liter), to inhibit both NO• and superoxide production by eNOS, or l-NMMA (100 μmol/liter), to inhibit NO• generation by eNOS, and stimulated with VEGF in the presence of these inhibitors. In SS-EC stimulated with VEGF, [3H]thymidine incorporation was decreased significantly in cells treated with l-NAME (53,874 ± 3,848 versus 41,932 ± 5,330 cpm, p < 0.01) or l-NMMA (53,874 ± 3,848 versus 44,187 ± 3,267 cpm, p < 0.01). Similarly, AS-EC stimulated with VEGF demonstrated a decrease in [3H]thymidine incorporation in the presence of l-NAME (32,156 ± 4,561 versus 26,987 ± 2,142 cpm, p < 0.05) and l-NMMA (32,156 ± 4,561 versus 26,116 ± 4,668 cpm, p < 0.05). We next sought to determine whether overexpression of G6PD would augment cell proliferation. To perform these studies, BAEC were infected with AdG6PD (AdG6PD-EC) at a multiplicity of infection = 10 plaque-forming units/cell with a 90–95% infection efficiency to increase G6PD activity 5-fold. Compared with Ad-transfected endothelial cells (Ad-EC), overexpression of G6PD markedly enhanced [3H]thymidine incorporation under basal conditions, and this response was enhanced further by VEGF (Fig. 2B). In AdG6PD-EC, the addition of l-NAME significantly decreased cell proliferation in unstimulated (52,178 ± 2,784 versus 38,146 ± 5,399 cpm, p < 0.009) and VEGF-treated cells (79,227 ± 4,110 versus 46,665 ± 5,294 cpm, p < 0.001), suggesting that increased G6PD activity promoted cell proliferation by increasing NO• levels. G6PD and Endothelial Cell Migration—To examine the influence of G6PD on VEGF-stimulated endothelial cell migration, we performed a cell wounding assay. SS-EC and AS-EC were incubated with VEGF, and cell migration across the midline was observed. After 12 h, VEGF markedly increased cell migration across the midline in SS-EC (44 ± 4 versus 142 ± 16 cells/hpf, p < 0.001), an effect that was not observed in AS-EC (27 ± 11 versus 35 ± 5 cells/hpf, p = not significant) (Fig. 3). These findings were confirmed utilizing a modified Boyden chamber assay to determine whether G6PD activity influenced directed VEGF-mediated endothelial cell migration. SS-EC and AS-EC were stimulated with VEGF (100 ng/ml) for 12 h during which time cells could migrate across the membrane. In SS-EC treated with VEGF, there was a significant increase in fluorescence (13.3 ± 2.6 versus 55.3 ± 6.6 units, p < 0.001), indicating an increase in cell migration, that was abrogated in AS-EC with deficient G6PD activity (14.1 ± 4.8 versus 20.2 ± 9.9 units, p = not significant). We next sought to determine the contribution of NO• to endothelial cell migration. By utilizing the modified Boyden chamber assay, SS-EC or AS-EC were stimulated with VEGF for 12 h in the presence or absence of l-NAME (1 mmol/liter) or l-NMMA (100 μmol/liter). In SS-EC, l-NAME significantly decreased cell migration in VEGF-stimulated cells (51.9 ± 4.2 versus 23.6 ± 7.0 units, p < 0.01) as did l-NMMA (51.9 ± 4.2 versus 21.6 ± 7.3 units, p < 0.003). Interestingly, in AS-EC, VEGF did not significantly increase endothelial cell migration, and l-NAME or l-NMMA did not alter this response. To determine whether overexpression of G6PD would promote directed endothelial cell migration, we overexpressed G6PD in endothelial cells and examined migration in a modified Boyden chamber assay. Compared with Ad-EC, AdG6PD-EC demonstrated a significant increase in fluorescence under basal conditions (16.4 ± 3.2 versus 35.1 ± 4.7 units, p < 0.009) and following stimulation with VEGF (51.9 ± 3.1 versus 98.8 ± 7.1 units, p < 0.001). In the presence of l-NAME, fluorescence was decreased in both unstimulated AdG6PD-EC (35.1 ± 4.7 versus 21.5 ± 3.4 units, p < 0.01) and VEGF-stimulated cells (98.8 ± 10.1 versus 36.4 ± 4.3 units, p < 0.001), suggesting that increased G6PD expression mediates endothelial cell migration by increasing NO• levels. G6PD and Endothelial Tube Formation—As endothelial cell proliferation and migration are processes integral to the formation of capillary-like tube structures, we examined the effect of G6PD activity on tube formation using an in vitro Matrigel assay. SS-EC and AS-EC were plated on Matrigel and stimulated with VEGF for 12 h. Under basal conditions, SS-EC plated on Matrigel formed tubes and networks, and the area occupied by SS-EC endothelial tubes was increased significantly following exposure to VEGF (9,911 ± 640 versus 24,729 ± 3,311 units, p < 0.001). Similarly, AS-EC formed endothelial tubes and the area occupied by these networks was not significantly different from that observed with SS-EC (9,911 ± 640 versus 8,424 ± 1,005 units, p = not significant); however, when stimulated with VEGF, AS-EC tube formation was decreased significantly compared with SS-EC (10,554 ± 895 versus 24,729 ± 3,311 units, p < 0.002) (Fig. 4A). To examine the role of NO• in endothelial tube formation, we performed the in vitro Matrigel assay in the presence of l-NAME or l-NMMA. In VEGF-stimulated SS-EC, the area occupied by endothelial tubes was decreased significantly in the presence of l-NAME (24,729 ± 3,311 versus 16,734 ± 2,396 units, p < 0.001) or l-NMMA (24,729 ± 3,311 versus 17,789 ± 1,923 units, p < 0.001). In contrast, there was no significant difference in the area occupied by endothelial tubes in VEGF-stimulated AS-EC in the presence of l-NAME or l-NMMA. As G6PD overexpression was associated with enhanced endothelial cell proliferation and migration, it is not surprising that compared with Ad-EC, tube formation was significantly increased in AdG6PD-EC under basal conditions (10,372 ± 1,286 versus 19,549 ± 1,502 units, p < 0.01). This response was augmented further following stimulation with VEGF (24,994 ± 2,533 versus 43,212 ± 4,208 units, p < 0.001) (Fig. 4B). Furthermore, NO• played a significant role in this effect as the addition of l-NAME to AdG6PD-EC decreased tube formation in unstimulated (19,549 ± 1,502 versus 14,889 ± 1,321 units, p < 0.005) as well as in VEGF-stimulated (43,212 ± 4,208 versus 21,956 ± 5,367 units, p < 0.001) cells. G6PD, eNOS Activity, and NO • Levels—These observations suggest that one mechanism by which G6PD influences endothelial cell proliferation, migration, and tube formation is to regulate eNOS activity and, thereby, bioavailable NO•. To examine the effect of G6PD expression on eNOS activity, we measured eNOS activity in intact cells (without the addition of exogenous cofactors) (Fig. 5A). In AS-EC, compared with SSEC, eNOS activity was significantly decreased (12,728 ± 488 versus 18,534 ± 250 cpm l-[3H]citrulline, p < 0.004) at base line and following stimulation with VEGF (18,622 ± 511 versus 29,749 ± 630 cpm l-[3H]citrulline, p < 0.001). In contrast, overexpression of G6PD significantly increased eNOS activity at base line compared with Ad-EC (14,621 ± 201 versus 42,766 ± 634 cpm l-[3H]citrulline, p < 0.001), and this response was augmented further following exposure to VEGF (23,772 ± 554 versus 62,858 ± 632 cpm l-[3H]citrulline, p < 0.001). This effect was not a consequence of G6PD on eNOS expression (Fig. 5D) (15Bouloumie A. Schini-Kerth V.B. Busse R. Cardiovasc. Res. 1999; 41: 773-780Crossref PubMed Scopus (231) Google Scholar). To determine the effect of decreased or increased G6PD expression on VEGF-mediated NO• production, we measured real time NO• levels using a nitric oxide probe (Fig. 5B). Peak NO• levels were significantly lower in AS-EC compared with SS-EC (14.5 ± 1.5 versus 64.2 ± 10.0 nmol/liter, p < 0.008) as was the integral of area under the curve (2,056.3 ± 471.1 versus 11,790.7 ± 3,104.1 nmol/liter/s, p < 0.04). In contrast, peak NO• levels were significantly higher in AdG6PD-EC compared with Ad-EC (59.8 ± 8.9 versus 211.1 ± 28.0 nmol/liter, p < 0.001) as was the integral of area under the curve (9,822.0 ± 2,992.5 versus 90,123.3 ± 2,667.4 nmol/liter, p < 0.001). To examine the mechanism(s) by which G6PD mediates NO• production, we first performed immunoprecipitation studies to determine whether G6PD co-localizes with eNOS. Under basal conditions, G6PD and eNOS co-localize, and following stimulation with VEGF, co-localization is enhanced (Fig. 5C). We next sought to determine whether G6PD influences activation of the VEGF receptor Flk-1/KDR to modulate Akt-eNOS activation. Interestingly, in AS-EC, tyrosine phosphorylation of a 230-kDa band, consistent with Flk-1/KDR, was decreased at both 5 and 15 min compared with what was observed in SS-EC resulting in decreased phosphorylation of Akt and eNOS in AS-EC compared with SS-EC (Fig. 5D). In contrast, in AdG6PD-EC, tyrosine phosphorylation of the 230-kDa band was enhanced at 5 and 15 min resulting in increased phosphorylation of Akt and eNOS compared with Ad-EC (Fig. 5D). G6PD, VEGF, and ROS Accumulation—It has been demonstrated recently that VEGF promotes angiogenesis in human umbilical vein endothelial cells by stimulating NAD(P)H oxidase to increase ROS formation (16Ushio-Fukai M. Tang Y. Fukai T. Dikalov S.I. Ma Y. Fujimoto M. Quinn M.T. Pagano P.J. Johnson C. Alexander R.W. Circ. Res. 2002; 91: 1160-1167Crossref PubMed Scopus (427) Google Scholar). Therefore, we measured ROS levels by 2′,7′-dichlorodihydrofluorescein diacetate fluorescence in endothelial cells with decreased or increased G6PD activity to determine whether our observations could be explained by changes in ROS levels. Interestingly, in AS-EC exposed to VEGF, ROS levels were significantly higher than in SS-EC (248 ± 27 versus 180 ± 22 units, p < 0.01), and conversely, in AdG6PD-EC, ROS levels were lower than in Ad-EC (156 ± 14 versus 197 ± 12 units, p < 0.01). These findings suggest that G6PD, which is recognized as an antioxidant enzyme in endothelial cells, may additionally modulate ROS accumulation by influencing NO• production to mediate the redox milieu to a level favorable for endothelial cell proliferation, migration, and tube formation. Ex Vivo Aorta Implants and Vessel Outgrowth—We next examined the effect of G6PD on VEGF-mediated angiogenesis using an in vivo model of G6PD deficiency, the Pretsch mouse. Compared with C3H background mice (WT), hemizygous male G6PD mice (HEMI) demonstrate only 31% of WT G6PD activity with a concomitant reduction in NADPH levels (0.6 ± 0.03 versus 0.3 ± 0.01 mmol/mg protein, p < 0.001). To examine specifically the effect of G6PD on aortic vessel outgrowth, we explanted thoracic aortas from WT and HEMI mice, sectioned them into rings, and embedded the rings in a collagen matrix. The rings were then observed over 6 days for vessel outgrowth. By day 3, there was significant outgrowth from the WT rings compared with the rings from the HEMI mice, and by day 6, there was a marked difference between outgrowth observed from WT and HEMI mice rings (Fig. 6). To quantify these observations, the area occupied by vessel outgrowth was determined using area image analysis with background subtraction. At day 3, the area occupied by vessel outgrowth was significantly greater in aortic rings from WT compared with HEMI mice (6,037 ± 975 versus 2,767 ± 391 units, p < 0.04), and this effect was more pronounced by day 6 (10,057 ± 713 versus 3,475 ± 295 units, p < 0.001). In Vivo Matrigel Assay—To examine the effect of G6PD activity on angiogenesis in vivo, we performed an in vivo Matrigel migration assay in WT and HEMI mice. Each mouse was injected on one side with either Matrigel alone or Matrigel supplemented with VEGF, and the plugs were examined after 14 days. In WT mice, there is noticeable cell migration into the Matrigel plug, and this response is increased in Matrigel supplemented with VEGF (12 ± 1 versus 52 ± 2 cells/hpf, p < 0.001). In contrast, in HEMI mice, there is a marked decrease in cell migration into the Matrigel plug compared with WT mice, and exposure to VEGF only modestly improved this response (6 ± 1 versus 9 ± 1 cells/hpf, p < 0.03) (Fig. 7A). These findings confirm that G6PD modulates angiogenesis in vivo. To determine that the cells that migrated into the Matrigel were endothelial cells, the M