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Originally published In Press as doi:10.1074/jbc.M300643200 on June 4, 2003

J. Biol. Chem., Vol. 278, Issue 33, 31000-31006, August 15, 2003
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AMP-activated Protein Kinase (AMPK) Signaling in Endothelial Cells Is Essential for Angiogenesis in Response to Hypoxic Stress*

Daisuke Nagata {ddagger}, Masaki Mogi and Kenneth Walsh §

From the Molecular Cardiology/Whitaker Cardiovascular Institute, Boston University School of Medicine, Boston, Massachusetts 02118

Received for publication, January 21, 2003 , and in revised form, May 29, 2003.


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
AMP-activated protein kinase (AMPK) is a stress-activated protein kinase that is regulated by hypoxia and other cellular stresses that result in diminished cellular ATP levels. Here, we investigated whether AMPK signaling in endothelial cells has a role in regulating angiogenesis. Hypoxia induced the activating phosphorylation of AMPK in human umbilical vein endothelial cells (HUVECs), and AMPK activation was required for the maintenance of pro-angiogenic Akt signaling under these conditions. Suppression of AMPK signaling inhibited both HUVEC migration to VEGF and in vitro differentiation into tube-like structures in hypoxic, but not normoxic cultures. Dominant-negative AMPK also inhibited in vivo angiogenesis in Matrigel plugs that were implanted subcutaneously in mice. These data identify AMPK signaling as a new regulator of angiogenesis that is specifically required for endothelial cell migration and differentiation under conditions of hypoxia. As such, endothelial AMPK signaling may be a critical determinant of blood vessel recruitment to tissues that are subjected to ischemic stress.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Angiogenesis is central feature of normal embryonic and post-natal development, and this process plays a critical role in the neovascularization that is associated with tumor growth and occlusive vascular diseases (1–3). A large body of evidence has shown that vascular angiogenic growth factors promote angiogenesis through activation of MAP kinase (4) and phosphatidylinositol 3-kinase (PI3K)-Akt/PKB (5) intracellular signaling pathways. However, the signaling molecules involved in angiogenic cellular responses under conditions of hypoxic stress is incompletely understood.

AMP-activated protein kinase (AMPK)1 is a metabolite-sensing protein kinase that shares amino acid sequence homology with yeast SNF1 (6, 7). In myocytes, AMPK is activated by increases in the AMP:ATP ratio, which is brought about by hypoxia/anoxia (8–10), vigorous exercise and muscle contraction (11–14) or pressure-overload hypertrophy (15). Under these conditions, AMPK is activated by a conformational change after binding AMP (16, 17), and by phosphorylation by its upstream kinase AMPK kinase (AMPKK) (17–19). Upon activation, AMPK phosphorylates and down-regulates several anabolic enzymes including 3-hydroxy-3-methylglutaryl-CoA (HMG-CoA) reductase (16, 20), acetyl-CoA carboxylase (ACC) (16, 21), sn-glycerol-3-phosphate acetyltransferase (22, 23), and glycogen synthase (24); thereby diminishing metabolite flux through synthetic pathways that consume ATP. AMPK activation also accelerates {beta}-oxidation of fatty acid, which promotes ATP production (25, 26).

Endothelial nitric-oxide synthase (eNOS) residue 1177 (in human) is a substrate for both Akt (27, 28) and AMPK (29). Phosphorylation of eNOS at this residue leads to enzyme activation and nitric oxide (NO) production. AMPK is reported to phosphorylate eNOS in cardiac myocytes under hypoxic conditions (29), but it is not clear whether NO production by endothelial cells is regulated by this reaction. Recently, AMPK was detected in human umbilical vein endothelial cells (HUVECs) (30). The pharmacological stimulator of AMPK, 5-aminoimidazole-4-carboxamide-1-{beta}-D-ribofuranoside (AICAR), protects HUVECs from apoptosis under hyperglycemic conditions (31). Interestingly, AICAR was found to reverse the inhibitory effect of hyperglycemia on mitogen-stimulated Akt phosphorylation.

In this study we analyzed whether AMPK signaling plays a role in angiogenesis. Specifically, we tested whether AMPK is essential for endothelial cell migration, differentiation into capillary-like structures, and NO production under both normoxic and hypoxic conditions. We also tested whether AMPK signaling is required for angiogenesis in Matrigel plugs that were subcutaneously implanted in mice. The results of this study suggest that cross-talk between AMPK and Akt is essential for angiogenesis under conditions of hypoxic stress, but dispensable for angiogenic cellular responses in normoxic endothelial cells.


   EXPERIMENTAL PROCEDURES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
RESULTS
 DISCUSSION
 REFERENCES
 
Materials—Phospho-AMPK (Thr-172), pan-{alpha}-AMPK, and phospho-Akt (Ser-473), phospho-eNOS (Ser-1177), and phospho-eNOS (Thr-495) antibody were purchased from Cell Signaling Technology (Beverly, MA). Phospho-Acetyl CoA Carboxylase (ACC) (Ser-79) and c-Myc tag antibody was purchased from Upstate Biotechnology (Lake Placid, NY). ACC protein antibody was from Alpha Diagnostic International (San Antonio, TX). Akt and eNOS antibodies were purchased from Santa Cruz Biotechnology. N{omega}-nitro-L-arginine methyl ester (L-NAME), sodium nitroprusside (SNP), and human vascular endothelial growth factor (VEGF) were purchased from Sigma.

Adenovirus—A replication-defective adenoviral vector expressing dominant-negative AMPK was a gift from Dr. Birnbaum (University of Pennsylvania). This vector expresses the rat AMPK {alpha}2-subunit cDNA, whose lysine 45 residue was changed to arginine (14), fused in-frame with the c-Myc epitope tag. Details of Ad-caAkt and Ad-dnAkt construction were previously described (32, 33). The encoded proteins are fused to the hemagglutinin (HA) epitope. Adenoviral vector expressing green fluorescence protein (GFP) was obtained from Qbiogene (Illkirch, France).

Cell Culture and Adenoviral Vectors Transduction—Human umbilical vein endothelium cells (HUVECs), endothelial cell basal medium (EBM), and endothelial cell growth medium (EGM) were purchased from Clonetics (San Diego, CA). HUVECs were cultured in EGM with 5% fetal bovine serum, 100 units/ml penicillin, and 100 µg/ml streptomycin. All experiments utilized HUVECs at passage 6 or less. HUVECs were transduced with the indicated replication-defective adenoviral vectors at a multiplicity of infection (m.o.i.) of 50 plaque-forming units for 1 day. The media was then changed to EBM containing 1% fetal bovine serum to reduce the effects of stimulation by serum mitogens. After a 1-day incubation in low mitogen media, transduced cells were placed in a hypoxia chamber (GasPak anaerobic system; BD Biosciences) according to the manufacturer's instructions for the indicated period of time. This system induces anaerobic conditions of less than 2% of oxygen and more than 4% of carbon dioxide by using carbon dioxidegenerating reagents (34). All experiments utilized HUVECs at passage 6 or less.

Western Blot Analysis—Western blot analysis was carried out as previously described (35). In brief, cell lysates were extracted with Nonidet P-40 lysis buffer (50 mM Tris-HCl pH 8.0, 150 mM NaCl, 1% Nonidet P-40, 2 µg/ml aprotinin, 2 µg/ml leupeptin, 1 mM phenylmethylsulfonyl fluoride, and 1 mM sodium orthovanadate) followed by SDS-PAGE. The membranes were immunoblotted with the indicated antibodies at a 1:1000 dilution followed by the secondary antibody conjugated with horseradish peroxidase at a 1:5000 dilution. ECL-PLUS Western blotting detection kit (Amersham Biosciences) was used for detection.

Migration Assay—Migration assays were performed as described previously using a modified Boyden chamber (Neuroprobe, Cabin John, MD) (36). HUVECs infected with the indicated adenoviral vectors were serum-starved overnight in serum-free media (EBM). Transduced cells were suspended with 0.05% trypsin-0.53 mM EDTA (Invitrogen), washed with phosphate-buffered saline, and resuspended in EBM. EBM containing VEGF 150 ng/ml was put into the wells of the lower chamber. Polycarbonate filters with 8-µm pores (PVPF; OSMONICS, Kent, WA) coated with 0.5% gelatin were used to separate the lower chamber and 1 x 105 suspended cells in the upper chamber. For some experiments, the indicated concentrations of SNP were added to both chambers. The chambers were then incubated for 8 h at 37 °C under normoxic or hypoxic conditions. Filters were carefully removed and the cells attached to the upper side were removed by wiping. Cells migrating through the filter (cells on the lower side of filter) were fixed with methanol for 15 min, stained with Giemsa stain solution (Sigma) and five random microscopic fields per well were quantified. Each experiment was performed in duplicate, and three separate experiments were performed.

Tube Formation Assay—The formation of vascular-like structures by HUVECs on growth factor-reduced Matrigel (BD Biosciences) was performed as previously described (37). 24-well culture plates were coated with Matrigel according to the manufacturer's instructions. The indicated adenovirus-transduced HUVECs were seeded on coated plates at 5 x 104 cells/well in EBM containing VEGF 50 ng/ml and incubated at 37 °C for 18 h under normoxic or hypoxic conditions. In some wells, SNP (1 µM-1 mM) was also added. Tube formation was observed using an inverted phase contrast microscope (Nikon, Tokyo, Japan). Images were captured with a video graphic system (DEI-750 CE Digital Output Camera, Optronics, Goleta, CA). The degree of tube formation was quantified by measuring the length of tubes in 3 randomly chosen low power fields (x100) from each well using the NIH Image Program. Each experiment was repeated for 4 times.

cGMP Assay—Intracellular cGMP concentration was measured using cGMP enzyme immunoassay system purchased from Amersham Biosciences according to the manufacturer's instruction. After adenovirus infection, cells were incubated overnight in low serum medium to reduce the effect of mitogens. Then VEGF (50 ng/ml) was added to the cultures in the absence or presence of NO synthase inhibitor, L-NAME (1 mg/ml). Cells were then incubated under hypoxic or normoxic conditions for 6 h prior to the cGMP measurement. Each experiment was performed in four independent samples, and two sets of separate experiments were performed.

Mouse Angiogenesis Assay—The formation of new vessels in vivo was evaluated by Matrigel plug assay as described previously (38, 39). For these experiments, 400 µl of Matrigel containing bFGF (250 ng/ml) and replication-defective adenoviral vectors encoding GFP, dominant-negative AMPK, or dominant-negative Akt (1 x 108 plaque forming units) was cooled on ice and then injected subcutaneously into the abdomen of C57BL mice. Mice were sacrificed 14 days after the injection. The Matrigel plugs with adjacent subcutaneous tissues were carefully recovered by en bloc resection, fixed in 4% paraformaldehyde, dehydrated with 30% sucrose, and embedded in OCT compound (GTI Microsystems, Tempe, AZ) in liquid nitrogen. Masson's trichrome staining and immunohistostaining for CD31 (PECAM-1: BD Biosciences) were performed on adjacent frozen sections. Primary antibody was used at a 1:50 dilution followed by incubation of secondary antibody (horseradish peroxidase-conjugated anti-rat IgG at a 1:100 dilution). The AEC Substrate Pack (Biogenex, San Ramon, CA) was used for detection. CD31-positive capillaries were counted in 4 randomly chosen low-power (x100) microscopic fields. Each experiment was repeated four times.

Statistical Analyses—Values were expressed as the mean ± S.E. Statistical comparisons were performed using analysis of variance (ANOVA) with Scheffe's F procedure for post-hoc analysis.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
DISCUSSION
 REFERENCES
 
AMPK Activation by Hypoxia in Endothelial Cells—To test whether AMPK is activated in endothelial cells under conditions of hypoxia, HUVEC cultures were incubated in a hypoxia chamber for different lengths of time and Western immunoblot analyses were performed on lysates using an antibody that is specific for AMPK phosphorylation at Thr-172 of the {alpha}-subunit. A basal level of AMPK phosphorylation was detected in cell cultured under normoxic conditions, and hypoxia produced a time-dependent increase in AMPK phosphorylation (Fig. 1A). A representative Western blot of this time course is shown (Fig. 1A). Thr-172 phosphorylation leads to AMPK activation, and the induction of AMPK signaling by hypoxia was indicated by the time-dependent phosphorylation of ACC at Ser-79 (Fig. 1B), which is a substrate for AMPK. Hypoxia also led to a time-dependent increase in AMPK activity as assessed by SAMS peptide phosphorylation with immunoprecipitated AMPK (data not shown). Basal and hypoxia-inducible phosphorylation of ACC was blocked when cells were transduced at 50 m.o.i. with an adenoviral vector expressing a dominant-negative mutant of AMPK (dominant-negative AMPK) that contains a c-Myc epitope tag (Fig. 1, A and B). Parallel cultures incubated with an adenoviral vector expressing the GFP marker protein at 50 m.o.i. revealed greater than a 95% transduction efficiency. Transduction with dominant-negative AMPK also inhibited the hypoxia-induced phosphorylation of endogenous AMPK (Fig. 1A). Hypoxia did not affect the total level of {alpha}-subunit expression in immunoblot analyses using a pan-{alpha}-AMPK antibody (Fig. 1A), nor did it affect the level of ACC protein expression (data not shown).



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  		FIG. 1.
Regulation of AMPK phosphorylation by hypoxia and dominant-negative AMPK in endothelial cells. A, Western blot analyses were performed on 75 µg of protein in each lane. Hypoxia was induced for the indicated lengths of time and AMPK phosphorylation at Thr-172 (p-AMPK) and ACC phosphorylation of Ser-79 (p-ACC) was assessed. Parallel cultures were transduced with an adenoviral vector expressing dominant-negative AMPK tagged with c-Myc (dnAMPK), 2 days prior to the initiation of hypoxia. Control cultures were transduced with an adenoviral vector expressing GFP under identical conditions. Representative blots of 3–4 experiments each are shown. B, phosphorylation levels of ACC in the presence (dashed lines, empty symbols) or absence (solid line, filled symbols) of dominant-negative AMPK were quantified using the NIH Image program. C, phosphorylation levels of Akt are shown. Immunoblots were normalized to total loaded protein. *, p < 0.01; #, p < 0.05 versus control.

 

Hypoxia led to a modest reduction in basal Akt phosphorylation at Ser-473, which is required for kinase activity (40, 41) (Fig. 1A). Transduction with dominant-negative AMPK further suppressed Akt phosphorylation, inhibiting Akt phosphorylation at every time point examined (Fig. 1C). The relative magnitude of the inhibition by dominant-negative AMPK increased as the length of exposure to hypoxia increased.

Regulation of eNOS Ser-1177 Phosphorylation and NO Production by AMPK—The production of endothelial NO is important for vascular homeostasis and angiogenesis (42–46). VEGF stimulation promotes Akt signaling which, in turn, produces NO through the activating phosphorylation of eNOS at residue 1177 (27). Because AMPK is also reported to phosphorylate eNOS at residue 1177 in cardiac myocytes (29), the contribution of AMPK to VEGF-stimulated eNOS phosphorylation was analyzed in HUVEC under normoxic and hypoxic conditions (Fig. 2A). In our assays, VEGF did not affect the level of AMPK expression or phosphorylation (data not shown). However, transduction with dominant-negative AMPK effectively blocked eNOS phosphorylation under conditions of hypoxia, whereas it had little effect in normoxic HUVEC cultures (Fig. 2B). While basal levels of AMPK in normoxic endothelial cells has little effect on the status of eNOS 1177 phosphorylation, AMPK signaling was essential for ACC phosphorylation under these conditions (Fig. 2A). A vector expressing dominant-negative Akt was used as a positive control in these experiments because it is known to inhibit VEGF-stimulated eNOS phosphorylation in normoxic endothelial cell cultures (27). In contrast to dominant-negative AMPK, transduction with dominant-negative Akt significantly inhibited eNOS phosphorylation at Ser-1177 in normoxic cultures. Transduction with dominant-negative Akt also inhibited eNOS phosphorylation in hypoxic cultures. However, it had no effect on ACC phosphorylation under any condition.



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  		FIG. 2.
AMPK-mediated regulation of eNOS Ser-1177 phosphorylation in hypoxic endothelial cells. Relative levels of eNOS phosphorylation at Ser-1177 and Thr-495, and ACC phosphorylation were assessed by Western blot analyses. Parallel cultures were maintained under normoxic conditions or exposed to hypoxia for 6 h. Some cultures were exposed to 50 ng/ml VEGF for 6 h prior to lysate preparation. Cultures were also transduced with adenoviral vectors expressing dominant-negative AMPK (dnAMPK), dominant-negative Akt (dnAkt), constitutively-active Akt (caAkt), or GFP (control). A, Western immunoblots were performed by loading 100 µg of lysate in each lane. Expression from the AMPK and Akt adenoviral vectors was checked by anti-c-Myc and anti-HA immunoblot, respectively. B, relative phosphorylation levels of eNOS Ser-1177 in the presence or absence of VEGF, dominant-negative AMPK or dominant-negative Akt were quantified using the NIH Image program. Immunoblots were normalized to total loaded protein. C, relative phosphorylation levels of eNOS Thr-495. *, p < 0.01 relative to VEGF alone.

 

Recently, it has been shown that eNOS activity is negatively regulated through phosphorylation of Thr-495 (29, 47, 48). In normoxic cultures, VEGF stimulation led to a decrease in Thr-495 phosphorylation (Fig. 2, A and C), consistent with a previous report (47). The reduction in Thr-495 phosphorylation by VEGF was reversed by transduction with dominant-negative Akt, while dominant-negative AMPK had no effect. Similar trends in Thr-495 phosphorylation were also seen in hypoxic cultures, but the overall magnitude of the VEGF effect was less than that seen in normoxic cultures. Furthermore, dominant-negative AMPK had no detectable effect on decline in Thr-495 under conditions of hypoxia.

The production of NO by eNOS was measured in intact HUVEC cultures by monitoring the accumulation of cGMP (39). Under the conditions of these assays, VEGF-induced cGMP accumulation was similar between HUVEC cultures incubated under normoxic and hypoxic conditions (Fig. 3). Under both conditions, cGMP accumulation was largely inhibited by the eNOS inhibitor L-NAME. Transduction with dominant-negative AMPK significantly inhibited NO output in hypoxic cultures. There was a trend toward lower NO output in normoxic cultures transduced with dominant-negative AMPK, but this was not statistically significant (p = 0.1). In contrast, transduction with dominant-negative Akt significantly inhibited NO output in both hypoxic and normoxic HUVEC cultures. Transduction with dominant-negative Akt or dominant-negative AMPK did not affect basal NO production in the absence of VEGF in either hypoxic or normoxic cultures.2



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  		FIG. 3.
AMPK-mediated regulation of NO production in hypoxic endothelial cells. Intracellular cGMP was measured to evaluate the regulation of VEGF-induced endothelial NO production by AMPK and Akt under normoxic or hypoxic conditions. Parallel cultures were maintained under conditions of normoxia or exposed to hypoxia for 6 h in the absence of VEGF (cross-hatch). Other cultures were stimulated with VEGF (50 ng/ml) in the presence (dots) or absence (solid) of the eNOS inhibitor L-NAME. VEGF-stimulated cultures were transduced with dominant-negative Akt (dnAkt, open) or dominant-negative AMPK (dnAMPK, hatch) 2 days prior to the experiment. Control, VEGF alone and VEGF + L-NAME cultures were transduced with an adenoviral vector expressing GFP. Four independent experiments were performed. *, p < 0.01 versus control.

 

AMPK Signaling Is Essential for Endothelial Cell Migration under Hypoxic Conditions—To evaluate the role of AMPK signaling on endothelial cell migration, HUVEC were transduced with Ad-dnAMPK and tested for their ability to migrate toward VEGF (150 ng/ml) in a Boyden chamber apparatus. Under hypoxic conditions, transduction with dominant-negative AMPK significantly suppressed cell migration relative to cells transduced with GFP control (Fig. 4). No differences in cell migration were detected between Ad-GFP-treated and nontransduced cells2. Furthermore, the reduction in cell migration did not appear to be due to decreased cell viability. Under these cell culture conditions dominant-negative AMPK-transduced cells did not display detectable increases in the frequencies of TUNEL- or pyknotic nuclei-positive cells, nor was there a detectable decrease in mitochondrial function (data not shown). Addition of 10 µM sodium nitroprusside (SNP), an NO donor, partially reversed the inhibitory action of dominant-negative AMPK on HUVEC migration to VEGF in the hypoxic cultures (Fig. 4). Similar effects were also observed at 1 and 100 µM SNP (data not shown). However, co-transfection with an adenoviral vector expressing a constitutively-active form of Akt was more effective than the NO donor at reversing the inhibition by dominant-negative AMPK in hypoxic cultures (p < 0.05 relative to dominant-negative AMPK plus SNP, and not significant relative to control). Transduction with dominant-negative AMPK had no effect on HUVEC migration to VEGF under normoxic conditions. In contrast, transduction with dominant-negative Akt suppressed migration under normoxic conditions, consistent with previous findings (49), and also exhibited this activity in hypoxic cultures.



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  		FIG. 4.
AMPK-mediated regulation of endothelial cell migration to VEGF under hypoxic and normoxic conditions. Cell migration was evaluated with a Boyden chamber assay. Parallel HUVEC cultures were evaluated for migration toward VEGF under normoxic and hypoxic conditions. Cultures were transduced with dominant-negative Akt (dominant-negative Akt), constitutively-active Akt (caAkt), dominant-negative AMPK (dnAMPK), or GFP (control) 2 days prior to the experiment. Sodium nitroprusside (SNP, 10 µM) was added to some cultures. Quantitative analysis of migration under the different experimental conditions was performed by counting cells on membranes that were stained with Giemsa solution. Each experiment was performed in duplicate, and four independent experiments were performed. Cells were counted in four randomly selected microscopic fields from each membrane. Results are shown as the mean ± S.E. Results are expressed relative to the normoxic control condition. Solid, control; open, dominant-negative Akt; hatch, dominant-negative AMPK; cross-hatch, dominant-negative AMPK plus 10 µM SNP; dots, dominant-negative AMPK plus caAkt. *, p < 0.0001 versus control; #, p < 0.01 versus dominant-negative AMPK alone.

 

AMPK Regulates Endothelial Cell Tube Formation under Hypoxic Conditions—To examine the role of AMPK signaling on endothelial cell differentiation into vascular structures in vitro, HUVECs were plated on matrigel in the presence of VEGF after transduction with GFP, dominant-negative AMPK or dominant-negative Akt, and tube formation was assessed under normoxic and hypoxic conditions (Fig. 5A). Tube structure length was quantified after 18 h using NIH imaging software (Fig. 5B). Transduction with dominant-negative AMPK suppressed tube formation to a similar extent as dominant-negative Akt under hypoxic conditions. The inhibition of tube formation by dominant-negative AMPK was partially reversed by the inclusion of 10 µM SNP in the culture media. Similar effects were observed at 1 and 100 µM SNP (data not shown). Co-transfection with an adenoviral vector expressing constitutively-active Akt was more effective than the NO donor in reversing the inhibition of tube formation by dominant-negative AMPK (p < 0.05 relative to dominant-negative AMPK plus SNP, and not significant relative to control). In contrast, transduction with dominant-negative Akt, but not dominant-negative AMPK, suppressed on tube formation in normoxic cultures (Fig. 5).



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  		FIG. 5.
AMPK-mediated regulation of endothelial cell differentiation into capillary structures in vitro under normoxic and hypoxic conditions. Parallel HUVEC cultures were evaluated for differentiation under normoxic and hypoxic conditions. Cultures were transduced with dominant-negative Akt (dnAkt), constitutively active Akt (caAkt), dominant-negative AMPK (dnAMPK) or GFP (control) 2 days prior to plating on Matrigel-coated culture dishes. Sodium nitroprusside (SNP, 10 µM) was added to some cultures. A, representative cultures for some experimental conditions are shown. B, quantitative analysis of tube formation under the different experimental conditions using NIH Image software. Four independent experiments were performed. Results are show as the mean ± S.E. Results are expressed relative to the normoxic control condition. Solid, control; open, dominant-negative Akt; hatch, dominant-negative AMPK; cross-hatch, dominant-negative AMPK plus 10 µM SNP; dots, dominant-negative AMPK plus caAkt. *, p < 0.0001 versus control; #, p < 0.01 versus dominant-negative AMPK alone.

 

AMPK Regulates Capillary Formation in Matrigel Plugs in Vivo—To assess the role of AMPK signaling on angiogenesis in an in vivo model, matrigel plugs containing adenoviral vectors were subcutaneously implanted on the abdomen of mice. In this assay the matrigel plug serves as a reservoir for the adenoviral vector, and endothelial cells that infiltrate the plug become transduced and express transgene (39). Two weeks after implantation, plugs were harvested and subjected to histochemical analysis using Masson's trichrome staining to identify capillary structures and immunohistochemical analysis using the endothelial cell marker CD31. Matrigel plugs formulated with Ad-dnAMPK showed fewer capillary structures than control (Fig. 6A). Plugs were predominantly comprised of CD31-positive endothelial cells, and plugs containing Ad-dnAMPK revealed fewer CD31-positive cells. Quantitative analyses of histological sections revealed a significantly lower density of CD31-positive cells in the plugs cast with Ad-dnAMPK (Fig. 6B). Similarly, matrigel plugs cast with Ad-dnAkt showed fewer capillary structures and a lower density of CD31-positive cells.



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  		FIG. 6.
AMPK-mediated regulation of angiogenesis in vivo. An in vivo Matrigel plug assay was performed to evaluate the role of AMPK and Akt signaling in angiogenesis. Matrigel plugs containing bFGF and adenoviral vectors expression dominant-negative AMPK (dnAMPK), dominant-negative Akt, or GFP (control) were injected subcutaneously into mice. A, after sacrifice, plugs were excised and stained with Masson's trichrome, to reveal cellularity, and the endothelial cell marker CD31. B, the frequency of CD31-positive cells in five low power fields (lpf) was determined for each Matrigel plug. This experiment was repeated four times. Results are expressed as the mean ± S.E. *, p < 0.01.

 


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
REFERENCES
 
AMPK is widely recognized as a regulator of cellular metabolism in response to hypoxia or vigorous contraction in muscle (8–13). Here, a new regulatory function for AMPK in the vascular endothelium is described. AMPK signaling is induced by oxygen deprivation in endothelial cells, and this induction is essential for angiogenic cellular responses under these conditions. Inhibition of AMPK signaling through overexpression of a dominant-negative mutant of the {alpha}2-subunit inhibited endothelial cell migration toward VEGF and impaired in vitro endothelial cell differentiation into tube-like structures in hypoxic cultures. Because migration and differentiation of endothelial cells are critical features of vessel formation, a mouse Matrigel angiogenesis assay was employed to examine the role of endothelial AMPK signaling in capillary formation in vivo. In this assay subcutaneous Matrigel serves as a reservoir for viral vectors that transduce endothelial cells as they infiltrate the plug, permitting the assessment of intracellular protein function on capillary formation (39). Dominant-negative AMPK significantly inhibited the vascularization of Matrigel plugs, providing evidence that this signaling step is essential for angiogenesis in vivo.

A key finding of this study is that AMPK signaling has little or no effect on endothelial cell migration, tube formation or NO production when cultures are exposed to normoxia. These findings indicate the specificity of the dominant-negative AMPK vector since this reagent only affects angiogenic cellular responses under cell culture conditions associated with robust AMPK induction. Furthermore, these data show that the basal level of AMPK signaling, detected in normoxic cells, contributes minimally to the angiogenic phenotype. Presumably, these relatively low levels of AMPK function in normoxic cells to regulate fatty acid oxidation and glucose metabolism in the normal endothelium (50). Consistent with this hypothesis, transduction of normoxic endothelial cells with dominant-negative AMPK significantly reduced the phosphorylation of ACC, a regulator of fatty acid metabolism (16, 21).

A vector expressing dominant-negative Akt was used as a positive control in this study because Akt signaling is recognized as an essential regulator of NO production (27, 28, 51), survival (32, 52), migration (49, 53, 54), and differentiation (37, 55) in normoxic endothelial cells. However, the role of Akt signaling in hypoxic endothelial cells has not been examined previously. Thus, a secondary finding of this study is that Akt signaling is essential for angiogenic cellular responses in hypoxic endothelial cells. Hypoxia produced a modest time-dependent reduction in Akt phosphorylation at Ser-473. Despite this reduction in Akt activation, signaling through this signaling step was essential for VEGF-mediated endothelial cell differentiation, migration and NO output under conditions of low oxygen tension. Transduction with dominant-negative Akt also inhibited capillary formation in Matrigel plugs implanted in mice. While a number of studies have shown that constitutive activation of Akt signaling is sufficient to promote angiogenesis in vivo (37, 56, 57), the data presented here provide the first direct evidence showing that Akt signaling is essential for neovascularization in an animal model.

Several lines of evidence suggest that cross-talk between AMPK and Akt signaling is an integral component of the proangiogenic cellular response to hypoxic stress (Fig. 7). It was found that inhibition of AMPK signaling suppresses the activating phosphorylation of Akt at Ser-473 in hypoxic endothelial cells. The observation that the inhibition of endothelial cell migration and differentiation by dominant-negative AMPK can be reversed by co-transduction with constitutively active Akt is also consistent with an AMPK-Akt cross-talk mechanism. AMPK-Akt cross-talk may also participate in the regulation of eNOS activation by phosphorylation, because transduction with either dominant-negative Akt or dominant-negative AMPK effectively suppressed eNOS Ser-1177 phosphorylation and NO output from hypoxic endothelial cells. Collectively, these data suggest that eNOS regulation is complex in hypoxic endothelial cells and depends upon the relative phosphorylation status of eNOS at Ser-1177 and Thr-495, the relative activities of Akt and AMPK, and the length of time that cells are exposed to low oxygen tension which causes relatively slow changes in the activities of both of these signaling steps. Under the conditions of our assays, the direct phosphorylation of eNOS by AMPK appeared to contribute minimally to eNOS phosphorylation or endothelial NO output (Fig. 7). However, direct AMPK-mediated eNOS phosphorylation at Ser-1177 may become significant when cultures are incubated under conditions of prolonged hypoxia or when peroxynitrite is produced during ischemia followed by reperfusion (58). Therefore, further analyses of AMPK and Akt signaling, and potential cross-talk between the molecules, may shed additional light on the process that controls angiogenesis in ischemic tissues.



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  		FIG. 7.
Proposed scheme for endothelial AMPK signaling in the control of angiogenesis. Hypoxia activates AMPK which, in turn, promotes mitogen-stimulated Akt activation and angiogenesis. Akt can be the predominant kinase involved in eNOS phosphorylation at Ser-1177 in hypoxic endothelial cells. However, direct eNOS phosphorylation by AMPK may become appreciable under conditions of prolonged hypoxia or peroxynitrite production (58).

 

The maintenance of NO output under hypoxic conditions may contribute partially to the pro-angiogenic activity of AMPK. Endothelium-derived NO is a positive modulator of angiogenesis (42–46), although the mechanism by which NO exerts this effect is unknown. In the study reported here, dominant-negative AMPK diminished eNOS phosphorylation on Ser-1177 and NO output under conditions of low oxygen tension, whereas AMPK-inhibition had little if any effect on these parameters in normoxic cultures. Furthermore, the inclusion of an NO donor to the media partially reversed the negative effects of dominant-negative AMPK on endothelial cell differentiation and migration to VEGF in hypoxic cultures. However, co-transduction with constitutively active Akt was more effective than the NO donor at reversing the inhibitory effects of dominant-negative AMPK. These data suggest that while NO has a stimulatory effect on angiogenic cellular processes, AMPK signaling can regulate angiogenesis independent of NO production (Fig. 7). Similar conclusions have recently been drawn from analyses of the NO-independent regulation of angiogenesis by glycogen synthase kinase-3{beta} (39).

In summary, we report that the activation of a pro-angiogenic phenotype in endothelial cells exposed to hypoxia is a new stress-related function for AMPK signaling. The finding that dominant-negative AMPK inhibits the vascularization of implanted Matrigel plugs suggests the physiological importance of AMPK in angiogenesis in vivo. Stimuli that may activate endothelial AMPK in vivo include decreases in oxygen tension that are associated with tumor growth or occlusive vessel diseases (29, 59). In contrast, angiogenesis associated with normal tissue growth and organ enlargement during development can be subject to regulatory controls that function independently of tissue oxygen gradients (56, 60). Collectively, these data suggest endothelial AMPK signaling may be an essential feature of the angiogenic response to ischemic stress, while having minimal effects on angiogenic endothelial cell responses during development or normal postnatal tissue growth. Therefore, endothelial AMPK may represent a pharmacological target for the selective inhibition of angiogenesis associated with pathological tissue growth.


   FOOTNOTES
 
* This work was supported by National Institutes of Health Grants AR40197, HL66957, AG15052, and AG17241 (to K. W.). The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact. Back

{ddagger} Supported by a Japan Heart Foundation grant and Ueda Memorial Foundation grant. Back

§ To whom correspondence should be addressed: Molecular Cardiology/Whitaker Cardiovascular Institute, Boston University School of Medicine, 715 Albany St., W611, Boston, MA 02118. Tel.: 617-414-2392; Fax: 617-414-2391; E-mail: kxwalsh{at}bu.edu.

1 The abbreviations used are: AMPK, AMP-activated kinase; AMPKK, AMP-activated kinase kinase; HMG-CoA, 3-hydroxy-3-methylglutaryl-CoA; ACC, acetyl-CoA; eNOS, endothelial nitric-oxide synthase; HUVECs, human umbilical vein endothelial cells; AICAR, 5-aminoimidazole-4-carboxamide-1-{beta}-D-ribofuranoside; GFP, green fluorescence protein; EBM, endothelial cell basal medium; EGM, endothelial cell growth medium; SNP, sodium nitroprusside; L-NAME, N{omega}-nitro-L-arginine methyl ester; m.o.i., multiplicity of infection; VEGF, vascular endothelial growth factor. Back


   ACKNOWLEDGMENTS
 
We thank Ann Bialik for technical help in the histological analysis. We thank Richard Cohen and Neil Ruderman for helpful discussion and advice.



   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 

  1. Breier, G., Damert, A., Plate, K. H., and Risau, W. (1997) Thromb. Haemost. 78, 678–683[Medline] [Order article via Infotrieve]
  2. Ferrara, N., and Alitalo, K. (1999) Nat. Med. 5, 1359–1364[CrossRef][Medline] [Order article via Infotrieve]
  3. Carmeliet, P. (2000) Nat. Med. 6, 389–395[CrossRef][Medline] [Order article via Infotrieve]
  4. Pages, G., Milanini, J., Richard, D. E., Berra, E., Gothie, E., Vinals, F., Pouyssegur, J. (2000) Ann. N. Y. Acad. Sci. 902, 187–200[Abstract/Free Full Text]
  5. Shiojima, I., and Walsh, K. (2002) Circ. Res. 90, 1243–1250[Abstract/Free Full Text]
  6. Carling, D., Aguan, K., Woods, A., Verhoeven, A. J., Beri, R. K., Brennan, C. H., Sidebottom, C., Davison, M. D., and Scott, J. (1994) J. Biol. Chem. 269, 11442–11448[Abstract/Free Full Text]
  7. Mitchelhill, K. I., Stapleton, D., Gao, G., House, C., Michell, B., Katsis, F., Witters, L. A., and Kemp, B. E. (1994) J. Biol. Chem. 269, 2361–2364[Abstract/Free Full Text]
  8. Kudo, N., Barr, A. J., Barr, R. L., Desai, S., and Lopaschuk, G. D. (1995) J. Biol. Chem. 270, 17513–17520[Abstract/Free Full Text]
  9. Kudo, N., Gillespie, J. G., Kung, L., Witters, L. A., Schulz, R., Clanachan, A. S., and Lopaschuk, G. D. (1996) Biochim. Biophys. Acta 1301, 67–75[Medline] [Order article via Infotrieve]
  10. Chen, Z., Heierhorst, J., Mann, R. J., Mitchelhill, K. I., Michell, B. J., Witters, L. A., Lynch, G. S., Kemp, B. E., and Stapleton, D. (1999) FEBS Lett. 460, 343–348[CrossRef][Medline] [Order article via Infotrieve]
  11. Winder, W. W., and Hardie, D. G. (1996) Am. J. Physiol. 270, E299-E304[Medline] [Order article via Infotrieve]
  12. Hutber, C. A., Hardie, D. G., and Winder, W. W. (1997) Am. J. Physiol. 272, E262-E266[Medline] [Order article via Infotrieve]
  13. Hayashi, T., Hirshman, M. F., Kurth, E. J., Winder, W. W., and Goodyear, L. J. (1998) Diabetes 47, 1369–1373[Abstract]
  14. Mu, J., Brozinick, J. T., Jr., Valladares, O., Bucan, M., and Birnbaum, M. J. (2001) Mol. Cell. 7, 1085–1094[CrossRef][Medline] [Order article via Infotrieve]
  15. Tian, R., Musi, N., D'Agostino, J., Hirshman, M. F., and Goodyear, L. J. (2001) Circulation 104, 1664–1669[Abstract/Free Full Text]
  16. Carling, D., Clarke, P. R., Zammit, V. A., and Hardie, D. G. (1989) Eur. J. Biochem. 186, 129–136[Medline] [Order article via Infotrieve]
  17. Moore, F., Weekes, J., and Hardie, D. G. (1991) Eur. J. Biochem. 199, 691–697[Medline] [Order article via Infotrieve]
  18. Weekes, J., Hawley, S. A., Corton, J., Shugar, D., and Hardie, D. G. (1994) Eur. J. Biochem. 219, 751–757[Medline] [Order article via Infotrieve]
  19. Hawley, S. A., Davison, M., Woods, A., Davies, S. P., Beri, R. K., Carling, D., and Hardie, D. G. (1996) J. Biol. Chem. 271, 27879–27887[Abstract/Free Full Text]
  20. Clarke, P. R., and Hardie, D. G. (1990) EMBO J. 9, 2439–2446[Medline] [Order article via Infotrieve]
  21. Davies, S. P., Sim, A. T., and Hardie, D. G. (1990) Eur. J. Biochem. 187, 183–190[Medline] [Order article via Infotrieve]
  22. Park, H., Kaushik, V. K., Constant, S., Prentki, M., Przybytkowski, E., Ruderman, N. B., and Saha, A. K. (2002) J. Biol. Chem. 277, 32571–32577[Abstract/Free Full Text]
  23. Muoio, D. M., Seefeld, K., Witters, L. A., and Coleman, R. A. (1999) Biochem. J. 338, 783–791[CrossRef][Medline] [Order article via Infotrieve]
  24. Carling, D., and Hardie, D. G. (1989) Biochim. Biophys. Acta 1012, 81–86[Medline] [Order article via Infotrieve]
  25. Velasco, G., Geelen, M. J., and Guzman, M. (1997) Arch. Biochem. Biophys. 337, 169–175[CrossRef][Medline] [Order article via Infotrieve]
  26. Ruderman, N. B., Saha, A. K., Vavvas, D., and Witters, L. A. (1999) Am. J. Physiol. 276, E1–E18[Medline] [Order article via Infotrieve]
  27. Fulton, D., Gratton, J. P., McCabe, T. J., Fontana, J., Fujio, Y., Walsh, K., Franke, T. F., Papapetropoulos, A., and Sessa, W. C. (1999) Nature 399, 597–601[CrossRef][Medline] [Order article via Infotrieve]
  28. Dimmeler, S., Fleming, I., Fisslthaler, B., Hermann, C., Busse, R., and Zeiher, A. Z. (1999) Nature 399, 601–605[CrossRef][Medline] [Order article via Infotrieve]
  29. Chen, Z. P., Mitchelhill, K. I., Michell, B. J., Stapleton, D., Rodriguez-Crespo, I., Witters, L. A., Power, D. A., Ortiz de Montellano, P. R., and Kemp, B. E. (1999) FEBS Lett. 443, 285–289[CrossRef][Medline] [Order article via Infotrieve]
  30. Dagher, Z., Ruderman, N., Tornheim, K., and Ido, Y. (1999) Biochem. Biophys. Res. Commun. 265, 112–115[CrossRef][Medline] [Order article via Infotrieve]
  31. Ido, Y., Carling, D., and Ruderman, N. (2002) Diabetes 51, 159–167[Abstract/Free Full Text]
  32. Fujio, Y., and Walsh, K. (1999) J. Biol. Chem. 274, 16349–16354[Abstract/Free Full Text]
  33. Fujio, Y., Guo, K., Mano, T., Mitsuuchi, Y., Testa, J. R., and Walsh, K. (1999) Mol. Cell. Biol. 19, 5073–5082[Abstract/Free Full Text]
  34. Nagata, D., Hirata, Y., Suzuki, E., Kakoki, M., Hayakawa, H., Goto, A., Ishimitsu, T., Minamino, N., Ono, Y., Kangawa, K., Matsuo, H., and Omata, M. (1999) Kidney Int. 55, 1259–1267[CrossRef][Medline] [Order article via Infotrieve]
  35. Nagata, D., Suzuki, E., Nishimatsu, H., Yoshizumi, M., Mano, T., Walsh, K., Sata, M., Kakoki, M., Goto, A., Omata, M., and Hirata, Y. (2000) Circ. Res. 87, 699–704[Abstract/Free Full Text]
  36. Witzenbichler, B., Kureishi, Y., Luo, Z., Le Roux, A., Branellec, D., and Walsh, K. (1999) J. Clin. Invest. 104, 1469–1480[Medline] [Order article via Infotrieve]
  37. Kureishi, Y., Luo, Z., Shiojima, I., Bialik, A., Fulton, D., Lefer, D. J., Sessa, W. C., and Walsh, K. (2000) Nat. Med. 6, 1004–1010[CrossRef][Medline] [Order article via Infotrieve]
  38. Muhlhauser, J., Merrill, M. J., Pili, R., Maeda, H., Bacic, M., Bewig, B., Passaniti, A., Edwards, N. A., Crystal, R. G., and Capogrossi, M. D. (1995) Circ. Res. 77, 1077–1086[Abstract/Free Full Text]
  39. Kim, H.-S., Skurk, C., Thomas, S. R., Bialik, A., Suhara, T., Kureishi, Y., Birnbaum, M., Keaney Jr., J. F., and Walsh, K. (2002) J. Biol. Chem. 277, 41888–41896[Abstract/Free Full Text]
  40. Alessi, D. R., Andjelkovic, M., Caudwell, B., Cron, P., Morrice, N., Cohen, P., and Hemmings, B. A. (1996) EMBO J. 15, 6541–6551[Medline] [Order article via Infotrieve]
  41. Toker, A., and Newton, A. C. (2000) J. Biol. Chem. 275, 8271–8274[Abstract/Free Full Text]
  42. Noiri, E., Lee, E., Testa, J., Quigley, J., Colflesh, D., Keese, C. R., Giaever, I., and Goligorsky, M. S. (1998) Am. J. Physiol. 274, C236-C244[Medline] [Order article via Infotrieve]
  43. Murohara, T., Asahara, T., Silver, M., Bauters, C., Masuda, H., Kalka, C., Kearney, M., Chen, D., Symes, J. F., Fishman, M. C., Huang, P. L., and Isner, J. M. (1998) J. Clin. Invest. 101, 2567–2578[Medline] [Order article via Infotrieve]
  44. Shizukuda, Y., Tang, S., Yokota, R., and Ware, J. A. (1999) Circ. Res. 85, 247–256[Abstract/Free Full Text]
  45. Fukumura, D., Gohongi, T., Kadambi, A., Izumi, Y., Ang, J., Yun, C. O., Buerk, D. G., Huang, P. L., and Jain, R. K. (2001) Proc. Natl. Acad. Sci. U. S. A. 98, 2604–2609[Abstract/Free Full Text]
  46. Rikitake, Y., Hirata, K., Kawashima, S., Ozaki, M., Takahashi, T., Ogawa, W., Inoue, N., and Yokoyama, M. (2002) Arterioscler. Thromb. Vasc. Biol. 22, 108–114[Abstract/Free Full Text]
  47. Michell, B. J., Chen, Z., Tiganis, T., Stapleton, D., Katsis, F., Power, D. A., Sim, A. T., and Kemp, B. E. (2001) J. Biol. Chem. 276, 17625–17628[Abstract/Free Full Text]
  48. Fleming, I., Fisslthaler, B., Dimmeler, S., Kemp, B. E., and Busse, R. (2001) Circ. Res. 88, E68–75[CrossRef][Medline] [Order article via Infotrieve]
  49. Morales-Ruiz, M., Fulton, G., Sowa, G., Languino, L. R., Fujio, Y., Walsh, K., and Sessa, W. C. (2000) Circ. Res. 86, 892–896[Abstract/Free Full Text]
  50. Dagher, Z., Ruderman, N., Tornheim, K., and Ido, Y. (2001) Circ. Res. 88, 1276–1282[Abstract/Free Full Text]
  51. Luo, Z., Fujio, Y., Kureishi, Y., Rudic, R. D., Daumerie, G., Fulton, D., Sessa, W. C., and Walsh, K. (2000) J. Clin. Invest. 106, 493–499[Medline] [Order article via Infotrieve]
  52. Gerber, H.-P., McMurtrey, A., Kowalski, J., Yan, M., Key, B. A., Dixit, V., and Ferrara, N. (1998) J. Biol. Chem. 273, 30336–30343[Abstract/Free Full Text]
  53. Dimmeler, S., Dernbach, E., and Zeiher, A. M. (2000) FEBS Lett. 477, 258–262[CrossRef][Medline] [Order article via Infotrieve]
  54. Radisavljevic, Z., Avraham, H., and Avraham, S. (2000) J. Biol. Chem. 275, 20770–20774[Abstract/Free Full Text]
  55. Schonherr, E., Levkau, B., Schaefer, L., Kresse, H., and Walsh, K. (2001) J. Biol. Chem. 276, 40687–40692[Abstract/Free Full Text]
  56. Takahashi, A., Kureishi, Y., Yang, J., Luo, Z., Guo, K., Mukhopadhyay, D., Ivashchenko, Y., Branellec, D., and Walsh, K. (2002) Mol. Cell. Biol. 22, 4803–4814[Abstract/Free Full Text]
  57. Jiang, B. H., Zheng, J. Z., Aoki, M., and Vogt, P. K. (2000) Proc. Natl. Acad. Sci. U. S. A. 97, 1749–1753[Abstract/Free Full Text]
  58. Zou, M. H., Hou, X. Y., Shi, C. M., Nagata, D., Walsh, K., and Cohen, R. A. (2002) J. Biol. Chem. 277, 32552–32557[Abstract/Free Full Text]
  59. Semenza, G. L. (2000) Genes Dev. 14, 1983–1991[Free Full Text]
  60. Oosthuyse, B., Moons, L., Storkebaum, E., Beck, H., Nuyens, D., Brusselmans, K., Van Dorpe, J., Hellings, P., Gorselink, M., Heymans, S., Theilmeier, G., Dewerchin, M., Laudenbach, V., Vermylen, P., Raat, H., Acker, T., Vleminckx, V., Van den Bosch, L., Cashman, N., Fujisawa, H., Drost, M. R., Sciot, R., Bruyninckx, F., Hicklin, D., Ince, C., Gressens, P., Lupu, F., Plate, K. H., Robberecht, W., Herbert, J. M., Collen, D., and Carmeliet, P. (2001) Nat. Genet. 28, 131–138[CrossRef][Medline] [Order article via Infotrieve]

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