In Mice With Type 2 Diabetes, a Vascular Endothelial Growth Factor (VEGF)-Activating Transcription Factor Modulates VEGF Signaling and Induces Therapeutic Angiogenesis After Hindlimb Ischemia

The VEGF-activating ZFP was delivered via a previously described genetically engineered plasmid that encoded a three–zinc finger DNA-binding domain, the nuclear localization signal from simian virus 40 large T antigen, and the transactivation domain from the p65 subunit of the human nuclear factor-κB, subcloned into pVAX1 (Invitrogen, Carlsbad, CA) with expression under direction of the cytomegalovirus promoter (7,16). Controls in prior studies included the identical plasmid without the ZFP VEGF insert, β-galactosidase, and the presence or absence of poloxomer (7,17). Therefore, for this current study, vials containing poloxomer with and without plasmid were identical in appearance and were coded to maintain blinding until data analysis was complete.

In total, 60 male C57BL/6 mice (Jackson Laboratory, Bar Harbor, ME) at 4 weeks old were fed a high-fat diet to induce type 2 diabetes (21). In the high-fat diet (D12451; Research Diet, New Brunswick, NJ), 45% of total calories were from fat, which has been shown to induce obesity with increased insulin levels and insulin resistance in the absence of genetic mutations (22,23). After 12 weeks on the high-fat diet, glucose tolerance testing was performed in which mice were fasted overnight, provided water ad libitum, and given a glucose load of 1 mg/kg body wt by intraperitoneal injection. Blood glucose concentrations were measured before and 30, 60, and 90 min after, using a OneTouch Ultra glucometer (LifeScan, Milpitas, CA). Glucose tolerance was determined as the area under the curve of blood glucose concentrations. All high-fat diet–fed mice met the criteria for diabetes (23).

Mice were anesthetized by intraperitoneal injection of ketamine (90 mg/kg) and xylazine (10 mg/kg) and underwent surgically induced unilateral hindlimb ischemia with ligation and excision of the femoral artery from its origin just above the inguinal ligament to its bifurcation at the origin of the saphenous and popliteal arteries, using methods described previously (24). The inferior epigastric, lateral circumflex, and superficial epigastric artery branches were also isolated and ligated. Mice were closely monitored during the postoperative period and underwent laser Doppler perfusion imaging immediately after surgery and at 10-day intervals until the termination of the study. The extent of necrosis, if any, in ischemic hindlimbs was recorded one time between days 3 and 5 after surgery: grade 0, no necrosis in ischemic limb; grade I, necrosis limited to toes; grade II, necrosis extending to dorsum pedis; grade III, necrosis extending to crus; and grade IV, necrosis extending to thigh.

Ten days after the operation, the mice were anesthetized with 2% isoflurane (VETEQUIP inhalation anesthesia system; VETEQUIP, Pleasanton, CA), and hindlimb perfusion was recorded as the 0 time point. Mice were randomized in a blinded manner into either a ZFP-VEGF treatment group (125 μg ZFP-VEGF plasmid in 125 μl 1% poloxamer/50 mmol/l NaCl) or control, no ZFP-VEGF group (125 μl 1% Poloxamer/50 mmol/l NaCl). The dose and method of intramuscular injections were previously described (7). Group assignments were made to accomplish different study objectives. Based upon their group assignment, the study was completed 3, 10, or 20 days after injection.

Ten age-matched wild-type C57BL/6 mice fed normal chow were used as normal controls for body weights, glucose level, VEGF, phospho-Akt, Akt protein level, and capillary density in skeletal muscle. Another 10 mice fed normal chow underwent hindlimb ischemic surgery to get the comparable perfusion recovery data.

A single operator (Y.L.) blinded to group assignments performed all of the surgeries and treatments. All protocols and procedures involving animals conformed with the “Guidelines for Use of Laboratory Animals” published by the U.S. Department of Health and Human Services and were approved by the Duke University Animal Care and Use Committee. All animals received care in accordance with “Principles of Laboratory Animal Care,” formulated by the National Society for Medical Research, and “Guide for the Care and Use of Laboratory Animals,” published by the National Institutes of Health (NIH Publication No. 86-23, revised 1985).

Mice were anesthetized, underwent laser Doppler imaging, and were killed, as previously described (7). The distal portion of the ischemic and contralateral, nonischemic muscles was placed in 30% sucrose-PBS solution and mounted in cross section in OCT. Cryostat sections (5 μm) were prepared on microscope slides, and the remaining tissues were snap frozen in liquid N₂ for RNA and/or protein extraction.

Total RNA was isolated and purified, and concentrations were determined by spectrophotometer, using methods previously described (7,17). For RT-PCR, primers, their sequences, and the amplifications conditions are shown in Table 1. Samples were subject to electrophoresis through an agarose gel, and water without primers served as the negative controls. For real-time PCR, 2 μg total RNA was used for first-strand cDNA synthesis by reverse transcription using SuperScript III First-Stand Synthesis System (Invitrogen), and 2 μl cDNA was quantitatively amplified using TaqMan Universal PCR Master Mix (AB Applied Biosystems, Foster City, CA). A single-tube PCR was optimized for the quantification of specific mouse primer and probe (Table 1) for VEGF. All real-time PCR data were measured and captured with a sequence detector, and mRNA concentrations were calculated using the Ct value representing the PCR cycle number at which fluorescence was detectable above an arbitrary threshold normalized against 18S rRNA, as previously described (17). Negative controls lacking cDNA were always included, and each sample was tested in triplicate.

Muscle samples were weighed, homogenized, centrifuged in Tris-saline buffer; protein concentrations were determined by Bradford assay; and total VEGF protein concentration was determined by a solid-state enzyme-linked immunoassay system, as previously described (17). Phospho-Akt and total Akt were measured using a Biosource AKT [pS473] and AKT ELISA kit, respectively (Invitrogen). Caspase activity was determined using Caspase-3 colorimetric assay kit (Biovision, Mountview, CA), and absorbance was normalized to the protein concentration in each sample. Data were expressed as units per milligrams of protein.

Protein samples were separated on SDS-polyacrylamide gels; transferred to a polyvinylidene difluoride membranes, which were treated with blocking buffer (5% skim milk); probed with a Ser473–phospho-Akt (1:1,000), Akt (1:1,000), anti–phospho–endothelial nitric oxide (NO) synthase (eNOS) Ser1177 (1:1,000), cyclooxygenase 2 (Cox-2) (1:500; Cell Signaling, Beverly, MA), or eNOS (1:1,000; Cell Signaling) antibodies overnight at 4°C; and then incubated with a horseradish peroxidase–conjugated anti-rabbit immunoglobulin (1:5,000; Cell Signaling) for 1 h at room temperature. Immune reactivity was visualized using ECL plus system. Quantification of results was performed by densitometry (ImageJ 1.36b; National Institutes of Health, Bethesda, MD) comparing the density of identically sized areas (corresponding to immunoreactive bands), and results were analyzed as total integrated density (arbitrary units).

Endothelial cells were identified by immunohistochemical staining using a rat anti-mouse CD31 antibody (1:200 dilution; Serotec, Raleigh, NC). Capillary density was measured by counting six random high-power (magnification ×200) fields or a minimum of 200 fibers from each ischemic and nonischemic limb on an inverted light microscope and was expressed by the number of CD31⁺ cells per square millimeter or per fiber. Area was measured with an NIH Image analysis system.

Cell proliferation and apoptosis in skeletal muscle was detected with immunohistochemical staining of proliferating cell nuclear antigen (PCNA, 1:50 dilution; Santa Cruz Biotechnology, Santa Cruz, CA) and transferase-mediated dUTP nick-end labeling (TUNEL) staining (ApopTag In Situ kit; Chemicon, Temecula, CA), respectively. The proliferation index and apoptotic index were expressed by the percentage of the number of positive nuclei divided by the total number of counted nuclei. The count was performed on three randomly selected fields (magnification ×200) and at least 200 nuclei per sample. A single reader (K.W. [see acknowledgments]) blinded to the sample type performed all the analysis.

Bilateral hindlimb perfusion was measured using a Laser Doppler Perfusion Imager system (PERIMED, Stockholm, Sweden). Briefly, mice received anesthesia with 2% isoflurane, and hair was removed from both legs using a depilatory cream, whereas mice were maintained on a heating plate to minimize temperature variations. The laser Doppler perfusion imager used a beam from a 2-mW helium-neon laser that sequentially scanned a 12- × 12-cm tissue surface to a depth of a few hundred microns. During the scanning procedure, the moving blood cells shifted the frequency of incident light according to the Doppler principle. The back-scattered light was collected by a photodiode collector, transformed into voltage variations, and expressed as a color-coded image representing the microvascular blood flow distribution. The results at any time point were expressed as a ratio of perfusion in the left (ischemic) versus right (normal) limb. Changes in perfusion were calculated as the ratio at the final time minus the baseline ratio, and this baseline (or day 0 value) was the immediate after- ligation value for the comparison of the mice with diabetes versus without diabetes and was the preinjection value for the effect of ZFP-VEGF treatment versus control over time.

Results are expressed as means ± SE. For comparison of mean data on ZFP-VEGF treatment versus control, at one time point (i.e., VEGF expression, phospho-Akt–to–Akt ratio, vascular density, proliferation, and apoptosis index), statistical significance was evaluated by use of two-tailed Student’s paired t test. To determine the effect of ZFP-VEGF treatment versus control on the change in perfusion ratio over time, a standard variance ANOVA repeated measure was performed. P values <0.05 were considered to be significant.

After 12 weeks of high-fat diet, C57BL/6 mice (n = 60) developed type 2 diabetes. Compared with age-matched normal chow–fed mice (n = 10), those with diabetes had significantly greater body weight (34.63 ± 4.67 vs. 26.18 ± 1.27 g; P < 0.05), fasting glucose levels (124.6 ± 29.3 vs. 92.6 ± 11.3 mg/dl; P < 0.001), and impaired glucose tolerance (22,585 ± 561 vs. 14,811 ± 541 mg · dl⁻¹ · min⁻¹; P < 0.001) (22).

Age- and sex-matched mice with and without diabetes (n = 10 per group) underwent surgically induced hindlimb ischemia. Although both groups demonstrated increases in the perfusion ratio over time (Fig. 1A), the recovery was significantly blunted in the mice with diabetes. The magnitude of the change in the perfusion ratio was assessed by subtracting the day-10, -20, or -30 perfusion ratio from the ratio immediately after ligation (day 0), and this change in perfusion was different between mice with type 2 diabetes and control at the day-10 (0.24 ± 0.07 vs. 0.35 ± 0.02; P < 0.05), day-20 (0.34 ± 0.07 vs. 0.53 ± 0.08; P < 0.05), and day-30 (0.33 ± 0.04 vs. 0.58 ± 0.02; P < 0.001) time points. A sizeable fraction (35%) of mice with type 2 diabetes showed evidence of grade I (28.33%), II (5.0%), or III (1.67%) limb necrosis, whereas necrosis was not evident in any control mice.

In the absence of ischemia, as shown in Fig. 1B, capillary density in the gastrocnemius muscle was similar between mice with type 2 diabetes versus mice without diabetes (433 ± 30/area vs. 435 ± 35/area, or 1.31 ± 0.04/fiber vs. 1.46 ± 0.08/fiber; NS). Although VEGF protein expression (Fig. 1B, right) was significantly greater (26.27 ± 1.16 vs. 22.69 ± 0.93 pg/mg; P < 0.05), the ratio of phospho-Akt to total Akt (Fig. 1C) was lower (0.38 ± 0.08 vs. 0.76 ± 0.13; P < 0.05).

Mice with type 2 diabetes were randomly assigned to receive either ZFP-VEGF plasmid or control (n = 10/group) injections into the ischemic hindlimb muscle 10 days after ischemia. Expression of VEGF mRNA, protein, and signaling (n = 6) was measured 3 days later. VEGF mRNA and protein levels were expressed as the copies of mRNA or concentration of protein in the ischemic gastronomes muscle minus the nonischemic muscle, for each mouse. As shown in Fig. 2A, VEGF mRNA copy number was significantly greater in the ZFP-VEGF treatment versus control (7.23 ± 2.60 vs. −0.89 ± 0.24; P < 0.05). Similarly, VEGF protein expression was significantly greater in the ZFP-VEGF treatment versus control (9.50 ± 3.53 vs. −3.95 ± 1.26 pg/mg; P < 0.05). By Western blotting (Fig. 2B, left), phospho-Akt was higher in ZFP-VEGF treatment compared with control. Quantitative ELISA assay showed that the phospho-Akt–to–Akt ratio (Fig. 2B, right) was significantly greater with ZFP-VEGF versus control treatment (0.74 ± 0.03 vs. 0.51 ± 0.15; P < 0.05). Finally, Fig. 2C, left, showed that more phosphorylated eNOS was present in ZFP-VEGF treatment compared with the control. Quantitative analysis with Image J showed that the phospho-eNOS–to–eNOS ratio was significant increase with ZFP-VEGF versus control treatment, 0.83 ± 0.06 vs. 0.48 ± 0.11; P < 0.05 (integrated density, Fig. 2C, right). By PCR, VEGF-A was greater in ZFP-VEGF versus control treatment, but there were no differences in VEGF-B, VEGF-C, VEGF-D, or VEGF receptor (VEGFR) 1 or 2 (Fig. 3A and B). To exclude differences in the inflammatory response, inducible NO synthase (iNOS) mRNA was similar with ZFP-VEGF or control treatment (Fig. 3C).

Next, 20 mice were randomly assigned to receive either ZFP-VEGF plasmid or control (n = 10/group) injection into the gastrocnemius muscle 10 days after hindlimb ischemia are harvested 10 days after gene transfer (20 days after hindlimb ischemia). Expression of VEGF protein (n = 8) and signaling (n = 7) were assessed as above. As shown in Fig. 4A, VEGF protein expression was significantly greater in the ZFP-VEGF treatment versus control treatment (6.57 ± 1.58 vs. −1.63 ± 2.3 pg/mg; P < 0.05). By ELISA, the phospho-Akt–to–Akt ratio (Fig. 4B) was significantly greater with ZFP-VEGF treatment, 0.56 ± 0.08 vs. 0.39 ± 0.07; P < 0.05. Although changes in the perfusion ratio (0.10 ± 0.03 vs. 0.069 ± 0.03; NS) were similar in the ZFP-VEGF treatment versus control (n = 10 for each), as shown in Fig. 5A, capsize activity was lower in the ZFP-VEGF treatment versus control treatment 0.06 ± 0.01 vs. 0.16 ± 0.04; P < 0.05 (n = 6/group). Although VEGFR1 and -2 mRNA levels were similar at 3 days after gene transfer in the ischemic gastrocnemius muscle, as shown in Fig. 5B, VEGF R1 was lower in the ZFP-VEGF versus control treatment with no difference in VEGF R2. Finally, as shown in Fig. 5C, protein levels of the inflammatory marker Cox-2 was similar between ZFP-VEGF treatment versus control.

Twenty mice were randomly assigned to receive either ZFP-VEGF plasmid or control (n = 10/group) injection into the ischemic gastrocnemius muscle 10 days after hindlimb ischemia. Laser Doppler perfusion imaging was performed before and at 10-day intervals after gene transfer. Before gene transfer, the perfusion ratio was similar between the ZFP-VEGF–treated and control-treated groups (0.62 ± 0.03 vs. 0.65 ± 0.03). By ANOVA, there was perfusion recovery over time in both the ZFP-VEGF and the control treatment groups. As we observed before (prior paragraph), there was a trend toward an increase in perfusion ratio in ZFP-VEGF versus control treatment (0.162 ± 0.022 vs. 0.103 ± 0.023; P = 0.08) at 10 days after injection (Fig. 6A). At 20 days after gene transfer, there was a clear and statistically significant increase in the change in the perfusion ratio with ZFP-VEGF treatment (0.241 ± 0.021 vs. 0.086 ± 0.016; P < 0.001, Fig. 6A).

Capillary density (Fig. 6B) in the ischemic gastrocnemius muscle was significantly higher in the ZFP-VEGF versus control-treated mice (528 ± 29 vs. 444 ± 17; P < 0.01). To exclude any potential differences induced by muscle edema or atrophy, the capillary–to–muscle fiber ratio in the ischemic muscle was also measured. Like capillary density, the capillary–to–muscle fiber ratio was significantly higher in the ZFP-VEGF versus control-treated mice (1.51 ± 0.06 vs. 1.28 ± 0.05; P < 0.01). The fraction of PCNA-positive cells (Fig. 6B) in the ischemic gastrocnemius muscle was significantly greater in ZFP-VEGF versus control-treated mice (2.56 ± 0.37 vs. 0.76 ± 0.28%; P < 0.01). Apoptosis as measured by TUNEL-positive nuclei in the ischemic muscle (Fig. 6B) was significantly decreased in ischemic muscle of ZFP-VEGF versus control-treated mice (1.50 ± 0.21 vs. 2.25 ± 0.25%; P < 0.05).

Type 2 diabetes is a major risk factor for the development of PAOD (2,3). When diabetes is superimposed on PAD, there is a poorer clinical prognosis, and surgical and/or catheter-based revascularization frequently cannot be used in a sizable fraction of these patients. Therefore, therapeutic angiogenesis is being actively explored as a future treatment for patients with PAD, many of whom will have superimposed type 2 diabetes (5,7,8). To the best of our knowledge, this report is the first to investigate the therapeutic efficiency of VEGF in a preclinical PAD model with diet-induced type 2 diabetes. The major findings of our study were that diet-induced diabetes was associated with: 1) an impaired perfusion recovery after hindlimb ischemia; 2) an increase in VEGF ligand but reduced VEGF signaling in skeletal muscle in the absence of ischemia; and 3) gene transfer with a transcription factor designed to increase VEGF expression into an ischemic limb, increase VEGF signaling, and improve the perfusion recovery.

A VEGF signaling defect may be one of the reasons for the reduced collateral blood vessel growth seen in patients with diabetes (18–20,25). Sasso et al. (19) showed that patients with diabetes had reduced VEGF signaling in the myocardium despite an increase in VEGF ligand (19). We found greater levels of the VEGF ligand with reduced VEGF signaling in hindlimb muscle before ligation, and after ligation this was associated with an impaired perfusion recovery by laser Doppler and a greater extent of tissue necrosis. Despite an increase in ligand levels, we found that this transcription factor was able to further increase VEGF ligand expression and substantially correct the VEGF signaling defect in ischemic muscle in the early days after gene transfer in mice with diabetes. Akt is a serine/threonine protein kinase and is activated by phosphorylation from phosphoinositide-dependent kinases, and the phosphorylation of eNOS is one of the targets of phospho-Akt (26,27). The correction in VEGF signaling was followed by improvements in tissue perfusion, greater amounts of cell proliferation, and reduced apoptosis in the treated limb. Interestingly, in a rat model of streptozotocininduced type 1 diabetes, the delivery of an engineered VEGF transcription factor was protective against diabetic neuropathy, and the authors also demonstrated that conditioned medium from transfected cells protected human neuroblastoma cells from serum starvation, indicating a direct cytoprotective effect (16). Take together these data suggest that modulating VEGF expression remains an option as a therapeutic target even in the setting of type 2 diabetes.

Ischemic injury is a potent stimulus for increasing VEGF and inflammation before initiating angiogenesis (28). However, in current study, we found an attenuated increase in VEGF after ischemic injury in mice with type 2 diabetes at both days 13 and 20 after ligation, as shown by a negative value when VEGF expression was compared with the opposite limb (Figs. 2A and 4A). This finding was consistent with the findings of Schiekofer et al. (29) who used Lepr^db/db mice and microarrays to identify hindlimb cDNA expression profiles before and after ischemia. Unlike the wild-type mice, VEGF-A was not upregulated in Lepr^db/db mice on days 1, 7, or 14 after surgery. The NOD model is characterized by a leukocytic infiltration of the pancreatic islets and immune dysfunctions (30). Rivard et al. (12) in NOD mice reported lower VEGF expression in ischemic muscle compared with wild-type mice from days 3 to 14 after surgery, even when glucose was controlled with insulin. They reported an impairment of perfusion recovery and neo-capillary formation in the NOD versus wild-type mice that was corrected with adeno-VEGF (murine VEGF₁₆₄) delivered immediately after surgery. Although in our study we showed an increase in VEGF expression after gene transfer, we did not find differences in iNOS or Cox-2 in the ZFP-VEGF versus control-treated muscle to implicate alterations in inflammation. Although poloxomer was present in our study, it was present in both groups, and prior studies from our laboratory did not find that the presence or absence of poloxomer was associated with differences in VEGF expression (7,17) or cellular infiltration (7).

VEGF has several isoforms that arise from differential splicing that are differentially regulated in the setting of ischemia. In mice with streptozotocin-induced type 1 diabetes, Roguin et al. (13) found no benefit on perfusion flow restoration from a continually delivery of rat VEGF-165 plasmid (90% homology of mouse VEGF) into ischemic muscle, although they did find an apparent increase in factor VIII–positive cells and smooth muscle actin–positive cells in the ischemic muscle of diabetic mice that received VEGF. Data from placebo-controlled human clinical trials have not been encouraging (14,15). Modulating endogenous VEGF transcription is one method to alter multiple VEGF isoforms, and we found that the ZFP-VEGF–activating transcription factor could upregulate VEGF-A in ischemic muscle from mice with diabetes. Interestingly, at 3 days after gene transfer, there was no difference in VEGF-B, VEGF-C, VEGF-D, and VEGFR1 or -2 in muscle treated with the ZFP-VEGF compared with control. At a later time point, we found more VEGFR2 relative to VEGFR1 mRNA. Studies from knockout mice have shown that VEGFR2 is an angiogenic VEGFR, and VEGFR1 may actually have an inhibitory role. Whereas VEGFR2-deficient mice die because of a failure to develop blood vessels or even endothelial cells at all, VEGFR1-deficient mice appear to have an overgrowth of endothelial cells that fail to properly pattern into normal vessels (31,32). These data suggested that a role of VEGFR1 is to limit endothelial cell proliferation. Further studies have shown that VEGFR1 limits VEGFR2-mediated endothelial cell proliferation through a pathway involving VEGFR1-mediated activation of phosphoinositide 3-kinase and subsequent NO production. (33,34). Whether modulating multiple VEGF isoforms will lead to better efficacy in humans compared with single isoform therapy remains to be determined.

This study was supported by the Iacoccoa Foundation. The ZFP-VEGF plasmid in poloxomer and the poloxomer control vials were provided by Edwards Life Sciences.

We acknowledge Shelley Odronic, Erika Reed, Karen Wu, and Conrad Ireland for their assistance. We thank Shoukang Zhu from Dr. Pascal Goldschmidt’s laboratory for help in the real-time PCR studies.