Id1 Gene Transfer Confers Angiogenic Property on Fully Differentiated Endothelial Cells and Contributes to Therapeutic Angiogenesis
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《循环学杂志》
the Departments of Cardiovascular Medicine (K.N., K.K., M.Y., H.O.) and Cardiovascular Surgery (K.T.), Graduate School of Medical Sciences, Kumamoto University, Kumamoto, Japan
Department of Physiological Chemistry and Metabolism, Division of Biochemistry and Molecular Biology (Y.K., H.K.), and Department of Developmental and Medical Technology (Sankyo) (K.N.), University of Tokyo Graduate School of Medicine, Tokyo, Japan
Department of Viral Infection and Vaccine Control, National Institute for Infectious Diseases (A.K.), Tokyo, Japan.
Abstract
Background— Transplantation of endothelial progenitor cells has been proposed as a potential strategy for therapeutic revascularization. However, the limited endogenous cell pool and the related technical difficulties constitute clinically important disadvantages to autologous transplantation. In this study we investigated whether fully differentiated endothelial cells (ECs) modified with gene transfer of Id1, a helix-loop-helix transcription factor involved in angiogenesis, have the potential to contribute to therapeutic angiogenesis.
Methods and Results— The Id1 gene was transferred into human umbilical vein ECs (HUVECs) via a Sendai virus vector. Id1 stimulated migration, proliferation, and capillary-like tube/cord formation of HUVECs. In addition, Id1 reduced serum deprivation–induced HUVEC apoptosis, as shown by FACS analysis with annexin V and TUNEL staining. Transplantation of Id1-overexpressing HUVECs accelerated recovery of blood flow as evaluated by laser-Doppler perfusion imaging, increased capillary density, and improved the rate of limb salvage compared with the transplantation of control HUVECs. Histochemical analysis revealed that the regenerated vascular networks of limbs transplanted with Id1-overexpressing HUVECs contained numerous HUVECs, some of which were in a proliferative state. Untransfected HUVECs were also incorporated with Id1-transfected HUVECs, suggesting the noncell autonomous effect of Id1. Finally, angiopoietin-1 was upregulated in Id1-overexpressing HUVECs and functionally contributed to the in vitro angiogenic effect of Id1.
Conclusions— Id1 gene transfer conferred HUVECs with an angiogenic property, contributing to neovascularization after transplantation into ischemic lesions. Transplantation of Id1-overexpressing mature ECs may serve as a novel and useful strategy for therapeutic angiogenesis.
Key Words: endothelial cells genes therapeutic angiogenesis
Introduction
The discovery of circulating endothelial progenitor cells (EPCs) has provided a new mechanistic understanding of neovascularization.1,2 In response to cytokines, EPCs are mobilized from the bone marrow, recruited to the site of wound healing, tissue ischemia, and tumor growth, and then differentiated into vascular endothelial cells (ECs).1–5 This process, termed postnatal vasculogenesis, is postulated, together with angiogenesis, to contribute to regional neovascularization.4 Several animal experiments and human trials under way are focused on the potential application of EPCs to vascular regenerative therapy.6
Although the usefulness of EPCs for therapeutic neovascularization is highly anticipated, several issues remain unresolved. Most notably, the limited cell pool and technical difficulties in expanding EPCs in vitro have prevented widespread clinical use of EPCs for autologous transplantation. Differentiated vascular ECs, on the other hand, can be isolated and expanded in vitro.7,8 Moreover, a large number of mature ECs can be differentiated from bone marrow–derived progenitor cells and embryonic stem cells.9,10 Thus, it would be advantageous for mature ECs to serve as cell sources for neovascularization. However, mature ECs are inefficiently incorporated into forming vessels, such that their contribution to therapeutic angiogenesis is limited.11 The fact that ECs in adult organs transform phenotypically into "angiogenic" cells under pathological circumstances and participate in neovascularization5 prompted us to hypothesize that quiescent ECs may become "angiogenic" via phenotypic modification with gene transfer.
The Id proteins are a family of helix-loop-helix (HLH) transcription factors.12 Id proteins lack a basic DNA binding domain and primarily act as inhibitors of basic HLH transcription factors, via heterodimerization.12 Through this mechanism, Id proteins have been implicated in regulating a variety of cellular processes including cell growth, senescence, differentiation, and neoplastic transformation.12–14 More recently, accumulating evidence has suggested that Id1 and Id3 play vital roles in regulating angiogenesis during embryonic development and tumorigenesis.15–17 Id1/Id3 double-knockout embryos display vascular malformations in the forebrain, leading to fatal hemorrhage.15 Partial Id1 and Id3 deficiencies impaired tumor angiogenesis in tumor-transplanted15,16 and spontaneously tumorigenic murine models.17,18 Id1 and Id3 are highly expressed in tumor vasculature ECs in comparison to ECs from normal tissue.17 Furthermore, ectopic expression of Id1 on cultured ECs enhanced capillary-like tube formation, reflecting angiogenesis, in vitro.19 Therefore, we speculated that transplantation of mature ECs, modified with Id1 gene transfer, would enhance neovascularization. To test this hypothesis, we examined whether Id1-transfected human umbilical vein ECs (HUVECs) improved the rate of limb salvage in a mouse model of hindlimb ischemia. For this purpose, we used a mononegavirus vector based on the Sendai virus (SeV), which achieved highly efficient gene transfer into HUVECs. The present results demonstrate that Id1 gene transfer conferred angiogenic properties on HUVECs in vitro and in vivo. Strikingly, transplantation of Id1-transfected HUVECs rescued ischemic limbs from autoamputation in athymic nude mice.
Methods
Antibodies
Polyclonal rabbit anti-SeV antibody was described previously.20 Rabbit anti-Id1 and goat anti–angiopoietin-1 (Ang1) antibodies were purchased form Santa Cruz Biotechnologies. Mouse antibodies against human CD31 were from PharMingen, mouse anti-human Ki67 antibodies from Zymed, mouse anti–-actin antibody from Sigma, rabbit anti–von Willebrand factor (vWF) antibody and biotinylated and horseradish peroxidase (HRP)–conjugated antibodies against goat immunoglobins from DAKO, and rhodamine- and HRP-conjugated antibodies against rabbit immunoglobins and HRP-conjugated antibody against mouse immunoglobins from Biosource.
Preparation of SeV Vector
The mouse Id1 cDNA was cloned by reverse transcription–polymerase chain reaction (RT-PCR) from total RNA from mouse E10.5 embryos with the use of the following primers: 5'AAgcggccgc-CGTACGCCGTAGTAAGAAAAACTTAGGGTGAAAGTTCTC-AGGATCATGAAGGTCGCCAG/TTgcggccgcGACGTCCGCCTCAGCGACACAAGATGCGATC3'. Underlined portions indicate a set of SeV E and S signals connected to the conserved intergenic trinucleotide; the lower case letters represent NotI restriction sites. The 510-bp fragments amplified with these primers were purified, digested with NotI, and introduced into the NotI site of pSeV18c(+).21 Then viruses were recovered with the use of LLCMK2 cells and hen eggs according to previously described procedures.21 Viral titers were determined with the use of a standard chicken red blood cell hemagglutination assay and a plaque assay on CV1 cells.
Cell Culture and Gene Transfer
HUVECs were purchased from Clonetics and maintained in endothelial cell basal medium (Clonetics, Inc) according to the supplier’s instructions. HUVECs (3 passages) were transfected with 5 to 10 multiplicities of infection of recombinant SeV encoding murine Id1 (SeV/Id1), recombinant SeV encoding murine Ang1 (SeV/Ang1), or the control empty vector (SeV/Null). After 1 hour, the viruses were removed, and the cells were cultured for 48 hours in complete medium before each experiment. For analyses of major angiogenic factors and receptors, HUVECs transfected with SeV were recovered in complete medium for 24 hours, after which the medium was substituted with fresh medium-199 containing 0.5% FCS, and the cells were incubated for 24 hours before the assays. HUVECs transfected with SeV were dissociated from culture dishes with the use of appropriate volumes of Hanks’-based cell dissociation buffer (Invitrogen). The conditioned media were collected, centrifuged, and used for Matrigel assay.
Reverse Transcription–Polymerase Chain Reaction
Gene expression was evaluated by semiquantitative RT-PCR analysis. Total RNA was extracted with the use of Isogen (Nippon gene), and samples (1 μg) were then reverse-transcribed with the use of Superscript II (Life Technologies) with random hexamer primers. The resultant cDNAs were amplified with Taq polymerase (Takara) in a thermocycler. Sequences of specific primers used were as follows: for SeV-delivered murine Id1, 5'CCAGTGGCAGTGCCGCAGCCGCTGCA/GCGACGTCC GCCTCAGCGACACAA 3' (449-bp PCR product); for vascular endothelial growth factor (VEGF), 5'TTCTGTATCAGTCTTTCCTGGTGAG/CGAAGTGGTGAAGTTCATGGATG3' (403-, 535-, and 607-bp PCR product corresponding to VEGF121, VEGF165, and VEGF189, respectively)22; for Ang1, 5'GGTCAGAAGAAAGGAGCAAG/TGGTAGCCGTGTGGTTCTGA3' (437-bp PCR product)23; for angiopoietin-2 (Ang2), 5'GGATCTGGGGAGAGAGGAAC/CTCTGCACCGAGTCATCGTA3' (516-bp PCR product)23; for Flk-1 (KDR), 5'CAACAAAGTCGGGAGAGGAG/ATGACGATGGACAAGTAGCC3'(816-bp PCR product)1; for Flt1, 5'CGACCTTGGTTGTGGCTGACT/CCCTTCTGGTTGGTGGCTTTG3' (656-bp PCR product)24; for TIE2, 5'GGTTCCTTCATCCATT/GTCCTTCCCATAAACC3' (275-bp PCR product)23; and for GAPDH, 5'TGAAGGTCGGAGTCAACGGATTTG/CATGTGGGCCATGAGGTCCACCAC3' (983-bp PCR product).1 Thermal cycling was performed for 25 to 30 cycles to maintain PCR conditions within the linear range of amplification before saturation was reached. Each cycle consisted of 30 seconds of denaturation at 94°C, 1 minute of annealing at each annealing temperature (54°C for TIE2; 60°C for Ang1 and Ang2; and 64°C for Id1, VEGF, Flk-1, Flt-1, and GAPDH), and 1 minute of extension at 72°C.
Western Blotting
HUVECs were washed twice with cold PBS and suspended in a cold lysis buffer (25 mmol/L HEPES, pH 7.5, 300 mmol/L NaCl, 1.5 mmol/L MgCl2, 0.2 mmol/L EDTA, 1% Triton X-100, 0.5 mmol/L dithiothreitol, 20 mmol/L -glycerophosphate, 0.1% SDS, 0.5% sodium deoxycholate, protease inhibitors [complete, Roche]) for Id1 protein assay or a cold lysis buffer (20 mmol/L HEPES, pH 7.5, 150 mmol/L NaCl, 1 mmol/L EDTA, 0.5% Triton X-100, protease inhibitors [complete, Roche]) for the Ang1 protein assay. Similar quantities of the soluble fractions were separated by SDS-PAGE, transferred onto a polyvinylidene difluoride membrane (Millipore), and immunoblotted with the first antibody. The membrane was then incubated with HRP-conjugated secondary antibody, and the signals were visualized with ECL-plus reagent (Amersham). After detection of Ang1 protein, the -actin protein levels were assayed with the use of the same membrane.
Immunocytochemistry
HUVECs were fixed in 4% paraformaldehyde or 5% dimethyl sulfoxide/methanol for 20 minutes at 4°C, permeabilized with 0.2% Triton X-100 for 10 minutes at room temperature, and blocked in 2% skim milk/PBS for 20 minutes at room temperature. Cells were then incubated with the appropriate antibody for 1 hour at room temperature or overnight at 4°C. After extensive washing with PBS-T, cells were incubated with the appropriate secondary antibody; for Ang1 staining, the signal was subsequently amplified with an avidin-biotin complex system with a Vectastain ABC kit (Vector). If needed, nuclei were stained with Hoechst 33258, and cells were then examined with a computer-assisted microscope (Olympus).
Migration Assay
EC migration was evaluated with the use of scratch wound assays.25 Confluent HUVECs in pretraced 24-well culture plates were serum-starved with medium-199 containing 0.5% FCS for 6 hours, after which a portion of the cell monolayer was scraped away with a sterile disposable rubber policeman. The remaining cells were gently washed with medium and incubated for 24 hours in medium-199 containing 0.5% FCS. EC migration from the edge of the injured monolayer was quantified by measuring the area between the wound edges, at a random position, before and after incubation with the use of a computer-assisted microscope (Olympus).
DNA Synthesis Assay
DNA synthesis was evaluated by performing a 5-bromo-2'deoxyuridine (BrdU) incorporation assay with a commercially available kit (Boehringer Mannheim) according to the supplier’s instructions. Briefly, 5x103 HUVECs were plated onto a 96-well plate and then incubated in complete medium for 24 hours. Next, HUVECs were serum-starved with medium-199 containing 0.5% FCS for 6 hours, after which BrdU was added to the medium, followed by further incubation for 8 hours for labeling of HUVECs with or without 10 ng/mL of VEGF (R&D). BrdU incorporation was evaluated by the ELISA method with anti-BrdU antibody and quantified by measuring absorbance at 405/492 nm with a microplate reader (M-Emax, Molecular Devices).
Apoptosis Assay
Apoptosis was induced in HUVECs by the serum deprivation method, as previously described.26 Briefly, HUVECs were seeded onto 0.2% gelatin-precoated 10-cm culture dishes or 2-well chamber slides to a density of 1x106 cells per well or 1x105 cells per well, respectively, and cultured in complete medium for 24 hours. The cells were extensively washed with serum-free medium-199 and then incubated for 24 hours in serum-free medium-199. All floating and adherent cells in a 10-cm culture dish were stained with fluorescein isothiocyanate (FITC)–conjugated annexin V and propidium iodide (PI) with the use of a commercially available kit (Amersham) as previously described.27 Thereafter, samples were analyzed by flow cytometry (FACScaliber, Becton Dickinson) for viable (annexin V–negative and PI-negative), early apoptotic (annexin V–positive, PI-negative), and late apoptotic/secondary necrotic (annexin V–positive and PI-positive) cells. The extent of the apoptosis was quantified as a percentage of annexin V–positive cells.27 Apoptosis was also assayed by a terminal deoxynucleotidyl transferase–mediated dUTP nick-end labeling (TUNEL) staining method with a commercial kit (Takara). Briefly, after removal of the floating cells, the adherent cells in a 2-well chamber slides were fixed and stained according to the supplier’s instructions. The mean number of apoptotic (TUNEL-positive) cells from 3 random fields (x40) in each well was calculated.
In Vitro Angiogenesis Model
Formation of capillary-like structures by HUVECs was assessed on growth factor–reduced Matrigel, a basement membrane matrix preparation (Becton Dickinson). HUVECs were incubated with medium-199 containing 0.5% FCS for 6 hours, after which the HUVECs were plated onto Matrigel (250 μL)-coated 24-well plates to a density of 5x104 cells per well and incubated in medium-199 containing 0.5% FCS for 1 hour. Unattached cells were then removed by gentle aspiration, and the medium was replaced with medium-199 recombinant human IgG1 Fc (IgG Fc) (R&D) or recombinant human Tie2/Fc chimeric protein (Tie2/Fc) (R&D). In another set of experiments, HUVECs were incubated with the conditioned media from SeV/Id1-HUVECs (SeV/Id1-CM) or SeV/Null-HUVECs (SeV/Null-CM) in the presence or absence of IgG Fc or Tie2/Fc. Formation of tube/cordlike structures was evaluated 8 hours later. The cultures were photographed with a CCD camera (Olympus), and with the use of NIH Image software, the total length of the tube/cordlike structures was quantified as an indicator of angiogenesis. The mean value from 5 random fields (x40) in each well was calculated.
In Vivo Angiogenesis Model
Animal care and use in our laboratory were in strict accordance with the guidelines for animal and recombinant DNA experiments put forth by Kumamoto University. Male athymic mice (BALB/c, 8 to 10 weeks old) were anesthetized with intraperitoneal injection of 160 mg/kg pentobarbital. To create the hindlimb ischemia model, the femoral artery and vein were excised, from just above the deep femoral arteries to the popliteal artery and vein. For cell transplantation intervention, 1x105 HUVECs were suspended in 50 μL of serum-free medium-199, and the solution was injected into 5 different areas of the leg muscles (3 in the thigh and 2 in the calf, 10 μL each) 1 day after the surgical procedures. Blood flow recovery after ischemia was serially evaluated with laser-Doppler perfusion imaging (LDPI) (Moor Instruments) over the course of 3 weeks, postoperatively, as previously described.28 To minimize signal variation caused by variations in ambient light and temperature, calculated blood perfusion (relative units) was expressed as the ischemic (right)/normal (left) limb blood perfusion ratio. At the end of the study, tissue samples were harvested from the lower calf and thigh muscles for histological analysis.
For transplanted HUVEC tracking, HUVECs were labeled with a red fluorescent marker, PKH26-RED (Sigma), according to the supplier’s instructions. The labeled HUVECs were rinsed, resuspended in serum-free medium-199, and then transplanted into the hindlimb muscles of a subgroup of mice as described above. For another cell-tracking study, a subgroup of mice received an intracardiac injection of FITC-conjugated Bandeiraea simplicifolia lectin I (BS-1; Sigma) and rhodamine-conjugated UEA-1 (Sigma) simultaneously or of FITC-conjugated UEA-1 (Sigma) only at 30 minutes before euthanasia.29
Histological Analysis
Tissue samples harvested from the hindlimb muscles were snap-frozen with precooled isopentane in liquid nitrogen, after which they were embedded in OCT compound (Miles) and cut into 5- to 7-μm-thick sections. For evaluation of capillary density, serving as an anatomic index of angiogenesis, capillary ECs were detected by staining tissue sections for alkaline phosphatase (AP) with the use of the iodoxyl tetrazolium method28 and counterstaining with eosin. Some additional sections were stained with anti-mouse CD31 antibody to further confirm the EC phenotype. The capillary ECs were then counted under light microscopy (x100) to determine capillary density. In each animal, 6 different fields of a section were randomly selected for EC counts, which were then averaged. Tissue sections obtained from hindlimb muscles transplanted with PKH26-RED–labeled HUVECs were also stained for AP to detect viable ECs immediately after being photographed with a CCD camera (Olympus). Tissue sections obtained from hindlimb muscles after premortem lectin staining were observed with a computer-assisted microscope (Nikon) (x400), and UEA-1–positive HUVECs were counted (x200) to quantify the incorporation of HUVECs into the murine vasculature. For each animal, 3 different fields of a section were randomly selected for cell counting. Tissue sections from hindlimb muscles treated only with UEA-1 were stained with anti-human vWF to further confirm the incorporation of HUVECs into the endothelial layer.
To evaluate the percentage of the SeV-transfected population in HUVECs incorporated into the ischemic limbs, we double-immunostained the limb section with anti-human CD31 and anti-SeV antibodies. The numbers of human CD31- and/or SeV-positive cells were counted for the sections randomly obtained from 3 different animals.
To evaluate the proliferative state of the transplanted HUVECs, we double-immunostained the limb section with an anti-human Ki67 antibody and UEA-1. The numbers of human Ki67- and/or UEA-1–positive cells were counted in the sections randomly obtained from 2 different animals.
Statistical Analysis
Each assay experiment was performed at least 3 times. Data are expressed as mean±SD. Student t test was used to compare continuous values between 2 groups. Differences in limb outcome between 2 groups were analyzed by the 2 test. Values of P<0.05 were considered to represent statistically significant differences.
Results
SeV-Mediated Id1 Gene Expression and Protein Synthesis
SeV-mediated Id1 gene expression and protein synthesis in HUVECs were examined by RT-PCR, Western blot, and immunocytochemical analyses. Immunostaining for SeV confirmed that &70% of HUVECs had been successfully infected with SeV within 2 days (data not shown). RT-PCR confirmed the presence of SeV-mediated Id1 mRNA in HUVECs infected with SeV/Id1 (SeV/Id1-HUVECs) 2 days after transfection but not in HUVECs transfected with SeV/Null (SeV/Null-HUVECs) (Figure 1A). Western blot analysis with anti-Id1 antibody detected a 14-kDa protein in the cell lysates of SeV/Id1-HUVECs (Figure 1B). In SeV/Id1-HUVECs, Id1 was predominantly localized to the nucleus (Figure 1C).
Id1 Gene Transfer Confers Angiogenic Activity on HUVECs
To examine the angiogenic effect of Id1 gene transfer, scratch wound assays were performed on HUVECs under serum-reduced and growth factor–free conditions. Id1 increased HUVEC migration up to 144% of the control level after 24 hours (P<0.05) (Figure 2A), as previously reported in other types of ECs.19,30 Id1 gene transfer also stimulated cell growth irrespective of the presence of VEGF as estimated by BrdU incorporation (Figure 2B).
In the Matrigel tube formation assay, SeV/Id1-HUVECs formed robust elongated tube/cordlike structures (Figure 2C). Quantitative analysis showed total tube/cord lengths to be significantly greater in the SeV/Id1-HUVECs than in the SeV/Null-HUVECs (P<0.01) (Figure 2D). These findings clearly suggest that Id1 gene transfer confers angiogenic activity on HUVECs in vitro.
Id1 Gene Transfer Protects Against HUVEC Apoptosis
Previous reports have suggested that Id1 can both positively and negatively regulate apoptosis depending on cell types.31,32 To examine the effects of Id1 on EC apoptosis, we cultured HUVECs in serum-free medium in the presence or absence of SeV/Id1 and evaluated apoptotic cells using flow cytometry. Id1 gene transfer reduced the number of annexin V–positive cells to 47% of the control level (Figure 3A). TUNEL staining also showed that Id1 significantly reduced the number of TUNEL-positive cells down to 40% of the control number (Figure 3B). Representative images of HUVECs with TUNEL staining are shown in Figure 3C. These results show clearly that Id1 can protect HUVECs against apoptosis in vitro.
Transplantation of Id1-Transfected HUVECs Augments Revascularization of Murine Ischemic Hindlimbs
We assessed the angiogenic potency of Id1-transfected HUVECs for therapeutic purposes by transplanting them into murine ischemic hindlimbs. One day after surgical induction of ischemia, 1x105 HUVECs were directly transplanted into the hindlimb muscles of the athymic mice. Blood perfusion recovery after transplantation was then serially assessed with LDPI. Ischemia/normal limb perfusion ratios before surgery were similar in the treatment groups: medium (n=14), SeV/Null-HUVECs (n=11), and SeV/Id1-HUVECs (n=14) (Figure 4A). Immediately after surgery, limb perfusion was severely reduced in all groups to the same extent (Figure 4A), indicating the ischemia severities to be comparable. Over the next 21 days, LDPI revealed gradual recovery of blood perfusion in all 3 groups. Remarkably, recovery was accelerated in the SeV/Id1-HUVECs group. Ischemia/normal limb perfusion ratios on postoperative days 14 and 21 (0.71±0.25 and 0.71±0.12, respectively) were significantly higher in transplanted hindlimbs than in those of the medium-only (0.46±0.18, P<0.01 and 0.48±0.20, P<0.01, respectively) and SeV/Null-HUVECs (0.52±0.20, P<0.05 and 0.54±0.13, P<0.01, respectively) groups (Figure 4A). Recovery of blood perfusion was similar in the SeV/Null-HUVECs and medium-only groups. We also examined increases in transplanted cell numbers up to 5x105 cells and obtained similar results (data not shown). Representative images of hindlimb blood perfusion recorded 21 days after surgical induction of ischemia are shown in Figure 4B.
We then examined whether the recovery of blood flow in this ischemic hindlimb model had resulted from neovascularization. Figure 4C represents photomicrographs of representative ischemic limb sections obtained on day 21. Consistent with the LDPI analysis, immunohistochemical staining for AP revealed larger numbers of capillary ECs in the hindlimbs of the SeV/Id1-HUVECs group (80.5±12.8 number per field) than in the medium-only (47.2±10.5, P<0.01) and in the SeV/Null-HUVECs (43.2±10.5, P<0.05) groups, but there was no significant difference between the latter 2 (Figure 4D).
Increased neovascularization in the mouse ischemic hindlimb can potentially lead to important biological consequences, including limb salvage. To examine limb outcome after induction of ischemia, the appearance of the ischemic hindlimb was observed on the last day of the study period (day 21 after surgery). Among 14 mice in the medium-only group, 7 (50.0%) showed toe necrosis, and 1 (7.1%) showed severe limb necrosis, including autoamputation (Figure 4E). Only 6 (42.9%) were devoid of necrotic lesions (Figure 4E). In contrast, transplantation of SeV/Id1-HUVECs significantly increased the rate of complete limb salvage up to 92% (Figure 4E). Transplantation of SeV/Null-HUVECs did not improve limb outcomes (Figure 4E). Taken together, these findings indicate that transplantation of SeV/Id1-HUVECs, but not of SeV/Null-HUVECs, can rescue ischemic lesions and thereby prevent catastrophic outcomes by augmenting neovascularization.
Evidence of Incorporation of Transplanted HUVECs Into Murine Hindlimb Vasculature
To determine whether transplanted HUVECs contribute to vascular structure regeneration, we labeled HUVECs with PKH26-RED before transplantation and examined their incorporation into capillaries on day 21. After inspection of tissue sections harvested from the ischemic limb for fluorescent-labeled HUVECs, the same sections were stained for AP to detect viable HUVECs. HUVECs double positive for PKH26-RED and AP were documented in the sites of hindlimb capillaries in SeV/Id1-HUVECs, showing incorporation of transplanted HUVECs into the murine hindlimb vasculature (Figure 5A, 5B). To further confirm and quantify these observations, HUVECs and murine ECs were identified with the use of premortem staining by intracardiac injection of fluorescent conjugated lectins (UEA-1 specific for human ECs and BS-1 specific for murine ECs). Stereomicroscopic examination of lectin-stained muscle blocks revealed numerous incorporated HUVECs in the hindlimb vasculature of the SeV/Id1-HUVECs group (Figure 5C to 5E). By contrast, only a few HUVECs were detected in the hindlimb vasculature in the SeV/Null-HUVECs group (Figure 5F). In addition, tissue sections were quantitatively analyzed for incorporated HUVECs. Representative photomicrographs of sections from both groups are presented in Figure 5G, 5H. The number of incorporated HUVECs in the SeV/Id1-HUVECs group was &5 times that of the SeV/Null-HUVECs group (Figure 5I). At higher magnification, the capillary wall proved to be partly composed of UEA-1–positive HUVECs (Figure 5J, 5K). In addition, UEA-1–positive HUVECs resided in the endothelial layer positive for vWF (Figure 5L, 5M). These results confirm that Id1-transfected HUVECs can be effectively incorporated into regenerating capillaries in ischemic lesions.
To evaluate whether Id1-transfected HUVECs were incorporated into the ischemic limbs, we double-immunostained the section obtained from 14 days after HUVEC transplantation with anti-human CD31 and anti-SeV antibodies. Cells positive for both human CD31 and SeV were regarded as SeV/Id1-transfected HUVECs. Among 243 HUVECs positive for human CD31, 211 cells (87%) were positive for SeV, indicating that the majority of the incorporated HUVECs overexpressed Id1. A representative photograph of human CD31- and SeV-positive cells is shown in Figure 5N to 5P.
To address the mechanism underlying the in vivo effects of Id1, we evaluated whether transplanted HUVECs are in a proliferative state by double-immunostaining the section obtained from hindlimb muscles 7 days after HUVEC transplantation with an anti-human Ki67 antibody and UEA-1. We observed that 6.5% of UEA-1–positive HUVECs were Ki67 positive in the section from the SeV/Id1-HUVEC group, whereas we did not detect any Ki67-positive HUVECs in the section from the SeV/Null-HUVEC group. A representative photograph of Ki67- and UEA-1–positive cells is shown in Figure 5Q to 5S.
Ang1 Is Upregulated and Functionally Contributes to In Vitro Angiogenesis on Id1-Transfected HUVECs
To address the mechanism of the angiogenic potencies in SeV/Id1-HUVECs, the gene expressions of several angiogenic factors and their receptors were assessed on HUVECs 2 days after Id1 gene transfer. Semiquantitative RT-PCR analysis showed the Ang1 gene, but not the VEGF, Ang2, KDR, Flt-1, or TIE2 gene, to be upregulated on SeV/Id1-HUVECs (Figure 6A). Western blot analysis revealed increased levels of Ang1 protein in the cell lysates of SeV/Id1-HUVECs (Figure 6B). In addition, coimmunostaining with anti-Ang1 and anti-CD31 antibodies detected Ang1 in the cytoplasm of SeV/Id1-HUVECs (Figure 6C). Staining for SeV/Ang1-transfected HUVECs served as a positive control (Figure 6D).
To evaluate whether upregulated Ang1 functionally contributes to the angiogenic property of SeV/Id1-HUVECs, we examined the effect of Tie2/Fc in a Matrigel tube formation assay. In Matrigel, the formation of elongated tube/cordlike structures in SeV/Id1-HUVECs was suppressed by Tie2/Fc (Figure 6E to 6H). Quantitative analysis showed total tube/cord lengths to be significantly smaller in SeV/Id1-HUVECs treated with Tie2/Fc than in those treated with IgG Fc (P<0.05) (Figure 6I). Total tube/cord length remained significantly greater in SeV/Id1-HUVECs treated with Tie2/Fc than in SeV/Null-HUVECs even if the used dosage of Tie2/Fc was increased (P<0.05) (Figure 6I). We next evaluated whether conditioned media from SeV/Id1-HUVECs (SeV/Id1-CM) also stimulate tube/cord formation of HUVECs in Matrigel. Total tube/cord lengths of HUVECs on Matrigel were significantly greater in the HUVECs treated with SeV/Id1-CM than in those with SeV/Null-CM (P<0.01) (Figure 6J); furthermore, the effects of SeV/Id1-CM on tube/cord formation of HUVECs were again partially suppressed by the presence of Tie2/Fc. A previous study reported that not only Ang1 but also Ang2 might activate Tie2 receptor.33 Because Ang1 but not Ang2 was upregulated in SeV/Id1-HUVECs in the present study, the inhibitory effects of tube/cord formation by Tie2/Fc were considered due to the suppression of Ang1/Tie2 signaling. Collectively, these results suggest that upregulated Ang1 functionally contributes to the angiogenic effects of Id1 and that additional angiogenic factor(s) may also be upregulated in SeV/Id1-HUVECs.
Discussion
In the present study we have demonstrated that gene transfer of Id1, a member of the HLH protein family, confers an angiogenic property on HUVECs in vitro and in vivo. Id1 gene transfer enhanced migration, proliferation, and capillary-like tube/cord formation of HUVECs. Id1 gene transfer also protected HUVECs from serum deprivation–induced apoptosis. Transplantation of SeV/Id1-HUVECs accelerated vascular regeneration in the murine ischemic limb and consequently led to limb salvage. Consistently, transplanted SeV/Id1-HUVECs were effectively incorporated into the regenerating vasculature. Finally, we documented upregulation of Ang1 in SeV/Id1-HUVECs, which functionally contributed to enhanced in vitro angiogenesis.
In previous studies,29,34,35 the angiogenic potency of EPC was investigated in hindlimb ischemia with the use of a similar mouse model, in which systemic transplantation of ex vivo expanded EPCs dramatically accelerated blood flow recovery and increased capillary densities in the ischemic limb. Beneficial effects of local transplantation of EPCs have also been shown in other animal models.36,37 Kim and colleagues11 transplanted ECs isolated form the aorta directly into the scar sites of rat hearts 2 weeks after cryoinjury. Transplanted ECs were incorporated into the new vessels and increased regional perfusion in myocardial scar tissue but failed to improve global function. Chekanov and colleagues38 administered autologous ECs into the ischemic myocardium of sheep using a fibrin matrix and demonstrated enhancement of neovascularization with improved left ventricular function. In tumor angiogenesis models,39,40 implanted mature ECs similarly contributed to host neovascularization and tumor growth; furthermore, genetic modification of the ECs with the antiapoptotic gene Bcl-2 enhanced the effects of cell implantation. These findings suggest that mature ECs have the potential to be incorporated into regenerating vessels in vivo under certain conditions.
The present results demonstrate that mature human ECs overexpressing the Id1 gene can be effectively incorporated into regenerating vessels in murine ischemic limbs. The extents of angiogenic and limb salvage effects were comparable to those of EPCs.29,34,35 These results indicate that mature ECs could potentially be optimized for therapeutic angiogenesis by inducing phenotypic changes through appropriate gene transfers. Id1 gene transfer stimulated migration, proliferation, and tube formation of HUVECs in vitro, as previously shown in bovine aortic ECs19 and HUVECs.41 Migration and proliferation of ECs is considered a critical step for physiological as well as therapeutic angiogenesis.5,34,35 In the present analysis, a part of transplanted SeV/Id1-HUVECs but not SeV/Null-HUVECs were in a proliferative state, indicating that stimulation of proliferation may contribute to the in vivo angiogenic effects of Id1. In addition, Id1 inhibited apoptosis of HUVECs under serum deprivation conditions. Id1 has been reported to both positively and negatively regulate apoptosis, depending on cell types and experimental conditions.31,32,41 Sakurai et al41 have reported that overexpression of Id3, but not Id1, inhibits HUVEC apoptosis induced by serum starvation. The discrepancy between their results and ours may be explained by differences in culture condition and/or in Id1 expression levels. A previous study showed Id1 to delay the onset of replicative senescence in human ECs42; Id1 may thereby increase the survival of SeV/Id1-HUVECs and contribute to angiogenesis.
In the present study, Id1 gene transfer upregulated the expression of Ang1 but not Ang2 in HUVECs. Furthermore, the present study revealed that upregulated Ang1 functionally contributed to the in vitro angiogenic effects of Id1 as an autocrine/paracrine factor. Ang1 is reported to augment EC migration and protect ECs against apoptosis partly via the activation of Tie2/PI3'-kinase/Akt signal transduction pathway.26,43,44 Upregulation of the signaling pathway may partly explain an increase in the survival of SeV/Id1-HUVECs. Although it remains unknown whether the effect of Id1 on Ang1 expression is direct, Ang1 upregulation may contribute to the mechanism underlying the in vivo angiogenic effect of Id1-transfected HUVECs, together with other possible contributions of metalloproteinases, integrins, and fibroblast growth factor receptor genes, as suggested by previous studies using Id gene–deleted mice15,18 and Id1-transfected ECs.41
We analyzed endogenous Id1 expression levels with immunohistochemical technique in the tissue section of murine ischemic limbs. However, we could not detect significant upregulation of Id1 in ECs (K. Nishiyama, MD, et al, unpublished data, 2003). In addition, our preliminary experiments showed that deferoxamine, an agent mimicking hypoxic condition, downregulated Id1 expression in cultured HUVECs, although it caused VEGF upregulation (K. Nishiyama, MD, et al, unpublished data, 2004), suggesting that Id1 expression is not induced in ischemic lesion. This result suggests that ischemia-induced suppression of Id1 expression may overcome the inductive effect of VEGF on Id1 expression. Therefore, Id1 gene transfer is expected to complement the downregulated activity of Id1 to confer angiogenic capacity on ECs.
Contrary to the beneficial angiogenic effect, it has been reported that the Id1 transgene may induce neoplastic transformation of ECs.13,45 In a previous study,42 however, Id1-transferred ECs were not immortalized and eventually underwent senescence despite high Id1 levels. In addition, no tumor formation was pathologically observed in the lesions with transplanted SeV/Id1-HUVECs in our present study (data not shown). In addition, the ex vivo gene transfer strategy may preclude recipients from being exposed to the Id1 gene–bearing vector. Thus, the combination of Id1 gene transfer and transplantation using mature ECs may serve as a novel and useful application for therapeutic revascularization with high efficiency and low risk.
Acknowledgments
This work was supported in part by grants from the Japan Society for the Promotion of Science Research for the Future Program; grants-in-aid for scientific research from the Ministry of Education, Culture, Sports, Science, and Technology, Japan; and a research grant for cardiovascular diseases (14C-1 and 14C-4) from the Ministry of Health, Labor, and Welfare.
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Department of Physiological Chemistry and Metabolism, Division of Biochemistry and Molecular Biology (Y.K., H.K.), and Department of Developmental and Medical Technology (Sankyo) (K.N.), University of Tokyo Graduate School of Medicine, Tokyo, Japan
Department of Viral Infection and Vaccine Control, National Institute for Infectious Diseases (A.K.), Tokyo, Japan.
Abstract
Background— Transplantation of endothelial progenitor cells has been proposed as a potential strategy for therapeutic revascularization. However, the limited endogenous cell pool and the related technical difficulties constitute clinically important disadvantages to autologous transplantation. In this study we investigated whether fully differentiated endothelial cells (ECs) modified with gene transfer of Id1, a helix-loop-helix transcription factor involved in angiogenesis, have the potential to contribute to therapeutic angiogenesis.
Methods and Results— The Id1 gene was transferred into human umbilical vein ECs (HUVECs) via a Sendai virus vector. Id1 stimulated migration, proliferation, and capillary-like tube/cord formation of HUVECs. In addition, Id1 reduced serum deprivation–induced HUVEC apoptosis, as shown by FACS analysis with annexin V and TUNEL staining. Transplantation of Id1-overexpressing HUVECs accelerated recovery of blood flow as evaluated by laser-Doppler perfusion imaging, increased capillary density, and improved the rate of limb salvage compared with the transplantation of control HUVECs. Histochemical analysis revealed that the regenerated vascular networks of limbs transplanted with Id1-overexpressing HUVECs contained numerous HUVECs, some of which were in a proliferative state. Untransfected HUVECs were also incorporated with Id1-transfected HUVECs, suggesting the noncell autonomous effect of Id1. Finally, angiopoietin-1 was upregulated in Id1-overexpressing HUVECs and functionally contributed to the in vitro angiogenic effect of Id1.
Conclusions— Id1 gene transfer conferred HUVECs with an angiogenic property, contributing to neovascularization after transplantation into ischemic lesions. Transplantation of Id1-overexpressing mature ECs may serve as a novel and useful strategy for therapeutic angiogenesis.
Key Words: endothelial cells genes therapeutic angiogenesis
Introduction
The discovery of circulating endothelial progenitor cells (EPCs) has provided a new mechanistic understanding of neovascularization.1,2 In response to cytokines, EPCs are mobilized from the bone marrow, recruited to the site of wound healing, tissue ischemia, and tumor growth, and then differentiated into vascular endothelial cells (ECs).1–5 This process, termed postnatal vasculogenesis, is postulated, together with angiogenesis, to contribute to regional neovascularization.4 Several animal experiments and human trials under way are focused on the potential application of EPCs to vascular regenerative therapy.6
Although the usefulness of EPCs for therapeutic neovascularization is highly anticipated, several issues remain unresolved. Most notably, the limited cell pool and technical difficulties in expanding EPCs in vitro have prevented widespread clinical use of EPCs for autologous transplantation. Differentiated vascular ECs, on the other hand, can be isolated and expanded in vitro.7,8 Moreover, a large number of mature ECs can be differentiated from bone marrow–derived progenitor cells and embryonic stem cells.9,10 Thus, it would be advantageous for mature ECs to serve as cell sources for neovascularization. However, mature ECs are inefficiently incorporated into forming vessels, such that their contribution to therapeutic angiogenesis is limited.11 The fact that ECs in adult organs transform phenotypically into "angiogenic" cells under pathological circumstances and participate in neovascularization5 prompted us to hypothesize that quiescent ECs may become "angiogenic" via phenotypic modification with gene transfer.
The Id proteins are a family of helix-loop-helix (HLH) transcription factors.12 Id proteins lack a basic DNA binding domain and primarily act as inhibitors of basic HLH transcription factors, via heterodimerization.12 Through this mechanism, Id proteins have been implicated in regulating a variety of cellular processes including cell growth, senescence, differentiation, and neoplastic transformation.12–14 More recently, accumulating evidence has suggested that Id1 and Id3 play vital roles in regulating angiogenesis during embryonic development and tumorigenesis.15–17 Id1/Id3 double-knockout embryos display vascular malformations in the forebrain, leading to fatal hemorrhage.15 Partial Id1 and Id3 deficiencies impaired tumor angiogenesis in tumor-transplanted15,16 and spontaneously tumorigenic murine models.17,18 Id1 and Id3 are highly expressed in tumor vasculature ECs in comparison to ECs from normal tissue.17 Furthermore, ectopic expression of Id1 on cultured ECs enhanced capillary-like tube formation, reflecting angiogenesis, in vitro.19 Therefore, we speculated that transplantation of mature ECs, modified with Id1 gene transfer, would enhance neovascularization. To test this hypothesis, we examined whether Id1-transfected human umbilical vein ECs (HUVECs) improved the rate of limb salvage in a mouse model of hindlimb ischemia. For this purpose, we used a mononegavirus vector based on the Sendai virus (SeV), which achieved highly efficient gene transfer into HUVECs. The present results demonstrate that Id1 gene transfer conferred angiogenic properties on HUVECs in vitro and in vivo. Strikingly, transplantation of Id1-transfected HUVECs rescued ischemic limbs from autoamputation in athymic nude mice.
Methods
Antibodies
Polyclonal rabbit anti-SeV antibody was described previously.20 Rabbit anti-Id1 and goat anti–angiopoietin-1 (Ang1) antibodies were purchased form Santa Cruz Biotechnologies. Mouse antibodies against human CD31 were from PharMingen, mouse anti-human Ki67 antibodies from Zymed, mouse anti–-actin antibody from Sigma, rabbit anti–von Willebrand factor (vWF) antibody and biotinylated and horseradish peroxidase (HRP)–conjugated antibodies against goat immunoglobins from DAKO, and rhodamine- and HRP-conjugated antibodies against rabbit immunoglobins and HRP-conjugated antibody against mouse immunoglobins from Biosource.
Preparation of SeV Vector
The mouse Id1 cDNA was cloned by reverse transcription–polymerase chain reaction (RT-PCR) from total RNA from mouse E10.5 embryos with the use of the following primers: 5'AAgcggccgc-CGTACGCCGTAGTAAGAAAAACTTAGGGTGAAAGTTCTC-AGGATCATGAAGGTCGCCAG/TTgcggccgcGACGTCCGCCTCAGCGACACAAGATGCGATC3'. Underlined portions indicate a set of SeV E and S signals connected to the conserved intergenic trinucleotide; the lower case letters represent NotI restriction sites. The 510-bp fragments amplified with these primers were purified, digested with NotI, and introduced into the NotI site of pSeV18c(+).21 Then viruses were recovered with the use of LLCMK2 cells and hen eggs according to previously described procedures.21 Viral titers were determined with the use of a standard chicken red blood cell hemagglutination assay and a plaque assay on CV1 cells.
Cell Culture and Gene Transfer
HUVECs were purchased from Clonetics and maintained in endothelial cell basal medium (Clonetics, Inc) according to the supplier’s instructions. HUVECs (3 passages) were transfected with 5 to 10 multiplicities of infection of recombinant SeV encoding murine Id1 (SeV/Id1), recombinant SeV encoding murine Ang1 (SeV/Ang1), or the control empty vector (SeV/Null). After 1 hour, the viruses were removed, and the cells were cultured for 48 hours in complete medium before each experiment. For analyses of major angiogenic factors and receptors, HUVECs transfected with SeV were recovered in complete medium for 24 hours, after which the medium was substituted with fresh medium-199 containing 0.5% FCS, and the cells were incubated for 24 hours before the assays. HUVECs transfected with SeV were dissociated from culture dishes with the use of appropriate volumes of Hanks’-based cell dissociation buffer (Invitrogen). The conditioned media were collected, centrifuged, and used for Matrigel assay.
Reverse Transcription–Polymerase Chain Reaction
Gene expression was evaluated by semiquantitative RT-PCR analysis. Total RNA was extracted with the use of Isogen (Nippon gene), and samples (1 μg) were then reverse-transcribed with the use of Superscript II (Life Technologies) with random hexamer primers. The resultant cDNAs were amplified with Taq polymerase (Takara) in a thermocycler. Sequences of specific primers used were as follows: for SeV-delivered murine Id1, 5'CCAGTGGCAGTGCCGCAGCCGCTGCA/GCGACGTCC GCCTCAGCGACACAA 3' (449-bp PCR product); for vascular endothelial growth factor (VEGF), 5'TTCTGTATCAGTCTTTCCTGGTGAG/CGAAGTGGTGAAGTTCATGGATG3' (403-, 535-, and 607-bp PCR product corresponding to VEGF121, VEGF165, and VEGF189, respectively)22; for Ang1, 5'GGTCAGAAGAAAGGAGCAAG/TGGTAGCCGTGTGGTTCTGA3' (437-bp PCR product)23; for angiopoietin-2 (Ang2), 5'GGATCTGGGGAGAGAGGAAC/CTCTGCACCGAGTCATCGTA3' (516-bp PCR product)23; for Flk-1 (KDR), 5'CAACAAAGTCGGGAGAGGAG/ATGACGATGGACAAGTAGCC3'(816-bp PCR product)1; for Flt1, 5'CGACCTTGGTTGTGGCTGACT/CCCTTCTGGTTGGTGGCTTTG3' (656-bp PCR product)24; for TIE2, 5'GGTTCCTTCATCCATT/GTCCTTCCCATAAACC3' (275-bp PCR product)23; and for GAPDH, 5'TGAAGGTCGGAGTCAACGGATTTG/CATGTGGGCCATGAGGTCCACCAC3' (983-bp PCR product).1 Thermal cycling was performed for 25 to 30 cycles to maintain PCR conditions within the linear range of amplification before saturation was reached. Each cycle consisted of 30 seconds of denaturation at 94°C, 1 minute of annealing at each annealing temperature (54°C for TIE2; 60°C for Ang1 and Ang2; and 64°C for Id1, VEGF, Flk-1, Flt-1, and GAPDH), and 1 minute of extension at 72°C.
Western Blotting
HUVECs were washed twice with cold PBS and suspended in a cold lysis buffer (25 mmol/L HEPES, pH 7.5, 300 mmol/L NaCl, 1.5 mmol/L MgCl2, 0.2 mmol/L EDTA, 1% Triton X-100, 0.5 mmol/L dithiothreitol, 20 mmol/L -glycerophosphate, 0.1% SDS, 0.5% sodium deoxycholate, protease inhibitors [complete, Roche]) for Id1 protein assay or a cold lysis buffer (20 mmol/L HEPES, pH 7.5, 150 mmol/L NaCl, 1 mmol/L EDTA, 0.5% Triton X-100, protease inhibitors [complete, Roche]) for the Ang1 protein assay. Similar quantities of the soluble fractions were separated by SDS-PAGE, transferred onto a polyvinylidene difluoride membrane (Millipore), and immunoblotted with the first antibody. The membrane was then incubated with HRP-conjugated secondary antibody, and the signals were visualized with ECL-plus reagent (Amersham). After detection of Ang1 protein, the -actin protein levels were assayed with the use of the same membrane.
Immunocytochemistry
HUVECs were fixed in 4% paraformaldehyde or 5% dimethyl sulfoxide/methanol for 20 minutes at 4°C, permeabilized with 0.2% Triton X-100 for 10 minutes at room temperature, and blocked in 2% skim milk/PBS for 20 minutes at room temperature. Cells were then incubated with the appropriate antibody for 1 hour at room temperature or overnight at 4°C. After extensive washing with PBS-T, cells were incubated with the appropriate secondary antibody; for Ang1 staining, the signal was subsequently amplified with an avidin-biotin complex system with a Vectastain ABC kit (Vector). If needed, nuclei were stained with Hoechst 33258, and cells were then examined with a computer-assisted microscope (Olympus).
Migration Assay
EC migration was evaluated with the use of scratch wound assays.25 Confluent HUVECs in pretraced 24-well culture plates were serum-starved with medium-199 containing 0.5% FCS for 6 hours, after which a portion of the cell monolayer was scraped away with a sterile disposable rubber policeman. The remaining cells were gently washed with medium and incubated for 24 hours in medium-199 containing 0.5% FCS. EC migration from the edge of the injured monolayer was quantified by measuring the area between the wound edges, at a random position, before and after incubation with the use of a computer-assisted microscope (Olympus).
DNA Synthesis Assay
DNA synthesis was evaluated by performing a 5-bromo-2'deoxyuridine (BrdU) incorporation assay with a commercially available kit (Boehringer Mannheim) according to the supplier’s instructions. Briefly, 5x103 HUVECs were plated onto a 96-well plate and then incubated in complete medium for 24 hours. Next, HUVECs were serum-starved with medium-199 containing 0.5% FCS for 6 hours, after which BrdU was added to the medium, followed by further incubation for 8 hours for labeling of HUVECs with or without 10 ng/mL of VEGF (R&D). BrdU incorporation was evaluated by the ELISA method with anti-BrdU antibody and quantified by measuring absorbance at 405/492 nm with a microplate reader (M-Emax, Molecular Devices).
Apoptosis Assay
Apoptosis was induced in HUVECs by the serum deprivation method, as previously described.26 Briefly, HUVECs were seeded onto 0.2% gelatin-precoated 10-cm culture dishes or 2-well chamber slides to a density of 1x106 cells per well or 1x105 cells per well, respectively, and cultured in complete medium for 24 hours. The cells were extensively washed with serum-free medium-199 and then incubated for 24 hours in serum-free medium-199. All floating and adherent cells in a 10-cm culture dish were stained with fluorescein isothiocyanate (FITC)–conjugated annexin V and propidium iodide (PI) with the use of a commercially available kit (Amersham) as previously described.27 Thereafter, samples were analyzed by flow cytometry (FACScaliber, Becton Dickinson) for viable (annexin V–negative and PI-negative), early apoptotic (annexin V–positive, PI-negative), and late apoptotic/secondary necrotic (annexin V–positive and PI-positive) cells. The extent of the apoptosis was quantified as a percentage of annexin V–positive cells.27 Apoptosis was also assayed by a terminal deoxynucleotidyl transferase–mediated dUTP nick-end labeling (TUNEL) staining method with a commercial kit (Takara). Briefly, after removal of the floating cells, the adherent cells in a 2-well chamber slides were fixed and stained according to the supplier’s instructions. The mean number of apoptotic (TUNEL-positive) cells from 3 random fields (x40) in each well was calculated.
In Vitro Angiogenesis Model
Formation of capillary-like structures by HUVECs was assessed on growth factor–reduced Matrigel, a basement membrane matrix preparation (Becton Dickinson). HUVECs were incubated with medium-199 containing 0.5% FCS for 6 hours, after which the HUVECs were plated onto Matrigel (250 μL)-coated 24-well plates to a density of 5x104 cells per well and incubated in medium-199 containing 0.5% FCS for 1 hour. Unattached cells were then removed by gentle aspiration, and the medium was replaced with medium-199 recombinant human IgG1 Fc (IgG Fc) (R&D) or recombinant human Tie2/Fc chimeric protein (Tie2/Fc) (R&D). In another set of experiments, HUVECs were incubated with the conditioned media from SeV/Id1-HUVECs (SeV/Id1-CM) or SeV/Null-HUVECs (SeV/Null-CM) in the presence or absence of IgG Fc or Tie2/Fc. Formation of tube/cordlike structures was evaluated 8 hours later. The cultures were photographed with a CCD camera (Olympus), and with the use of NIH Image software, the total length of the tube/cordlike structures was quantified as an indicator of angiogenesis. The mean value from 5 random fields (x40) in each well was calculated.
In Vivo Angiogenesis Model
Animal care and use in our laboratory were in strict accordance with the guidelines for animal and recombinant DNA experiments put forth by Kumamoto University. Male athymic mice (BALB/c, 8 to 10 weeks old) were anesthetized with intraperitoneal injection of 160 mg/kg pentobarbital. To create the hindlimb ischemia model, the femoral artery and vein were excised, from just above the deep femoral arteries to the popliteal artery and vein. For cell transplantation intervention, 1x105 HUVECs were suspended in 50 μL of serum-free medium-199, and the solution was injected into 5 different areas of the leg muscles (3 in the thigh and 2 in the calf, 10 μL each) 1 day after the surgical procedures. Blood flow recovery after ischemia was serially evaluated with laser-Doppler perfusion imaging (LDPI) (Moor Instruments) over the course of 3 weeks, postoperatively, as previously described.28 To minimize signal variation caused by variations in ambient light and temperature, calculated blood perfusion (relative units) was expressed as the ischemic (right)/normal (left) limb blood perfusion ratio. At the end of the study, tissue samples were harvested from the lower calf and thigh muscles for histological analysis.
For transplanted HUVEC tracking, HUVECs were labeled with a red fluorescent marker, PKH26-RED (Sigma), according to the supplier’s instructions. The labeled HUVECs were rinsed, resuspended in serum-free medium-199, and then transplanted into the hindlimb muscles of a subgroup of mice as described above. For another cell-tracking study, a subgroup of mice received an intracardiac injection of FITC-conjugated Bandeiraea simplicifolia lectin I (BS-1; Sigma) and rhodamine-conjugated UEA-1 (Sigma) simultaneously or of FITC-conjugated UEA-1 (Sigma) only at 30 minutes before euthanasia.29
Histological Analysis
Tissue samples harvested from the hindlimb muscles were snap-frozen with precooled isopentane in liquid nitrogen, after which they were embedded in OCT compound (Miles) and cut into 5- to 7-μm-thick sections. For evaluation of capillary density, serving as an anatomic index of angiogenesis, capillary ECs were detected by staining tissue sections for alkaline phosphatase (AP) with the use of the iodoxyl tetrazolium method28 and counterstaining with eosin. Some additional sections were stained with anti-mouse CD31 antibody to further confirm the EC phenotype. The capillary ECs were then counted under light microscopy (x100) to determine capillary density. In each animal, 6 different fields of a section were randomly selected for EC counts, which were then averaged. Tissue sections obtained from hindlimb muscles transplanted with PKH26-RED–labeled HUVECs were also stained for AP to detect viable ECs immediately after being photographed with a CCD camera (Olympus). Tissue sections obtained from hindlimb muscles after premortem lectin staining were observed with a computer-assisted microscope (Nikon) (x400), and UEA-1–positive HUVECs were counted (x200) to quantify the incorporation of HUVECs into the murine vasculature. For each animal, 3 different fields of a section were randomly selected for cell counting. Tissue sections from hindlimb muscles treated only with UEA-1 were stained with anti-human vWF to further confirm the incorporation of HUVECs into the endothelial layer.
To evaluate the percentage of the SeV-transfected population in HUVECs incorporated into the ischemic limbs, we double-immunostained the limb section with anti-human CD31 and anti-SeV antibodies. The numbers of human CD31- and/or SeV-positive cells were counted for the sections randomly obtained from 3 different animals.
To evaluate the proliferative state of the transplanted HUVECs, we double-immunostained the limb section with an anti-human Ki67 antibody and UEA-1. The numbers of human Ki67- and/or UEA-1–positive cells were counted in the sections randomly obtained from 2 different animals.
Statistical Analysis
Each assay experiment was performed at least 3 times. Data are expressed as mean±SD. Student t test was used to compare continuous values between 2 groups. Differences in limb outcome between 2 groups were analyzed by the 2 test. Values of P<0.05 were considered to represent statistically significant differences.
Results
SeV-Mediated Id1 Gene Expression and Protein Synthesis
SeV-mediated Id1 gene expression and protein synthesis in HUVECs were examined by RT-PCR, Western blot, and immunocytochemical analyses. Immunostaining for SeV confirmed that &70% of HUVECs had been successfully infected with SeV within 2 days (data not shown). RT-PCR confirmed the presence of SeV-mediated Id1 mRNA in HUVECs infected with SeV/Id1 (SeV/Id1-HUVECs) 2 days after transfection but not in HUVECs transfected with SeV/Null (SeV/Null-HUVECs) (Figure 1A). Western blot analysis with anti-Id1 antibody detected a 14-kDa protein in the cell lysates of SeV/Id1-HUVECs (Figure 1B). In SeV/Id1-HUVECs, Id1 was predominantly localized to the nucleus (Figure 1C).
Id1 Gene Transfer Confers Angiogenic Activity on HUVECs
To examine the angiogenic effect of Id1 gene transfer, scratch wound assays were performed on HUVECs under serum-reduced and growth factor–free conditions. Id1 increased HUVEC migration up to 144% of the control level after 24 hours (P<0.05) (Figure 2A), as previously reported in other types of ECs.19,30 Id1 gene transfer also stimulated cell growth irrespective of the presence of VEGF as estimated by BrdU incorporation (Figure 2B).
In the Matrigel tube formation assay, SeV/Id1-HUVECs formed robust elongated tube/cordlike structures (Figure 2C). Quantitative analysis showed total tube/cord lengths to be significantly greater in the SeV/Id1-HUVECs than in the SeV/Null-HUVECs (P<0.01) (Figure 2D). These findings clearly suggest that Id1 gene transfer confers angiogenic activity on HUVECs in vitro.
Id1 Gene Transfer Protects Against HUVEC Apoptosis
Previous reports have suggested that Id1 can both positively and negatively regulate apoptosis depending on cell types.31,32 To examine the effects of Id1 on EC apoptosis, we cultured HUVECs in serum-free medium in the presence or absence of SeV/Id1 and evaluated apoptotic cells using flow cytometry. Id1 gene transfer reduced the number of annexin V–positive cells to 47% of the control level (Figure 3A). TUNEL staining also showed that Id1 significantly reduced the number of TUNEL-positive cells down to 40% of the control number (Figure 3B). Representative images of HUVECs with TUNEL staining are shown in Figure 3C. These results show clearly that Id1 can protect HUVECs against apoptosis in vitro.
Transplantation of Id1-Transfected HUVECs Augments Revascularization of Murine Ischemic Hindlimbs
We assessed the angiogenic potency of Id1-transfected HUVECs for therapeutic purposes by transplanting them into murine ischemic hindlimbs. One day after surgical induction of ischemia, 1x105 HUVECs were directly transplanted into the hindlimb muscles of the athymic mice. Blood perfusion recovery after transplantation was then serially assessed with LDPI. Ischemia/normal limb perfusion ratios before surgery were similar in the treatment groups: medium (n=14), SeV/Null-HUVECs (n=11), and SeV/Id1-HUVECs (n=14) (Figure 4A). Immediately after surgery, limb perfusion was severely reduced in all groups to the same extent (Figure 4A), indicating the ischemia severities to be comparable. Over the next 21 days, LDPI revealed gradual recovery of blood perfusion in all 3 groups. Remarkably, recovery was accelerated in the SeV/Id1-HUVECs group. Ischemia/normal limb perfusion ratios on postoperative days 14 and 21 (0.71±0.25 and 0.71±0.12, respectively) were significantly higher in transplanted hindlimbs than in those of the medium-only (0.46±0.18, P<0.01 and 0.48±0.20, P<0.01, respectively) and SeV/Null-HUVECs (0.52±0.20, P<0.05 and 0.54±0.13, P<0.01, respectively) groups (Figure 4A). Recovery of blood perfusion was similar in the SeV/Null-HUVECs and medium-only groups. We also examined increases in transplanted cell numbers up to 5x105 cells and obtained similar results (data not shown). Representative images of hindlimb blood perfusion recorded 21 days after surgical induction of ischemia are shown in Figure 4B.
We then examined whether the recovery of blood flow in this ischemic hindlimb model had resulted from neovascularization. Figure 4C represents photomicrographs of representative ischemic limb sections obtained on day 21. Consistent with the LDPI analysis, immunohistochemical staining for AP revealed larger numbers of capillary ECs in the hindlimbs of the SeV/Id1-HUVECs group (80.5±12.8 number per field) than in the medium-only (47.2±10.5, P<0.01) and in the SeV/Null-HUVECs (43.2±10.5, P<0.05) groups, but there was no significant difference between the latter 2 (Figure 4D).
Increased neovascularization in the mouse ischemic hindlimb can potentially lead to important biological consequences, including limb salvage. To examine limb outcome after induction of ischemia, the appearance of the ischemic hindlimb was observed on the last day of the study period (day 21 after surgery). Among 14 mice in the medium-only group, 7 (50.0%) showed toe necrosis, and 1 (7.1%) showed severe limb necrosis, including autoamputation (Figure 4E). Only 6 (42.9%) were devoid of necrotic lesions (Figure 4E). In contrast, transplantation of SeV/Id1-HUVECs significantly increased the rate of complete limb salvage up to 92% (Figure 4E). Transplantation of SeV/Null-HUVECs did not improve limb outcomes (Figure 4E). Taken together, these findings indicate that transplantation of SeV/Id1-HUVECs, but not of SeV/Null-HUVECs, can rescue ischemic lesions and thereby prevent catastrophic outcomes by augmenting neovascularization.
Evidence of Incorporation of Transplanted HUVECs Into Murine Hindlimb Vasculature
To determine whether transplanted HUVECs contribute to vascular structure regeneration, we labeled HUVECs with PKH26-RED before transplantation and examined their incorporation into capillaries on day 21. After inspection of tissue sections harvested from the ischemic limb for fluorescent-labeled HUVECs, the same sections were stained for AP to detect viable HUVECs. HUVECs double positive for PKH26-RED and AP were documented in the sites of hindlimb capillaries in SeV/Id1-HUVECs, showing incorporation of transplanted HUVECs into the murine hindlimb vasculature (Figure 5A, 5B). To further confirm and quantify these observations, HUVECs and murine ECs were identified with the use of premortem staining by intracardiac injection of fluorescent conjugated lectins (UEA-1 specific for human ECs and BS-1 specific for murine ECs). Stereomicroscopic examination of lectin-stained muscle blocks revealed numerous incorporated HUVECs in the hindlimb vasculature of the SeV/Id1-HUVECs group (Figure 5C to 5E). By contrast, only a few HUVECs were detected in the hindlimb vasculature in the SeV/Null-HUVECs group (Figure 5F). In addition, tissue sections were quantitatively analyzed for incorporated HUVECs. Representative photomicrographs of sections from both groups are presented in Figure 5G, 5H. The number of incorporated HUVECs in the SeV/Id1-HUVECs group was &5 times that of the SeV/Null-HUVECs group (Figure 5I). At higher magnification, the capillary wall proved to be partly composed of UEA-1–positive HUVECs (Figure 5J, 5K). In addition, UEA-1–positive HUVECs resided in the endothelial layer positive for vWF (Figure 5L, 5M). These results confirm that Id1-transfected HUVECs can be effectively incorporated into regenerating capillaries in ischemic lesions.
To evaluate whether Id1-transfected HUVECs were incorporated into the ischemic limbs, we double-immunostained the section obtained from 14 days after HUVEC transplantation with anti-human CD31 and anti-SeV antibodies. Cells positive for both human CD31 and SeV were regarded as SeV/Id1-transfected HUVECs. Among 243 HUVECs positive for human CD31, 211 cells (87%) were positive for SeV, indicating that the majority of the incorporated HUVECs overexpressed Id1. A representative photograph of human CD31- and SeV-positive cells is shown in Figure 5N to 5P.
To address the mechanism underlying the in vivo effects of Id1, we evaluated whether transplanted HUVECs are in a proliferative state by double-immunostaining the section obtained from hindlimb muscles 7 days after HUVEC transplantation with an anti-human Ki67 antibody and UEA-1. We observed that 6.5% of UEA-1–positive HUVECs were Ki67 positive in the section from the SeV/Id1-HUVEC group, whereas we did not detect any Ki67-positive HUVECs in the section from the SeV/Null-HUVEC group. A representative photograph of Ki67- and UEA-1–positive cells is shown in Figure 5Q to 5S.
Ang1 Is Upregulated and Functionally Contributes to In Vitro Angiogenesis on Id1-Transfected HUVECs
To address the mechanism of the angiogenic potencies in SeV/Id1-HUVECs, the gene expressions of several angiogenic factors and their receptors were assessed on HUVECs 2 days after Id1 gene transfer. Semiquantitative RT-PCR analysis showed the Ang1 gene, but not the VEGF, Ang2, KDR, Flt-1, or TIE2 gene, to be upregulated on SeV/Id1-HUVECs (Figure 6A). Western blot analysis revealed increased levels of Ang1 protein in the cell lysates of SeV/Id1-HUVECs (Figure 6B). In addition, coimmunostaining with anti-Ang1 and anti-CD31 antibodies detected Ang1 in the cytoplasm of SeV/Id1-HUVECs (Figure 6C). Staining for SeV/Ang1-transfected HUVECs served as a positive control (Figure 6D).
To evaluate whether upregulated Ang1 functionally contributes to the angiogenic property of SeV/Id1-HUVECs, we examined the effect of Tie2/Fc in a Matrigel tube formation assay. In Matrigel, the formation of elongated tube/cordlike structures in SeV/Id1-HUVECs was suppressed by Tie2/Fc (Figure 6E to 6H). Quantitative analysis showed total tube/cord lengths to be significantly smaller in SeV/Id1-HUVECs treated with Tie2/Fc than in those treated with IgG Fc (P<0.05) (Figure 6I). Total tube/cord length remained significantly greater in SeV/Id1-HUVECs treated with Tie2/Fc than in SeV/Null-HUVECs even if the used dosage of Tie2/Fc was increased (P<0.05) (Figure 6I). We next evaluated whether conditioned media from SeV/Id1-HUVECs (SeV/Id1-CM) also stimulate tube/cord formation of HUVECs in Matrigel. Total tube/cord lengths of HUVECs on Matrigel were significantly greater in the HUVECs treated with SeV/Id1-CM than in those with SeV/Null-CM (P<0.01) (Figure 6J); furthermore, the effects of SeV/Id1-CM on tube/cord formation of HUVECs were again partially suppressed by the presence of Tie2/Fc. A previous study reported that not only Ang1 but also Ang2 might activate Tie2 receptor.33 Because Ang1 but not Ang2 was upregulated in SeV/Id1-HUVECs in the present study, the inhibitory effects of tube/cord formation by Tie2/Fc were considered due to the suppression of Ang1/Tie2 signaling. Collectively, these results suggest that upregulated Ang1 functionally contributes to the angiogenic effects of Id1 and that additional angiogenic factor(s) may also be upregulated in SeV/Id1-HUVECs.
Discussion
In the present study we have demonstrated that gene transfer of Id1, a member of the HLH protein family, confers an angiogenic property on HUVECs in vitro and in vivo. Id1 gene transfer enhanced migration, proliferation, and capillary-like tube/cord formation of HUVECs. Id1 gene transfer also protected HUVECs from serum deprivation–induced apoptosis. Transplantation of SeV/Id1-HUVECs accelerated vascular regeneration in the murine ischemic limb and consequently led to limb salvage. Consistently, transplanted SeV/Id1-HUVECs were effectively incorporated into the regenerating vasculature. Finally, we documented upregulation of Ang1 in SeV/Id1-HUVECs, which functionally contributed to enhanced in vitro angiogenesis.
In previous studies,29,34,35 the angiogenic potency of EPC was investigated in hindlimb ischemia with the use of a similar mouse model, in which systemic transplantation of ex vivo expanded EPCs dramatically accelerated blood flow recovery and increased capillary densities in the ischemic limb. Beneficial effects of local transplantation of EPCs have also been shown in other animal models.36,37 Kim and colleagues11 transplanted ECs isolated form the aorta directly into the scar sites of rat hearts 2 weeks after cryoinjury. Transplanted ECs were incorporated into the new vessels and increased regional perfusion in myocardial scar tissue but failed to improve global function. Chekanov and colleagues38 administered autologous ECs into the ischemic myocardium of sheep using a fibrin matrix and demonstrated enhancement of neovascularization with improved left ventricular function. In tumor angiogenesis models,39,40 implanted mature ECs similarly contributed to host neovascularization and tumor growth; furthermore, genetic modification of the ECs with the antiapoptotic gene Bcl-2 enhanced the effects of cell implantation. These findings suggest that mature ECs have the potential to be incorporated into regenerating vessels in vivo under certain conditions.
The present results demonstrate that mature human ECs overexpressing the Id1 gene can be effectively incorporated into regenerating vessels in murine ischemic limbs. The extents of angiogenic and limb salvage effects were comparable to those of EPCs.29,34,35 These results indicate that mature ECs could potentially be optimized for therapeutic angiogenesis by inducing phenotypic changes through appropriate gene transfers. Id1 gene transfer stimulated migration, proliferation, and tube formation of HUVECs in vitro, as previously shown in bovine aortic ECs19 and HUVECs.41 Migration and proliferation of ECs is considered a critical step for physiological as well as therapeutic angiogenesis.5,34,35 In the present analysis, a part of transplanted SeV/Id1-HUVECs but not SeV/Null-HUVECs were in a proliferative state, indicating that stimulation of proliferation may contribute to the in vivo angiogenic effects of Id1. In addition, Id1 inhibited apoptosis of HUVECs under serum deprivation conditions. Id1 has been reported to both positively and negatively regulate apoptosis, depending on cell types and experimental conditions.31,32,41 Sakurai et al41 have reported that overexpression of Id3, but not Id1, inhibits HUVEC apoptosis induced by serum starvation. The discrepancy between their results and ours may be explained by differences in culture condition and/or in Id1 expression levels. A previous study showed Id1 to delay the onset of replicative senescence in human ECs42; Id1 may thereby increase the survival of SeV/Id1-HUVECs and contribute to angiogenesis.
In the present study, Id1 gene transfer upregulated the expression of Ang1 but not Ang2 in HUVECs. Furthermore, the present study revealed that upregulated Ang1 functionally contributed to the in vitro angiogenic effects of Id1 as an autocrine/paracrine factor. Ang1 is reported to augment EC migration and protect ECs against apoptosis partly via the activation of Tie2/PI3'-kinase/Akt signal transduction pathway.26,43,44 Upregulation of the signaling pathway may partly explain an increase in the survival of SeV/Id1-HUVECs. Although it remains unknown whether the effect of Id1 on Ang1 expression is direct, Ang1 upregulation may contribute to the mechanism underlying the in vivo angiogenic effect of Id1-transfected HUVECs, together with other possible contributions of metalloproteinases, integrins, and fibroblast growth factor receptor genes, as suggested by previous studies using Id gene–deleted mice15,18 and Id1-transfected ECs.41
We analyzed endogenous Id1 expression levels with immunohistochemical technique in the tissue section of murine ischemic limbs. However, we could not detect significant upregulation of Id1 in ECs (K. Nishiyama, MD, et al, unpublished data, 2003). In addition, our preliminary experiments showed that deferoxamine, an agent mimicking hypoxic condition, downregulated Id1 expression in cultured HUVECs, although it caused VEGF upregulation (K. Nishiyama, MD, et al, unpublished data, 2004), suggesting that Id1 expression is not induced in ischemic lesion. This result suggests that ischemia-induced suppression of Id1 expression may overcome the inductive effect of VEGF on Id1 expression. Therefore, Id1 gene transfer is expected to complement the downregulated activity of Id1 to confer angiogenic capacity on ECs.
Contrary to the beneficial angiogenic effect, it has been reported that the Id1 transgene may induce neoplastic transformation of ECs.13,45 In a previous study,42 however, Id1-transferred ECs were not immortalized and eventually underwent senescence despite high Id1 levels. In addition, no tumor formation was pathologically observed in the lesions with transplanted SeV/Id1-HUVECs in our present study (data not shown). In addition, the ex vivo gene transfer strategy may preclude recipients from being exposed to the Id1 gene–bearing vector. Thus, the combination of Id1 gene transfer and transplantation using mature ECs may serve as a novel and useful application for therapeutic revascularization with high efficiency and low risk.
Acknowledgments
This work was supported in part by grants from the Japan Society for the Promotion of Science Research for the Future Program; grants-in-aid for scientific research from the Ministry of Education, Culture, Sports, Science, and Technology, Japan; and a research grant for cardiovascular diseases (14C-1 and 14C-4) from the Ministry of Health, Labor, and Welfare.
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