Differentiation of canine adipose tissue–derived mesenchymal stem cells towards endothelial progenitor cells Byung-Jae Kang, DVM, PhD; Seung Hoon Lee, DVM; Oh-Kyeong Kweon, DVM, PhD; Je-Yoel Cho, DVM, PhD
Objective—To determine the differentiation of canine adipose tissue–derived mesenchymal stem cells (ASCs) into endothelial progenitor cells (EPCs). Animals—3 healthy adult Beagles. Procedures—Canine ASCs were isolated and cultured from adipose tissue, and endothelial differentiation was induced by culturing ASCs in differentiation medium. Morphological and immunophenotypic changes were monitored. Expression of endothelial-specific markers was analyzed by conventional and real-time RT-PCR assay. The in vitro and in vivo functional characteristics of the endothelial-like cells induced from canine ASCs were evaluated by use of an in vitro solubilized basement membrane tube assay, low-density lipoprotein uptake assay, and in vivo solubilized basement membrane plug assay. Results—After differentiation culture, the cells developed morphological changes, expressed EPC markers such as CD34 and vascular endothelial growth factor receptor 2, and revealed functional characteristics in vitro. Furthermore, the induced cells allowed vessel formation in solubilized basement membrane plugs in vivo. Conclusion and Clinical Relevance—Results indicated that canine ASCs developed EPC characteristics when stimulated by differentiation medium with growth factors. Thus, differentiated canine ASC-EPCs may have the potential to provide vascularization for constructs used in regenerative medicine strategies. (Am J Vet Res 2014;75:685–691)
T
issue engineering is the use of a combination of cells, scaffolds, and growth factors to restore, maintain, and improve biological tissue function.1 Recently, various applications of tissue engineering have been reported in veterinary medicine.2,3 One of the main difficulties in tissue engineering is that the vascularization of the engineered tissues is inevitable. Previous studies4,5 reveal that vascularization in engineered tissues with mature endothelial cells improves blood perfusion, cell viability, and survival of the tissues after transplantation. Furthermore, neovascularization is a critical initial step for functional rehabilitation and wound healing. However, mature endothelial cells have limited capacity to vascularize damaged tissues because these cells have low proliferation potential. Thus, it is necessary to find and investigate alternative sources of these cells to regenerate injured tissues. Endothelial proReceived January 6, 2014. Accepted March 10, 2014. From the Departments of Veterinary Biochemistry, BK21Plus Program and Research Institute for Veterinary Science (Kang, Cho) and Veterinary Surgery (Kang, Lee, Kweon), College of Veterinary Medicine, Seoul National University, Seoul 151-742, Republic of Korea. Supported by the National Research Foundation (NRF), Ministry of Science, ICT & Future Planning (2012M3A9C6049716 and 20110019355). The authors thank Dr. Hyung-Sik Kim for technical assistance with fluorescent-activated cell sorting. Address correspondence to Dr. Cho ([email protected]). AJVR, Vol 75, No. 7, July 2014
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ASC EGM-2 FACS EPC FBS LDL MSC VEGF VEGFR2 vWF
ABBREVIATIONS
Adipose tissue–derived mesenchymal stem cell Endothelial growth medium-2 Fluorescence-activated cell sorting Endothelial progenitor cell Fetal bovine serum Low-density lipoprotein Mesenchymal stem cell Vascular endothelial growth factor Vascular endothelial growth factor receptor 2 Von Willebrand factor
genitor cells are immature cells with some self-renewal capacity and can differentiate into mature endothelial cells in vitro and in vivo.6,7 Results of several studies6,8 suggest the effectiveness of EPCs for re-endothelialization and neovascularization in humans and other animals. However, EPCs do not provide sufficient numbers of cells for therapeutic application owing to loss of their proliferation potential after extended expansion.9 Recently, several studies10–15 have found that MSCs derived from various tissues, including bone marrow, neonatal tissue, and adipose tissue, are able to differentiate into endothelial lineages in vitro. In particular, ASCs have received extensive attention as a possible 685
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candidate for tissue regeneration because these cells can be easily obtained in high yield with minimal cell death.14,16 Although mouse, rat, and human ASCs have already been reported to differentiate into endothelial lineages14,17–19 and the cell thrombogenicity of canine autologous ASCs seeded onto vascular grafts in experimental canine studies has been evaluated,19 in vitro differentiation of canine ASCs into endothelial-like cells has never been described to our knowledge. The clinical information obtained from dogs may be useful for studying new therapies for human disease such as cardiovascular or ischemic disease, compared with rodents, because canine studies provide research data that may be directly transferable to human medicine.20,21 In vitro differentiation of canine ASCs into endothelial lineages will have the potential to treat various tissue injuries in dogs and will provide the opportunity to evaluate efficacy and safety of cell transplantation for the treatment of human cardiovascular disease. On the basis of findings in studies of human and rodent ASCs, we hypothesized that canine ASCs can differentiate into endothelial lineages. Therefore, the purpose of the study reported here was to evaluate whether canine ASCs differentiate into endothelial lineages, as determined by use of morphological, immunophenotypic, RT-PCR assay, and functional analyses. Materials and Methods Isolation and cultivation of canine ASCs—Canine ASCs were obtained according to the methods described in previous articles.16,22 Briefly, adipose tissues were aseptically collected from the gluteal region of the dogs during general anesthesia. This procedure was conducted with the approval of the Institutional Animal Care and Use Committee of Seoul National University (SNU-120306-5). The adipose tissues were extensively washed with PBS solution and then minced with scissors. The minced tissues were digested with collagenase type Ia (1 mg/mL) for 2 hours at 37°C. The tissue samples were washed with PBS solution and then centrifuged at 300 X g for 10 minutes. The resulting pellet of stromal vascular fraction was resuspended, filtered through a 100-µm nylon mesh, and incubated overnight in mediumb with 10% FBSc at 37C in a 5% CO2 humidified atmosphere. After 24 hours, the unattached cells and residual nonadherent RBCs were removed by washing with PBS solution. The medium was changed at 48-hour intervals until the cells became confluent. After the cells reached 90% confluence, they were subcultured. At passage 2, the cells were used for the following experiments. In vitro differentiation into endothelial lineage—To induce endothelial differentiation in vitro, canine ASCs at passage 2 were cultured in a commercially available EGM-2d on 6-well culture plates at a density of 3,000 cells/ cm2. Endothelial growth media-2 con686
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tained epidermal growth factor, hydrocortisone, antimicrobials (gentamicin and amphotericin-B), basic fibroblast growth factor, VEGF, insulin-like growth factor, ascorbic acid, and heparin with 2% FBS at the manufacturer’s concentrations. Medium was changed every 2 to 3 days, and the degree of differentiation was assessed after 7 and 14 days. Canine ASCs cultured in medium with 10% FBS were used as a control. Morphology—Morphological changes of the cells were observed and captured by use of a live-cell movie analyzere after 3, 7, and 14 days of differentiation. The areas and perimeters of the cells were measured at each time point with software,f and the circularity index was calculated as (4π X area/[perimeter]2), as reported23; 3 frames were captured, and the areas and perimeters of 5 cells/frame were measured for calculating the circularity index. Flow cytometric analysis—Trypsinized cells were suspended in PBS solution at a concentration of 5 X 105 cells/30 µL. The cells were stained with fluorescein isothiocyanate–conjugated antibodies specific for CD31g and CD34h at 20°C for 1 hour. Expressions of the corresponding cell surface markers were evaluated with an FACS flow cytometeri by use of software.j RT-PCR and real-time PCR assays—Total RNAs were isolated from canine ASCs, ASC-EPCs, and cephalic vein tissue (positive control) by means of a chemical solutionk under conditions recommended by the manufacturer. Extracted RNA was dissolved in diethypyrocarbonate-treated water, and cDNAs were synthesized from 2 µg of total RNA for the cells and 1 µg of total RNA for the positive control with a reverse transcription kit.l Then, the PCR assay was performed by use of specific primers for canine CD31, vWF, VEGFR2, and glyceraldehyde-3-phosphate dehydrogenase. Primers for PCR were follows: CD31 (forward, 5′-GCACACAAGAGGCATGGTAAC-3′; reverse,
Figure 1—Phase contrast microscopic views of undifferentiated canine ASCs (A) and ASC-EPCs differentiated in EGM-2 for 3 days (B), 1 week (C), and 2 weeks (D). Bar = 250 µm. AJVR, Vol 75, No. 7, July 2014
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5′-GAATGGAGCACCACAGGTTT-3′), vWF (forward, 5′-GCAATGTCTCCTCTGATGAAG-3′; reverse, 5′-GTACAAGACAACCCCCTGCT-3′), VEGFR2 (forward, 5′-GGTATGGTCCTTGCCTCAGA-3′; reverse, 5′-CAGTGGTATCCGTGTCATCG-3′), and glyceraldehyde-3-phosphate dehydrogenase (forward, 5′-CATTGCCCTCAATGACCACT-3′; reverse, 5′-TCCTTGGAGGCCATGTAGAC-3′). Twenty-five microliters of PCR product was prepared with 2 µL of cDNA, 10 pmol of each primer, 1.25 units of Taq polymerase,m PCR buffer, MgCl2, and a final concentration of 0.2mM dNTPs. Cycle conditions for conventional and real time PCR assays were as follows: conventional PCR assay (95°C for 3 minutes, 35 cycles at 95°C for 30 seconds, 53.5°C for 30 seconds, 72°C for 1.5 minutes, and 72°C for 5 minutes), real time PCR assay (95°C for 2 minutes, 35 cycles at 95°C for 10 seconds, 53.5°C for 30 seconds, 72°C for 30 seconds, and 72°C for 5 minutes). The con-
ventional PCR assay products were analyzed via 1.5% agarose gel electrophoresis and visualized with ethidium bromide. Real-time PCR reaction for VEGFR2 was performed with nucleic acid gel stainn and a real-time PCR detection system.o In vitro solubilized basement membrane tube assay—Solubilized basement membrane preparationp was allowed to thaw at 4°C overnight and then added to 24well culture plates at a concentration of 50 µL/cm2. The coated plates were then incubated at 37°C for 45 minutes to allow for solidification. The ASC-EPCs cultured for 14 days in EGM-2 and undifferentiated ASCs were dissociated with trypsin, reseeded at 2.5 X 104 cells/cm2 on the solubilized basement membrane substrate, and incubated at 37°C for 24 hours. After 24 hours incubation, tube-like network formation was examined and recorded with a phase contrast microscope. LDL uptake assay—The LDL uptake assay was performed with an assay kit,q according to the manufacturer’s instructions. The ASC-EPCs were cultured for 14 days in EGM-2, and undifferentiated ASCs were harvested with trypsin, replated in 48-well culture plates at 2.5 X 104 cells/cm2, and incubated at 37°C for 4 hours in EGM-2 containing LDL working solution. At the end of the LDL uptake incubation, the culture medium was aspirated and replaced with PBS solution. The degree of LDL uptake was determined with a fluorescent microscope.
Figure 2—Quantitative flow cytometric analysis expression of CD31 and CD34 by undifferentiated canine ASCs (A) and ASC-EPCs differentiated in EGM-2 for 1 week (B) and 2 weeks (C). The colored peak indicates the distribution of cells stained with fluorescein isothiocyanate-conjugated antibodies, and the black peak represents a control. The percentage values indicate percentage increase versus controls. FL1-H = Fluorescence intensity of fluorescein isothiocyanate–conjugated antibodies. AJVR, Vol 75, No. 7, July 2014
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In vivo solubilized basement membrane plug assay—Two healthy Beagles were used for subcutaneous implantation of ASC-EPCs and undifferentiated ASCs. All animal procedures were performed in accordance with the guidelines of the Institutional Animal Care and Use Committee of Seoul National University (SNU-131212-1-1). Anesthesia was induced by IV administration of tiletamine hydrochlorider at a dose of 7.5 mg/kg and maintained with isofluranes in oxygen. Intravenously administered tramadolt at a dose of 2 mg/kg was used as an analgesic. Solubilized basement membrane preparation (500 µL) was used to trap 2.5 X 105 ASC-EPCs cultured for 14 days in EGM-2 and 2.5 X 105 undifferentiated ASCs before SC injection. Cell-free plugs of the same size were used as negative controls, and plugs containing recombinant human VEGFu were used as positive controls. After 7 days, plugs were harvested for examination of vessel formation. The sample tissues were fixed in 4% paraformaldehyde for 1 day, dehydrated by use of a graded alcohol series, and embedded in paraffin. Five-micrometerthick slices were mounted on glass slides. 687
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The sections were deparaffinized, hydrated, and stained with H&E. The stained slides were observed via light microscopy to examine vessel formation.
Results
Morphological and immunophenotypic changes—Endothelial differentiation was induced by culStatistical analysis—Data are expressed as mean turing canine ASCs in EGM-2 media. Undifferenti± SD. Statistical analysis was performed with software.v ated ASCs had typical mesenchymal morphology, with The student t test was performed to assess differences a fibroblast-like shape (Figure 1). In contrast, after 3 between 2 groups. A value of P < 0.05 was considered days of culture in an endothelial differentiation medito be significant. um, these cells became less elongated, compared with undifferentiated ASCs. After 1 week of differentiation culture, the cells changed into a more polygonal shape. After 2 weeks of culture, ASC-EPCs acquired a cobblestone-like morphological appearance typical for endothelial lineage. On the basis of a circularity index between 0 and 1 (the circularity index of 1 represents round objects, and the lower the value is, the less round is the object), the circularity index of ASC-EPCs was significantly (P = 0.01) higher after 2-week differentiation, compared with undifferentiated ASCs (n = 15 frames; 0.79 ± 0.10 vs 0.13 ± 0.02). Quantitative flow cytometric analysis of CD31 and CD34 expression was performed to confirm the immunophenotypic changes that occurred under the endothelial differentiation condition. In general, CD31 characterizes mature endothelial cells, and CD34 is used as an EPC marker. Undifferentiated ASCs had no specific expression for CD31 and CD34 (Figure 2). The ASC-EPCs had increased CD34 expression after 1 and 2 weeks of differentiation. The expression Figure 3—Agarose gel electrophoretogram resulting from RT-PCR assay of endothe- of mature endothelial marker CD31 was lial-specific genes (CD31, vWF, and VEGFR2) and glyceraldehyde-3-phosphate dehydrogenase (GAPDH) of undifferentiated canine ASCs and ASC-EPCs differentiated in also mildly increased at the first week, EGM-2 for 1 week and 2 weeks. although it was decreased at 2 weeks of differentiation. However, changes of CD31 expression were not significant, compared with those of CD34. The immunophenotypic changes suggested that alteration of these cells is possible from mesenchymal to endothelial lineage, especially endothelial progenitor type, under endothelial differentiation condition.
Figure 4—Phase contrast microscopic views of capillary network formation by undifferentiated canine ASCs (A) and differentiated ASC-EPCs (B), and fluorescence imaging of LDL uptake by ASCs (C) and ASC-EPCs (D). Notice that network formation and LDL uptake are observed only in ASC-EPCs. Bar = 250 mm. 688
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EPC and EC markers in cells differentiated from canine ASCs—The expression of endothelial specific markers CD31, vWF, and VEGFR2 was analyzed by conventional RT-PCR assay (Figure 3). Positive controls (vessel tissues) expressed the 3 markers. Undifferentiated and differentiated ASCs did not express CD31 and vWF, whereas these cells expressed VEGFR2. The expression of VEGFR2 was also observed in undifferentiated ASCs but was lower than that in differentiated ASC-EPCs. Real-time PCR assay was performed to quantify the mRNA expression levels of VEGFR2 AJVR, Vol 75, No. 7, July 2014
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tional characteristics of endothelial lineage in vitro and in vivo. They formed tubular networks on solubilized basement membrane substrate, integrated LDL in vitro culture, and contributed to vessel-like structures in vivo. Analysis of some surface markers and gene expressions was also performed to assess endothelial differentiation. The differentiated cells had increased expression of CD34 and VEGFR2 after 1 and 2 weeks of endothelial differentiation. Interestingly, however, increased mRNA expressions of CD31 and vWF were not observed after induction, but a small increase of the surface CD31 protein was detected by use of FACS. Expression of CD31 was slightly increased after the first week of endothelial differentiation but was again decreased after the second week of induction. Although this finding Figure 5—Photomicrographic views of blood vessel formation by canine cells in solubilized basement membrane plugs containing no cells (A), VEGF (B), ASCs (C), and of FACS analysis somewhat suggested ASC-EPCs (D). Arrows indicate blood vessels. Notice that little or no blood vessel CD31 expression, this expression was formation is observed in cell-free plugs. H&E stain; bar = 50 mm. less than that of CD34. Therefore, we concluded that canine ASCs expressed during the endothelial differentiation process of the cano or little CD31 after endothelial differentiation by nine ASCs. Quantitative real-time RT-PCR analysis reuse of EGM-2 alone. Similarly, several previous studvealed a consistent increase with time in the transcripies11,12,15,24 found that the cells expressed no or little tion of VEGFR2 during ASC differentiation in EGM-2. CD31 after endothelial differentiation. However, in The expression of VEGFR2 in 1-week differentiated other studies,14,17 the CD31 expression level had apparcells increased approximately 4-fold (P < 0.05), and the ently increased after endothelial differentiation. CD31 expression in 2-week samples increased approximateis a mature endothelial differentiation marker; thus, ly 10-fold (P < 0.05), compared with undifferentiated elongation of differentiation time may lead to an upcells. regulation of CD31.12,15 In addition, the differentiation condition used in the present study may not fully lead Functional characteristics of ASC-EPCs—Tube to mature endothelial differentiation of canine ASCs formation was assessed by seeding undifferentiated and could maintain the endothelial progenitor potenASCs and ASC-EPCs differentiated in EGM-2 on solutial. Indeed, in this study, canine ASCs also maintained bilized basement membrane preparation. Undifferenproliferation potential during the endothelial differentiated cells developed little or no tube-like network tiation culture, although these cells had morphological formation (Figure 4). In contrast, differentiated cells and immunophenotypic changes and functional angiodeveloped clear tube-like structures. genesis characteristics such as LDL uptake and tube In vitro functional characterization was also performation. The results of the present study were conformed by assessing LDL uptake. Undifferentiated sistent with previous research on acquisition of endoASCs had little or no LDL uptake (Figure 4), whereas thelial cell characteristics by human ASCs.19 The preASC-EPCs after cultivation in EGM-2 for 2 weeks had vious study used endothelial cell growth supplement high LDL uptake. as the main stimulus of endothelial differentiation, but The solubilized basement membrane plug assay this supplement alone did not promote the expression was performed to investigate whether ASC-EPCs can of molecular markers. However, CD31 expression and promote new blood vessel formation in vivo. One week LDL uptake were elicited after shear stress was applied after implantation, the plugs were harvested and vesas an additional stimulus. Shear stress is defined as the sel formation was observed. Little or no blood vessel frictional force per unit area, and it acts on endothelial formation was observed in cell-free plugs (Figure 5). cells as blood flows throughout the vascular system.10 Vessels were observed in the positive control containRecent evidence suggests that stem cells differentiing VEGF. Smaller but high numbers of vessels were ate into endothelial cells when exposed to fluid shear found in the plugs seeded with ASCs and ASC-EPCs. stress.10,19 Eventually, the effect of EGM-2 alone on the Differences among groups (except the negative control) differentiation of canine ASCs into mature endothelial were not significant. cells appeared limited. Therefore, further investigations should be carefully performed to evaluate whether Discussion mechanotransduction can modulate mature endothelial differentiation of canine ASCs. Results of the present study revealed that canine Generally, EPCs are characterized by 3 surface ASCs were capable of differentiating into endothelialmarkers including CD133, CD34, and VEGFR2.6 In like cells in vitro. The differentiated cells had the funcAJVR, Vol 75, No. 7, July 2014
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the present study, the differentiated ASC-EPCs had increased expression of CD34 and VEGFR2 as well as functional vascular characteristics. Some previous studies25,26 revealed that human ASCs had a low level of expression of VEGFR2 and could gradually differentiate into endothelial-like cells with increased expression of CD34 and VEGFR2. Therefore, we termed endothelial-differentiated ASCs as ASC-EPCs. However, further investigations are needed to determine why some cells were responsive to the differentiation induction and what the conditions are for differentiating ASCs into mature endothelial cells. A previous report10 suggested that a reliable endothelial differentiation protocol for MSCs was not yet established, whereas standardization of other differentiation procedures such as osteogenesis and adipogenesis was uncomplicated and in agreement with the published protocols. Therefore, protocols for differentiation towards EPCs or endothelial cells should be established for improving the clinical application of these cells. Subcutaneous implantation of solubilized basement membrane plugs containing ASCs, ASC-EPCs, or VEGF resulted in the formation of new blood vessels. Blood vessel density was slightly greater in VEGF and ASC-EPC plugs. As already known, VEGF alone in solubilized basement membrane plugs stimulated cells to form blood vessels. From this, it is speculated that ASCs or ASC-EPCs might also stimulate VEGF secretion to induce blood vessel formation in vivo. However, it can be concluded that differentiation of ASCs into ASC-EPCs did not impair but rather increased their ability to induce development of new blood vessels. However, blood vessel density is known to be dependent on the number of implanted cells24 and may be different according to the period of implantation. Therefore, further studies are required to verify whether differentiation of canine ASCs prior to application can further improve the performance of these cells for neovascularization. The EPCs or CD34-positive cells may be a possible therapeutic option for accelerated re-endothelization and neovascularization. However, a substantial problem in the therapeutic use of these cells is the limited number of autologous cells that can be obtained from adult blood. Furthermore, the number of circulating EPCs is reduced in patients with risk factors for ischemic cardiovascular disease.17,27 In the present study, we successfully obtained CD34-positive endothelial progenitor-like cells by differentiation of canine ASCs with EGM-2. The differentiated cells have the advantage that they can be expanded from ASCs, which are easily obtained in large numbers. Therefore, ASC-EPCs could be a useful cell source for cell-based therapies of ischemic disease and vascularization of engineered tissues. Endothelial growth media-2 is used to maintain endothelial cell growth, but has also been used to differentiate MSCs derived from humans or mice into endothelial lineages in many studies.14,23,24,28 Fibroblast growth factor and VEGF included in EGM-2 are known to play a critical role in the induction of endothelial lineages.14 We attempted to determine whether canine ASCs could differentiate into endothelial-like cells in EGM-2 and confirmed that these cells responded to the 690
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induction of growth factors included in EGM-2. To our knowledge, this study revealed for the first time that ASCs derived from dogs are capable of differentiating towards endothelial-like cells. Generally, rodents have been widely used in preclinical studies for human disease, but clinical data obtained from larger animals such as dogs provide more reliable information with regard to human disease.21 Therefore, the establishment of endothelial differentiation of stem cells derived from dogs will provide the opportunity to evaluate the efficacy and safety of cell therapy for human cardiovascular disease such as myocardial infarction and limb ischemia. In addition, canine ASC-EPCs may have the potential for use in treating dogs with various tissue injuries and ischemic diseases. a. b. c. d. e. f. g. h. i. j. k. l. m. n. o. p. q. r. s. t. u. v.
Collagenase type I, Sigma-Aldrich Inc, St Louis, Mo. Dulbecco modified Eagle medium, Gibco, Billings, Mont. FBS, Gibco, Billings, Mont. EGM-2, Lonza, Walkersville, Mich. JuLI, NanoEnTek, Seoul, Korea. ImageJ software, NIH, Bethesda, Md. Rabbit anti-CD31 polyclonal antibody, Bioss, Beijing, China. Mouse anti-dog CD34 monoclonal antibody, Serotec, Oxford, England. FACS Calibur flow cytometer, BD Biosciences, San Jose, Calif. CELL Quest Pro, BD Biosciences, San Jose, Calif. Trizol, Invitrogen Co, Carlsbad, Calif. Omniscript Reverse Transcription Kit, Qiagen Inc, Valencia, Calif. Taq polymerase, Promega, Madison, Wis. SYBR Green I nucleic acid gel stain, Invitrogen Co, Carlsbad, Calif. CFX ConnectT Real-Time System, Bio-Rad Laboratories Inc, Hercules, Calif. Matrigel, BD Biosciences, San Jose, Calif. LDL Uptake Cell-Based Assay Kit, Cayman Chemical Co, Ann Arbor, Mich. Tiletamine hydrochloride, Virbac, Carros, France. Isoflurane, Baxter Healthcare, Deerfield, Ill. Tramadol, Samsung Pharmaceutical, Seoul, Korea. Recombinant human VEGF, R&D systems, Minneapolis, Minn. SPSS, version 20, SPSS Inc, Chicago, Ill.
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Objective—To determine the differentiation of canine adipose tissue–derived mesenchymal stem cells (ASCs) into endothelial progenitor cells (EPCs). Animals—3 healthy adult Beagles. Procedures—Canine ASCs were isolated and cultured from adipose tissue, and endothelial differentiation was induced by culturing ASCs in differentiation medium. Morphological and immunophenotypic changes were monitored. Expression of endothelial-specific markers was analyzed by conventional and real-time RT-PCR assay. The in vitro and in vivo functional characteristics of the endothelial-like cells induced from canine ASCs were evaluated by use of an in vitro solubilized basement membrane tube assay, low-density lipoprotein uptake assay, and in vivo solubilized basement membrane plug assay. Results—After differentiation culture, the cells developed morphological changes, expressed EPC markers such as CD34 and vascular endothelial growth factor receptor 2, and revealed functional characteristics in vitro. Furthermore, the induced cells allowed vessel formation in solubilized basement membrane plugs in vivo. Conclusion and Clinical Relevance—Results indicated that canine ASCs developed EPC characteristics when stimulated by differentiation medium with growth factors. Thus, differentiated canine ASC-EPCs may have the potential to provide vascularization for constructs used in regenerative medicine strategies. (Am J Vet Res 2014;75:685–691)
T
issue engineering is the use of a combination of cells, scaffolds, and growth factors to restore, maintain, and improve biological tissue function.1 Recently, various applications of tissue engineering have been reported in veterinary medicine.2,3 One of the main difficulties in tissue engineering is that the vascularization of the engineered tissues is inevitable. Previous studies4,5 reveal that vascularization in engineered tissues with mature endothelial cells improves blood perfusion, cell viability, and survival of the tissues after transplantation. Furthermore, neovascularization is a critical initial step for functional rehabilitation and wound healing. However, mature endothelial cells have limited capacity to vascularize damaged tissues because these cells have low proliferation potential. Thus, it is necessary to find and investigate alternative sources of these cells to regenerate injured tissues. Endothelial proReceived January 6, 2014. Accepted March 10, 2014. From the Departments of Veterinary Biochemistry, BK21Plus Program and Research Institute for Veterinary Science (Kang, Cho) and Veterinary Surgery (Kang, Lee, Kweon), College of Veterinary Medicine, Seoul National University, Seoul 151-742, Republic of Korea. Supported by the National Research Foundation (NRF), Ministry of Science, ICT & Future Planning (2012M3A9C6049716 and 20110019355). The authors thank Dr. Hyung-Sik Kim for technical assistance with fluorescent-activated cell sorting. Address correspondence to Dr. Cho ([email protected]). AJVR, Vol 75, No. 7, July 2014
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ASC EGM-2 FACS EPC FBS LDL MSC VEGF VEGFR2 vWF
ABBREVIATIONS
Adipose tissue–derived mesenchymal stem cell Endothelial growth medium-2 Fluorescence-activated cell sorting Endothelial progenitor cell Fetal bovine serum Low-density lipoprotein Mesenchymal stem cell Vascular endothelial growth factor Vascular endothelial growth factor receptor 2 Von Willebrand factor
genitor cells are immature cells with some self-renewal capacity and can differentiate into mature endothelial cells in vitro and in vivo.6,7 Results of several studies6,8 suggest the effectiveness of EPCs for re-endothelialization and neovascularization in humans and other animals. However, EPCs do not provide sufficient numbers of cells for therapeutic application owing to loss of their proliferation potential after extended expansion.9 Recently, several studies10–15 have found that MSCs derived from various tissues, including bone marrow, neonatal tissue, and adipose tissue, are able to differentiate into endothelial lineages in vitro. In particular, ASCs have received extensive attention as a possible 685
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candidate for tissue regeneration because these cells can be easily obtained in high yield with minimal cell death.14,16 Although mouse, rat, and human ASCs have already been reported to differentiate into endothelial lineages14,17–19 and the cell thrombogenicity of canine autologous ASCs seeded onto vascular grafts in experimental canine studies has been evaluated,19 in vitro differentiation of canine ASCs into endothelial-like cells has never been described to our knowledge. The clinical information obtained from dogs may be useful for studying new therapies for human disease such as cardiovascular or ischemic disease, compared with rodents, because canine studies provide research data that may be directly transferable to human medicine.20,21 In vitro differentiation of canine ASCs into endothelial lineages will have the potential to treat various tissue injuries in dogs and will provide the opportunity to evaluate efficacy and safety of cell transplantation for the treatment of human cardiovascular disease. On the basis of findings in studies of human and rodent ASCs, we hypothesized that canine ASCs can differentiate into endothelial lineages. Therefore, the purpose of the study reported here was to evaluate whether canine ASCs differentiate into endothelial lineages, as determined by use of morphological, immunophenotypic, RT-PCR assay, and functional analyses. Materials and Methods Isolation and cultivation of canine ASCs—Canine ASCs were obtained according to the methods described in previous articles.16,22 Briefly, adipose tissues were aseptically collected from the gluteal region of the dogs during general anesthesia. This procedure was conducted with the approval of the Institutional Animal Care and Use Committee of Seoul National University (SNU-120306-5). The adipose tissues were extensively washed with PBS solution and then minced with scissors. The minced tissues were digested with collagenase type Ia (1 mg/mL) for 2 hours at 37°C. The tissue samples were washed with PBS solution and then centrifuged at 300 X g for 10 minutes. The resulting pellet of stromal vascular fraction was resuspended, filtered through a 100-µm nylon mesh, and incubated overnight in mediumb with 10% FBSc at 37C in a 5% CO2 humidified atmosphere. After 24 hours, the unattached cells and residual nonadherent RBCs were removed by washing with PBS solution. The medium was changed at 48-hour intervals until the cells became confluent. After the cells reached 90% confluence, they were subcultured. At passage 2, the cells were used for the following experiments. In vitro differentiation into endothelial lineage—To induce endothelial differentiation in vitro, canine ASCs at passage 2 were cultured in a commercially available EGM-2d on 6-well culture plates at a density of 3,000 cells/ cm2. Endothelial growth media-2 con686
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tained epidermal growth factor, hydrocortisone, antimicrobials (gentamicin and amphotericin-B), basic fibroblast growth factor, VEGF, insulin-like growth factor, ascorbic acid, and heparin with 2% FBS at the manufacturer’s concentrations. Medium was changed every 2 to 3 days, and the degree of differentiation was assessed after 7 and 14 days. Canine ASCs cultured in medium with 10% FBS were used as a control. Morphology—Morphological changes of the cells were observed and captured by use of a live-cell movie analyzere after 3, 7, and 14 days of differentiation. The areas and perimeters of the cells were measured at each time point with software,f and the circularity index was calculated as (4π X area/[perimeter]2), as reported23; 3 frames were captured, and the areas and perimeters of 5 cells/frame were measured for calculating the circularity index. Flow cytometric analysis—Trypsinized cells were suspended in PBS solution at a concentration of 5 X 105 cells/30 µL. The cells were stained with fluorescein isothiocyanate–conjugated antibodies specific for CD31g and CD34h at 20°C for 1 hour. Expressions of the corresponding cell surface markers were evaluated with an FACS flow cytometeri by use of software.j RT-PCR and real-time PCR assays—Total RNAs were isolated from canine ASCs, ASC-EPCs, and cephalic vein tissue (positive control) by means of a chemical solutionk under conditions recommended by the manufacturer. Extracted RNA was dissolved in diethypyrocarbonate-treated water, and cDNAs were synthesized from 2 µg of total RNA for the cells and 1 µg of total RNA for the positive control with a reverse transcription kit.l Then, the PCR assay was performed by use of specific primers for canine CD31, vWF, VEGFR2, and glyceraldehyde-3-phosphate dehydrogenase. Primers for PCR were follows: CD31 (forward, 5′-GCACACAAGAGGCATGGTAAC-3′; reverse,
Figure 1—Phase contrast microscopic views of undifferentiated canine ASCs (A) and ASC-EPCs differentiated in EGM-2 for 3 days (B), 1 week (C), and 2 weeks (D). Bar = 250 µm. AJVR, Vol 75, No. 7, July 2014
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5′-GAATGGAGCACCACAGGTTT-3′), vWF (forward, 5′-GCAATGTCTCCTCTGATGAAG-3′; reverse, 5′-GTACAAGACAACCCCCTGCT-3′), VEGFR2 (forward, 5′-GGTATGGTCCTTGCCTCAGA-3′; reverse, 5′-CAGTGGTATCCGTGTCATCG-3′), and glyceraldehyde-3-phosphate dehydrogenase (forward, 5′-CATTGCCCTCAATGACCACT-3′; reverse, 5′-TCCTTGGAGGCCATGTAGAC-3′). Twenty-five microliters of PCR product was prepared with 2 µL of cDNA, 10 pmol of each primer, 1.25 units of Taq polymerase,m PCR buffer, MgCl2, and a final concentration of 0.2mM dNTPs. Cycle conditions for conventional and real time PCR assays were as follows: conventional PCR assay (95°C for 3 minutes, 35 cycles at 95°C for 30 seconds, 53.5°C for 30 seconds, 72°C for 1.5 minutes, and 72°C for 5 minutes), real time PCR assay (95°C for 2 minutes, 35 cycles at 95°C for 10 seconds, 53.5°C for 30 seconds, 72°C for 30 seconds, and 72°C for 5 minutes). The con-
ventional PCR assay products were analyzed via 1.5% agarose gel electrophoresis and visualized with ethidium bromide. Real-time PCR reaction for VEGFR2 was performed with nucleic acid gel stainn and a real-time PCR detection system.o In vitro solubilized basement membrane tube assay—Solubilized basement membrane preparationp was allowed to thaw at 4°C overnight and then added to 24well culture plates at a concentration of 50 µL/cm2. The coated plates were then incubated at 37°C for 45 minutes to allow for solidification. The ASC-EPCs cultured for 14 days in EGM-2 and undifferentiated ASCs were dissociated with trypsin, reseeded at 2.5 X 104 cells/cm2 on the solubilized basement membrane substrate, and incubated at 37°C for 24 hours. After 24 hours incubation, tube-like network formation was examined and recorded with a phase contrast microscope. LDL uptake assay—The LDL uptake assay was performed with an assay kit,q according to the manufacturer’s instructions. The ASC-EPCs were cultured for 14 days in EGM-2, and undifferentiated ASCs were harvested with trypsin, replated in 48-well culture plates at 2.5 X 104 cells/cm2, and incubated at 37°C for 4 hours in EGM-2 containing LDL working solution. At the end of the LDL uptake incubation, the culture medium was aspirated and replaced with PBS solution. The degree of LDL uptake was determined with a fluorescent microscope.
Figure 2—Quantitative flow cytometric analysis expression of CD31 and CD34 by undifferentiated canine ASCs (A) and ASC-EPCs differentiated in EGM-2 for 1 week (B) and 2 weeks (C). The colored peak indicates the distribution of cells stained with fluorescein isothiocyanate-conjugated antibodies, and the black peak represents a control. The percentage values indicate percentage increase versus controls. FL1-H = Fluorescence intensity of fluorescein isothiocyanate–conjugated antibodies. AJVR, Vol 75, No. 7, July 2014
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In vivo solubilized basement membrane plug assay—Two healthy Beagles were used for subcutaneous implantation of ASC-EPCs and undifferentiated ASCs. All animal procedures were performed in accordance with the guidelines of the Institutional Animal Care and Use Committee of Seoul National University (SNU-131212-1-1). Anesthesia was induced by IV administration of tiletamine hydrochlorider at a dose of 7.5 mg/kg and maintained with isofluranes in oxygen. Intravenously administered tramadolt at a dose of 2 mg/kg was used as an analgesic. Solubilized basement membrane preparation (500 µL) was used to trap 2.5 X 105 ASC-EPCs cultured for 14 days in EGM-2 and 2.5 X 105 undifferentiated ASCs before SC injection. Cell-free plugs of the same size were used as negative controls, and plugs containing recombinant human VEGFu were used as positive controls. After 7 days, plugs were harvested for examination of vessel formation. The sample tissues were fixed in 4% paraformaldehyde for 1 day, dehydrated by use of a graded alcohol series, and embedded in paraffin. Five-micrometerthick slices were mounted on glass slides. 687
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The sections were deparaffinized, hydrated, and stained with H&E. The stained slides were observed via light microscopy to examine vessel formation.
Results
Morphological and immunophenotypic changes—Endothelial differentiation was induced by culStatistical analysis—Data are expressed as mean turing canine ASCs in EGM-2 media. Undifferenti± SD. Statistical analysis was performed with software.v ated ASCs had typical mesenchymal morphology, with The student t test was performed to assess differences a fibroblast-like shape (Figure 1). In contrast, after 3 between 2 groups. A value of P < 0.05 was considered days of culture in an endothelial differentiation medito be significant. um, these cells became less elongated, compared with undifferentiated ASCs. After 1 week of differentiation culture, the cells changed into a more polygonal shape. After 2 weeks of culture, ASC-EPCs acquired a cobblestone-like morphological appearance typical for endothelial lineage. On the basis of a circularity index between 0 and 1 (the circularity index of 1 represents round objects, and the lower the value is, the less round is the object), the circularity index of ASC-EPCs was significantly (P = 0.01) higher after 2-week differentiation, compared with undifferentiated ASCs (n = 15 frames; 0.79 ± 0.10 vs 0.13 ± 0.02). Quantitative flow cytometric analysis of CD31 and CD34 expression was performed to confirm the immunophenotypic changes that occurred under the endothelial differentiation condition. In general, CD31 characterizes mature endothelial cells, and CD34 is used as an EPC marker. Undifferentiated ASCs had no specific expression for CD31 and CD34 (Figure 2). The ASC-EPCs had increased CD34 expression after 1 and 2 weeks of differentiation. The expression Figure 3—Agarose gel electrophoretogram resulting from RT-PCR assay of endothe- of mature endothelial marker CD31 was lial-specific genes (CD31, vWF, and VEGFR2) and glyceraldehyde-3-phosphate dehydrogenase (GAPDH) of undifferentiated canine ASCs and ASC-EPCs differentiated in also mildly increased at the first week, EGM-2 for 1 week and 2 weeks. although it was decreased at 2 weeks of differentiation. However, changes of CD31 expression were not significant, compared with those of CD34. The immunophenotypic changes suggested that alteration of these cells is possible from mesenchymal to endothelial lineage, especially endothelial progenitor type, under endothelial differentiation condition.
Figure 4—Phase contrast microscopic views of capillary network formation by undifferentiated canine ASCs (A) and differentiated ASC-EPCs (B), and fluorescence imaging of LDL uptake by ASCs (C) and ASC-EPCs (D). Notice that network formation and LDL uptake are observed only in ASC-EPCs. Bar = 250 mm. 688
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EPC and EC markers in cells differentiated from canine ASCs—The expression of endothelial specific markers CD31, vWF, and VEGFR2 was analyzed by conventional RT-PCR assay (Figure 3). Positive controls (vessel tissues) expressed the 3 markers. Undifferentiated and differentiated ASCs did not express CD31 and vWF, whereas these cells expressed VEGFR2. The expression of VEGFR2 was also observed in undifferentiated ASCs but was lower than that in differentiated ASC-EPCs. Real-time PCR assay was performed to quantify the mRNA expression levels of VEGFR2 AJVR, Vol 75, No. 7, July 2014
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tional characteristics of endothelial lineage in vitro and in vivo. They formed tubular networks on solubilized basement membrane substrate, integrated LDL in vitro culture, and contributed to vessel-like structures in vivo. Analysis of some surface markers and gene expressions was also performed to assess endothelial differentiation. The differentiated cells had increased expression of CD34 and VEGFR2 after 1 and 2 weeks of endothelial differentiation. Interestingly, however, increased mRNA expressions of CD31 and vWF were not observed after induction, but a small increase of the surface CD31 protein was detected by use of FACS. Expression of CD31 was slightly increased after the first week of endothelial differentiation but was again decreased after the second week of induction. Although this finding Figure 5—Photomicrographic views of blood vessel formation by canine cells in solubilized basement membrane plugs containing no cells (A), VEGF (B), ASCs (C), and of FACS analysis somewhat suggested ASC-EPCs (D). Arrows indicate blood vessels. Notice that little or no blood vessel CD31 expression, this expression was formation is observed in cell-free plugs. H&E stain; bar = 50 mm. less than that of CD34. Therefore, we concluded that canine ASCs expressed during the endothelial differentiation process of the cano or little CD31 after endothelial differentiation by nine ASCs. Quantitative real-time RT-PCR analysis reuse of EGM-2 alone. Similarly, several previous studvealed a consistent increase with time in the transcripies11,12,15,24 found that the cells expressed no or little tion of VEGFR2 during ASC differentiation in EGM-2. CD31 after endothelial differentiation. However, in The expression of VEGFR2 in 1-week differentiated other studies,14,17 the CD31 expression level had apparcells increased approximately 4-fold (P < 0.05), and the ently increased after endothelial differentiation. CD31 expression in 2-week samples increased approximateis a mature endothelial differentiation marker; thus, ly 10-fold (P < 0.05), compared with undifferentiated elongation of differentiation time may lead to an upcells. regulation of CD31.12,15 In addition, the differentiation condition used in the present study may not fully lead Functional characteristics of ASC-EPCs—Tube to mature endothelial differentiation of canine ASCs formation was assessed by seeding undifferentiated and could maintain the endothelial progenitor potenASCs and ASC-EPCs differentiated in EGM-2 on solutial. Indeed, in this study, canine ASCs also maintained bilized basement membrane preparation. Undifferenproliferation potential during the endothelial differentiated cells developed little or no tube-like network tiation culture, although these cells had morphological formation (Figure 4). In contrast, differentiated cells and immunophenotypic changes and functional angiodeveloped clear tube-like structures. genesis characteristics such as LDL uptake and tube In vitro functional characterization was also performation. The results of the present study were conformed by assessing LDL uptake. Undifferentiated sistent with previous research on acquisition of endoASCs had little or no LDL uptake (Figure 4), whereas thelial cell characteristics by human ASCs.19 The preASC-EPCs after cultivation in EGM-2 for 2 weeks had vious study used endothelial cell growth supplement high LDL uptake. as the main stimulus of endothelial differentiation, but The solubilized basement membrane plug assay this supplement alone did not promote the expression was performed to investigate whether ASC-EPCs can of molecular markers. However, CD31 expression and promote new blood vessel formation in vivo. One week LDL uptake were elicited after shear stress was applied after implantation, the plugs were harvested and vesas an additional stimulus. Shear stress is defined as the sel formation was observed. Little or no blood vessel frictional force per unit area, and it acts on endothelial formation was observed in cell-free plugs (Figure 5). cells as blood flows throughout the vascular system.10 Vessels were observed in the positive control containRecent evidence suggests that stem cells differentiing VEGF. Smaller but high numbers of vessels were ate into endothelial cells when exposed to fluid shear found in the plugs seeded with ASCs and ASC-EPCs. stress.10,19 Eventually, the effect of EGM-2 alone on the Differences among groups (except the negative control) differentiation of canine ASCs into mature endothelial were not significant. cells appeared limited. Therefore, further investigations should be carefully performed to evaluate whether Discussion mechanotransduction can modulate mature endothelial differentiation of canine ASCs. Results of the present study revealed that canine Generally, EPCs are characterized by 3 surface ASCs were capable of differentiating into endothelialmarkers including CD133, CD34, and VEGFR2.6 In like cells in vitro. The differentiated cells had the funcAJVR, Vol 75, No. 7, July 2014
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the present study, the differentiated ASC-EPCs had increased expression of CD34 and VEGFR2 as well as functional vascular characteristics. Some previous studies25,26 revealed that human ASCs had a low level of expression of VEGFR2 and could gradually differentiate into endothelial-like cells with increased expression of CD34 and VEGFR2. Therefore, we termed endothelial-differentiated ASCs as ASC-EPCs. However, further investigations are needed to determine why some cells were responsive to the differentiation induction and what the conditions are for differentiating ASCs into mature endothelial cells. A previous report10 suggested that a reliable endothelial differentiation protocol for MSCs was not yet established, whereas standardization of other differentiation procedures such as osteogenesis and adipogenesis was uncomplicated and in agreement with the published protocols. Therefore, protocols for differentiation towards EPCs or endothelial cells should be established for improving the clinical application of these cells. Subcutaneous implantation of solubilized basement membrane plugs containing ASCs, ASC-EPCs, or VEGF resulted in the formation of new blood vessels. Blood vessel density was slightly greater in VEGF and ASC-EPC plugs. As already known, VEGF alone in solubilized basement membrane plugs stimulated cells to form blood vessels. From this, it is speculated that ASCs or ASC-EPCs might also stimulate VEGF secretion to induce blood vessel formation in vivo. However, it can be concluded that differentiation of ASCs into ASC-EPCs did not impair but rather increased their ability to induce development of new blood vessels. However, blood vessel density is known to be dependent on the number of implanted cells24 and may be different according to the period of implantation. Therefore, further studies are required to verify whether differentiation of canine ASCs prior to application can further improve the performance of these cells for neovascularization. The EPCs or CD34-positive cells may be a possible therapeutic option for accelerated re-endothelization and neovascularization. However, a substantial problem in the therapeutic use of these cells is the limited number of autologous cells that can be obtained from adult blood. Furthermore, the number of circulating EPCs is reduced in patients with risk factors for ischemic cardiovascular disease.17,27 In the present study, we successfully obtained CD34-positive endothelial progenitor-like cells by differentiation of canine ASCs with EGM-2. The differentiated cells have the advantage that they can be expanded from ASCs, which are easily obtained in large numbers. Therefore, ASC-EPCs could be a useful cell source for cell-based therapies of ischemic disease and vascularization of engineered tissues. Endothelial growth media-2 is used to maintain endothelial cell growth, but has also been used to differentiate MSCs derived from humans or mice into endothelial lineages in many studies.14,23,24,28 Fibroblast growth factor and VEGF included in EGM-2 are known to play a critical role in the induction of endothelial lineages.14 We attempted to determine whether canine ASCs could differentiate into endothelial-like cells in EGM-2 and confirmed that these cells responded to the 690
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induction of growth factors included in EGM-2. To our knowledge, this study revealed for the first time that ASCs derived from dogs are capable of differentiating towards endothelial-like cells. Generally, rodents have been widely used in preclinical studies for human disease, but clinical data obtained from larger animals such as dogs provide more reliable information with regard to human disease.21 Therefore, the establishment of endothelial differentiation of stem cells derived from dogs will provide the opportunity to evaluate the efficacy and safety of cell therapy for human cardiovascular disease such as myocardial infarction and limb ischemia. In addition, canine ASC-EPCs may have the potential for use in treating dogs with various tissue injuries and ischemic diseases. a. b. c. d. e. f. g. h. i. j. k. l. m. n. o. p. q. r. s. t. u. v.
Collagenase type I, Sigma-Aldrich Inc, St Louis, Mo. Dulbecco modified Eagle medium, Gibco, Billings, Mont. FBS, Gibco, Billings, Mont. EGM-2, Lonza, Walkersville, Mich. JuLI, NanoEnTek, Seoul, Korea. ImageJ software, NIH, Bethesda, Md. Rabbit anti-CD31 polyclonal antibody, Bioss, Beijing, China. Mouse anti-dog CD34 monoclonal antibody, Serotec, Oxford, England. FACS Calibur flow cytometer, BD Biosciences, San Jose, Calif. CELL Quest Pro, BD Biosciences, San Jose, Calif. Trizol, Invitrogen Co, Carlsbad, Calif. Omniscript Reverse Transcription Kit, Qiagen Inc, Valencia, Calif. Taq polymerase, Promega, Madison, Wis. SYBR Green I nucleic acid gel stain, Invitrogen Co, Carlsbad, Calif. CFX ConnectT Real-Time System, Bio-Rad Laboratories Inc, Hercules, Calif. Matrigel, BD Biosciences, San Jose, Calif. LDL Uptake Cell-Based Assay Kit, Cayman Chemical Co, Ann Arbor, Mich. Tiletamine hydrochloride, Virbac, Carros, France. Isoflurane, Baxter Healthcare, Deerfield, Ill. Tramadol, Samsung Pharmaceutical, Seoul, Korea. Recombinant human VEGF, R&D systems, Minneapolis, Minn. SPSS, version 20, SPSS Inc, Chicago, Ill.
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