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GROWTH, DIFFERENTIATION, AND APOPTOSIS
Cardiology Division and Center of Excellence on Aging, "G. d'Annunzio" University, Chieti, Italy
Submitted 3 April 2008 ; accepted in final form 9 September 2008
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ABSTRACT |
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 TOP ABSTRACT MATERIALS AND METHODSRESULTSDISCUSSIONGRANTSREFERENCES |
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adipogenesis; neovasculogenesis; progenitor cells; stromal cells; biopolymers
1:10,000 nucleated cells (43)] coresiding with other types of stem cells (e.g., mesenchymal and hematopoietic stem cells) in a microenvironment rich in growth factors (39, 40). However, the relatively low abundance, the small tissue volume retrieved by sampling, and the difficult accessibility hamper the clinical usefulness of bone marrow-derived EPCs. The mesoderm-derived stromal vascular fraction freshly isolated from adipose tissue has recently attracted much attention because of its capacity to differentiate into mature endothelial cells and participate in the formation of new blood vessels (29, 37). Indeed, the expansion of adipose tissue in adult life is one of the few examples of active postnatal neovascularization (3, 10). Adipose tissue is available from several sources and in large amounts and is relatively easy to obtain through biopsy or liposuction, and its isolation causes minimal discomfort or morbidity at the donor site. Adipose tissue has been recently shown to contain multipotent stem cells, able to differentiate in vitro and in vivo into both mesenchymal and nonmesenchymal lineages, including chondrocytes, adipocytes, osteoblasts, myocytes, and neurons (19, 47). The potential of adipose tissue vessels to grow and regress together with the expansion and regression of the adipocyte mass is related to the plasticity and remodeling capacities of the adipose tissue vasculature (41). Unlike mature vessels, which are typically quiescent and resistant to angioactive agents, neovessels, as occurring in the expanding adipose tissue or in tumors, can readily remodel and respond to such factors (15). This neovascularization potential has been proposed as a therapeutic tool to facilitate the revascularization and healing of ischemic tissues (29, 33, 37). However, the origin of endothelial cells that form the neovasculature in the expanding adipose tissue remains largely unknown.
In the present study we therefore analyzed neovasculogenesis surface antigen expression and kinetics in an in vitro model of developing adipose tissue based on the use of biopolymers such as Matrigel and methylcellulose and determined the relative contribution of circulating progenitor cells and adipose tissue-resident progenitor cells in the formation of new vessels in developing adipose tissue.
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MATERIALS AND METHODS |
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TOPABSTRACTMATERIALS AND METHODSRESULTSDISCUSSIONGRANTSREFERENCES |
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Cell isolation and culture. For the isolation of murine adipose tissue-derived stromal cells (ADSCs), we used 6-wk-old male BALB/c mice (mean weight 28 ± 3 g) from Charles River Laboratories (Margate, UK). For the isolation of human ADSCs, visceral abdominal (omental) adipose tissue was obtained from four individuals without inflammatory conditions or hematologic malignancies undergoing abdominal surgery (body mass index 29 ± 6, age 56 ± 7 yr) under a protocol approved by the local Institutional Review Board.
Animals were anesthetized with an intraperitoneal injection of pentobarbital sodium (50 mg/kg) and sodium heparin (1,000 U/kg). After anesthesia, a laparotomy was performed, periepididymal and visceral (omental) abdominal adipose tissues were harvested, and ADSCs were isolated with a modification of published methods (47). In brief, the human or murine adipose tissues were mechanically minced and digested with type I collagenase. After adipocyte removal by centrifugation at 1,200 rpm for 5 min, the vascular stromal fraction was plated at a density of 1,000 cells/cm2 in Dulbecco's modified Eagle's medium (DMEM)-F-12 medium supplemented with penicillin (100 U/ml), streptomycin sulfate (100 µg/ml), and 10% fetal calf serum (FCS). After 24 h, nonadherent cells were removed, and subconfluent adherent, nonpassaged ADSCs were used for the experiments. To exclude possible contamination of ADSCs with mesothelial cells from the abdominal cavity, which might theoretically also form capillary-like structures in Matrigel and might participate in neoangiogenesis in vivo, the expression profile of the specific markers for mesothelial cells cytokeratin 8 and 18 was analyzed in murine and human ADSCs by flow cytometry (see Supplemental Table S1; Refs. 7, 8).1 For the isolation of murine aortic endothelial cells (mAECs), 6-wk-old male BALB/c mice were anesthetized as described above. After the induction of anesthesia, a thoracotomy was performed and the heart was excised from each animal. The upper end of the thoracic aorta was rapidly cannulated in situ with a 24-gauge blunt-ended needle, while the lower end was ligated with a 7-0-type silk thread. Through the cannula attached to the aorta and a syringe type II collagenase [1% in phosphate-buffered saline (PBS)] was placed into the aorta, and chemical digestion of the vessel intima was performed for 10 min. Cells harvested enzymatically from a pool of six aortas were centrifuged for 5 min at 1,200 rpm and then plated in 12-well plates and cultured in medium 199 (GIBCO BRL, Life Technologies, Milan, Italy) containing HEPES (25 mmol/l), heparin (50 U/ml), endothelial cell growth factor (50 µg/ml), L-glutamine (2 mmol/l), penicillin (100 U/ml), streptomycin sulfate (100 µg/ml), and 15% FCS (BioWhittaker, Bergamo, Italy). The occurrence of satisfactory (>90%) purity of cultures was evaluated by von Willebrand factor (vWF) immunostaining. Cells at passage 1 were used for further experiments.
Mononuclear cell preparation. Peripheral blood, collected in ethylenediaminetetraacetic acid (EDTA)-treated tubes from four individuals at the time of omental adipose tissue harvesting, was diluted with PBS and centrifuged over Ficoll for 30 min at 400 g (4). Mononuclear cells recovered from the interface were washed and used for flow cytometry.
Methylcellulose cultures and colony-forming unit assays. Methylcellulose cultures were performed with mSMCs, adherent murine ADSCs, or adherent murine bone marrow-derived stromal cells (BMSCs), which were trypsinized once after isolation and then introduced into the methylcellulose medium (MethoCult MG3534, StemCell Technologies, Vancouver, BC, Canada), all at 1.5 x 104 cells/ml by single-cell plating. Total BMSCs were obtained by flushing the tibias and femurs of 6-wk-old male BALB/c mice. Plates were examined under phase-contrast microscopy, and colonies were scored after a minimum of 4–14 days from triplicate cultures by two independent investigators. Branched alignments and tubelike structures from ADSC cultures were observed after 10 days and fixed for immunostaining. MethoCult, a culture procedure with limited cell culture and no further plating, proved to be a useful and efficient tool to reveal the presence of endothelial, hematopoietic, and adipogenic progenitors (16). For the immunostaining of tubelike structures, cells in methylcellulose cultures were eluted from the matrix by washing the slides three times with PBS and fixed in 4% paraformaldehyde for 10 min at room temperature. Slides were then blocked in PBS-2% FCS and incubated for 1 h on ice with the following antibodies: 1) fluorescein isothiocyanate (FITC)-conjugated mouse anti-CD34 monoclonal antibody (Miltenyi Biotec, Bergisch Gladbach, Germany) and 2) purified monoclonal goat anti-CD31 antibody (Santa Cruz Biotechnology, Santa Cruz, CA). Isotype-matched negative controls were also used to detect nonspecific staining. After washing with PBS-2% FCS, purified primary antibody were incubated with a FITC-conjugated anti-goat IgG secondary antibody (BD Biosciences, San Jose, CA). The slides were washed and mounted with a solution containing the fluorochrome 4',6-diamidino-2-phenylindole (DAPI, VectaShield, Vector Labs, Burlingame, CA) and viewed through a fluorescence microscope.
Oil Red O staining. ADSCs in methylcellulose were washed with PBS, fixed in a 10% solution of formaldehyde (Sigma) for 1 h, washed with 60% isopropanol (Sigma), and stained with an Oil Red O solution (stock solution from Sigma, diluted in 60% isopropanol) for 10 min. Cells were washed with tap water, destained in 100% isopropanol for 15 min, and then counterstained with the nuclear counterstain hematoxylin QS (Vector Labs).
Tube formation assays. Tube formation assays were performed with a minimal volume of Matrigel in 96-well plates (BD Biosciences), which allows the formation of both tubules and a vascular network. We used nonpassaged human and murine ADSCs, human umbilical vein endothelial cells (HUVECs, at passage 3) cultured as previously described (11) as positive controls, and mSMCs (passage 3) as negative controls. Before the assay, the various cell types were grown for 72 h in endothelial basal medium (EBM, BD Biosciences) supplemented with growth factors [2.5% FCS, 1 µg/ml hydrocortisone, 10 ng/ml human epidermal growth factor, 100 ng/ml vascular endothelial growth factor (VEGF), 3 ng/ml basic fibroblast growth factor, and 15 IU/ml heparin]. At subconfluence, cells were incubated with 1,1'-dioctadecyl-3,3,3',3'-tetramethylindocarbocyanine-labeled acetylated low-density lipoprotein (DiI-acLDL, Molecular Probes, Eugene, OR) at 5 µg/ml for 7 h, then shifted to the same medium enriched with 10% FCS, and plated on Matrigel (at a density of 2 x 105 cells/50 µl Matrigel) for a further 24 h. The uptake of DiI-acLDL by nonpassaged ADSCs characterizes cells of the endothelial lineage (14). Images were acquired in Nomarski and Texas red channels with a x10 objective and an immunofluorescence microscope. Three microscopic fields were selected at random and photographed, and tube areas, tube length, and tube number were quantified with National Institutes of Health (NIH) Image software.
To better determine the progenitor population responsible for the neovascularization, a parallel set of tube formation assays was performed by using each of the following types of ADSCs: total unfractionated ADSCs, DiI-acLDL+ ADSCs, and DiI-acLDL– ADSCs. Before the assay, positive cells for DiI-acLDL uptake were determined by incubation of cells with 5 µg/ml DiI-acLDL for 7 h and subsequent detachment by scraping. DiI-acLDL+ ADSCs were sorted from DiI-acLDL– ADSCs with a fluorescence-based cell sorter (Beckman-Coulter, Miami, FL) and used for tube formation assays (see Supplemental Fig. S1).
Immunoblotting. Total proteins from nonpassaged murine ADSCs or mAECs were isolated in an ice-cold radioimmunoprecipitation assay (RIPA) buffer (50 mM Tris·HCl, 1% NP-40, 0.25% sodium deoxycholate, 150 mmol/l sodium chloride, 1 mmol/l EDTA, and a cocktail of protease inhibitors including aprotinin, leupeptin, and pepstatin, each at 1 µg/ml), separated under reducing conditions, and electroblotted onto polyvinylidene fluoride (PVDF) membranes (Immobilon-P, Millipore, Bedford, MA). After blocking, membranes were incubated overnight at 4°C with the following primary antibodies: 1) monoclonal goat anti-CD31 antibody (Santa Cruz Biotechnology), 2) monoclonal rabbit anti-CD34 antibody (Santa Cruz Biotechnology), 3) monoclonal goat anti-VEGF receptor-2/kinase insert domain receptor (KDR) antibody (Santa Cruz Biotechnology), 4) polyclonal rabbit anti-vWF (DAKO, Carpinteria, CA), and 5) polyclonal rabbit anti-perilipin antibody (Intergen Chemicon, Temecula, CA). Blots were incubated with horseradish peroxidase-coupled secondary antibodies, washed, and developed with a SuperSignal West Pico Chemiluminescent Substrate kit (Pierce, Rockford, IL). The intensity of each immunoreactive protein band was quantified by densitometric analysis (Kodak Digital Science ID Image Analysis Software, Eastman Kodak, Rochester, NY). To verify equal loading of proteins, membranes were stripped and reprobed with a monoclonal anti-β-actin antibody (Sigma).
Flow cytometry analysis of ADSCs and peripheral blood mononuclear cells.
For flow cytometry analysis of EPC markers, human and murine ADSCs were washed with PBS and detached by scraping in 3 mmol/l EDTA-Hanks’ balanced salt solution without trypsin. Peripheral blood mononuclear cells (PBMNCs) or ADSCs (1 x 106 resuspended in 100 µl PBS-2% FCS-2 mmol/l EDTA) were incubated for 20 min at 4°C with 10 µl of the following antibodies: 1) FITC-conjugated mouse anti-CD34 monoclonal antibody (Miltenyi Biotec), which detects a class III epitope on all glycoforms of the CD34 antigen and 2) phycoerythrin (PE)-conjugated mouse anti-CD133 monoclonal antibody (Miltenyi Biotec). For flow cytometry analysis of mesenchymal stem cell and monocyte/macrophage markers (see Supplemental Table S1), human and murine ADSCs were incubated with the following antibodies: 1) pan-CD45 peridinin-chlorophyll-protein (PerCP)-conjugated antibody (detecting all isoforms and glycoforms of CD45; from BD Biosciences); 2) purified rat anti-mouse CD105 (endoglin) monoclonal antibody (BD Biosciences), cross-specific for the analogous human antigen; 3) purified rat anti-mouse CD44 monoclonal antibody (Cedarlane Laboratories, Burlington, ON, Canada), cross-specific for the analogous human antigen; 4) FITC-conjugated anti-human CD29 (ImmunoTools, Friesoythe, Germany); 5) FITC-conjugated anti-human CD71 (ImmunoTools); 6) FITC-conjugated anti-human CD29 (ImmunoTools); 7) FITC-conjugated anti-human CD106 (Santa Cruz Biotechnology); and 8) FITC-conjugated anti-human CD14 (ImmunoTools). For flow cytometry analysis of pericytes and smooth muscle cell lineages (Refs. 17, 21; see Supplemental Table S1), human and murine ADSCs were incubated with the following antibodies: 1) FITC-conjugated anti-human desmin (Sigma-Aldrich); 2) purified human 
-smooth muscle cell actin (-SMA, Sigma-Aldrich). All antibodies were used at the manufacturers’ recommended concentrations after in-house verification of reactivity. Isotype-matched negative controls were also used to detect nonspecific events. Such events were subsequently subtracted from those attributed to positive events estimated in the samples. After washing with PBS-2% FCS-2 mmol/l EDTA, purified primary antibodies were incubated with a PE-conjugated goat anti-rat IgG secondary antibody (BD Biosciences). After a further washing with PBS-2% FCS-2 mmol/l EDTA, labeled cells were analyzed by flow cytometry with a FACSCalibur flow cytometer (BD Biosciences). The instrument was calibrated daily with CaliBRITE beads (BD Biosciences) with the aid of FACSComp software (BD Biosciences). Additionally, compensation was checked visually in each run. A forward scatter (FSC) vs. side scatter (SSC) live gate was set in the acquisition to exclude debris and cellular aggregates. FSC and SSC were recorded by linear amplification with a first fluorescence detector (FL1) and a second fluorescence detector (FL2), respectively, and a logarithmic amplification. For each sample, a total of 30,000 events were acquired and analyzed with CellQuest software (BD Biosciences). For the flow cytometry analysis of cytokeratins 8/18, ADSCs (n = 1 x 106) were fixed with cold methanol for 15 min and then permeabilized in PBS-2% FCS-0.1% Triton X-100 for 10 min. Cells were incubated for 60 min at 4°C directly with a purified guinea pig anti-cytokeratin 8/18 polyclonal antibody (Progen Biotechnik, Heidelberg, Germany) cross-specific for both the human and the mouse antigens. After washing with PBS-2% FCS-2 mmol/l EDTA, labeled cells were analyzed by flow cytometry as described above.
Flow cytometry analysis of whole peripheral blood. To assess the quality and reliability of the CD34+/CD133+ cell quantification estimate by the previously described flow cytometry protocol, we compared it with the CD34+/CD133+ cell quantification estimate obtained by the International Society of Hematotherapy and Graft Engineering (ISHAGE) protocol. This has been designed to provide guidelines for the accurate detection of human blood-derived circulating CD34+ cells, based on five flow cytometry parameters: CD45 conjugated with PerCP, CD34-FITC, CD133-PE staining, SSC, and FSC. Importantly, this approach allows the discrimination of hematopoietic stem cells (which express relatively low levels of CD45 on their surface) from lymphocytes and monocytes, thus allowing the assessment of "true" CD34+ cells, which are "dim" for CD45 fluorescence and have a low SSC (CD45dim, SSClow).
An exact volume of 100 µl of accurately mixed human peripheral blood, collected in EDTA tubes from each of the four individuals tested in this analysis at the time of harvesting of adipose tissue, was placed on the bottom of the tubes with a repeater pipette (Eppendorf 4780, Brinkmann Instruments, Mississauga, ON, Canada) and incubated for 20 min at 4°C with 10 µl of the pan-CD45 PerCP-conjugated antibody (detecting all isoforms and glycoforms of CD45; from BD Biosciences) and 10 µl of the previously described anti-CD34 and anti-CD133 antibodies. Isotype-matched negative controls were also used to detect nonspecific events exhibiting fluorescence and light scatter characteristics of CD45+, CD34+, and CD133+ cells. Such events were subsequently subtracted from the number of CD45+, CD34+, and CD133+ cells estimated in the labeled samples. Lysis of red blood cells was subsequently performed by adding 2 ml of ammonium chloride (BD Pharm-Lyse lysing solution, BD Biosciences) All tubes were gently vortexed and incubated at room temperature for 15 min in the dark, washed twice in PBS (by centrifugation at 500 g for 5 min), and resuspended in 1 ml of PBS. Samples were then stored on ice at 4°C in the dark and analyzed by flow cytometry within 1 h. Flow cytometry analysis was performed on a FACSCalibur flow cytometer (BD Biosciences). Calibration and compensation were performed as described above. FL1, FL2, and FL3 were recorded with logarithmic amplification. To satisfy statistical requirements, 100 CD34+ events or 70,000 total events were acquired and analyzed with CellQuest software (BD Biosciences).
According to the ISHAGE protocol, we used the following gating strategy: an R1 region, representing CD45+ events, included all nucleated white blood cells. Red blood cells, nucleated red blood cells, platelets, and other cellular debris, identified as the R7 region, were excluded from R1, since they do not express CD45. CD45+ events in the R1 region were then analyzed for CD34 staining, and positive events were gated into an R2 region. Events defined by R1 and R2 regions were analyzed on dot plots describing granularity vs. CD45 staining. Events representing true blast cells, which were identified as SSClow, CD45dim, were gated in the R3 region. Events in the R4 region were defined as cells with characteristics of blast cells and lymphocytes. Cells from the R4 region were named true CD34+ blast cells. Events in the R5 region identified lymphocytes and were counted on dot plot 5 as R1 and R5. Events in the R6 region identified monocytes and were counted on dot plot 1 as R1 and R8. True CD34+ blasts were finally plotted against CD133+ events recorded in the FL2 channel.
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RESULTS |
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TOPABSTRACTMATERIALS AND METHODSRESULTSDISCUSSIONGRANTSREFERENCES |
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Deprivation of the DiI-acLDL+ cell population from ADSCs resulted in a total inhibition of tube or network formation (Fig. 2, I and L). By comparing quantitative parameters, such as tube area, of total ADSCs with dioctadecyl-3,3,3',3'-tetramethylindocarbocyanine (DiD)-acLDL+ ADSCs substantial differences were found, since an increase in this parameter was observed by using total ADSCs (Fig. 2M). Together, these results indicate that the DiD-acLDL+ cell population might be considered as the main cellular part of adipose tissue functionally active in terms of vessel formation and physically involved in vessel formation, while the remaining cell populations including pericytes, smooth muscle cells, and preadipocytes, which do not show any uptake of DiD-acLDL (i.e., DiD-acLDL– ADSCs), might modulate this angiogenic activity of DiI-acLDL ADSCs, most probably by producing angiogenic and antiangiogenic factors (33, 38) or by stabilizing endothelial networks (42).
Neovasculogenesis-specific gene expression profile in ADSCs and role of "circulating" progenitor cells in formation of adipose tissue neovessels. We next determined the neovasculogenesis-specific surface antigen profile of nonpassaged ADSCs. By immunoblotting, we analyzed the expression profile of early markers for endothelial differentiation (i.e., CD34 and KDR) and for mature endothelial cells (i.e., vWF and CD31) (16, 32) in ADSCs compared with adult mAECs. ADSCs clearly showed positive staining for CD34 and KDR but expressed almost no CD31, whereas mAECs showed strong expression of CD31. By contrast, ADSCs featured lower levels of vWF than mAECs (Fig. 3).
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-SMA and human desmin in primary cultures of ADSCs was 47.9 ± 4% and 7.4 ± 5%, respectively (see Supplemental Table S2). Figure 5 shows two different methods to quantify CD34+/CD133+ progenitor cells in peripheral blood, based on the presence (Fig. 5A) or the absence (Fig. 5B) of CD45 expression. We compared the number of progenitor cells from ADSCs with the number of progenitor cells from peripheral blood, quantified either as CD34+/CD133+ cells (Fig. 5A) or CD45dim/CD34+/CD133+ cells (Fig. 5B and Table 3). With the use of more stringent gating strategies for peripheral blood, based on the absence of CD45 expression, the percentage of CD45–/CD34+/CD133+ cells within ADSCs was markedly higher than CD45dim/CD34+/CD133+ cells in the blood. The specificity of immunostaining was checked by using isotype-matched murine IgG conjugated with FITC, PE, or PerCP (Supplemental Fig. S3, A and B).
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DISCUSSION |
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TOPABSTRACTMATERIALS AND METHODSRESULTSDISCUSSIONGRANTSREFERENCES |
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We here addressed the issue of mechanisms regulating vascular growth and remodeling from fat-derived stromal tissue in an in vitro model of adipose tissue development from ADSCs based on biopolymers such as Matrigel or methylcellulose. Immunofluorescence microscopy and flow cytometry analyses revealed that a substantial number of ADSCs express early markers of endothelial differentiation and low levels of markers for mature endothelial cells, therefore suggesting the presence of a population of progenitor cells as an intrinsic characteristic of ADSCs. Animal studies using hindlimb ischemia or myocardial ischemia models in immune-deficient rodents have demonstrated that transplantation of 106 peripheral blood-derived EPCs (25) can enhance angiogenesis. The rarity of EPCs in peripheral blood would require large amounts to permit the isolation of sufficient numbers of cells to achieve a proangiogenic effect. Such amounts of blood are not readily obtainable in a clinical setting. For the bone marrow, traditional procedures of tissue procurement may yield low numbers of stem cells upon processing (1 bone marrow-derived stem cell per 105 adherent stromal cells; Ref. 43). On the basis of recently published studies that demonstrate the importance of using stringent gating strategies to quantitatively analyze EPCs, such as the lack of expression of CD45 together with the expression of CD34 and CD133 (5), we here compared the yield of EPCs from adipose tissue and peripheral blood. We found that adipose tissue is capable of yielding a higher number of EPCs than peripheral blood, thus supporting the contention that it may be a better source of EPCs for revascularization-based therapies.
ADSCs, traditionally used to study adipocyte differentiation, were shown here also to be able to differentiate into endothelial cells. Indeed, Matrigel, rich in extracellular matrix largely from basement membranes and derived from a mouse sarcoma line enriched with proangiogenic growth factors, including basic fibroblast growth factor, hydrocortisone, human epidermal growth factor, and VEGF, allowed ADSCs to form new vessels in vitro, producing networks of branched tubelike structures. Cultures in methylcellulose and immunofluorescence staining for CD34 and DiI-acLDL, two markers of both mature and progenitor endothelial cells (13, 21, 26, 35), and CD31, a marker for mature endothelial cells (16, 32), demonstrated that the kinetics of neovascularization in ADSCs strictly paralleled the formation of intracellular lipid vacuoles. This is a demonstration of the existence of extensive bidirectional cross talks between adipocytes and endothelial cells in the developing adipose tissue, where extracellular matrix components likely play a role in coordinating adipogenesis and angiogenesis (3, 44).
The vascularized adipose tissue may also contain other types of cells, such as smooth muscle cells and multipotent stem cells, such as pericytes and bone marrow-derived mesenchymal stem cells (19, 47). Disruption of the blood supply during fat tissue isolation may result in the release of circulating bone marrow-derived mesenchymal stem cells and pericytes, known to possess the potential for multilineage differentiation (36). It was therefore plausible that a population of EPCs in the adipose tissue stromal fraction consists of circulating progenitor cells originating from the bone marrow or other sources. The presence of such cells in the circulation and within ischemic tissues has been clearly established (1, 2, 25), but the question remained as to the extent and relevance of their contribution to newly forming blood vessels in the adipose tissue. Three lines of evidence from our study suggest that such cells do not play a major role in the formation of new vessels in the developing adipose tissue. First, EPCs in the peripheral blood and in the microcirculation circulate at low levels (27), and the volume of blood in the adipose tissue is small, making the presence of such cells in sufficient number to produce neovessels unlikely. Second, and more cogently, the total ADSC fraction isolated from the adipose tissue in clonal assays in our studies always failed, at variance from positive controls from BMSCs, to support hematopoiesis. One would expect the development of hematopoietic colonies in the case of significant infiltration with circulating progenitor cells. This strongly argues against the hypothesis that the angiogenic potential of adipose tissue derives from an infiltration with circulating progenitor cells. The growth pattern and the expression of differentiation markers for EPCs in the stromal population from the adipose tissue rather indicate a relative abundance of tissue-resident progenitor cells, niching in the stromal compartment of the adipose tissue and capable of forming new vessels. Third, we have excluded the possibility that cells in the stromal population from adipose tissue, which are negative for DiI-acLDL uptake (i.e., all of nonendothelial cell lineages) could be responsible for adipose tissue-derived neovascularization, since we have shown that DiI-acLDL+ ADSCs, but not DiI-acLDL– ADSCs, possess the capacity for tube/network formation of adipose tissue. However, we admittedly cannot totally exclude the possibility that ADSC-induced vessel formation and growth might also be related to the contribution of preadipocytes and resident microvascular endothelial cells through the secretion of proangiogenic factors (28, 31, 33, 34, 38), or through the contribution of periendothelial pericytes capable of interacting with and stabilizing endothelial networks (42). This still, however, would point out the important advance derived from our observations that non-BMSCs, resident in adipose tissue and capable of evolving toward adipocytes, have the potential of growing new vessels in appropriate conditions, while losing the ability to undergo adipocyte differentiation. Certainly, the use of an in vitro model of developing adipose tissue, such as our culture system in methylcellulose medium with limited cell amplification and no subfractioning of ADSCs, where a number of variables can be controlled, appears to be an appropriate tool to investigate such factors. Of note, we report much lower percentages of CD29-positive cells, marking mesenchymal stem cells, compared with other reports in the literature (18, 30, 47). Differences in the source of adipose tissue (visceral adipose tissue in our case vs. subcutaneous adipose tissue in all other reports) and cell passaging (nonpassaged cells in our case) and the avoidance in our case of the use of formalin or paraformaldehyde, decreasing background fluorescence, may account for such discrepancies.
The findings of these in vitro studies admittedly suffer from limitations in not precisely reproducing the environment within a living ischemic heart or other ischemic tissues. We are currently conducting in vivo experiments in which ADSCs derived from adipose tissue are transplanted into animal hearts in models of myocardial infarction or ischemia. Such in vivo preclinical investigations could generate important information that may, in turn, warrant clinical trials of ADSCs in selected patients with heart disease.
Biocompatible materials are an emerging area of research in cardiac tissue engineering, showing promise in the treatment of myocardial infarction (6, 12, 22, 45). Injectable matrix biopolymers, such as fibrin, collagen I, and Matrigel, have been shown to improve left ventricular remodeling by providing structural support to aneurismal thinning of the left ventricular wall (23, 24). When used in combination with cells such as skeletal myoblasts, cardiomyocytes, endothelial cells, embryonic stem cells, and bone marrow-derived stem cells, biopolymers may serve as cell delivery vehicles by providing a favorable matrix environment that confers properties of improved viability, migration, and proliferation on cells (23, 45). The results of our study therefore also highlight the potential of using three-dimensional biopolymers, such as Matrigel or methylcellulose, as cell delivery vehicles capable of enhancing neovascularization-based tissue repair by ADSCs.
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FOOTNOTES |
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The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
1 The online version of this article contains supplemental material. 
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