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INTRODUCTION: IFATS COLLECTION: TISSUE-SPECIFIC STEM CELLS

IFATS Collection: The Role of Human Adipose-Derived Stromal Cells in Inflammatory Microvascular Remodeling and Evidence of a Perivascular Phenotype

^aDepartment of Biomedical Engineering and
^bDepartment of Plastic Surgery, University of Virginia, Charlottesville, Virginia, USA

Key Words. Adipose-derived stromal cells • Microcirculation • Pericyte • Angiogenesis

Correspondence: Correspondence: Shayn M. Peirce, Ph.D.,Dept. of Biomedical Engineering, University of Virginia, P.O. Box 800759, 415 Lane Rd., Charlottesville, VA 22908, USA. Telephone: 434-243-5795; Fax: 434-982-3870; e-mail: speirce{at}virginia.edu

Received on January 11, 2008; accepted for publication on April 7, 2008.

First published online in STEM CELLS EXPRESS  April 24, 2008.

	ABSTRACT

Top Footnotes Abstract Introduction Materials and Methods Results Discussion Conclusions Disclosure of Potential... Acknowledgments References

 
A growing body of literature suggests that human adipose-derived stromal cells (hASCs) possess developmental plasticity both in vitro and in vivo, and might represent a viable cell source for therapeutic angiogenesis and tissue engineering. We investigate their phenotypic similarity to perivascular cell types, ability to contribute to in vivo microvascular remodeling, and ability to modulate vascular stability. We evaluated hASC surface expression of vascular and stem/progenitor cell markers in vitro, as well as any effects of platelet-derived growth factor B chain (PDGF-BB) and vascular endothelial growth factor 165 on in vitro hASC migration. To ascertain in vivo behavior of hASCs in an angiogenic environment, hASCs were isolated, expanded in culture, labeled with a fluorescent marker, and injected into adult nude rat mesenteries that were stimulated to undergo microvascular remodeling. Ten, 30, and 60 days after injection, tissues from anesthetized animals were harvested and processed with immunohistochemical techniques to determine hASC quantity, positional fate in relation to microvessels, and expression of endothelial and perivascular cell markers. After 60 days, 29% of hASCs exhibited perivascular morphologies compared with 11% of injected human lung fibroblasts. hASCs exhibiting perivascular morphologies also expressed markers characteristic of vascular pericytes: smooth muscle -actin (10%) and neuron-glia antigen 2 (8%). In tissues treated with hASCs, vascular density was significantly increased over age-matched controls lacking hASCs. This study demonstrates that hASCs express pericyte lineage markers in vivo and in vitro, exhibit increased migration in response to PDGF-BB in vitro, exhibit perivascular morphology when injected in vivo, and contribute to increases in microvascular density during angiogenesis by migrating toward vessels.

Disclosure of potential conflicts of interest is found at the end of this article.

	INTRODUCTION

Top Footnotes Abstract Introduction Materials and Methods Results Discussion Conclusions Disclosure of Potential... Acknowledgments References

 
Much attention has been devoted to the plasticity of different adult cell types, termed "adult stem cells" or "progenitor cells," in injury responses and as a therapeutic tool. Circulating [1, 2], bone marrow-derived [3–5] and tissue-derived [6] progenitor cells have been implicated in the process of microvascular remodeling. An expanding body of literature suggests that cells within the stromal-vascular fraction (SVF) of human adipose tissue possess substantial developmental plasticity, both in vitro [7] and in vivo [8, 9]. These stromal cells have alternatively been referred to as processed lipoaspirate cells, adipose-derived stem cells, and adipose-derived stromal cells. We refer to these cells as adherent adipose-derived stromal cells (ASCs) to distinguish them from SVF cells, which have not been separated based on adherence to tissue culture plastic. The cell surface phenotype of adherent human ASCs (hASCs) differs from freshly isolated SVF cells, undergoing changes in response to variables such as exposure to tissue culture plastic, duration in culture, and exposure to specific medias/supplements [8–14]. Previous studies suggest that stromal cells isolated from adipose tissue on the basis of adherence to tissue culture plastic have a remarkably consistent molecular and cell surface profile [8, 15, 16]. Interestingly, this profile is quite similar to that of stromal cells isolated from bone marrow (also referred to as mesenchymal stem cells) [7, 11, 17–19], yet an easily definable phenotype remains elusive.

hASCs are readily available as they can be harvested in large quantities using minimally invasive techniques, and they can be expanded in vitro [20]. In addition, previous work has shown that hASCs can be genetically modified to secrete proangiogenic proteins [9], making this cell population an appealing and practical candidate for translation of autologous transplantation strategies into the clinical setting. These cells have been shown to differentiate into chondrogenic, myogenic, osteogenic, and adipogenic cells in the presence of lineage-specific induction factors in culture [20]. Moreover, adipose-derived stromal cells have been shown to differentiate into endothelial cells [8, 10, 14, 21], form vascular-like sprouts in matrigel [8], enhance neovascularization in an ischemic hind limb model [8–10], and secrete angiogenic and antiapoptotic growth factors [10], suggesting a potential for this cell population in therapeutic vascularization and tissue engineering of vascularized constructs. It has been hypothesized that the proangiogenic activity of human adipose-derived stromal cells is a combined result of their ability to produce angiogenic growth factors and to differentiate into endothelial cells [8–10, 14, 21]. In addition, several recent studies have shown in vitro evidence that hASCs can assume a pericyte role; however, data supporting functional benefit to the vasculature in vivo have not yet been produced [22–24], leaving the in vivo role of hASCs as perivascular cells in question.

Although most previous work has focused on the role of endothelial cell migration and proliferation during angiogenesis, a critical component of microvascular growth is the recruitment of perivascular support cells (such as pericytes and smooth muscle cells) to the abluminal surface of the microvessel wall. This step is important for vessel maintenance via prevention of microvascular regression [15], physical guidance of capillary sprouts [25], and regulation of capillary permeability [26]. Furthermore, it has been suggested that pericytes can differentiate into vascular smooth muscle cells in response to growth factor signals and function to transform a capillary into a contractile arteriole, thus participating in the process of arteriogenesis [27, 28].

Since it has been suggested that pericytes contribute to microvessel growth [25] and maintenance [15], we tested the hypothesis that hASCs function as microvascular support cells by analyzing their perivascular investment in relation to changes in total vascular density. We show on a single-cell level that hASCs are capable of expressing perivascular cell markers in vitro and in vivo, responding to platelet-derived growth factor B chain (PDGF-BB) with increased migration in vitro, exhibiting pericyte-like morphologies in vivo by migrating to the abluminal surface of microvessels and conforming to the curvature of the microvessel in a manner that is consistent with pericyte (and not endothelial) cell behavior, and increasing total microvascular length when injected into remodeling rat mesenteries compared with mesenteries receiving vehicle control (no cells) or human lung fibroblasts (hLFs). Therefore, we present pericyte-like behavior as a role for human adipose-derived stromal cells in the promotion of vascular stability and the enhancement of microvascular growth in vivo.

	MATERIALS AND METHODS

Top Footnotes Abstract Introduction Materials and Methods Results Discussion Conclusions Disclosure of Potential... Acknowledgments References

 
Isolation, Culture, and Labeling of hASCs
Subcutaneous adipose tissue was obtained from 14 female patients undergoing elective surgical procedures in the Department of Plastic Surgery, University of Virginia (age range: 30–56 years; mean, 42 years old) for in vivo experiments. The University of Virginia's Human Investigation Committee approved tissue harvest protocols. Adipose tissue came either from intraoperative suction lipectomy (n = 6) or from laboratory liposuction of panniculectomy specimens (n = 8). Cells used in scratch test experiments were isolated by intraoperative suction lipectomy from a single 38-year-old female donor.

Cells were isolated from adipose tissue using methods previously described [11, 20]. Briefly, harvested tissue was washed several times and enzymatically dissociated [11, 29]. Dissociated tissue was filtered to remove debris, and the resulting cell suspension was centrifuged. Pelleted stromal cells were recovered and washed several times. Contaminating erythrocytes were lysed with an osmotic buffer, and the stromal cells were plated onto tissue culture plastic. Cultures were washed with buffer 24–48 hours after plating to remove unattached cells, and then refed with fresh medium. Plating and expansion medium consisted of Dulbecco's modified Eagle's medium (DMEM)/F12 (Invitrogen, Carlsbad, CA, http://www.invitrogen.com) with 10% fetal bovine serum (FBS; Invitrogen) and 1% antibiotic-antimycotic (Invitrogen). Cultures were maintained at 37°C with 5% CO₂ and were fed three times per week.

Cells were grown to confluence after the initial plating (passage 0; p = 0), typically within 10–14 days. Adherent cells were released with either 0.5% trypsin-EDTA or Accutase cell detachment medium (Innovative Cell Technologies, San Diego, http://www.innovativecelltech.com/) and replated at 2,000 cells/cm² (p = 1). Cell cultures were passaged every 7–8 days until analysis. All cells used for injection studies were between passage 2 and 4, corresponding to approximately 11 or fewer total population doublings.

One day prior to injection, cells were labeled with the fluorescent marker 1,1'-dioetadeeyl-3,3,3',3'-tetramethylindocarboeyanine perchlorate (DiI, 5 µM) according to the manufacturer's instructions (Molecular Probes, Eugene, OR, http://probes.invitrogen.com). Cells were rinsed, trypsinized, counted, and resuspended for injection. The use of DiI as a label for identifying progenitor cells in vivo has been well documented [16, 30–32]. Furthermore, plated hASCs were labeled with DiI and maintained in culture to confirm that DiI fluorescence did not diminish visibly over time or with cell division.

Immunophenotypic Characterization of hASCs
The vascular-related cell surface phenotype of hASCs was analyzed using flow cytometry. Freshly isolated (SVF) cells and cultured hASCs were evaluated for their expression of the following cell surface proteins over time in culture: CD31 (PECAM-1) (BD Biosciences, San Diego, http://www.bdbiosciences.com), CD34 (BD Biosciences; Santa Cruz Biotechnology, Inc., Santa Cruz, CA, http://www.scbt.com; and Caltech Labs, Carlsbad, CA, http://www.invitrogen.com), CD133 (Miltenyi Biotech, Auburn, CA, http://www.miltenyibiotec.com), CD144 (VE-Cadherin) (BD Biosciences), CD140b (PDGF-β receptor) (BD Biosciences), and neuron-glia antigen 2 (NG2) (Beckman Coulter, Fullerton, CA, http://www.beckmancoulter.com). Flow cytometry was performed on a Becton Dickinson FACSCalibur (Becton, Dickinson and Company, Franklin Lakes, NJ, http://www.bd.com), as previously described [11]. A minimum of 10,000 events were counted for each analysis. Human leukocyte antigen-ABC was used as a positive control for each flow trial. Isotype-matched (negative) controls were also performed in all cases.

In Vitro Scratch Test
Isolated ASCs were allowed to grow to confluence on culture-treated plastic (Corning Inc., Corning, NY, http://www.corning.com/index.aspx) at p = 3 (37°C, 5% CO₂). Cells were cultured for 24 hours prior to experimentation in DMEM/F12 with 0% FBS and 1% antibiotic/antimycotic. A 1,000-µl pipette tip was used to scratch each dish five times before cells were washed with Dulbecco's phosphate-buffered saline (DPBS). The cells were then subjected to culture in DMEM/F12 containing no serum, 1% antibiotic/antimycotic, and either 10 ng/ml recombinant human PDGF-BB (R&D Systems, Minneapolis, http://www.rndsystems.com) or 20 ng/ml recombinant human vascular endothelial growth factor 165 (VEGF₁₆₅; Biovision Inc., Mountain View, CA, http://www.biovision.com/). In a separate experiment, scratch wounds were allowed to heal with either 10 ng/ml PDGF-BB, 500 ng/ml PDGF-BB antibody (Chemicon Int., Temecula, CA, http://www.chemicon.com), 1 µg/ml PDGF receptor β antibody (Sigma Biosciences, St. Louis, http://www.sigma-aldrich.com), 10 ng/ml PDGF-BB plus 500 ng/ml PDGF-BB antibody, or 10 ng/ml PDGF-BB plus 1 µg/ml PDGF-β receptor antibody. Individual untreated controls containing no additives were run in parallel for each of these two experimental designs. All cells were used at passage 3.

Dishes were imaged at 0, 12, 20, and 30 hours after introduction of scratches. In all cases, scratch healing was determined by measuring the shortest distance between scratch edges in each field of view. At least 16 fields of view were analyzed for each condition at each time point comparing PDGF and VEGF effects on scratch closure, and at least 51 fields of view were analyzed for each condition at each time point comparing the abilities of PDGF antibody and PDGF-β receptor antibody to abrogate scratch closure in the presence of PDGF.

Animal Studies
Experiments were performed using sterile techniques according to the guidelines of the University of Virginia Animal Care and Use Committee. Seventy-two male nude rats (National Cancer Institute, Bethesda, MD, http://www.cancer.gov/) weighing 250 ± 20 g were divided into six study groups: (a) hASC injection (1 x 10⁶ cells), (b) hASC injection (1 x 10⁶ cells) and 48/80 stimulation, (c) hLF injection (1 x 10⁶ cells), (d) hLF injection (1 x 10⁶ cells) and 48/80 stimulation, (e) vehicle control (sterile phosphate-buffered saline [PBS]), and (f) vehicle control and 48/80 stimulation.

Stimulation of Microvascular Remodeling with Compound 48/80 and Cell Injection
Compound 48/80 (condensation product of N-methyl-p-methoxyphenylethylamine with formaldehyde; Sigma Biosciences) is a pharmacological agent known to act specifically on mast cells by inducing degranulation. Injection of compound 48/80 into the rat mesentery stimulates well-characterized microvascular growth and remodeling in the mesenteric vasculature [25, 33], and this small-animal model is a well-established assay for studying cellular and molecular mechanisms of angiogenesis [34, 35]. Compound 48/80 was injected i.p. (1 ml/100-g animal weight) in 0.9% sterile NaCl on the first five consecutive study days. Two doses of each concentration (100, 200, 300, and 400 µg/ml) were administered per day separated by 8 hours on the first four days. On day 5, rats in these study groups received a single dose of 500 µg/ml. To rule out any direct effects of compound 48/80 on hASC survival, proliferation, and differentiation, cultured hASCs (p = 2) were exposed to either 100 or 5 µg/ml of compound 48/80 in the media for 6 days.

On day 4 of 48/80 injections, 1 x 10⁶ DiI-labeled hASCs or hLFs (WI-38 cell line, no. CCL-75; American Type Culture Collection, Manassas, VA, http://www.atcc.org) in 0.5 ml sterile PBS were injected i.p. using syringes with 25-G, 0.5-inch needles.

Harvesting of Mesenteric Tissue
Rats were anesthetized with intramuscular injections of ketamine (80 mg/kg body weight [bw]), atropine (0.08 mg/kg bw), and xylazine (8 mg/kg bw). Six mesenteric windows were harvested from each animal 10, 30, or 60 days after cell injection. Tissues were whole-mounted on gelatin-coated slides.

Immunohistochemistry
To determine whether injected hASCs expressed markers consistent with a perivascular cell phenotype, tissues were immunostained for an array of markers known to be expressed by smooth muscle cells and pericytes: NG2, smooth muscle -actin (SMA), PDGF-β receptor, and desmin [36, 37]. Tissues were washed in PBS + 0.1% saponin three times for 10 minutes and immunolabeled with lectin from Bandeiraea simplicifolia (BSI-lectin) fluorescein isothiocyanate (FITC)-conjugate (1:100; Sigma Biosciences) or Alexa Fluor 647-conjugate (1:100; Molecular Probes), and/or antibody to SMA using purified FITC-conjugated clone 1A4 mouse monoclonal anti-SMA (1:500; Sigma Biosciences), diluted in PBS buffer containing 0.1% saponin and 2% bovine albumin (Fisher Scientific, Hampton, NH, http://www.fisherscientific.com) at pH 7.4 (incubation for 1 hour at room temperature). Tissues were also stained with perivascular cell markers [36], including antibodies to the following: NG2 (1:150, rabbit polyclonal; Chemicon Int.), desmin (1:100, mouse antihuman clone D33; DAKO, Glostrup, Denmark, http://www.dako.com), and PDGF-β receptor (1:100, rabbit polyclonal; Santa Cruz Biotechnology Inc.). Cytochrome 2-conjugated secondary antibodies were applied for 1 hour at room temperature: NG2 and PDGF-β receptor goat anti-rabbit IgG, and desmin goat anti-mouse IgG-fab fragment (1:100; Jackson Immunoresearch Laboratories, West Grove, PA, http://www.jacksonimmuno.com). In the final wash cycle, Hoechst 33342 (1 x 10^–6 mM) or TOTO-3 (1:1000; Molecular Probes) was added for visualization of nuclei.

Image Acquisition and Data Analysis
Mesenteric tissues were examined with a Nikon Eclipse TE2000-E microscope equipped with confocal accessories (Nikon D-Eclipse C1) using x20 Nikon water/oil immersion and x60 Nikon oil immersion objectives (Melville, NY, http://www.nikonusa.com/). Images were digitized and analyzed using Scion Image software version 4.0.2 (Scion Corporation, Frederick, MD, http://www.scioncorp.com). The number of DiI-positive cells per tissue area and total microvessel length were quantified. Nuclei were visualized with TOTO-3 and Hoechst 33342 to confirm the presence of DiI-labeled hASCs and hLFs.

Although there is no single marker/protein that defines a pericyte phenotype, NG2, SMA, and desmin are considered to be supportive of, and consistent with, this lineage [36]. Pericytes are perhaps best defined by their histological and anatomical positions and shape, with the sine qua non being their close physical contact with microvascular endothelial cells.

Therefore, to compliment and expand upon our immunophenotypic findings, detected cells were examined for pericyte-like morphology, defined here as cells whose processes extend along vessels in a manner that conforms to the curvature of the vessel and whose cell bodies are no more than 5 µm from the abluminal surface of the endothelium. Morphometrists were blinded to the treatment group at the time of analysis. The cell behavior quantified using this method is distinct from smooth muscle cell morphology, which is characterized by wrapping of the smooth muscle cell around the abluminal endothelial surface in a direction perpendicular to the vessel axis and parallel to adjacent smooth muscle cells. Therefore, we term the cell morphology observed here "pericyte-like" as opposed to "perivascular-like," which encompasses both pericyte and smooth muscle cell morphology.

Scratch test data were collected using a Nikon TE2000-E2 equipped with confocal accessories (Nikon D-Eclipse C1) using a x10 Nikon air objective and an Olympus Microfire digital camera (Tokyo, http://www.olympus-global.com). These data were analyzed using ImageJ ver. 1.37 software (National Institutes of Health, Bethesda, MD, http://www.nih.gov/).

Statistical Analysis
Results are presented in the form of mean ± standard error. Comparisons for in vivo data were made using the statistical analysis tools provided by SigmaPlot 5.0 (Systat Software, Inc., Chicago, http://www.systat.com/). Data were tested for normality and analyzed by one-way analysis of variance followed by nonpaired Tukey's t test. Significance was asserted at p < .05. In vitro experiments were analyzed using SigmaStat 3.5 (Systat Software, Inc.) and are presented in the form of ± one standard deviation. These data were analyzed by unpaired t test or Mann-Whitney rank sum test when normality tests failed.

	RESULTS

Top Footnotes Abstract Introduction Materials and Methods Results Discussion Conclusions Disclosure of Potential... Acknowledgments References

 
Immunocharacterization of hASCs
The vascular-related cell surface profile of human SVF (hSVF) and hASCs was analyzed using flow cytometry and/or immunohistochemistry (SMA, desmin). A very small percentage (2.5% or less) of early passage hASCs stain positively for markers of mature and/or activated endothelial cells (CD31, CD144), and these phenotypic markers become undetectable by passage 5 (Fig. 1A). Under the described isolation and culture conditions, less than 1% of hASCs stain for the progenitor/stem cell marker CD133 (data not shown), whereas 9%–97% of them stain positively for CD34, with specific expression levels depending on both the length in culture and the specific antibody used (Fig. 1B). hASCs consistently stain for several markers consistent with a perivascular phenotype [36] including NG2 (Fig. 1C, mean of p = 1 through p = 4 equals 15% ± 6%, n = 3), PDGF-β receptor (CD140b; 76%–97% by flow; mean, 90%), and SMA (approximately 25% by immunocytochemistry; data not shown). Flow cytometry data showed that compound 48/80 did not enhance NG2 expression (data not shown).

View larger version (28K): [in this window] [in a new window]   Figure 1. In vitro characterization of human adipose-derived stromal cell surface marker expression. (A): Average expression (± standard deviation) of CD31 (white) and CD144 (black) is minimal and declines to 0%. (B): Detected expression of CD34 depends on the specific antibody used (8G12: white, 581: black, BI-3C5: gray). (C): Average expression (± standard deviation) of neuron-glia antigen 2 using flow cytometry.

View larger version (38K): [in this window] [in a new window]   Figure 2. In vitro migration of human adipose-derived stromal cells (hASCs) in the presence of platelet-derived growth factor B chain (PDGF-BB) or vascular endothelial growth factor 165 (VEGF165). (A): Relative hASC migration (± standard deviation; *, p < .001) into untreated scratch wounds or wounds treated with VEGF165 or PDGF-BB. (B): Inhibition of PDGF-BB-induced migration using antibodies to PDGF-BB and PDGF-β receptor (± standard deviation; *, p < .001; , p < .005; , p < .01; #, p < .05).

View larger version (43K): [in this window] [in a new window]   Figure 3. Pericyte-like localization and surface marker expression of human adipose-derived stromal cells (hASCs) in rat mesentery. Rat mesenteric tissues injected with human cells were harvested 60 days later and immunostained to visualize microvessels. (A): Some 1,1'-dioetadeeyl-3,3,3',3'-tetramethylindocarboeyanine perchlorate-positive (DiI+) hASCs express neuron-glia antigen 2 (yellow, arrows), some do not (red), and some exhibit pericyte-like morphologies (vessel wrapping) and are aligned with capillaries that express lectin from Bandeiraea simplicifolia (BSI-lectin, blue). (B): Some hASCs expressing smooth muscle -actin (SMA, yellow) and exhibiting pericyte-like morphologies (arrows) along BSI-lectin-positive capillaries (blue). (C–D): Examples of DiI-labeled SMA-expressing hASCs (yellow) in mesenteric tissue costained with SMA and BSI-lectin (both stains are green). (E): Some human lung fibroblasts (hLFs, red) exhibit pericyte-like morphologies (arrows) along BSI-lectin-positive capillary (blue) and SMA-positive venules (green), although hLFs do not express SMA and total vessel density is reduced compared with hASC-injected tissue, as seen in (B). (F): No DiI+ cells are present in control, uninjected tissues (green: BSI-lectin). Scale bar = 25 µm in (A, B, E, F); scale bar = 20 µm in (C, D).

View larger version (32K): [in this window] [in a new window]   Figure 4. Quantification of in vivo pericyte-like localization and surface marker expression in 48/80-stimulated tissues. (A): Percentage of human adipose-derived stromal cells (hASCs) and human lung fibroblasts (hLFs) that exhibit pericyte-like morphologies regardless of pericyte marker expression. (B): Percentage of hASCs and hLFs in the tissue that exhibit pericyte-like morphologies (but not pericyte markers). (C): Percentage of total hASCs in the tissue that express pericyte markers (but not morphology). (D): Percentage of total hASCs in the tissue that both exhibit pericyte-like morphologies and express pericyte markers. Error bars represent standard error; *, p .05.

Vascular Density of Microvessels Is Affected by hASCs at Early and Late Time Points
Since some hASCs exhibited pericyte-like morphologies in vivo, and native pericytes in the microvasculature have been shown to contribute to microvessel maintenance and prevent vascular regression, we investigated the functional effect of adipose-derived cells on microvascular length density in networks stimulated to undergo remodeling. Compound 48/80 stimulation alone (no cells) is known to evoke peak vascular remodeling responses between 14 and 20 days after delivery [33]. Accordingly, our data shows that even by day 10 compound 48/80 increases vascular length density approximately 35% in the absence of cells compared with vascular length density in mesenteries receiving neither compound 48/80 nor hASCs (Fig. 5; compare 10-day white bars between Figure 5A and 5B). Ten days after hASC injection, length density, or total vascular length, increased significantly in tissues treated with hASCs relative to tissues treated with hLFs (Fig. 5A; compare black bar to gray bar). This observation is independent of compound 48/80 treatment (Fig. 5B). After 30 days, there was no significant difference in length density between treatment groups in 48/80-stimulated (Fig. 5A) or unstimulated tissues (Fig. 5B). By day 60, the presence of hASCs in both compound 48/80-stimulated (Fig. 5A) and unstimulated tissues (Fig. 5B) evokes significant length density increases compared with that of hLF-treated and vehicle control-treated tissues. In stimulated tissues, this increase in length density coincides temporally with the elevated percentage of hASCs exhibiting a pericyte-like morphology compared with hLFs (Fig. 4A). Furthermore, in tissues receiving hASCs, vascular length density at day 60 is elevated to levels comparable to those measured on day 10, regardless of compound 48/80 treatment (Fig. 5A, 5B).

View larger version (24K): [in this window] [in a new window]   Figure 5. Functional impact of human adipose-derived stromal cells (hASCs) on vascular length density in vivo. Vascular length density for vehicle control (no cells: white) versus hASC (black) versus human lung fibroblast (hLF, gray) in (A) 48/80-treated tissues and (B) tissues not treated with 48/80. Error bars represent standard error; *, p .05.

View larger version (17K): [in this window] [in a new window]   Figure 6. Distribution of human adipose-derived stromal cells (hASCs) with respect to length density quartile. (A): Percentage of total hASCs in the tissue that exhibit pericyte-like morphologies at day 60, and (B) percentage of total hASCs that exhibit pericyte-like morphologies and also express pericyte markers for each quartile of length density at day 60. Error bars represent standard error; *, significantly different (p .05).

	DISCUSSION

Top Footnotes Abstract Introduction Materials and Methods Results Discussion Conclusions Disclosure of Potential... Acknowledgments References

The reason for the apparent increase in pericyte marker expression relative to endothelial marker expression may be as simple as the fact that the hASCs in this study were cultured before use. It is possible that culturing hASCs could select for pericyte-like cells over endothelial cells via their relative abilities to adhere to culture plastic. Alternatively, the genetic expression profiles of the hASCs may be altered by the artificial conditions of culture (adhesion mechanisms/forces, media ingredients, oxygenation, pH), causing the cells to truly differentiate into pericyte-like cells.

As reported both in these experiments and those performed by other groups [13, 14, 39], time in culture is a factor for expression levels of CD34, a molecule generally recognized as a marker of hematopoietic stem/progenitor cells, as well as microvascular endothelial cells, bone marrow stromal progenitors, and fibroblast-like dendritic cells from adult dermis and adipose tissue [40]. Approximately 1% of early passage hASCs were found to be CD133+, a protein expressed by hematopoietic stem cells and/or endothelial progenitor cells [41]. This is consistent with Miranville et al., who reported 1.5%–5.3% of freshly isolated SVF cells to express the CD133 marker, with exact levels varying depending on the source depot of adipose tissue [10]. The low level of CD133 expression in culture and loss of CD34 expression over time in culture could mean that time in culture causes hASC lineage commitment instead of subselection through adhesion.

It has been shown that detection and quantification of CD34 on a given cell population can be highly variable depending on the class of antibody that is used, as well as the particular antibody conjugate [42]. Our findings for hASCs are consistent with this work, demonstrating higher levels of CD34 expression/detection in early passage cells compared with later passage cells, and when using class III antibodies (8G12 and 581) compared with class I antibody (BI-3C5) [42]. The qualitative implication of these quantitative discrepancies is not clear at this time, however; the potential significance of distinct CD34-positive hASC subpopulations deserves further evaluation.

hASCs Participate in Microvascular Growth and Maintenance
The fact that neither hASCs nor hLFs are detectable at day 10 in tissues untreated with compound 48/80 suggests that either compound 48/80 or the proangiogenic, inflamed tissue environment created by this stimulus promotes hASC retention/survival in vivo early on. The former explanation can be ruled out by in vitro studies (data not shown), indicating that if compound 48/80 has any direct effect on hASCs, it is to moderately reduce, not enhance, their proliferative capabilities. This suggests, instead, that it is the environment created by compound 48/80 that initially enhances hASC presence in the mesentery (Table 1, Day 10).

The lack of a significant difference in length density between 48/80-stimulated and unstimulated tissues at day 30 is to be expected based on the transient nature of the remodeling response to compound 48/80, whereby newly sprouted vessels gradually regress after the peak remodeling response, causing length density to return to control levels after the inflammatory response has subsided. The subsequent increase in length density at day 60 in 48/80-stimulated tissues coincides temporally with the elevated percentage of hASCs exhibiting a pericyte-like phenotype compared with hLFs (Figs. 4, 5), suggesting that the presence of hASCs with pericyte-like phenotypes and the increase in vascular density are causally related. This supports the possibility that the presence of hASCs in the tissue at late time points provides longer lasting support to vessels that would normally regress in vehicle control- or hLF-treated tissues. Figure 6 shows that stimulated tissues with the highest length densities also have the highest percentages of hASCs exhibiting pericyte-like morphology and expressing pericyte markers. This suggests that there is a significant, and possibly causative, correlation between vascular length density and pericyte-like behavior in the inflamed environment.

Although ASC counts (Table 1) suggest that the inflamed tissue environment created by compound 48/80 stimulation is supportive of cell proliferation or cell survival/migration, the vascular density results suggest that, in tissues unstimulated by compound 48/80, ASCs are able to persist in the in vivo environment and elicit a potent vascular density response at later time points, even though relatively few ASCs are invested in the mesenteric tissue. It is possible that although ASCs are not invested in the mesenteric tissue (and, therefore, are undetected) in animals untreated by compound 48/80, they may be affecting the angiogenic environment through a paracrine mechanism. Thus, the biphasic nature of the length density measurements and ASC investment in rat mesenteries could be a superposition of tissue remodeling in response to inflammation from initial injections and the paracrine angiogenic secretions of injected ASCs over time.

These two proposed mechanisms for increasing vascular length density are not identical, so we may hypothesize that the inflamed environment caused by compound 48/80 treatment provides a cue for ASCs to assume a pericyte-like phenotype and associate directly with vessels in the mesenteric tissue instead of maintaining a default paracrine support phenotype. In comparison, the less inflamed environment of peritoneal cavities untreated with compound 48/80 may be experiencing increased length densities at late time points due primarily to the angiogenic environment (and activation of native pericytes) provided by secretions from ASCs that are performing paracrine instead of pericyte-like roles. If large enough in magnitude, paracrine secretions may account for increases in vascular length density without a significant presence of ASCs in the mesenteric tissue itself.

	CONCLUSIONS

Top Footnotes Abstract Introduction Materials and Methods Results Discussion Conclusions Disclosure of Potential... Acknowledgments References

 
This study presents a description of the in vivo behavior of unsorted, adherent hASCs on the single-cell level and suggests that this behavior is indicative of a vascular support or maintenance role. We show that injected hASCs show increased migration in response to VEGF₁₆₅ and PDGF-BB in vitro, express markers of perivascular cells both in vivo and in vitro, persist over time in vivo, exhibit pericyte-like morphologies around microvessels, and contribute to vascular growth and/or maintenance, as reflected by increases in vascular density compared with vehicle or cell control-treated tissues. This work suggests that hASCs might be an effective source of perivascular cells for therapeutic delivery or in the engineering of tissue equivalents, where perivascular cell support signals are likely to be necessary for long-term vessel maintenance [43].

Future work is needed to determine whether these behaviors are consistent with a vascular growth or vascular maintenance role. However, it has been shown that these cells produce antiapoptotic growth factors as well as angiogenic growth factors, including granulocyte-macrophage colony-stimulating factor, hepatocyte growth factor, basic fibroblast growth factor, VEGF, and transforming growth factor β, suggesting that both roles are possible and consistent with those of native pericytes [9, 19]. Whether injected hASCs are able to differentiate into perivascular cells, whether and how this is mediated by the in vivo tissue environment, and the relative importance of isolated hASCs expressing a perivascular phenotype prior to injection remain to be determined through future studies. Future work should also aim to decouple the presence of ASCs from their angiogenic secretions either by injecting appropriate amounts of those secretions over the 60-day period or by silencing key proangiogenic secretions prior to ASC injection.

	DISCLOSURE OF POTENTIAL CONFLICTS OF INTEREST

Top Footnotes Abstract Introduction Materials and Methods Results Discussion Conclusions Disclosure of Potential... Acknowledgments References

 
The authors indicate no potential conflicts of interest.

	ACKNOWLEDGMENTS

Top Footnotes Abstract Introduction Materials and Methods Results Discussion Conclusions Disclosure of Potential... Acknowledgments References

 
This work was supported by the National Institutes of Health grants HL72141 (A.J.K.) and HL031912 (S.M.P.) and by a charitable gift from Dr. and Mrs. Peyton Weary (A.J.K.). The authors thank the faculty and staff of the Department of Plastic Surgery for facilitating the harvest of adipose tissue and Peter Stapor for assistance with image analysis.

	FOOTNOTES

 
Author contributions: P.J.A.: conception and design, administrative support, collection and assembly of data, data analysis and interpretation, manuscript writing; H.S., A.M.B. and A.T.: collection and assembly of data; A.J.K.: financial support, provision of study material and patients; S.M.P.: conception and design, financial support, administrative support, provision of study material, collection and assembly of data, data analysis and interpretation, final approval of manuscript.

	REFERENCES

Top Footnotes Abstract Introduction Materials and Methods Results Discussion Conclusions Disclosure of Potential... Acknowledgments References

 

1. Iba O, Matsubara H, Nozawa Y et al. Angiogenesis by implantation of peripheral blood mononuclear cells and platelets into ischemic limbs. Circulation 2002;106:2019–2025.

2. Peichev M, Naiyer AJ, Zhu Z et al. Expression of VEGFR-2 and AC133 by circulating human CD34+ cells identifies a population of functional endothelial precursors. Hemostas Thomb Vasc Biol 2000;95:952–958.

3. Asahara T, Haruchika M, Tomono T et al. Bone marrow origin of endothelial progenitor cells responsible for postnatal vasculogenesis in physiological and pathological neovascularization. Circ Res 1999;85:221–228.[Abstract/Free Full Text]

4. Rajantie I, Ilmonen M, Alminaite A et al. Adult bone marrow-derived cells recruited during angiogenesis comprise precursors for periendothelial vascular mural cells. Blood 2004;104:2084–2086.

5. Carmeliet P, Luttun A. The role of the bone marrow-derived stem cells in (therapeutic) angiogenesis. Thromb Haemostas 2001;86:289–297.[Medline]

6. Majka SM, Jackson KA, Kienstra KA et al. Distinct progenitor populations in skeletal muscle are bone marrow derived and exhibit different cell fates during vascular regeneration. J Clin Invest 2003;111:71–79.[CrossRef][Medline]

7. Gronthos S, Franklin DM, Leddy HA et al. Surface protein characterization of human adipose tissue-derived stromal cells. J Cell Physiol 2001;189:54–63.[CrossRef][Medline]

8. Planat-Benard V, Silvestre JS, Cousin B et al. Plasticity of human adipose lineage cells toward endothelial cells. Circulation 2004;109:r23–r30.[CrossRef]

9. Rehman J, Traktuev D, Li J et al. Secretion of angiogenic and antiapoptotic factors by human adipose stromal cells. Circulation 2004;109:r52–r58.

10. Miranville A, Heeschen C, Sengenes C et al. Improvement of postnatal neovascularization by human adipose tissue-derived stem cells. Circulation 2004;110:349–355.

11. Katz AJ, Tholpady A, Tholpady S et al. Cell surface and transcriptional characterization of human adipose-derived adherent stromal (hADAS) cells. STEM CELLS 2005;23:412–423.[Abstract/Free Full Text]

12. Boquest AC, Shahdadfar A, Frønsdal K et al. Isolation and transcription profiling of purified uncultured human stromal stem cells: Alteration of gene expression after in vitro cell culture. Mol Biol Cell 2005;16:1131–1141.[Abstract/Free Full Text]

13. Meyerrose TE, De Ugarte DA, Hofling AA et al. In vivo distribution of human adipose-derived mesenchymal stem cells in novel xenotransplantation models. STEM CELLS 2007;25:220–227.[Abstract/Free Full Text]

14. Mitchell JB, McIntosh K, Zvonic S et al. Immunophenotype of human adipose-derived cells: temporal changes in stromal-associated and stem cell-associated markers. STEM CELLS 2006;24:376–385.[Abstract/Free Full Text]

15. Benjamin LE, Hemo I, Keshet E. A plasticity window for blood vessel remodeling is defined by pericyte coverage of the preformed endothelial network and is regulated by PDGF-B and VEGF. Development 1998;125:1591–1598.[Abstract]

16. Barresi V, Belluardo N, Sipione S et al. Transplantation of prodrug-converting neural progenitor cells for brain tumor therapy. Cancer Gene Therapy 2003;10:396–402.[CrossRef][Medline]

17. De Ugarte DA, Alfonso Z, Zuk PA et al. Differential expression of stem cell mobilization-associated molecules on multi-lineage cells from adipose tissue and bone marrow. Immunol Letters 2003;89:267–270.[CrossRef][Medline]

18. Ibberson D, Tremain N, Gray A. What is in a name? Defining the molecular phenotype of marrow stromal cells and their relationship to other stem/progenitor cells. Cytotherapy 2001;35:409–411.

19. Kilroy GE, Foster SJ, Wu X et al. Cytokine profile of human adipose-derived stem cells: Expression of angiogenic, hematopoietic, and pro-inflammatory factors. J Cell Physiol 2007;212:702–709.[CrossRef][Medline]

20. Zuk PA, Zhu M, Mizuno H et al. Multilineage cells from human adipose tissue: Implications for cell-based therapies. Tissue Eng 2001;7:211–227.[CrossRef][Medline]

21. Sengenes C, Miranville A, Maumus M et al. Chemotaxis and differentiation of human adipose tissue CD34+/CD31- progenitor cells: Role of SDF-1 released by adipose tissue capillary endothelial cells. STEM CELLS 2007;25:2269–2276.[Abstract/Free Full Text]

22. Kang YJ, Jeon ES, Song HY et al. Role of c-Jun N-terminal kinase in the PDGF-induced proliferation and migration of human adipose tissue-derived mesenchymal stem cells. J Cell Biochem 2005;95:1135–1145.[CrossRef][Medline]

23. Traktuev D, Merfeld-Clauss S, Li J et al. A population of multipotent CD34-positive adipose stromal cells share pericyte and mesenchymal surface markers, reside in a periendothelial location, and stabilize endothelial networks. Circ Res 2008;102:77–85.[Abstract/Free Full Text]

24. Zannettino ACW, Paton S, Arthur A et al. Multipotent human adipose-derived stromal stem cells exhibit a perivascular phenotype in vitro and in vivo. J Cell Physiol 2008;214:413–421.[CrossRef][Medline]

25. Nehls V, Denzer K, Drenckhahn D. Pericyte involvement in capillary sprouting during angiogenesis in situ. Cell Tissue Res 1992;270:469–474.[CrossRef][Medline]

26. Hellstrom M, Gerhardt H, Kalen M et al. Lack of pericytes leads to endothelial hyperplasia and abnormal vascular morphogenesis. J Cell Biol 2001;153:543–553.[Abstract/Free Full Text]

27. Hirschi KK, Rohovsky SA, Beck LH et al. Endothelial cells modulate the proliferation of mural cell precursors via platelet-derived growth factor-BB and heterotypic cell contact. Circ Res 1998;84:298–305.

28. Peirce SM, Skalak TC. Microvascular remodeling: A complex continuum spanning angiogenesis to arteriogenesis. Microcirculation 2003;10:99–111.[CrossRef][Medline]

29. Katz AJ. In: Atala A, Lanza RP, eds. Methods of Tissue Engineering. Mesenchymal Cell Culture: Adipose Tissue. Burlington, MA: Academic Press, 2002:277-286.

30. Iwaguro H, Yamaguchi J, Kalka C et al. Endothelial progenitor cell vascular endothelial growth factor gene transfer for vascular regeneration. Circulation 2002;105:732–738.

31. Weber A, Pedrosa I, Kawamoto A et al. Magnetic resonance mapping of transplanted endothelial progenitor cells for therapeutic neovascularization in ischemic heart disease. Eur J Cardio Thoracic Surg 2004;26:137–143.[CrossRef]

32. Mothe AJ, Tator CH. Proliferation, migration, and differentiation of endogenous ependymal region stem/progenitor cells following minimal spinal cord injury in the adult rat. Neuroscience 2005;131:177–187.[CrossRef][Medline]

33. Norrby K, Jakobsson A, Sorbo J. Mast-cell-mediated angiogenesis: A novel experimental model using the rat mesentery. Virchows Archiv B Cell Pathol 1986;52:195–206.[CrossRef]

34. Anderson CR, Ponce AM, Price RJ. Immunohistochemical identification of an extracellular matrix scaffold that microguides capillary sprouting in vivo. J Histochem Cytochem 2004;52:1063–1072.[Abstract/Free Full Text]

35. Bocci G, Danesi R, Benelli U et al. Inhibitory effect of suramin in rat models of angiogenesis in vitro and in vivo. Cancer Chemother Pharmacol 1999;43:205–212.[CrossRef][Medline]

36. Gerhardt H, Betsholtz C. Endothelial-pericyte interactions in angiogenesis. Cell Tissue Res 2003;314:15–23.[CrossRef][Medline]

37. Murfee WL, Skalak TC, Peirce SM. Differential arterial/venous expression of NG2 proteoglycan in perivascular cells along microvessels: Identifying a venule-specific phenotype. Microcirculation 2005;12:151–160.[Medline]

38. Hasumi Y, Klosowska-Wardega A, Furuhashi M et al. Identification of a subset of pericytes that respond to combination therapy targeting PDGF and VEGF signaling. Int J Cancer 2007;121:2606–2614.[CrossRef][Medline]

39. Oedayrajsingh-Varma MJ, Breuls RGM, Schouten TE et al. Phenotypical and functional characterization of freshly isolated adipose tissue-derived stem cells. Stem Cells Dev 2007;16:91–104.[CrossRef][Medline]

40. Lanza F, Healy L, Dutherland DR. Structural and functional features of the CD34 antigen: An update. J Biol Reg And Homeostatic Agents 2001;15:1–13.

41. Hristov M, Erl W, Weber PC. Endothelial progenitor cells isolation and characterization. Trends In Cardiovas Med 2003;13:201–206.[CrossRef]

42. Croockewit AJ, Raymakers RAP, Preijers FWMB et al. The role of different CD34 epitopes in detection and positive selection of CD31+ bone marrow and peripheral blood stem cells. Scand J Immunol 1998;47:82–90.[CrossRef][Medline]

43. Bouhadir JH, Mooney DJ. Promoting angiogenesis in engineered tissues. J Drug Targeting 2001;9:397–406.

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