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TISSUE-SPECIFIC STEM CELLS

Three-Dimensional Perfusion Culture of Human Adipose Tissue-Derived Endothelial and Osteoblastic Progenitors Generates Osteogenic Constructs with Intrinsic Vascularization Capacity

Departments of Surgery and Research, Institute for Surgical Research and Hospital Management, University Hospital Basel, Basel, Switzerland

Key Words. Mesenchymal stem cells • Fat tissue • Bioreactor • Osteoinductive • Bone graft • Angiogenesis

Correspondence: Ivan Martin, Ph.D., Tissue Engineering Group, Institute for Surgical Research and Hospital Management, University Hospital Basel, Hebelstrasse 20, CH-4031 Basel, Switzerland. Telephone: 41-61-265-2384; Fax: 41-61-265-3990; e-mail: imartin{at}uhbs.ch

Received on February 13, 2007; accepted for publication on April 8, 2007.

First published online in STEM CELLS EXPRESS  April 19, 2007.

	ABSTRACT

Top Abstract Introduction Materials and Methods Results Discussion Conclusion Disclosures of Potential... Acknowledgments References

 
In this study, we aimed at generating osteogenic and vasculogenic constructs starting from the stromal vascular fraction (SVF) of human adipose tissue as a single cell source. SVF cells from human lipoaspirates were seeded and cultured for 5 days in porous hydroxyapatite scaffolds by alternate perfusion through the scaffold pores, eliminating standard monolayer (two-dimensional [2D]) culture. The resulting cell-scaffold constructs were either enzymatically treated to extract and characterize the cells or subcutaneously implanted in nude mice for 8 weeks to assess the capacity to form bone tissue and blood vessels. SVF cells were also expanded in 2D culture for 5 days and statically loaded in the scaffolds. The SVF yielded 5.9 ± 3.5 x 10⁵ cells per milliliter of lipoaspirate containing both mesenchymal progenitors (5.2% ± 0.9% fibroblastic colony forming units) and endothelial-lineage cells (54% ± 6% CD34⁺/CD31⁺ cells). After 5 days, the total cell number was 1.8-fold higher in 2D than in three-dimensional (3D) cultures, but the percentage of mesenchymal- and endothelial-lineage cells was similar (i.e., 65%–72% of CD90⁺ cells and 7%–9% of CD34⁺/CD31⁺ cells). After implantation, constructs from both conditions contained blood vessels stained for human CD31 and CD34, functionally connected to the host vasculature. Importantly, constructs generated under 3D perfusion, and not those based on 2D-expanded cells, reproducibly formed bone tissue. In conclusion, direct perfusion of human adipose-derived cells through ceramic scaffolds establishes a 3D culture system for osteoprogenitor and endothelial cells and generates osteogenic-vasculogenic constructs. It remains to be tested whether the presence of endothelial cells accelerates construct vascularization and could thereby enhance implanted cell survival in larger size implants.

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

	INTRODUCTION

Top Abstract Introduction Materials and Methods Results Discussion Conclusion Disclosures of Potential... Acknowledgments References

 
In the context of tissue engineering and regenerative medicine, there is increasing recognition of the importance of coculturing different cell types in a three-dimensional (3D) environment in order to generate constructs with increased functionality and engraftment capacity [1, 2]. In particular, the coculture of tissue-specific cells together with endothelial cells has been proposed as a tool to address one of the main limitations of tissue engineered grafts, namely their rapid vascularization [3, 4]. For example, 3D coculture of myoblasts together with endothelial cells improved blood perfusion and survival of the muscle tissue constructs after transplantation [2]. In addition, capillary-like structures formed in vitro by coculture of keratinocytes and endothelial cells were able to create a connection with the host vessels upon implantation, thus accelerating blood supply of hypoxic zones within the graft [5]. The approach is currently being extended to generate a prevascular network in engineered bone constructs in order to possibly favor an accelerated vascularization and increased cell survival upon implantation [6, 7]. In all these studies, however, the tissue-specific and endothelial cells are derived from different sources or even species, and thus have limited relevance regarding potential clinical applications.

The stromal vascular fraction (SVF) of adipose tissue is known to include progenitor cells with extensive plasticity and a capacity to differentiate into various musculoskeletal lineages as well as into neuronal and endothelial cells [8]. In particular, typical osteogenic culture supplements were shown to induce osteoblastic cell differentiation [9, 10], whereas the endothelial component in SVF was reported to produce vascular-like structures on Matrigel in vitro and to produce neovascularization in an in vivo ischemic mouse model [11]. Thus, adipose tissue could be seen as a common source of osteoblastic and endothelial cells for the engineering of bone grafts. However, beyond some general traits of osteoblastic differentiation in vitro, the actual capacity of human SVF cells to generate bone tissue in vivo is still controversial and appears to require either preculture in osteogenic medium [12] or transfection with bone morphogenic protein (BMP)-2 [13]. Moreover, although the endothelial component is present in the SVF, it is rapidly lost in culture [11, 14] and can be induced and/or maintained only by serial changes in substratum and culture medium [15] or by culture in specific media [11, 16].

We recently reported that bone marrow-derived mesenchymal stromal cells can be expanded by direct perfusion of fresh bone marrow cells through the pores of 3D ceramic scaffolds, avoiding monolayer (two-dimensional [2D]) expansion in plastic dishes [17]. The perfusion culture system, as compared with typical 2D expansion, yields a higher fraction of clonogenic mesenchymal cells and generates more reproducibly osteogenic constructs. Moreover, upon medium supplementation with specific additives, it allows the maintenance in culture of nonmesenchymal cell populations, including early hematopoietic progenitors [17]. Thus, a 3D culture system under perfusion has the potential to support coculture of cell types of different lineages, likely providing appropriate "niches" for their interaction.

In this study, we aimed at using the SVF of human adipose tissue as a common cell source to (a) establish a 3D coculture system for osteoblast- and endothelial-lineage cells and (b) generate constructs that are both osteogenic and vasculogenic upon ectopic implantation in nude mice. In order to achieve this goal, freshly isolated SVF cells were directly perfused through the pores of 3D ceramic scaffolds, bypassing the typical phase of monolayer culture in plastic dishes.

	MATERIALS AND METHODS

Top Abstract Introduction Materials and Methods Results Discussion Conclusion Disclosures of Potential... Acknowledgments References

 
Cell Isolation
Subcutaneous adipose tissue in the form of lipoaspirates was obtained from 12 healthy donors (20–63 years old, body mass index 19.5–25.1 kg/m²) during routine lipoaspirations of the inner and/or outer thighs after informed consent from the patient and following protocol approval by the local ethical committee. The tissue was digested in 0.075% collagenase type 2 (Worthington, Lakewood, NJ, http://www.worthington-biochem.com) for 45 minutes at 37°C on an orbital shaker. The suspension was thereafter centrifuged at 300g for 10 minutes, and the resulting SVF pellet was washed once with phosphate-buffered saline (PBS), resuspended in -minimal essential medium (MEM) (Gibco, Grand Island, NY, http://www.invitrogen.com), and finally filtered through a 100-µm strainer (BD Falcon; BD Biosciences, San Diego, http://www.bdbiosciences.com). Bone marrow-derived nucleated cells from donors with matched age and body mass indexes were obtained from the iliac crest during routine orthopedic surgical procedures as previously described [18].

Monolayer (2D) Cell Cultures
SVF cells were seeded onto Petri dishes at a density of 20 x 10³ cells per cm² in -MEM containing 10% fetal bovine serum (basic medium) supplemented with 5 ng/ml fibroblast growth factor-2 (FGF-2). Addition of FGF-2 was introduced following preliminary experiments, indicating its capacity to sustain reproducible cell proliferation during expansion, in analogy with previous observations using bone marrow-derived mesenchymal cells [19]. The number of clonogenic cells, generally referred to as colony forming units-fibroblastic (CFU-f), was determined by plating 500 or 1,000 SVF cells in 56-cm² dishes followed by staining with crystal violet and counting of colonies after 2 weeks of culture. The proliferation rate was determined by counting the number of cells at the first plating and at the end of this passage by using a Neubauer chamber. The total number of doublings during the culture period, defined as the base 2 logarithm of the ratio between final cell number and initial cell number, was divided by the number of days in culture to obtain the number of cell doublings per day. The initial number of CFU-f plated was calculated from the number of nucleated cells plated by using the independently determined clonogenic frequency.

Osteogenic Differentiation
Freshly harvested SVF cells or the same cells expanded up to three passages, hereafter referred to as adipose tissue stromal cells (ATSC), were cultured in osteogenic medium for 3 weeks, with medium changes twice per week. Osteogenic differentiation medium consisted of basic medium supplemented with 100 nM dexamethasone, 10 mM ß-glycerophosphate, and 0.05 mM ascorbic acid-2-phosphate. After 3 weeks, cell layers were washed twice with PBS, fixed for 10 minutes with 4% formalin, and washed twice with water. Cells were then either incubated for 10 minutes with alizarin red and washed extensively with water or stained with 5% AgNO₃ under UV for 20 minutes (von Kossa staining) and washed thereafter with water and 5% Na₂S₂O₃, 5 H₂O.

Adipogenic Differentiation
Adipogenic differentiation was induced in 2D cultures using cycles of treatment with different media as previously described [20]. Briefly, cells were seeded and cultured until confluence. Cells were then treated with 10 µg/ml insulin, 10^–5 M dexamethasone, 100 µM indomethacin, and 500 µM 3-isobutyl-1-methyl xanthine (adipogenic induction medium) for 72 hours and subsequently with 10 µg/ml insulin (adipogenic maintenance medium) for 24 hours. The 96-hour treatment cycle was repeated four times, and cells were then cultured for an additional week in adipogenic maintenance medium.

Three-Dimensional Perfusion Cultures
Cell seeding and subsequent culture of freshly isolated cells from adipose tissue in 3D scaffolds were performed using a previously described bioreactor system [21], which is based on the principle of continuous direct perfusion of cell suspensions or culture medium through the pores of 3D scaffolds in alternate directions. SVF cells (8.6 x 10⁵ cells per milliliter) were perfused at a velocity of 3 ml/minute through porous hydroxyapatite ceramic scaffolds (ENGIpore; Fin-Ceramica Faenza, Faenza, Italy, http://www.finceramica.it) with an average porosity of 83% ± 3% and the size of 8-mm diameter, 4-mm-thick disks (3 x 10⁶ cells per disk, corresponding to approximately 2.5 x 10³ cells per cm²) in the same medium as for 2D cultures. After 3 days, the initial cell suspension was totally removed and replaced by fresh medium. After two additional days of perfusion with cell-free medium, the resulting constructs were either treated enzymatically to extract the cells or implanted in nude mice as described below.

Cell Extraction from the Constructs
After culture in the 3D perfusion system, cells were extracted from the generated constructs by perfusing a solution of 0.3% collagenase type 2 for 45 minutes followed by perfusion with 0.05% trypsin/0.53 mM EDTA (Gibco) for 15 minutes, both at a velocity of 6 ml/minute. Extracted cells were collected in culture medium, counted by using a Neubauer chamber, and characterized by flow cytometry as described below. This technique was previously shown to result in a >95% extraction of the cells included in the scaffolds, as assessed by DNA quantification of the enzymatically treated constructs [17].

Flow Cytometry Analysis
Suspensions of SVF, 2D-, or 3D-expanded cells were incubated for 30 minutes at 4°C with fluorochrome-conjugated antibodies against the indicated proteins or an isotype control. The antibodies used were from (a) Becton, Dickinson and Company (Franklin Lakes, NJ, http://www.bd.com): CD44-FITC (catalog number 347943), CD90-FITC (catalog number 555595), CD45-FITC/CD34-PE (catalog number 341071), and CD31-PE (catalog number 555446); (b) Miltenyi Biotec (Bergisch Gladbach, Germany, http://www.miltenyibiotec.com): CD133-APC (catalog number 120-001-123); (c) Serotec Ltd. (Oxford, U.K., http://www.serotec.com): CD105-FITC (catalog number MCA1557F); or (d) R&D Systems Inc. (Minneapolis, http://www.rndsystems.com): CD144-PE (VE-cadherin, catalog number FAB9381P) and VEGFR-2-PE (catalog number FAB357P). Cells were washed and resuspended in PBS and analyzed with a FACSCalibur flow cytometer.

Assessment of Bone Formation Following In Vivo Implantation
The osteogenicity of scaffold-cell constructs was assessed by ectopic implantation in nude mice (CD-1 nu/nu, 1-month old; Charles River Laboratories, Wilmington, MA, http://www.criver.com) in accordance with institutional guidelines. Constructs were generated starting from 3 x 10⁶ SVF cells by (a) 3D perfusion culture for 5 days directly within ceramic scaffolds, (b) 2D expansion for 5 days and static loading into the scaffolds of the resulting progeny, or (c) immediate static loading into the scaffolds of the freshly harvested cells, with no further culture. We previously reported that the fraction of cells retained in the scaffolds after seeding by static loading was similar to that obtained using the described perfusion device (i.e., 70%–85%), although cells seeded statically were less uniformly distributed [21]. For each experiment (i.e., using cells from each of five different donors) and experimental group, duplicate constructs were implanted in two different mice.

Eight weeks after implantation, mice were sacrificed, and the constructs were harvested and fixed overnight in 4% formalin, completely decalcified by incubation with Osteodec (Bio-Optica, Milan, Italy, http://www.bio-optica.it) for 3 hours at 37°C under gentle mixing onto an orbital shaker, paraffin embedded, and sectioned at different levels (7-µm-thick sections). Sections were then stained by hematoxylin and eosin (H&E) and observed microscopically for the formation of bone tissue. The amount of bone tissue normalized to the total available pore space was quantified as previously described [22]. For polarized light microscopy, H&E-stained sections were analyzed under transmitted light with crossed polarizers by using a petrographic Leica DMRX microscope (Heerbrugg, Switzerland, http://www.leica.com). Immunostaining for bone sialoprotein (BSP) was performed with an anti-human BSP antibody (catalog number A4232.1/A4232.2; Immundiagnostik AG, Bensheim, Germany, http://www.immundiagnostik.com) followed by incubation with ABC-alkaline phosphatase complex kit (Dako, Glostrup, Denmark, http://www.dako.com) and counterstaining with H&E. Some sections were also stained with Safranin O to assess the formation of cartilaginous tissue.

Assessment of Human Blood Vessels Following In Vivo Implantation
The presence of blood vessels of human origin was determined in 7-µm-thick paraffin-embedded sections of the same constructs described above 8 weeks after implantation in nude mice. Human endothelial cells were specifically stained with anti-human CD31 (Dako) and biotin-conjugated anti-human CD34 (Chemicon, Temecula, CA, http://www.chemicon.com) antibodies. For the anti-CD31 antibody, the M.O.M. Kit from Vector Laboratories (Burlingame, CA, http://www.vectorlabs.com) was used to reduce the unspecific binding of the secondary biotin-conjugated anti-mouse antibody on the sections. After incubation with ABC-alkaline phosphatase complex, specific staining was revealed by using Fast Red (Dako). Sections were counterstained with hematoxylin and mounted.

	RESULTS

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Characterization of Adipose Tissue-Derived Cells In Vitro
Following the digestion of adipose tissue and subsequent centrifugation to remove the differentiated adipocytes, 5.9 ± 3.5 x 10⁵ cells per milliliter of lipoaspirate were obtained, of which 5.2% ± 0.9% were clonogenic, as demonstrated by CFU-f assays (n = 7 donors). The proliferation rate of ATSC during monolayer expansion in tissue culture plastic after the first passage averaged 0.92 ± 0.38 cell doublings per day (n = 3 donors) and was in the range of that typically measured for human bone marrow stromal cells (data not shown). Freshly isolated adipose cells from the SVF were able to deposit abundant mineralized matrix when cultured for 3 weeks in osteogenic medium, as demonstrated by a strong alizarin red and von Kossa staining (Fig. 1A). No changes in the capacity to deposit mineralized matrix were observed in cells expanded for up to three passages (data not shown). SVF cells were also efficiently differentiated toward the adipogenic lineage, as indicated by the presence of lipid droplets in the cytoplasm following culture in adipogenic medium (Fig. 1B).

View larger version (47K): [in this window] [in a new window]   Figure 1. Characterization of adipose tissue-derived cells. (A): Alizarin red and von Kossa staining of freshly isolated SVF cells cultured with osteogenic medium for 3 weeks. (B): Phase contrast microscopic picture of SVF cells cultured with adipogenic differentiation supplements for 3 weeks. (C): Number of cells after 5 days of culture, starting from 3 x 106 freshly isolated SVF nucleated cells expanded either in monolayer (2D) culture or by 3D perfusion within the pores of a hydroxyapatite ceramic scaffold. (D): Flow cytometry analysis for the indicated surface markers of SVF cells, freshly isolated or after 5 days of either 2D or 3D culture. Results are given as % of positive cells ± standard deviation in the indicated number of independent experiments (n = 2 to n = 6). (E): Flow cytometry histograms of freshly isolated SVF cells from one representative experiment showing, on the left, the CD34+/CD90+ mesenchymal cells and, on the right, the CD34+/CD31+ endothelial-lineage cells. Abbreviations: 2D, two dimensional; 3D, three dimensional; SVF, stromal vascular fraction.

Assessment of In Vivo Formation of Bone Tissue by Adipose Tissue-Derived Cells
Cell-ceramic constructs generated starting from the same number of SVF cells, either directly cultured under 3D perfusion or loaded into the scaffolds following 2D expansion for the same time (i.e., 5 days), were subcutaneously implanted in nude mice for 8 weeks in order to determine their osteogenic and vasculogenic capacity (n = 5 donors). For four of the five donors, constructs generated by 3D cultures displayed formation of bone tissue, ranging between 5% and 27% of the available pore space, organized in structures typically referred to as "ossicles," formed starting from the ceramic surface of the pores (Fig. 2A) and including vascular elements (Fig. 2B). Instead, all constructs generated by 2D-expanded cells contained loose interstitial tissue, with no evidence of frank bone tissue formation (Fig. 2C, 2D). Similarly, when SVF cells were loaded onto the scaffold directly after isolation and implanted without culture, only a loose connective tissue was observed (data not shown, n = 3 donors). Safranin O staining (Fig. 2E) of the explanted constructs was negative, indicating that cells in the constructs did not acquire a chondrogenic phenotype, and that bone formation likely did not follow an endochondral ossification process. The newly formed ossicles were positively stained for bone sialoprotein (Fig. 2F) and under polarized light microscopy displayed a moderately dense network of collagen fibrils, organized in partially stratified bundles, characteristic of a rather immature bone tissue (Fig. 2G, 2H).

View larger version (90K): [in this window] [in a new window]   Figure 2. In vivo bone formation by adipose tissue-derived cells. Representative light microscopy pictures of hematoxylin and eosin-stained sections of constructs ectopically implanted for 8 weeks in nude mice and generated by three-dimensional (3D) perfusion culture of stromal vascular fraction (SVF) cells in ceramic scaffolds for 5 days (A, B) or by static loading in the same scaffolds of SVF cells expanded in two-dimensional culture for 5 days (C, D). Bone tissue in explants generated by 3D perfusion culture is also displayed following Safranin O staining (E), following immunostaining for bone sialoprotein (F) or under polarized light (G, H). Abbreviations: b, bone matrix; b.v., blood vessel; f.t., fibrous tissue.

View larger version (164K): [in this window] [in a new window]   Figure 3. In vivo formation of human blood vessels by three-dimensional (3D) cultured adipose-derived cells. Representative light microscopy pictures of sections of constructs ectopically implanted for 8 weeks in nude mice and generated by 3D perfusion culture of stromal vascular fraction cells in ceramic scaffolds for 5 days. Sections are stained with an anti-human CD34 (B–D) or anti-human CD31 (F) antibody or with the corresponding isotype controls (A, E).

View larger version (75K): [in this window] [in a new window]   Figure 4. In vivo formation of human blood vessels by fresh or two-dimensional (2D) expanded adipose-derived cells. Representative light microscopy pictures of sections of constructs ectopically implanted for 8 weeks in nude mice and generated by static loading into ceramic-based scaffolds of stromal vascular fraction cells expanded in 2D culture for 5 days (A) or freshly harvested (B). Sections are stained with an anti-human CD34 antibody.

	DISCUSSION

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A suitable cell source for bone tissue engineering applications requires the availability of a large number of functional cells under minimally invasive harvest conditions. In this regard, assuming that a clonogenic mesenchymal cell (CFU-f) represents the functional unit for generation of bone tissue [23], our results indicate that the quantity of CFU-f that can be derived from 1 ml of human adipose tissue (i.e., approximately 30,000) is approximately 50–100-fold higher than the number of CFU-f typically available from 1 ml of human bone marrow (considering a yield of 3–6 million nucleated cells after Ficoll-gradient isolation and a clonogenicity of 0.01% [17]). Moreover, since harvesting of more than 200 ml of marrow aspirate is challenging without increasing the related morbidity, whereas up to 1 l of lipoaspirate can be collected with a minimal morbidity, the quantity of CFU-f available for clinical applications could be up to 500-fold higher using adipose tissue as compared with bone marrow as a cell source. Obviously, an equivalence in osteogenic function between bone marrow- and adipose tissue-derived CFU-f remains to be demonstrated. However, the finding that only 5 days of preculture, corresponding to a minimal extent of cell expansion, were sufficient to obtain osteogenic grafts starting from SVF cells is an indication of the importance of the initially high amount of CFU-f available from adipose tissue.

Our result showing that adipose tissue-derived cells expanded in 2D were not capable to generate bone tissue upon ectopic implantation is consistent with previous studies proposing preculture in osteogenic medium [12] or transfection with BMP-2 [13] as possible strategies to use human lipoaspirate cells for bone repair. Interestingly, even constructs immediately implanted following scaffold seeding with SVF cells were not osteogenic, whereas 5 days of 3D perfusion culture were sufficient to achieve reproducible osteogenic properties. Since during the 5 days of 3D perfusion culture a limited extent of expansion was achieved, this result suggests the possibility of a specific osteogenic commitment imparted by the exposure to fluid-induced shear [24], by the integrin-mediated signaling triggered by a mineralized substrate [25], by the establishment of 3D cell-cell interactions [26], or by a combination of these factors.

The SVF of human adipose tissue is known to comprise highly heterogeneous cell populations, including adipose stromal (mesenchymal) cells, blood derived cells, vascular (endothelial) cells [27], smooth muscle cells and pericytes [28], and most likely several other yet unidentified cell types. Our phenotypic analysis of SVF cells is in general accordance with previous studies [29, 30], although with some differences in the proportions of the different cell types, which indeed are known to greatly vary from patient to patient and with parameters such as duration of collagenase digestion and donor site or storage duration [31–33]. A precise identification of all different cell types generated after 2D and 3D culture was beyond the scope of the present study, and the analyses and interpretations were restricted to the two cell populations of interest, namely the stromal mesenchymal/osteoprogenitor cells (characterized by the expression of CD90, CD105, and CD44) and the cells of the endothelial lineage. With regard to the latter cell population, our results indicate that they are neither frank endothelial progenitor cells, which are described to express not only CD34 and CD31 but also CD133 and VEGFR2 [34], nor fully differentiated endothelial cells, which typically express VE-cadherin (CD144). It is possible that the CD34⁺/CD31⁺ cells that we define as "endothelial-lineage cells" are a population of endothelial progenitors, since they were functional to generate the endothelial layer of the human blood vessels in vivo, but possibly more committed than those described to be present in peripheral blood [34].

During skeletal development and regeneration, interactions between endothelial cells (EC) and osteoblasts are thought to play a key role [35]. Although transplanted EC have been reported to enhance orthotopic bone formation by bone marrow stromal cells in vivo [36], in vitro studies about the influence of endothelial cells on osteoblastic differentiation have been so far controversial [37–40]. In this context, our perfusion-based culture system represents a relevant model to study the 3D interactions between endothelial and osteoblastic cells as well as to investigate the possibility that mutual conditioning of the two cell populations, including a precise control of their self-renewing and differentiation capacity, requires the formation of an appropriate niche.

	CONCLUSION

Top Abstract Introduction Materials and Methods Results Discussion Conclusion Disclosures of Potential... Acknowledgments References

 
In conclusion, our study provides a prospective for the streamlined manufacturing of osteogenic and vasculogenic grafts in 3D perfusion systems starting from a single cell source, namely adipose tissue cells, and bypassing the constraining 2D culture step, which is typically used in tissue engineering approaches. Since cells from the SVF of adipose tissue are being proposed for the regeneration of other tissues with stringent vascularization requirements (e.g., cardiac [41] and skeletal muscle [42]), the approach may be extended to other areas of regenerative medicine. It remains to be tested whether in larger, clinically relevant-sized constructs the coculture of mesenchymal and endothelial progenitors will accelerate vascularization inside the implanted graft and thereby enhance cell survival and improve engraftment.

	DISCLOSURES OF POTENTIAL CONFLICTS OF INTEREST

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The authors indicate no potential conflicts of interest.

	ACKNOWLEDGMENTS

Top Abstract Introduction Materials and Methods Results Discussion Conclusion Disclosures of Potential... Acknowledgments References

 
We are grateful to Dr. I. Farhadi (Schwerzenbach, Switzerland) for the kind supply of lipoaspirates, to Dr. David Wendt for the support in 3D perfusion cultures and manuscript proofreading, to Silvia Francioli for performing the human bone sialoprotein immunostaining, to Roberta Martinetti (Fin-Ceramica Faenza S.p.a, Italy) for the supply of hydroxyapatite scaffolds, to Dr. Franz Leander (Geological Institute, Basel, Switzerland) for support with polarized light microscopy, and to Paolo Bianco for helpful discussions. We acknowledge the European Union for financial support (European project STEPS, FP6, contract number NMP3-CT-2005-500465, http://www.stepsproject.com).

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