Published online before print May 14, 2009, doi: 10.1161/CIRCRESAHA.108.190926
| |||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
© 2009 American Heart Association, Inc.
Integrative Physiology |
Robust Functional Vascular Network Formation In Vivo by Cooperation of Adipose Progenitor and Endothelial Cells
From the Department of Medicine (D.O.T., S.M.-C., A.R.S., B.H.J., K.L.M.); Indiana Center for Vascular Biology and Medicine (D.O.T., D.N.P., S.M.-C., A.R.S., M.R.S., M.M., B.H.J., D.A.I., K.L.M.); Department of Pediatrics (D.N.P., D.A.I.), Herman B. Wells Center for Pediatric Research; Department of Surgery (M.R.S., M.M.); Department of Biochemistry and Molecular Biology (D.A.I.); Department of Cellular and Integrative Physiology (K.L.M.), Indiana University School of Medicine; and R. L. Roudebush Veterans Affairs Medical Center (K.L.M.), Indianapolis, Ind.
Correspondence to Keith L. March, MD, PhD, Indiana Center for Vascular Biology, Indiana University, School of Medicine, 975 W Walnut St, IB441, Indianapolis, IN 46202. E-mail kmarch{at}iupui.edu
 |
Abstract |
|---|
 Top Abstract IntroductionMaterials and MethodsResultsDiscussionReferences |
|---|
Rapid induction and maintenance of blood flow through new vascular networks is essential for successfully treating ischemic tissues and maintaining function of engineered neo-organs. We have previously shown that human endothelial progenitor cells (EPCs) form functioning vessels in mice, but these are limited in number and persistence; and also that human adipose stromal cells (ASCs) are multipotent cells with pericytic properties which can stabilize vascular assembly in vitro. In this study, we tested whether ASCs would cooperate with EPCs to coassemble vessels in in vivo implants. Collagen implants containing EPCs, ASCs, or a 4:1 mixture of both were placed subcutaneously into NOD/SCID mice. After a range of time periods, constructs were explanted and evaluated with regard to vascular network assembly and cell fate; and heterotypic cell interactions were explored by targeted molecular perturbations. The density and complexity of vascular networks formed by the synergistic dual-cell system was many-fold higher than found in implants containing either ASCs or EPCs alone. Coimplantation of ASCs and EPCs with either pancreatic islets or adipocytes produced neoorgans populated by these parenchymal cells, as well as by chimeric human vessels conducting flow. This study is the first to demonstrate prompt and consistent assembly of a vascular network by human ASCs and endothelial cells and vascularization by these cells of parenchymal cells in implants. Mixture of these 2 readily available, nontransformed human cell types provides a practical approach to tissue engineering, therapeutic revascularization, and in vivo studies of human vasculogenesis.
Key Words: adipose stromal cells • vasculogenesis • tissue engineering • regenerative medicine • vascular networks
|
Introduction |
|---|
|
TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
|---|
Assembly of new vascular networks for therapeutic purposes has been critical but elusive for tissue engineering and angiogenesis. A general requirement for preserving viability within a regenerating region is that a vascular bed is assembled or expanded sufficiently to ensure adequate tissue perfusion. Also important to success of such applications is the ability of any network to anastomose or inosculate promptly with the vessels of immediately adjacent tissues. Cell-based revascularization therapies were tested in patients with various ischemic diseases. These studies used progenitor cells from bone marrow1,2 and skeletal muscle3,4 to treat patients with myocardial infarction,1,2 heart failure,3–6 and peripheral vascular disease.7,8 Despite accumulating data and recent metaanalyses9,10 that support the hypothesis that progenitor cells have high potential for promoting tissue revascularization and functional recovery, invasive methods of cells harvest and their low abundance may limit adoption some of progenitor cells for therapies. Recent approaches were focused on evaluating cells derived from more available tissues including adipose tissue11,12 and umbilical cord blood.13,14
Endothelial progenitor cells (EPCs) isolated from adult peripheral blood,15 bone marrow, umbilical cord blood,13 and vessel wall16 were intensively studied over the past decade. Recently, we have shown that umbilical cord blood contains a population of EPCs with particularly high proliferative potential, which we have termed endothelial colony forming cell.13 Implantation of endothelial colony forming cells in a matrix in mice resulted in in vivo functional vessels formation.17 Although the presence of blood cells within capillary networks formed by such human EPCs confirmed anastomoses with host vasculature, the neovessels were limited in frequency and size. This extended a prior observation for implants containing untransformed adult endothelial cells (ECs), which yielded vessels characterized as narrow-caliber with single-layer walls.18 With nontransformed ECs, failure to establish stable, mature vasculature may be attributable to prolonged absence of a stabilizing layer of mural cells such as pericytes or smooth muscle cells (SMCs). Although EPCs secrete multiple angiogenic factors, conditions within the gel matrix may not attract sufficient host mural cells within an appropriate time frame to promote stability of neovasculature before competing forces act to disassemble the vessels. Although coimplantation of pericytes with EPCs might promote stabilization and maturation of the vessels, an adequate and easily-accessible source of pericytes has not been previously recognized. However, network stabilization by mural cells has been shown by the coimplantation of ECs with human saphenous vein-derived SMCs19 and murine C3H-10T1/2 cells.20 Unfortunately, obtaining the former requires excision of a vein, whereas the latter is not appropriate for therapies.
We have recently discovered that adipose stromal cells (ASCs), a population of pluripotent mesenchymal cells that are readily available in large numbers from adipose tissue, are predominantly associated with vessels in vivo, and phenotypically and functionally equivalent to pericytes associated with microvessels.21 Recognition of this ASC function provides a new context for understanding the observations that local or systemic ASC delivery restores blood flow in ischemic muscle tissues,22–24 an activity that has been proposed to occur via several mechanisms, including paracrine support24,25 of ECs with suppression of their apoptosis22,23 and ASC integration into host vasculature.23 Our previous finding that ASC possess properties of pericytes and are able to stabilize endothelial networks in vitro21 suggested the possibility that provision of ASCs along with ECs in vivo might promote and stabilize new vessel formation, thus potentially providing an approach to enhance development of vascular networks.
|
Materials and Methods |
|---|
|
TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
|---|
An expanded version of Materials and Methods section is available in online data supplement at http://circres.ahajournals.org and includes detailed methods for the following: isolation and cultivation of EPCs from umbilical cord blood and ASCs, formation and implantation of collagen/fibronectin gels containing embedded cells, fabrication of artificial tissues carrying adipocytes and islets, implantation of cells into mouse ears, isolation of porcine islets, evaluation of vascular endothelial growth factor (VEGF) accumulation in ASC-conditioned media, immunohistochemical evaluation of cellular implants and mouse ear tissue, proliferation and apoptosis assays, and statistical analysis.
|
Results |
|---|
|
TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
|---|
Vasculogenesis by Human Primary ASCs and EPCs
We previously showed in vitro that cocultures of ASCs and ECs on Matrigel in a ratio of 1:3 to 1:7 ASC:EC produced a stable vascular network with inner layers formed by ECs and outer layers formed by ASCs.21 Based on this observation, in vivo experiments in the present study were conducted with a ratio of 1:4 (ASC:EPC).
To evaluate the potential for ASCs to assist in vessel formation by ECs and stabilization of neovasculature, we conducted studies with collagen/fibronectin matrix containing: (1) EPCs, (2) ASCs, or (3) a 1:4 mixture of ASCs to EPCs. A clear difference was found in the appearance of the implants harvested from mice at 2 weeks after implantation (Figure 1a through 1c). Whereas implants containing EPCs or ASCs alone were whitish in color with superficial, thin vascular structures, matrices containing the combination of the 2 cell types were consistently red because of the presence of blood-filled vessels. Additionally, it was observed that implants containing both cell types were tightly associated with the muscle fascia, whereas implants with either ASCs or EPCs were loosely attached to host tissue.
|
The visible differences in blood content of implants with human ASCs and EPCs indicated that this combination formed an extensive network of vessels that connected with the host vasculature. Microscopic examination of implant sections stained with hematoxylin/eosin (Figure 1d through 1f) was used to identify vessels as luminal structures that were further classified according to their size, presence of single or multiple layers of cells in the vascular wall, and the presence or absence of contained blood elements. Analysis of implants combine from all independent studies revealed that among implants containing only EPCs: 20% had at least 1 multilayered vessel, 40% had only single-layer vessels, and the remaining 40% had no vessels. Among implants containing only ASCs, none of the implants contained complex multilayered vessels, 30% contained small simple vessels, and 70% possessed no visible vessels. Remarkably, all implants containing both cell types contained numerous vessels comprised of an endothelial layer surrounded by a layer of mural cells, with connections to the host vasculature evidenced by the presence of erythrocytes within the lumens (Figure 1f).
Vessel density in the implants was further assessed by staining section for human ECs (CD31/PECAM) and smooth muscle cells (
-SMA) antigens (Figure 2a through 2f and Online Figure I, a and b). Vessels characterized as complete circular structures possessing distinct lumina and formed by human ECs or cells staining for -SMA were quantified (Figure 2g and 2
h). EPC-containing implants gave rise to 26.6±5.8 CD31+ and 13.1±3.6 -SMA+ vessels per millimeter squared, the latter indicating that host mural cells invaded the implants and contributed to vessel formation. ASC implants possessed 10.2±3.5 -SMA+ vessels per millimeter squared, which were presumably derived from the input human ASCs. Vessels containing human CD31-expressing cells were not detected in any of the implants containing only ASCs, indicating that the observed vessels either incorporated host ECs or were pseudovessels formed by ASCs but lacking an endothelial layer. By comparison to these groups, Mix implants contained remarkably more vessels as enumerated by both CD31 (122.4±9.8 vessels per millimeter squared) and -SMA (124.7±19.7 vessels per millimeter squared) staining (P<0.001). The observed similarity in density of CD31+ and -SMA+ vessels detected in implants formed by the ASC-EPC combination led to hypothesis of routine joint participation of both cell types in the neovessel formation. Analysis of the vascular networks with respect to vessel diameter revealed that the dual cell implants gave rise to a broader distribution of vascular dimension, compared with implants containing only EPCs (Online Figure I, c).
|
To confirm that mixed implants formed multilayered vessels, sections were double-stained with antibodies directed against the endothelial marker CD31 and against the ASC/mural cell marker -SMA. Analysis of sections by confocal immunofluorescence technique (Figure 3a through 3d) confirmed the presence of bilaminar vessels with an inner layer formed by donor human EPCs surrounded by an outer layer of -SMA+ cells (presumably ASCs). Moreover, the presence of autofluorescent erythrocytes (green) in the lumen was apparent.
|
To test the origin of the mural layer of the newly formed vessels, experiments were conducted in which ASCs transduced with lentiviral vectors encoding green fluorescent protein (GFP) were coembedded with EPCs and implanted into mice. Immunodetection of GFP at day 14 revealed that vessels were routinely coated by GFP-expressing ASCs (Figure 3e and 3f), confirming human donor origin of the mural cells of the assembled vessels. Also, analysis of the sections probed for GFP and -SMA antigens confirmed that the majority of ASCs in implants robustly expressed -SMA (Figure 3g through 3j).
Donor-Derived Neovascular Networks Link to Host Vasculature
It is apparent from the above data that ASCs and EPCs in the matrix operate in concert to assemble a vascular network with a range of diameters in these implants. To determine the degree to which these vessels inosculated with the host vasculature, the CD31-positive vessels that clearly contained erythrocytes were scored at 14 days postimplantation (Figure 4a). In the implants containing solely EPCs, 3% of the total vessels detected contained erythrocytes, whereas none was observed in ASC implants. Conversely, nearly 75% (92.5±16.2 per millimeter squared) of CD31+ vessels observed in Mix implants were functional and blood-filled, demonstrating connections with host (mouse) vasculature and incorporation into the circulatory system. To support our hypothesis that the neovessels were functional, implants were imaged by ultrasound following IV injection of microbubbles into host mice. These dynamic images revealed multiple functional vessels traversing the implant region (see Online Movie). To determine whether the cord blood source of EPC was critical for the vascular assembly process and inosculation with the host vasculature, implants containing ASC in combination with ECs either derived from human umbilical vein (HUVEC), human placenta (Pl-EC), or adult human adipose tissue (AT-EC), were evaluated. The density of CD31+ vessels containing erythrocytes in implants containing ASC+HUVEC, ASC+Pl-EC, or ASC+AT-EC was 33.3±12.6, 31.5±9.0, and 61.1±1.0 vessels per millimeter squared, respectively (Online Figure II), confirming that ECs from several sources were able to partner with ASCs in vascular network assembly.
|
The dynamics of vessel formation by the combination of cells in vivo were evaluated in implants harvested at 2, 4, and 6 days after placement. At day 2 following implantation, ECs had assembled into tubes, which had not formed apparent connections with host vasculature (Figure 4b and 4e). By day 4, a significant number of newly formed vessels (100.1±8.1 per millimeter squared) were filled with erythrocytes (Figure 4c and 4e) followed by an additional increase by 45% by day 6 (147.5±22.1 per millimeter squared; Figure 4d and 4e). Moreover, the vessels had formed branching networks throughout the implants (Figure 4d). Thus, the cooperative formation of vessels by ASCs and EPCs occurs quickly in vivo and is followed by connection with the host vasculature.
Cooperative Vasculogenesis by ASCs and EPCs Is Associated With Cell Proliferation and ASC-Mediated Reduction of EPC Apoptosis
5-Bromodeoxyuridine (BrdUrd) labeling was used to determine the cycling status of cells comprising vessels within the matrices containing both cell types. Probing sections against BrdUrd revealed that a significant number of cells underwent DNA synthesis during the first 6 days following matrix insertion and were identified throughout the implants, with numerous BrdUrd-positive cells located in vessel walls (Figure 5a) in both the luminal (EPCs) and abluminal layer (ASCs). To further confirm the identity and location of cycling cells, sections were simultaneously probed for Ki67 and either CD31 or -SMA antigens. As shown in Figure 5e and 5i, Ki67 expression colocalized with both human CD31-positive luminal cells (EPCs) and -SMA–positive ASCs contributing to the vascular mural layer.
|
Because we previously observed that implants containing solely EPCs formed transient vessels, we next determined whether ASCs served to prevent vessel regression by affecting apoptosis of ECs. Matrices containing ASCs and EPCs alone or both were analyzed for apoptotic cells by TUNEL staining at day 14 postimplantation (Online Figure III, b). Many apoptotic cells were observed in matrices implanted with only EPCs. Conversely, implants with only ASCs had few apoptotic cells and importantly, apoptosis was suppressed to very low levels in combination implants.
Crucial Role for Platelet-Derived Growth Factor in Cooperative Vessel Assembly by ASCs and EPCs
We previously described in vitro interaction of ASCs and ECs, accompanied by secretion of complementary growth factors, including platelet-derived growth factor (PDGF)-BB by ECs and VEGF by ASCs.21 To determine whether the in vivo process of vasculogenesis conducted by the combined cells depended on signaling by PDGF-BB, gels were implanted with the addition of either control or anti-PDGF-BB neutralizing antibodies. Staining the sections for CD31 (Figure 6a and 6b) and -SMA (Figure 6c and 6d) revealed the specific disruption of vascular assembly by antagonism of PDGF-BB, whereas both ASCs and ECs survive within these gels to the same extend based on the TUNEL staining (Online Figure IV), their assembly into lumen-containing structures is notably absent in the presence of anti–PDGF-BB antibodies.
|
0.01). f, Analysis of VEGF accumulation in media of ASCs (24 hours) exposed to basal media, EPC-conditioned media (EPC_CM), or EPC_CM media pretreated with either 1 µg/mL anti–PDGF-BB or control goat IgG. ||P
To evaluate for bidirectional signaling between EPCs and ASCs, we determined the effect of PDGF-BB, a factor secreted by EPCs, on ASC secretion of VEGF, in turn a key factor for EPCs physiology. PDGF-BB exposure upregulated ASC secretion of VEGF by 2.4-fold (Figure 6e). At the same time, EPC-conditioned media upregulated VEGF secretion by ASCs by more than 9-fold in comparison with control media (EBM-2/5% FBS), whereas preincubation of EPC-conditioned media with PDGF-BB neutralizing IgGs partially (
50%) suppressed this effect (Figure 6f). Neither EBM-2/5% FBS nor EPC-conditioned media revealed any signal for hVEGF protein.
Vasculogenesis by ASCs and EPCs Without Exogenous Matrix
To test the ability of directly injected ASCs and EPCs to establish vasculogenesis in host tissue, and the necessity of exogenous collagen/fibronectin matrix for this process, we injected distal ear pinnae of mice with the cell mixture after suspension in either collagen/fibronectin gel (n=4) or media (n=4). Fourteen days postdelivery, immunohistochemical analysis of ear sections revealed human CD31-positive, functional (RBC-filled) vessels in both treatment groups (Figure 7a and 7b).
|
Collagen Implants Carrying Adipocytes and Islets
To evaluate the potential of ASCs and EPCs to establish a functional vascular network in the context of an admixed parenchymal or secretory cell population, collagen/fibronectin gels were fabricated carrying ASCs and EPCs in a 1:4 ratio, combined with either freshly isolated mature human adipocytes or pig pancreatic islets. Immunohistochemical analysis of these constructs harvested after 2 weeks revealed that implants carrying adipocytes had typical morphology of vascularized adipose tissue (Figure 8a and 8b), whereas implants carrying islets contained vascular networks with clusters of cells staining positive for insulin (Figure 8d through 8f). Quantitation of vessels in implants containing adipocytes revealed a density of 86.2±2.9 vessels per millimeter squared, and in implants with islets, a density of 38.6±2.9 vessels per millimeter squared.
|
|
Discussion |
|---|
|
TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
|---|
Engineering of tissue constructs with thickness greater than accommodated by gas or nutrient diffusion will require practical means for provision of vascular components that invest constructs and provide blood flow promptly at implantation. Additionally, local augmentation of vascular network development has been a goal for therapy of disorders such as myocardial infarction and peripheral vascular diseases. This study demonstrates for the first time the ability of 2 readily available, genetically unmodified primary human cell types, ASCs and ECs isolated from umbilical cord blood and from umbilical vein, placenta, and adult adipose tissue, to synergize in de novo formation of vascular networks in implanted collagen matrices, as well as in the context of admixed parenchymal nonvascular cells. Important to potential utility of this approach is the fact that ASCs and EPCs represent cells that can be obtained from humans with minimal difficulty and can be rapidly amplified after initial isolation if required.13,26
Previous studies demonstrated that human adult or umbilical cord blood–derived ECs alone after embedding into the matrices developed low-caliber vessel networks with limited persistence. However, their coimplantation with adult human SMCs27 or C3H10T1/2 cells bearing characteristics of SMCs20 resulted in vascular networks composed of vessels with greater caliber and thicker walls. Our recent finding that ASCs are expressing markers (CD140a, CD140b, NG2) and physiological characteristics of pericytes21 prompted in vivo evaluation of ASC and EC interaction. The present study confirmed previous observations that implants comprised only of EPCs demonstrate limited vascular network formation. At the same time, combination of EC-ASC in implants led to development remarkably dense and stable vascular network, which supported the main hypothesis of our study that ASC are able to behave as pericytes in vivo. An important effect of ASCs on ECs clearly involves abrogation of marked apoptosis present in implants containing only ECs. This is also consistent with our prior findings that factors released from ASCs can protect ECs from apoptosis,24 as well as stabilize EC cord formation on Matrigel in vitro settings.21 Several molecular mechanisms may be involved in these effects of ASCs on ECs, including the secretion by ASCs of diffusible proangiogenic and antiapoptotic factors (such as VEGF and angiopoietin-1) and direct contact with forming endothelial tubes.
Given this role of ASCs in supporting ECs survival during the process of vasculogenesis, we explored whether ECs play a complementary role in modulating ASCs behavior via factors secreted by ECs. PDGF-BB is a key factor secreted by ECs28 and EPCs (data not shown), for which we have previously demonstrated functional receptors on ASCs and their proliferative response.21 Interruption of vascular assembly by local blockade of PDGF-BB signaling (with neutralizing antibodies) suggested a crucial role for this factor in signaling from EPCs to ASCs during vasculogenesis (Figure 6). We thus hypothesized that blockade of PDGF-BB signaling prevented formation of mature vessels by reducing ASCs migration toward EPCs involved in lumen formation, with consequent diminution of ASC support for EPC, because of either loss of proximity or alteration of the ASCs secretory profile. This concept is supported by our finding that PDGF-BB secreted by EPCs plays a dominant but not solitary role in the stimulation of VEGF secretion by ASC (Figure 6); this in turn is important for EC survival29 and proliferation. This is also consistent with previous findings that exposure of human tenocytes, C3H10T1/2, or human myometrial SMCs to PDGF-BB increases VEGF expression.29–31
Both in the context of an engineered implant and therapeutic augmentation of tissue perfusion, timely provision of functional circulation is essential. Accordingly, it is notable that the implanted cells organized into vessels and established communication with the host circulation by day 4 following implantation in the dual cell system (Figure 4). Analysis of cell cycle revealed active proliferation of the cells in both vascular layers in the implants, suggesting involvement of proliferation in neovessels remodeling (Figure 5). The extent to which input cells are initially capable of expansion following implantation is not clear, but stabilization of the vascular density between days 7 and 14 postimplant in the collagen gels, suggests mechanisms controlling proliferation, concurrently with vascular remodeling in the context of flow.
One potential application of this approach is in fabrication of soft tissue or neoorgans, such as adipose implants for reconstructive surgery, as well as for metabolic or secretory tissues such as liver and pancreas. Although the sufficiency and persistence of the vascular network provided by ASCs and EPCs requires further evaluation, the present experiments (Figure 8) provide an initial demonstration that coimplantation of the cell mixture together with adipocytes or islets can establish neoorgans populated by such parenchymal cells.
The "2-cell system" also provides a means for evaluating the role of matrix in vasculogenesis. Although the collagen/fibronectin matrix was used in most experiments to provide a supportive scaffold, we speculate that cell delivery in a range of matrices may assist both in restricting cells redistribution and augmenting their survival, particularly in ischemic environments, which may be hostile to implanted cells. In addition to delivery within exogenous matrix, results of cell injection into mouse pinnae (Figure 7) show that the mixture of cells is capable of assembly into vascular structures without exogenous extracellular matrix proteins.
The ready availability of ASCs and EPCs from clinically feasible sources, and their simple, well-defined preparation provide attractive features compare with previous approaches. Although use of human SMCs, primarily from dermis,18,32 has been described experimentally, the need for significant expansion because of the limited cell number practically available distinguishes this type of mural cell from ASCs.
Another approach to vascularization of tissue-engineered constructs is suggested by observations that both ASCs and ECs are present within enzymatically dissociated adipose tissue,22–24 although in variable proportions depending on the preparation used.33–35 It has been shown previously and is consistent with the observations of our study (Online Figure III) that these heterogeneous adipose-derived cell mixtures are able to establish functional vasculature when transplanted to in vivo models.36,37 Although these systems are attractive in the use of solely adipose tissue as the source of vascular components,37,38 their utility may be hampered by the variability in viability and composition of the product of tissue harvesting and digestion without additional cell characterization. In addition, it has recently been demonstrated that mature ECs possess decreased ability to partner with mural cells to establish functional vasculature than cord blood–derived cells.19,20
A potential limitation of the clinical application of the approach described in this study is the use of allogeneic EPCs, which may require HLA host/donor matching for implants engineering or tissue revascularization. Our prior description of cord blood as a rich source of high proliferative EPCs13,39 suggests that extensive EPC expansion and banking may provide adequate source of cells as a component of vasculogenic therapeutic mixture for proposed applications. It is intriguing that ASCs were characterized as possessing immunosuppressive properties40; we speculate that factors secreted by ASCs could alter immune response caused by heterologous ECs.
We conclude that the EC-ASC combination provides a clinically practical approach for engineering constructs composed of stable vascular networks and functional parenchymal cellular components that could be used for secretion of endogenous or specifically overexpressed bioactive factors and for augmentation of perfusion of ischemic tissues.
|
Acknowledgments |
|---|
We thank Dr Ken Cornetta for providing pCSCGW-EGFP construct, James Solomon (Indiana University) for technical assistance, and Michael Green and Kelley Nonweiler (Vitacyte) for providing porcine islets.
Sources of Funding
Supported by NIH grant R01 HL77688-01 and VA Merit Review grants (to K.L.M.); NIH grant T32 HL0799905 and American Heart Association Postdoctoral Award 0727777Z (to D.T.); and the Cryptic Masons’ Medical Research Foundation.
Disclosures
Dmitry Traktuev, Brian Johnstone, and Keith March disclose an ownership interest in intellectual property related to findings covered within this manuscript.
|
Footnotes |
|---|
Original received June 20, 2008; resubmission received November 18, 2008; revised resubmission received April 29, 2009; accepted May 4, 2009.
|
References |
|---|
|
TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
|---|
1. Assmus B, Schachinger V, Teupe C, Britten M, Lehmann R, Dobert N, Grunwald F, Aicher A, Urbich C, Martin H, Hoelzer D, Dimmeler S, Zeiher AM. Transplantation of Progenitor Cells and Regeneration Enhancement in Acute Myocardial Infarction (TOPCARE-AMI). Circulation. 2002; 106: 3009–3017.
2. Strauer BE, Brehm M, Zeus T, Kostering M, Hernandez A, Sorg RV, Kogler G, Wernet P. Repair of infarcted myocardium by autologous intracoronary mononuclear bone marrow cell transplantation in humans. Circulation. 2002; 106: 1913–1918.
3. Menasche P, Hagege AA, Scorsin M, Pouzet B, Desnos M, Duboc D, Schwartz K, Vilquin JT, Marolleau JP. Myoblast transplantation for heart failure. Lancet. 2001; 357: 279–280.[CrossRef][Medline] [Order article via Infotrieve]
4. Menasche P, Hagege AA, Vilquin JT, Desnos M, Abergel E, Pouzet B, Bel A, Sarateanu S, Scorsin M, Schwartz K, Bruneval P, Benbunan M, Marolleau JP, Duboc D. Autologous skeletal myoblast transplantation for severe postinfarction left ventricular dysfunction. J Am Coll Cardiol. 2003; 41: 1078–1083.
5. Perin EC, Dohmann HF, Borojevic R, Silva SA, Sousa AL, Mesquita CT, Rossi MI, Carvalho AC, Dutra HS, Dohmann HJ, Silva GV, Belem L, Vivacqua R, Rangel FO, Esporcatte R, Geng YJ, Vaughn WK, Assad JA, Mesquita ET, Willerson JT. Transendocardial, autologous bone marrow cell transplantation for severe, chronic ischemic heart failure. Circulation. 2003; 107: 2294–2302.
6. Fuchs S, Satler LF, Kornowski R, Okubagzi P, Weisz G, Baffour R, Waksman R, Weissman NJ, Cerqueira M, Leon MB, Epstein SE. Catheter-based autologous bone marrow myocardial injection in no-option patients with advanced coronary artery disease: a feasibility study. J Am Coll Cardiol. 2003; 41: 1721–1724.
7. Koshikawa M, Shimodaira S, Yoshioka T, Kasai H, Watanabe N, Wada Y, Seto T, Fukui D, Amano J, Ikeda U. Therapeutic angiogenesis by bone marrow implantation for critical hand ischemia in patients with peripheral arterial disease: a pilot study. Curr Med Res Opin. 2006; 22: 793–798.[CrossRef][Medline] [Order article via Infotrieve]
8. Tateishi-Yuyama E, Matsubara H, Murohara T, Ikeda U, Shintani S, Masaki H, Amano K, Kishimoto Y, Yoshimoto K, Akashi H, Shimada K, Iwasaka T, Imaizumi T. Therapeutic angiogenesis for patients with limb ischaemia by autologous transplantation of bone-marrow cells: a pilot study and a randomised controlled trial. Lancet. 2002; 360: 427–435.[CrossRef][Medline] [Order article via Infotrieve]
9. Jolicoeur EM, Granger CB, Fakunding JL, Mockrin SC, Grant SM, Ellis SG, Weisel RD, Goodell MA. Bringing cardiovascular cell-based therapy to clinical application: perspectives based on a National Heart, Lung, and Blood Institute Cell Therapy Working Group meeting. Am Heart J. 2007; 153: 732–742.[CrossRef][Medline] [Order article via Infotrieve]
10. Abdel-Latif A, Bolli R, Tleyjeh IM, Montori VM, Perin EC, Hornung CA, Zuba-Surma EK, Al-Mallah M, Dawn B. Adult bone marrow-derived cells for cardiac repair: a systematic review and meta-analysis. Arch Intern Med. 2007; 167: 989–997.
11. Zuk PA, Zhu M, Ashjian P, De Ugarte DA, Huang JI, Mizuno H, Alfonso ZC, Fraser JK, Benhaim P, Hedrick MH. Human adipose tissue is a source of multipotent stem cells. Mol Biol Cell. 2002; 13: 4279–4295.
12. Zuk PA, Zhu M, Mizuno H, Huang J, Futrell JW, Katz AJ, Benhaim P, Lorenz HP, Hedrick MH. Multilineage cells from human adipose tissue: implications for cell-based therapies. Tissue Eng. 2001; 7: 211–228.[CrossRef][Medline] [Order article via Infotrieve]
13. Ingram DA, Mead LE, Tanaka H, Meade V, Fenoglio A, Mortell K, Pollok K, Ferkowicz MJ, Gilley D, Yoder MC. Identification of a novel hierarchy of endothelial progenitor cells using human peripheral and umbilical cord blood. Blood. 2004; 104: 2752–2760.
14. Gang EJ, Jeong JA, Hong SH, Hwang SH, Kim SW, Yang IH, Ahn C, Han H, Kim H. Skeletal myogenic differentiation of mesenchymal stem cells isolated from human umbilical cord blood. Stem Cells. 2004; 22: 617–624.[CrossRef][Medline] [Order article via Infotrieve]
15. Asahara T, Murohara T, Sullivan A, Silver M, van der Zee R, Li T, Witzenbichler B, Schatteman G, Isner JM. Isolation of putative progenitor endothelial cells for angiogenesis. Science. 1997; 275: 964–967.
16. Ingram DA, Mead LE, Moore DB, Woodard W, Fenoglio A, Yoder MC. Vessel wall-derived endothelial cells rapidly proliferate because they contain a complete hierarchy of endothelial progenitor cells. Blood. 2005; 105: 2783–2786.
17. Ingram DA, Krier TR, Mead LE, McGuire C, Prater DN, Bhavsar J, Saadatzadeh MR, Bijangi-Vishehsaraei K, Li F, Yoder MC, Haneline LS. Clonogenic endothelial progenitor cells are sensitive to oxidative stress. Stem Cells. 2007; 25: 297–304.[CrossRef][Medline] [Order article via Infotrieve]
18. Schechner JS, Nath AK, Zheng L, Kluger MS, Hughes CC, Sierra-Honigmann MR, Lorber MI, Tellides G, Kashgarian M, Bothwell AL, Pober JS. In vivo formation of complex microvessels lined by human endothelial cells in an immunodeficient mouse. Proc Natl Acad Sci U S A. 2000; 97: 9191–9196.
19. Melero-Martin JM, Khan ZA, Picard A, Wu X, Paruchuri S, Bischoff J. In vivo vasculogenic potential of human blood-derived endothelial progenitor cells. Blood. 2007; 109: 4761–4768.
20. Au P, Daheron LM, Duda DG, Cohen KS, Tyrrell JA, Lanning RM, Fukumura D, Scadden DT, Jain RK. Differential in vivo potential of endothelial progenitor cells from human umbilical cord blood and adult peripheral blood to form functional long-lasting vessels. Blood. 2008; 111: 1302–1305.
21. Traktuev DO, Merfeld-Clauss S, Li J, Kolonin M, Arap W, Pasqualini R, Johnstone BH, March KL. 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.
22. Miranville A, Heeschen C, Sengenes C, Curat CA, Busse R, Bouloumie A. Improvement of postnatal neovascularization by human adipose tissue-derived stem cells. Circulation. 2004; 110: 349–355.
23. Planat-Benard V, Silvestre JS, Cousin B, Andre M, Nibbelink M, Tamarat R, Clergue M, Manneville C, Saillan-Barreau C, Duriez M, Tedgui A, Levy B, Penicaud L, Casteilla L. Plasticity of human adipose lineage cells toward endothelial cells: physiological and therapeutic perspectives. Circulation. 2004; 109: 656–663.
24. Rehman J, Traktuev D, Li J, Merfeld-Clauss S, Temm-Grove CJ, Bovenkerk JE, Pell CL, Johnstone BH, Considine RV, March KL. Secretion of angiogenic and antiapoptotic factors by human adipose stromal cells. Circulation. 2004; 109: 1292–1298.
25. Nakagami H, Maeda K, Morishita R, Iguchi S, Nishikawa T, Takami Y, Kikuchi Y, Saito Y, Tamai K, Ogihara T, Kaneda Y. Novel autologous cell therapy in ischemic limb disease through growth factor secretion by cultured adipose tissue-derived stromal cells. Arterioscler Thromb Vasc Biol. 2005; 25: 2542–2547.
26. Merfeld-Clauss S, Johnstone BH, Ingram DA, March KL. A hierarchy of proliferative cells is observed in populations of non-hematopoietic, CD34+ pluripotent cells isolated from adipose tissue. Adipocytes. 2005; 1: 195.
27. Wu X, Rabkin-Aikawa E, Guleserian KJ, Perry TE, Masuda Y, Sutherland FW, Schoen FJ, Mayer JE Jr, Bischoff J. Tissue-engineered microvessels on three-dimensional biodegradable scaffolds using human endothelial progenitor cells. Am J Physiol. 2004; 287: H480–H487.
28. Aromatario C, Sterpetti AV, Palumbo R, Patrizi AL, Di Carlo A, Proietti P, Guglielmi MB, Cavallaro A, Santoro-D'Angelo L, Cucina A. Fluid shear stress increases the release of platelet derived growth factor BB (PDGF BB) by aortic endothelial cells. Minerva Cardioangiol. 1997; 45: 1–7.[Medline] [Order article via Infotrieve]
29. Reinmuth N, Liu W, Jung YD, Ahmad SA, Shaheen RM, Fan F, Bucana CD, McMahon G, Gallick GE, Ellis LM. Induction of VEGF in perivascular cells defines a potential paracrine mechanism for endothelial cell survival. FASEB J. 2001; 15: 1239–1241.
30. Petersen W, Pufe T, Zantop T, Tillmann B, Mentlein R. Hypoxia and PDGF have a synergistic effect that increases the expression of the angiogenetic peptide vascular endothelial growth factor in Achilles tendon fibroblasts. Arch Orthop Trauma Surg. 2003; 123: 485–488.[CrossRef][Medline] [Order article via Infotrieve]
31. Taniguchi Y, Morita I, Kubota T, Murota S, Aso T. Human uterine myometrial smooth muscle cell proliferation and vascular endothelial growth-factor production in response to platelet-derived growth factor. J Endocrinol. 2001; 169: 79–86.[Abstract]
32. Nor JE, Peters MC, Christensen JB, Sutorik MM, Linn S, Khan MK, Addison CL, Mooney DJ, Polverini PJ. Engineering and characterization of functional human microvessels in immunodeficient mice. Lab Invest. 2001; 81: 453–463.[Medline] [Order article via Infotrieve]
33. Yoshimura K, Shigeura T, Matsumoto D, Sato T, Takaki Y, Aiba-Kojima E, Sato K, Inoue K, Nagase T, Koshima I, Gonda K. Characterization of freshly isolated and cultured cells derived from the fatty and fluid portions of liposuction aspirates. J Cell Physiol. 2006; 208: 64–76.[CrossRef][Medline] [Order article via Infotrieve]
34. Oedayrajsingh-Varma MJ, van Ham SM, Knippenberg M, Helder MN, Klein-Nulend J, Schouten TE, Ritt MJ, van Milligen FJ. Adipose tissue-derived mesenchymal stem cell yield and growth characteristics are affected by the tissue-harvesting procedure. Cytotherapy. 2006; 8: 166–177.[CrossRef][Medline] [Order article via Infotrieve]
35. Astori G, Vignati F, Bardelli S, Tubio M, Gola M, Albertini V, Bambi F, Scali G, Castelli D, Rasini V, Soldati G, Moccetti T. "In vitro" and multicolor phenotypic characterization of cell subpopulations identified in fresh human adipose tissue stromal vascular fraction and in the derived mesenchymal stem cells. J Transl Med. 2007; 5: 55.[CrossRef][Medline] [Order article via Infotrieve]
36. Scherberich A, Galli R, Jaquiery C, Farhadi J, Martin I. Three-dimensional perfusion culture of human adipose tissue-derived endothelial and osteoblastic progenitors generates osteogenic constructs with intrinsic vascularization capacity. Stem Cells. 2007; 25: 1823–1829.[CrossRef][Medline] [Order article via Infotrieve]
37. Hoying JB, Boswell CA, Williams SK. Angiogenic potential of microvessel fragments established in three-dimensional collagen gels. In Vitro Cell Dev Biol. 1996; 32: 409–419.[CrossRef]
38. Shepherd BR, Chen HY, Smith CM, Gruionu G, Williams SK, Hoying JB. Rapid perfusion and network remodeling in a microvascular construct after implantation. Arterioscler Thromb Vasc Biol. 2004; 24: 898–904.
39. Yoder MC, Mead LE, Prater D, Krier TR, Mroueh KN, Li F, Krasich R, Temm CJ, Prchal JT, Ingram DA. Redefining endothelial progenitor cells via clonal analysis and hematopoietic stem/progenitor cell principals. Blood. 2007; 109: 1801–1809.
40. Puissant B, Barreau C, Bourin P, Clavel C, Corre J, Bousquet C, Taureau C, Cousin B, Abbal M, Laharrague P, Penicaud L, Casteilla L, Blancher A. Immunomodulatory effect of human adipose tissue-derived adult stem cells: comparison with bone marrow mesenchymal stem cells. Br J Haematol. 2005; 129: 118–129.[CrossRef][Medline] [Order article via Infotrieve]
Related Article:
Circ. Res. 2009 104: 1330-1332.
This article has been cited by other articles:
 |
|
 |
|
  J. Saif, C. Heeschen, and A. Aicher Add Some Fat to Vascular Progenitor Cell Therapy Circ. Res., June 19, 2009; 104(12): 1330 - 1332. [Full Text] [PDF]
|
|
| |||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||