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World J Stem Cells. Jul 26, 2014; 6(3): 312-321
Published online Jul 26, 2014. doi: 10.4252/wjsc.v6.i3.312
Adipose-derived stem cells: Implications in tissue regeneration
Wakako Tsuji, J Peter Rubin, Kacey G Marra
Wakako Tsuji, J Peter Rubin, Kacey G Marra, Adipose Stem Cell Center, Department of Plastic Surgery, University of Pittsburgh, Pittsburgh, PA 15213, United States
J Peter Rubin, Kacey G Marra, McGowan Institute for Regenerative Medicine, University of Pittsburgh, Pittsburgh, PA 15213, United States
J Peter Rubin, Kacey G Marra, Department of Bioengineering, University of Pittsburgh, Pittsburgh, PA 15213, United States
Author contributions: Tsuji W and Marra KG wrote the manuscript; Rubin JP and Marra KG supervised the paper.
Correspondence to: Kacey G Marra, Associate Professor, Adipose Stem Cell Center, Department of Plastic Surgery, University of Pittsburgh, 200 Lothrop Street, Pittsburgh, PA 15213, United States.
Telephone: +1-412-3838924 Fax: +1-412-3614643
Received: February 23, 2014
Revised: April 16, 2014
Accepted: June 10, 2014
Published online: July 26, 2014


Adipose-derived stem cells (ASCs) are mesenchymal stem cells (MSCs) that are obtained from abundant adipose tissue, adherent on plastic culture flasks, can be expanded in vitro, and have the capacity to differentiate into multiple cell lineages. Unlike bone marrow-derived MSCs, ASCs can be obtained from abundant adipose tissue by a minimally invasive procedure, which results in a high number of cells. Therefore, ASCs are promising for regenerating tissues and organs damaged by injury and diseases. This article reviews the implications of ASCs in tissue regeneration.

Key Words: Mesenchymal stem cells, Adipose-derived stem cells, Differentiation, Growth factors, Tissue engineering, Clinical trials

Core tip: This review article provides an overview on adipose-derived stem cells (ASCs) for implications in tissue regeneration. ASCs are obtained in high yields from abundant adipose tissue in the body and have multi-lineage differentiation ability. This article focuses on ASC characterization, growth factor secretion from ASCs, differentiation ability in vitro and in vivo, and the potential clinical applications.


Mesenchymal stem cells (MSCs) are adult stem cells that were originally identified in bone marrow as multi-potent cells[1,2]. Stem cells are characterized by their self-renewal ability and multi-potency. Bone marrow-derived stem cells are most broadly studied for therapeutic potentials since their discovery in the 1960s[1]. After the discovery of bone marrow-derived MSCs, MSCs have been isolated from nearly every tissue in the body[3], for example, adipose tissue[4], umbilical cord blood[5], peripheral blood[6], dental pulp[7], dermis[8], and amniotic fluid[9], and even in tumors[10]. Adipose-derived stem cells (ASCs) were first identified as MSCs in adipose tissue in 2001[11], and since then adipose tissue has been studied as a cell source for tissue engineering and regenerative medicine. There are multiple terms for stem cells derived from adipose tissue, for example, preadipocytes, adipose-derived stromal cells, processed lipoaspirated cells, adipose-derived mesenchymal stem cells, adipose-derived adult stem cells. In 2004, the consensus was reached the term as ASCs.

There are several types of adipose tissue, with subcutaneous as the most clinically relevant source. ASCs can be isolated from subcutaneous adipose tissue of the abdomen, thigh, and arm. Because adipose tissue is typically abundant in the human body, ASCs can potentially be isolated in high numbers. The multi-lineage capacity of ASCs offers the potential to repair, maintain or enhance various tissues. This review article will focus on source, isolation, and characterization of ASCs, secretion of growth factors from ASCs, in vitro and in vivo differentiation ability of ASCs, and the potential clinical application.


There are mainly two types of adipose tissue: white adipose tissue and brown adipose tissue. They are morphologically and functionally different. Brown adipose tissue much less abundant than white adipose tissue, but can be found in the neck, mediastinum, and interscapular areas in neonates. However, brown adipose tissue undergoes a morphologic transformation with aging. The appearance of brown adipose tissue is literally brown. Brown adipocytes are multilocular and retain small lipid vacuoles compared to white adipocytes. Vascularization is obvious because brown adipose tissue requires much more oxygen consumption compared to other tissues. Brown adipocytes have no known correlation with insulin resistance. The main function of brown adipose tissue is thermogenesis[12,13]. Brown adipose tissue contains a large number of mitochondria and expresses uncoupling protein 1 (UCP1). UCP1 is a brown adipose tissue-specific marker, not expressed within white adipose tissue. UCP1 is expressed in the inner membrane of mitochondria, mainly regulated by adrenergic signaling through sympathetic innervations, and this signaling is responsible for thermogenesis[12,13]. Brown adipose tissue is activated by thyroid hormone, cold temperatures, thiazolidinediones, and activated brown adipose tissue is inversely correlated with body mass index, adipose tissue mass and insulin resistance.

White adipose tissue is found throughout the body, representatively in subcutaneous and visceral adipose tissue. The appearance of white adipose tissue is yellow or ivory. White adipocytes are unilocular and contain large lipid vacuoles. White adipose tissue function is to store excess energy in the form of triglycerides, and its hyperplasia causes obesity and dysfunction of metabolic pathways as insulin resistance. UCP1 is not expressed in white adipocytes but the isoform UCP2 is expressed in parts of white adipocytes.

Recently, beige adipocytes have been discovered within white adipose tissue, especially inguinal white adipose tissue[14]. Beige adipocytes have the characteristics of both brown and white adipocytes. Beige adipocytes contain both unilocular large and multiple small lipid vacuoles. Its function is adaptive thermogenesis. In response to cold temperature exposure, beige cells transform into cells which have brown adipose tissue-like characteristics, such as UCP1 expression and small lipid vacuoles[15]. It is still controversial whether the beige adipocytes arise through the transdifferentiation of white adipocytes or by de novo adipogenesis from a subgroup of precursor cells[16,17].

ASCs isolated from white adipose tissue have different characteristics from those isolated from brown adipose tissue, just as ASCs from different anatomical areas have different characteristics. Subcutaneous tissues are easily obtained via lipoaspiration and usually discarded after the surgery. The lipoaspiration technique does not affect function of ASCs, but the vacuum process does damage mature adipocytes[18].

Zuk et al[4] developed a widely used method for isolating ASCs from white adipose tissue in 2001. Adipose tissues are minced and then undergo enzymatic digestion with collagenase type II. After centrifugation, the resulting pellet is called the stroma vascular fraction (SVF). Approximately 2 to 6 million cells in SVF can be obtained from one milliliter of lipoaspirate[19]. SVF contains ASCs, endothelial cells, endothelial progenitor cells, pericytes, smooth muscle cells, leukocytes, and erythrocytes[20]. ASCs are obtained as the plastic-adherent population after overnight culturing.

Stem cell yield is higher from adipose tissue than bone marrow-both for aspirated and excised adipose tissues. One gram of aspirated adipose tissue yields approximately 3.5 × 105 to 1 × 106 ASCs. This is compared to 5 hundred to 5 × 104 of bone marrow-derived MSCs (BM-MSCs) isolated from one gram of bone marrow aspirate[21]. However, ASC yield from lipoaspirated adipose tissue has been reported to be approximately one half that isolated from whole, excised adipose tissue[22]. ASCs are isolated from the SVF after plating, as ASCs adhere fairly quickly to the surface of tissue culture-treated flasks. ASCs are easily cultured and expanded in vitro; average doubling time of cultured ASC varies between 2 to 5 d, depending on passage number and culture medium[23,24]. ASCs can be easily cryopreserved in a media containing serum and dimethylsulfoxide. Proliferation and differentiation of ASCs are not affected by cryopreservation[25]. The morphology of ASCs is spindle-shaped, very similar to BM-MSCs.

One notable characteristic of ASCs is that they are not homogenous population[11,26]. Many studies have attempted to characterize ASCs using cell surface markers via flow cytometry analysis, but a unique single marker has yet to be identified. ASCs have a positive expression of CD34 at the first passage of culture, but CD34 expression decreases after passaging[20,23]. ASCs express typical mesenchymal markers such as CD13, CD29, CD44, CD63, CD73, CD90, and CD105, and ASCs are negative for hematopoietic antigens such as CD14, CD31, CD45, and CD144[4,23,27]. After culturing and passaging, ASC’s surface markers can change with passaging. The expression of hematopoietic markers such as CD11, CD14, CD34, and CD45 dissipates or are lost[28]. On the other hand, the expression level of CD29, CD73, CD90, and CD166 increase from the SVF to passage 2[23]. Passaging is considered to select cell population with more homogenous cell surface markers compared to SVF.

Further characterization of the heterogeneous ASC population has been recently reported. Li et al[26] categorized four ASC subpopulations: pericytes as CD146+/CD31-/CD34-, mature endothelial cells as CD31+/CD34-, premature endothelial cells as CD31+CD34+, and preadipocytes as CD31-/CD34+. The highest subpopulation was preadipocytes with 67.6%, premature endothelial cell was the second highest subpopulation with 5.2%, and the percentage of pericytes and mature endothelial cells were less than 1%. The cells with CD31-/CD34+ expression demonstrated the greatest proliferation and highest adipogenic differentiation. The localization of ASCs within adipose tissue is not totally clarified yet, but the niche of ASCs is suggested to be in the vasculature of adipose tissue[29]. Histological analysis also suggested that ASCs reside within adipose tissue in a perivascular location[30,31]. Traktuev et al[31] concluded that the location of the ASCs in the vessel is at interface between endothelium and adipocytes, and ASCs have the ability to both support vascular structure and generate adipocytes.


ASCs are considered to be a mediator of tissue regeneration through the secretion of specific soluble factors. ASCs secrete multiple growth factors, including basic fibroblast growth factor (bFGF), vascular endothelial growth factor (VEGF), insulin-like growth factor 1, hepatocyte growth factors (HGF), and transforming growth factor (TGF)-β1[32]. The expression of cytokines renders ASCs promising therapies for transplantation and ischemia patients. Transplanted tissues and organs are exposed to hypoxia soon after transplantation due to a lack of initial vasculature and tend to undergo apoptosis[33]. The levels of VEGF, bFGF, and HGF secreted by ASCs are reported to be upregulated by hypoxia. VEGF was secreted at the concentrations of 70.17 and 200.17 pg/mL under normoxia and hypoxia, respectively. Basic FGF was secreted at the concentrations of 10.62 and 24.75 pg/mL under normoxia and hypoxia, respectively[34]. Similarly, Rehman et al[35] reported that ASCs secreted significant levels of VEGF and HGF under hypoxia, which induced the healing mice hindlimb ischemia. ASCs are considered to be a cell source that induces angiogenesis, which is actually used for human ischemia treatment[36].

In addition to growth factor secretion, ASCs are responsive to growth factors, including enhancing proliferation by bFGF, and platelet-derived growth factor (PDGF). Basic FGF is released from an injured extracellular matrix[37]. PDGF is released from activated platelets on bleeding[38]. When ASCs are exposed growth factors, tissues can be regenerated more effectively. Kaewsuwan et al[39] studied effect of six growth factors on the proliferation of ASCs, and found that PDGF-BB had the highest stimulatory effect at the concentration of 10 ng/mL. PDGF receptors α and β are expressed in ASCs, and PDGF-BB and PDGF receptor β signaling is involved in the stimulation of ASCs[39]. Besides PDGF receptors α and β, ASCs express VEGF, HGF, epidermal growth factor (EGF), and bFGF receptors[40,41]. VEGF increases migration and promotes chondrogenic differentiation[41], and HGF promotes hepatogenic differentiation of ASCs in vitro[42]. EGF inhibits ASC adipogenic differentiation, and bFGF increase ASC proliferation, promotes adipogenic and chondrogenic differentiation, and inhibit osteogenic differentiation in vitro[43-47].

ASCs possess unique paracrine characteristics. ASCs secrete growth factors that stimulate recovery of damaged tissue. Furthermore, ASCs express several kinds of growth factor receptors and are sensitive to growth factors. Therefore, ASCs mediate tissue regeneration.


ASCs can be differentiated into multiple lineages under culturing with specific conditions[11], which results in the potential of ASCs for multiple clinical applications. The induction of ASC differentiation in vitro is achieved by culture with media containing selective lineage-specific induction factors. ASCs have been shown to be differentiated into cells of ectodermal, endodermal and mesodermal origin[4,48,49]. Less controversial is the differentiation of ASCs into adipogenic, chondrogenic, and osteogenic cells, because ASCs are of mesodermal origin. With a combination of morphological observation, immunofluorescence, and polymerase chain reaction (PCR) analysis in vitro, adipogenic, osteogenic, and chondrogenic potentials of ASCs has been reported[4,11]. As mentioned above, MSCs from different anatomical sources demonstrate some differences. ASCs have prominent adipogenic differentiation ability compared to BM-MSCs in vitro[50,51]. BM-MSCs have been shown to have higher osteogeneic differentiation ability compared to ASCs[51-53].

ASCs can be differentiated into adipocytes when cultured in adipogenic differentiation media, which typically contains isobuthyl-methylxanthine, insulin, and indomethacin[54]. ASCs develop multiple lipid droplets about 7 d following exposure to the induction media, and the number of lipid droplets gradually increases. By 2 to 3 wk, the lipid droplets begin to form a unilocular lipid. During differentiation into mature adipocytes, ASCs express several types of extracellular matrix (ECM) proteins, including fibronectin, laminin, and various types of collagen. During adipogenesis, a fibronectin network develops first, and a type-I collagen network is formed last[55]. These ECMs allow ASCs to differentiate into mature adipocytes. ASCs show promise for soft-tissue applications. Lipid droplets contain triglycerides, and can be easily confirmed histologically using Oil red O and Sudan III staining. Gene expression that is specific to mature adipocutes includes peroxisome proliferator activated receptor (PPAR)-γ2, leptin, aP2, and glucose transporter type 4[56]. The real-time PCR study showed that the expression levels of PPAR-γ2 in ASCs isolated from female mice were higher than in those from male mice, suggesting that adipogenic differentiation of ASCs is closely related to gender[57].

When ASCs are cultured in osteogenic differentiation media, which may contain 1,25-dihydroxyvitamin D3, ascorbate-2-phosphate, and bone morphogenetic protein-2 (BMP)-2, for 2 to 4 wk, the cells differentiate into osteoblast-like cells in vitro[58]. After differentiation, the osteoblast-like cells start to produce calcium phosphate within the ECM which can be assessed with Alizarin Red or von Kossa staining to reveal osteocytes. Alkaline phosphatase, type I collagen, osteoponin, osteocalcin, bone sialoprotein, Runx-1, BMP-2, BMP-4, parathyroid hormone receptor, BMP receptor 1 and 2 are common genes that are up-regulated during osteogenesis[56]. Furthermore, male ASCs differentiate into bone more rapidly and more effectively than female ASCs[59].

For chondrogenic differentiation, cells typically require a 3D environment, such as an “aggregate culture” or “micromass pellet culture”. The micromass pellet culture model mimics precartilage condensation during embryonic development, which increases the cell-to-cell interaction and leads to the production of a cartilage-like matrix[60]. Chondrogenic differentiation requires the use of a defined media supplemented with TGF-β1, insulin, dexamethasone, ascorbate-2-phosphate, and BMP-6. Basic FGF can be used to expand ASCs, and at the same time, down-regulate chondrogenic markers during cell expansion[61]. Differentiated chondrocytes express type II collagen, type IV collagen, aggrecan, prolyl endopeptidase-like, and sulfate-proteoglycan[62]. Alcian blue and collagen type II staining indicate chondrocytes.

Although somewhat controversial, ASCs may possess ectodermal differentiation capacity, e.g., neurogenesis. Many studies have been reported[48,63,64]. Under culture conditions with media containing butylated acid, valproic acid, and insulin, ASCs become morphologically similar to neurons, and express markers of both neuronal (neuron-specific enolase, nestin, and NeuN) and glial lineages [S100, p75, nerve growth factor (NGF), receptorm, and NG2][48]. The differentiation of ASCs into Schwann cells that are capable of myelinating peripheral neurons has been reported[48]. Human ASCs form nestin-positive neurospheres and express Schwann cell markers including S100, glial fibrillary acidic protein, and the p75 NGF receptor after dissociation.

In addition to mesodermal and ectodermal capacity, the endodermal differentiation of ASCs has been reported. Numerous studies reported differentiation of ASCs into hepatocytes and beta islet cells[42,65,66]. In an environment with the differentiation factors activin-A, exendin-4, HGF, and pentagastrin, ASCs were demonstrated to differentiate into insulin-producing cells in vitro[66]. Meanwhile, adding HGF, oncostatin M, and dimethyl sulfoxide in the culture media resulted in the ability of ASCs to gain hepatocytic functions in vitro, including albumin and alpha-fetoprotein expression and urea production[42].


While in vitro differentiation of ASCs into multiple phenotypes has been reported, the in vivo translation can be challenging. While ASCs have many advantages as a cell source, (e.g., easily harvested, abundant, and easy to culture), there remains the challenge of cell survival in vivo. Poor cell survival after in vivo injection or implantation is common. This is in part due to the hypoxic environment, particularly if cells are transplanted into ischemic tissues. ASCs have been shown to survive in ischemic tissues, whereas mature adipocytes die easily under ischemic conditions[67], ASCs also secrete angiogenic factors under hypoxic conditions[34,35]. For certain clinical applications, ASC implantation may require suitable biomaterial scaffolds that support cell attachment, proliferation, and differentiation. The scaffold should be selected based numerous characteristics, such as porosity, bioactivity, mechanical integrity, biodegradability, and low immunogenicity. Ideal scaffolds can provide cells with an environment suitable for cell survival[68]. The environment immediately following implantation can be severe for the cells to survive because oxygen and nutrients are insufficient. Implanted cells need to survive until effective angiogenesis occurs. As described above, ASCs secrete significant levels of angiogenic factors under hypoxia. ASCs can survive in an ischemic environment, and provide a reservoir of growth factors that are necessary for angiogenesis.

ASCs have immense potential in wound healing applications. Altman et al[69] grafted an acellular dermal matrix construct seeded with human ASCs into a murine injury model, and found that ASCs enhanced the wound healing at day 7. Most of the ASCs were viable 2 wk after the engraftment. An appropriate scaffold contributed to ASC homing and surviving. ASCs grafted in the wound can result in the augmentation of the local blood supply and in an improvement of regeneration capacity.

As ASC differentiation into adipocytes is well established, adipose tissue regeneration using ASCs in vivo has been investigated. Clinical applications include soft tissue augmentation after injury, surgical resection, and congenital malformations. Among the strategies to generate adipose tissue are the combination of ASCs and scaffolds, the use of acellular scaffolds, and the addition of drugs or growth factors to the scaffolds that have been examined include type I collagen, fibrin, silk fibroin, alginate, hyaluronic acid, and matrigel[70-72]. Injectable scaffolds are an attractive option, as minimally invasive therapies would be widely adapted by surgeons. Methods of drug delivery include using polymeric microspheres to control the release of factors such as bFGF, insulin, and dexamethasone[73-75].

Regarding osteogenic potential, ASCs show promise for bone tissue regeneration after injection or congenital malformations. Since ASCs were discovered to have osteogenic potential, many in vivo studies have combined ASCs with biodegradable scaffold materials to promote bone growth. Immuno-deficient animal models for nonweight-bearing bone formation have become a common model to assess human ASC osteogenic potential in vivo. Because bone is composed of hydroxyapatite (HA) crystals, bioceramics such as HA and beta-tricalcium phosphate are used for bone regeneration. The ceramic biomaterials used in these applications should mimic the natural bone architecture to ensure ASC attachment and migration within porous materials. In addition, ceramic biomaterials should be absorbed overtime or integrated with the surrounding tissue and eventually replaced by new or existing host tissue. As collagen is the other main component of bone tissue, it has been widely studied as a natural biomaterial scaffold for bone regeneration. In contrast, synthetic polymers such as poly (L-lactic acid-co-glycolic acid), and poly-e-caprolactone (PCL), can also be utilized for bone engineering. The advantage of these polymers is that they are readily reproducible, and have flexible mechanical, chemical, and biological properties that allow them to be tailored to suit specific functions[76].

There has been much interest in examining ASCs for the cartilage tissue engineering required to remedy osteoarthritis (OA), which affects millions of patients all over the world. A cure for OA remains elusive, because, in part, while the cartilage ECM is maintained by a sparse population of chondrocytes, it exhibits little capacity for self-repair owing to the lack of a tissue blood supply. Researchers have investigated a variety of scaffold materials including alginate, agarose, fibrin, gelatin, and chondroitin sulfate to evaluate their ability to support chondrogenic differentiation of ASCs in vivo[77-79]. Several studies have demonstrated that ASCs were able to differentiate into chondrocytes in vivo when seeded within any of these scaffolds, but different construct materials can significantly influence the differentiation of ASCs and functional properties of the tissue-engineered construct[77,78].

Finally, ASCs also have potential in neural applications. Peripheral nerves can be regenerated if injuries are small, and bioengineering strategies are focused on alternatives to the nerve autograft[80]. Properties of the ideal nerve conduit should include biodegradability, controlled release of growth factors, incorporation of support cells, electrical activity, intraluminal channels, and oriented nerve substratum. Santiago et al[81] was among the first to report that implanted ASCs into the lumen of PCL-based nerve conduits in a rat sciatic nerve defect model was shown to promote the formation of a more robust nerve. However, both the endodermal and ectodermal transdifferentiation of ASCs remains to be validated.


A number of clinical applications using ASCs can be found through searches and on clinical trial websites. ASCs are mainly used for cell-based therapy, and the combination of ASCs with biomaterials or drugs is still to be studied. Most studies use adipose tissue as the scaffold. Garcia-Olmo et al[82-86] performed phase I-III clinical trials to investigate the efficacy and safety of expanded ASCs in the treatment of complex perianal fistulae including Crohn’s disease. Autologous ASCs were mixed with fibrin glue then injected into the fistulous tract. As a result, patients who received ASCs demonstrated a better rate of healing compared to the patients who received fibrin glue without ASCs. ASCs with fibrin glue therapy were determined to be a safe and effective for treating complex perianal fistulae. Two mechanisms of ASCs to treat fistulae are speculated: one was that ASCs induced immunosuppressive activity, and the other was that ASCs might help healing through the expression of matrix proteins[83].

One of the first clinical reports using stem cells derived from adipose tissue in a patient was reported in 2004. Lendeckel et al[87] reported a case of a 7-year-old girl suffering from widespread calvarial defects after severe head injury with multifragment calvarial fractures. This is among the first reports of bone tissue engineering using autologous stromal vascular fraction and fibrin glue, although it was a case study. Fibrin glue was manufactured from the patient’s plasma 2 d prior to the surgery. SVF was kept in place using autologous fibrin glue, and computed tomography scans showed new bone formation 3 mo after the reconstruction. It was noted that ASCs have a great advantage in the point of cell yield compared to BM-MSCs especially for pediatric patients. Indeed, 295 × 106 mononuclear cells were extracted from 42.3 g adipose tissue, and about 2%-3% of these cells are expected to be stem cells[87].

The disadvantages associated with the implantation of synthetic materials or autologous fat grafts could be overcome by engineered adipose tissue. Stillaert et al[88] attempted adipose tissue engineering in 12 volunteers. Hyaluronic acid-based scaffolds were implanted in the sub-umbilical area with and without ASCs. Unlike successful results with nude mice[89], the hyaluronic acid-based scaffolds didn’t support ASC survival and were not inductive towards adipose tissue formation in humans. Meanwhile, ASC enriched lipotransfer has been studied for facial lipoatrophy and breast augmentation[90,91]. Yoshimura et al[92] enrolled 15 patients, transplanted SVF containing lipoaspirate after removing artificial breast implants, and followed for 12 mo. It was concluded that ASC-rich lipotransfer is effective to enhance the volume of injected adipose tissue[90-92]. The increased volume of adipose tissue may not be due to ASC differentiation but paracrine support of the tissue through the secretion of angiogenic and adipogenic factors. However, the interaction between ASCs and cancer cells are not fully elucidated. ASCs may promote cancer growth and metastasis through paracrine properties, epithelial-mesenchymal transition[93,94], and immunosuppressive mechanisms[95,96]. Higher risk of local recurrence was observed in early stage breast cancer patients following lipoinjection[97]. ASCs have not only bright side for regenerative medicine but dark side as cancer promotion.

In the field of wound healing, Rigotti et al[98] showed ASCs are effective on severe symptoms such as atrophy, retraction, fibrosis, or ulcers induced by radiation therapy. Twenty patients were recruited and received lipoaspirate containing ASCs repeatedly, and followed-up to 31 mo. Patients demonstrated an improvement of ultrastructual tissue characteristics with neovessel formation as well as significant clinical improvements. The authors concluded that the treatment with ASC-containing lipoaspirates is potentially extended to other forms of microangiopathies.

Regarding the potential of ASCs to generate immune tolerance for transplant patients, ASCs have been reported to have an immunomodulatory effect[99]. It has been shown that ASCs don’t possess human leucocyte antigen class II antigens, and ASCs can suppress inflammatory cytokines, stimulate anti-inflammatory cytokine interleukin-10, and induce antigen-specific regulatory T cells[100]. In a case study, the intravenous infusion of allogenic ASCs in treating severe refractory acute graft-versus-host disease has proven to be effective[101]. Fang et al[102,103] treated patients with hematologic and immunologic disorders such as idiopathic thrombocytopenic purpura and refractory pure red cell aplasia, with allogenic ASC infusions, and reported significant improvements with these patients. From these results, ASCs are suggested to have immunomodulatory.


ASCs have prominent implications in tissue regeneration due to their high cell yield in adipose tissue, the ability to differentiate into multiple lineages and secrete various cytokines, and immunomodulatory effects. A large number of clinical trials using ASCs have already performed and many of them are ongoing. However, very few phase III clinical studies have been published. ASCs are a promising cell source for regenerative medicine, and more research is needed to warrant the safety of ASCs and the efficacy of tissue engineering using ASCs.


P- Reviewers: Bonetti B, Ho I, Phinney DG S- Editor: Song XX L- Editor: A E- Editor: Liu SQ

1.  Friedenstein AJ, Petrakova KV, Kurolesova AI, Frolova GP. Heterotopic of bone marrow. Analysis of precursor cells for osteogenic and hematopoietic tissues. Transplantation. 1968;6:230-247.  [PubMed]  [DOI]
2.  Pittenger MF, Mackay AM, Beck SC, Jaiswal RK, Douglas R, Mosca JD, Moorman MA, Simonetti DW, Craig S, Marshak DR. Multilineage potential of adult human mesenchymal stem cells. Science. 1999;284:143-147.  [PubMed]  [DOI]
3.  da Silva Meirelles L, Chagastelles PC, Nardi NB. Mesenchymal stem cells reside in virtually all post-natal organs and tissues. J Cell Sci. 2006;119:2204-2213.  [PubMed]  [DOI]
4.  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.  [PubMed]  [DOI]
5.  Erices A, Conget P, Minguell JJ. Mesenchymal progenitor cells in human umbilical cord blood. Br J Haematol. 2000;109:235-242.  [PubMed]  [DOI]
6.  Roufosse CA, Direkze NC, Otto WR, Wright NA. Circulating mesenchymal stem cells. Int J Biochem Cell Biol. 2004;36:585-597.  [PubMed]  [DOI]
7.  Gronthos S, Mankani M, Brahim J, Robey PG, Shi S. Postnatal human dental pulp stem cells (DPSCs) in vitro and in vivo. Proc Natl Acad Sci USA. 2000;97:13625-13630.  [PubMed]  [DOI]
8.  Haniffa MA, Wang XN, Holtick U, Rae M, Isaacs JD, Dickinson AM, Hilkens CM, Collin MP. Adult human fibroblasts are potent immunoregulatory cells and functionally equivalent to mesenchymal stem cells. J Immunol. 2007;179:1595-1604.  [PubMed]  [DOI]
9.  Sessarego N, Parodi A, Podestà M, Benvenuto F, Mogni M, Raviolo V, Lituania M, Kunkl A, Ferlazzo G, Bricarelli FD. Multipotent mesenchymal stromal cells from amniotic fluid: solid perspectives for clinical application. Haematologica. 2008;93:339-346.  [PubMed]  [DOI]
10.  Yan XL, Fu CJ, Chen L, Qin JH, Zeng Q, Yuan HF, Nan X, Chen HX, Zhou JN, Lin YL. Mesenchymal stem cells from primary breast cancer tissue promote cancer proliferation and enhance mammosphere formation partially via EGF/EGFR/Akt pathway. Breast Cancer Res Treat. 2012;132:153-164.  [PubMed]  [DOI]
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.  [PubMed]  [DOI]
12.  Elabd C, Chiellini C, Carmona M, Galitzky J, Cochet O, Petersen R, Pénicaud L, Kristiansen K, Bouloumié A, Casteilla L. Human multipotent adipose-derived stem cells differentiate into functional brown adipocytes. Stem Cells. 2009;27:2753-2760.  [PubMed]  [DOI]
13.  Tseng YH, Kokkotou E, Schulz TJ, Huang TL, Winnay JN, Taniguchi CM, Tran TT, Suzuki R, Espinoza DO, Yamamoto Y. New role of bone morphogenetic protein 7 in brown adipogenesis and energy expenditure. Nature. 2008;454:1000-1004.  [PubMed]  [DOI]
14.  Wu J, Boström P, Sparks LM, Ye L, Choi JH, Giang AH, Khandekar M, Virtanen KA, Nuutila P, Schaart G. Beige adipocytes are a distinct type of thermogenic fat cell in mouse and human. Cell. 2012;150:366-376.  [PubMed]  [DOI]
15.  Barbatelli G, Murano I, Madsen L, Hao Q, Jimenez M, Kristiansen K, Giacobino JP, De Matteis R, Cinti S. The emergence of cold-induced brown adipocytes in mouse white fat depots is determined predominantly by white to brown adipocyte transdifferentiation. Am J Physiol Endocrinol Metab. 2010;298:E1244-E1253.  [PubMed]  [DOI]
16.  Wang QA, Tao C, Gupta RK, Scherer PE. Tracking adipogenesis during white adipose tissue development, expansion and regeneration. Nat Med. 2013;19:1338-1344.  [PubMed]  [DOI]
17.  Himms-Hagen J, Melnyk A, Zingaretti MC, Ceresi E, Barbatelli G, Cinti S. Multilocular fat cells in WAT of CL-316243-treated rats derive directly from white adipocytes. Am J Physiol Cell Physiol. 2000;279:C670-C681.  [PubMed]  [DOI]
18.  Fraser J, Wulur I, Alfonso Z, Zhu M, Wheeler E. Differences in stem and progenitor cell yield in different subcutaneous adipose tissue depots. Cytotherapy. 2007;9:459-467.  [PubMed]  [DOI]
19.  Aust L, Devlin B, Foster SJ, Halvorsen YD, Hicok K, du Laney T, Sen A, Willingmyre GD, Gimble JM. Yield of human adipose-derived adult stem cells from liposuction aspirates. Cytotherapy. 2004;6:7-14.  [PubMed]  [DOI]
20.  Yoshimura K, Shigeura T, Matsumoto D, Sato T, Takaki Y, Aiba-Kojima E, Sato K, Inoue K, Nagase T, Koshima I. Characterization of freshly isolated and cultured cells derived from the fatty and fluid portions of liposuction aspirates. J Cell Physiol. 2006;208:64-76.  [PubMed]  [DOI]
21.  De Ugarte DA, Morizono K, Elbarbary A, Alfonso Z, Zuk PA, Zhu M, Dragoo JL, Ashjian P, Thomas B, Benhaim P. Comparison of multi-lineage cells from human adipose tissue and bone marrow. Cells Tissues Organs. 2003;174:101-109.  [PubMed]  [DOI]
22.  Eto H, Suga H, Matsumoto D, Inoue K, Aoi N, Kato H, Araki J, Yoshimura K. Characterization of structure and cellular components of aspirated and excised adipose tissue. Plast Reconstr Surg. 2009;124:1087-1097.  [PubMed]  [DOI]
23.  Mitchell JB, McIntosh K, Zvonic S, Garrett S, Floyd ZE, Kloster A, Di Halvorsen Y, Storms RW, Goh B, Kilroy G. Immunophenotype of human adipose-derived cells: temporal changes in stromal-associated and stem cell-associated markers. Stem Cells. 2006;24:376-385.  [PubMed]  [DOI]
24.  Guilak F, Lott KE, Awad HA, Cao Q, Hicok KC, Fermor B, Gimble JM. Clonal analysis of the differentiation potential of human adipose-derived adult stem cells. J Cell Physiol. 2006;206:229-237.  [PubMed]  [DOI]
25.  Gonda K, Shigeura T, Sato T, Matsumoto D, Suga H, Inoue K, Aoi N, Kato H, Sato K, Murase S. Preserved proliferative capacity and multipotency of human adipose-derived stem cells after long-term cryopreservation. Plast Reconstr Surg. 2008;121:401-410.  [PubMed]  [DOI]
26.  Li H, Zimmerlin L, Marra KG, Donnenberg VS, Donnenberg AD, Rubin JP. Adipogenic potential of adipose stem cell subpopulations. Plast Reconstr Surg. 2011;128:663-672.  [PubMed]  [DOI]
27.  Katz AJ, Tholpady A, Tholpady SS, Shang H, Ogle RC. Cell surface and transcriptional characterization of human adipose-derived adherent stromal (hADAS) cells. Stem Cells. 2005;23:412-423.  [PubMed]  [DOI]
28.  McIntosh K, Zvonic S, Garrett S, Mitchell JB, Floyd ZE, Hammill L, Kloster A, Di Halvorsen Y, Ting JP, Storms RW. The immunogenicity of human adipose-derived cells: temporal changes in vitro. Stem Cells. 2006;24:1246-1253.  [PubMed]  [DOI]
29.  Zimmerlin L, Donnenberg VS, Pfeifer ME, Meyer EM, Péault B, Rubin JP, Donnenberg AD. Stromal vascular progenitors in adult human adipose tissue. Cytometry A. 2010;77:22-30.  [PubMed]  [DOI]
30.  Zannettino AC, Paton S, Arthur A, Khor F, Itescu S, Gimble JM, Gronthos S. Multipotential human adipose-derived stromal stem cells exhibit a perivascular phenotype in vitro and in vivo. J Cell Physiol. 2008;214:413-421.  [PubMed]  [DOI]
31.  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.  [PubMed]  [DOI]
32.  Salgado AJ, Reis RL, Sousa NJ, Gimble JM. Adipose tissue derived stem cells secretome: soluble factors and their roles in regenerative medicine. Curr Stem Cell Res Ther. 2010;5:103-110.  [PubMed]  [DOI]
33.  Bhang SH, Cho SW, Lim JM, Kang JM, Lee TJ, Yang HS, Song YS, Park MH, Kim HS, Yoo KJ. Locally delivered growth factor enhances the angiogenic efficacy of adipose-derived stromal cells transplanted to ischemic limbs. Stem Cells. 2009;27:1976-1986.  [PubMed]  [DOI]
34.  Lee EY, Xia Y, Kim WS, Kim MH, Kim TH, Kim KJ, Park BS, Sung JH. Hypoxia-enhanced wound-healing function of adipose-derived stem cells: increase in stem cell proliferation and up-regulation of VEGF and bFGF. Wound Repair Regen. 2009;17:540-547.  [PubMed]  [DOI]
35.  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.  [PubMed]  [DOI]
36.  Lee HC, An SG, Lee HW, Park JS, Cha KS, Hong TJ, Park JH, Lee SY, Kim SP, Kim YD. Safety and effect of adipose tissue-derived stem cell implantation in patients with critical limb ischemia: a pilot study. Circ J. 2012;76:1750-1760.  [PubMed]  [DOI]
37.  Muthukrishnan L, Warder E, McNeil PL. Basic fibroblast growth factor is efficiently released from a cytolsolic storage site through plasma membrane disruptions of endothelial cells. J Cell Physiol. 1991;148:1-16.  [PubMed]  [DOI]
38.  Werner S, Grose R. Regulation of wound healing by growth factors and cytokines. Physiol Rev. 2003;83:835-870.  [PubMed]  [DOI]
39.  Kaewsuwan S, Song SY, Kim JH, Sung JH. Mimicking the functional niche of adipose-derived stem cells for regenerative medicine. Expert Opin Biol Ther. 2012;12:1575-1588.  [PubMed]  [DOI]
40.  Kilroy GE, Foster SJ, Wu X, Ruiz J, Sherwood S, Heifetz A, Ludlow JW, Stricker DM, Potiny S, Green P. Cytokine profile of human adipose-derived stem cells: expression of angiogenic, hematopoietic, and pro-inflammatory factors. J Cell Physiol. 2007;212:702-709.  [PubMed]  [DOI]
41.  Song SY, Chung HM, Sung JH. The pivotal role of VEGF in adipose-derived-stem-cell-mediated regeneration. Expert Opin Biol Ther. 2010;10:1529-1537.  [PubMed]  [DOI]
42.  Seo MJ, Suh SY, Bae YC, Jung JS. Differentiation of human adipose stromal cells into hepatic lineage in vitro and in vivo. Biochem Biophys Res Commun. 2005;328:258-264.  [PubMed]  [DOI]
43.  Hauner H, Röhrig K, Petruschke T. Effects of epidermal growth factor (EGF), platelet-derived growth factor (PDGF) and fibroblast growth factor (FGF) on human adipocyte development and function. Eur J Clin Invest. 1995;25:90-96.  [PubMed]  [DOI]
44.  Kakudo N, Shimotsuma A, Kusumoto K. Fibroblast growth factor-2 stimulates adipogenic differentiation of human adipose-derived stem cells. Biochem Biophys Res Commun. 2007;359:239-244.  [PubMed]  [DOI]
45.  Kang YJ, Jeon ES, Song HY, Woo JS, Jung JS, Kim YK, Kim JH. 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.  [PubMed]  [DOI]
46.  Quarto N, Longaker MT. FGF-2 inhibits osteogenesis in mouse adipose tissue-derived stromal cells and sustains their proliferative and osteogenic potential state. Tissue Eng. 2006;12:1405-1418.  [PubMed]  [DOI]
47.  Serrero G. EGF inhibits the differentiation of adipocyte precursors in primary cultures. Biochem Biophys Res Commun. 1987;146:194-202.  [PubMed]  [DOI]
48.  Radtke C, Schmitz B, Spies M, Kocsis JD, Vogt PM. Peripheral glial cell differentiation from neurospheres derived from adipose mesenchymal stem cells. Int J Dev Neurosci. 2009;27:817-823.  [PubMed]  [DOI]
49.  Timper K, Seboek D, Eberhardt M, Linscheid P, Christ-Crain M, Keller U, Müller B, Zulewski H. Human adipose tissue-derived mesenchymal stem cells differentiate into insulin, somatostatin, and glucagon expressing cells. Biochem Biophys Res Commun. 2006;341:1135-1140.  [PubMed]  [DOI]
50.  Pachón-Peña G, Yu G, Tucker A, Wu X, Vendrell J, Bunnell BA, Gimble JM. Stromal stem cells from adipose tissue and bone marrow of age-matched female donors display distinct immunophenotypic profiles. J Cell Physiol. 2011;226:843-851.  [PubMed]  [DOI]
51.  Sakaguchi Y, Sekiya I, Yagishita K, Muneta T. Comparison of human stem cells derived from various mesenchymal tissues: superiority of synovium as a cell source. Arthritis Rheum. 2005;52:2521-2529.  [PubMed]  [DOI]
52.  Noël D, Caton D, Roche S, Bony C, Lehmann S, Casteilla L, Jorgensen C, Cousin B. Cell specific differences between human adipose-derived and mesenchymal-stromal cells despite similar differentiation potentials. Exp Cell Res. 2008;314:1575-1584.  [PubMed]  [DOI]
53.  Bochev I, Elmadjian G, Kyurkchiev D, Tzvetanov L, Altankova I, Tivchev P, Kyurkchiev S. Mesenchymal stem cells from human bone marrow or adipose tissue differently modulate mitogen-stimulated B-cell immunoglobulin production in vitro. Cell Biol Int. 2008;32:384-393.  [PubMed]  [DOI]
54.  Brayfield C, Marra K, Rubin JP. Adipose stem cells for soft tissue regeneration. Handchir Mikrochir Plast Chir. 2010;42:124-128.  [PubMed]  [DOI]
55.  Kubo Y, Kaidzu S, Nakajima I, Takenouchi K, Nakamura F. Organization of extracellular matrix components during differentiation of adipocytes in long-term culture. In Vitro Cell Dev Biol Anim. 2000;36:38-44.  [PubMed]  [DOI]
56.  Gentile P, Orlandi A, Scioli MG, Di Pasquali C, Bocchini I, Cervelli V. Concise review: adipose-derived stromal vascular fraction cells and platelet-rich plasma: basic and clinical implications for tissue engineering therapies in regenerative surgery. Stem Cells Transl Med. 2012;1:230-236.  [PubMed]  [DOI]
57.  Ogawa R, Mizuno H, Watanabe A, Migita M, Hyakusoku H, Shimada T. Adipogenic differentiation by adipose-derived stem cells harvested from GFP transgenic mice-including relationship of sex differences. Biochem Biophys Res Commun. 2004;319:511-517.  [PubMed]  [DOI]
58.  Halvorsen YD, Franklin D, Bond AL, Hitt DC, Auchter C, Boskey AL, Paschalis EP, Wilkison WO, Gimble JM. Extracellular matrix mineralization and osteoblast gene expression by human adipose tissue-derived stromal cells. Tissue Eng. 2001;7:729-741.  [PubMed]  [DOI]
59.  Aksu AE, Rubin JP, Dudas JR, Marra KG. Role of gender and anatomical region on induction of osteogenic differentiation of human adipose-derived stem cells. Ann Plast Surg. 2008;60:306-322.  [PubMed]  [DOI]
60.  Wei Y, Sun X, Wang W, Hu Y. Adipose-derived stem cells and chondrogenesis. Cytotherapy. 2007;9:712-716.  [PubMed]  [DOI]
61.  Khan WS, Tew SR, Adesida AB, Hardingham TE. Human infrapatellar fat pad-derived stem cells express the pericyte marker 3G5 and show enhanced chondrogenesis after expansion in fibroblast growth factor-2. Arthritis Res Ther. 2008;10:R74.  [PubMed]  [DOI]
62.  Hildner F, Albrecht C, Gabriel C, Redl H, van Griensven M. State of the art and future perspectives of articular cartilage regeneration: a focus on adipose-derived stem cells and platelet-derived products. J Tissue Eng Regen Med. 2011;5:e36-e51.  [PubMed]  [DOI]
63.  Anghileri E, Marconi S, Pignatelli A, Cifelli P, Galié M, Sbarbati A, Krampera M, Belluzzi O, Bonetti B. Neuronal differentiation potential of human adipose-derived mesenchymal stem cells. Stem Cells Dev. 2008;17:909-916.  [PubMed]  [DOI]
64.  Yu JM, Bunnell BA, Kang SK. Neural differentiation of human adipose tissue-derived stem cells. Methods Mol Biol. 2011;702:219-231.  [PubMed]  [DOI]
65.  Bonora-Centelles A, Jover R, Mirabet V, Lahoz A, Carbonell F, Castell JV, Gómez-Lechón MJ. Sequential hepatogenic transdifferentiation of adipose tissue-derived stem cells: relevance of different extracellular signaling molecules, transcription factors involved, and expression of new key marker genes. Cell Transplant. 2009;18:1319-1340.  [PubMed]  [DOI]
66.  Okura H, Komoda H, Fumimoto Y, Lee CM, Nishida T, Sawa Y, Matsuyama A. Transdifferentiation of human adipose tissue-derived stromal cells into insulin-producing clusters. J Artif Organs. 2009;12:123-130.  [PubMed]  [DOI]
67.  Suga H, Eto H, Aoi N, Kato H, Araki J, Doi K, Higashino T, Yoshimura K. Adipose tissue remodeling under ischemia: death of adipocytes and activation of stem/progenitor cells. Plast Reconstr Surg. 2010;126:1911-1923.  [PubMed]  [DOI]
68.  Tabata Y. Biomaterial technology for tissue engineering applications. J R Soc Interface. 2009;6 Suppl 3:S311-S324.  [PubMed]  [DOI]
69.  Altman AM, Matthias N, Yan Y, Song YH, Bai X, Chiu ES, Slakey DP, Alt EU. Dermal matrix as a carrier for in vivo delivery of human adipose-derived stem cells. Biomaterials. 2008;29:1431-1442.  [PubMed]  [DOI]
70.  Choi JH, Gimble JM, Lee K, Marra KG, Rubin JP, Yoo JJ, Vunjak-Novakovic G, Kaplan DL. Adipose tissue engineering for soft tissue regeneration. Tissue Eng Part B Rev. 2010;16:413-426.  [PubMed]  [DOI]
71.  Tsuji W, Inamoto T, Yamashiro H, Ueno T, Kato H, Kimura Y, Tabata Y, Toi M. Adipogenesis induced by human adipose tissue-derived stem cells. Tissue Eng Part A. 2009;15:83-93.  [PubMed]  [DOI]
72.  Ito R, Morimoto N, Liem PH, Nakamura Y, Kawai K, Taira T, Tsuji W, Toi M, Suzuki S. Adipogenesis using human adipose tissue-derived stromal cells combined with a collagen/gelatin sponge sustaining release of basic fibroblast growth factor. J Tissue Eng Regen Med. 2012;Epub ahead of print.  [PubMed]  [DOI]
73.  Marra KG, Defail AJ, Clavijo-Alvarez JA, Badylak SF, Taieb A, Schipper B, Bennett J, Rubin JP. FGF-2 enhances vascularization for adipose tissue engineering. Plast Reconstr Surg. 2008;121:1153-1164.  [PubMed]  [DOI]
74.  Kimura Y, Ozeki M, Inamoto T, Tabata Y. Time course of de novo adipogenesis in matrigel by gelatin microspheres incorporating basic fibroblast growth factor. Tissue Eng. 2002;8:603-613.  [PubMed]  [DOI]
75.  Rubin JP, DeFail A, Rajendran N, Marra KG. Encapsulation of adipogenic factors to promote differentiation of adipose-derived stem cells. J Drug Target. 2009;17:207-215.  [PubMed]  [DOI]
76.  Zanetti AS, Sabliov C, Gimble JM, Hayes DJ. Human adipose-derived stem cells and three-dimensional scaffold constructs: a review of the biomaterials and models currently used for bone regeneration. J Biomed Mater Res B Appl Biomater. 2013;101:187-199.  [PubMed]  [DOI]
77.  Awad HA, Wickham MQ, Leddy HA, Gimble JM, Guilak F. Chondrogenic differentiation of adipose-derived adult stem cells in agarose, alginate, and gelatin scaffolds. Biomaterials. 2004;25:3211-3222.  [PubMed]  [DOI]
78.  Wei Y, Hu Y, Hao W, Han Y, Meng G, Zhang D, Wu Z, Wang H. A novel injectable scaffold for cartilage tissue engineering using adipose-derived adult stem cells. J Orthop Res. 2008;26:27-33.  [PubMed]  [DOI]
79.  Dragoo JL, Carlson G, McCormick F, Khan-Farooqi H, Zhu M, Zuk PA, Benhaim P. Healing full-thickness cartilage defects using adipose-derived stem cells. Tissue Eng. 2007;13:1615-1621.  [PubMed]  [DOI]
80.  Schmidt CE, Leach JB. Neural tissue engineering: strategies for repair and regeneration. Annu Rev Biomed Eng. 2003;5:293-347.  [PubMed]  [DOI]
81.  Santiago LY, Clavijo-Alvarez J, Brayfield C, Rubin JP, Marra KG. Delivery of adipose-derived precursor cells for peripheral nerve repair. Cell Transplant. 2009;18:145-158.  [PubMed]  [DOI]
82.  García-Olmo D, García-Arranz M, García LG, Cuellar ES, Blanco IF, Prianes LA, Montes JA, Pinto FL, Marcos DH, García-Sancho L. Autologous stem cell transplantation for treatment of rectovaginal fistula in perianal Crohn’s disease: a new cell-based therapy. Int J Colorectal Dis. 2003;18:451-454.  [PubMed]  [DOI]
83.  Garcia-Olmo D, Garcia-Arranz M, Herreros D. Expanded adipose-derived stem cells for the treatment of complex perianal fistula including Crohn’s disease. Expert Opin Biol Ther. 2008;8:1417-1423.  [PubMed]  [DOI]
84.  García-Olmo D, García-Arranz M, Herreros D, Pascual I, Peiro C, Rodríguez-Montes JA. A phase I clinical trial of the treatment of Crohn’s fistula by adipose mesenchymal stem cell transplantation. Dis Colon Rectum. 2005;48:1416-1423.  [PubMed]  [DOI]
85.  Garcia-Olmo D, Herreros D, Pascual I, Pascual JA, Del-Valle E, Zorrilla J, De-La-Quintana P, Garcia-Arranz M, Pascual M. Expanded adipose-derived stem cells for the treatment of complex perianal fistula: a phase II clinical trial. Dis Colon Rectum. 2009;52:79-86.  [PubMed]  [DOI]
86.  Garcia-Olmo D, Herreros D, Pascual M, Pascual I, De-La-Quintana P, Trebol J, Garcia-Arranz M. Treatment of enterocutaneous fistula in Crohn’s Disease with adipose-derived stem cells: a comparison of protocols with and without cell expansion. Int J Colorectal Dis. 2009;24:27-30.  [PubMed]  [DOI]
87.  Lendeckel S, Jödicke A, Christophis P, Heidinger K, Wolff J, Fraser JK, Hedrick MH, Berthold L, Howaldt HP. Autologous stem cells (adipose) and fibrin glue used to treat widespread traumatic calvarial defects: case report. J Craniomaxillofac Surg. 2004;32:370-373.  [PubMed]  [DOI]
88.  Stillaert FB, Di Bartolo C, Hunt JA, Rhodes NP, Tognana E, Monstrey S, Blondeel PN. Human clinical experience with adipose precursor cells seeded on hyaluronic acid-based spongy scaffolds. Biomaterials. 2008;29:3953-3959.  [PubMed]  [DOI]
89.  von Heimburg D, Zachariah S, Low A, Pallua N. Influence of different biodegradable carriers on the in vivo behavior of human adipose precursor cells. Plast Reconstr Surg. 2001;108:411-420; discussion 421-422.  [PubMed]  [DOI]
90.  Yoshimura K, Sato K, Aoi N, Kurita M, Hirohi T, Harii K. Cell-assisted lipotransfer for cosmetic breast augmentation: supportive use of adipose-derived stem/stromal cells. Aesthetic Plast Surg. 2008;32:48-55; discussion 56-57.  [PubMed]  [DOI]
91.  Yoshimura K, Sato K, Aoi N, Kurita M, Inoue K, Suga H, Eto H, Kato H, Hirohi T, Harii K. Cell-assisted lipotransfer for facial lipoatrophy: efficacy of clinical use of adipose-derived stem cells. Dermatol Surg. 2008;34:1178-1185.  [PubMed]  [DOI]
92.  Yoshimura K, Asano Y, Aoi N, Kurita M, Oshima Y, Sato K, Inoue K, Suga H, Eto H, Kato H. Progenitor-enriched adipose tissue transplantation as rescue for breast implant complications. Breast J. 2010;16:169-175.  [PubMed]  [DOI]
93.  Zimmerlin L, Park TS, Zambidis ET, Donnenberg VS, Donnenberg AD. Mesenchymal stem cell secretome and regenerative therapy after cancer. Biochimie. 2013;95:2235-2245.  [PubMed]  [DOI]
94.  Battula VL, Evans KW, Hollier BG, Shi Y, Marini FC, Ayyanan A, Wang RY, Brisken C, Guerra R, Andreeff M. Epithelial-mesenchymal transition-derived cells exhibit multilineage differentiation potential similar to mesenchymal stem cells. Stem Cells. 2010;28:1435-1445.  [PubMed]  [DOI]
95.  Bertolini F, Lohsiriwat V, Petit JY, Kolonin MG. Adipose tissue cells, lipotransfer and cancer: a challenge for scientists, oncologists and surgeons. Biochim Biophys Acta. 2012;1826:209-214.  [PubMed]  [DOI]
96.  Fraser JK, Hedrick MH, Cohen SR. Oncologic risks of autologous fat grafting to the breast. Aesthet Surg J. 2011;31:68-75.  [PubMed]  [DOI]
97.  Petit JY, Rietjens M, Botteri E, Rotmensz N, Bertolini F, Curigliano G, Rey P, Garusi C, De Lorenzi F, Martella S. Evaluation of fat grafting safety in patients with intraepithelial neoplasia: a matched-cohort study. Ann Oncol. 2013;24:1479-1484.  [PubMed]  [DOI]
98.  Rigotti G, Marchi A, Galiè M, Baroni G, Benati D, Krampera M, Pasini A, Sbarbati A. Clinical treatment of radiotherapy tissue damage by lipoaspirate transplant: a healing process mediated by adipose-derived adult stem cells. Plast Reconstr Surg. 2007;119:1409-1422; discussion 1423-1424.  [PubMed]  [DOI]
99.  Puissant B, Barreau C, Bourin P, Clavel C, Corre J, Bousquet C, Taureau C, Cousin B, Abbal M, Laharrague P. Immunomodulatory effect of human adipose tissue-derived adult stem cells: comparison with bone marrow mesenchymal stem cells. Br J Haematol. 2005;129:118-129.  [PubMed]  [DOI]
100.  Gonzalez-Rey E, Gonzalez MA, Varela N, O’Valle F, Hernandez-Cortes P, Rico L, Büscher D, Delgado M. Human adipose-derived mesenchymal stem cells reduce inflammatory and T cell responses and induce regulatory T cells in vitro in rheumatoid arthritis. Ann Rheum Dis. 2010;69:241-248.  [PubMed]  [DOI]
101.  Fang B, Song Y, Lin Q, Zhang Y, Cao Y, Zhao RC, Ma Y. Human adipose tissue-derived mesenchymal stromal cells as salvage therapy for treatment of severe refractory acute graft-vs.-host disease in two children. Pediatr Transplant. 2007;11:814-817.  [PubMed]  [DOI]
102.  Fang B, Song Y, Li N, Li J, Han Q, Zhao RC. Mesenchymal stem cells for the treatment of refractory pure red cell aplasia after major ABO-incompatible hematopoietic stem cell transplantation. Ann Hematol. 2009;88:261-266.  [PubMed]  [DOI]
103.  Fang B, Mai L, Li N, Song Y. Favorable response of chronic refractory immune thrombocytopenic purpura to mesenchymal stem cells. Stem Cells Dev. 2012;21:497-502.  [PubMed]  [DOI]