Immunophenotype of Human Adipose-Derived Cells: Temporal Changes in Stromal-Asso - 《stemcells》干细胞学杂志干细胞之家



免疫细胞治疗专区	欢迎关注干细胞微信公众号

12 3 4 5 6 7 8 9 10 ... 25 下一页

返回列表

查看: 474434\|回复: 245	go Immunophenotype of Human Adipose-Derived Cells: Temporal Changes in Stromal-Asso [复制链接]

江边孤钓

版主

Rank: 7 Rank: 7 Rank: 7

积分: 0
威望: 0
包包: 483

楼主

发表于 2009-3-5 00:11 |只看该作者 |倒序浏览 |打印

作者：James B. Mitchella, Kevin McIntosha, Sanjin Zvonicb, Sara Garretta, Z. Elizabeth Floydb, Amy Klosterc, Yuan Di Halvorsenc,d, Robert W. Stormse, Brian Gohb, Gail Kilroyb, Xiying Wub, Jeffrey M. Gimbleb,c $ {% v* n3 P5 p4 m
   , s1 {& Z$ R5 E7 h! ?4 v
   * D7 }* D; D: h3 w9 w

                       v( j& `/ s  b0 X# e7 M* i
         ' c- r" H/ t3 t8 |% \7 `2 n8 D


         * S% [; i% r7 B3 e8 ?3 x

      7 [+ w6 O0 e* l- J
        v9 r1 ~7 U1 \1 R6 n, J

      【摘要】
   Adipose tissue represents an abundant and accessible source of multipotent adult stem cells and is used by many investigators for tissue engineering applications; however, not all laboratories use cells at equivalent stages of isolation and passage. We have compared the immunophenotype of freshly isolated human adipose tissue-derived stromal vascular fraction (SVF) cells relative to serial-passaged adipose-derived stem cells (ASCs). The initial SVF cells contained colony-forming unit fibroblasts at a frequency of 1:32. Colony-forming unit adipocytes and osteoblasts were present in the SVF cells at comparable frequencies (1:28 and 1:16, respectively). The immunophenotype of the adipose-derived cells based on flow cytometry changed progressively with adherence and passage. Stromal cell¨Cassociated markers (CD13, CD29, CD44, CD63, CD73, CD90, CD166) were initially low on SVF cells and increased significantly with successive passages. The stem cell¨Cassociated marker CD34 was at peak levels in the SVF cells and/or early-passage ASCs and remained present, although at reduced levels, throughout the culture period. Aldehyde dehydrogenase and the multidrug-resistance transport protein (ABCG2), both of which have been used to identify and characterize hematopoietic stem cells, are expressed by SVF cells and ASCs at detectable levels. Endothelial cell¨Cassociated markers (CD31, CD144 or VE-cadherin, vascular endothelial growth factor receptor 2, von Willebrand factor) were expressed on SVF cells and did not change significantly with serial passage. Thus, the adherence to plastic and subsequent expansion of human adipose-derived cells in fetal bovine serum-supplemented medium selects for a relatively homogeneous cell population, enriching for cells expressing a stromal immunophenotype, compared with the heterogeneity of the crude SVF.
      【关键词】 Adipocyte Adipose-derived stem cells Aldehyde dehydrogenase Alkaline phosphatase Bone Colony-forming unit Osteoblast Stromal vascular fraction
   INTRODUCTION4 k5 p) F/ M& A4 h" v

Until recently, bone marrow has been the major tissue source of human adult stem cells. Bone marrow contains hematopoietic stem cells (HSCs), which have served as the prototypic example of an adult stem cell. The defining characteristics of a stem cell, such as surface immunophenotype and the capacity for multipotent differentiation and self renewal, are based primarily on studies of HSCs ., f' Q; J( A# @4 K4 [
/ p- V8 z# G0 N/ w
Adipose tissue represents an alternative source of stem cells. Subcutaneous adipose depots are accessible, abundant, and replenishable, thereby providing a potential adult stem cell reservoir for each individual. Many groups working independently have shown that adult stem cells derived from white adipose tissues can differentiate along multiple pathways in vitro, including the adipocyte, chondrocyte, endothelial, epithelial, hematopoietic support, hepatocyte, neuronal, myogenic, and osteoblast lineages . In accordance with a consensus reached by investigators attending the Second Annual International Fat Applied Technology Society meeting (October 3¨C5, 2004, Pittsburgh, PA), we will refer to these cells as ASCs from now on. Although some groups have focused their attention exclusively on the minimally processed SVF cell population, others have worked with the expanded plastic adherent ASC subpopulation at various stages of passage. It is likely that the SVF cells and ASCs exhibit different features and properties. This investigation set out to determine the frequency of adherent and lineage progenitor cells in the SVF cell by colony-forming unit (CFU) assays and to define the immunophenotype of human adipose-derived cells at various stages of isolation, purification, and expansion, based on flow cytometric assays using markers associated with endothelial cell, stem cell, and MSC.

MATERIALS AND METHODS
$ h6 F2 b6 i9 ~
Materials/ Y( d9 m  W1 i
0 j$ y  _. i: C) M2 h
All materials were obtained from Sigma-Aldrich (St. Louis, http://www.sigmaaldrich.com) or Fisher Scientific (Pittsburgh, http://www.fisherscientific.com) unless otherwise noted.) p/ h2 u5 z4 b1 p3 {" n

ADAS Cell Isolation and Expansion6 S( |9 f' P! K
; }! q5 ?) ^' u$ }3 h5 k8 r* t
All protocols were reviewed and approved by the Pennington Biomedical Research Center Institutional Research Board before the study. Liposuction aspirates from subcutaneous adipose tissue sites were obtained from male and female subjects undergoing elective procedures in local plastic surgical offices. Tissues were washed three to four times with phosphate-buffered saline (PBS) and suspended in an equal volume of PBS supplemented with 1% bovine serum and 0.1% collagenase type I (Worthington Biochemical Corporation, Lakewood, NJ, http://www.worthington-biochem.com) prewarmed to 37¡ãC. The tissue was placed in a shaking water bath at 37¡ãC with continuous agitation for 60 minutes and centrifuged for 5 minutes at 300 to 500g at room temperature. The supernatant, containing mature adipocytes, was aspirated. The pellet was identified as the SVF cell. Portions of the SVF cells were resuspended in cryopreservation medium (10% dimethylsulfoxide, 10% Dulbecco¡¯s modified Eagle¡¯s medium , 100 U penicillin/100 µg streptomycin/0.25 µg fungizone) at a density of 0.156 ml of tissue digest/sq cm of surface area for expansion and culture. This initial passage of the primary cell culture was referred to as passage 0 (P0). After the first 48 hours of incubation at 37¡ãC at 5% CO2, the cultures were washed with PBS and maintained in stromal media until they achieved 75%¨C90% confluence (approximately 6 days in culture). The cells were passaged by trypsin (0.05%) digestion and plated at a density of 5,000 cells/cm2 (passage 1). Cell viability and numbers at the time of passage were determined by trypan blue exclusion and hemacytometer cell counts. Cells were passaged repeatedly after achieving a density of 75%¨C90% (approximately 6 days in culture) until passage 4., r- }2 O8 {9 y

Adipogenesis
: g5 f/ d$ b8 e  z
Confluent cultures of primary ASCs were induced to undergo adipogenesis by replacing the stromal media with adipocyte induction medium composed of DMEM/F-12 with 3% FBS, 33 µM biotin, 17 µM pantothenate, 1 µM bovine insulin, 1 µM dexamethasone, 0.5 mM isobutylmethylxanthine (IBMX), 5 µM rosiglitazone, and 100 U penicillin/100 µg streptomycin/0.25 µg fungizone. After 3 days, media were changed to adipocyte maintenance media that were identical to induction media except for the deletion of both IBMX and rosiglitazone. Cells were maintained in culture for up to 9 days, with 90% of the maintenance media replaced every 3 days. Cultures were rinsed with PBS and fixed in formalin solution, and adipocyte differentiation was determined by staining of neutral lipids with oil red O.
# S' v. Y' Y/ d/ C. O! Z+ t% T
Osteogenesis* g9 l6 A# d8 I8 E) X) E

Confluent cultures of primary ASCs were induced to undergo osteogenesis by replacing the stromal medium with osteogenic induction medium composed of DMEM/F-12 Ham¡¯s with 10% FBS, 10 mM ß-glycerophosphate, 10 nM dexamethasone, 50 µg/ml sodium ascorbate 2-phosphate, and 100 U penicillin/100 µg streptomycin/0.25 µg fungizone. Cultures were fed with fresh osteogenic induction medium every 3 to 4 days for a period of up to 3 weeks. Cultures were rinsed in 0.9% NaCl and fixed in 70% ethanol, and osteogenic differentiation was determined by staining for calcium phosphate with Alizarin red., ]+ J& z% N* o4 v+ b

Colony-Forming Unit Assays8 C& ^1 H# |* M) z( }  T

The frequency of CFUs was determined by limit dilution assay with the assumption that the number of progenitor cells follows a Poisson distribution . By combining this approach with the knowledge of the total number of cells seeded per well, it is possible to calculate the CFU frequency for any specific lineage by linear regression analysis. The second 96-well plate was committed to a CFU-alkaline phosphatase (CFU-ALP) assay. The plate was rinsed with PBS, fixed in 100% ethanol, incubated for 1 hour in the presence of a solution containing 36 mM sodium metaborate, 0.46 mM 5-bromo-4-chloro-3-indoxyl phosphate, 1.2 mM nitroblue tetrazolium, and 8.3 mM magnesium sulfate (pH 9.3), and rinsed with water, and the number of wells that did not contain colonies > 20 ALP  cells was determined for each cell concentration; these data were used to determine the number of CFU-ALP according to the above formula. The remaining two 96-well plates were induced to undergo adipogenesis or osteogenesis, respectively, as described above. The CFU-adipocyte (CFU-Ad) was determined by oil red O staining 9 days after induction, whereas the CFU-osteoblast (CFU-O) was determined by Alizarin red staining >14 days after induction. Similar CFU analyses were performed on serial dilutions of ASCs from passages P0 to P4 isolated from a group of n = 4 donors.

Flow Cytometry
- A8 P, ]" k- }2 n9 Z( ~
Flow cytometry was performed on cells from the SVF as well as from ASCs cultured from passages 0 through 4. All cells were cryopreserved at concentrations of 0.5 to 2.0 million cells per ml in 80% FBS, 10% dimethylsulfoxide, and 10% DMEM/F-12 Ham¡¯s medium for periods of ~1 to 4 months before analysis. Immediately before flow cytometric analysis, individual vials of cells were rapidly thawed in a 37¡ãC water bath (1 to 2 minutes of agitation), resuspended in 5 to 10 ml of medium containing 10% FBS, centrifuged at room temperature, and used for immunostaining. At this time, cell viability for all donors was at least 50% or greater based on trypan blue exclusion. Cells were analyzed for phenotypic markers falling within three general categories (hematopoietic, stromal, and stem cell) as well as aldehyde dehydrogenase (ALDH) activity (Stem Cell Technologies, Vancouver, British Columbia, Canada, http://www.stemcell.com). The cells were analyzed using both conjugated and unconjugated mouse monoclonal antibodies. Briefly, approximately 4 to 8 x 106 ASCs were acquired from each population. A total of 1 x 106 cells were removed for ALDH analysis, and 1 to 2 x 106 cells were removed for staining with the unconjugated monoclonals. Ten thousand events were acquired per antibody set, and a minimum of 25,000 events was acquired for the ALDH assay on a Becton, Dickinson and Company FACS-Caliber flow cytometer using CELLQuest acquisition software (Becton, Dickinson and Company, Franklin Lakes, NJ, http://www.bd.com). Data analysis was performed using Flow Jo analysis software (Tree Star, Ashland, OR, http://www.treestar.com).& }: l" Q% U% C- f5 T
# n% q) P0 r# i$ d4 w: w- U
Conjugated Monoclonal Antibodies

The cells were washed once in flow wash buffer (1x Dulbecco¡¯s phosphate-buffered saline, 0.5% BSA, and 0.1% sodium azide), resuspended in blocking buffer (wash buffer with 25 µg/ml mouse immunoglobulin G), and incubated for 10 minutes on ice. A total of 100 µl of cell suspension (approximately 5 x 105 cells) was aliquoted per tube, and appropriately labeled monoclonal antibodies were added for tricolor analysis (fluorescein isothiocyanate , and antigen-presenting cell). Appropriate isotype control combinations were performed to reflect the monoclonal isotype combinations. Antibodies directed against the following antigens (catalog no.) were purchased from BD Pharmingen (San Diego, http://www.bdbiosciences.com/pharmingen) unless otherwise indicated and used at the vendor-recommended quantities: CD13 PE (#555394), CD29 FITC (Caltag #CD2901), CD31 FITC (Caltag #MHCD3101), CD34 PE (#348057), CD44 FITC (Cell Sciences #852.601.010), CD49a PE (#559596), CD63 FITC (#557288), CD73 PE (#550257), CD90 FITC (#555595), CD105 PE (Caltag #MHCD10504), CD144 (Chemicon #MAB1989), CD146 PE (#550315), CD166 PE (#559263), ABCG2 FITC (Chemicon #MAB4155F), vascular endothelial growth factor receptor 2 (VEGFr2) (Chemicon #MAB1667), and von Willebrand factor (Chemicon MAB3442). All tubes were incubated on ice and protected from light for 30 minutes. The cells were washed once in wash buffer and fixed in 200 µl of 1% paraformaldehyde./ G0 D# y/ R: Q; Q. i/ ^; r

Unconjugated Monoclonal Antibodies

The cells were washed as stated above, blocked in wash buffer containing 5% goat serum, incubated for 10 minutes, and distributed into 100-µl aliquots. The primary antibodies (CD144, anti-VEGFr2 , and anti-von Willebrand factor) were added (10 µg/ml), and the cells were incubated for 30 minutes on ice. The cells were washed once in wash buffer and resuspended in wash buffer without serum. Goat anti-mouse PE-conjugated secondary antibody was added (5 µg/ml) to the suspensions containing primary antibody as well as a secondary-only control. The cells were incubated on ice and protected from light for 15 minutes. The cells were then washed in flow wash buffer and fixed with 1% paraformaldehyde.

RESULTS2 C" r/ N$ d5 A& j/ E3 _
' r* U3 d: N. C: y$ B- n
Cell Yield6 U0 u) b! Q% t! ?/ d( m1 k

Subcutaneous adipose tissue lipoaspirates obtained from a total of 44 donors were processed by collagenase digestion and differential centrifugation. The age (mean ¡À standard deviation . To assess the frequency of progenitor cells in the adipose tissue, the mean number of nucleated cell numbers present in the SVF was determined as 308,849 per ml of lipoaspirate tissue (Table 1). Based on these calculations, CFU assays were established in 96-well plates by limiting dilution assays to determine the CFU frequency for specific lineage phenotypes based on histochemical staining characteristics and the Poisson distribution as outlined in Materials and Methods (Tables 2, 3). After 9 days in the culture, the number of wells containing cells staining positive for toluidine blue or alkaline phosphatase was used to determine the frequency of CFU-F and CFU-ALP, respectively (Fig. 1). At that time, identical plates were induced to undergo adipogenesis or osteogenesis. The number of wells staining positive for neutral lipids by oil red O or for calcium phosphate by Alizarin red was determined after an additional 9 days or >14 days, respectively. The resulting mean CFU frequencies in the SVF isolated from n = 7 to 12 donors were as follows: CFU-F, 1 per 32 cells; CFU-ALP, 1 per 328 cells; CFU-Ad, 1 per 28 cells; and CFU-Ob, 1 per 16 cells (Table 2). The effect of successive passage on the CFU frequency was determined using a subset of donors (n = 4) (Table 3). The lineage frequencies increased by as much as 10-fold by passage 4, as evidenced by the number of CFU-ALP and CFU-Ad.

Table 1. Cell yields per milliliter of lipoaspirate tissue

Table 2. Frequency of colony-forming units within the nucleated stromal vascular fraction cell population: Q% ~# Y; @# [8 m4 i
4 q4 ?. L. l8 a7 k, O
Table 3. Frequency of colony-forming units with successive passage (n = 4 donors)

Figure 1. Colony-forming unit assays. The photomicrographs display staining profiles representative of the following colonies: bottom left, toluidine blue  colony-forming unit fibroblasts; top right, alkaline phosphatase  colony-forming unit alkaline phosphatase; top left, oil red O  colony-forming unit adipocytes; bottom right, Alizarin red  colony-forming unit osteoblasts.3 F0 d: a$ V8 o
& a, f, u* j* c) a4 x) A: D0 m
After the initial plating, cells were maintained in culture for a mean of 6 days (Table 4) to yield the P0 population. Upon harvest by trypsin digestion, a mean of 247,401 adherent P0 cells (Table 1) were obtained per milliliter of lipoaspirate tissue. These values are comparable to previous studies . Cells were maintained through an additional four successive passages of 6 to 7 days each. During each passage, the cell doubling times ranged between 3.6 and 4.7 days (Table 4).2 h5 r% c/ [; Z) q5 [

Table 4. Mean cell doubling times and passage lengths
0 j/ r8 F7 _/ \$ C  N
Immunophenotype
9 g  j7 Q+ n* S3 i+ l" V. R
Flow cytometric analysis was performed on cells cryopreserved after each stage of purification and passage (Table 5); representative flow histograms are shown in Figure 2. The initial SVF cells contained a subset of cells that were positive for a panel of endothelial cell¨Cassociated markers, including CD31, CD144 (VE-cadherin), VEGFr2, and von Willebrand factor (Table 5, Fig. 2). The levels of these markers did not change significantly through passage 4 (P4).
% Q: k0 O$ Z1 R; @. f1 N
Table 5. Phenotypic characterization of human adipose-derived cells at progressive stages of isolation and passagea
( \8 b2 p9 V/ I; w* L' t
Figure 2. Flow cytometry histogram of adipose-derived cells. The flow cytometry histograms for selected hematopoietic, stem cell, and stromal cell markers from a representative donor are displayed at the stromal vascular fraction (SVF) and passage 2 (P2) stages. The percentage of cells staining positive is indicated in the upper right corner of each panel. The blue line indicates the positive staining cells, whereas the red line indicates the isotype-matched monoclonal antibody control.- Z6 R/ \9 f" A: U* h
. F# f/ K; G% C! e, v0 p& Y
Only a subset of the initial SVF cell population expressed stromal cell-associated markers (Table 5, Fig. 3). Less than 1% of the SVFs expressed the activated lymphocyte common adhesion molecule (CD166), whereas 64% of the SVFs expressed the hyaluronate receptor (CD44); the levels of CD29, CD73, CD90, and CD105 were intermediate to these values. With successive passages, the percentage of cells staining positive for each of these markers increased, rising to between 69% (CD166) and 98% (CD44) by passage 4 (P4).

Figure 3. Relative change in the immunophenotype of adipose-derived cells as a function of purification and passage. The percentage of positive-staining cells is displayed relative to the isolation stage and passage number. (A): Stromal cell¨Cassociated markers CD166, CD73, CD44, and CD29. (Note that the order of the passage numbers is reversed in panel A relative to panel B.) (B): Stem cell¨Cassociated markers ABCG2 and CD34.7 h/ }2 o3 @+ h: k+ t# ]

The initial SVF contained a subpopulation of cells positive for stem cell¨Cassociated markers. A mean of 60% of the SVFs expressed the HSC-associated marker CD34, a sialomucin and L-selectin ligand . Although these levels increased during passage P1 and decreased in subsequent passages, the changes were not statistically significant relative to the SVFs.

High levels of the enzyme ALDH (ALDHbr) have proven to be a novel marker for the identification and isolation of HSCs . Based on flow cytometric analysis using a fluorescent substrate, the adipose-derived cells contained an ALDHbr sub-population (Table 5, Fig. 4). Although the ALDH levels were low in the SVF cells, the percentage of ALDHbr reached >70% between passages P0 to P4, with mean fluorescent intensities of 114 to 306 (data not shown). The percentage of ALDHbr ASCs fell to 10% when the cells were maintained in culture up to P9 (data not shown).4 f: q% z4 h8 q# ?8 U8 `* n
2 s, [2 m# g  F8 B
Figure 4. Aldehyde dehydrogenase staining of adipose-derived cells as a function of purification and passage.
. f  H, f1 Q/ F2 I8 G! E6 n6 s
DISCUSSION
5 |2 |6 L# u$ m+ g+ P* R4 \
Previous studies from our laboratory and by other groups have defined the immunophenotype of plastic adherent ASCs at passage 2 or later . Since these authors used an enrichment step with the STRO-1 antibody, these are at least three orders of magnitude less than those currently reported for human adipose tissue. Thus, the abundance of CFU-F in adipose tissue is substantially greater than that of bone marrow.8 i% Z0 ~) a. U4 _. _

The frequencies of CFU-Ad and CFU-Ob in adipose tissue were comparable to those of the CFU-F; however, the incidence of CFU-ALP was approximately one order of magnitude less frequent. Alkaline phosphatase enzyme activity has been used as a defining characteristic of bone marrow osteoblast progenitors and Westin-Bainton stromal cells . The current study measured alkaline phosphatase activity after 9 days in culture, whereas Alizarin red staining was performed after an additional 14 to 21 days. Since robust alkaline phosphatase staining was associated with multi-tiered cell layers (Fig. 1), it is possible that the frequency of CFU-ALP would have been closer to that of CFU-F and CFU-Ob if it had been assessed after an extended culture period. Progressive passage increased the frequency of CFUs for all lineages examined. By passage 4, CFU-F and CFU-Ob accounted for 33% of the adherent cells whereas CFU-Ad and CFU-ALP represented 16% and 8% of the population, respectively (Table 3). These findings indicate that the adhesion and expansion process enriches for a select functional cell phenotype.4 M& R/ K% p! @( e) y5 y
$ `7 }5 @( A4 }" s1 a% N
Multiple groups have begun to isolate adipose-derived cells for both in vitro and in vivo applications; however, the degree of consistency between laboratories with respect to the isolation and characterization of the cell population under investigation remains unclear. Recent studies have focused on adipose tissue-derived cells at earlier stages of isolation, using the SVF or adherent cells at early passage number . Likewise, we find that the SVF cell population includes hematopoietic lineage cells based on their expression of CD11, CD14, CD45, and other markers (data reported in accompanying paper); however, their expression is lost with progressive passage, suggesting that they do not account for the adherent cell population.

The levels of stem cell¨Cassociated markers (CD34, ABCG2, ALDHbr) reach their peak in the earliest stages of culture (passages 0/1). Consistent with one of these findings, we have documented the presence of mitochondrial ALDH by tandem mass spectroscopy proteomic analysis of undifferentiated and adipocytes-differentiated human ASCs .
9 d  ?4 r% G: w/ ?+ g$ O' F
In the earliest stages of isolation, the cells of the SVF exhibit low levels of stromal-associated markers (CD13, CD29, CD44, CD73, CD90, CD105, CD166). By the later stages of culture (passages 3/4), the cells assume a more homogeneous profile with consistently high levels of stromal markers. Overall, this temporal expression pattern resembles that reported for human bone marrow¨Cderived MSCs . These findings are consistent with the current immunophenotypic characterization of the adipose-derived cells at various stages of isolation and expansion.
# M; v* l4 I& S% N, g/ d
CONCLUSION5 {" A( m0 E2 S* w0 Z% h
3 o: b* X5 B% j. A% W# s! I/ @7 l! c
We have examined cells derived from human adipose tissue based on adherence characteristics and immunophenotype. The initially isolated SVF cells are heterogeneous; however, only 1 of 30 cells actually adheres to plastic and can account for the subsequent expansion of those cells now termed ASCs. The frequency of adipocyte and osteoblast progenitors in the SVF is comparable to that of the adherent cell population. This close correlation between CFU-F, CFU-Ad, and CFU-Ob data are consistent with studies from our laboratory and others demonstrating the presence of bipotent and tripotent clonal cells in human adipose tissue , these findings have implications concerning the potential utility of human adipose tissue as a source of adult stem cells for regenerative medical therapies.$ z9 U& B% e, R- G6 K+ F2 a

ACKNOWLEDGMENTS
: J9 I' w+ S% ?
We thank the Pennington Biomedical Research Foundation for financial support; our colleagues Drs. Barbara Kozak, Ken Eilertsen, Rachel Power, Randy Mynatt, and Farshid Guilak for critical review of the manuscript; and Drs. Elizabeth Clubb and James Wade, their office and nursing staffs, and their patients for donating and providing liposuction waste material to these studies." ?5 T. N( C6 _0 L9 N# _( j- w

DISCLOSURES

J.G. and X.W. have financial interest in Artecel Sciences. S.G., J.G., K.M., J.M., and X.W. have financial interest in Cognate Therapeutics.
      【参考文献】9 A7 }& k$ p3 u( B

Kondo M, Wagers AJ, Manz MG et al. Biology of hematopoietic stem cells and progenitors: Implications for clinical application. Annu Rev Immunol 2003;21:759¨C806.6 Q; A( N. g" @! Q5 s; D  F

Caplan AI. Mesenchymal stem cells. J Orthop Res 1991;9:641¨C650.6 ?( [0 v) F6 y" a9 q2 R
1 o6 Q: Y% Z+ \  j7 B) M
Friedenstein AJ. Precursor cells of mechanocytes. Int Rev Cytol 1976; 47:327¨C359./ j0 O  u6 J- \3 ?( Y! g) b
7 ]# N+ R5 u1 \1 w+ w
Gimble JM, Nuttall ME. Bone and fat: Old questions, new insights. Endocrine 2004;23:183¨C188.
  ?4 y' T( ?4 [3 ^7 d, k, g) J, s
Jiang Y, Vaessen B, Lenvik T et al. Multipotent progenitor cells can be isolated from postnatal murine bone marrow, muscle, and brain. Exp Hematol 2002;30:896¨C904.0 @" a/ Z( H0 ]

Ferraris C, Chevalier G, Favier B et al. Adult corneal epithelium basal cells possess the capacity to activate epidermal, pilosebaceous and sweat gland genetic programs in response to embryonic dermal stimuli. Development 2000;127:5487¨C5495.

Jones PH, Harper S, Watt FM. Stem cell patterning and fate in human epidermis. Cell 1995;80:83¨C93.
, G. [2 o0 j1 q
Fuchs E, Segre JA. Stem cells: A new lease on life. Cell 2000;100:143¨C155.

Gawronska-Kozak B. Regeneration in the ears of immunodeficient mice: Identification and lineage analysis of mesenchymal stem cells. Tissue Eng 2004;10:1251¨C1265.: A/ D5 V' j5 I9 t( m) K0 ^# U. v- G
% e7 e2 i: A8 y/ l
Gage FH, Ray J, Fisher LJ. Isolation, characterization, and use of stem cells from the CNS. Annu Rev Neurosci 1995;18:159¨C192.. l$ q$ A( X7 G+ K  F
% `) V+ m  v9 B0 e
Lechner A, Habener JF. Stem/progenitor cells derived from adult tissues: Potential for the treatment of diabetes mellitus. Am J Physiol Endocrinol Metab 2003;284:E259¨CE266.
! e4 r/ g# j; L
Jankowski RJ, Deasy BM, Huard J. Muscle-derived stem cells. Gene Ther 2002;9:642¨C647.( P! h, |, T# M$ K9 v
  w* _, h$ k1 B& x0 O. [* d8 C
Cao B, Zheng B, Jankowski RJ et al. Muscle stem cells differentiate into haematopoietic lineages but retain myogenic potential. Nat Cell Biol 2003;5:640¨C646.
4 z; |; d2 c: c% O1 \/ Z: }
Gimble JM, Guilak F. Differentiation potential of adipose derived adult stem (ADAS) cells. Curr Top Dev Biol 2003;58:137¨C160.

Halvorsen YC, Wilkison WO, Gimble JM. Adipose-derived stromal cells¨Ctheir utility and potential in bone formation. Int J Obes Relat Metab Disord 2002;24(suppl 4):S41¨CS44." K+ j$ `$ a3 i" n
) p  q  ]& W+ s: H6 u5 P+ w
Halvorsen YD, Bond A, Sen A et al. Thiazolidinediones and glucocorticoids synergistically induce differentiation of human adipose tissue stromal cells: Biochemical, cellular, and molecular analysis. Metabolism 2001;50:407¨C413.

Halvorsen YD, Franklin D, Bond AL et al. Extracellular matrix mineralization and osteoblast gene expression by human adipose tissue-derived stromal cells. Tissue Eng 2001;7:729¨C741.

Hicok KC, Du Laney TV, Zhou YS et al. Human adipose-derived adult stem cells produce osteoid in vivo. Tissue Eng 2004;10:371¨C380.
& y+ y2 {( l- u* T+ G, y, n, {) Q
Justesen J, Pedersen SB, Stenderup K et al. Subcutaneous adipocytes can differentiate into bone-forming cells in vitro and in vivo. Tissue Eng 2004;10:381¨C391.
2 a* E4 t' K+ E! O) V8 ]" Y
Erickson GR, Gimble JM, Franklin DM et al. Chondrogenic potential of adipose tissue-derived stromal cells in vitro and in vivo. Biochem Biophys Res Commun 2002;290:763¨C769.
$ I- c6 v/ n$ k
Safford KM, Hicok KC, Safford SD et al. Neurogenic differentiation of murine and human adipose-derived stromal cells. Biochem Biophys Res Commun 2002;294:371¨C379.
1 ^7 G/ M" m9 W. U% C: F
Safford KM, Safford SD, Gimble JM et al. Characterization of neuronal/glial differentiation of murine adipose-derived adult stromal cells. Exp Neurol 2004;187:319¨C328.

Zuk PA, Zhu M, Ashjian P et al. Human adipose tissue is a source of multipotent stem cells. Mol Biol Cell 2002;13:4279¨C4295.

Zuk PA, Zhu M, Mizuno H et al. Multilineage cells from human adipose tissue: Implications for cell-based therapies. Tissue Eng 2001;7:211¨C228.3 l; q* V# T: b9 E6 q) [  z
: K! C# B( e- J$ C
Mizuno H, Zuk PA, Zhu M et al. Myogenic differentiation by human processed lipoaspirate cells. Plast Reconstr Surg 2002;109:199¨C209.9 s6 Y8 @1 ~6 y8 N0 i$ u+ l

Seo MJ, Suh SY, Bae YC et al. Differentiation of human adipose stromal cells into hepatic lineage in vitro and in vivo. Biochem Biophys Res Commun 2005;328:258¨C264.
# i: o+ `5 s7 f5 N5 h3 L1 |) h
Gimble J, Guilak F. Adipose-derived adult stem cells: Isolation, characterization, and differentiation potential. Cytotherapy 2003;5:362¨C369.

Brzoska M, Geiger H, Gauer S et al. Epithelial differentiation of human adipose tissue-derived adult stem cells. Biochem Biophys Res Commun 2005;330:142¨C150.6 E" S1 B: ?: [; u) e5 v; L* g. I

Rodbell M. Metabolism of isolated fat cells, II: The similar effects of phospholipase C (Clostridium perfringens alpha toxin) and of insulin on glucose and amino acid metabolism. J Biol Chem 1966;241:130¨C139.

Rodbell M. The metabolism of isolated fat cells, IV: Regulation of release of protein by lipolytic hormones and insulin. J Biol Chem 1966;241:3909¨C3917.8 D$ j) P1 w2 u# H% C
# G6 v0 J- R* L8 V$ O, k
Rodbell M, Jones AB. Metabolism of isolated fat cells, 3: the similar inhibitory action of phospholipase C (Clostridium perfringens alpha toxin) and of insulin on lipolysis stimulated by lipolytic hormones and theophylline. J Biol Chem 1966;241:140¨C142.

Hauner H, Entenmann G, Wabitsch M et al. Promoting effect of glucocorticoids on the differentiation of human adipocyte precursor cells cultured in a chemically defined medium. J Clin Invest 1989;84:1663¨C1670.
5 |# P. ~% [" L. E5 q
Katz AJ, Tholpady A, Tholpady SS et al. Cell surface and transcriptional characterization of human adipose-derived adherent stromal (hADAS) cells. STEM CELLS 2005;23:412¨C423.

Miranville A, Heeschen C, Sengenes C, et al. Improvement of postnatal neovascularization by human adipose tissue-derived stem cells. Circulation 2004;110:349¨C355.4 V. I; R& n9 U7 o4 z! C* {

Bellows CG, Aubin JE. Determination of numbers of osteoprogenitors present in isolated fetal rat calvaria cells in vitro. Dev Biol 1989;133: 8¨C13.

Wu X, Peters JM, Gonzalez FJ et al. Frequency of stromal lineage colony forming units in bone marrow of peroxisome proliferator-activated receptor-alpha-null mice. Bone 2000;26:21¨C26.! v7 E& W& R: @5 e* b, l, L6 b

Castro-Malaspina H, Gay RE, Resnick G et al. Characterization of human bone marrow fibroblast colony-forming cells (CFU-F) and their progeny. Blood 1980;56:289¨C301.! [( e; H+ Z2 V
( X$ V5 u/ D- I/ Z  K2 F0 l
Aust L, Devlin B, Foster SJ et al. Yield of human adipose-derived adult stem cells from liposuction aspirates. Cytotherapy 2004;6:7¨C14.5 J1 F; R2 C+ f6 r( O$ h
2 B8 R/ F; U2 v/ l! n. ~; u
Sen A, Lea-Currie YR, Sujkowska D et al. Adipogenic potential of human adipose derived stromal cells from multiple donors is heterogeneous. J Cell Biochem 2001;81:312¨C319.

Shailubhai K, Streeter PR, Smith CE et al. Sulfation and sialylation requirements for a glycoform of CD34, a major endothelial ligand for L-selectin in porcine peripheral lymph nodes. Glycobiology 1997;7:305¨C314.$ R0 c  e7 P- G

McIntosh K, Zvonic S, Garrett S et al. The immunogenicity of human adipose-derived cells: Temporal changes in vitro. STEM CELLS (in press).# ]* V/ W0 J/ L

Goodell MA, Brose K, Paradis G et al. Isolation and functional properties of murine hematopoietic stem cells that are replicating in vivo. J Exp Med 1996;183:1797¨C1806.

Storms RW, Trujillo AP, Springer JB et al. Isolation of primitive human hematopoietic progenitors on the basis of aldehyde dehydrogenase activity. Proc Natl Acad Sci U S A 1999;96:9118¨C9123.& f$ z$ g+ m- ^# }
, A. p3 k3 c2 c" `) ^4 ~" Z
Fallon P, Gentry T, Balber AE et al. Mobilized peripheral blood SS-CloALDHbr cells have the phenotypic and functional properties of primitive haematopoietic cells and their number correlates with engraftment following autologous transplantation. Br J Haematol 2003;122:99¨C108.% I; C0 @& S  U$ X. ?
! I' N, o+ u, o& a+ U
Storms RW, Green PD, Safford KM et al. Distinct hematopoietic progenitor compartments are delineated by the expression of aldehyde dehydrogenase and CD34. Blood 2005;106:95¨C102.% R/ ?% o" Q% ^
. F6 K, s. N: R! {3 N
Gronthos S, Franklin DM, Leddy HA et al. Surface protein characterization of human adipose tissue-derived stromal cells. J Cell Physiol 2001;189:54¨C63.
/ c2 R( {9 h! ]1 ^/ a. }
Pittenger MF, Mackay AM, Beck SC et al. Multilineage potential of adult human mesenchymal stem cells. Science 1999;284:143¨C147.  x- L% v' ], l7 ^6 L
- T9 }# l9 h0 Y+ m0 W! V3 T
Guilak F, Lott KE, Awad HA et al. Clonal analysis of the differentiation potential of human adipose-derived adult stem cells. J Cell Physiol 2006;206:229¨C237.
" N: y$ x3 d& z
Stenderup K, Justesen J, Eriksen EF et al. Number and proliferative capacity of osteogenic stem cells are maintained during aging and in patients with osteoporosis. J Bone Miner Res 2001;16:1120¨C1129.; N( Z+ r+ A$ k
' S5 E5 B8 w9 o+ w
Friedenstein AY. Induction of bone tissue by transitional epithelium. Clin Orthop Relat Res 1968;59:21¨C37.& g7 f) ~6 M, h" W2 k7 K) a

Westen H, Bainton DF. Association of alkaline-phosphatase-positive reticulum cells in bone marrow with granulocytic precursors. J Exp Med 1979;150:919¨C937.! b, X6 M& d; H( ^7 T
+ ~, y: E. Y! V3 O/ e$ x
Planat-Benard V, Silvestre JS, Cousin B et al. Plasticity of human adipose lineage cells toward endothelial cells: Physiological and therapeutic perspectives. Circulation 2004;109:656¨C663.7 b& w' T$ q" [& k  h1 W
; |  e, L: O, |3 U4 P
Rehman J, Traktuev D, Li J et al. Secretion of angiogenic and antiapoptotic factors by human adipose stromal cells. Circulation 2004;109: 1292¨C1298.
" _2 m( n5 s1 M, l' g, w0 h( A
Martinez-Estrada OM, Munoz-Santos Y, Julve J et al. Human adipose tissue as a source of Flk-1  cells: New method of differentiation and expansion. Cardiovasc Res 2005;65:328¨C333.% o4 `% x+ d) y% c

Cousin B, Andre M, Arnaud E et al. Reconstitution of lethally irradiated mice by cells isolated from adipose tissue. Biochem Biophys Res Commun 2003;301:1016¨C1022.
* C1 }7 i8 `; n# t2 e
Cousin B, Andre M, Casteilla L et al. Altered macrophage-like functions of preadipocytes in inflammation and genetic obesity. J Cell Physiol 2001;186:380¨C386.

Cousin B, Munoz O, Andre M et al. A role for preadipocytes as macrophage-like cells. FASEB J 1999;13:305¨C312.3 D- V+ T% v+ D$ {
3 i) K" }2 K4 b3 ]/ K8 u% r7 Z
Charriere G, Cousin B, Arnaud E et al. Preadipocyte conversion to macrophage: Evidence of plasticity. J Biol Chem 2003;278:9850¨C9855., a( Q$ y/ _4 }9 M

DeLany J, Floyd ZE, Zvonic S et al. Proteomic analysis of primary cultures of human adipose derived stem cells: Modulation by adipogenesis. Mol Cell Proteomics 2005;4:731¨C740.+ P3 `; Q- ^. z  j1 ]
3 N8 h/ z' J; j% I
Barry F, Boynton R, Murphy M et al. The SH-3 and SH-4 antibodies recognize distinct epitopes on CD73 from human mesenchymal stem cells. Biochem Biophys Res Commun 2001;289:519¨C524.; _* J; m, w2 A' j: T

Barry FP, Boynton RE, Haynesworth S et al. The monoclonal antibody SH-2, raised against human mesenchymal stem cells, recognizes an epitope on endoglin (CD105). Biochem Biophys Res Commun 1999; 265:134¨C139.
& }+ u8 g+ ]. W& k8 g% G
Rubio D, Garcia-Castro J, Martin MC et al. Spontaneous human adult stem cell transformation. Cancer Res 2005;65:3035¨C3039.

收藏0 分享0 顶0 踩0