JOURNAL OF CELLULAR PHYSIOLOGY 208:64–76 (2006)

Characterization of Freshly Isolated and Cultured Cells
Derived From the Fatty and Fluid Portions of
Liposuction Aspirates
KOTARO YOSHIMURA,1* TOMOKUNI SHIGEURA,2 DAISUKE MATSUMOTO,1 TAKAHIRO SATO,2YASUYUKI TAKAKI,1 EMIKO AIBA-KOJIMA,1 KATSUJIRO SATO,1 KEITA INOUE,1 TAKASHI NAGASE,1
ISAO KOSHIMA,1 AND KOICHI GONDA1
1
Department of Plastic Surgery, University of Tokyo School of Medicine,
Bunkyo-ku, Tokyo, Japan2
Department of Research and Development, Biomaster, Inc.,
Imperial Tower 12F, Uchisaiwai-cho, Chiyoda-ku, Tokyo, Japan
Liposuction aspirates (primarily saline solution, blood, and adipose tissue fragments) separate into fatty and fluid portions. Cells
isolated from the fatty portion are termed processed lipoaspirate (PLA) cells and contain adipose-derived adherent stromalcells (ASCs). Here we define cells isolated from the fluid portion of liposuction aspirates as liposuction aspirate fluid (LAF) cells.
Stromal vascular fractions (SVF) were isolated separately from both portions and characterized under cultured and non-cultured
conditions. A comparable number of LAF and PLA cells were freshly isolated, but fewer LAF cells were adherent. CD34þCD45
cells from fresh LAF isolates were expanded by adherent culture, suggesting that LAF cells contain ASCs. Although freshly isolated
PLA and LAF cells have distinct cell surface marker profiles, adherent PLA and LAF cells have quite similar characteristics withregard to growth kinetics, morphology, capacity for differentiation, and surface marker profiles. After plating, both PLA and LAF cells
showed significant increased expression of CD29, CD44, CD49d, CD73, CD90, CD105, and CD151 and decreased
expression of CD31 and CD45. Multicolor FACS analysis revealed that SVF are composed of heterogeneous cell populations
including blood-derived cells (CD45þ), ASCs (CD31CD34þCD45CD90þCD105CD146), endothelial (progenitor) cells(CD31þCD34þCD45 CD90þCD105lowCD146þ), pericytes (CD31CD34CD45CD90þCD105CD146þ), and other cells.
After plating, ASCs showed a dramatic increase in CD105 expression. Although some adherent ASCs lost CD34 expression withincreasing culture time, our culture method maintained CD34 expression in ASCs for at least 10–20 weeks. These results suggestthat liposuction-derived cells may be useful and valuable for cell-based therapies. J. Cell. Physiol. 208: 64–76, 2006.
ß 2006 Wiley-Liss, Inc.

The stromal vascular fraction (SVF) isolated from
adipose tissue contains adipogenic progenitors withfibroblast-like morphologies (Van et al., 1976). Thesecells have been referred to by various names including
preadipocytes and vascular stromal cells. In this study,
we refer to the adherent stromal cells isolated from
adipose tissue as adipose-derived stromal (or stem) cells(ASCs). Monoclonal (Zuk et al., 2002) and polyclonal (Zuket al., 2001) culture studies have shown that humanASCs can be obtained from liposuction aspirates and candifferentiate into multiple lineages of mesodermal or
ectodermal origin. In both in vitro and in vivo studies,human ASCs have been shown to differentiate not only
into mesenchymal lineages, such as adipogenic, chondrogenic (Erickson et al., 2002; Awad et al., 2003; Huang
et al., 2004), osteogenic (Dragoo et al., 2003; Cowan et al.,
2004; Halvorsen et al., 2004; Hicok et al., 2004; Peterson
et al., 2005), myogenic (Mizuno et al., 2002; Rodriguez
et al., 2005), and cardiomyogenic (Planat-Benard et al.,2004a; Strem et al., 2005) lines, but also into neurogenic(Safford et al., 2002; Ashjian et al., 2003; Kang et al.,2003), angiogenic (Miranville et al., 2004; Rehman et al.,
2004; Planat-Benard et al., 2004b; Cao et al., 2005), andhepatic (Seo et al., 2005) lineages.
Liposuction is one of the most popular cosmetic
surgical procedures; worldwide, an estimated one
million liposuctions are performed annually. Althoughliposuction yields a large volume (e.g., 1 L) of adipose
tissue and is considered the typical method for clinicallyharvesting ASCs, liposuction aspirates have not beenß 2006 WILEY-LISS, INC.

well researched for use in clinical situations. Liposuction aspirates are comprised of fatty and fluid portions(Fig. 1a). The fatty portion consists of suctioned adipose
tissue that has been ‘‘shredded’’ by the reciprocal
movement of a metal canulla and vacuum pressure(500–700 mm Hg), while the fluid portion is the liquid
aspirated along with the fatty portion. The fluid portionis primarily composed of (1) a saline solution preoperatively injected into the site to prevent nerve andblood vessel damage, (2) peripheral blood, and (3) cells or
tissue fractions derived from adipose tissue. AlthoughThis article includes Supplementary Material available fromthe authors upon request or via the Internet at http://www.
interscience.wiley.com/jpages/0021-9541/suppmat
Kotaro Yoshimura and Tomokuni Shigeura contributed equally tothe study.
Contract grant sponsor: Japanese Ministry of Education, Culture,
Sports, Science, and Technology (MEXT); Contract grant number:
B2-16390507.
*Correspondence to: Kotaro Yoshimura, Department of Plastic
Surgery, University of Tokyo School of Medicine, 7-3-1, Hongo,
Bunkyo-ku, Tokyo 113-8655, Japan.
E-mail: yoshimura-pla@h.u-tokyo.ac.jp
Received 12 November 2005; Accepted 31 January 2006
DOI: 10.1002/jcp.20636

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65

Fig. 1. Isolation of ASCs from the fatty and fluid portions ofliposuction aspirates. a: The two portions of liposuction aspirates ina bottle. The fatty portion floats above the denser fluid portion. Cells
isolated from the fatty portion were named PLA cells; those isolated
from the liquid portion were termed LAF cells. b: Primary cultures offreshly isolated PLA and LAF cells. Scale bar ¼ 100 mm. c: Cell yields
of freshly isolated PLA and LAF cells. No statistically significant

difference in yield was detected. d: Adherent cell yields of cultured
PLA and LAF cells at 1 week. A significantly higher yield of adherent
PLA cells was detected. *P < 0.05. e: Doubling times of adherent PLA
and LAF cells. A statistically significant difference was seen between
the two serum concentrations (10% and 15%) in both PLA and LAF
cells, but not between PLA and LAF cells at either serum concentration, *P < 0.05.

only the fatty portion of liposuction aspirates has
been investigated to date, we recognized through apreliminary survey that a significant amount of progenitor cells can be isolated from the liquid portion as
well. Cells isolated from the fatty portion have beentermed processed lipoaspirate (PLA) cells (Zuk et al.,2001, 2002), and here, we refer to cells isolated fromthe fluid portion as liposuction aspirate fluid (LAF)
cells.
ASCs are currently being used in clinical trials
including studies investigating bone defect (Lendeckel
et al., 2004) and rectovaginal fistula (Garcia-Olmo et al.,2005) treatments and soft tissue augmentation (ourunpublished data). ASCs can be used clinically withoutcell expansion because a sufficient number can beobtained directly by processing liposuction aspirates,

which are usually of a large volume. Furthermore, theuse of minimally manipulated fresh cells may lead tohigher safety and efficacy in actual treatments. Thus,
because freshly isolated ASCs are preferable to cultured
ones for cell-based therapies, especially in the initialstage of regenerative medicine, minimally manipulated
SVF from adipose tissue must be characterized in moredetail. The SVF is known to be comprised of a heterogeneous cell population, but the exact cell composition
remains to be determined.
In this study, the fatty and fluid portions of liposuction
aspirates were investigated as sources for ASCs. Cells
isolated from both portions were characterized underfresh and cultured conditions. In addition, long-termchanges in cell surface marker expression profiles were
determined for both cell populations.

Journal of Cellular Physiology DOI 10.1002/jcp

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YOSHIMURA ET AL.

MATERIALS AND METHODS
Human tissue sampling

After informed consent, we obtained liposuction aspirates
using an IRB-approved protocol from healthy female donors
aged 21 to 59 years who underwent liposuction of the abdomenor thighs. Liposuction aspirates were divided into two portions:a floating adipose portion (also called lipoaspirate) and adenser fluid portion (Fig. 1a). Both portions were used assources for PLA and LAF cells. For harvesting human vascularendothelial cells (HUVEC) and dermal fibroblasts, umbilicalcords and skin were obtained from separate donors underinformed consent. Human mesenchymal stem cells derived
from bone marrow (frozen at Passage 2) were purchased from
Cambrex Bio Science Walkersville, Inc. (Walkersville, MD)
and cultured with the same medium used for adipose-derived
cells.
Cell isolation

All chemicals were purchased from Wako Pure Chemicals(Osaka, Japan), unless otherwise stated.
PLA cells were separated from the fatty portions ofliposuction aspirates using a procedure modified from Zuk
et al. (2001). Briefly, the suctioned fat was digested with
0.075% collagenase in PBS for 30 min on a shaker at 378C.
Mature adipocytes and connective tissues were separated frompellets by centrifugation (800g, 10 min). Pellets were resuspended in erythrocyte lysis buffer (155 mM NH4Cl, 10 mM
KHCO3, 0.1 mM EDTA) and incubated for 5 min at room
temperature. The pellets were resuspended and passed
through a 100-mm mesh filter (Millipore, Billerica, MA).
LAF cells were harvested from the fluid portions ofliposuction aspirates. The suctioned fluid was centrifuged(400g, 10 min), and the pellets were resuspended in erythrocyte lysis buffer. After 5 min at room temperature, lysates werepassed through a 100-mm mesh filter. The pellets were thenprocessed for density gradient centrifugation with Ficoll (GEHealthcare Bio-Sciences, Piscataway, NJ). After centrifugation (800g, 20 min), cells at the gradient interface werecollected, washed with PBS, and passed through a 100-mm
mesh filter. For flow cytometry of freshly isolated LAF cells,
density gradient centrifugation was not conducted. Nucleatedcell counts were performed using a NucleoCounter (Chemometec, Allerod, Denmark).
Cell culture of adherent cells from adipose tissue

Freshly isolated PLA or LAF cells were plated in medium ata density of 5  106 nucleated cells/100-mm gelatin-coateddish. Cells were cultured at 378C, 5% CO2, in humid air. The
culture medium was M-199 containing 10% FBS, 100 IU
penicillin, 100 mg/ml streptomycin, 5 ng/ml heparin, and 2 ng/
ml acidic FGF. For measurement of doubling time, the same
medium was used with a serum concentration of 10% or 15%.
Primary cells were cultured for 7 days and were defined as‘‘Passage 0.’’ The medium was replaced every 3 days, and cells
were passaged every week. After primary culture for 7 days,
attached cells were passaged by trypsinization and plated in
the same medium at a density of 2,000 cells/cm2.
Induced differentiation of cultured PLA and LAF cells

Capacities to differentiate along adipogenic, chondrogenic,
and osteogenic lineages were examined. Seven days after
seeding PLA or LAF cells at Passage 3–5, cell differentiation
was initiated by replacing the M-199 culture medium. Cells
cultured in control medium (DMEM plus 100 IU penicillin and100 mg/ml streptomycin) containing 10% FBS were used as
negative controls. For adipogenic differentiation, confluent
cultures were incubated for 4 weeks in the control medium
containing 10% FBS supplemented with 0.5 mM isobutylmethylxanthine (Sigma, St. Louis, MO), 1 mM dexamethasone,
10 mM insulin (Sigma), and 200 mM indomethacin. Fixed cells(4% paraformaldehyde for 10 min) were washed with 60%
isopropanol and incubated for 15 min with Oil-Red O tovisualize lipid droplets. Cells were then washed with isopropanol and counterstained with hematoxylin.
Journal of Cellular Physiology DOI 10.1002/jcp

For chondrogenic differentiation, two types of evaluations
were performed. First, confluent cultures were incubatedfor 4 weeks with the control medium containing 1% FBS
supplemented with 6.25 mg/ml insulin, 10 ng/ml TGFb1, and
50 nM ascorbate-2-phosphate. Fixed cells (4% paraformaldehyde for 10 min) were washed with 3% acetic acid andincubated for 30 min with 1% Alcian Blue 8 GX (Sigma), 3%
acetic acid to visualize the extracellular matrix. Cells werethen washed with 3% acetic acid and counterstained with 0.1%
Nuclear Fast Red, 5% Al2(SO4)3 solution. Second, a micromass
culture system was used as previously reported (Johnstone
et al., 1998). Cells were pelleted in a 15-ml tube and cultured
with the chondrogenic medium for 3 weeks.
For osteogenic differentiation, cells were incubated for
4 weeks in the control medium containing 10% FBS supplemented with 0.1 mM dexamethasone, 50 mM ascorbate-2phosphate, and 10 mM b-glycerophosphate (Nacalai Tesque,
Kyoto, Japan). Fixed cells (4% paraformaldehyde for 10 min)were washed and incubated with 2.5% silver nitrate for 20 min
in the dark. The cells were then washed, placed in the light for
15 min, and incubated for 2 min with 0.5% hydroquinone. Cells
were incubated for 2 min with 5% sodium thiosulphate tovisualize calcified deposits. The cells were then washed andcounterstained with 0.1% Nuclear Fast Red, 5% Al2(SO4)3
solution.
For adipogenic differentiation, colony-forming unit analysis
was also performed to quantify colonies with intracellularlipids. One hundred fifty cells were plated on a 100-mm culture
disk and cultured for 10–14 days, followed by incubation for
2 weeks with adipogenic medium and staining with Oil-Red O
as described above. The numbers of colonies positive or
negative for staining were counted under a microscope.
Flow cytometry and sorting

Freshly isolated PLA and LAF cells were examined forsurface and intracellular molecule expression using flow
cytometry. In addition, adherent PLA and LAF cells were
examined at weeks 1, 2, 4, 6, 8, 10, and 20 of cell culture. The
following monoclonal antibodies (MAbs) conjugated to fluorochromes were used: anti-CD4-FITC, CD10-PE, CD13-PE,
CD16-PE, CD29-PE, CD31-PE, CD34-PE, CD34-FITC, CD34PE Cy7, CD36-PE, CD44-PE, CD45-PE, CD45-FITC, CD49dPE, CD49e-PE, CD54-PE, CD56-PE, CD57-FITC, CD62E-PE,
CD62P-PE, CD69-FITC, CD73-PE, CD90-PE, CD106-FITC,
CD117-PE, CD135-PE, CD146-PE, CD-151-PE, HLA-A,B,CPE, Tie-2-PE (BD Biosciences, San Diego, CA), CD31-APC(eBioscience, San Diego, CA), CD144-PE (Beckman Coulter,
Fullerton, CA), CD59-PE (Ancell, Bayport, MN), CD71-PE,
CD105-PE (Serotec, Oxford, UK), CD133-PE, and Flk-1-PE(Techne, Minneapolis, MN). Irrelevant control MAbs were
included for all fluorochromes. Cells were incubated withdirectly conjugated MAbs for 30 min, then washed and fixed in
1% paraformaldehyde. Cells were analyzed using a LSR II(Becton Dickinson, San Jose, CA) or FACS Vantage SE (Becton
Dickinson) flow cytometry system. Data acquisition andanalysis were then performed (Cell Quest software, Becton
Dickinson). Gates were set based on staining with combinations of relevant and irrelevant MAbs so that no more than
0.1% of cells were positive using irrelevant antibodies. Cellsorting and subsequent analyses were performed using a
FACSAria cell sorter (Becton Dickinson).
Statistical analyses

Results were expressed as mean  SEM. Welch’s t-test wasused to compare each parameter.

RESULTS
Isolation and expansion of stromal cells from fatty
and fluid portions of liposuction aspirates

Compared to the fatty portion of liposuction aspirates,
the fluid portion contains more peripheral blood discharged from the suctioned site during liposuction. The
volume of peripheral blood varied among patients. The
adipose portion was subjected to collagenase digestion
followed by filtration for exclusion of extracellularCELLS DERIVED FROM LIPOSUCTION ASPIRATES

matrix (ECM) fragments and debris, while the fluid
portion was centrifuged and processed for lysis ofcontaminating erythrocytes. After plating on culture
dishes, non-adherent cells were discarded by changing
the culture medium. Both adherent PLA and LAF cells
had fibroblast-like morphologies and proliferated withsimilar doubling times, although a smaller number ofadherent cells were harvested from LAF cells than from
PLA cells (Fig. 1b).
Cell yields were normalized by dividing the isolated
cell number by the volume (in liters) of the fatty portionof the liposuction aspirate. Normalized numbers of
nucleate cells in SVF from the adipose (fresh PLA cells;
n ¼ 28) and fluid (fresh LAF cells; n ¼ 28) portions were1.31  0.50  109 and 1.55  0.79  109 per 1 L of adipose
portion (P ¼ 0.401), respectively (Fig. 1c). However, thecell number varied considerably among patients. Erythrocyte contamination was seen in both fresh PLA andLAF cells, although LAF cells apparently contained amuch greater number of cells derived from peripheral
blood. After 1 week of cell culture, normalized numbersof adherent PLA (n ¼ 28) and LAF (n ¼ 28) cells were
9.7  1.7  107 and 3.0  0.6  107 (P < 0.001), respectively (Fig. 1d). Thus, there was no significant differencein cell yield between freshly isolated PLA and LAF cells,
but there was a difference between adherent PLA andLAF cells cultured for 1 week.
PLA and LAF cells were cultured in medium with 10%(n ¼ 14) or 15% FBS (n ¼ 15), and doubling times were
measured using cells at Passage 0. Doubling times of
PLA and LAF cells were 28.5  1.7 h and 31.0  2.6 h,
respectively, when cultured with 15% FBS, and 43.3 
2.8 h and 40.2  3.0 h, respectively, when cultured with
10% FBS. A statistically significant difference indoubling time was observed between the two serum
concentrations in both PLA (P < 0.001) and LAF(P < 0.05) cells, but not between PLA and LAF cells at
either serum concentration (Fig. 1e).
In vitro differentiation of PLA and LAF cells

To compare the multipotency of PLA and LAF cells,
cell differentiation was induced using cells at Passage
3–5 by culturing the cells for 4 weeks with adipogenic,chondrogenic, or osteogenic medium. The results showed that both cell populations have similar capacities todifferentiate along the adipogenic (Fig. 2a), chondrogenic (Fig. 2b), and osteogenic lineages (Fig. 2c). In
addition, similar cartilage formation was observed by
the micromass system (Fig. 2d). Colony-forming unit
analysis showed that the percentage of cells staining
positive for Oil-Red O was 29.0  7.6% for PLA cells and24.1  4.3% for LAF cells (P ¼ 0.12) (Fig. 2d).
Flow cytometric analysis of PLA and LAF cells

Flow cytometric analysis revealed that freshly isolated LAF cells differ significantly in cell surface marker
expression from freshly isolated PLA cells. Compared to
fresh LAF cells, fresh PLA cells contained higher
percentages of cells positive for CD29 (b1-integrin),
CD34, and CD90 (Thy-1) expression and a decreased
percentage of CD45þ cells of hematopoietic origin.
Conversely, in fresh LAF cells, there were higher
percentages of CD31 (PECAM-1)þ cells and CD45þ
cells, suggesting that fresh LAF cells contained a larger
number of blood-derived cells than fresh PLA cells. In
the FACS plot of forward and side scatter characteristics(FSC and SSC) of fresh LAF cells, there were three
CD45þ cell clusters corresponding to granulocytes,
monocytes/macrophages, and lymphocytes (Figs. 3a,
Journal of Cellular Physiology DOI 10.1002/jcp

67S1). CD34þ cells and CD45 cells were located in
the cluster corresponding to monocytes/macrophages(Figs. 3a, S1). Double color analysis of fresh LAF cells for
CD34 and CD45 indicated that most of the CD34þ cells
were CD45, suggesting that most of the CD34þ cells
were not derived from peripheral blood but from adipose
tissue (Fig. 3b). In addition, CD34þCD45 cells, but not
CD34þCD45þ cells, proliferated on a culture dish,suggesting that the freshly isolated LAF cells containedASCs derived from adipose tissue (Fig. 3b). CD34þ cells(P2) were sorted and cultured using the protocol
described above, but after 2 weeks in culture, approximately half of the cells were negative for CD34 (Fig. 3b),suggesting that some CD34þ cells lost CD34 expression
with increased culture time. The CD34 cells did notreacquire CD34 expression by further culturing. The
doubling time of CD34 cells was significantly shorter
than that of CD34þ cells (Fig. 3c).
Subsequently, we performed multicolor FACS assays
to investigate the SVF (fresh PLA and LAF cells) in moredetail, especially with regard to cell composition. Even
after processing with hypotonic erythrocyte lysis buffer,
the SVF contained a large number of erythrocytes, soerythrocytes and platelets were excluded from analysis
by gating them out by cell size. Consequently, only
nucleated cells were analyzed (Figs. 4 and 5). The results
are summarized in Table 1. We classified freshly isolated
cells derived from liposuction aspirates into 11 cell
populations [4 blood-derived (CD45þ) and 7 adiposederived (CD45)] as follows. There were three major(>1%) and one minor (<1%) cell populations derived
from blood with regard to expression patterns of CD31,
CD34, CD45, CD105 (Endoglin), CD14, and CD15
(Figs. 5a, S2): (1) CD31lowCD34CD45þCD105low
CD14CD15þ cells (corresponding to granulocytes),(2) CD31lowCD34CD45þCD105lowCD14þCD15 cells(corresponding to monocytes/macrophages), (3) CD31
CD34CD45þCD105 cells (corresponding to lymphocytes), and (4) CD31CD34þCD45þCD105 cells(corresponding to hematopoietic stem cells). Cell compositions varied among samples, with the percentages ofthe first three populations dependent on the compositionof the peripheral blood of each sample. Furthermore, thetotal percentage of blood-derived populations in SVF
depends on the intraoperative hemorrhage volume ofeach sample.
There were seven adipose-derived populations withregard to expression patterns of CD31, CD34, CD45,
CD90, CD105, and CD146: (1) CD31CD34þCD45
CD90þCD105CD146 cells (corresponding to ASCs),(2) CD31þCD34þCD45CD90þCD105low CD146þ cells,(3) CD31CD34þCD45CD90þCD105CD146þ cells,(4) CD31CD34CD45CD90þCD105CD146þ cells,(5) CD31CD34þCD45CD90þCD105low cells, (6) CD31low
CD34þCD45CD90þCD105lowCD146þ cells, and (7)CD31CD34CD45CD90þCD105CD146 cells. The
first population corresponded to ASCs and comprised
70–90% of the total adipose-derived (CD45) cells. Most
ASCs were CD29þ, CD117 (c-kit), CD133/AC133,CD144 (VE-cadherin), and Flk-1(VEGFR-2) (Fig. 4a).
The second population was positive for both CD31 andCD146 (identified as P1 in Fig. 5a; also identified in
Figs. 4b and 5b), suggesting that it is composed ofvascular endothelial cells or endothelial progenitor cells
derived from adipose. The third (CD34þ) and fourth(CD34) populations were CD31CD146þ (Fig. 4a,b),
suggesting that they may correspond to pericyte
progenitors (CD34þ) and pericytes (CD34), respectively. The fifth population was very small and seemed

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YOSHIMURA ET AL.

Fig. 2. Cell differentiation analysis of adherent PLA and LAF cells.
Representative results are shown in a–d: PLA and LAF cells at
Passage 3–5 were similarly induced to differentiate into adipogenic,chondrogenic, and osteogenic lineages. a: Cultures under adipogenic
conditions. Adipogenic differentiation was induced, and lipid droplets
were visualized with Oil-Red O staining at 4 weeks. b: Cultures underchondrogenic conditions. Chondrogenic differentiation was induced
and visualized with Alcian blue staining at 4 weeks. c: Cultures under

osteogenic conditions. Osteogenic differentiation was induced andvisualized with von Kossa staining at 4 weeks. Scale bar ¼ 100 mm.
d: Micromass culture of PLA and LAF cells. Cartilage formation was
similarly observed using the micromass system. e: Colony-forming
unit analysis under adipogenic conditions. Between adherent PLA
and LAF cells, the difference in percentages of colonies thatdifferentiated into an adipogenic lineage was not statisticallysignificant.

to correspond to CD105-positive ASCs (P1 in Fig. 5b).
The sixth population was also small, with moderate
expression of CD31 (P2 in Fig. 5a). Finally, the seventh
population was positive only for CD90, suggesting that it
may be fibroblasts or ASCs that lost CD34 expression(P3 in Fig. 5a).
Additionally, a multi-color FACS assay was performed for cultured PLA cells (at 5 and 10 days;
Fig. 6a,b) and cultured BM-MSCs (Passage 3; Fig. 6c)
for comparison. Most PLA cells cultured for 5 days wereCD31CD34þCD45CD90þCD105þCD146, but small
percentages of other cell populations, such as CD31þ
CD34þCD45CD90þCD105þCD146þ cells (endothelial
cells or endothelial progenitors), CD31CD34CD45
CD90þCD105þCD146þ cells (possibly pericyte progenitors), and CD31CD34CD45CD90þCD105þCD146þ

cells (pericytes) were also observed. Unlike freshly
isolated ASCs, the major ASC population expressed
CD105. Compared to PLA cells cultured for 5 days, the10-day cultures showed decreased percentages of
CD31þ endothelial cells and CD34þ cells and increased
percentages of CD31CD34þCD146þ cells and CD31
CD34CD146þ cells. Cultured BM-MSCs showed similar surface marker expression patterns. Most cultured
BM-MSCs were identified as CD31CD34CD45
CD90þCD105þ cells with variable expression of
CD146. BM-MSCs contained only a very small percentage (<1%) of CD34þ cells, which was the only difference
from ASCs clearly detected in this assay.
Representative single-color FACS data for freshly
isolated and cultured (at 2 weeks) PLA and LAF cells areshown in Figure 7 and sequential changes in cell surface

Journal of Cellular Physiology DOI 10.1002/jcp

CELLS DERIVED FROM LIPOSUCTION ASPIRATES

69

Fig. 3. CD34 expression in freshly isolated and adherent LAF cells.
a: Flow cytometric analysis of freshly isolated LAF cells for cell size,
granularity, and expression of CD34 or CD45. In the FACS plot of SSC(granularity) versus FSC (cell size) for freshly isolated LAF cells,
CD34þ cells (left), and CD45 cells (right) are shown in green. Bothwere located in the same area as monocytes/macrophages, while
CD45þ cells (middle) were located in the areas typical for blood cells
such as granulocytes, monocytes/macrophages, and lymphocytes (seeFig. S1). b: Double color flow cytometric cell sorting of fresh LAF cellsby CD34 and CD45 expression, and the fates of sorted cells after
plating. Left: Most CD34þ cells from fresh LAF cells were CD45.
Middle: The CD34þCD45 (P1) and CD34þCD45þ (P2) cell populations were sorted and plated; only CD34þCD45 cells grew on the dish.
Scale bar ¼ 100 mm. Right: After 2 weeks’ culture, approximately halfof the CD34þCD45 cells had lost CD34 expression. c: Comparison of
doubling time between CD34þ and CD34 adherent LAF cells. The
doubling time of CD34þ adherent LAF cells was significantly shorter
than that of CD34 cells, *P < 0.05.

marker expression are shown in Figure 8. After plating,the surface marker expression profiles for both PLA andLAF cells changed markedly, and adherent cells of thetwo cell populations showed quite similar expression
profiles. The percentage of CD34þ cells increased in PLA
cells, which uniformly expressed mesenchymal markers, such as CD13, CD29, CD44, CD73, and CD90. In
PLA cells cultured for more than 1 week, CD10, CD49e,
CD59, and CD151 were also uniformly expressed. Oneweek culture of freshly isolated PLA or LAF cells
resulted in a dramatic enrichment in CD105 from

1.2  0.6 to 64.1  9.7 (P < 0.001) or from 1.4  0.7 to74.6  7.6 (P < 0.001), respectively. No statisticallysignificant difference in CD105 expression was observed
between adherent PLA and LAF cells. After 1 week inculture, CD45, Flk-1, Tie-2, CD31, CD117, and CD133/
AC133 expression had decreased significantly in PLA
and LAF cells. Both cell populations were negative for
CD4, CD45, CD62E (E selectin), CD62P (P selectin),
CD69, CD135, and CD144 at 1 week and were negative
for CD16, CD31, CD57, CD106 (VCAM-1), CD133, Flk-1,
and Tie-2 after culture for more than 2 weeks.

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Fig. 4. Multicolor FACS analysis of SVF (1). a: SVF from the adipose
portion of liposuction aspirates was analyzed for CD34, CD45, and oneof the following markers: CD29, CD90, CD105, CD117, CD133,
CD144, CD146, or Flk-1. CD34 and CD45 expression is shown in thegraph at left, with blood-derived CD45þ cells in red and adiposederived CD45 cells in blue. Only adipose-derived CD45 cells (bluedots) were plotted in each of the eight graphs at right. Most adiposederived CD34þ cells were CD29þ, CD90þ, CD105, CD117, CD133,CD144, and Flk-1. It should be noted that there are two populationsof adipose-derived CD34þ cells with regard to CD146 expression.
b: SVF from the adipose portion was analyzed for CD31, CD34, CD45,

and one of the following markers: CD90, CD105, or CD146. Only
adipose-derived CD45-cells were plotted. There are two major and one
minor populations of CD34þ cells. One major population (endothelial
cells or endothelial progenitors) was CD31þCD34þCD90þ
CD105lowCD146þ and comprised 2–8% of CD34þ cells. The largest
population (ASCs) was CD31CD34þCD90þCD105CD146. The
minor population was CD31CD34þCD90þCD105CD146þ. There
were two small populations of CD34 cells; one was CD31CD34
CD90þCD105CD146þ, and the other was CD31CD34CD90þ
CD105CD146.

CD34 expression decreased with increased culture
time, but 10–20% of cells maintained CD34 expression
up to 20 and 10 weeks’ culture in PLA and LAF cells,
respectively. In addition, consistent expression of themesenchymal markers (CD13, CD29, CD44, CD73, andCD90) as well as other markers (CD 49d, CD59, CD105,
and CD151) was observed in adherent PLA and LAF
cells up to 20 and 10 weeks, respectively. After the initial
2 weeks, no remarkable changes in surface marker
expression were seen between adherent PLA and LAF
cells throughout the culture periods. Taken together,
these data suggest that adherent PLA and LAF cells canbe expanded using our culture method without losing
stem cell-associated surface markers. Our studies
further revealed that after the initial 1–2 weeks inculture, adherent PLA and LAF cells have quite similar
surface marker expression profiles throughout theculture periods, suggesting that both cell populations
can be considered ASCs.

Journal of Cellular Physiology DOI 10.1002/jcp

DISCUSSION

Adipose tissue is comprised predominantly of matureadipocytes, connective tissue, ASCs, blood-derived cells,
vascular cells, such as endothelial (progenitor) cells,smooth muscle cells, and pericytes. Adipocytes represent roughly two-thirds of the total cell number andmore than 90% of the tissue volume (van Harmelen et al.,2005). The ratio of adipocytes to ASCs is constant in
humans, independent of body mass index (BMI) and age(van Harmelen et al., 2003). In the present study, acorrelation between ASC cell yield and age or BMI was
not detected (data not shown). ASC cell yield varies
among patients and is affected by many factors including donor site and storage duration. It also depends

Fig. 5. Multicolor flow cytometric analysis of SVF (2). Freshly
isolated SVF from the adipose portion was analyzed with multicolor
flow cytometry for CD31, CD34, CD45, and CD105. a: CD31 and CD45
expression is shown in the graph at left, with six major populations(P1–P6) identified. Each population was individually plotted for CD34
and CD105 expression in the six graphs at right. P1 was composed of
CD31þCD34þCD45CD105low cells, which are fresh endothelial cells
or their progenitors. P2 was an unknown minor population(CD31lowCD34CD45CD105low cells; possibly CD31low endothelial
cells). P3 contained ASCs, which are CD31CD34þCD45CD105.

P4 consisted predominantly of granulocytes and monocytes/macrophages, with a small number of lymphocytes. P5 and P6 consisted oflymphocytes (Fig. S2). b: CD45 and CD105 expression is shown in thegraph at left, and two adipose-derived (CD45) populations (P1:
CD34þ, P2: CD34) are identified. Each population was individuallyplotted for CD31 and CD34 expression in the graphs at right. P1 wascomprised of CD31þCD34þCD45CD105low cells (fresh endothelial
cells or progenitors) and CD31CD34þCD45CD105low cells. P2 wascomposed of other adipose-derived cells including ASCs.

TABLE 1. Summary of cell composition in SVF derived from liposuction aspirates

HSC, hematopoietic stem cell; ASC, adipose-derived stromal cell; fEC, fresh endothelial cell; EPC, endothelial progenitor cell; PC, pericytes; PPC, pericyte progenitor
cell.
Results of multi-color FACS assays are summarized. We classified freshly isolated cells from liposuction aspirates into 11 populations according to surface marker
expression profiles; 4 blood-derived (CD45þ; above) and 7 adipose-derived (CD45; below) cell populations. Cell composition percentages varied among samples, so a
range of values is presented.

Journal of Cellular Physiology DOI 10.1002/jcp

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YOSHIMURA ET AL.

Fig. 6. Multicolor FACS analysis of cultured PLA cells and cultured
BM-MSCs. PLA cells cultured for 5 days (a) and 10 days (b) and BMMSC (Passage 3) (c) were analyzed for CD31, CD34, CD45, and one offollowing markers: CD90, CD105, or CD146. a, b: In the graphs at far
left, all cells were plotted, while only CD45 cells (blue dots) are shownin the six graphs at right. There are several differences in surface
marker expressions between freshly isolated and cultured PLA cells;
Almost all ASCs started expressing CD105 after plating, and someASCs lost CD34 expression with increased culture time. CD31þ

endothelial cells decreased in percentage with culture time, while
CD31CD34CD146þ cells (possibly pericyte progenitors) and CD31
CD34CD146þ cells (pericytes) increased. CD31þCD90þ endothelial
cells observed in fresh SVF lost CD90 expression with increased
culture time, as seen in HUVEC (Fig. S3a, b). c: Nearly all BM-MSCs
were CD34. Red and blue dots are CD34þ and CD34 cells,
respectively. Cultured BM-MSCs and ASCs showed similar expression of CD31, CD45, CD90, CD105, and CD146, with the only
detectable difference being in CD34 expression.

strongly on the isolation method, for example, durationof collagenase digestion (Aust et al., 2004; Bakker et al.,
2004; von Heimburg et al., 2004).
Although only the adipose portion of liposuction
aspirates has been used as a source of ASCs, we also
isolated cells from the fluid portion of liposuction
aspirates and found that a comparable amount of ASCs
can be harvested. By flow cytometric analysis, fresh PLA
and LAF cells showed distinct surface marker profiles.
Higher percentages of CD31þ and CD45þ cells wereobserved in fresh LAF cells than in PLA cells, suggestingthat fresh LAF cells contain a larger amount of bloodderived cells. However, adherent LAF cells cultured for
1 week showed surface marker profiles quite similar to
cultured PLA cells, suggesting that the fluid portion ofliposuction aspirates also contains ASCs. In addition,
our results showed that most of the CD34þ cells infreshly isolated LAF cells were derived from adipose
tissue (CD45), and these CD34þCD45 cells expandedin culture dishes, suggesting that they correspond to thesame ASC population derived from PLA cells. Why asignificant amount of ASCs are isolated from the fluid
portion of liposuction aspirates has yet to be determined.
Nor is the location of ASCs in adipose tissue clearly
understood. Some ASCs are thought to be located in the
adipose connective tissue and others between adipocytes or around micro- or macro-vasculature. ASCs
located between adipocytes might be released into thefluid by mechanical injury during liposuction procedures, and other ASCs might be released by endogenous

Journal of Cellular Physiology DOI 10.1002/jcp

CELLS DERIVED FROM LIPOSUCTION ASPIRATES

73

Fig. 7. Comparison of flow cytometric data for freshly isolated and 2-week-cultured PLA and LAF cells.
Freshly isolated PLA and LAF cells displayed distinct cell surface marker profiles, while PLA and LAF
cells cultured for 2 weeks showed quite similar expression profiles. Adherent, but not freshly isolated,
PLA, and LAF cells expressed CD105. Results are representative of samples from five patients.

proteases during surgery or subsequent storage periods.
The extent of mechanical injury may vary among
patients because it can be affected by the size of thesuction canulla, vacuum pressure strength during
liposuction, suction procedure (manual or powered),and other factors.
One of the reasons why adipose tissue is thought to bea promising source of stem cells is that a large volume of
adipose tissue can be harvested with minimal morbidity.
Thus, ASCs can be used clinically after minimal
manipulation (without cell culture). Therefore, examJournal of Cellular Physiology DOI 10.1002/jcp

ination and characterization of freshly isolated PLA andLAF cells have significant clinical implications. Some
recent studies examined freshly isolated SVF fromhuman adipose tissue using magnetic cell sorting(Boquest et al., 2005; Sengenès et al., 2005), and theinvestigators partly characterized some cell populationsin the SVF. CD31CD34þCD45CD105þ cells were
proposed as typical proliferating ASCs (Boquest et al.,2005). Other studies showed that CD34þCD31
cells differentiated into endothelial cells and contributed to neovascularization (Miranville et al., 2004;

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YOSHIMURA ET AL.

Fig. 8. Sequential changes in representative cell surface marker expression in fresh and cultured PLA
and LAF cells. Statistically significant differences in expression of some cell surface markers (data notshown for CD16 and CD49e) were observed between freshly isolated PLA and LAF cells. These differenceswere not apparent after 1–2 weeks in culture. Data are shown as mean þ (LAF) or  (PLA) standard
error.

Planat-Benard et al., 2004b). For analysis of SVF, weused multicolor (four or five colors) flow cytometry
assays and showed that there are several major andminor cell populations derived from blood and adipose.
The typical surface marker expression pattern of freshASCs was CD31CD34þCD45CD90þCD105CD146,Journal of Cellular Physiology DOI 10.1002/jcp

which was distinct from that of cultured ASCs in CD105
expression only. This profile of fresh ASCs differs from aprevious study (Boquest et al., 2005), possibly because
the magnetic cell sorting procedure used in that study
might have affected CD105 expression. Our results also
showed that most fresh ASCs are CD29þ, CD90þ,CELLS DERIVED FROM LIPOSUCTION ASPIRATES

CD117, CD133, CD144, and Flk-1. It should be
noted that there are significant numbers of CD146þ andCD146 cells in both CD34þ and CD34 cell populationsin SVF. Given that CD31 is an endothelial cell marker
and that CD146 is a known marker of endothelial cells
and pericytes, CD31CD34CD146þ cells in SVF arelikely pericytes, and CD31CD34þCD146þ cells may
function as pericyte progenitors. Nearly all adiposederived CD31þ cells in SVF were found to be CD34þ, andthis population was previously reported as endothelial
cells (Sengenès et al., 2005). Our results showed that
this population was CD90þCD105lowCD146þ. However,a major proportion of cultured HUVECs (Passage 2)
were CD34 and CD90 (Fig. S3b), suggesting that freshbut not cultured endothelial cells express CD34 andCD90. Indeed, freshly isolated HUVEC cells wereCD34þ with variable expression of CD90 (Fig. S3a).
CD31CD34CD45CD90þCD105CD146 cells in
SVF may be fibroblasts, as suggested by comparison
with cultured dermal fibroblasts (Fig. S3c).
PLA and LAF cells showed similar surface marker
profiles after being cultured for 1 week. Notable changes
between fresh and cultured states include decreased
expression of CD31 and CD45 and increased expressionof CD29 and CD105. These changes suggest that cells
other than ASCs, such as vascular endothelial cells andblood-derived cells, are selectively excluded during
culturing on plastic plates. Even when cultured inendothelial cell growth medium, ASCs quickly outgrow
CD31þ endothelial cells (Hutley et al., 2001). As noted
above, a high percentage of ASCs began to express
CD105 after plating. Our results showed that theexpression of most surface markers is sustained
throughout the culture period (up to 20 weeks for PLA
and 10 weeks for LAF cells), and that significant changes
seen after the initial 2 weeks are limited to a gradualincrease in CD49d and a decrease in CD71.
Because CD34 is one of the most well established stem
cell markers, CD34 expression may indicate ASC
clinical usefulness. CD105, known as a mesenchymal
stem cell-associated marker, was highly expressed incultured ASCs and may reflect the capacity of ASCs todifferentiate into lineages of mesenchymal origin suchas adipose, cartilage, and bone. Flk-1, known to be
expressed in hemangioblasts, was recently shown to be
expressed in ASCs under certain culture conditions (Caoet al., 2005; Martinez-Estrada et al., 2005). These factors
suggest that ASCs may have clinical potential for cellbased therapies. A number of studies characterizing
human ASCs have reported that ASCs (freshly isolated
or cultured for less than 2 weeks) are CD34þ (Gronthos
et al., 2001; Miranville et al., 2004; Rehman et al., 2004;
Planat-Benard et al., 2004b; Boquest et al., 2005).
However, CD34 expression was not found in humanASCs cultured for more than 2 weeks with conventional
culture methods (Gronthos et al., 2001; Zuk et al., 2002;
De Ugarte et al., 2003; Lee et al., 2004; Rehman et al.,
2004; Boquest et al., 2005; Katz et al., 2005). In contrast,
our results showed that CD34þ ASCs were present at
10–20% of the cell population even after 10–20 weeks ofculture (Fig. 7). CD34þ cells were sorted and cultured,
but about half of the cells became CD34 after 2 weeks inculture (Fig. 3b), suggesting that cultured ASCs may
exist in a variety of stages ranging from CD34þ
undifferentiated cells to CD34 partially differentiatedcells. The result that CD34þ cells proliferated more
quickly than CD34 cells may explain the sustained
percentage of CD34þ cells in cultured PLA or LAF cells.
Thus, using our culture protocol, it is possible to expand
Journal of Cellular Physiology DOI 10.1002/jcp

75

CD34þ ASCs taken from liposuction aspirates up to
104 –107 times after 4 weeks’ culture. Together with thefact that Flk-1þ ASCs can be obtained in high percentages using another culture protocol (Cao et al., 2005;
Martinez-Estrada et al., 2005), ASCs may dramaticallychange their surface marker profiles depending on theculture media and methods.
It was reported that human ASCs differ from human
BM-MSCs in expression of CD49d (expressed only inASCs) and CD106 (expressed only in BM-MSCs) (Zuket al., 2002; De Ugarte et al., 2003). In our study using
multicolor assays with limited surface markers, the only
difference between cultured ASCs and cultured BMMSCs was CD34 expression (positive in ASCs andnegative in BM-MSCs). Human dermal fibroblasts havea surface marker expression profile similar to ASCs butlack expression of CD34, CD105, and CD146 (Fig. S3c).
Because BM-MSCs (Zuk et al., 2002; De Ugarte et al.,2003) and other stromal progenitors cultured for long
periods do not express CD34, ASCs may constitute a
unique mesenchymal cell population in view of their
CD34 and CD146 expression. This characteristic may
contribute to potentialities of ASCs other than mesenchymal progenitors, such as endothelial (Miranville et al.,
2004; Rehman et al., 2004; Planat-Benard et al., 2004b)
or pericyte progenitors.
In summary, adherent LAF cells have quite similar
characteristics with respect to growth kinetics, morphology, surface marker profiles, and capacity for
differentiation to adherent PLA cells. A significant
amount of ASCs can be isolated from the fluid portionof liposuction aspirates, although in smaller amounts
than from the fatty portion. In addition, we found that
SVF are composed of heterogeneous cell populations
including blood-derived cells, ASCs, endothelial (progenitor) cells, pericytes (and progenitors), and otherunknown progenitors. A major population of ASCs
in SVF was identified as CD31CD34þCD45CD90þ
CD105CD146 cells but began to express CD105 after
plating. Adipose-derived CD34þ ASCs can be expanded
for at least 20 weeks using our culture method. These
results suggest that liposuction-derived human ASCs
may have significant clinical utility for cell-based
therapies.
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JCP-05-0688(Suppl)
JOURNAL OF CELLULAR PHYSIOLOGY 9999:1–2 (2006)

Author Proof

Characterization of Freshly Isolated and Cultured Cells
Derived From the Fatty and Fluid Portions of
Liposuction Aspirates

A

COLOR

KOTARO YOSHIMURA,1* TOMOKUNI SHIGEURA,2 DAISUKE MATSUMOTO,1 TAKAHIRO SATO,2YASUYUKI TAKAKI,1 EMIKO AIBA,1 KATSUJIRO SATO,1 KEITA INOUE,1 TAKASHI NAGASE,1
ISAO KOSHIMA,1 AND KOICHI GONDA1
1
Department of Plastic Surgery, University of Tokyo School of Medicine,
Bunkyo-ku, Tokyo, Japan2
Department of Research and Development, Biomaster,Inc. Imperial Tower 12F, Uchisaiwai-cho, Chiyoda-ku, Tokyo, Japan

COLOR

Fig. S1. FSC and SSC analysis of peripheral blood. FACS analysis of peripheral blood by FSC (cell size)
and SSC (granularity). Blue dots are cells positive for each CD antigen: CD4 (helper T lymphocytes), CD8(cytotoxic T lymphocytes), CD11b (both granulocytes and monocytes/macrophages), CD14 (monocytes/
macrophages), CD15 (granulocytes), and CD19 (B lymphocytes). Granulocytes, monocytes/macrophages,
and lymphocytes are located predominantly in the upper, middle, and lower populations, respectively.

Kotaro Yoshimura and Tomokuni Shigeura contributed equally tothe study.
*Correspondence to: Kotaro Yoshimura, Department of Plastic
Surgery, University of Tokyo School of Medicine, 7-3-1, Hongo,
Bunkyo-ku, Tokyo 113-8655, Japan.
E-mail: yoshimura-pla@h.u-tokyo.ac.jp

Fig. S2. Multicolor FACS analysis of peripheral blood. Blue dots arecells positive for CD4 (helper T lymphocytes), CD8 (cytotoxic T
lymphocytes), CD14 (monocytes/macrophages), and CD15 (granulocytes). All blood-derived cells were CD45þ. Granulocytes and monocytes/macrophages were CD31þ, while lymphocytes were CD31.
ß 2006 WILEY-LISS, INC.

Received 12 November 2005; Accepted 31 January 2006
Published online in Wiley InterScience(www.interscience.wiley.com.), 00 Month 2006.
DOI: 10.1002/jcp.Suppl

2

YOSHIMURA ET AL.

COLOR

Author ProofA

Fig. S3. Multicolor FACS analysis of freshly isolated HUVEC,
cultured HUVEC, and cultured dermal fibroblasts. Freshly isolated
HUVEC (a), cultured HUVEC (EGM-2; Passage 2) (b), and cultured
dermal fibroblasts (DMEMþ10%FBS, Passage 4) (c) were analyzed for
CD31, CD34, CD45, and one of following markers: CD90, CD105, orCD146. a: All freshly isolated cells from human umbilical cord vein
were plotted in the graph at far left, showing contamination byblood-derived CD45þ cells (red dots). Only adipose-derived CD45
cells (blue dots) were shown in the six graphs at right. Most freshly
isolated HUVEC were CD31þCD34þCD45CD105þCD146þ, while

Journal of Cellular Physiology DOI 10.1002/jcp

CD90 expression varied. b: Most cultured HUVEC (Passage 2) wereCD31þCD34CD45CD90CD105þCD146þ. Small percentages of
CD31þCD34þCD45CD90CD105þCD146þ and CD31þCD34CD45
CD90þCD105þCD146þ cells were also detected. Red and blue dots are
CD34þ and CD34 cells, respectively. c: Most of the cultured dermal
fibroblasts were CD31CD34CD45CD90þCD105CD146, and asmall percentage was CD31CD34þCD45CD90þCD105CD146.
Red and blue dots are CD34þ and CD34 cells, respectively, gated in
the graph at far left.