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Higher chondrogenic potential of fibrous synovium and adipose synoviumderived cells compared with subcutaneous fatderived cellsDistinguishing properties of mesenchymal stem cells in humans.

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ARTHRITIS & RHEUMATISM
Vol. 54, No. 3, March 2006, pp 843–853
DOI 10.1002/art.21651
© 2006, American College of Rheumatology
Higher Chondrogenic Potential of Fibrous Synovium– and
Adipose Synovium–Derived Cells Compared With
Subcutaneous Fat–Derived Cells
Distinguishing Properties of Mesenchymal Stem Cells in Humans
Tomoyuki Mochizuki, Takeshi Muneta, Yusuke Sakaguchi, Akimoto Nimura, Akiko Yokoyama,
Hideyuki Koga, and Ichiro Sekiya
Results. Fibrous synovium– and adipose
synovium–derived cells were higher in STRO-1 and
CD106 and lower in CD10 compared with subcutaneous
fat–derived cells. Cells derived from fibrous and adipose
synovium had higher proliferative potential and colonyforming efficiency compared with subcutaneous fat–
derived cells, both in mixed-population and in singlecell–derived cultures. In chondrogenic assays, pellets
from fibrous synovium– and adipose synovium–derived
cells produced more cartilage matrix than did cell
pellets from subcutaneous fat. Osteogenic ability was
also higher in fibrous synovium– and adipose
synovium–derived cells, whereas adipogenic potential
was nearly indistinguishable among the 3 populations.
Differentiation potential of the cells was similar between
young and elderly donors.
Conclusion. Cells derived from the fibrous synovium and from the adipose synovium demonstrate
comparable chondrogenic potential. Adipose synovium–
derived cells are more similar to fibrous synovium–
derived cells than to subcutaneous fat–derived cells.
Objective. Mesenchymal stem cells from synovium have a greater proliferation and chondrogenic
potential than do those from bone marrow, periosteum,
fat, and muscle. This study was undertaken to compare
fibrous synovium and adipose synovium (components of
the synovium with subsynovium) to determine which is
a more suitable source for mesenchymal stem cells,
especially for cartilage regeneration, and to examine the
features of adipose synovium–derived cells, fibrous
synovium–derived cells, and subcutaneous fat–derived
cells to determine their similarities.
Methods. Human fibrous synovium, adipose synovium, and subcutaneous fat were harvested from 4 young
donors and 4 elderly donors. After digestion, the nucleated
cells were plated at a density considered proper to expand
at a maximum rate without colony-to-colony contact. The
surface epitopes, proliferative capacity, cloning efficiency,
and chondrogenic, osteogenic, and adipogenic differentiation potentials of the cells were compared.
Supported in part by the Japan Society for the Promotion of
Science (grant 16591478), the Japan Orthopaedics and Traumatology
Foundation, and the Japan Sports Medicine Foundation. Dr. Muneta’s
work was supported by the Japan Society for the Promotion of Science
(grant 16591477), the Japan Latest Osteoarthritis Society, and the
Center of Excellence Program for Frontier Research on Molecular
Destruction and Reconstruction of Tooth and Bone in Tokyo Medical
and Dental University. Dr. Sekiya’s work was supported by the
Nakatomi Foundation.
Tomoyuki Mochizuki, MD, PhD, Takeshi Muneta, MD, PhD,
Yusuke Sakaguchi, MD, PhD, Akimoto Nimura, MD, Akiko
Yokoyama, MD, Hideyuki Koga, MD, Ichiro Sekiya, MD, PhD: Tokyo
Medical and Dental University, Tokyo, Japan.
Address correspondence and reprint requests to Ichiro
Sekiya, MD, PhD, Section of Orthopedic Surgery, Graduate School,
Tokyo Medical and Dental University, 1-5-45 Yushima, Bunkyo-ku,
Tokyo 113-8519, Japan. E-mail: sekiya.orj@tmd.ac.jp.
Submitted for publication May 27, 2005; accepted in revised
form November 21, 2005.
For treatment of articular cartilage injury, one of
the promising procedures is the transplantation of autologous cultured chondrocytes (1). However, surgical
invasion of normal articular cartilage and limited ex vivo
expansion of the chondrocytes lead to difficulties in
repairing large defects. Mesenchymal stem cells (MSCs)
have been a fascinating source for use in regenerative
medicine because they can be harvested in a less invasive
manner. Moreover, MSCs are easily isolated and expanded, with multipotential capabilities, including chondrogenesis (2,3).
An MSC is defined as being derived from mesenchymal tissue and having the functional capacity for self843
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MOCHIZUKI ET AL
renewal, commonly identified by colony-forming unit fibroblast assay (4) and generation of a number of differentiated
progeny (5). Increasing evidence suggests that postnatal
stem cells are not exclusive to bone marrow, but also are
present in various other tissues. We previously compared
MSCs derived from bone marrow, synovium, periosteum,
adipose tissue, and muscle, demonstrating that synovium
was a better cell source for MSCs with regard to cartilage
regeneration, in that synovium-derived MSCs had a greater
proliferative capacity and chondrogenic potential (6).
Synovium is a thin layer of tissue that lines the
joint space and covers a subsynovium. Depending on its
anatomic position, subsynovium comprises either a fibrous or an adipose connective tissue. There have been
other reports describing human synovium–derived
MSCs; however, details regarding the harvest site of
synovium and the histologic characteristics were not
mentioned (7,8). Most likely, synovium with subsynovium was used in these studies, because separation of
only the synovium layer from the subsynovial tissue is
difficult. Our first aim in the present study was to
distinguish MSCs by their sources, that is, synovium with
fibrous subsynovium, referred to as fibrous synovium,
and synovium with adipose subsynovium, referred as
adipose synovium.
MSCs derived from adipose synovium, also commonly called the infrapatellar fat pad, have been reported
to have multidifferentiation potential. MSCs of the adipose
synovium have been regarded as similar to liposuctionderived cells (9), except that the infrapatellar fat pad is
covered with synovium. Our second aim in the present
study was to identify whether infrapatellar fat pad–derived
cells are more closely related to fibrous synovium–derived
cells than to subcutaneous fat–derived cells.
In this study, we collected fibrous synovium, adipose synovium, and subcutaneous fat and performed
patient-matched quantitative comparisons of the properties of the 3 MSC populations. The properties examined
were surface epitopes, proliferative capacity, cloning efficiency, and chondrogenic, osteogenic, and adipogenic differentiation potentials. The goal of this study was to
characterize the suitability of fibrous synovium–derived
MSCs as compared with adipose synovium– or subcutaneous fat–derived MSCs for cartilage regeneration, from the
standpoint of the properties of each MSC population and
the accessibility of the MSC sources.
PATIENTS AND METHODS
Harvest of fibrous synovium, adipose synovium, and
subcutaneous fat. Tissues were harvested from 8 human
donors during knee operations. These donors were 4 young
patients with anterior cruciate ligament injury (mean ⫾ SD age
17.2 ⫾ 0.7 years) and 4 elderly patients with osteoarthritis
(mean ⫾ SD age 70.5 ⫾ 9.2 years). Young patients underwent
arthroscopic anterior cruciate ligament reconstructions, and
elderly patients received total knee arthroplasties. Fibrous
synovium was harvested from the inner side of the lateral joint
capsule, which overlays the noncartilaginous areas of the
lateral condyles of the femur. Adipose synovium was harvested
from the inner side of the infrapatellar fat pad, which overlays
the patellar tendon. Subcutaneous fat tissue under skin incisions over the tibiae was harvested. The study was approved by
an institutional review board, and informed consent was obtained from all study subjects.
Histologic analysis. A portion of the tissues was fixed
overnight at 4°C in 10% formalin, embedded in paraffin, and
sectioned at 5 ␮m. Tissue sections were then stained with
hematoxylin and eosin to identify histologic features.
Isolation and culture of MSCs. The tissue was minced
to pieces with a surgical knife, washed thoroughly with phosphate buffered saline (PBS) to remove hematopoietic cells,
and digested in a collagenase solution (3 mg/ml collagenase D;
Roche Diagnostics, Mannheim, Germany) in ␣–minimum essential medium (Invitrogen, Carlsbad, CA) at 37°C. After 3
hours, digested cells were filtered through a 70-␮m nylon filter
(Becton Dickinson, Franklin Lakes, NJ). Nucleated cells from
the tissues were plated at 103, 104, or 105 cells/cm2, each in 6
dishes, and cultured for 14 days at passage 0. After each of 3
dishes was stained with 0.5% crystal violet, optimal initial cell
densities were determined; this was decided according to the
supposition that colony size was not affected by colony-tocolony contact and that greater numbers of colonies were
therefore obtained. Total cell yields were thus established, and
the cells were used in further experiments (6).
Flow cytometry. One million cells (at passage 3) were
suspended in 500 ␮l PBS containing 20 ␮g/ml of antibody.
After incubation for 30 minutes at 4°C, the cells were washed
with PBS and suspended in 1 ml PBS for the analysis.
Fluorescein isothiocyanate (FITC)– or phycoerythrin (PE)–
coupled antibodies against CD34, CD45, CD90, and CD147
and anti–nerve growth factor receptor (anti-NGFR) antibody
were from Becton Dickinson, CD31,CD44, CD54 (intracellular adhesion molecule 1 [ICAM-1]), CD106 (vascular cell
adhesion molecule 1 [VCAM-1]), and CD117 were from
eBioscience (San Diego, CA), CD105 and CD166 (activated
leukocyte cell adhesion molecule [ALCAM]) were from Ancell
(Bayport, MN), STRO-1 and vascular endothelial cell growth
factor receptor 2 (VEGFR-2) were from Genzyme-Techne
(Minneapolis, MN), and CD10 was from DakoCytomation
(Copenhagen, Denmark). For isotype control, FITC- or PEcoupled nonspecific mouse IgG (Becton Dickinson) was substituted for the primary antibody.
Cell fluorescence was evaluated by flow cytometry
using a FACSCalibur instrument (Becton Dickinson), and data
were analyzed using CellQuest software (Becton Dickinson).
All analyses were performed on cells from 2 young donors and
2 elderly donors. For STRO-1 staining, the cells were incubated for 30 minutes with an antibody against STRO-1 (mouse
IgM; Genzyme-Techne). The cells were then incubated with a
secondary antibody (fluorescein-conjugated goat anti-mouse
IgM; Vector, Burlingame, CA) for 30 minutes. For the isotype
control against STRO-1, anti-mouse IgM (eBioscience) was
substituted. Positive expression was defined as a level of
PROPERTIES OF HUMAN SYNOVIAL MESENCHYMAL STEM CELLS
fluorescence ⬎99% of that observed with the corresponding
isotype-matched control antibodies (6,10).
Colony-forming efficiency. The cells at passage 0 were
replated at 100 cells per 60-cm2 dish, incubated for 14 days, and
stained with 0.5% crystal violet in 4% paraformaldehyde for 5
minutes. The cells were washed twice with distilled water, and the
number of colonies per dish was determined. Colonies ⬍2 mm in
diameter and faintly stained colonies were ignored (11).
In vitro chondrogenesis. Two hundred thousand cells
were placed in a 15-ml polypropylene tube (Becton Dickinson)
and centrifuged at 450g for 10 minutes. The pellet was cultured at
37°C with 5% CO2 in 400 ␮l of chondrogenic medium that
contained 500 ng/ml bone morphogenetic protein 2 (Yamanouchi
Pharmaceutical, Tokyo, Japan) in high-glucose Dulbecco’s modified Eagle’s medium (DMEM; Sigma-Aldrich, St. Louis, MO)
supplemented with 10 ng/ml transforming growth factor ␤3
(R&D Systems, Minneapolis, MN), 100 nM dexamethasone
(Sigma-Aldrich), 50 ␮g/ml ascorbate-2-phosphate, 40 ␮g/ml proline, 100 ␮g/ml pyruvate, and 1:100 diluted ITS⫹ Premix (6.25
␮g/ml insulin, 6.25 ␮g/ml transferrin, 6.25 ng/ml selenious acid,
1.25 mg/ml bovine serum albumin, and 5.35 mg/ml linoleic acid;
BD Biosciences, Bedford, MA). The medium was replaced every
3–4 days for 21 days. For microscopy, the pellets were embedded
in paraffin, cut into 5-␮m sections, and stained with toluidine blue
(12,13).
Reverse transcription–polymerase chain reaction (RTPCR). Cartilage pellets were digested with 3 mg/ml collagenase
for 3 hours to collect cells, and total RNA was prepared using the
RNAqueous kit (Ambion, Austin, TX). RNA was converted to
complementary DNA (cDNA) and amplified with the Titan One
Tube RT-PCR System (Roche Diagnostics) according to the
manufacturer’s recommendations. RT was performed with a
30-minute incubation at 50°C, followed by a 2-minute incubation
at 94°C to inactivate the RT. PCR amplification of the resulting
cDNA was performed under the following conditions: 35 cycles at
94°C for 30 seconds, 58°C for 45 seconds, and 68°C for 45 seconds,
the latter of which was increased by 5 seconds every cycle after
10 cycles. Forty cycles of these steps were performed for SOX5
and SOX9.
The reaction products were resolved by electrophoresis on a 1.5% agarose gel and visualized with ethidium
bromide (14). PCR primers were as follows: for COL2A1,
5⬘-TTCAGCTATGGAGATGACAATC-3⬘ (forward) and 5⬘AGAGTCCTAGAGTGACTGAG-3⬘ (reverse) (472 bp); for
aggrecan, 5⬘-GCAGAGACGCATCTAGAAATT-3⬘ (forward) and 5⬘-GGTAATTGCAGGGAACATCAT-3⬘ (reverse)
(505 bp); for decorin, 5⬘-CCTTTGGTGAAGTTGGAACG-3⬘
(forward) and 5⬘-AAGATGTAATTCCGTAAGGG-3⬘ (reverse) (300 bp); for biglycan, 5⬘-TGCAGAACAACGACATCTCC-3⬘ (forward) and 5⬘-AGCTTGGAGTAGCGAAGCAG-3⬘
(reverse) (475 bp); for link protein, 5⬘-CCTATGATGAAGCGGTGC-3⬘ (forward) and 5⬘-TTGTGCTTGTGGAACCTG-3⬘ (reverse) (618 bp); for SOX5, 5⬘-AGCCAGAGTTAGCACAATAGG-3⬘ (forward) and 5⬘-CATGATTGCCTTGTATTC-3⬘ (reverse) (619 bp); for SOX6, 5⬘-ACTGTGGCTGAAGCACGAGTC-3⬘ (forward) and 5⬘-TCCGCCATCTGTCTTCATACC-3⬘ (reverse) (562 bp); for SOX9,
5⬘-GAACGCACATCAAGACGGAG-3⬘ (forward) and 5⬘TCTCGTTGATTTCGCTGCTC-3⬘ (reverse) (631 bp); for
chondroitin 4-sulfotransferase (C4ST), 5⬘-CATCTACTGCTACGTGCCCA-3⬘ (forward) and 5⬘-CTTCAGGTAGCTGCCCACTC-3⬘ (reverse) (547 bp); for C6ST, 5⬘-GACTTT-
845
GTGCACAGCCTGAA-3⬘ (forward) and 5⬘-CCCTGCTGGTTGAAGAACTC-3⬘ (reverse) (431 bp); and for ␤-actin,
5⬘-CCAAGGCCAACCGCGAGAAGATGAC-3⬘ (forward)
and 5⬘-AGGGTACATGGTGGTGCCGCCAGAC-3⬘ (reverse) (587 bp).
Analysis of glycosaminoglycans (GAGs). The GAG content was quantified according to a previously described method
(15). Cartilage pellets were digested with 3 mg/ml collagenase in
0.25 ml of DMEM for 3 hours at 37°C, and a fraction of the cells
was removed. The supernatant was digested with chondroitinase
ABC (Chase ABC; Seikagaku, Tokyo, Japan) and hyaluronidase
derived from Streptococcus dysgalactonase (HAase SD; Seikagaku) for 2 hours at 37°C. After ultrafiltration, the filtrate was
analyzed by high-performance liquid chromatography. The levels
of chondroitin 4-sulfate (C4S), chondroitin 6-sulfate (C6S), and
hyaluronic acid were evaluated (14).
Osteogenesis in a colony-forming assay. One hundred
cells were plated in 60-cm2 dishes and cultured in complete
medium for 14 days. The medium was switched to calcification
medium that consisted of complete medium supplemented with 1
nM dexamethasone (Sigma-Aldrich), 20 mM glycerol phosphate
(Wako, Osaka, Japan), and 50 ␮g/ml ascorbate-2-phosphate for
an additional 21 days. Dishes were subsequently stained with
fresh 0.5% alizarin red solution, and the number of alizarin
red–positive colonies was determined. Colonies ⬍2 mm in diameter and faintly stained colonies were ignored. The same calcification cultures were then stained with crystal violet, and the total
number of cell colonies was determined (6,10).
Adipogenesis in a colony-forming assay. One hundred
cells were plated in 60-cm2 dishes and cultured in complete
medium for 14 days. The medium was then switched to
adipogenic medium that consisted of complete medium supplemented with 10 nM dexamethasone, 0.5 mM isobutylmethylxanthine (Sigma-Aldrich), and 50 ␮M indomethacin (Wako)
for an additional 21 days. The adipogenic cultures were fixed in
4% paraformaldehyde, stained with fresh 0.5% oil red O
solution, and the number of oil red O–positive colonies was
determined. Colonies ⬍2 mm in diameter and faintly stained
colonies were ignored. The same adipogenic cultures were
subsequently stained with crystal violet, and the total number
of cell colonies was determined (16).
Statistical analysis. Analysis of variance was used for
assessing differences between cell populations. P values less
than 0.05 were considered significant.
RESULTS
Macroscopic and histologic features of fibrous
synovium, adipose synovium, and subcutaneous fat. On
macroscopic analysis, fibrous synovium appeared relatively white, whereas both adipose synovium and subcutaneous fat were yellowish white (Figure 1A). Interestingly, fibrous synovium sunk, adipose synovium floated
partially, and subcutaneous fat floated completely in PBS,
indicating a difference in specific gravities (Figure 1B).
Histologically, fibrous synovium was composed of mostly
fibrous tissues, whereas adipose synovium consisted of
both fibrous tissues and subsynovial fatty tissue. It appeared that samples from the elderly patients had more
fibrous tissue than did those from the young donors, both
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MOCHIZUKI ET AL
Figure 1. Comparison of fibrous synovium–, adipose synovium–, and subcutaneous fat–derived cells. A and B, Macroscopic features of cells from an
elderly donor were examined on a 1-mm scale (A) and in phosphate buffered saline (B). C, Histologic analysis was performed on cells (stained with
hematoxylin and eosin) from both young and elderly donors. D, To determine the nucleated cell number per tissue weight, materials were digested with
collagenase, nucleated cells were counted, and the mean (and SEM) number of nucleated cells per tissue weight was calculated (n ⫽ 4). E, Representative
cell colonies from an elderly donor are shown. Nucleated cells were plated at 103, 104, and 105 cells/cm2 in 60-cm2 dishes, cultured for 14 days, and stained
with crystal violet. Representative morphologic features of the cells shown in E are magnified (⫻200) in F.
in fibrous synovium and in adipose synovium (Figure 1C).
Few, if any, erythrocytes were observed.
Nucleated cell number per tissue weight. The mean
nucleated cell number per tissue weight is shown in Figure
1D. The greatest number of nucleated cells was found in
fibrous synovium, an intermediate amount was found in
adipose synovium, and the fewest number of nucleated
cells was found in subcutaneous fat. The nucleated cell
PROPERTIES OF HUMAN SYNOVIAL MESENCHYMAL STEM CELLS
Figure 2. Flow cytometric analysis of fibrous synovium–derived
cells (solid bars), adipose synovium–derived cells (shaded bars),
and subcutaneous fat–derived cells (open bars). All analyses were
performed on cells from 2 young donors and 2 elderly donors.
Values are the mean and SD percentage expression of each
cell-surface protein. Data were analyzed using one-way analysis of
variance to assess the effect of cell sources. ⴱ ⫽ P ⬍ 0.05.
VEGFR2 ⫽ vascular endothelial growth factor receptor 2;
NGFR ⫽ nerve growth factor receptor.
847
number per tissue weight in the fibrous synovium of elderly
donors was substantially larger than that in the fibrous
synovium of young donors.
Colony formation and morphologic features of
the cells. In order to gain maximum yields per nucleated
cells, we examined the effect of plating density on nucleated cells. Larger single-cell–derived colonies were observed when the nucleated cells were plated at 103 cells/cm2
in all populations (Figure 1E). When plated at 104 or 105
cells/cm2, colony size decreased or became indistinct, possibly due to colony-to-colony contact inhibition. These
observations indicated an optimal initial cell density at 103
cells/cm2 to maximize cell yields per dish. Thus, we were
able to identify the optimal initial cell density for nucleated
cells, and this was used in further experiments.
On morphologic analysis, fibrous synovium–
Figure 3. A, Proliferation potential of fibrous synovium–, adipose synovium–, and subcutaneous fat–derived cells from 1 young donor. Cells at
passage 1 were plated at 10, 50, and 100 cells/cm2 and the fold increase (n ⫽ 3) was calculated after 7 and 14 days. Data were analyzed using
one-factor analysis of variance (ANOVA) to assess the effect of plating density on fold increase. B, The cells at passage 1 from 4 young and 4 elderly
donors were plated in triplicate at 50 cells/cm2 and the fold increase was calculated after 14 days. Data were analyzed using two-factor ANOVA to
assess the effect of cell sources and donor age. C, One hundred cells at passage 1 from 4 young and 4 elderly donors were plated in triplicate in 60-cm2
dishes and the colony-forming efficiency was calculated after 14 days. D, For analyses of single-cell–derived cultures, nucleated cells from 1 elderly
donor and 1 young donor were plated at 103 cells/cm2 and cultured for 14 days, and 3 single-cell–derived colonies were collected. Three clones from
fibrous synovium–, adipose synovium–, and subcutaneous fat–derived cells were replated at 50 cells/cm2, each in triplicate, and the fold increase was
calculated after 14 days. E, One hundred cells from the clones were plated in triplicate in 60-cm2 dishes and the colony-forming efficiency was
calculated after 14 days. Bars show the mean and SD. ⴱ ⫽ P ⬍ 0.05; ⴱⴱ ⴝ P ⬍ 0.01; ⴱⴱⴱ ⫽ P ⬍ 0.005.
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MOCHIZUKI ET AL
Figure 4. A and B, Chondrogenic potential of fibrous synovium–, adipose synovium–, and subcutaneous fat–derived cells. The cells were pelleted
and cultured in chondrogenic medium for 21 days. Representative macroscopic findings from a young donor are shown on a 1-mm scale (A), and
the histologic features were examined by staining with toluidine blue (B). C, The wet weight of the pellets from 3 young and 3 elderly donors was
measured in triplicate; data are expressed as the mean and SD, analyzed using two-factor analysis of variance to assess the effect of cell sources and
donor age. D, To compare glycosaminoglycans in the extracellular matrix, 3 cell pellets from 1 young donor were digested to remove the cells, and
chondroitin 4-sulfate (C4S), chondroitin 6-sulfate (C6S), and hyaluronic acid (HA) in the supernatant were evaluated. E, To compare the mRNA
expression for chondrogenic genes, total RNA was prepared from 3 cell pellets from 1 young donor, and reverse transcription–polymerase chain
reaction was performed. C4ST ⫽ chondroitin 4-sulfotransferase.
derived cells and adipose synovium–derived cells appeared
similar, in that they were small and spindle-shaped. In
contrast, subcutaneous fat–derived cells were larger and
flatter, especially when cultured at higher densities (Figure
1F).
Epitope profile. Of the 15 antibodies examined,
the rate of positivity for CD45 (hematopoietic cell
marker), CD31 (endothelial cell marker), CD117 (C-kit,
stem cell factor receptor), and CD34 (hematopoietic
progenitor cell antigen) (3,6,10,17–20) was ⬍2%, and
PROPERTIES OF HUMAN SYNOVIAL MESENCHYMAL STEM CELLS
that for VEGFR-2 (Flk-1) (6,10,18) and for NGFR
(6,10,18) was ⬍4% in fibrous synovium–, adipose
synovium–, and subcutaneous fat–derived cells (Figure
2). However, rates of the STRO-1 (6,10,18,21) and
CD106 (VCAM-1) positivity (3,6,10,22) in fibrous
synovium– and adipose synovium–derived cells were
⬍6%, which was significantly higher than that in subcutaneous fat–derived cells. In contrast, the rate of CD10
positivity (17,18) in subcutaneous fat–derived cells was
40%, which was higher than that in fibrous synovium–
and adipose synovium–derived cells, both of which had
⬃10% positivity for CD10. The rates of positivity for
CD54 (ICAM-1) (3,6,10,23) and CD166 (ALCAM,
SB10) (23) were 20–40%, that for CD90 (Thy1) (17) was
40–70%, that for CD105 (SH-2) (3,24) was 60–80%, that
for CD44 (hyaluronan receptor) (3) was 70–90%, and
that for CD147 (neuroregulin) (18) was ⬎90% in all 3
MSC populations.
Proliferative potential. Initial cell density affected increases in proliferation of fibrous synovium–,
adipose synovium–, and subcutaneous fat–derived cells
(Figure 3A). In young donors, fibrous synovium–derived
cells had higher proliferative potential than did adipose
synovium–derived cells; however, the proliferative potential of the 2 populations was similar in elderly donors
(Figure 3B). The proliferative potential was lowest in
subcutaneous fat–derived cells, in young donors and in
elderly donors. The colony-forming efficiency of subcutaneous fat–derived cells was significantly lower than
that of fibrous synovium–derived cells, in young donors
and in elderly donors (Figure 3C).
Single-cell–derived cultures. To compare the
cells and better define each population, clones of fibrous
synovium–, adipose synovium–, and subcutaneous fat–
derived cells were prepared. Both the magnitude of
increase and the colony-forming efficiency of the fibrous
synovium– and adipose synovium–derived cells were
higher than those of the subcutaneous fat–derived cells
in both young and elderly donors (Figures 3D and E).
The results from single-cell–derived cultures generally
appeared similar to those obtained in a mixed population of the cells.
Chondrogenesis. To compare the chondrogenic
potential of the MSC populations, cells were differentiated into cartilage in vitro. After 21 days of culture, the
cell pellets became spherical (Figure 4A). During in
vitro chondrogenesis, the pellets increased in size, which
was attributable to production of extracellular matrix
(12). Pellets from fibrous synovium– and adipose
synovium–derived cells had greater amounts of cartilage
matrix than did pellets from subcutaneous fat, as shown
by staining with toluidine blue (Figure 4B). Pellets from
849
fibrous synovium– and adipose synovium–derived cells
were also heavier than those from subcutaneous fat–
derived cells (Figure 4C).
Determination of GAGs in the extracellular matrix of the pellets showed that the amount of C4S, C6S,
and hyaluronic acid differed among the cell pellets. Cells
derived from fibrous synovium and adipose synovium
exhibited greater amounts of GAGs than did those from
subcutaneous fat (Figure 4D).
RT-PCR demonstrated that expression of messenger RNA (mRNA) for the chondrogenic genes
COL2A1, link protein, SOX6, SOX9, and C4ST in cell
pellets derived from fibrous synovium and adipose synovium was higher than that in cells from subcutaneous fat
(Figure 4E). Expression of mRNA for the other chondrogenic genes, aggrecan, decorin, biglycan, SOX5, and
C6ST, appeared similar among the cell pellets from the
3 populations. These results indicate that fibrous
synovium– and adipose synovium–derived cells had
higher chondrogenic potential than did subcutaneous
fat–derived cells.
Osteogenesis. To evaluate the osteogenic potential of the MSC populations, cells were cultured in
osteogenic medium. All cells were calcified and stained
with alizarin red (Figure 5A). The ratios of alizarin
red–positive (6,10) colonies in fibrous synovium– and
adipose synovium–derived cells were higher than that in
subcutaneous fat–derived cells (Figure 5B), indicating
that there was a difference in osteogenic potential
among these cells. Results in the 3 populations were
similar between cells from young donors and those from
elderly donors.
Adipogenesis. To compare the adipogenic potential of the cells in the 3 populations, the ratio of oil red
O–positive colonies to total colonies was evaluated
(6,10,11,16). The oil red O–positive colony rate was
similar among the 3 populations (Figures 6A and B).
There were no differences between cells from young
donors and those from elderly donors.
DISCUSSION
In this study, we compared fibrous synovium–,
adipose synovium–, and subcutaneous fat–derived cells
from the perspective of common properties of MSCs. In
the 3 populations, epitope profiles of the cells were
similar, in that the rate of positivity for CD34 and CD45
(hematopoietic cell markers) was low and the rate of
positivity for CD44 (hyaluronan receptor) and CD105
(SH-2) was high. These results coincide with the phenotypic properties of bone marrow–derived MSCs
(3,6,10,18,25).
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MOCHIZUKI ET AL
marrow (26) and has shown some promise for use in
immunophenotyping MSCs (27). The rate of CD106
(VCAM-1) positivity in fibrous synovium– and adipose
synovium–derived cells was 5%, which was higher than
that in subcutaneous fat–derived cells (1%). VCAM-1 is
Figure 5. Osteogenic potential of fibrous synovium–, adipose
synovium–, and subcutaneous fat–derived cells. The cells were plated
at 100 cells per 60-cm2 dish and cultured for 14 days to make cell
colonies. The cells were then incubated in osteogenic medium for an
additional 21 days. A, Calcified colonies stained with alizarin red are
shown as red colonies. The same dishes were then stained with crystal
violet to count total colony number. B, Analyses were performed in
triplicate on cells from 3 young and 3 elderly donors, and the ratios of
alizarin red–positive colonies to total colonies were calculated. Data
were analyzed using two-factor analysis of variance to assess the effect
of cell source and donor age; bars show the mean and SD. ⴱ ⫽ P ⬍
0.05; ⴱⴱⴱ ⫽ P ⬍ 0.005.
Interestingly, the rate of STRO-1 positivity in
fibrous synovium– and adipose synovium–derived cells
was higher than that in subcutaneous fat–derived cells.
STRO-1 was originally reported to identify colonyforming osteogenic precursor cells isolated from bone
Figure 6. Adipogenic potential of fibrous synovium–, adipose
synovium–, and subcutaneous fat–derived cells. The cells were plated
at 100 cells per 60-cm2 dish and cultured for 14 days to make cell
colonies. The cells were then incubated in adipogenic medium for an
additional 21 days. A, Adipocyte colonies stained with oil red O are
shown as red colonies. The same dishes were then stained with crystal
violet to count total colony number. B, Analyses were performed in
triplicate on cells from 3 young and 3 elderly donors, and the ratios of
oil red O–positive colonies to the total colonies were calculated. Data
were analyzed using two-factor analysis of variance to assess the effect
of cell source and donor age; bars show the mean and SD.
PROPERTIES OF HUMAN SYNOVIAL MESENCHYMAL STEM CELLS
a cell-surface glycoprotein that is produced by cytokineactivated endothelium, and is expressed primarily on
lining layer cells in synovial tissue (28).
In contrast, the rate of CD10 positivity in subcutaneous fat–derived cells was 40%, which was higher
than that in fibrous synovium– and adipose synovium–
derived cells (10%). CD10 is also known as common
acute lymphocytic leukemia antigen or human
membrane–associated neutral endopeptidase. Colter et
al previously demonstrated that single-cell–derived colonies of bone marrow–derived MSCs contained 3 morphologically distinct cell types: large flat cells, small
spindle-shaped cells, and extremely small, rapidly dividing cells, and showed that samples enriched for the small
and extremely small cells had a greater ability for
multipotential differentiation than did samples enriched
for the large cells. Those authors found that CD10 was a
negative marker for small and extremely small cells (18).
In this study, fibrous synovium– and adipose synovium–
derived cells expressed lower levels of CD10, showed
increased proliferation, and had higher chondrogenic
and osteogenic potential than did subcutaneous fat–
derived cells, which seems to support the findings in
bone marrow–derived MSCs reported by Colter et al.
Several reports have described MSCs derived
from human adipose tissue. To harvest the cells, great
amounts of liposuction tissue were collected, digested
with collagenase, separated by stromal–vascular fraction,
and expanded (29–31). The processed lipoaspiratederived cells contaminate endothelial, smooth muscle,
and pericyte cell populations (31). In our study, subcutaneous fat–derived cells lacked robust chondrogenic
activity. We collected only an ⬃100-mg fat tissue, and
after digestion, the cells were plated without gradient
separation. The quantity of tissue and procedure for
fractionation may explain the difference in properties
observed in the adipose-derived cells in this study.
MSCs derived from the infrapatellar fat pad
(adipose synovium) have also been described (9,32).
Dragoo et al (9) regarded infrapatellar fat pad–derived
cells as adipose-derived cells; however, the infrapatellar
fat pad is composed of synovium and subsynovial adipose tissues. In our study, the results from morphologic
study of infrapatellar fat pad tissue, morphologic study
of expanded cells, examination of surface epitopes, and
studies of the proliferation, colony-forming efficiency,
and chondrogenetic and osteogenetic potential indicate
that adipose synovium–derived cells are more similar to
fibrous synovium–derived cells than to subcutaneous
fat–derived cells.
We evaluated the properties of MSCs in the 3
populations in young and elderly donors separately,
851
because we had initial concerns that synovium-derived
MSCs from elderly donors might lack the ability for
expansion and differentiation. There are several reports
describing the influence of aging on the properties of
bone marrow–derived MSCs, and this topic remains
controversial. Some studies have shown that aging does
not affect colony-forming efficiency (33–35), adipogenesis (34), and calcification (33,34,36). In contrast, others
have reported that aging affects the proliferative capacity at passage 1 (36) as well as the chondrogenic (36),
osteogenic (35,37), and adipogenic (36) differentiation
ability of bone marrow–derived MSCs.
Our previous study indicated no obvious differences between bone marrow–derived MSCs from young
and elderly donors in terms of the yields of cells at
passage 0, the colony-forming efficiency at passage 0,
surface-cell antigens, and chondrocyte, adipocyte, and
osteoblast differentiation potentials. However, the proliferative ability of passage 1 cells decreased with age,
with results observed as a decrease in cell numbers per
colony (10). Apparently, this discrepancy can be attributable to the differences in donor status, site, differentiation protocol, and evaluation method. For example, D’Ippolito et al demonstrated that the number of
MSCs with osteogenic potential decreased during
aging. They collected bone marrow from vertebral
bodies, switched to osteogenic medium 1 day after
plating, and evaluated osteogenic potential as the ratios
of alkaline phosphatase–positive colonies (35). We collected bone marrow from the proximal tibiae, switched
osteogenic medium 14 days after plating, and evaluated
osteogenic potential by the ratios of alizarin red–positive
colonies (10).
Oreffo et al demonstrated a significant decrease
in the ratio of alkaline phosphatase–positive colonies in
elderly donors with osteoporosis as compared with
young donors, whereas there was no difference in this
ratio between elderly donors with osteoarthritis and
young donors (37). Murphy et al demonstrated a significant reduction in chondrogenic and adipogenic activity
of bone marrow–derived MSCs from elderly osteoarthritis patients compared with those from younger donors
(36). Their chondrogenic medium did not contain bone
morphogenetic proteins, whereas ours included bone
morphogenetic protein 2, which enhanced the in vitro
chondrogenesis of MSCs (13). Furthermore, they evaluated chondrogenic potential by the amount of GAG
standardized to DNA content, while we compared cartilage pellet weight. With regard to adipogenesis, Murphy et al plated MSCs at high density, differentiated the
MSCs into adipocytes, and quantitated nile red fluorescence standardized to 4⬘,6-diamidino-2-phenylindole,
852
whereas we plated at low density, formed cell colonies,
differentiated cells into adipocytes, and evaluated oil red
O–positive colony rates.
This study showed no remarkable differences
between young and elderly donors in terms of the
proliferative ability and colony-forming efficiency of the
cells at passage 1, or the chondrocyte, osteoblast, and
adipocyte differentiation potential in each MSC population derived from fibrous synovium, adipose synovium,
and subcutaneous fat. Contrary to our initial predictions,
the nucleated cell number per tissue weight in the
fibrous synovium of elderly donors was larger than that
in young donors. Revell et al reported that synovium
from elderly patients with osteoarthritis is likely to be
fibrous (38), and our results were consistent with this
observation (as shown in Figure 1C).
Osteoarthritis comprises a common, age-related
heterogeneous group of disorders that are characterized
pathologically by focal areas of loss of articular cartilage
in synovial joints, associated with varying degrees of
osteophyte formation, subchondral bone changes, and
synovitis (39). The secondary inflammation in the synovium could alter its cellular composition and could be
responsible for changes to the stem cell populations.
Detailed pathologic investigation of synovium will be
important for clarifying the role of the disease and the
role of aging in the characteristics of stem cells derived
from the synovium. This general information will be
valuable for comparison with other studies.
An age-related decrease in the chondrogenic
differentiation potential has been reported in rabbit
fibrous synovium (40) and periosteum (41) in an ex vivo
organ culture (42). In contrast, De Bari et al demonstrated that the chondrogenic potential of synoviumderived cells was independent of donor age (7), which is
similar to our result. These findings suggest that aging
may affect the chondrogenic differentiation potential in
organ culture but does not affect the cells expanded
in vitro.
We demonstrated that both fibrous synovium–
and adipose synovium–derived MSCs had a better chondrogenic capacity than did subcutaneous fat–derived
MSCs. Given this difference, important biologic questions are raised, and we have developed 2 hypotheses: 1)
The observed differences between synovium and subcutaneous fat may be due to differences in the number of
ancestral MSCs, or alternatively, 2) the observed differences in chondrogenesis could have arisen as a result of
different MSC propensities to follow a chondrogenic
pathway, suggesting that the local tissue microenvironment may be directing the “fate” of the MSCs toward a
particular lineage.
MOCHIZUKI ET AL
We also demonstrated that synovium-derived
MSCs had a higher osteogenetic ability than did adiposederived MSCs. Furthermore, single-cell–derived cultures
as well as mixed synovial cells showed higher colonyforming efficiency and expansion ability than did those
from adipose tissue. These findings may implicate the role
of the local tissue microenvironment in directing the
“fate” of the MSCs. The adipogenic capacity of MSCs
derived from the synovium and those derived from the
subcutaneous fat was similar, which would support the
first hypothesis described above. However, MSCs derived from adipose tissue could have been preconditioned via the microenvironment, and thus slightly predisposed toward an adipocyte lineage, which would be
consistent with the second hypothesis.
The important consideration in tissue engineering is to harvest the greatest amount of MSCs with the
highest potential while minimizing the amounts of mesenchymal tissues needed, resulting in less-invasive treatments. Fibrous synovium– and adipose synovium–
derived MSCs were similar in terms of their cell
morphologic features, epitope profiles, colony-forming
efficiency, chondrogenesis, osteogenesis, and adipogenesis potentials. The nucleated cell number per tissue
weight was higher in fibrous synovium than in adipose
synovium, which may be an advantage of fibrous synovium. However, adipose synovium cells also have an
advantage due to their high chondrogenic potential and
accessibility, in that sufficient amounts of adipose synovium can be harvested with possibly fewer complications.
We therefore conclude that both fibrous synovium and
adipose synovium are suitable MSC sources for cartilage
regeneration.
ACKNOWLEDGMENTS
We thank Kenichi Shinomiya, MD, PhD, for continuous support, Kazuyoshi Yagishita, MD, PhD, for sample
collection, Izumi Nakagawa for excellent technical assistance,
Miyoko Ojima for expert help with histology, Kyosuke
Miyazaki for analysis of chondroitin sulfate, and Benjamin L.
Larson for proofreading.
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stem, fibrous, fatderived, synoviumderived, properties, synovium, cellsdistinguishing, adipose, human, cells, potential, mesenchymal, compare, higher, subcutaneous, chondrogenic
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