Specific markers and properties of synovial mesenchymal stem cells in the surface, stromal, and perivascular regions
Mizuno et al. Stem Cell Research & Therapy
Specific markers and properties of synovial mesenchymal stem cells in the surface, stromal, and perivascular regions
Mitsuru Mizuno 0
Hisako Katano 0
Yo Mabuchi 2
Yusuke Ogata 2
Shizuko Ichinose 0
Shizuka Fujii 0
Koji Otabe 0
Keiichiro Komori 0
Nobutake Ozeki 0
Hideyuki Koga 1
Kunikazu Tsuji 4
Chihiro Akazawa 2
Takeshi Muneta 3
Ichiro Sekiya 0
0 Center for Stem Cell and Regenerative Medicine, Tokyo Medical and Dental University , 1-5-45 Yushima, Bunkyo-ku, Tokyo 113-8510 , Japan
1 Department of Joint Surgery and Sports medicine, Tokyo Medical and Dental University , Tokyo , Japan
2 Department of Biochemistry and Biophysics, Tokyo Medical and Dental University , Tokyo , Japan
3 National Hospital Organization, Disaster Medical Center , Tokyo , Japan
4 Department of Cartilage Regeneration, Tokyo Medical and Dental University , Tokyo , Japan
Background: Synovial mesenchymal stem cells (MSCs) are an attractive cell source for cartilage and meniscus regeneration. Synovial tissue can be histologically classified into three regions; surface, stromal and perivascular region, but the localization of synovial MSCs has not been fully investigated. We identified markers specific for each region, and compared properties of MSCs derived from each region in the synovium. Methods: The intensity of immunostaining with 19 antibodies was examined for surface, stromal, and perivascular regions of human synovium from six osteoarthritis patients. Specific markers were identified and synovial cells derived from each region were sorted. Proliferation, surface marker expression, chondrogenesis, calcification and adipogenesis potentials were compared in synovial MSCs derived from the three regions. Results: We selected CD55+ CD271− for synovial cells in the surface region, CD55− CD271− in the stromal region, and CD55− CD271+ in the perivascular region. The ratio of the sorted cells to non-hematopoietic lineage cells was 5% in the surface region, 70% in the stromal region and 15% in the perivascular region. Synovial cells in the perivascular fraction had the greatest proliferation potential. After expansion, surface marker expression profiles and adipogenesis potentials were similar but chondrogenic and calcification potentials were higher in synovial MSCs derived from the perivascular region than in those derived from the surface and stromal regions. Conclusions: We identified specific markers to isolate synovial cells from the surface, stromal, and perivascular regions of the synovium. Synovial MSCs in the perivascular region had the highest proliferative and chondrogenic potentials among the three regions.
Mesenchymal stem cells (MSCs) are an attractive cell
source for cell therapies. These cells participate in tissue
homoeostasis, remodeling, and repair by ensuring
replacement of mature cells that are lost during the course
of physiological turnover, senescence, injury, or disease
]. Along with preclinical studies, a large number of
clinical trials have been conducted for cardiovascular
diseases, osteoarthritis, liver disorders, graft versus host
disease (GvHD), respiratory disorders, spinal cord injury,
and others [
]. MSCs are found not only in bone
marrow but multiple adult tissues [
MSCs are defined as non-hematopoietic-lineage,
plastic-adherent, self-renewing cells that can differentiate
into chondrocytes, adipocytes and osteoblasts in vitro [
]. Traditionally, the isolation of MSCs has relied on
their adherence to plastic dishes and colony-forming
ability in an unfractionated cell population. This
technique may give rise to heterogeneous cell populations in
MSCs. To better characterize this heterogeneity, surface
markers have been investigated for bone marrow MSCs
from the osteoblast region , endosteum region [
and perivascular region [
Synovial MSCs have a higher chondrogenic potential
than bone marrow MSCs [
]. Transplantation of
synovial MSCs regenerated cartilage [
] and meniscus [
Synovial MSCs are clinically used for cartilage
]. To prepare synovial MSCs, synovium is
digested, and unfractionated synovial cells are expanded
to form cell colonies of synovial MSCs [
tissue can be histologically classified into three regions;
surface, stromal, and perivascular regions . If
synovial cells can be obtained and synovial MSCs can be
prepared from each region separately, more attractive
synovial MSCs can be used in clinical therapies. This
also provides important information on the physiological
roles of cells in the synovium. The purpose of the
present study was to identify specific markers for the
isolation of synovial cells in the surface, stromal, and
perivascular regions, and to compare properties of MSCs
sorted by the specific markers.
This study was approved by the Medical Research Ethics
Committee of Tokyo Medical and Dental University and
all human study subjects provided informed consent.
Human synovium was harvested from the knees of ten
donors (59–85 years) with osteoarthritis during total
Transmission electron microscopy (TEM)
The specimens of synovial tissues were rapidly fixed in
2.5% glutaraldehyde in 0.1 M phosphate buffer for 2 h.
The samples were washed with 0.1 M phosphate buffer,
post-fixed in 1% OsO4 buffered with 0.1 M phosphate
buffer for 2 h, dehydrated in a graded series of ethanol
and embedded in Epon 812. Ultrathin sections at 90 nm
were collected on copper grids, double-stained with
uranyl acetate and lead citrate, and then examined by
transmission electron microscopy (H-7100, Hitachi,
Tokyo, Japan) [
Synovial tissues were rapidly embedded in OCT
compound (Sakura Finetec Japan, Tokyo, Japan) and 4%
carboxymethyl cellulose and were washed with 0.1%
Tween-TBS. After blocking with Protein Block
SerumFree (Dako, Glostrup, Denmark), sections (5 μm thick)
were incubated with 19 antibodies; CD90 (Becton,
Dickinson and Company; BD, Franklin Lakes, NJ, USA),
CD44 (BD), CD73 (BD), CD105 (BD), CD271 (Miltenyi
Biotec, Bergisch Gladbach, Germany), CD140a (BD),
CD140b (BD), CD29 (Merck Millipore, Darmstadt,
Germany), CD49f (Merck Millipore), Ki67 (Dako),
Proliferating Cell Nuclear Antigen (PCNA; Santa Cruz
Biotechnology, Inc., Santa Cruz, CA, USA), CD55
(Miltenyi Biotec), CD31 (antibody derived from mouse
(Dako) for IHC and sheep (R&D Systems, Minneapolis,
MN, USA) for IF), CD146(BD), Laminin (Dako),
Collagen type IV (Dako), Proteoglycan 4/Lubricin (PRG4;
Santa Cruz Biotechnology), Hyaluronan synthase 1
(HAS-1; Santa Cruz Biotechnology) and HAS-2 (Santa
Cruz Biotechnology), at 4 °C overnight. After washing
three times, secondary antibodies (Chemmate Envision
HRP-polymer, Dako) or anti-goat horseradish peroxidase
(HRP)-conjugated secondary antibody (Dako) were
added, followed by incubation for 30 min at room
temparature. Staining was simultaneously developed in DAB
+ solution (Dako), with counterstaining by hematoxylin.
Samples were analyzed with a light microscope. The
DAB positive intensity of surface and matrix protein
expression in synovium was quantified with Image J
software after immunostaining [
For fluorescence microscopy images, sections were
first washed then incubated with Alexa Fluor 488- and/
or 594-conjugated secondary antibodies (1:500; Thermo
Fisher Scientific, Waltham, MA, USA) specific for the
appropriate species for 1 h at room temperature.
Samples were counterstained with
4,6-diamidino-2-phenylindole (DAPI; Wako, Osaka, Japan) and analyzed with a
microscope (BZ-X700, Keyence Co., Ltd., Osaka, Japan).
Flow cytometric isolation and analysis
Synovium was digested in a solution of 3 mg/mL collagenase
(Sigma-Aldrich Japan, Tokyo, Japan) at 37 °C. After 3 h, the
digested cells were filtered through a 70-μm cell strainer
(Greiner Bio-One GmbH, Kremsmunster, Austria). The cells
from six donors were harvested using a cell-dissociation
buffer. Cells were suspended in HBSS at a density of 5 × 105
cells/mL and stained for 30 min on ice with the antibodies.
For cell isolation, cells were stained with CD31-PE-Cy7 (BD),
CD45-PE-Cy7 (BD), CD235a-PE-Cy7 (BD), CD55-FITC
(Miltenyi Biotec), CD90-PE (BD) and CD271-APC (Miltenyi
Biotec) were used at day 0. Flow cytometric isolation of cell
surface antigens were performed by a double-laser Aria 2
system (BD). For cell surface analysis, cells were stained with
CD31-FITC (BD), CD45-FITC (BD), CD44-APC-H7 (BD),
CD73-BV421 (BD), CD90-PE (BD), CD105-PerCP-Cy5.5
(BD), CD55-PE (BD), CD271-APC (Miltenyi Biotec),
CD140b-PerCP-Cy5.5 (BD) and CD146-FITC (BD) at
passage 3. Flow cytometric analysis of cell surface antigens was
performed by a triple-laser FACS Verse system (BD). These
data were analyzed using FlowJo software (Tree Star Inc.,
Ashland, OR, USA). Flow cytometric analyses were also
performed for expanded cells at passage 3.
Colony formation and proliferation ability of synovial
For proliferation assays, bulk and sorted synovial cells
from four donors were plated on six wells at 2000 cells
per 10 cm2 wells for 10 days in complete culture medium
with 10% FBS (Thermo Fisher Scientific, Inc.) and 1%
penicillin/streptomycin in alpha MEM (Thermo Fisher
Scientific, Inc.). Cultured cells were harvested with 0.25%
trypsin and 1 mM ethylenediaminetetraacetic acid (EDTA)
(Thermo Fisher Scientific, Inc) at 37 °C for 5 min and
counted with cell-counting plates. For colony formation
assays, bulk and sorted synovial cells from four donors were
plated as above. The dishes were stained with crystal violet
at 14 days and the colony number was counted.
Differentiation assay of synovial MSCs
For chondrogenic differentiation, cultured synovial MSCs
from four donors at passage 2 were harvested using a
celldissociation buffer as time 0 or cells at 48 h were
harvested from preservation tubes after preservation. Then 2.
5 × 105 cells were transferred to a 15 mL tube (BD Falcon)
and cultured in chondrogenic induction medium
containing 10 ng/mL transforming growth factor-β3
(Miltenyi Biotec) and 1 μg/mL bone morphogenetic protein 2
(Medtronic, Minneapolis, MN, USA), in high glucose
DMEM with 1% antibiotic-antimycotic (Thermo Fisher
Scienific) which was changed every 3–4 days. After
21 days, chondrogenic differentiated cells were analyzed
by safranin-o (Wako) staining.
For calcification induction, 100 cells were transferred
to a 60 cm2 dish and cultured for 14 days in complete
culture medium. Adherent cells were then cultured in
osteogenic induction medium containing 50 μg/mL
ascorbic acid 2-phosphate (Wako), 10 nM
dexamethasone (Wako), and 10 mM β-glycerophosphate
(SigmaAldrich), in complete culture medium, which was
changed every 3–4 days. After 21 days, the differentiation of
these cells into osteoblasts was assessed by alizarin red
staining (Merck Millipore). To quantify the amount of
alizarin red, the deposition was extracted by 10% (w/v)
cetylpyridinium chloride (Sigma-Aldrich) in 10 mM
sodium phosphate (pH 7.0) at room temperature for 1 h
and the alizarin red stain in the extraction buffer was
determined by measuring the optical density of the
solution at 560 nm absorbance [
For adipogenic differentiation, adherent cells were
cultured in adipogenic induction medium (Lonza, Basel,
Switzerland), which was changed every 3–4 days. After
21 days, oil red-o staining (Muto Pure Chemicals,
Tokyo, Japan) confirmed the differentiation of these cells
into adipocytes. To quantify adipogenic ability, the
amount of triglyceride was measured by adipogenesis
assay kit (BioVision, Milpitas, CA, USA) in accordance
with manufacturer’s instructions.
RNA isolation and RT-PCR analysis
For chondrogenesis, six pellets from each donor were
digested together. For calcification and adipogenesis,
MSCs were plated at 50 cells/ cm2 in 145 cm2 plates
and extracted from two dishes. Total RNA was extracted
using RNeasy Mini Kit (Qiagen N.V., Venlo,
Netherlands). Concentration and quality of the RNA
were verified on a Quantus Fluorometer (Promega Co.,
Madison, WI, USA). The cDNA was synthesized with
random hexamer primers from total RNA using the
Transcriptor High Fidelity cDNA Synthesis kit (Roche
Diagnostics, Basel, Switzerland). Real-time PCR was
performed in a LightCycler 480 instrument (Roche
Diagnostics). PCR reaction used the LightCycler 480 Probes
Master. Relative amounts of mRNA were calculated and
standardized as previously described [
]. The following
TaqMan gene expression assay kits (Integrated DNA
Technologies, IA, USA) were used as Hs.PT.39a.
22214847 for ACTB, Hs.PT.58.38984663 for SOX9, Hs.
PT.58.38672730 for COL10A1, Hs.PT.56a.742783 for
ACAN, Hs.PT.56a.40555206 for ALP, Hs.PT.56a.
19568141 for RUNX2, Hs.PT.58.25464465 for PPARG,
Hs.PT.58.4022335.g for CEBPA, Hs.PT.58.3040231 for
GTF3A, and Hs.PT.58.20087469 for LPL.
All data were statistically evaluated with GraphPad
Prism 6 (GraphPad Software, La Jolla, CA, USA). Data
are expressed as mean ± SD. Each statistical analysis
method is described in the legend. Two-tailed P values
of < 0.05 were considered to be significant.
TEM images for surface, stromal, and perivascular regions of synovium
Synovium can be histologically classified into three
regions; surface, stroma, and perivascular regions (Fig. 1a).
These three regions had different ultra-microstructures.
The surface region of the synovial membrane primarily
consisted of macrophage and fibroblast cell components
(Fig. 1b). The stromal region, defined as subsynovial
tissue excluding the perivascular region, consisted of
stromal cells with collagen fibrils. The perivascular
region in synovial tissue contained blood vessels with
Immunological characterization for surface, stromal, and perivascular region of synovium
To characterize the three regions immunologically,
immunostaining with 19 antibodies was performed. We
were able to detect the following proteins in the surface,
stroma or perivascular region; MSC markers (CD90,
CD44 and CD73), growth factor receptors of MSC
markers (CD105, CD271, CD140a and CD140b),
integrins (CD29 and CD49f ), proliferative markers (Ki67,
PCNA), complement inhibitor (CD55), endothelial
markers (CD31 and CD146), and extracellular matrix
(Laminin, Col4, PRG4, HAS-1 and HAS-2) (Fig. 2).
The expression of proteins was region specific.
Generally, in the surface region, MSC markers, proliferation
markers, complement inhibitor, and extracellular
matrices were highly expressed. In the stromal region, only
MSC markers were highly expressed. In the perivascular
region, most proteins we examined, with the exception
of complement inhibitor, were highly expressed (Fig. 3a).
Specific markers for isolation of synovial cells from surface, stromal, and perivascular regions
As specific markers for isolation of synovial cells from
each region, we selected CD55 as a positive marker for
synovial cells derived from the surface region, CD271
and CD55 as negative markers for synovial cells derived
from the stromal region, and CD271 as a positive
marker for synovial cells derived from the perivascular
region (Fig. 3a). We confirmed the two markers by
fluorescent immunostaining (Fig. 3b). CD55+ cells were
found in the surface region and were colocalized with
basement membrane positive for laminin. CD55 was
also expressed on endothelial cells, but the level was
lower than that of the surface region cells. CD271+ cells
were confirmed in perivascular cells positive for CD31,
while CD271 was not expressed in stromal cells negative
Ratio of synovial cells in the three regions by flowcytometric isolation
From the non-hematopoietic lineage cells, synovial cells
in the surface regions were sorted for CD55, those in the
stromal region were negatively sorted for CD55 and
CD271, and those in the perivascular region were sorted
for CD271 (Fig. 4a). The ratio of the gated cells to PI−
cells was 25% in the hematopoietic cells and 70% in the
non-hematopoietic cells (Fig. 4b). The ratio of the sorted
cells to other non-hematopoietic lineage cells was
approximately 5% in the surface region, 70% in the stromal
region, and 15% in the perivascular region (Fig. 4c). The
ratio of synovial cells derived from the stroma region
was statistically higher than from the surface or
Proliferation and colony formation in the bulk and sorted synovial cells
We prepared five fractions of synovial cells; the bulk
fraction, the non-hematopoietic and endothelial fraction,
the surface fraction, the stromal fraction, and the
perivascular fraction. No obvious morphological
differences were observed among the five fractions (Fig. 5a).
Proliferation was the highest in the perivascular fraction
(Fig. 5b). Colony morphology appeared similar and no
significant differences in colony number were obtained
among the five fractions (Fig. 5c and d).
Surface markers of colony-forming cells derived from the bulk and sorted synovial cells
Passage 3 colony-forming cells expressed CD44, CD73,
and CD90 at high rates (over 90%); CD55 and CD105 at
a moderate rate (50–90%); CD140b and CD271 at low
rates (approximately 10%); and did not express CD31,
CD45, and CD146 (less than 3%) (Fig. 5e). This
expression pattern was similar to MSCs. Though synovial cells
in the perivascular fraction were obtained after sorting
with CD 271, the positive rate decreased to only 5% after
Differentiation of colony-forming cells derived from the bulk and sorted synovial cells
After condrogenic induction, cell pellets from the five
fractions were differentiated into cartilage that stained
positive for safranin-o (Fig. 6a). The diameter of the
cartilage pellets was the largest in the perivascular fraction
among the five fractions (Fig. 6b). The perivascular
fraction showed higher mRNA expression of SOX9,
Aggrecan (ACAN), and COL10A1 in the cartilage pellets
After calcification induction, the colony-forming cells
derived from synovial cells stained positive for alizarin
red in the five fractions (Fig. 6c). Absorbance of alizarin
red, used to quantify differentiation was also the highest
in the perivascular fraction (Fig. 6d). Real-time RT-PCR
analyses showed similar expression levels of ALP and
RUNX2 for the calcification observed for each region
(Fig. 6f ). No expression of osteogenic genes, such as
OPN, OCN, OSX, and DLX5, was detected (data not
After adipogenic induction, colony-forming cells
derived from synovial cells stained positive for oil red-o in
the five fractions (Fig. 6e). No significant differences
were observed in triglyceride production among the five
fractions (Fig. 6f ). Real-time RT-PCR analyses showed
similar expression levels of PPARG and LPL for the
adipogenesis derived from each region (Fig. 6i). GTF3A and
CEBPA expression was lower for the surface fraction
than for the other four regions.
We used histological evaluations to classify the
synovium into three regions: the surface, stroma, and
perivascular regions. We then characterized the three
regions immunologically by immunostaining with 19
antibodies. Based on the results, we selected CD55 as a
positive marker for synovial cells derived from the
surface region, CD271 and CD55 as negative markers
for synovial cells derived from the stromal region,
and CD271 as a positive marker for synovial cells
derived from the perivascular region. The number of
synovial cells derived from the perivascular region
was much smaller than that derived from the stromal
region, but the colony-forming cells derived from the
perivascular region had higher proliferative and
We selected CD55, a complement inhibitor, as a specific
marker for the surface region because activation of
complement in the synovial membrane plays an important role
in the pathogenesis of osteoarthritis [
]. CD55 is also
recognized as a decay-accelerating factor (DAF) and is
expressed in the synovial lining during inflammation.
Synovial tissues derived from patients with osteoarthritis are
exposed to inflammatory conditions in the knee joint, so
the surface region is in constant contact with synovial
fluid containing inflammatory cytokines. Analysis of
MSCs positive for CD55 will therefore provide a better
clarification of the pathology of osteoarthritis.
We selected CD271 as a specific marker for the
perivascular region. CD271 is a low-affinity nerve growth
factor receptor (LNGFR) and serves as one of the two
receptor types for neurotrophins, a family of protein
growth factors that stimulate the survival and
differentiation of neuronal cells. These nerve cells in the
intraarticular tissues are thought to be important in
cartilage repair after cartilage damage, as demonstrated in
mouse studies [
]. CD271 also serves as a marker of
MSCs with high colony-forming ability in bone marrow
and synovium [
]. The role of CD271 in the
perivascular region of the synovium is unknown, but CD271
expression is possibly related to the pathophysiological
response of vascular developments/neurogenesis associated
with the synovitis of osteoarthritis. Functional analysis
CD271 expression by the CD271+ cells in the synovial
tissues will contribute to a greater understanding of the
healthy and diseased state.
Though synovial cells in the perivascular fraction were
obtained after sorting with CD 271, the positive rate
decreased to only 5% after colony formation. This means
that positive rate of CD271 decrease after expanding of
the sorted cells. There are some reports describing that
positive rate of CD271 decreased after expanding of
CD271+ MSCs [
]. Although the reason for the
reduction of CD271 expression is not clear, one
possibility is that the alteration of the environment from
in vivo three-dimensional surroundings, in which CD
271-positive cells located at synovial perivascular region,
to in vitro two-dimensional surroundings, in which CD
271-positive cells were expanded on culture dishes.
We selected CD55 and CD271 as negative specific
markers for the stroma region, as our initial attempts to
identify positive markers specific for the stroma region
were unsuccessful. Other negative markers specific for
the stroma region were CD55 (complement inhibitor),
Ki67, and Col4 (extracellular matrix marker).
The surface marker expression pattern in the
colonyforming cells derived from the bulk and sorted synovial
cells in each group was similar to that observed for
MSCs. These colony-forming cells also demonstrated
multi-potentiality, indicating that indicate they were
MSCs. This finding further confirmed that MSCs can be
obtained from synovium, irrespective of sorting. Even
without sorting, a small number of
hematopoieticlineage cells adhere to plastic dishes, and the rate of
adhesion of hematopoietic-lineage cells further decreases
after colony formation of the non-hematopoietic cells.
The MSCs derived from the perivascular fraction
showed the largest cartilage pellets and the highest
expression levels of SOX9 and Aggrecan mRNA during
chondrogenesis. The pellet size reflects the
chondrogenic potential for each population of MSCs, whereas
SOX9 and Aggrecan mRNA expression reflects the
chondrogenic potential of each individual MSC. Thus,
the MSCs derived from the perivascular fraction had the
highest chondrogenic potential, whether expressed per
population or per single cell. However, these MSCs also
showed the highest COL10A1 mRNA expression,
suggesting that their potential for hypertrophic chondrocyte
differentiation though this was not evident in
The perivascular fraction also showed the highest
absorbance of alizarin red, indicating pronounced
calcification. This high absorbance might reflect the high
number of cells present just before calcification
induction, because MSCs derived from the perivascular
fraction had a high proliferation capability. By contrast, the
other five fractions showed equivalent mRNA expression
of ALP and RUNX2 and no mRNA expression of other
osteogenic genes, such as DLX5, OSX, OPN, and OCN
(data not shown). Based on these findings, we assume
that this differentiation assay does not mimic the
osteogenesis of MSCs, as we mentioned previously [
this reason, we have used the term “calcification” rather
than “osteogenesis” [
The five fractions also showed equivalent triglyceride
production and mRNA expression of PPARG and LPL,
indicating similar adipogenesis activity. Conversely,
mRNA expression levels of GTF3A and CEBPA were
higher in the surface fraction than in the other fractions.
PPARG and CEBPA are early markers and LPL and
GTF3A are late markers of adipogenesis, so these
contradictory expression patterns cannot be explained
by different developmental stages. They may possibly
reflect the complexity of molecular regulation of
The perivascular fraction contained MSCs that were
superior in proliferation and cartilage differentiation
when compared to the MSCs in the other fractions.
Caplan et al. recently advocated that MSCs are derived
from pericytes, and our results support this hypothesis
]. However, our results also demonstrated that MSCs
are present in the surface and stromal regions, as well as
in the perivascular region, although the MSCs from the
surface and stromal regions showed poorer potentials
for proliferation and cartilage differentiation.
The MSCs from the three synovial regions showed
similar expression patterns of ten surface markers after
expansion, but demonstrated different patterns of
proliferation and differentiation. One reason for these
differences might be that the three populations are still
distinct, even though they show similar expression
patterns for the ten surface markers. Other surface markers
will therefore be useful to further distinguish these three
populations. For example, Mabuchi et al. reported that
the CD90+ and CD271+ fractions of bone marrow MSCs
differed in their proliferation ability with and without
expression of CD106 and CD49d [
]. Identification of
surface markers or transcription factors to distinguish
differentiation potentials among the three groups will be
important for clarifying synovial MSC biology.
A higher proliferation and chondrogenic potential was
observed in the present study MSCs derived from the
perivascular region, although the number of sorted cells
in the perivascular region was only approximately 20%
of that in the stromal region. The MSCs derived from
the stromal region showed poorer chondrogenic
potentials; however, because greater numbers of MSCs could
be prepared from that region than from the perivascular
region, these differences in chondrogenic potential might
be minimized. Therefore, from the standpoint of total
cells harvested, the synovial stroma region could be a
suitable MSC source.
The current study has three limitations that should be
considered. First, the staining for the immunological
characterization of surface, stromal, and perivascular
regions of human synovium was graded subjectively
and not objectively. Second, we selected CD55+ as the
marker for synovial cells from the surface region, but
because we investigated synovium derived from
osteoarthritis patients, these cells may be representative of
the inflammatory condition. Third, in vivo chondrogenesis
was not evaluated.
In conclusion, we selected CD55 as a positive marker for
synovial cells derived from the surface region, CD271
and CD55 as negative markers for synovial cells derived
from the stromal regions, and CD271 as a positive
marker for synovial cells derived from the perivascular
region. The number of synovial cells derived from the
perivascular region was much lower than that derived
from the stromal region, but the colony-forming cells
derived from the perivascular region had higher
proliferative and chondrogenic potentials.
We would like to thank Ms. Mika Watanabe and Ms. Kimiko Takanashi for the
management of our laboratory and Ms. Ellen Roider for English editing. Yuji
Kohno, Yusuke Nakagawa, Tomomasa Nakamura, Masafumi Horie, and
Toshifumi Watanabe contributed to the recruitment of the patients and the
acquisition of the written informed consent from the patients.
This study was supported by the Japan Society for the Promotion of Science
(JSPS) to MM (16H06262) and by the Highway Program for Realization of
Regenerative Medicine from the Japan Agency for Medical Research and
Development (AMED) to IS (JP16bm0504001).
Availability of data and materials
All the data supporting the results can be found in this manuscript and
supplemental data. Please contact the corresponding author for more data
MM and IS contributed to study conception and design. MM, YM, YO, SI, KK,
and SF contributed to acquisition of data. KO, NO, HKa, HKo, KT, CA, and TM
contributed to analysis and interpretation of data. All authors were involved
in drafting the article or revising it critically for important intellectual content,
and all authors approved the final version to be published. IS had full access
to all of the data in the study and takes responsibility for the integrity of the
data and the accuracy of the data analysis.
Ethics approval and consent to participate
This study was properly approved and certificated to be in compliance with
the Helsinki Declaration by the institutional review board of Tokyo Medical
and Dental University (reference number: M2000–2121). Written informed
consent forms were submitted by all the participating patients.
The authors declare that they have no competing interests.
Springer Nature remains neutral with regard to jurisdictional claims in
published maps and institutional affiliations.
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