Platelet-rich plasma enhances the proliferation of human adipose stem cells through multiple signaling pathways
Lai et al. Stem Cell Research & Therapy
Platelet-rich plasma enhances the proliferation of human adipose stem cells through multiple signaling pathways
Fangyuan Lai 0
Natsuko Kakudo 0
Naoki Morimoto 0
Shigeru Taketani 2
Tomoya Hara 0 1
Takeshi Ogawa 0
Kenji Kusumoto 0
0 Department of Plastic and Reconstructive Surgery, Kansai Medical University , 2-5-1 Shin-machi, Hirakata, Osaka 573-1010 , Japan
1 Department of Oral Implantology, Osaka Dental University , Osaka 573-1121 , Japan
2 Department of Microbiology, Kansai Medical University , Osaka 573-1010 , Japan
Background: Platelet-rich plasma (PRP) is an autologous blood product that contains a high concentration of several growth factors. Platelet-derived growth factor (PDGF)-BB is a potential mitogen for human adipose-derived stem cells (hASCs). PRP stimulates proliferation of hASCs; however, the signaling pathways activated by PRP remain unclear. Methods: hASCs were cultured with or without PRP or PDGF-BB, and proliferation was assessed. hASCs were also treated with PRP or PDGF-BB with or without imatinib, which is a PDGF receptor tyrosine kinase inhibitor, or sorafenib, which is a multikinase inhibitor. Inhibition of cell proliferation was examined using anti-PDGF antibody (Abcam, Cambridge, UK), by cell counting. We assessed the effects of inhibitors of various protein kinases such as ERK1/2, JNK, p38, and Akt on the proliferation of hASCs. Results: The proliferation was remarkably promoted in cells treated with either 1% PRP or 10 ng/ml PDGF-BB, and both imatinib and sorafenib inhibited this proliferation. Anti-PDGF antibody (0.5 and 2 μg/ml) significantly decreased the proliferation of hASCs compared with control. PRP-mediated hASC proliferation was blocked by inhibitors of ERK1/2, Akt, and JNK, but not by an inhibitor of p38. Conclusions: PRP promotes hASC proliferation, and PDGF-BB in PRP plays a major role in inducing the proliferation of hASCs. PRP promotes hASC proliferation via ERK1/2, PI3K/Akt, and JNK signaling pathways.
Human adipose-derived stem cells (hASCs) were first
isolated from human adipose tissue and identified by
Zuk et al. in 2001 [
]. These cells can differentiate
toward multiple lineages, such as osteogenic [
], adipogenic [
], cardiac [
], epidermal [
] lineages. hASCs are used widely in the
field of regenerative medicine, including to promote
bone regeneration [
], tooth and periodontal
], cartilage regeneration [
], wound healing [
], and nerve regeneration to cure Parkinson’s disease
, as well as to suppress aging [
]. Due to the
advantages of the autologous source of these cells and their
relative abundance and ease of isolation, hASCs have
also been widely used in the fields of plastic surgery and
regenerative medicine [
However, the proliferation and differentiation
capacities of hASCs decrease with age [
], body mass
index , diabetes mellitus [
], radiation exposure
, and tamoxifen treatment [
]. hASCs account for
about 16–30% of the stromal vascular fraction [
obtain a sufficient amount of cells for therapeutic
purposes, in-vitro proliferation of the cells is required. Fetal
bovine serum (FBS) is widely used for this purpose in
multiple types of cells in vitro. However, due to the risk
of heterologous immunization and zoonosis, FBS has
limited clinical use.
Platelet-rich plasma (PRP) is a blood portion that is
enriched with platelets [
]. Upon activation, platelets in
PRP release granules containing molecules including
growth factors and regulatory proteins, such as
plateletderived growth factor (PDGF), epidermal growth factor
(EGF), insulin-like growth factors (IGFs), transforming
growth factor beta (TGF-β), vascular endothelial growth
factor (VEGF), and others [
]. These growth factors
play important roles in cell proliferation, migration, and
Our previous study revealed that activated PRP has a
potential effect on the proliferation of hASCs and
human dermal fibroblasts (hDFs) compared with
nonactivated PRP [
]. Furthermore, we also reported that
activated PRP induces hDF proliferation via the
activation of ERK1/2 signaling [
]. Recently, other
investigators reported that PDGF also enhances proliferation of
hASCs through the JNK pathway [
]. However, the
signaling pathways involved in PRP-stimulated proliferation
of hASCs have not been clarified.
In the present study, we show that PRP stimulated cell
proliferation by ERK1/2, JNK, and Akt activation. We
compared this effect with the proliferative effect of
PDGF-BB, a major growth factor in PRP.
Preparation of activated PRP
Activated PRP was obtained using the double-spin
method as described previously [
]. Briefly, after
obtaining informed consent from healthy adult
volunteers (n = 3), blood was collected in tubes containing an
acid-citrate-dextrose solution formula A anticoagulant,
and spun in a standard laboratory centrifuge for 7 min
at 450×g. The yellow plasma with buffy coat, containing
platelets, leukocytes, and some erythrocytes from two
tubes, was collected in a monovette via a long cannula
and centrifuged for 5 min at 1600×g. Platelets that
accumulated in the thrombocyte pellet in 1.0 ml plasma were
used as PRP. A separate sample of 8 ml blood was
allowed to stand for 30 min at room temperature in a tube
without anticoagulant and then spun for 8 min at 2015×g.
The supernatant was collected as an autologous thrombin.
A 1:1 (v/v) mixture of 0.5 M CaCl2 and autologous
thrombin was prepared in advance as an activator. A 10:1 (v/v)
mixture of PRP and activator was incubated for 10 min at
room temperature. Activated PRP was centrifuged at 90×g
and then 9000×g for 10 min each; the supernatant was
filtered through a 0.22-μm membrane (Millex GP; Merck
Millipore, Tullagreen, Carrigtwohill, Co. Cork, Ireland)
and stored at −80 °C until use.
Measurement of platelet concentrations and growth factor levels
The number of platelets in whole plasma and PRP was
counted using an XE-2100 automated hematology
system (Sysmex Corp., Tokyo, Japan). PDGF-BB, IGF, and
EGF levels in whole plasma and activated PRP were
determined using commercially available ELISA kits (R&D
Systems, Minneapolis, MN, USA), according to the
Isolation of hASCs
Unnecessary adipose tissue was obtained from a
61year-old male patient who had previously provided
informed consent and underwent plastic surgery.
hASCs were isolated using a method described
]. After washing extensively with
phosphatebuffered saline (PBS), the adipose tissues were cut
into small pieces and incubated with 3 volumes of 0.
1% collagenase (Sigma-Aldrich, St. Louis, MO, USA)
solution with constant shaking at 40 °C for 40 min.
After adding DMEM containing 10% FBS (Hyclone,
Logan, UT, USA) and antibiotics (complete medium),
the tissue was centrifuged at 400×g for 3 min. After
removing cellular debris through a 100-μm nylon
mesh (BD Falcon, Bedford, MA, USA), the cells were
incubated in complete medium in a dish. The
primary hASCs were cultured for 4–5 days until they
reached confluence. These cells were defined as
passage “0”. For all experiments, cells from passages 7–
9 were used.
Cell proliferation assay
For the cell proliferation assays, hASCs were seeded at a
density of 1.0 × 104 cells/well in 24-well culture plates
and incubated in complete medium overnight. The cell
medium was then replaced with serum-free DMEM.
After 6 h of incubation, hASCs were treated with PRP or
human recombinant PDGF-BB (PeproTech EC Ltd, London,
UK) at the stated concentrations in serum-free DMEM for
48 h. Inhibitors included the PDGF receptor tyrosine kinase
inhibitor imatinib (Wako Co., Ltd, Tokyo, Japan), the
multikinase inhibitor sorafenib (AdooQ, Irvine, CA, USA),
the MEK inhibitor PD98059, the
phosphatidylinositol-3kinase-Akt inhibitor LY294002, the p38 inhibitor SB203580
(Calbiochem-Novabiochem, San Diego, CA, USA), and the
JNK inhibitor SP600125 (Sigma). Inhibitors were added 1 h
before the incubation with PRP or PDGF-BB. Cell
proliferation was determined using Cell Counting Kit-8 (Dojindo
Molecular Technologies, Kumamoto, Japan), according to
the manufacturer’s instructions. Absorbance was read at
450 nm on a multiwell plate reader (EnSpire 2300 Multilabel
Reader; PerkinElmer, Inc., Waltham, MA, USA).
To estimate the cell number from the absorbance, a
standard curve was established. hASCs were seeded at
densities of 0, 6250, 12,500, 25,000, 50,000, and 100,000
cells/well in 24-well plates for 3 h with 10% FBS in
DMEM. The cells were then incubated with Cell
Counting Kit-8 solution for 1 h, and the absorbance was read.
The standard curve was established by plotting the
number of cells versus the absorbance.
hASCs were seeded at a density of 2 × 103 cells/well in
96-well culture plates containing complete medium.
After overnight incubation, the hASCs were first starved
in serum-free DMEM for 6 h and then treated with PRP,
PDGF-BB, human recombinant IGF, or human
recombinant EGF (PeproTech EC Ltd) in serum-free DMEM
for 48 h. Inhibitors were added 1 h before incubation
with PRP or PDGF-BB. Quantification of cell
proliferation was determined using the Cell Proliferation ELISA
BrdU kit (Roche, Mannheim, Germany).
Cell cycle assay
hASCs (1 × 106 cells) were seeded in a 10-cm culture
dish containing complete medium and cultured
overnight. The medium was then replaced with serum-free
DMEM for 6 h, and the cells were treated with reagents
at the stated concentrations for 48 h. Treated cells were
collected by trypsinization. After washing with ice-cold
PBS twice, cells were fixed in 70% ethanol at −20 °C for
3 h. The fixed cells were then stained with Muse™ Cell
Cycle reagent (Millipore) in the dark at room
temperature for 30 min. Cell cycle phases were analyzed
by flow cytometric quantification of DNA with the
Muse™ Cell Analyzer (Millipore).
Western blot analysis
The cells were treated with the indicated compounds
and lysed. Extracted cellular proteins (20 μg) were
separated by sodium dodecyl sulfate-polyacrylamide
gel electrophoresis (SDS-PAGE) and then transferred
to a polyvinylidene difluoride (PVDF) membrane. The
membrane was first blocked with Blocking One-P reagent
(Nacalai Tesque, Kyoto, Japan) for 30 min at room temperature,
and then incubated with the following primary antibodies:
rabbit anti-phospho-ERK1/2 (1:1000; Epitomics Inc.,
Burlingame, CA, USA), rabbit anti-phospho-Akt, rabbit
anti-Akt (1:5000; Abcam, Cambridge, UK), rabbit
antiERK1/2 (1:1000; Cell Signaling Technology, Beverly, MA,
USA), or rabbit anti-β-actin (1:1000; Cell Signaling
Technology) at 4 °C overnight. This was followed by
incubation with peroxidase-linked secondary antibody (1:
20000; GE Healthcare, Little Chalfont, UK) at room
temperature for 30 min. The labeled proteins were
detected with enhanced chemiluminescence using the Prime
Western blotting detection system (GE Healthcare).
In-vitro JNK activity assay
Activity of the JNK pathway was analyzed with the
SAPK/JNK Kinase Assay Kit (Cell Signaling
Technology). Briefly, cell lysates were immunoprecipitated with
the anti-phospho-JNK antibody coupled to JNK
sepharose beads. The concentrated active JNK protein was
then reacted with the substrate, c-Jun fusion protein, in
the presence of ATP. The reaction mixture was
separated with SDS-PAGE and transferred to a PDVF
membrane. The membrane was incubated with rabbit
antiphospho-c-Jun (1:1000) at 4 °C overnight, followed by
incubation with peroxidase-linked anti-rabbit IgG at
room temperature for 30 min. The labeled proteins were
also detected with enhanced chemiluminescence using
the Prime Western blotting detection system. All of the
experiments were replicated three times.
Data are presented as the mean ± standard deviation
(SD). The Mann–Whitney U test was used to evaluate
differences among groups. P < 0.05 and P < 0.01 were
considered statistically significant.
Concentrations of platelet and growth factors in PRP and blood
Compared with whole plasma, PRP showed a 10.1-fold
enrichment in platelets and a 25.9-fold enrichment in
PDGF-BB. In contrast, the concentrations of EGF were
comparable between PRP and whole plasma. IGF in PRP
was present at a concentration of only 60% that in whole
plasma (Table 1).
PRP stimulated proliferation of hASCs
Cell proliferation was increased by treatment with 0.2%
PRP (P < 0.01 vs control), and 1% PRP stimulated cell
proliferation to a greater extent (P < 0.01 vs control and
P < 0.01 vs 0.2% PRP). Thus, PRP stimulated
proliferation of hASCs in a dose-dependent manner between 0
and 1% PRP (Fig. 1a). The proliferation was decreased
when cells were treated with 3% and 5% PRP, compared
with 1% (data not shown). The cell growth stimulated by
PRP was confirmed by observation with phase-contrast
microscopy (Fig. 1b).
PRP containing PDGF-BB promoted hASC proliferation
Cell proliferation of hASCs was also enhanced by
treatment with 2 ng/ml PDGF-BB (P < 0.05 vs control).
PDGF-BB at a concentration of 10 ng/ml markedly
stimulated cell proliferation (P < 0.01 vs control and P < 0.01
vs 0.2% PRP). PDGF-BB displayed a dose-dependent
stimulation of hASC proliferation between 0 and 10 ng/
ml (Fig. 2a). Treatment with imatinib (5 μM) or
sorafenib (5 μM) reduced the PRP-stimulated hASC
proliferation (Fig. 2b). Similarly, both imatinib and sorafenib
significantly inhibited the proliferation of hASCs
induced by PDGF-BB (10 ng/ml). Inhibition with sorafenib
was more potent than that with imatinib (Fig. 2c).
Furthermore, treatment with anti-PDGF antibody inhibited
PRP-stimulated growth of hASCs in a dose-dependent
manner (Fig. 2d).
226 ± 6
145 ± 4
PDGF platelet-derived growth factor, EGF epidermal growth factor, IGF insulin-like growth factor, PRP platelet-rich plasma
PDGF-BB stimulated DNA synthesis
Cell proliferation of hASCs was also evaluated using
BrdU incorporation assays. Compared with control, PRP
induced a 5.42-fold increase in BrdU incorporation, and
imatinib decreased the incorporation 2.97-fold. Thus,
PRP significantly stimulated DNA synthesis in hASCs,
and imatinib inhibited DNA synthesis (Fig. 3a).
We also tested the stimulating effect of growth factors.
PDGF-BB markedly enhanced DNA synthesis (10.08-fold vs
control). In contrast, treatment with either IGF-I (10 ng/ml)
(3.33-fold vs control) or EGF (10 ng/ml) (2.02-fold vs
control) showed a minimal effect on DNA synthesis in hASCs.
PDGF-BB plus either IGF or EGF stimulated DNA synthesis
to a greater extent than IGF or EGF alone (Fig. 3b).
Promotion of cell cycle transition from G0/G1 to S phase by PRP and PDGF-BB
When treated with PRP compared to control, the flow
cytometry showed a trend in which cells in the S and
G2/M phases increased (Fig. 4a). A histogram of the
flow cytometry results is shown in Fig. 4b. The percent
of cells treated with PRP that were in the S phase (15.44
± 7.31%) was significantly higher compared with control
(3.67 ± 0.91%). Similarly, the percent of cells treated with
PRP that were in the G2/M phase (32.11 ± 5.5%) was
also significantly higher compared with control (13.61 ±
6.63%). The percent of cells treated with PRP plus
imatinib in the S and G2/M phases (8.81 ± 3.27% and 17.28
± 3.15%, respectively) and the percent treated with
PDGF-BB (9.01 ± 4.54% and 19.75 ± 2.97%, respectively)
were not significantly different compared to control (3.
67 ± 0.91% and 13.61 ± 6.63%, respectively). PRP
stimulated cell progression to the S and G2/M phases, and
imatinib inhibited this effect. PDGF-BB showed a small
effect on the cell cycle (Fig. 4a, b).
Activation of ERK1/2, Akt, and JNK signaling pathways with PRP and PDGF-BB
To examine the signaling pathways involved in
stimulation of hASCs by PRP, cells were treated with an
ERK1/2 inhibitor (PD98059, 20 μM), PI3K/Akt
inhibitor (LY294002, 10 μM), JNK inhibitor (SP600125,
20 μM), or p38 inhibitor (SB203580, 10 μM).
PRPinduced cell proliferation was suppressed by PD98059,
LY294002, and SP600125, but not SB203580 (Fig. 5).
The signaling pathways activated by these treatments
were further analyzed in hASCs with western blotting and
a JNK activity assay. Phosphorylation of ERK1/2 and Akt
increased following treatment with PRP or PDGF-BB.
Imatinib inhibited the phosphorylation of these enzymes
in the presence of PRP (Fig. 6a, b). Phosphorylation of
JNK was not detected under these conditions. Next, we
measured the activity of JNK using the substrate c-Jun. As
shown in Fig. 6b, PRP markedly activated JNK. Thus,
stimulation of cell growth by PRP was mediated through
multiple signaling pathways.
We demonstrated that PRP enhanced the proliferation
of hASCs through multiple signaling pathways by
activating ERK1/2, JNK, and Akt. The proliferative effect
of PRP, which was similar to the proliferative effect of
PDGF-BB alone, was inhibited by the tyrosine kinase
inhibitor imatinib and by the multikinase inhibitor
sorafenib. The proliferative effect of PRP was also lowered by
adding anti-PDGF antibody to the medium, indicating
that PDGF-BB, which was abundant in activated PRP,
played a major role in the proliferative effect of PRP. In
addition, PRP induced the proliferation of cells in the S
phase of the cell cycle, concomitant with an increase in
BrdU uptake. Addition of PRP activated ERK, JNK, and
Akt, and PRP-mediated proliferation was blocked by
inhibitors of these signal transduction enzymes. On the
other hand, PDGF-BB alone only slightly activated ERK,
JNK, and Akt (Fig. 6a, b). These results indicated that
other factors in PRP function in the additive effect on
PRP is enriched in platelets, which were collected by
centrifugation of autologous blood. Cytokines including
PDGF, TGF-β, VEGF, IGF, EGF, and basic fibroblast
growth factor (bFGF) are contained in α-granules of
platelets. PRP was collected without coagulation, and
was then activated by adding autologous thrombin and
calcium chloride. Growth factors in activated PRP are
indispensable for the proliferation of various types of
]. We now demonstrated that PRP was a potent
inducer of proliferation of hASCs.
Competence activity is known to be stimulated by
factors that can make cells become “competent” to replicate
their DNA and divide. Competence growth factors
include PDGF [
] and FGF-2 [
]. PDGF and FGF-2
alone act on cells that are in either the G0 or early G1
phase of the cell cycle, rendering them competent to
initiate DNA replication [
]. In contrast, progression
activity refers to activity mediated by factors that can
dictate the ultimate fraction of competent cells that
enters the S phase [
]. These typical progression growth
factors are EGF [
] and IGF-I [
]. The progression
growth factors allow cells to progress through the
prereplicative phase of the cycle, inducing cells to enter the
S, G2, and M phases. PRP comprises a large amount of
competence growth factors, such as PDGF and FGF-2,
and the progression growth factors, EGF and IGF-I. This
study strongly suggests that these competence and
progression growth factors act on hASCs in a concerted
and compounding manner, and progress the cell cycle
from the G0 phase to G1 and S phases.
Cell cycle progression is regulated by the expression of
cyclins. Cyclins are factors that bind to and activate the
cyclin-dependent kinases (CDKs). There are
approximately 20 kinds of cyclins, such as cyclin A2, B1, and
D1, and several types of CDKs such as CDK1, CDK2,
and CDK4. These factors and kinases are known to
control cell cycle progression by binding with each other in
different combinations. Cyclin D is expressed in
response to mitogens, and then binds with CDK4 or
CDK6. The formed cyclin D complex phosphorylates the
target protein, progressing the cell cycle from the G1 to
the S phase. It was reported that the expression of cyclin
D1 in hASCs increased with the transition of the cell
cycle from the G1 to the S phase [
]. In addition, the
cdc2/cyclin B complex was reported to regulate the G2/
M phase transition [
]. This study found that the
addition of PRP led to an increase in the proliferation of
hASCs in the S and M phases, implying a possible
involvement of cyclin D1 and B1.
The proliferative effect of PRP on preadipocytes [
], and bone marrow mesenchymal stem cells
] has been reported. Furthermore, PRP can enhance
the proliferative effect of mesenchymal progenitor cells by
activating the ERK signaling pathway, and PDGF-BB is a
key factor in this stimulation [
]. Also, the proliferative
effect on chondrocytes can be significantly stimulated by
PRP via the ERK signaling pathway, and platelet-derived
adenosine diphosphate in PRP is a key mediator of
]. We found that PRP induces the proliferation
of hDFs by the activation of ERK1/2 signaling [
Proliferation of hASCs induced by PRP was reported by
Kakudo et al. [
] and Gersch et al. [
], but the signaling
pathways remain unclear. PDGF-BB stimulates DNA
synthesis in hASCs and cell proliferation, and these effects
are mediated by JNK activation [
] or Akt activation [
Also, FGF-2 [
], EGF [
], or VEGF [
] induces the
proliferation of hASCs through ERK1/2 activation. Our
present study showed that the addition of PRP to hASCs
activated JNK, ERK1/2, and Akt, and the addition of
inhibitors of these kinases reduced the proliferative activity.
Because PRP contains abundant PDGF-BB, FGF-2, EGF,
and VEGF, the interaction among these growth factors
may stimulate cell proliferation through multiple signaling
pathways. The addition of PD98059, SP600125, or LY294002
to PRP-treated hASCs partially inhibited cell proliferation,
which supports this conclusion. Hye Kim et al. [
that an isoform of PDGF, PDGF-D, showed a strong
proliferation effect on hASCs, and thus PDGF-D present in PRP
may also induce proliferation of hASCs through the ERK1/2
and Akt pathways.
It is noted that the use of stem cells for therapeutic
applications is influenced by their proliferative and
differentiation potential, which is affected by the age of the
donors. It was previously reported that compared with
young cells, aged hASCs exhibited increased cellular
senescence features [
], a decline in both stromal
vascular fraction (SVF) cell yield [
] and hASC
proliferation rate [
], a decreased differentiation potential
 toward adipogenic [
41, 43, 44
], osteogenic [
and chondrogenic  lineages, negative effects on
hASC frequency [
], fewer progenitor cell numbers
], and impaired migration ability [
]. Based on these
reports, we believe that hASCs obtained from older
patients may have limitations in clinical application. Our
study had limited sources of adipose acquisition due to
the lack of younger patients, and thus it was impossible
to compare the differences between young hASCs and
aged hASCs. However, based on our experimental
results, as proliferation of aged hASCs can still be
stimulated by PRP, we speculate that young hASCs will have
higher proliferative ability by PRP stimulation.
In future studies, we will use young hASCs to examine
the effects of PRP on proliferation promotion.
We found that both PRP and PDGF-BB can induce the
proliferation of hASCs by activating ERK1/2, Akt, and JNK
signaling pathways. This study clarified that PDGF-BB
present in PRP plays an essential role in the proliferation
via multiple signaling pathways, and is not limited to
stimulation by PDGF-BB. The reason for the potent effect of
PRP may be due to the presence of various factors involved
in a variety of proliferative activities.
Thus, PRP is a powerful promoter to proliferate
hASCs in vitro. Future studies are required to clarify the
interaction of these factors that are present in PRP.
The authors thank the Central Research of Laboratory of the Kansai Medical
University for their technical assistance.
This work was supported by the research grant D2 from Kansai Medical University.
Availability of data and materials
All data generated and/or analyzed during this study are included in this
FL conceived the work, acquired data, drafted the manuscript, and approved
the final version. NK and ST conceived the work, revised the manuscript, and
approved the final version. TH and TO acquired data, and approved the final
version. NM and KK revised the manuscript, and approved the final version.
All authors read and approved the final manuscript.
Ethics approval and consent to participate
The study was approved by the Ethics Review Board of Kansai Medical
University in accordance with the ethical guidelines of the Helsinki
Declaration of 1975. All specimens were collected and used with informed
consent from the donors.
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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