- Research article
- Open Access
Characterization of adipocyte differentiation from human mesenchymal stem cells in bone marrow
© Qian et al; licensee BioMed Central Ltd. 2010
- Received: 31 December 2009
- Accepted: 7 May 2010
- Published: 7 May 2010
Adipocyte hyperplasia is associated with obesity and arises due to adipogenic differentiation of resident multipotent stem cells in the vascular stroma of adipose tissue and remote stem cells of other organs. The mechanistic characterization of adipocyte differentiation has been researched in murine pre-adipocyte models (i.e. 3T3-L1 and 3T3-F442A), revealing that growth-arrest pre-adipocytes undergo mitotic clonal expansion and that regulation of the differentiation process relies on the sequential expression of three key transcription factors (C/EBPβ, C/EBPα and PPARγ). However, the mechanisms underlying adipocyte differentiation from multipotent stem cells, particularly human mesenchymal stem cells (hBMSCs), remain poorly understood. This study investigated cell cycle regulation and the roles of C/EBPβ, C/EBPα and PPARγ during adipocyte differentiation from hBMSCs.
Utilising a BrdU incorporation assay and manual cell counting it was demonstrated that induction of adipocyte differentiation in culture resulted in 3T3-L1 pre-adipocytes but not hBMSCs undergoing mitotic clonal expansion. Knock-down and over-expression assays revealed that C/EBPβ, C/EBPα and PPARγ were required for adipocyte differentiation from hBMSCs. C/EBPβ and C/EBPα individually induced adipocyte differentiation in the presence of inducers; PPARγ alone initiated adipocyte differentiation but the cells failed to differentiate fully. Therefore, the roles of these transcription factors during human adipocyte differentiation are different from their respective roles in mouse.
The characteristics of hBMSCs during adipogenic differentiation are different from those of murine cells. These findings could be important in elucidating the mechanisms underlying human obesity further.
- Adipocyte Differentiation
- Adipogenic Differentiation
- Vinylidene Difluoride
- Cell Cycle Alteration
- FABP4 Expression
Increased adipose tissue mass associated with obesity is due to the increased number and size of adipocytes [1, 2]. Adipocyte differentiation from mesenchymal stem cells plays an important role in the hyperplasia of adult adipose tissue. A population of cells resident in the vascular stroma of adipose tissue can differentiate into adipocytes in vitro and in vivo . Recent studies indicate that pericytes in blood vessel walls have adipogenic potential, express mesenchymal stem cell (MSC) markers and are multipotent . In addition to resident stem cells, non-resident stem cells can serve as a source of adipocyte precursors; bone marrow MSCs can be recruited to adipose tissue and generate new adipocytes in response to treatment with thiazolidinediones (TZDs) or high fat stimulation .
The characteristics and molecular mechanism underlying adipocyte differentiation have been extensively investigated in the murine pre-adipocyte cell lines 3T3-L1 and 3T3-F442A [6, 7]. Growth-arrested pre-adipocytes have been shown to re-enter the cell cycle synchronously and undergo mitotic clonal expansion in response to MDI (M: methyl-isobutyl-xanthine, D: dexamethasone, I: insulin) treatment, before exiting the cell cycle and terminally differentiating . The transcription factors C/EBPβ (CCAAT/enhancer binding protein β), C/EBPα (CCAAT/enhancer binding protein α) and PPARγ (peroxisome proliferator-activated receptor γ) act sequentially during 3T3-L1 pre-adipocyte differentiation . C/EBPβ is induced immediately after exposure to the differentiation cocktail, resulting in phosphorylation and activation [10, 11], and it transactivates the expression of C/EBPα and PPARγ . C/EBPα and PPARγ, together or in isolation, can initiate differentiation without inducers [13–15]. C/EPBα is believed to be relevant to the acquisition of insulin sensitivity .
MSCs have been isolated and induced to differentiate into adipocytes in a variety of organs [17–22]. However, the differentiation procedure and the roles of adipose-related genes in that procedure have not been characterized completely owing to the heterogeneity, low proliferation ability and ineffective ectopic gene transfection of hBMSCs [23, 24]. Human primary cells are of great interest because of their biological and therapeutic potential, therefore this study extends the research carried out in murine 3T3-L1 cells to hBMSCs from bone marrow.
Isolation and adipogenic differentiation of hBMSCs
Cell cycle alteration during adipocyte differentiation from hBMSCs
BrdU incorporation assays were performed to investigate whether DNA synthesis occurs during adipocyte differentiation from hBMSCs. We found that differentiated hBMSCs were BrdU negative, while differentiated 3T3-L1 cells were BrdU positive (Figure 3C). Confocal microscopy verified the positional relationship between nuclei (as indicated by BrdU incorporation into DNA) and cells with lipid droplets in the cytoplasm (Figure 3D, 3E). These results demonstrate that hBMSCs did not undergo mitotic clonal expansion during adipogenic differentiation under culture conditions.
Role of C/EBPβ in adipocyte differentiation of hBMSCs
C/EBPβ was over-expressed in hBMSCs using an adenovirus expression system (Figure 4D) to investigate its function during differentiation. Control cells expressing Lac Z didn't differentiate, while expression of exogenous C/EBPβ alone induced adipogenesis (Figure 4E), and some cells presented with small intracellular fat droplets that could not be adequately stained using oil red O. However, FABP4 expression was detected by western blotting (Figure 4F) and was significantly up-regulated by the addition of inducers, the highest levels of expression being evident when indomethacin (PPARγ agonist) was included (Figure 4E, 4F).
Role of C/EBPα in adipocyte differentiation from hBMSCs
Role of PPARγ in adipocyte differentiation from hBMSCs
HBMSCs are more difficult to handle than mouse stem cell lines but their importance and therapeutic potential necessitate their use in research of the type outlined herein. The previous studies are focused on the mouse stem cell lines but the regulation of them could be different in some aspects, and results of murine cells would be less convincing in interpreting the onset of human disease. On the other hand, adipocytes differentiated from HBMSCs would be of better immuno-compatibility in autograft for plastic purpose. So, in this study, a comprehensive analysis of adipocyte differentiation from multipotent human stem cells was carried out.
HBMSCs were isolated from bone marrow and induced to differentiate into adipocytes under culture conditions. The PPARγ agonist, indomethacin, was added as well as the conventional inducers used in adipocyte differentiation protocols for murine pre-adipocytes. HBMSCs behaved differently from 3T3-L1 pre-adipocytes, with only a small number of cells differentiating into adipocytes after one cycle of treatment; approximately 60%~70% of hBMSCs differentiated into adipocytes after three cycles of treatment (Figure 1B). A long G0 phase and a lack of contact inhibition (Figure 2C) meant that hBMSCs did not synchronize at the time when differentiation was initiated (Figure 2B). Growth arrest is a prerequisite for adipocyte differentiation , so it was concluded that only a minority of hBMSCs were growth arrested when differentiation was induced.
MCE (mitotic clonal expansion) is an essential event associated with adipocyte differentiation from mouse pre-adipocyte cell lines [8, 11]. However, it is not known whether MCE is required for adipocyte differentiation from all cell types. We have previously demonstrated that committed C3H10T1/2 cells treated with BMP4 divide when induced to differentiate , and primary cultures of mouse embryonic fibroblasts (MEF) undergo MCE when differentiating into adipocytes . In this study, hBMSCs from bone marrow did not undergo division during differentiation (Figure 3), which is in agreement with other reports showing that adipose precursor cells prepared from human adipose tissue (hADSCs) did not divide during differentiation under culture conditions . The authors argued that hADSCs had completed division before being isolated; however, hADSCs are multipotent and can differentiate into other cell lineages including adipocytes ex vivo [31, 32]. HADSCs could behave similarly to hBMSCs from bone marrow under culture conditions and remain uncommitted. The diversity of cell cycle alterations during adipocyte differentiation could be species-specific.
Murine proteins and comparable human proteins can function differently in the same context. In this study, C/EBPβ expression in hBMSCs did not alter significantly during the early stages of induction whereas expression was up-regulated immediately following induction and declined after two days in 3T3-L1 pre-adipocytes . The decline of C/EBPβ at 14 day might result from most of cells being terminal differentiated. However, C/EBPβ was required for adipocyte differentiation in hBMSCs as its knock-down expression impaired differentiation (Figure 4B, 4C). C/EBPβ has important roles in mitosis and terminal adipocyte differentiation [34, 35], but mitosis did not occur during differentiation of hBMSCs (Figure 3) and that could possibly explain the lack of differential expression of C/EBPβ upon induction. It is likely that the role of C/EBPβ during adipocyte differentiation from hBMSCs relates to its modification and not its expression levels, although the importance of C/EBPβ phosphorylation requires further investigation.
C/EBPβ or C/EBPα is sufficient to induce 3T3-L1 pre-adipocytes to differentiate into mature adipocytes without using inducers [36, 37]. Over-expression of C/EBPβ alone stimulated differentiation of hBMSCs, as evidenced by FABP4 expression (Figure 4E, 4F). C/EBPα was less effective than C/EBPβ as expression of C/EBPα alone did not stimulate differentiation (Figure 5E, 5F). C/EBPβ and C/EBPα individually enhanced adipocyte differentiation of hBMSCs dependent on exogenous hormone agent treatment, particularly in the presence of a PPARγ activator (Figure 4E, 4F, 5E, 5F). HBMSCs may lack endogenous PPARγ ligands; however, this cannot be determined at this time because the results concerning the study of natural PPARγ ligands are indecisive .
PPARγ plays pivotal roles in adipocyte differentiation as it induces adipogenesis in cultured mouse fibroblasts . With the use of high affinity, selective PPARγ agonists, PPARγ activation stimulates 3T3-F442A cells to develop into mature fat cells with a phenotype that includes morphological changes, lipid accumulation, and the acquisition of insulin sensitivity . In addition, ectopic expression of PPARγ in hBMSCs initiates adipocyte differentiation. However, these cells were immature adipocytes, as demonstrated by morphological observations and the expression of some adipocyte-specific genes (Figure 6G, 6H). In humans, PPARγ functions to regulate a part of genes required for adipocyte maturation, as demonstrated by its ability to induce FABP4 but not GLUT4 expression (Figure 6H). In addition, PPARγ could play a role in cytoskeletal alterations associated with the morphological changes during differentiation, as the cells rounded up when PPARγ was over-expressed and elongated when expression of PPARγ was knocked down.
This study demonstrates that the characteristics of hBMSCs during adipogenic differentiation are different from those of mouse cells. HBMSCs do not undergo mitotic clonal expansion during adipocyte differentiation. C/EBPβ, C/EBPα, and PPARγ are all required but not sufficient for adipocyte differentiation from hBMSCs. The ability of the transcription factors to stimulate adipocyte differentiation differed between human and murine cells. Further studies concerning on how C/EBPβ, C/EBPα and PPARγ regulating human adipocyte differentiation could help to elucidate the molecular mechanism of adipocyte differentiation from human stem cells, help to elucidate the mechanisms underlying human obesity and identify therapeutic targets.
Bone marrow was obtained from the iliums of patients undergoing iliac crest bone grafts following informed consent. Five samples were obtained from male patients between the ages of 25 and 55 years who did not suffer from obesity and/or diabetes. The sample collection procedure and related research work was approved by the ethics committee of Institutes of Biomedical Sciences, Fudan University. Results were reproducible between donors, and the data presented in the results section were from a 32-year-old male donor.
Isolation and adipogenic differentiation of hBMSCs
HBMSCs were isolated by density gradient centrifugation with Ficoll-Paque (GE Healthcare) and plastic adherence and grown in DMEM (low glucose, Invitrogen) containing 10% fetal bovine serum and 1% antibiotics; cells from passages 3-5 were used experimentally. A published protocol was followed to induce adipogenic differentiation of hBMSCs . HBMSCs were cultured at a density of 5000~6000 cells/cm2. After reaching confluence, hBMSCs were cultured for one more week and induced in adipogenic medium containing 0.5 mM isobutyl-methylxanthine (Sigma-Aldrich), 1 μM dexamethasone (Sigma-Aldrich), 10 μM insulin (Roche), 100 μM indomethacin (Sigma-Aldrich) for three days and maintained in medium with 10 μM insulin for one day. The treatment was repeated two or three times, after which the cells were maintained in DMEM with 10 μM insulin until day 21 and subjected to oil red O staining to detect cytoplasmic triglyceride.
Oil red O staining
Cells were washed three times with PBS and then fixed for 2 min with 3.7% formaldehyde. Oil red O (0.5% in isopropanol) was diluted with water (3:2) filtered through a 0.45 μm filter and incubated with the fixed cells for 1 h at room temperature. Cells were washed with water and the stained fat droplets in the cells were visualized by light microscopy and photographed. The percentage of differentiated cells was determined by counting cells based on oil red staining in the lipid vacuoles and 4',6'-diamidino-2-phenylindole staining of DNA.
At various time points cells were washed with cold PBS (pH 7.4) and lysed with lysis buffer (2% SDS, 60 mM Tris-Cl, pH 6.8). The lysates were heated to 100°C for 10 min and clarified by centrifugation; equal amounts of protein were separated by SDS-PAGE. Proteins were transferred to poly(vinylidene difluoride) membranes and immunoblotted with antibodies to FABP4(422/aP2), C/EBPβ, C/EBPα, and PPARγ [antibodies to 422/aP2, C/EBPβ and C/EBPα were provided by Dr. M Daniel Lane (Johns Hopkins University School of Medicine, Baltimore) and the antibody to PPARγ was purchased from Cell Signalling Technology ].
Cell cycle analysis by propidium iodide staining and flow cytometry
Cells were trypsinized, washed with PBS and fixed with 2% (wt/vol) paraformaldehyde in PBS. They were treated with 0.5 mg/ml RNase A for 1 h at room temperature and incubated with 0.1 mg/ml propidium iodide (Sigma) for 45 min at 37°C. DNA content was determined by flow cytometry (Bio-Rad).
BrdU labelling and immunofluorescence microscopy
BrdU labeling of hBMSCs and 3T3-L1 cells (kindly provided by Dr. M Daniel Lane, Johns Hopkins University School of Medicine, Baltimore) was performed following the procedure published by Tang  with modifications. Cells were plated on to cover-slips and maintained in DMEM containing 10% FBS for several days after confluence and induced to differentiate. Regarding the growth kinetics differennce (hBMSCs have a longer G0/G1 phase than 3T3-L1, the entry of hBMSCs into S phase is ~20h at passage 3 ), BrdU for 3T3-L1, BrdU (10 μg/ml) was added at 18 h after induction (during S phase) until 48 h and then shifted to maintain medium (with insulin only); for hBMSCs, BrdU was added at 24 h until 72 h. After differentiation, the cover-slips were fixed in 70% ethanol for 30 min followed by 100% methanol for 10 min at room temperature. The fixed cells were treated for 30 min with 1.5 M HCl, blocked with 0.5% Tween 20 in PBS with 10% FBS for 5 min, incubated with anti-BrdU (1:100, Sigma) or anti-perilipin (1:50, Santa Cruz) primary antibodies in the same buffer overnight, and incubated with FITC/TRITC-conjugated secondary antibodies for 1-2 h. Nuclei were counterstained with 4˜,6-diamidino-2-phenylindole (DAPI). Images were taken on a confocal microscope.
Adenoviral expression vectors and infection
The adenoviral expression vectors pAd/CMV/V5-DEST (Invitrogen) encoding human C/EBPβ, C/EBPα, PPARγ and Lac Z (control) were constructed according to the manufacturer's protocols. shRNAs for C/EBPα, PPARγ and Lac Z were cloned into pBlock-it (Invitrogen). The sequences of the shRNAs were as follows: C/EBPα, CACCAGGAGGATGAAGCCAAGCAGCTCGAAAGCTGCTTGGCTTCATCCTCCT. PPARγ, CACCGGGTGAAACTCTGGGAGATTCCGAAGAATCTCCCAGAGTTTCACCC. Confluent hBMSCs were infected with the adenovirus at MOI (multiplicity of infection) of 10 for 4 h; the expression of human C/EBPβ, C/EBPα, PPARγ was assessed by real-time PCR at 24 h or by immunoblotting with antibodies against human C/EBPβ, C/EBPα, PPARγ and FLAG at 48 h. For adipocyte differentiation, various combinations of inducers were added to the infected cells for three days. Oil red O staining was used to demonstrate fat lipid accumulation on day eight and western blotting was used to demonstrate FABP4 (422/aP2 in mouse) expression on day four.
RNAi of C/EBPβ with siRNA
SiRNA oligonucleotides specific for C/EBPβ mRNA (5'-CCCUGCGGAACUUGUUCAAGCAGCU-3') were synthesized by Invitrogen. The silencing effect was verified by real-time PCR for C/EBPβ expression. HBMSCs in 60 mm dishes at 60-70% confluence were transfected with Negative and C/EBPβ siRNA oligonucleotides by using Lipofectamine RNAiMAX (Invitrogen). After 24 h the cells were trypsynized and plated into 35 mm dishes in order to reach confluence immediately. After a further 24 h they were induced to differentiate by three cycles of treatment, and subjected to oil red O staining at day 14.
Real-time quantitative PCR
Real-time quantitative PCRs were performed with 2× PCR Master Mix (Power SYBR® Green, ABI) on a Bio-Rad Q5 instrument (Bio-Rad). The threshold cycles (Ct) for the target genes and the 18S rRNA control signals were determined in triplicate experiments, and the relative RNA quantity was calculated using the comparative Ct method. Primers were as follows:
18S rRNA: Forward 5'-CGGCTACCACATCCAAGGAA-3', Reverse 5'-GCTGGAATTACCGCGGCT-3'.
C/EBPβ: Forward 5'-GCAAGAGCCGCGACAAG-3', Reverse 5'-GGCTCGGGCAGCTGCTT-3'.
C/EBPα: Forward 5'-AAGAAGTCGGTGGACAAGAACAG-3', Reverse 5'-TGCGCACCGCGATGT-3'.
PPARγ: Forward 5'-GATACACTGTCTGCAAACATATCACAA-3', Reverse 5'-CCACGGAGCTGATCCCAA-3'.
FABP4: Forward 5'-GCTTTGCCACCAGGAAAGTG-3', Reverse 5'-ATGGACGCATTCCACCACCA-3'.
GLUT4: Forward 5'-GCCGGACGTTTGACCAGAT-3', Reverse 5'-TGGGTTTCACCTCCTGCTCTA-3'.
This research is supported by National Key Basic Research Project Grant 2006CB943704, National Natural Science Foundation for Distinguished Scholars Grant 30625015, National Natural Science Foundation Grant 30700121 and 30870510, Program for Outstanding Medical Academic Leader Grant B-LJ06032, Shanghai Key Science and Technology Research Project 08dj1400603, Program for New Century Excellent Talents in University NCET-08-0130 and Shanghai Rising Star Program 08QA14012. The Department is supported by Shanghai Leading Academic Discipline Project, Project Number: B110.
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