The roles of integrin β1 in phenotypic maintenance and dedifferentiation in chondroid cells differentiated from human adipose-derived stem cells
- Simin Luo†1, 2,
- Qiping Shi†1,
- Zhengang Zha1, 2Email author,
- Ping Yao3,
- Hongsheng Lin1, 2,
- Ning Liu1, 2,
- Hao Wu1, 2,
- Jiye Cai4 and
- Shangyun Sun1
© Luo et al.; licensee Springer. 2013
Received: 15 February 2013
Accepted: 10 March 2013
Published: 24 March 2013
The aim of this study is to probe the intrinsic mechanism of chondroid cell dedifferentiation in order to provide a feasible solution for this in cell culture.
Morphological and biomechanical properties of cells undergoing chondrogenic differentiation from human adipose-derived stem cells (ADSCs) were measured at the nanometer scale using atomic force microscopy and laser confocal scanning microscopy. Gene expression was determined by real-time quantitative polymerase chain reaction.
The expression of COL II, SOX9, and Aggrecan mRNA began to increase gradually at the beginning of differentiation and reach a peak similar to that of normal chondrocytes on the 12th day, then dropped to the level of the 6th day at 18th day. Cell topography and mechanics trended resembled those of the genes’ expression. Integrin β1 was expressed in ADSCs and rapidly upregulated during differentiation but downregulated after reaching maturity.
The amount and distribution of integrin β1 may play a critical role in mediating both chondroid cell maturity and dedifferentiation. Integrin β1 is a possible new marker and target for phenotypic maintenance in chondroid cells.
KeywordsIntegrin β1 Adipose-derived stem cells Dedifferentiation Atomic force microscope
Adipose-derived stem cells (ADSCs) are multipotent cells that can differentiate into cells of multiple tissue lineages, such as osteocytes, chondrocytes, adipocytes, or neuronal cells. Recent research has indicated that ADSCs can differentiate into chondrocytes in vitro, but chondroid cells ultimately lose their phenotype, or dedifferentiate, in long-term culture through a poorly understood mechanism[1, 2]. Over the past several years, in order to maintain or reinstate differentiation of chondrocytes, cultures were supplemented with exogenous cytokines, such as PTHrP, exogenous bone morphogenetic protein (BMP)-2, triiodothyronine (T3), fibroblast growth factor 18, and electroporation-mediated transfer of SOX trio genes (SOX-5, SOX-6, and SOX9) to mesenchymal cells. Additional methods to prevent dedifferentiation include changing culture systems to those similar to microcarriers, high-density micromass culture, three-dimensional (3D) cultures in hydrogels, in pellet culture using centrifuge tubes, and 3D dynamic culture using 3D-stirred suspension bioreactor (spinner-flask) culture system.
The cell membrane plays an important role in cell physiology and in regulating processes such as material transport, energy conversion, signal transduction, cell survival, apoptosis, and differentiation[13–15]; so alteration of the cell surface ultrastructure can directly influence cellular function. Despite its importance, there are still many unanswered questions about the role of the cell membrane in differentiation: whether there are changes or defects on cellular membrane later in differentiation, whether these defects during late stage differentiation cause dedifferentiation by disturbing cellular homeostasis, and whether the biophysical properties in plasma membrane could be manipulated to maintain differentiation or redifferentiate the cell.
Atomic force microscopy (AFM) has recently emerged as an implement to image the cell membrane and detect mechanical properties at nanometer scale. We are the first to use AFM to observe the change in morphological and biomechanical properties between chondroid cells and normal chondrocytes, leading to the detection of plasma membrane proteins at the molecular scale. We also used flow cytometry and laser confocal scanning microscopy (LCSM) to analyze integrin β1 expression during chondrogenic differentiation of ADSCs. We used these techniques to probe the intrinsic mechanism of chondroid cell dedifferentiation in order to provide a feasible solution for this in cell culture.
ADSCs isolation, culture, and identification
Subcutaneous adipose tissue was resected from seven patients (mean age, 26 years; range, 12 ~ 32 years) undergoing inguinal herniorrhaphy. Research ethics board approval for this study was obtained from Jinan University. Isolation and identification of ADSCs was performed as described previously with modifications. Cells were cultured in DMEM/F12 (Gibco, Invitrogen, Carlsbad, CA, USA) supplemented with 10% fetal bovine serum (FBS, Gibco, USA) and 1% antibiotic (100 U/ml penicillin and 0.1 mg/ml streptomycin, Sigma-Aldrich Corporation, St. Louis, MO, USA) in an incubator (5% CO2, 37°C). The medium was refreshed every 3 days, and cells were split 1:3 after reaching 90% confluence.
ADSCs (passage 3) were seeded at a high-cell density (2 × 105/10 ml), then the medium was changed to DMEM/F12 supplemented with chondrogenic medium: 1% FBS, 6.25 μg/ml insulin + ITS (Sigma, USA), 10 ng/ml TGF-β1 (Peprotech, Rocky Hill, NJ, USA), 10 to 7 M dexamethasone (Sigma, USA), 50 μg/ml ascorbic acid (Sigma, USA), 100 U/ml penicillin, and 0.1 mg/ml streptomycin as previously described. Twenty-one days after induction, lipid accumulations in adipocytes were visualized by staining with oil red-O as follows: cells were fixed in 10% formalin for 1 h and stained for lipid with 0.3% oil red-O for 15 min. After rinsing three times with double distilled H2O, the red-staining cells in six random areas of 1 mm2 were counted in each well and presented as an average ± standard deviation for 3 to 6 replicate wells.
Chondrocytes isolation and culture
Cartilage was obtained from six patients (mean age, 58 years; range, 40 ~ 78 years) undergoing total hip replacement at the First Affiliated Hospital of Jinan University, with femoral neck fracture. Chondrocytes were isolated and collected according to the procedure proposed by Malicev et al., with slight modifications. Culture medium contains DMEM/F12 supplement with 10% FBS.
Sequences of primers for real-time PCR
Forward primer (5′-3′)
Reverse primer (5′-3′)
Product size (bp)
5 ′ -CTGCCCCAGAAGTGAGTGGAG-3 ′
5 ′ -TGGTGCTGATGACAACGCCC-3 ′
5 ′ -CACCTGCAGAGACCTGAAA-3 ′
5 ′ -CAAGTCTCGCCAGTCTCCAT-3 ′
5 ′ -AACGCCATCTTCAAGGCG-3 ′
5 ′ -CTCTCGCTTCAGGTCAGCCTT-3 ′
5 ′ -CCTGGATGCCATCAAAGTCT-3 ′
5 ′ -ACTGCAACTGGAATCCATCG-3 ′
5 ′ -CCACCATGGAGAAGGCTG-3 ′
5 ′ -GGTGCTAAGCAGTTGGTCCT-3 ′
RNA isolation and real-time-polymerase chain reaction analysis
Total RNA was extracted using Trizol (Invitrogen, USA) protocol. Two micrograms of total RNA was used for reverse transcription reaction with the RevertAid First Strand cDNA synthesis kit (Fermentas, Thermo Fisher Scientific Waltham, MA, USA) and random oligo(dT) primer (Fermentas), according to the manufacturer’s instructions. The cell was collected at different time points after differentiation (0, 3, 6, 9, 12, 15, 18, and 21 days), and expression of Aggrecan, COLII, SOX9, COLI, and GAPDH genes in the regenerated fragments was measured by real-time polymerase chain reaction (RT-PCR). Samples were set up in duplicate with the Power SYBR® Green and analyzed with the ABI 7500 Real-Time PCR System (Applied Biosystems, Life Technologies Corp., Carlsbad, CA, USA). RT-PCR was performed using PCR Taq core kit (Takara Bio Inc., Dalian, China).
Single cell atomic force microscopy measurement
The cells were fixed with 2.5% glutaraldehyde for 15 min, then washed three times with distilled water. Morphology and mechanical response of cells were obtained by AFM (Autoprobe CP Research, Veeco, Plainview, NY, USA) imaging under contact mode. All data were analyzed with the instrument-equipped software IP2.1. silicon nitride tips (UL20B, Park Scientific Instruments, Suwon, South Korea) were used in all AFM measurements. In each group, single-cell imaging was repeated for six cells, and each cell was scanned three times. The nominal tip curvature radius was less than 10 nm; a spring constant of silicon cantilevers was 0.01 N/m; a resonance frequency was 285 kHz; the loading force was adjusted to below 1 ~ 2 nN. All parameters were obtained from manufacturer. R a is the average roughness in analytical area, and R q means the root mean square roughness.
After scanning of cellular topographic images, various locations on a cell were selected to obtain the force-distance curves by the force-modulate mode AFM. All force-distance curve experiments were performed at the same loading rate. Twenty force-distance curves were acquired from each cell; five different cells should be detected in each group.
Laser confocal scanning microscopy and observation
ADS, 12DD, 21DD, and normal chondrocytes (NC) were washed with phosphate buffered solution (PBS) three times, fixed in 4% paraformaldehyde for 15 min at room temperature, then washed with PBS again and blocked with unimmunized goat serum for 10 min at 37°C before incubating with primary antibodies (rabbit anti-human integrin β1) for 20 min. After washing with PBS, the cells were incubated with rhodamine-conjugated rat anti-rabbit (1:100) secondary antibody (Biotium Inc., Hayward, CA, USA) at 37°C for 1 h to label integrin β1. Then the cells were identified by counterstaining with 4′,6-diamidino-2-phenylindole (DAPI) for 10 min in the dark. After washing with PBS, the labeled cells were observed using a laser confocal scanning microscopy (LCM 510 Meta Duo Scan, Carl Zeiss, Oberkochen, Germany).
ADS, 12DD, 21DD, and NC were prepared for integrin β1 marker. A number of 1 × 106 cells were incubated with PE-conjugated integrin β1 antibodies at 37°C for 1 h in the dark. Then the cells were centrifuged and washed in PBS three times. Finally, cells were acquired by use of a FACScan (Becton Dickinson, Franklin Lakes, NJ, USA) flow cytometer running its accompanying CellQuest software.
All data were mean values ± standard deviation (SD). Statistical analysis was performed using one-way analysis of variance test (SPSS17.0), with P < 0.05 regarded as statistical significance.
Detection of SOX9, COL II, COL I, and Aggrecan genes by real-time RT-PCR
Atomic force microscopy analysis
Further scanning for local within small scale was conducted (scanning area 5 × 5 μm2). Membrane surface particles were clustered in ADS (Figure2, A3 and A4), and the particle sizes were generally between 50 and 250 nm (Figure2, A5). Surface particles of 3DD and 6DD were between 100 and 400 nm (Figure2, B5 and C5) and clustered, but they were sparse and distributed randomly (Figure2, B3, B4, C3, and C4). In contrast, the surface of 9DD was flat and uniform. Particle numbers were reduced, but the size range was narrower, between 250 and 300 nm (Figure2, D3, D4, and D5). Some shallow and uniform cavities were observed on 12DD (Figure2, E3 and E4), and the particles were between 200 and 300 nm. NC had a similar porous arrangement, but cavities were deeper and more irregular with larger particle size, between 300 and 400 nm (Figure2, I3 and I4). Porous structure disappeared in 15DD, 18DD, and 21DD. The particle size was reduced and they were distributed in a line in 15DD and 18DD (Figure2, F3, F4, G3, and G4). In 21DD (Figure2, H3, and H4), membrane surface particles returned to a clustered distribution, while the sizes varied from 20 to 450 nm.
Morphological and biomechanical parameters of differentiated cells detected by AFM
Surface average roughness (R a) (nm)
Root mean square roughness (R q) (nm)
Adhesive force (pN)
Young’s modulus (kPa)
46.700 ± 4.495b
72.450 ± 7.246b
182.326 ± 18.229a
1.597 ± 0.110b
71.155 ± 7.096a,b
106.448 ± 12.070a,b
200.254 ± 17.138a
2.059 ± 0.179a,b
72.407 ± 7.621a,b
106.721 ± 13.489a,b
261.688 ± 19.416a,b
2.314 ± 0.207a,b
85.044 ± 7.170a,b
104.311 ± 11.333a,b
301.049 ± 22.776a,b
2.405 ± 0.213a
220.847 ± 21.308a,b
300.940 ± 29.248a,b
410.440 ± 28.638a,b
2.711 ± 0.236a
169.844 ± 16.589a,b
218.186 ± 17.884 a,b
369.682 ± 26.958a,b
2.996 ± 0.233a
154.426 ± 12.985a,b
180.992 ± 18.232a,b
306.807 ± 23.506a,b
3.090 ± 0.234a
116.913 ± 12.361a,b
151.729 ± 13.340a,b
181.895 ± 18.648b
3.518 ± 0.381a,b
303.205 ± 29.475a
362.011 ± 35.296a
639.197 ± 47.678a
2.742 ± 0.200a
Young’s modulus is another valuable way to describe mechanical properties of cell membranes, and the value is calculated as described in the ‘Methods’ section. A larger Young’s modulus indicates that the cell was more difficult to deform, implying lower cell elasticity and greater stiffness. A comparison of the Young’s modulus of the samples is listed in Table 2. The value increased gradually during chondrogenic differentiation of ADSCs. Young’s modulus of 12DD was about twofold higher than ADS, equivalent to NC (P > 0.05). The maximum value of 3.518 ± 0.381 kPa was reached at 21DD.
Laser confocal scanning microscopy and observation
Integrin β1 content flow cytometry
Flow cytometry was used for the quantification of integrin β1 of four groups (ADS, 12DD, 21DD, and NC). Integrin β1 content of NC was the highest, up to 90.53%, followed by 12DD, which is 75.36%, and then 21DD and ADS had only 43.02% and 39.84%, respectively.
The RT-PCR results showed that ADSCs could be differentiated into chondroid cells expressing chondrocyte-specific markers such as COL II, Aggrecan, and SOX9. When differentiated to the 12th day, the expression of COL II, Aggrecan, and SOX9 was close to that in normal chondrocytes, but subsequently fell. Therefore, through our PCR results, we inferred that ADSCs might be differentiated to mature chondroid cells at 12th day after induction, but after that their differentiated state is not maintained. Additionally, expression of the dedifferentiated marker gene COL I increased, behaving in an opposite manner to the differentiation markers. From this, we see that the extension of differentiation time does not improve the differentiation rate and indeed leads to dedifferentiation. Because no clear morphological markers of dedifferentiation are apparent under an inverted microscope, we employed other methods to observe the sequential morphological variation over the course of differentiation at nanometer scale. Because the cell membrane is not only a barrier between the intracellular environment and extracellular world but also a regulator of many important biological processes such as signal transduction, material transportation, and energy exchange, we looked for variation in the cell membrane structure accompanying with the change of cellular function; in this case, the level of differentiation.
AFM is a powerful tool for nanobiological studies, so we first used AFM to compare the ultrastructure of chondroid cells and NC and attempt to explain the relationship between cell dysfunction and its ultrastructure.
We obtained visual data of appearance and size, as well as dynamic changes of R a and R q on the nanometer scale using this method. In our experiment, we observed that ADSCs were irregular, long spindle shape with a round and extruded nucleus, but 12DD and NC were triangular or polygonal with flat and compact nuclei and endochylema. Both R a and R q in 12DD were close to those NC. Though there was no obvious morphological change with 21DD, we still obtained the change of R a data. The R a value of 21DD was reduced distinctly and membrane protein arrangement changed from regular porous arrangement to more of line and clusters. Taking the PCR data, we conclude that dedifferentiation after the 12th day is responsible for the ultrastructure changes. We hope the visual and quantitative data will be helpful in analyzing the differentiation process of ADSCs to mature chondroid cells and revealing a mechanism of cell destabilization in the late stage.
Obtaining of cell biomechanical data was another strength of AFM. Recent studies found that mechanical properties of a cell may be used as phenotypic biomarkers. Therefore, we inferred that the functional change of cells caused by late stage dedifferentiation may also be observed through the cellular mechanics. To test this, we measured adhesion force and Young’s modulus across the whole differentiation process to further support the changes in function and cell surface ultrastructure.
Adhesion force mostly represents the number and distribution of cell surface adhesion molecules. Our force-distance curve shows that during chondrogenic differentiation, adhesion force gradually increases to the maximum at the 12th day, but this value is slightly lower than that of NC, and then the value decreases as differentiation continues. Adhesion force corresponds to the change of R a. Our data demonstrate a trend of adhesion force that is in accordance with R a in the process of chondrogenic differentiation. Quantity and distribution of cell surface proteins directly affects R a data. Surface particle numbers increased, causing the cell membrane to be uneven and rough thereby increasing R a. The higher adhesion force and R a value of 12th day are due to the increase of biomacromolecule particles on the mature chondroid cells, which interact more with the AFM needle. Likewise, as differentiation continued, there were fewer cell surface adhesion proteins, and the adhesion force and R a decreased. Thus, the dedifferentiation of chondroid cells was relative to the decrease of cell surface proteins.
Expression of adequate adhesion proteins is important for cells to attach in cartilage lacuna, which is necessary for stable synthesis and secretion of extracellular matrix (ECM) proteins. It is crucial for chondrocytes to remain differentiated to function properly. We chose integrin β1 as a representative adhesion protein for this experiment because it is widely expressed and is the main adhesion molecule in chondrocytes[26, 27]. Then, we detected the distribution of integrin β1 through LCSM. We found integrin β1 on the cell membrane and the dynamic tracing of integrin β1 revealed a maximum fluorescence intensity of integrin β1 on the 12th day. In parallel, we used flow cytometry to test the quantity of integrin β1, and this supported the maximum at day 12, although the quantity did not reach that of NC. The qualitative and quantitative changes of integrin β1 in these groups correspond to R a and adhesion force results, so we conclude that dedifferentiation of chondroid cells may be directly related to loss or involution of integrin β1.
Acting as a bridge between ECM and the cytoskeleton, integrin not only transmits signals between the cell and the ECM but also regulates cytoskeletal arrangement and therefore cell rigidity[28, 29]. We then wanted to test if the change of integrin β1 is accompanied with the change of cell rigidity, and we did so using AFM to measure cell Young’s modulus of each differentiation stage. We found that Young’s modulus increased gradually throughout the differentiation process. It came to the maximum at 21DD and was higher than NC in 15DD, 18DD, and 21DD. Young’s modulus of 12DD was similar to that of NC, having no statistically significant difference. Our data imply that 12DD had the most ideal stiffness and elasticity for chondrocytes.
The stiffness of cells is related to their physiological roles, and cartilage cells in particular require stiffness to bear and transmit a stress load. Reduction in elasticity would prevent the cartilage from buffering the vibrations from stress loads. We observed that the stiffness of chondroid cells increased continuously in the late stage differentiation, reducing cell deformability and perhaps causing cell degeneration. This is an important consideration in tissue engineering of cartilage as opposed to normal cartilage, because the continual increase in stiffness could negate the therapeutic effect of regenerative cartilage tissue. We speculate the improper rigidity of 21DD chondroid cells might be an objective manifestation and the intrinsic factor of degeneration.
In general, the process of differentiating ADSCs into chondroid cells involves the synthetic process of integrin β1. We considered that chondroid cells mature when integrin β1 reaches its peak value. Degeneration and structural changes of integrin β1 distribution lead to dedifferentiation of chondroid cells. Therefore, integrin β1 may be responsible for the maturation and degeneration of chondrogenic differentiation of ADSCs.
This work was supported by Guangdong Provincial Science and Technology Project of China (2011B031800066 and 2010B031600105), Guangdong Provincial Medical Scientific Research Foundation (B2011161), the Fundamental Research Funds for the Central Universities, the Science and Technology Development Fund of Macau (025/2010/A), and Natural Science Foundation of Guangdong Province (10151063201000052).
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