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Effect of cell density on adipogenic differentiation of mesenchymal stem cells
Biochemical and Biophysical Research Communications 381 (2009) 322–327
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Biochemical and Biophysical Research Communications journal homepage: www.elsevier.com/locate/ybbrc
Effect of cell density on adipogenic differentiation of mesenchymal stem cells Hongxu Lu a,b,1, Likun Guo b,c,1, Michal J. Wozniak b, Naoki Kawazoe b,d, Tetsuya Tateishi b, Xingdong Zhang c, Guoping Chen a,b,d,* a
Graduate School of Pure and Applied Sciences, University of Tsukuba, 1-1-1 Tennodai, Tsukuba, Ibaraki 305-8577, Japan Biomaterials Center, National Institute for Materials Science, 1-1 Namiki, Tsukuba, Ibaraki 305-0044, Japan c National Engineering Research Center for Biomaterials, Sichuan University, 29 Wangjiang Road, Chengdu 610064, China d International Center for Materials Nanoarchitectonics (MANA), National Institute for Materials Science, 1-1 Namiki, Tsukuba, Ibaraki 305-0044, Japan b
a r t i c l e
i n f o
Article history: Received 23 January 2009 Available online 4 February 2009
Keywords: Mesenchymal stem cells Adipogenic differentiation Cell density Micropattern Tissue engineering
a b s t r a c t The effect of cell density on the adipogenic differentiation of human bone marrow-derived mesenchymal stem cells (MSCs) was investigated by using a patterning technique to induce the formation of a cell density gradient on a micropatterned surface. The adipogenic differentiation of MSCs at a density gradient from 5 103 to 3 104 cells/cm2 was examined. Lipid vacuoles were observed at all cell densities after 1–3 weeks of culture in adipogenic differentiation medium although the lipid vacuoles were scarce at the low cell density and abundant at the high cell density. Real-time RT-PCR analysis showed that adipogenesis marker genes encoding peroxisome proliferator-activated receptor c2 (PPARc2), lipoprotein lipase (LPL), and fatty acid binding protein-4 (FABP4) were detected in the MSCs cultured at all cell densities. The results suggest that there was no apparent effect of cell density on the adipogenic differentiation of human MSCs. Ó 2009 Elsevier Inc. All rights reserved.
Mesenchymal stem cells (MSCs) are a prospective source of cells for tissue engineering because they are relatively easy to obtain from a small aspirate of bone marrow and are multipotent, able to differentiate into different cell lineages such as osteoblasts, chondrocytes, adipose cells, and neural cells [1–3]. MSCs have been used for tissue engineering of various tissues such as cartilage [4], bone [5], muscle [6], tendon [7], ligament [8], and fat [9,10]. Manipulation of stem cell differentiation remains a great challenge in tissue engineering. Many factors affect the differentiation of mesenchymal stem cells. These include soluble growth factors and cytokines [11,12], mechanical stimuli [13], surface properties [14], and culture conditions [15]. Cell density has also been reported [16,17] as affecting cell functions such as proliferation and differentiation. McBeath et al. [18] has reported that human mesenchymal stem cells (MSCs) plated at low density have a high potential to become osteoblasts, whereas cells plated at high density have a propensity to become adipocytes. The effect of cell seeding density on differentiation remains controversial. Both et al. [19] and Colter et al. [20] have reported the effect of increased expansion of MSCs cultured at lower densities, respectively. However, the results of cell differentiation from the two groups are dif-
* Corresponding author. Address: Biomaterials Center, National Institute for Materials Science, 1-1 Namiki, Tsukuba, Ibaraki 305-0044, Japan. Fax: +81 29 860 4714. E-mail address: [email protected] (G. Chen). 1 These two authors are contributed equally to this work. 0006-291X/$ - see front matter Ó 2009 Elsevier Inc. All rights reserved. doi:10.1016/j.bbrc.2009.01.174
ferent. Both et al. indicated that the osteogenic differentiation potential of MSCs both in vitro and in vivo seems to decrease with repeated passages while Colter et al. reported that MSCs retained their multipotentiality for differentiation during expansion. For these studies, the effects of cell density on cell functions were compared by separately culturing cells at different cell densities. It is difficult to completely avoid the influence of other factors induced during separate cell culture from the results. It is easier to compare the effect of cell density on cell differentiation more directly if cells at various cell densities can be cultured simultaneously on a single surface. In this study, we used a patterning technique to prepare a micropatterned surface that allowed the formation of a cell density gradient on a single surface. A cell density gradient of human MSCs was formed when MSCs were cultured on a micropatterned surface, enabling direct comparison of the effect of cell density on adipogenic differentiation of MSCs on a single surface. Materials and methods Preparation of micropatterned surface. Azidophenyl-derivatized poly(vinyl alcohol) conjugate (AzPhPVA) was synthesized by coupling poly(vinyl alcohol) with 4-azidobenzoic acid as previously described [21]. DMSO solution containing 234.09 mg dicyclohexylcarbodiimide (2 mL) was added dropwise to 5 mL DMSO solution containing 185.37 mg 4-azidobenzoic acid under stirring at room temperature in the dark. Then, 2 mL of DMSO solution dissolving 16.84 mg 4-(1-pyrrolidinyl) pyridine was added dropwise to the
H. Lu et al. / Biochemical and Biophysical Research Communications 381 (2009) 322–327
reaction mixture under stirring. After 10 min, 8 mL of DMSO solution containing 100 mg of poly(vinyl alcohol) was added dropwise to the above reaction mixture under stirring in the dark, and the reaction was allowed to proceed overnight. After 24 h, dicyclohexylurea, which formed during the reaction, was filtered off. The filtrate was collected and purified by dialysis against Milli-Q water. After purification, the product was freeze-dried and stored at room temperature in the dark. The number of azidophenyl groups in the AzPhPVA was determined by 1H NMR from the peak intensities of the azidophenyl protons at around 7 ppm, and those of the methylene and methylidyne protons of the polymer main chain at 1.5 and 3.9 ppm, respectively. A polystyrene plate (2 2 cm) was cut from a cell culture polystyrene flask. The AzPhPVA was dissolved in water (200 lg/mL). The solution (100 lL) was placed on the polystyrene plate and air-dried at room temperature in the dark. The plate was covered with a patterned photomask and irradiated with ultraviolet light at an intensity of 5.0 103 lJ/cm2 from a distance of 15 cm for 25 s. After irradiation, the plate was immersed in Milli-Q water and then sonicated to completely remove any unreacted polymer from the unirradiated areas. After complete washing, PVA-micropatterned surfaces were obtained. Observation by scanning probe microscopy (SPM). An SPA-400 multi-function SPM unit (SII NanoTechnology Inc., Tokyo, Japan) connected to an SPI4000 controller was used for measurements in air and liquid in contact mode. The cantilevers used were OTR8 silicon nitride probes for measurement in air and OTR4 for measurement in liquid (Veeco Instruments, Santa Barbara, CA) having spring constants of 0.15 N/m and 0.02 N/m, respectively. All SPM measurements were made at room temperature. The polystyrene plates with the PVA-micropattern grafted surfaces were observed in a dry state in air and in a wet state in Milli-Q water. A 100 100 lm area of the sample was observed by SPM. The heights of the micropatterned PVA were measured from the SPM images. Nine spots from the topographic images of each kind of grafted pattern were used to measure the mean heights and standard deviations. The data were expressed as the average ± standard deviation of the nine spots. Cell culture. Human bone marrow-derived MSCs (MSCs) were obtained from Osiris (Worthington Biochemical, Lakewood, NJ) at passage 2. The cells were seeded in T-75 culture flasks (Iwaki Glass, Tokyo, Japan) using proliferation medium purchased from Lonza (Walkersville, MD). The proliferation medium contained 440 mL MSC basal medium, 50 mL mesenchymal cell growth supplement, 10 mL 200 mM L-glutamine, and 0.5 mL penicillin/streptomycin mixture. After reaching confluence, the cells were further subcultured once and used at passage 4. The cells were collected by treatment with trypsin/EDTA solution and suspended in serum (control) medium at a density of 2.50 104 cells/mL. The serum medium was composed of Dulbecco’s modified Eagle’s medium (Sigma–Aldrich, St. Louis, MO) supplemented with 4500 mg/L glucose, 584 mg/L glutamine, 100 U/mL penicillin, 100 lg/mL streptomycin, 0.1 mM nonessential amino acid, 0.4 mM proline, 50 mg/L ascorbic acid, and 10% fetal bovine serum (FBS). The micropatterned polystyrene plates were put in a cell culture dish and a glass cylinder (10 mm in both bore diameter and height) was placed over each PVA-micropatterned polystyrene plate. Cell suspension solution (0.157 mL/well) was added into the glass cylinder (initial cell density: 5.00 103 cells/cm2). The MSCs were cultured in the serum medium for 3 days. To induce adipogenic differentiation, the MSCs were cultured in adipogenic differentiation medium [22] consisting of DMEM serum medium supplemented with 1 lM dexamethasone and 0.5 mM methyl-isobutylxanthine, insulin (10 lg/mL), and 100 lM indomethacin. The cells on the micropatterned surfaces were incubated in the serum medium or the serum control medium for 1, 2, and 3 weeks and used for cell stain-
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ing. Alternatively, for analyzing gene expression, the MSCs were seeded into 6-well cell culture plates at respective densities of 5.00 103, 1.00 104, 2.00 104, and 3.00 104 cells/cm2 and cultured in the adipogenic differentiation medium or the serum control medium for 1, 2, and 3 weeks. Oil Red O staining. The cultured cells were rinsed with phosphate buffer saline (PBS) three times, fixed with 4% paraformaldehyde for 1 h at 4 °C, and then stained with fresh Oil Red O solution for 2 h. The Oil Red O solution was prepared by mixing three parts stock solution (0.5% in isopropanol; Sigma) with two parts water and filtering through a 0.2-lm filter. The stained cells were washed three times with PBS to remove any background Oil Red O stain. Photomicrographs were captured by a optical microscope with a DP-70 CCD camera (Olympus, Tokyo, Japan). RNA isolation and quantitative real-time RT-PCR analysis. After culture for 1 and 3 weeks in the 6-well cell-culture plates, the cells were washed with PBS three times, and 1 mL of Isogen reagent (Nippon Gene, Toyama, Japan) was added to each well. Total RNA was extracted following the manufacturer’s protocol. Total RNA (1 lg) was reversely transcribed into cDNA using a random hexamer primer (Applied Biosystems, Foster City, CA) in a 20 ll reaction. An aliquot (1 ll) of 10-times diluted reaction solution was used for each 25 ll real-time PCR reaction together with 300 nM forward and reverse primers, 150 nM probes, and qPCR MasterMix (Eurogentec, Seraing, Belgium). TaqmanÒ probes and primer pairs for fatty acid binding protein-4 (FABP4, assay identification number Hs00609791_m1) [23], lipoprotein lipase (LPL, assay identification number Hs00173425_m1) [23], and peroxisome proliferatoractivated receptor c2 (PPARc2, assay identification number Hs01115510_m1), which are assay-on-demand gene expression products, were obtained from Applied Biosystems. Real-time quantitative RT-PCR analysis was performed using a housekeeping gene GAPDH and 18S ribosomal RNA, as previously described. After an initial incubation step of 2 min at 50 °C and denaturation for 10 min at 95 °C, 40 cycles of PCR (95 °C for 15 s, 60 °C for 1 min) were performed. Reactions were performed in triplicate. 18S ribosomal RNA levels were used as endogenous controls and gene expression levels relative to GAPDH were calculated using the comparative Ct method. To calculate the means and standard deviations, three samples under each condition were measured. A oneway analysis of variance (ANOVA) with Tukey’s post hoc test for multiple comparisons was used for statistical analysis. A value of p < 0.05 was considered statistically significant.
Results and discussion PVA-micropatterned surface Poly(vinyl alcohol) was micropatterned on polystyrene plates cut from cell culture flasks. First, photoreactive AzPhPVA was synthesized by coupling poly(vinyl alcohol) with 4-azidobenzoic acid. The introduction of photoreactive azido groups in AzPhPVA was confirmed by the appearance of peaks assigned to the phenylazido proton around 7 ppm in the 1H NMR spectrum. The percentage of the hydroxyl groups in the PVA coupled with the azidophenyl groups was 2.1%. Subsequently, the photoreactive AzPhPVA was micropatterned on polystyrene plates by photolithography (Fig. 1A). An aqueous solution of AzPhPVA was eluted on a polystyrene plate and air-dried in the dark. The cast plate was covered with a photomask and photoirradiated. The photomask pattern was composed of alternate UV-transparent and UV-nontransparent stripes (Fig. 1B). The UV-nontransparent stripes were 200 lm wide and the UV-transparent stripes had gradient widths from 20 to 1000 lm. AzPhPVA in the irradiated areas should be intermolecularly and intramolecularly crosslinked and grafted to the poly-
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Fig. 1. The poly(vinyl alcohol) micropattern and cell distribution. Preparation scheme of poly(vinyl alcohol)-micropatterned surface (A). Photomicrograph of the photomask (B). The white stripes were UV-transparent and the dark stripes UV-nontransparent. The photomask had a pattern of UV-transparent stripes with widths varying from 20 to 1000 lm separated by 200 lm UV-nontransparent stripes. Three-dimensional topographic images of scanning probe microscopy (SPM) of poly(vinyl alcohol)-micropatterned surface in air (C) and in Milli-Q water (D). Photomicrographs of MSCs cultured on poly(vinyl alcohol)-micropatterned surface in serum medium after 10 min (E) and 1 day (F). The number indicates the theoretical cell density based on the ratio of the cell adhesive polystyrene area to the nonadhesive PVA area.
styrene surface. In the other areas, AzPhPVA should not be crosslinked and could be removed by washing with Milli-Q water. After washing, a pattern of PVA stripes having gradient widths was prepared. Observation with an optical microscope demonstrated that the PVA formed the same striped pattern as that of the photomask. The PVA-micropatterned surfaces in the dry and wet states were observed by scanning probe microscopy (Fig. 1C and D). The tiered structures of the micropatterned PVA in air and MilliQ water were confirmed by the three-dimensional images. The step height of the micropatterned PVA was 95.75 ± 9.74 nm in air and 143.00 ± 26.50 nm in water. The micropatterned PVA surfaces were smoother and higher in water than they were in air, indicating that the micropatterned PVA swelled when immersed in water. The micropatterned surfaces were used for cell culture in an aqueous medium. Therefore, the SPM images of the micropatterned surface in water reflected the true state of the micropatterned surfaces during cell culture. Adipogenic differentiation of MSCs on the micropatterned surface Mesenchymal stem cells (MSCs) were seeded on the PVA-micropatterned surface at a cell density of 5.00 103 cells/cm2 and cultured in serum medium for 3 days. Immediately after cell seeding, the cells were distributed evenly on the PVA-micropatterned surfaces (Fig. 1E). During cell culture, the cells falling on the PVA stripes moved to the polystyrene stripes because PVA did not support cell adhesion. PVA has been reported to inhibit protein
adsorption and thus inhibit cell adhesion. The MSCs adhered to the polystyrene stripes. After 1 day of culture, the cells were observed only on the polystyrene stripes and formed a striped pattern (Fig. 1F). According to the surface area ratio of the polystyrene stripes and PVA stripes, the cell density from the right to left polystyrene stripes should theoretically be 1.10, 1.10, 1.30, 1.50, 1.75, 2.00, 2.50, 3.00, 3.50, 4.00, 4.50, 5.00, 5.50, 6.00, and 6.00 times that of 5.00 103 cells/cm2 assuming that the cells moved randomly from the nonadhesive PVA stripes to the polystyrene stripes. The cell number on the polystyrene stripes was counted from the three areas of each stripe and cell density was calculated. The cell density on each stripe from right to left was 4.84 ± 0.37, 3.91 ± 0.26, 6.71 ± 0.65, 6.95 ± 2.05, 7.58 ± 2.48, 8.82 ± 0.78 103 and 1.17 ± 0.45, 1.25 ± 0.36, 1.49 ± 0.50, 1.92 ± 0.54, 2.16 ± 0.36, 2.40 ± 0.34, 2.63 ± 0.30, 2.79 ± 0.33, 2.98 ± 0.27 104 cells/cm2, respectively. The result indicates that cell density could be adjusted by controlling the ratio of the cell adhesion and nonadhesion areas and cell density gradient was formed on the PVA-micropatterned surface. After the MSCs were cultured on the PVA-micropatterned surfaces in the serum medium for 3 days, the culture medium was changed to an adipogenic differentiation medium and the cells were further cultured for 1, 2, and 3 weeks. Some of the cells were further cultured in serum medium for 1, 2, and 3 weeks as a control. Lipid vacuoles were observed after culture for 1 week (Fig. 2A–C). The number of lipid vacuoles was low at the low density cells and abundant at the high density cells. The number of li-
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Fig. 2. Photomicrographs of MSCs cultured on poly(vinyl alcohol)-micropatterned surface for 1 (A, D), 2 (B, E), and 3 (C, F) weeks in adipogenic differentiation medium without staining (A–C) and stained with Oil Red O (D–F). Scale bar = 500 lm.
pid vacuoles increased after 2 and 3 weeks of culture. The cells at low density (right stripes of Fig. 2) did not become confluent. Some cells existed alone. The cells at high density (left stripes of Fig. 2) became confluent and came into contact with each other. Culture in adipogenic differentiation medium arrested cell proliferation. The lipid vacuoles were further stained with Oil Red O (Fig. 2D– F). Oil Red O positive cells were observed after 1 week of culture in the adipogenic differentiation medium. The cells at high cell density were more densely stained by Oil Red O than were the cells at low cell density. Oil Red O staining also increased with culture
time. No lipid vacuoles were detected on the MSCs cultured in the control medium (Fig. 3). The cells proliferated and became confluent when cultured in the serum control medium. Although the density of the lipid vacuoles and Oil Red O staining increased with the increase of cell density, the MSCs at all cell densities differentiated into adipocyte-like cells. Cell density did not affect adipogenic differentiation of MSCs in the cell density range used in the present study. To check the effect of cell density on the expression of genes relative to adipogenic differentiation, the MSCs were cultured in 6-
Fig. 3. Photomicrographs of Oil Red O staining of MSCs cultured on poly(vinyl alcohol)-micropatterned surface in serum control medium for 1 (A), 2 (B), and 3 (C) weeks. Scale bar = 500 lm.
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well cell culture polystyrene plates. The seeded cell densities were changed from 1.00, 2.00, 4.00, and 6.00 times that 5.00 103 cells/ cm2. The cells were cultured in adipogenic differentiation and serum control mediums for 1 and 3 weeks. The adipogenesis marker genes encoding peroxisome proliferator-activated receptor c2 (PPARc2), lipoprotein lipase (LPL), and fatty acid binding protein4 (FABP4) were analyzed (Fig. 4). The MSCs cultured in adipogenic differentiation medium expressed the genes while the MSCs cultured in control medium did not. All the genes were detected in the MSCs cultured at different cell densities. The expression level of the genes increased with culture time. There was no significant
difference in the expression level of these genes among the four cell densities. The cell density did not show any evidence of an effect on the expression of the genes. It has been reported that a high density of MSCs facilitates adipogenic differentiation [18]. Conventionally, before adipogenic differentiation induction, cells are cultured to confluence because the adipogenic induction medium inhibits cell proliferation [22,24]. On the other hand, it has also been reported [25] that single-cell derived colonies generated by plating early passage human MSCs at extremely low densities show adipogenic differentiation. The passage cell density [20,26] has been reported to affect MSC proliferation and differentiation potential. Lower cell density results in a higher proliferation rate during cell passage, but the multilineage differentiation potential of the passaged cells is not affected by the passage cell density. In the present study, MSCs of different densities were cultured on a single surface. The MSCs of different densities showed similar adipogenic differentiation. No apparent effect of cell density on adipogenic differentiation was observed. The cell density range in the present study was from 5.00 103 to 3.00 104 cells/cm2. A wider range is possible if the ratio of polystyrene and PVA surface areas is adjusted by changing the photomask. The patterning technique will provide a powerful tool for creating cell arrays in a density gradient to directly compare the effect of cell density on cell functions. Acknowledgments This work was supported in part by the World Premier International Research Center (WPI) Initiative on Materials Nanoarchitectonics, MEXT, Japan, and in part by the New Energy and Industrial Technology Development Organization of Japan. References
Fig. 4. Real-time PCR results of mRNA expression of PPARc2 (A), LPL (B), and FABP4 (C) genes of MSCs cultured at various cell densities in 6-well cell culture polystyrene plates with adipogenic differentiation and serum control medium for 1 and 3 weeks. Cell density is indicated under the horizontal axis. MSCs indicate cells used for cell seeding. Control means the cells were cultured in serum medium. The data are normalized to GAPDH and represent means ± SD (n = 3).
[1] R.O. Oreffo, C. Cooper, C. Mason, M. Clements, Mesenchymal stem cells: lineage, plasticity, and skeletal therapeutic potential, Stem Cell Rev. 1 (2005) 169–178. [2] C. Lange, J. Schroeder, M.V. Lioznov, A.R. Zander, High-potential human mesenchymal stem cell, Stem Cells Dev. 14 (2005) 70–80. [3] M.F. Pittenger, Mesenchymal stem cells from adult bone marrow, Methods Mol. Biol. 449 (2008) 27–44. [4] G. Chen, D. Liu, M. Tadokoro, R. Hirochika, H. Ohgushi, J. Tanaka, T. Tateishi, Chondrogenic differentiation of human mesenchymal stem cells cultured in a cobweb-like biodegradable scaffold, Biochem. Biophys. Res. Commun. 322 (2004) 50–55. [5] L. Meinel, V. Karageorgiou, R. Fajardo, B. Snyder, V. Shinde-Patil, L. Zichner, D. Kaplan, R. Langer, G. Vunjak-Novakovic, Bone tissue engineering using human mesenchymal stem cells: effects of scaffold material and medium flow, Ann. Biomed. Eng. 32 (2004) 112–122. [6] K. Natsu, M. Ochi, Y. Mochizuki, H. Hachisuka, S. Yanada, Y. Yasunaga, Allogeneic bone marrow-derived mesenchymal stromal cells promote the regeneration of injured skeletal muscle without differentiation into myofibers, Tissue Eng. 10 (2004) 1093–1112. [7] H.A. Awad, D.L. Butler, M.T. Harris, R.E. Ibrahim, Y. Wu, R.G. Young, S. Kadiyala, G.P. Boivin, In vitro characterization of mesenchymal stem cell-seeded collagen scaffolds for tendon repair: effects of initial seeding density on contraction kinetics, J. Biomed. Mater. Res. 51 (2000) 233–240. [8] H. Fan, H. Liu, S.L. Toh, J.C. Goh, Enhanced differentiation of mesenchymal stem cells co-cultured with ligament fibroblasts on gelatin/silk fibroin hybrid scaffold, Biomaterials 29 (2008) 1017–1027. [9] M. Neubauer, M. Hacker, P. Bauer-Kreisel, B. Weiser, C. Fischbach, M.B. Schulz, A. Goepferich, T. Blunk, Adipose tissue engineering based on mesenchymal stem cells and basic fibroblast growth factor in vitro, Tissue Eng. 11 (2005) 1840–1851. [10] Y.S. Choi, S. Park, H. Suh, Adipose tissue engineering using mesenchymal stem cells attached to injectable PLGA spheres, Biomaterials 26 (2005) 5855–5863. [11] T.C. Otto, M.D. Lane, Adipose development: from stem cell to adipocyte, Crit. Rev. Biochem. Mol. Biol. 40 (2005) 229–242. [12] R.R. Bowers, M.D. Lane, A role for bone morphogenetic protein-4 in adipocyte development, Cell Cycle 6 (2007) 385–389. [13] I.C. Lee, J.H. Wang, Y.T. Lee, T.H. Young, The differentiation of mesenchymal stem cells by mechanical stress or/and co-culture system, Biochem. Biophys. Res. Commun. 352 (2007) 147–152. [14] L. Guo, N. Kawazoe, Y. Fan, Y. Ito, J. Tanaka, T. Tateishi, X. Zhang, G. Chen, Chondrogenic differentiation of human mesenchymal stem cells on photoreactive polymer-modified surfaces, Biomaterials 29 (2008) 23–32.
H. Lu et al. / Biochemical and Biophysical Research Communications 381 (2009) 322–327 [15] S.J. Song, O. Jeon, H.S. Yang, D.K. Han, B.S. Kim, Effects of culture conditions on osteogenic differentiation in human mesenchymal stem cells, J. Microbiol. Biotechnol. 17 (2007) 1113–1119. [16] B. Neuhuber, S.A. Swanger, L. Howard, A. Mackay, I. Fischer, Effects of plating density and culture time on bone marrow stromal cell characteristics, Exp. Hematol. 36 (2008) 1176–1185. [17] M. Bitar, R.A. Brown, V. Salih, A.G. Kidane, J.C. Knowles, S.N. Nazhat, Effect of cell density on osteoblastic differentiation and matrix degradation of biomimetic dense collagen scaffolds, Biomacromolecules 9 (2008) 129–135. [18] R. McBeath, D.M. Pirone, C.M. Nelson, K. Bhadriraju, C.S. Chen, Cell shape cytoskeletal tension, and RhoA regulate stem cell lineage commitment, Dev. Cell 6 (2004) 483–495. [19] S.K. Both, A.J. van der Muijsenberg, C.A. van Blitterswijk, J. de Boer, J.D. de Bruijn, A rapid and efficient method for expansion of human mesenchymal stem cells, Tissue Eng. 13 (2007) 3–9. [20] D.C. Colter, R. Class, C.M. DiGirolamo, D.J. Prockop, Rapid expansion of recycling stem cells in cultures of plastic-adherent cells from human bone marrow, Proc. Natl. Acad. Sci. USA 97 (2000) 3213–3218.
327
[21] N. Kawazoe, L. Guo, M.J. Wozniak, Y. Imaizumi, T. Tateishi, X. Zhang, G. Chen, Adipogenic differentiation of mesenchymal stem cells on micropatterned polyelectrolyte surfaces, J. Nanosci. Nanotechnol. 9 (2009) 230–239. [22] J.R. Mauney, V. Volloch, D.L. Kaplan, Matrix-mediated retention of adipogenic differentiation potential by human adult bone marrow-derived mesenchymal stem cells during ex vivo expansion, Biomaterials 26 (2005) 6167–6175. [23] M.F. Pittenger, A.M. Mackay, S.C. Beck, R.K. Jaiswal, R. Douglas, J.D. Mosca, M.A. Moorman, D.W. Simonetti, S. Craig, D.R. Marshak, Multilineage potential of adult human mesenchymal stem cells, Science 284 (1999) 143–147. [24] K.D. Murphy, J.O. Lee, D.N. Herndon, Current pharmacotherapy for the treatment of severe burns, Expert Opin. Pharmacother. 4 (2003) 369–384. [25] I. Sekiya, B.L. Larson, J.T. Vuoristo, J.G. Cui, D.J. Prockop, Adipogenic differentiation of human adult stem cells from bone marrow stroma (MSCs), J. Bone Miner. Res. 19 (2004) 256–264. [26] P.A. Sotiropoulou, S.A. Perez, M. Salagianni, C.N. Baxevanis, M. Papamichail, Characterization of the optimal culture conditions for clinical scale production of human mesenchymal stem cells, Stem cells 24 (2006) 462–471.
Contents lists available at ScienceDirect
Biochemical and Biophysical Research Communications journal homepage: www.elsevier.com/locate/ybbrc
Effect of cell density on adipogenic differentiation of mesenchymal stem cells Hongxu Lu a,b,1, Likun Guo b,c,1, Michal J. Wozniak b, Naoki Kawazoe b,d, Tetsuya Tateishi b, Xingdong Zhang c, Guoping Chen a,b,d,* a
Graduate School of Pure and Applied Sciences, University of Tsukuba, 1-1-1 Tennodai, Tsukuba, Ibaraki 305-8577, Japan Biomaterials Center, National Institute for Materials Science, 1-1 Namiki, Tsukuba, Ibaraki 305-0044, Japan c National Engineering Research Center for Biomaterials, Sichuan University, 29 Wangjiang Road, Chengdu 610064, China d International Center for Materials Nanoarchitectonics (MANA), National Institute for Materials Science, 1-1 Namiki, Tsukuba, Ibaraki 305-0044, Japan b
a r t i c l e
i n f o
Article history: Received 23 January 2009 Available online 4 February 2009
Keywords: Mesenchymal stem cells Adipogenic differentiation Cell density Micropattern Tissue engineering
a b s t r a c t The effect of cell density on the adipogenic differentiation of human bone marrow-derived mesenchymal stem cells (MSCs) was investigated by using a patterning technique to induce the formation of a cell density gradient on a micropatterned surface. The adipogenic differentiation of MSCs at a density gradient from 5 103 to 3 104 cells/cm2 was examined. Lipid vacuoles were observed at all cell densities after 1–3 weeks of culture in adipogenic differentiation medium although the lipid vacuoles were scarce at the low cell density and abundant at the high cell density. Real-time RT-PCR analysis showed that adipogenesis marker genes encoding peroxisome proliferator-activated receptor c2 (PPARc2), lipoprotein lipase (LPL), and fatty acid binding protein-4 (FABP4) were detected in the MSCs cultured at all cell densities. The results suggest that there was no apparent effect of cell density on the adipogenic differentiation of human MSCs. Ó 2009 Elsevier Inc. All rights reserved.
Mesenchymal stem cells (MSCs) are a prospective source of cells for tissue engineering because they are relatively easy to obtain from a small aspirate of bone marrow and are multipotent, able to differentiate into different cell lineages such as osteoblasts, chondrocytes, adipose cells, and neural cells [1–3]. MSCs have been used for tissue engineering of various tissues such as cartilage [4], bone [5], muscle [6], tendon [7], ligament [8], and fat [9,10]. Manipulation of stem cell differentiation remains a great challenge in tissue engineering. Many factors affect the differentiation of mesenchymal stem cells. These include soluble growth factors and cytokines [11,12], mechanical stimuli [13], surface properties [14], and culture conditions [15]. Cell density has also been reported [16,17] as affecting cell functions such as proliferation and differentiation. McBeath et al. [18] has reported that human mesenchymal stem cells (MSCs) plated at low density have a high potential to become osteoblasts, whereas cells plated at high density have a propensity to become adipocytes. The effect of cell seeding density on differentiation remains controversial. Both et al. [19] and Colter et al. [20] have reported the effect of increased expansion of MSCs cultured at lower densities, respectively. However, the results of cell differentiation from the two groups are dif-
* Corresponding author. Address: Biomaterials Center, National Institute for Materials Science, 1-1 Namiki, Tsukuba, Ibaraki 305-0044, Japan. Fax: +81 29 860 4714. E-mail address: [email protected] (G. Chen). 1 These two authors are contributed equally to this work. 0006-291X/$ - see front matter Ó 2009 Elsevier Inc. All rights reserved. doi:10.1016/j.bbrc.2009.01.174
ferent. Both et al. indicated that the osteogenic differentiation potential of MSCs both in vitro and in vivo seems to decrease with repeated passages while Colter et al. reported that MSCs retained their multipotentiality for differentiation during expansion. For these studies, the effects of cell density on cell functions were compared by separately culturing cells at different cell densities. It is difficult to completely avoid the influence of other factors induced during separate cell culture from the results. It is easier to compare the effect of cell density on cell differentiation more directly if cells at various cell densities can be cultured simultaneously on a single surface. In this study, we used a patterning technique to prepare a micropatterned surface that allowed the formation of a cell density gradient on a single surface. A cell density gradient of human MSCs was formed when MSCs were cultured on a micropatterned surface, enabling direct comparison of the effect of cell density on adipogenic differentiation of MSCs on a single surface. Materials and methods Preparation of micropatterned surface. Azidophenyl-derivatized poly(vinyl alcohol) conjugate (AzPhPVA) was synthesized by coupling poly(vinyl alcohol) with 4-azidobenzoic acid as previously described [21]. DMSO solution containing 234.09 mg dicyclohexylcarbodiimide (2 mL) was added dropwise to 5 mL DMSO solution containing 185.37 mg 4-azidobenzoic acid under stirring at room temperature in the dark. Then, 2 mL of DMSO solution dissolving 16.84 mg 4-(1-pyrrolidinyl) pyridine was added dropwise to the
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reaction mixture under stirring. After 10 min, 8 mL of DMSO solution containing 100 mg of poly(vinyl alcohol) was added dropwise to the above reaction mixture under stirring in the dark, and the reaction was allowed to proceed overnight. After 24 h, dicyclohexylurea, which formed during the reaction, was filtered off. The filtrate was collected and purified by dialysis against Milli-Q water. After purification, the product was freeze-dried and stored at room temperature in the dark. The number of azidophenyl groups in the AzPhPVA was determined by 1H NMR from the peak intensities of the azidophenyl protons at around 7 ppm, and those of the methylene and methylidyne protons of the polymer main chain at 1.5 and 3.9 ppm, respectively. A polystyrene plate (2 2 cm) was cut from a cell culture polystyrene flask. The AzPhPVA was dissolved in water (200 lg/mL). The solution (100 lL) was placed on the polystyrene plate and air-dried at room temperature in the dark. The plate was covered with a patterned photomask and irradiated with ultraviolet light at an intensity of 5.0 103 lJ/cm2 from a distance of 15 cm for 25 s. After irradiation, the plate was immersed in Milli-Q water and then sonicated to completely remove any unreacted polymer from the unirradiated areas. After complete washing, PVA-micropatterned surfaces were obtained. Observation by scanning probe microscopy (SPM). An SPA-400 multi-function SPM unit (SII NanoTechnology Inc., Tokyo, Japan) connected to an SPI4000 controller was used for measurements in air and liquid in contact mode. The cantilevers used were OTR8 silicon nitride probes for measurement in air and OTR4 for measurement in liquid (Veeco Instruments, Santa Barbara, CA) having spring constants of 0.15 N/m and 0.02 N/m, respectively. All SPM measurements were made at room temperature. The polystyrene plates with the PVA-micropattern grafted surfaces were observed in a dry state in air and in a wet state in Milli-Q water. A 100 100 lm area of the sample was observed by SPM. The heights of the micropatterned PVA were measured from the SPM images. Nine spots from the topographic images of each kind of grafted pattern were used to measure the mean heights and standard deviations. The data were expressed as the average ± standard deviation of the nine spots. Cell culture. Human bone marrow-derived MSCs (MSCs) were obtained from Osiris (Worthington Biochemical, Lakewood, NJ) at passage 2. The cells were seeded in T-75 culture flasks (Iwaki Glass, Tokyo, Japan) using proliferation medium purchased from Lonza (Walkersville, MD). The proliferation medium contained 440 mL MSC basal medium, 50 mL mesenchymal cell growth supplement, 10 mL 200 mM L-glutamine, and 0.5 mL penicillin/streptomycin mixture. After reaching confluence, the cells were further subcultured once and used at passage 4. The cells were collected by treatment with trypsin/EDTA solution and suspended in serum (control) medium at a density of 2.50 104 cells/mL. The serum medium was composed of Dulbecco’s modified Eagle’s medium (Sigma–Aldrich, St. Louis, MO) supplemented with 4500 mg/L glucose, 584 mg/L glutamine, 100 U/mL penicillin, 100 lg/mL streptomycin, 0.1 mM nonessential amino acid, 0.4 mM proline, 50 mg/L ascorbic acid, and 10% fetal bovine serum (FBS). The micropatterned polystyrene plates were put in a cell culture dish and a glass cylinder (10 mm in both bore diameter and height) was placed over each PVA-micropatterned polystyrene plate. Cell suspension solution (0.157 mL/well) was added into the glass cylinder (initial cell density: 5.00 103 cells/cm2). The MSCs were cultured in the serum medium for 3 days. To induce adipogenic differentiation, the MSCs were cultured in adipogenic differentiation medium [22] consisting of DMEM serum medium supplemented with 1 lM dexamethasone and 0.5 mM methyl-isobutylxanthine, insulin (10 lg/mL), and 100 lM indomethacin. The cells on the micropatterned surfaces were incubated in the serum medium or the serum control medium for 1, 2, and 3 weeks and used for cell stain-
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ing. Alternatively, for analyzing gene expression, the MSCs were seeded into 6-well cell culture plates at respective densities of 5.00 103, 1.00 104, 2.00 104, and 3.00 104 cells/cm2 and cultured in the adipogenic differentiation medium or the serum control medium for 1, 2, and 3 weeks. Oil Red O staining. The cultured cells were rinsed with phosphate buffer saline (PBS) three times, fixed with 4% paraformaldehyde for 1 h at 4 °C, and then stained with fresh Oil Red O solution for 2 h. The Oil Red O solution was prepared by mixing three parts stock solution (0.5% in isopropanol; Sigma) with two parts water and filtering through a 0.2-lm filter. The stained cells were washed three times with PBS to remove any background Oil Red O stain. Photomicrographs were captured by a optical microscope with a DP-70 CCD camera (Olympus, Tokyo, Japan). RNA isolation and quantitative real-time RT-PCR analysis. After culture for 1 and 3 weeks in the 6-well cell-culture plates, the cells were washed with PBS three times, and 1 mL of Isogen reagent (Nippon Gene, Toyama, Japan) was added to each well. Total RNA was extracted following the manufacturer’s protocol. Total RNA (1 lg) was reversely transcribed into cDNA using a random hexamer primer (Applied Biosystems, Foster City, CA) in a 20 ll reaction. An aliquot (1 ll) of 10-times diluted reaction solution was used for each 25 ll real-time PCR reaction together with 300 nM forward and reverse primers, 150 nM probes, and qPCR MasterMix (Eurogentec, Seraing, Belgium). TaqmanÒ probes and primer pairs for fatty acid binding protein-4 (FABP4, assay identification number Hs00609791_m1) [23], lipoprotein lipase (LPL, assay identification number Hs00173425_m1) [23], and peroxisome proliferatoractivated receptor c2 (PPARc2, assay identification number Hs01115510_m1), which are assay-on-demand gene expression products, were obtained from Applied Biosystems. Real-time quantitative RT-PCR analysis was performed using a housekeeping gene GAPDH and 18S ribosomal RNA, as previously described. After an initial incubation step of 2 min at 50 °C and denaturation for 10 min at 95 °C, 40 cycles of PCR (95 °C for 15 s, 60 °C for 1 min) were performed. Reactions were performed in triplicate. 18S ribosomal RNA levels were used as endogenous controls and gene expression levels relative to GAPDH were calculated using the comparative Ct method. To calculate the means and standard deviations, three samples under each condition were measured. A oneway analysis of variance (ANOVA) with Tukey’s post hoc test for multiple comparisons was used for statistical analysis. A value of p < 0.05 was considered statistically significant.
Results and discussion PVA-micropatterned surface Poly(vinyl alcohol) was micropatterned on polystyrene plates cut from cell culture flasks. First, photoreactive AzPhPVA was synthesized by coupling poly(vinyl alcohol) with 4-azidobenzoic acid. The introduction of photoreactive azido groups in AzPhPVA was confirmed by the appearance of peaks assigned to the phenylazido proton around 7 ppm in the 1H NMR spectrum. The percentage of the hydroxyl groups in the PVA coupled with the azidophenyl groups was 2.1%. Subsequently, the photoreactive AzPhPVA was micropatterned on polystyrene plates by photolithography (Fig. 1A). An aqueous solution of AzPhPVA was eluted on a polystyrene plate and air-dried in the dark. The cast plate was covered with a photomask and photoirradiated. The photomask pattern was composed of alternate UV-transparent and UV-nontransparent stripes (Fig. 1B). The UV-nontransparent stripes were 200 lm wide and the UV-transparent stripes had gradient widths from 20 to 1000 lm. AzPhPVA in the irradiated areas should be intermolecularly and intramolecularly crosslinked and grafted to the poly-
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Fig. 1. The poly(vinyl alcohol) micropattern and cell distribution. Preparation scheme of poly(vinyl alcohol)-micropatterned surface (A). Photomicrograph of the photomask (B). The white stripes were UV-transparent and the dark stripes UV-nontransparent. The photomask had a pattern of UV-transparent stripes with widths varying from 20 to 1000 lm separated by 200 lm UV-nontransparent stripes. Three-dimensional topographic images of scanning probe microscopy (SPM) of poly(vinyl alcohol)-micropatterned surface in air (C) and in Milli-Q water (D). Photomicrographs of MSCs cultured on poly(vinyl alcohol)-micropatterned surface in serum medium after 10 min (E) and 1 day (F). The number indicates the theoretical cell density based on the ratio of the cell adhesive polystyrene area to the nonadhesive PVA area.
styrene surface. In the other areas, AzPhPVA should not be crosslinked and could be removed by washing with Milli-Q water. After washing, a pattern of PVA stripes having gradient widths was prepared. Observation with an optical microscope demonstrated that the PVA formed the same striped pattern as that of the photomask. The PVA-micropatterned surfaces in the dry and wet states were observed by scanning probe microscopy (Fig. 1C and D). The tiered structures of the micropatterned PVA in air and MilliQ water were confirmed by the three-dimensional images. The step height of the micropatterned PVA was 95.75 ± 9.74 nm in air and 143.00 ± 26.50 nm in water. The micropatterned PVA surfaces were smoother and higher in water than they were in air, indicating that the micropatterned PVA swelled when immersed in water. The micropatterned surfaces were used for cell culture in an aqueous medium. Therefore, the SPM images of the micropatterned surface in water reflected the true state of the micropatterned surfaces during cell culture. Adipogenic differentiation of MSCs on the micropatterned surface Mesenchymal stem cells (MSCs) were seeded on the PVA-micropatterned surface at a cell density of 5.00 103 cells/cm2 and cultured in serum medium for 3 days. Immediately after cell seeding, the cells were distributed evenly on the PVA-micropatterned surfaces (Fig. 1E). During cell culture, the cells falling on the PVA stripes moved to the polystyrene stripes because PVA did not support cell adhesion. PVA has been reported to inhibit protein
adsorption and thus inhibit cell adhesion. The MSCs adhered to the polystyrene stripes. After 1 day of culture, the cells were observed only on the polystyrene stripes and formed a striped pattern (Fig. 1F). According to the surface area ratio of the polystyrene stripes and PVA stripes, the cell density from the right to left polystyrene stripes should theoretically be 1.10, 1.10, 1.30, 1.50, 1.75, 2.00, 2.50, 3.00, 3.50, 4.00, 4.50, 5.00, 5.50, 6.00, and 6.00 times that of 5.00 103 cells/cm2 assuming that the cells moved randomly from the nonadhesive PVA stripes to the polystyrene stripes. The cell number on the polystyrene stripes was counted from the three areas of each stripe and cell density was calculated. The cell density on each stripe from right to left was 4.84 ± 0.37, 3.91 ± 0.26, 6.71 ± 0.65, 6.95 ± 2.05, 7.58 ± 2.48, 8.82 ± 0.78 103 and 1.17 ± 0.45, 1.25 ± 0.36, 1.49 ± 0.50, 1.92 ± 0.54, 2.16 ± 0.36, 2.40 ± 0.34, 2.63 ± 0.30, 2.79 ± 0.33, 2.98 ± 0.27 104 cells/cm2, respectively. The result indicates that cell density could be adjusted by controlling the ratio of the cell adhesion and nonadhesion areas and cell density gradient was formed on the PVA-micropatterned surface. After the MSCs were cultured on the PVA-micropatterned surfaces in the serum medium for 3 days, the culture medium was changed to an adipogenic differentiation medium and the cells were further cultured for 1, 2, and 3 weeks. Some of the cells were further cultured in serum medium for 1, 2, and 3 weeks as a control. Lipid vacuoles were observed after culture for 1 week (Fig. 2A–C). The number of lipid vacuoles was low at the low density cells and abundant at the high density cells. The number of li-
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Fig. 2. Photomicrographs of MSCs cultured on poly(vinyl alcohol)-micropatterned surface for 1 (A, D), 2 (B, E), and 3 (C, F) weeks in adipogenic differentiation medium without staining (A–C) and stained with Oil Red O (D–F). Scale bar = 500 lm.
pid vacuoles increased after 2 and 3 weeks of culture. The cells at low density (right stripes of Fig. 2) did not become confluent. Some cells existed alone. The cells at high density (left stripes of Fig. 2) became confluent and came into contact with each other. Culture in adipogenic differentiation medium arrested cell proliferation. The lipid vacuoles were further stained with Oil Red O (Fig. 2D– F). Oil Red O positive cells were observed after 1 week of culture in the adipogenic differentiation medium. The cells at high cell density were more densely stained by Oil Red O than were the cells at low cell density. Oil Red O staining also increased with culture
time. No lipid vacuoles were detected on the MSCs cultured in the control medium (Fig. 3). The cells proliferated and became confluent when cultured in the serum control medium. Although the density of the lipid vacuoles and Oil Red O staining increased with the increase of cell density, the MSCs at all cell densities differentiated into adipocyte-like cells. Cell density did not affect adipogenic differentiation of MSCs in the cell density range used in the present study. To check the effect of cell density on the expression of genes relative to adipogenic differentiation, the MSCs were cultured in 6-
Fig. 3. Photomicrographs of Oil Red O staining of MSCs cultured on poly(vinyl alcohol)-micropatterned surface in serum control medium for 1 (A), 2 (B), and 3 (C) weeks. Scale bar = 500 lm.
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well cell culture polystyrene plates. The seeded cell densities were changed from 1.00, 2.00, 4.00, and 6.00 times that 5.00 103 cells/ cm2. The cells were cultured in adipogenic differentiation and serum control mediums for 1 and 3 weeks. The adipogenesis marker genes encoding peroxisome proliferator-activated receptor c2 (PPARc2), lipoprotein lipase (LPL), and fatty acid binding protein4 (FABP4) were analyzed (Fig. 4). The MSCs cultured in adipogenic differentiation medium expressed the genes while the MSCs cultured in control medium did not. All the genes were detected in the MSCs cultured at different cell densities. The expression level of the genes increased with culture time. There was no significant
difference in the expression level of these genes among the four cell densities. The cell density did not show any evidence of an effect on the expression of the genes. It has been reported that a high density of MSCs facilitates adipogenic differentiation [18]. Conventionally, before adipogenic differentiation induction, cells are cultured to confluence because the adipogenic induction medium inhibits cell proliferation [22,24]. On the other hand, it has also been reported [25] that single-cell derived colonies generated by plating early passage human MSCs at extremely low densities show adipogenic differentiation. The passage cell density [20,26] has been reported to affect MSC proliferation and differentiation potential. Lower cell density results in a higher proliferation rate during cell passage, but the multilineage differentiation potential of the passaged cells is not affected by the passage cell density. In the present study, MSCs of different densities were cultured on a single surface. The MSCs of different densities showed similar adipogenic differentiation. No apparent effect of cell density on adipogenic differentiation was observed. The cell density range in the present study was from 5.00 103 to 3.00 104 cells/cm2. A wider range is possible if the ratio of polystyrene and PVA surface areas is adjusted by changing the photomask. The patterning technique will provide a powerful tool for creating cell arrays in a density gradient to directly compare the effect of cell density on cell functions. Acknowledgments This work was supported in part by the World Premier International Research Center (WPI) Initiative on Materials Nanoarchitectonics, MEXT, Japan, and in part by the New Energy and Industrial Technology Development Organization of Japan. References
Fig. 4. Real-time PCR results of mRNA expression of PPARc2 (A), LPL (B), and FABP4 (C) genes of MSCs cultured at various cell densities in 6-well cell culture polystyrene plates with adipogenic differentiation and serum control medium for 1 and 3 weeks. Cell density is indicated under the horizontal axis. MSCs indicate cells used for cell seeding. Control means the cells were cultured in serum medium. The data are normalized to GAPDH and represent means ± SD (n = 3).
[1] R.O. Oreffo, C. Cooper, C. Mason, M. Clements, Mesenchymal stem cells: lineage, plasticity, and skeletal therapeutic potential, Stem Cell Rev. 1 (2005) 169–178. [2] C. Lange, J. Schroeder, M.V. Lioznov, A.R. Zander, High-potential human mesenchymal stem cell, Stem Cells Dev. 14 (2005) 70–80. [3] M.F. Pittenger, Mesenchymal stem cells from adult bone marrow, Methods Mol. Biol. 449 (2008) 27–44. [4] G. Chen, D. Liu, M. Tadokoro, R. Hirochika, H. Ohgushi, J. Tanaka, T. Tateishi, Chondrogenic differentiation of human mesenchymal stem cells cultured in a cobweb-like biodegradable scaffold, Biochem. Biophys. Res. Commun. 322 (2004) 50–55. [5] L. Meinel, V. Karageorgiou, R. Fajardo, B. Snyder, V. Shinde-Patil, L. Zichner, D. Kaplan, R. Langer, G. Vunjak-Novakovic, Bone tissue engineering using human mesenchymal stem cells: effects of scaffold material and medium flow, Ann. Biomed. Eng. 32 (2004) 112–122. [6] K. Natsu, M. Ochi, Y. Mochizuki, H. Hachisuka, S. Yanada, Y. Yasunaga, Allogeneic bone marrow-derived mesenchymal stromal cells promote the regeneration of injured skeletal muscle without differentiation into myofibers, Tissue Eng. 10 (2004) 1093–1112. [7] H.A. Awad, D.L. Butler, M.T. Harris, R.E. Ibrahim, Y. Wu, R.G. Young, S. Kadiyala, G.P. Boivin, In vitro characterization of mesenchymal stem cell-seeded collagen scaffolds for tendon repair: effects of initial seeding density on contraction kinetics, J. Biomed. Mater. Res. 51 (2000) 233–240. [8] H. Fan, H. Liu, S.L. Toh, J.C. Goh, Enhanced differentiation of mesenchymal stem cells co-cultured with ligament fibroblasts on gelatin/silk fibroin hybrid scaffold, Biomaterials 29 (2008) 1017–1027. [9] M. Neubauer, M. Hacker, P. Bauer-Kreisel, B. Weiser, C. Fischbach, M.B. Schulz, A. Goepferich, T. Blunk, Adipose tissue engineering based on mesenchymal stem cells and basic fibroblast growth factor in vitro, Tissue Eng. 11 (2005) 1840–1851. [10] Y.S. Choi, S. Park, H. Suh, Adipose tissue engineering using mesenchymal stem cells attached to injectable PLGA spheres, Biomaterials 26 (2005) 5855–5863. [11] T.C. Otto, M.D. Lane, Adipose development: from stem cell to adipocyte, Crit. Rev. Biochem. Mol. Biol. 40 (2005) 229–242. [12] R.R. Bowers, M.D. Lane, A role for bone morphogenetic protein-4 in adipocyte development, Cell Cycle 6 (2007) 385–389. [13] I.C. Lee, J.H. Wang, Y.T. Lee, T.H. Young, The differentiation of mesenchymal stem cells by mechanical stress or/and co-culture system, Biochem. Biophys. Res. Commun. 352 (2007) 147–152. [14] L. Guo, N. Kawazoe, Y. Fan, Y. Ito, J. Tanaka, T. Tateishi, X. Zhang, G. Chen, Chondrogenic differentiation of human mesenchymal stem cells on photoreactive polymer-modified surfaces, Biomaterials 29 (2008) 23–32.
H. Lu et al. / Biochemical and Biophysical Research Communications 381 (2009) 322–327 [15] S.J. Song, O. Jeon, H.S. Yang, D.K. Han, B.S. Kim, Effects of culture conditions on osteogenic differentiation in human mesenchymal stem cells, J. Microbiol. Biotechnol. 17 (2007) 1113–1119. [16] B. Neuhuber, S.A. Swanger, L. Howard, A. Mackay, I. Fischer, Effects of plating density and culture time on bone marrow stromal cell characteristics, Exp. Hematol. 36 (2008) 1176–1185. [17] M. Bitar, R.A. Brown, V. Salih, A.G. Kidane, J.C. Knowles, S.N. Nazhat, Effect of cell density on osteoblastic differentiation and matrix degradation of biomimetic dense collagen scaffolds, Biomacromolecules 9 (2008) 129–135. [18] R. McBeath, D.M. Pirone, C.M. Nelson, K. Bhadriraju, C.S. Chen, Cell shape cytoskeletal tension, and RhoA regulate stem cell lineage commitment, Dev. Cell 6 (2004) 483–495. [19] S.K. Both, A.J. van der Muijsenberg, C.A. van Blitterswijk, J. de Boer, J.D. de Bruijn, A rapid and efficient method for expansion of human mesenchymal stem cells, Tissue Eng. 13 (2007) 3–9. [20] D.C. Colter, R. Class, C.M. DiGirolamo, D.J. Prockop, Rapid expansion of recycling stem cells in cultures of plastic-adherent cells from human bone marrow, Proc. Natl. Acad. Sci. USA 97 (2000) 3213–3218.
327
[21] N. Kawazoe, L. Guo, M.J. Wozniak, Y. Imaizumi, T. Tateishi, X. Zhang, G. Chen, Adipogenic differentiation of mesenchymal stem cells on micropatterned polyelectrolyte surfaces, J. Nanosci. Nanotechnol. 9 (2009) 230–239. [22] J.R. Mauney, V. Volloch, D.L. Kaplan, Matrix-mediated retention of adipogenic differentiation potential by human adult bone marrow-derived mesenchymal stem cells during ex vivo expansion, Biomaterials 26 (2005) 6167–6175. [23] M.F. Pittenger, A.M. Mackay, S.C. Beck, R.K. Jaiswal, R. Douglas, J.D. Mosca, M.A. Moorman, D.W. Simonetti, S. Craig, D.R. Marshak, Multilineage potential of adult human mesenchymal stem cells, Science 284 (1999) 143–147. [24] K.D. Murphy, J.O. Lee, D.N. Herndon, Current pharmacotherapy for the treatment of severe burns, Expert Opin. Pharmacother. 4 (2003) 369–384. [25] I. Sekiya, B.L. Larson, J.T. Vuoristo, J.G. Cui, D.J. Prockop, Adipogenic differentiation of human adult stem cells from bone marrow stroma (MSCs), J. Bone Miner. Res. 19 (2004) 256–264. [26] P.A. Sotiropoulou, S.A. Perez, M. Salagianni, C.N. Baxevanis, M. Papamichail, Characterization of the optimal culture conditions for clinical scale production of human mesenchymal stem cells, Stem cells 24 (2006) 462–471.