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Matrix elasticity, replicative senescence and DNA methylation patterns of mesenchymal stem cells
Biomaterials xxx (2014) 1e8
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Matrix elasticity, replicative senescence and DNA methylation patterns of mesenchymal stem cells Anne Schellenberg a, Sylvia Joussen a, Kristin Moser a, b, Nico Hampe b, Nils Hersch b, Hatim Hemeda a, Jan Schnitker c, Bernd Denecke d, Qiong Lin a, e, Norbert Pallua f, Martin Zenke e, Rudolf Merkel b, Bernd Hoffmann b, Wolfgang Wagner a, * a Helmholtz-Institute for Biomedical Engineering, Stem Cell Biology and Cellular Engineering, RWTH Aachen University Medical School, Pauwelsstrasse 20, Aachen 52074, Germany b Institute of Complex Systems, ICS-7: Biomechanics, Forschungszentrum Jülich GmbH, Jülich 52425, Germany c Institute of Complex Systems, ICS-8: Bioelectronics, Forschungszentrum Jülich GmbH, Jülich 52425, Germany d Interdisciplinary Centre for Clinical Research (IZKF) Aachen, RWTH Aachen University Medical School, 52074 Aachen, Germany e Institute for Biomedical Technology e Cell Biology, RWTH Aachen University Medical School, Aachen, Germany f Department of Plastic and Reconstructive Surgery, RWTH Aachen University Medical School, 52074 Aachen, Germany
a r t i c l e i n f o
a b s t r a c t
Article history: Received 28 March 2014 Accepted 22 April 2014 Available online xxx
Matrix elasticity guides differentiation of mesenchymal stem cells (MSCs) but it is unclear if these effects are only transient e while the cells reside on the substrate e or if they reflect persistent lineage commitment. In this study, MSCs were continuously culture-expanded in parallel either on tissue culture plastic (TCP) or on polydimethylsiloxane (PDMS) gels of different elasticity to compare impact on replicative senescence, in vitro differentiation, gene expression, and DNA methylation (DNAm) profiles. The maximal number of cumulative population doublings was not affected by matrix elasticity. Differentiation towards adipogenic and osteogenic lineage was increased on soft and rigid biomaterials, respectively e but this propensity was no more evident if cells were transferred to TCP. Global gene expression profiles and DNAm profiles revealed relatively few differences in MSCs cultured on soft or rigid matrices. Furthermore, only moderate DNAm changes were observed upon culture on very soft hydrogels of human platelet lysate. Our results support the notion that matrix elasticity influences cellular behavior while the cells reside on the substrate, but it does not have major impact on cellintrinsic lineage determination, replicative senescence or DNAm patterns. Ó 2014 Elsevier Ltd. All rights reserved.
Keywords: Mesenchymal stem cells Elasticity Long-term culture Platelet lysate DNA-methylation Epigenetic
1. Introduction Cells are normally embedded in a complex microenvironment of specific topography and matrix elasticity [1e4]. The elastic modulus of extracellular matrix (ECM), also known as Young’s modulus, ranges from 0.1 kPa in brain, some few kPa in fat tissue, to several hundred MPa in calcified bone [5,6]. In contrast, in vitro cell culture is usually performed on very rigid polystyrene surfaces of several GPa. Various studies have indicated that substrate elasticity is an important regulator for cellular function and these were mostly performed on polyacrylamide hydrogels [5,7,8]. However,
* Corresponding author. Helmholtz-Institute for Biomedical Engineering, Stem Cell Biology and Cellular Engineering, RWTH Aachen University Medical School, Pauwelsstrasse 20, 52074 Aachen, Germany. Tel.: þ49 241 8088611. E-mail address: [email protected] (W. Wagner).
changes in topography upon surface creasing of polyacrylamide substrates can significantly influence cell behavior [9]. Alternatively, elastomers of cross-linked polydimethylsiloxane (PDMS), a viscoelastic organic polymer, display rather smooth surfaces [10]. PDMS is widely used in biomedical engineering and cell culture due to good biocompatibility, optical transparency and tunable stiffness e it is therefore a suitable substrate for mechanotransduction experiments [11]. Effects of matrix elasticity were often addressed in mesenchymal stem cells (MSCs). These cells comprise a multipotent subset capable of differentiation potential towards osteocytes, chondrocytes and adipocytes and they are concurrently tested in a wide variety of clinical trials [12,13]. It has been suggested that matrix elasticity alone can direct MSC faith: soft (0.1e1 kPa), intermediate (8e17 kPa), and rigid (>34 kPa) polyacrylamide hydrogels supported differentiation towards neurogenic, myogenic and osteogenic lineages, respectively [5]. MSCs reveal directed
http://dx.doi.org/10.1016/j.biomaterials.2014.04.079 0142-9612/Ó 2014 Elsevier Ltd. All rights reserved.
Please cite this article in press as: Schellenberg A, et al., Matrix elasticity, replicative senescence and DNA methylation patterns of mesenchymal stem cells, Biomaterials (2014), http://dx.doi.org/10.1016/j.biomaterials.2014.04.079
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A. Schellenberg et al. / Biomaterials xxx (2014) 1e8
migration towards stiffer substrates, called durotaxis [14]. Cell adhesion, cell spreading, and proliferation rates are significantly increased on rigid biomaterials [8,15,16]. It has even been suggested that stiffness gradients, rather than stiffness alone, might be crucial for regulation of MSC behavior [14]. This is of particular relevance for tissue engineering approaches with MSCs embedded in matrix or scaffolds of specific elasticity. If matrix elasticity also entails persistent lineage conversion it would be relevant for generation of tailored cell products, too. Cellular differentiation is reflected by the epigenetic makeup. DNA methylation (DNAm) is the best characterized epigenetic modification: cytosine guanine dinucleotides (CpGs) can be methylated at cytosine moieties and this governs chromatin structure and gene regulation [17]. It is yet unclear if matrix elasticity impacts on DNAm profiles e which would be expected if matrix elasticity induces persistent cell-intrinsic modifications. All primary cells enter replicative senescence after a certain number of cell divisions, the so called Hayflick-Limit [18]. This process is accompanied by large-spread morphology, loss of differentiation potential, and unequivocal proliferation arrest [19e 22]. We have demonstrated that long-term culture of MSCs is associated with highly reproducible modifications in the DNAm pattern, particularly in developmental genes [23,24]. Notably, these senescence-associated DNAm changes are reversed upon reprogramming of MSCs into induced pluripotent stem cells (iPSCs) e and iPSCs do not reveal replicative senescence while in pluripotent state [25]. Recently, it has been reported that tissue stiffness and stress increase lamin-A levels, which stabilizes the nucleus [26]. Furthermore, hypomethylation upon long-term culture is enriched in lamin-associated domains [27] (Hänzelmann S. et al., manuscript in preparation). Hence it might be speculated that substrate stiffness, which alters cellular and nuclear morphology, contributes to cellular senescence. In this study, we have continuously culture expanded MSCs on PDMS of different Young’s modulus until they reached replicative senescence. Thereby, we wanted to determine if matrix elasticity affects cellular aging, whether it induces persistent lineage commitment, and if this is also reflected on DNAm level. 2. Methods 2.1. Fabrication of elastomeric substrates Polydimethylsiloxan substrates (Sylgard 184 Silicone Elastomer Kit, Dow Corning, MI, USA) were generated by mixing base and cross linker at ratios of 70:1 (1.5 kPa), 60:1 (6.5 kPa), 50:1 (15 kPa), 40:1 (50 kPa) and 30:1 (100 kPa). The mixture was then degassed and 2 ml were added in wells of 6-well plates. Cross-linking was performed at 60 C for 16 h. Characterization of elastomer material properties were performed by stretching cylindrical test pieces and macroscopic indentation tests as described previously [28,29]. For generation of human platelet lysate (hPL) gels 5 platelet units were pooled as described before [30]. Culture medium with 10% hPL gelatinized in the absence of anticoagulants due to the plasma components. The cells were then seeded on these hPL-gels in the same culture medium with 10% hPL supplemented with heparin (1 IU/ml) [31]. For passaging, MSCs were harvested together with the hPL-gel by pipetting, diluted with culture medium, and then plated onto new plates with hPLgel as described previously [30]. To determine cell numbers, fractions of the cell suspension were plated on TCP and conventionally counted upon adhesion to TCP (within 1 day). 2.2. Isolation and culture of MSCs Mesenchymal stem cells were isolated from lipoaspirates after patient’s written consent using guidelines approved by the Ethic Committee of the RWTH Aachen University Medical School (Permit Number EK163/07). Lipoaspirates were digested with 2 g/L collagenase type I (PAA, Pasching, Austria) [32] and seeded in culture medium consisting of Dulbecco’s Modified Eagles Medium-Low Glucose (DMEM-LG; PAA) with 2 mM L-glutamine (Sigma Aldrich, St. Louis, MO, USA), 100 U/mL penicillin/ streptomycin (pen/strep; Lonza, Basel, Switzerland) and 10% hPL [33]. Prior to cell seeding PDMS substrates were coated with 2.5 mg/cm2 fibronectin (Biochrom Berlin, Germany) for 45 min at 37 C. Cultures were maintained at 37 C in a humidified atmosphere containing 5% CO2 with medium changes twice per week. Fibroblastoid
colony-forming unit (CFU-f)-frequencies were determined 14 days after seeding of MSCs using crystal violet staining [32]. Upon 80% confluent growth MSCs were harvested from the substrates (either TCP or PDMS) using trypsin, counted with a Neubauer chamber (Brand, Wertheim, Germany), and re-seeded at a density of 10,000 cells/cm2 in 6-well-plates (Nunc Thermo Fisher Scientific, Langenselbold, Germany). Cell population doublings were analyzed for each passage and cumulative population doublings (cPD) were calculated from the first passage onward as described before [32]. 2.3. Characterization of MSCs Surface marker expression was analyzed on a FACSCanto II (Beckton Dickinson, BD) upon staining with the following antibodies as described before [32]: CD14allophycocyanin (APC, clone M5E2, BD), CD29-phycoerythrin (PE, clone MAR4, BD), CD31-PE (clone WM59, BD), CD34-APC (clone 8G12, BD), CD45-APC (clone HI30, BD), CD73-PE (clone AD2, BD), CD90-APC (clone 5E10, BD), CD105-fluorescein isothiocyanate (FITC, clone MEM-226 Immuno Tools). Osteogenic and adipogenic differentiation potential of MSCs was assessed in parallel on the different substrates as described before [22,34]. Alternatively, some of the cells were harvested from the PDMS substrates and cultured on TCP for two passages before induction of differentiation. Osteogenic differentiation was estimated by Alizarin Red staining and quantified with a Tecan infinite M200 platereader (405 nm) [35]. Adipogenic differentiation was estimated by counting cells with lipid-droplets in phase contrast images from five randomly chosen areas (Lipophilic dyes could not be used on PDMS) with counter staining with DAPI (40 ,6Diamidin-2-phenylindol; both Molecular Probes, Eugene, Oregon, USA) [32,36]. Images were captured using a Leica DM IL LED microscope (Leica, Wetzlar, Germany). 2.4. Analysis of senescence-associated b-galactosidase Expression of pH-dependent senescence associated b-galactosidase (SA-b-gal) activity was analyzed by flow cytometry as described before [37]: MSCs were pretreated with Bafilomycin A1 (Sigma, St Louis, MO, USA) and 5dodecanoylaminofluorescein di-b-D-galactopyranoside (C12FDG, Invitrogen, Eugene, OR, USA) was subsequently used as fluorogenic substrate for bgalactosidase. 2.5. Apoptosis assay Apoptosis was estimated by flow cytometry using Annexin V and propidium iodid (PI) double staining. In brief, MSCs were harvested from the substrates, washed twice in phosphate buffered saline (PBS), and 1 105 cells were stained in 5 ml Annexin V with 100 ml binding buffer (Becton Dickinson [BD], San Jose, USA) for 15 min at RT in the dark. Subsequently, 400 ml Annexin V binding puffer (BD) and 2 ml PI were added prior to analysis on a FACSCanto II (BD). 2.6. Immunofluorescence and scanning electron microscopy MSCs were grown on glass or PDMS substrates for 8 days, fixed using 3.7% formaldehyde (Merck, Germany) and stained for vinculin and actin expression as previously described [38]. For scanning electron microscopy of cells on PDMS, 5000 cells were grown on cross-linked PDMS substrates exhibiting elasticities of 1.5 and 100 kPa, respectively. After 24 h of growth cells were treated as described before [39]. Substrates were sputtered with a thin layer of gold and analyzed by a LEO-1550 scanning electron microscope (Zeiss, Germany). Scanning electron microscopy of hPL gels was performed as described before [30]. 2.7. Gene expression analysis Gene expression profiles were analyzed in three MSC preparations (always at passage 4) upon continuous culture-expansion on PDMS (1.5 kPa or 50 kPa) or TCP. RNA was isolated from 106 cells using the NucleoSpin RNA Isolation Kit (Macherey Nagel, Düren, Germany) and quality control was carried out using the Agilent 2100 Bioanalyzer (Agilent Technologies, Inc., Santa Clara, CA, USA). RNA was amplified and hybridized to the GeneChip Human Gene 1.0 ST Arrays (Affymetrix, Santa Clara, CA, USA) according to the manufacturer’s instructions. Raw data were normalized by RMA (Affymetrix Power Tools). The microarray data are accessible in NCBIs Gene Expression Omnibus (GEO, http://www.ncbi.nlm.nih.gov/geo/) through accession number GSE55867. Pairwise limma-T-test analysis was calculated in R to select for differentially expressed genes (adjusted p-value < 0.05). Quantitative real time PCR (RT-qPCR) of selected genes was performed using the High Capacity cDNA Reverse Transcription Kit and a StepOnePlus instrument (both Applied Biosystems, Carlsbad, CA, USA) in a MicroAmp optical 96-well reaction plate. TaqMan Gene Expression Assays (Applied Biosystems) used in this study are listed in Supplemental Table 1: Gene expression levels were normalized to GAPDH expression. 2.8. DNA-methylation profiling DNA methylation profiles were analyzed in four MSC preparations (always at passage 4) upon continuous culture-expansion on PDMS (1.5 kPa or 50 kPa) or TCP.
Please cite this article in press as: Schellenberg A, et al., Matrix elasticity, replicative senescence and DNA methylation patterns of mesenchymal stem cells, Biomaterials (2014), http://dx.doi.org/10.1016/j.biomaterials.2014.04.079
A. Schellenberg et al. / Biomaterials xxx (2014) 1e8 Furthermore, we compared DNAm profiles of 4 MSC preparations (passage 4) cultured either on TCP or hPL-gel. Genomic DNA was isolated from 106 cell using the QIAamp DNA Blood Midi Kit (Qiagen, Hilden, Germany) and subsequently bisulfiteconverted (EZ DNA MethylationTM Kit, Zymo, Irvine, USA). DNAm profiles were analyzed using the HumanMethylation450 BeadChip (Illumina, San Diego, USA) as described before [40]. In brief, about 200 ng of bisulfite converted DNA was hybridized according to the manufacturer’s instructions. After single-base extension using DNP- and Biotin-labeled ddNTPs, the array was fluorescently stained, scanned, and the intensities of non-methylated and methylated bead types measured. Hybridization and initial data analysis with the BeadStudio Methylation Module was performed at the DKFZ Gene Core Facility (Heidelberg, Germany). The complete dataset is accessible through GEO Series accession number GSE55888. Hierarchical clustering of DNAm profiles was performed using the MultiExperiment Viewer (MeV, TM4; Euclidian distance). Pairwise limma-T-test analysis was calculated in R to select for CpGs with significant different DNAm levels (adjusted p-value < 0.05). Under these parameters no CpGs reached the significance threshold and therefore we alternatively selected CpGs with at least 20% differential DNAm in MSCs isolated on different substrates. Genes associated with the differentially methylated CpG sites were classified by Gene Ontology analysis using DAVID Bioinformatics Resources 6.7 (http://david.abcc.ncifcrf.gov/home.jsp). 2.9. Statistics Results are expressed as mean standard deviation (SD) of at least three independent experiments. To estimate the probability of differences we have adopted the paired two-sided Student’s T-test.
3. Results 3.1. Elasticity and replicative senescence of MSCs Mesenchymal stem cells were isolated from adipose tissue and continuously cultured in parallel either on TCP or on PDMS of different stiffness (1.5 kPa, 6.5 kPa, 15 kPa, 50 kPa and 100 kPa). Scanning electron microscopy revealed that PDMS substrates were
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evenly flat and cells attached on this substrate (Fig. 1A). Initial colony formation was higher on TCP than on PDMS (Fig. 1B). After 7 days (passage 0) all cell preparations displayed the typical immunophenotypic pattern of MSCs (CD14-, CD29þ, CD31, CD34þ/-, CD45, CD73þ, CD90þ, CD105þ and CD140; Supplemental Fig. 1), regardless of the substrate they were isolated on. Proliferation rate was slightly higher on TCP and there was no significant difference in the long-term growth curves (Fig. 1C): MSCs continuously expanded in parallel on TCP, PDMS 1.5 kPa, or PDMS 50 kPa entered replicative senescence within 90 days with a similar maximal number of cumulative population doublings (cPDs; Fig. 1C). However, senescent cells on elastic substrates did not reveal typical large “fried-egg” morphology and remained overall smaller (Fig 1D; Supplemental Fig. 2). Furthermore, neither flow cytometric measurement of senescence associated b-galactosidase (SA-b-gal) activity (Fig. 1E, F), nor apoptosis rates (Fig. 1G) revealed significant differences upon expansion on the various substrates. Thus, matrix elasticity affects initial outgrowth and morphology of MSCs but it does not impact on replicative senescence. 3.2. Impact of elasticity on in vitro differentiation MSCs culture expanded on the different substrate elasticities were differentiated towards osteogenic or adipogenic lineage: osteogenic differentiation, reflected by calcium phosphate depositions in Alizarin Red staining after 14 days, markedly increased with substrate stiffness (Fig. 2A); whereas adipogenic differentiation, estimated by the percentage of cells with lipid droplets, was more pronounced on softer PDMS substrates (Fig. 2B). These tendencies have also been described in previous studies [5,8,15,41].
Fig. 1. Long-term culture of MSCs on elastic PDMS substrates. (A) Scanning electron microscopy of MSCs cultured on PDMS 1.5 kPa (scale bar 20 mm). (B) Initial fibroblastoid colony forming unit (CFU-f) frequency of adipose tissue derived MSCs was higher on tissue culture plastic (TCP) than on PDMS substrates (n ¼ 5; *p < 0.05). (C) MSCs of 3 donors were continuously culture expanded on soft PDMS (1.5 kPa), stiffer PDMS (50 kPa) and rigid TCP until replicative senescence. Cumulative population doublings (cPDs) were calculated from the first passage onward (transient inclination of proliferation after 30 d might be due to change of hPL supplements). (D) No significant differences were observed in the maximum number of cPDs. (E) Cells of late passages on elastic substrates did not show the typical “fried egg morphology” of senescent cells (scale bar 100 mm). (F, G) However, flow cytometric analysis of senescence-associated b-galactosidase by C12FDG was not affected by the biomaterials (n ¼ 3). (H) Analysis of apoptotic cells by flow cytometric analysis of Annexin V did not reveal any differences, too (n ¼ 3).
Please cite this article in press as: Schellenberg A, et al., Matrix elasticity, replicative senescence and DNA methylation patterns of mesenchymal stem cells, Biomaterials (2014), http://dx.doi.org/10.1016/j.biomaterials.2014.04.079
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Fig. 2. Transient effect of substrate elasticity on differentiation of MSCs. (A) Osteogenic differentiation was induced for two weeks in MSCs cultured on either tissue culture plastic (TCP) or on cross-linked PDMS substrates with indicated Young’s moduli. Semiquantitative analysis of calcium phosphate precipitates with a plate reader upon Alizarin red staining revealed increased differentiation with substrate stiffness (*p < 0.05). (B) In contrast, adipogenic differentiation, determined by the percentage of cells containing lipid-droplets, was increased on softer matrices (*p < 0.05). (C) Staining for actin (green) and vinculin (red) in MSCs on PDMS substrates and glass. If indicated, cells were additionally differentiated towards osteogenic and adipogenic lineages for two weeks (size bar ¼ 50 mm). (D, F) MSCs which were continuously cultured on the substrates as indicated were re-seeded on TCP before induction of in vitro differentiation e thereby the above mentioned propensity of lineage-specific differentiation was lost. (For interpretation of the references to color in this figure legend, the reader is referred to the web version of this article.)
Immunostaining of vinculin and actin revealed heterogeneous cell morphologies on all elasticities. Upon osteogenic differentiation the cells enlarged with more stress fibers which could be visualized on all substrates. In contrast adipogenic differentiation resulted in rather round cellular morphology with less pronounced actin structures and accumulation of lipid droplets was particularly observed on soft substrates (Fig. 2C). Subsequently, MSCs were harvested from the different substrates for two additional passages on TCP. Now the initially observed propensities for in vitro differentiation e osteogenic differentiation on stiff and adipogenic differentiation on soft matrices e were not evident any more (Fig. 2D, F). Thus matrix elasticity appears to affect in vitro differentiation only while the cells are on the matrix, but it does not result in cellintrinsic fate conversion which remains upon passaging on other substrate elasticities. 3.3. Effects of elasticity on gene expression profiles Global gene expression profiles were compared in MSCs continuously cultured on different substrates (TCP, PDMS 1.5 kPa, PDMS 50 kPa; all passage 4). Hierarchical cluster analysis revealed close relationship of MSC preparations derived from the same donor (Fig. 3A). However, gene expression profiles of MSCs cultured on PDMS 1.5 kPa versus PDMS 50 kPa did not reveal any significant differences (adjusted limma paired-T-test < 0.05). Only nine genes were differentially expressed in MSCs expanded on PDMS 1.5 kPa versus TCP (Fig 3B): sulfatase 1 (SULF1), fibronectin type III domain containing 1 (FNDC1), retinoic acid receptor
responder 2 (RARRES2), myosin heavy chain 11 (MYH11), and vascular cell adhesion protein 1 (VCAM1) were higher expressed on TCP, whereas ATP-binding cassette C3 (ABCC3), chemokine C-X-C motif ligand 1 (CXCL1), ribosomal protein SA pseudogene 52 (RPSAP52), and interleukin 8 (IL-8) were higher expressed on PDMS 1.5 kPa. Furthermore, only seven genes were differential expressed in MSCs expanded on PDMS 50 kPa versus TCP (Fig. 3C): SULF1, FNDC1, RARRES2, MYH11, lymphoid enhancer-binding factor-1 (LEF1), and thymocyte expressed, positive selection associated 1 (KIAA0748) are higher expressed on TCP, whereas ABCC3 was higher expressed on PDMS 50 kPa. Differential expression was confirmed by RT-qPCR (Fig. 3D, E). These results were somewhat unexpected, as previous studies indicated substrate elasticity has major impact on gene expression profiles [5]. We have subsequently focused on the described lineage-specific gene expression changes but the effects of matrix elasticity were not recapitulated in our data (Supplemental Fig. 3). 3.4. Effects of elasticity on DNA-methylation profiles Subsequently, we analyzed DNAm profiles of MSCs expanded on different substrates (TCP, PDMS 1.5 kPa, or PDMS 50 kPa) using the Illumina HumanMethylation450 platform which assays more than 480,000 CpG sites at single base resolution (covering 99% of RefSeq genes and 96% of CpG islands) [40]. Hierarchical cluster analysis of DNAm profiles demonstrated donor-specific differences as described before [42] (Fig. 4A). MSCs did not cluster according to the substrate elasticities and this was also observed in principal
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Fig. 3. Gene expression profiles of MSCs on different substrates. (A) Gene expression profiles of MSCs cultured for 4 passages in parallel on PDMS substrates (1.5 kPa or 50 kPa) or tissue culture plastic (TCP) were analyzed on Affymetrix 1.0 ST microarrays. The three individual donor samples clustered closely together. (B) Nine genes with differential gene expression in MSCs on PDMS 1.5 kPa versus TCP, (C) and seven genes with differential expression on PDMS 50 kPa versus TCP are depicted (Limma paired t-test, adjusted pvalue < 0.05). (D, E) Differential expression of these genes was validated by RT-qPCR.
component analysis (PCA, not shown). Significance Analysis of Microarray (SAM) or limma paired T-test did not reveal any CpG with differential DNAm between the growth conditions (adjusted p-value < 0.05). Alternatively, we focused on the four CpGs with more than 20% difference in DNAm level. Culture of MSCs on stiffer substrates resulted in hypermethylation of two CpGs associated with the homeobox genes HOXA5 and HOXC4 (Fig. 4BeD). However, these differences were neither significant, nor were they correlated
with differential gene expression. Alternatively, we have focused on CpGs which reflect senescence-associated DNAm changes in MSCs (1702 CpGs with >20% hypermethylated; 2116 CpGs with >20% hypomethylation in MSCs of late passage [25]), but there was no association with DNAm patterns of MSCs expanded on different substrate elasticities (Supplemental Fig. 4). Overall, elasticity therefore does not evoke significant differences in global DNAm profiles.
Fig. 4. DNA methylation profiles of MSCs on elastic substrates. (A) DNAm profiles of MSCs cultured in parallel on PDMS 1.5 kPa, PDMS 50 kPa and TCP (passage 4) were compared by HumanMethylation450 BeadChips. Hierachical clustering revealed separation according to the four donor samples. (B, C, D) Scatterplots depict CpGs with more than 20% differential methylation between MSCs cultured on the different substrates and the corresponding gene IDs are highlighted. None of these CpGs revealed significant DNAm changes (adjusted pvalue < 0.05).
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3.5. DNA-methylation patterns on platelet lysate-hydrogel To further verify the effects of substrate elasticity on DNAm we performed additional experiments with MSCs cultured on hPL-gel. These hydrogels consist of a fibrin-network which forms in the absence of anticoagulants and possess an extremely low Young’s modulus of about 30 Pa [30] (Fig. 5A). Notably, hPL-gel consists of the same components as the over-layered culture medium, apart from heparin, and thus the cells are embedded in a matrix without immediate contact to artificial biomaterials (Fig. 5B, C). DNAm profiles of cells expanded for 4 passages on hPL-gel did not reveal significant differences in comparison to those expanded on TCP. Alternatively, we used the cutoff of 20% differential methylation: 149 CpG sites were more than 20% hypermethylated and 106 CpG sites were more than 20% hypomethylated on hPL-gel (Fig. 5D; Supplemental Table 2) demonstrating that there was more variation than on PDMS. Differentially methylated CpGs were particularly associated with homeobox genes such as HOXB3, HOXB6, HOXB7, HOXC10 and HOXD3. In fact, the HOX clusters reflected some coherent DNAm changes in neighboring CpGs upon culture on hPLgel versus TCP indicating that this pattern might be regulated (Supplemental Fig. 5). Gene ontology analysis of genes with differentially methylated CpGs revealed significant enrichment in GO terms for cartilage development and skeletal system morphogenesis (Fig. 5E). Furthermore, senescence-associated DNAm changes were slightly enhanced in MSCs cultured on hPL-gel as compared to TCP (Fig. 5F). This moderate effect can be attributed to increased proliferation and more population doublings on hPL-gel.
Overall, even on very soft hPL-gel the DNAm profiles of MSCs were hardly different to those cultured on TCP - despite the different culture methods and growth patterns throughout cultureexpansion. 4. Discussion It is commonly accepted that matrix elasticity has major impact on cellular function. So far, this has been particularly addressed by seeding of cells on different substrates and functional comparison whilst on the substrate [5]. Here, we describe that elasticity of PDMS has no sustained effect on replicative senescence, cellintrinsic lineage commitment, or DNAm profiles of cells which are continuously cultured on these substrates. Tissue culture plastic is biofunctionalized by plasma treatment to facilitate better cell attachment and this may contribute to the higher CFU-f frequency in comparison to PDMS. Plasma treatment of PDMS would vitrify the surface and distort the elasticity moduli and therefore hydrophobic PDMS substrates were pre-coated with fibronectin [8]. Furthermore, adsorption of serum proteins e such as fibrin e notoriously takes place during cell culture. Different response of such matrix proteins to the different PDMS substrates may contribute to differences in cell adhesion [43]. We did not observe sharp folds in the gel surface, as described for polyacrylamide substrates after swelling [9]. Absence of topographical features on PDMS may therefore contribute to discrepancies to previous described studies on polyacrylamide gels [5,15].
Fig. 5. DNA methylation profiles on MSCs on human platelet lysate gel. (A) Histological sections of MSCs cultured for 4 days on hPL-gel (10% hPL) reveal multilayered growth at the interface to the culture medium (also with 10% hPL; size bar ¼ 100 mm). (B) Scanning electron microscopy (SEM) of the reticular fibrin scaffold in hPL-gel (size bar ¼ 2 mm). (C) SEM image of MSCs cultured on hPL-gel (size bar ¼ 20 mm). (D) DNAm profiles were compared in four individual MSC preparations which were cultured in parallel either on tissue culture plastic or hPL-gel (up to passage 4). Scatterplot analysis reveals mean DNAm levels: 149 CpGs are >20% hypermethylated on hPL-gel (red); 106 CpGs are > 20% hypomethylated on hPL-gel (green). (E) The corresponding genes to these CpGs were classified by Gene ontology categories. (F) Alternatively, we have highlighted CpGs with either senescence-associated hypermethylation (red), or hypomethylation (green) upon long-term culture in MSCs [25]. Overall, there is only a very moderate effect of the substrates on senescence-associated DNAm which might be attributed to higher proliferation on hPL-gel. (For interpretation of the references to color in this figure legend, the reader is referred to the web version of this article.)
Please cite this article in press as: Schellenberg A, et al., Matrix elasticity, replicative senescence and DNA methylation patterns of mesenchymal stem cells, Biomaterials (2014), http://dx.doi.org/10.1016/j.biomaterials.2014.04.079
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We demonstrate that substrate elasticity does not affect replicative senescence. Other authors have demonstrated that decreased hydrophobic properties of poly(ethylene-co-vinyl alcohol) impair long-term culture and induces typical senescence-associated phenotypic changes in fibroblasts but this effect was compensated by absorption of ECM proteins to the biomaterials [44]. Furthermore, proliferation of adherent cells increases with higher substrate stiffness and may even recruit quiescent cells into cell cycle [8,15,16]. We observed a moderate growth-stimulatory effect on rigid PDMS substrates whereas the maximal number of cumulative population doublings did not differ in MSCs continuously culture expanded on the different substrate elasticities. Notably, senescent cells were smaller and did not reveal the typical “fried egg” morphology on soft PDMS. This is in line with previous reports that cells grown on soft matrix have smaller cell size [45] and reduced spreading [15,45,46]. Either way, the very high stiffness of TCP, which may entail alterations of nuclear morphology [26], is not the driving force of replicative senescence. Differentiation of MSCs is influenced by the elasticity of the substrate they reside on [5,7,8,41]. In analogy to previous studies we observed that adipogenic differentiation was increased on soft substrates corresponding to the elastic modulus of adipose tissue [47]. In contrast, osteogenic differentiation was more pronounced on stiffer substrates mimicking the elastic modulus of bone [48] and it was associated with increased stress fiber formation [49,50]. Higher stiffness may provide better anchorage for these stress fibers and thereby support osteogenic differentiation. However, the propensity for differentiation towards adipogenic or osteogenic lineage was not maintained when reseeded on TCP. These results indicate that matrix elasticity supports differentiation only while the cells are in contact to the substrate. Biomaterials have impact on gene expression of MSCs [51e54] and it has been suggested that matrix elasticity up-regulates particularly lineages-specific genes [5]. These effects were not recapitulated in our study. In fact, no significant differences were observed in MSCs cultured on softer versus stiffer PDMS. Furthermore, only few genes were significantly differentially expressed on TCP as compared to PDMS e they might therefore rather be evoked by the different biomaterial than specifically by substrate elasticity. Many of the previous studies did not involve biological replica for sophisticated bioinformatics analysis of gene expression changes on different substrate elasticities. The discrepancy may also be due to creasing of polyacrylamide substrates [9]. Our results do not reveal up-regulation of lineage-specific gene expression by substrate elasticity and conversely this aspect deserves further analysis in other studies. Substrate properties can influence the epigenetic makeup: micropatterns, microgrooves and mechanical strain on substrate can modulate nuclear shape, HDAC activity, and histone acetylation [55]. Recently, Downing et al. reported that micro-grooved PDMS substrates induce pronounced changes in histone acetylation and methylation patterns in fibroblasts [56]. So far, effects of only matrix elasticity on the epigenetic makeup have not been analyzed. In this study, we demonstrate that DNAm profiles are not significantly influenced by the Young’s modulus of PDMS. Alternatively, we analyzed DNAm profiles of MSCs cultured on hPL-hydrogels. These hydrogels are very soft and facilitate multilayered cell growth at the interface of the fibrin network. Notably, even on hPL-gel none of the CpGs revealed significant DNAm changes. Several CpGs reached a threshold of at least 20% differential DNAm and these modifications can partially be attributed to senescence-associated DNAm changes e due to higher proliferation rates on hPL-gel. The high enrichment of DNAm changes in homeobox genes may suggest further impact on cellular differentiation [57-59] even though reproducibility was not high enough to reach statistical significance. Taken together,
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matrix elasticity did hardly induce reproducible DNAm changes in MSCs e other epigenetic modifications, such as the histone code, need to be further analyzed in future studies. 5. Conclusions Mesenchymal stem cells which are continuously cultured on the TCP or PDMS of different elasticity reveal very similar cell-intrinsic differentiation potential, gene expression profiles, and DNAm profiles. Furthermore, we demonstrate that substrate elasticity does not affect replicative senescence and it has relatively little impact on the DNAm profiles even on the very soft hPL-gel. This indicates that neither specific subpopulations are selected by the different biomaterials e which might be anticipated given to the heterogeneous composition of MSCs [60,61] e nor the cells are primed for specific lineages by matrix elasticity. In this regard, matrix elasticity needs to be taken into account for tissue engineering approaches where cells directly interact with biomaterials, but it appears to be is less relevant for cellular therapy with cells harvested from their substrates. Acknowledgments The authors would like to thank Sonja Hänzelmann (Computational Biology Lab, IZKF, RWTH Aachen University) for support in graphical presentation of gene expression results. This work was supported by the German Research Foundation (DFG; WA 1706/32), within the Boost-Fund Project “MechCell” of the excellence initiative of RWTH Aachen University, by the Else Kröner-Fresenius Stiftung and by the Stem Cell Network North Rhine Westphalia. Appendix A. Supplementary data Supplementary data related to this article can be found at http:// dx.doi.org/10.1016/j.biomaterials.2014.04.079. References [1] Hynes RO. The extracellular matrix: not just pretty fibrils. Science 2009;326: 1216e9. [2] Chai C, Leong KW. Biomaterials approach to expand and direct differentiation of stem cells. Mol Ther 2007;15:467e80. [3] Lim JY, Donahue HJ. Cell sensing and response to micro- and nanostructured surfaces produced by chemical and topographic patterning. Tissue Eng 2007;13:1879e91. [4] Hinz B. Matrix mechanics and regulation of the fibroblast phenotype. Periodontol 2000 2013;63:14e28. [5] Engler AJ, Sen S, Sweeney HL, Discher DE. Matrix elasticity directs stem cell lineage specification. Cell 2006;126:677e89. [6] Samani A, Bishop J, Luginbuhl C, Plewes DB. Measuring the elastic modulus of ex vivo small tissue samples. Phys Med Biol 2003;48:2183e98. [7] Sharma RI, Snedeker JG. Biochemical and biomechanical gradients for directed bone marrow stromal cell differentiation toward tendon and bone. Biomaterials 2010;31:7695e704. [8] Rowlands AS, George PA, Cooper-White JJ. Directing osteogenic and myogenic differentiation of MSCs: interplay of stiffness and adhesive ligand presentation. Am J Physiol Cell Physiol 2008;295:C1037e44. [9] Saha K, Kim J, Irwin E, Yoon J, Momin F, Trujillo V, et al. Surface creasing instability of soft polyacrylamide cell culture substrates. Biophys J 2010;99: L94e6. [10] Palchesko RN, Zhang L, Sun Y, Feinberg AW. Development of polydimethylsiloxane substrates with tunable elastic modulus to study cell mechanobiology in muscle and nerve. PLoS ONE 2012;7:e51499. [11] Rosenzweig DH, Matmati M, Khayat G, Chaudhry S, Hinz B, Quinn TM. Culture of primary bovine chondrocytes on a continuously expanding surface inhibits dedifferentiation. Tissue Eng Part A 2012;18:2466e76. [12] Dominici M, Le Blanc K, Mueller I, Slaper-Cortenbach I, Marini F, Krause D, et al. Minimal criteria for defining multipotent mesenchymal stromal cells. The international society for cellular therapy position statement. Cytotherapy 2006;8:315e7. [13] Horwitz EM, Le Blanc K, Dominici M, Mueller I, Slaper-Cortenbach I, Marini FC, et al. Clarification of the nomenclature for MSC: the international society for cellular therapy position statement. Cytotherapy 2005;7:393e5.
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Matrix elasticity, replicative senescence and DNA methylation patterns of mesenchymal stem cells Anne Schellenberg a, Sylvia Joussen a, Kristin Moser a, b, Nico Hampe b, Nils Hersch b, Hatim Hemeda a, Jan Schnitker c, Bernd Denecke d, Qiong Lin a, e, Norbert Pallua f, Martin Zenke e, Rudolf Merkel b, Bernd Hoffmann b, Wolfgang Wagner a, * a Helmholtz-Institute for Biomedical Engineering, Stem Cell Biology and Cellular Engineering, RWTH Aachen University Medical School, Pauwelsstrasse 20, Aachen 52074, Germany b Institute of Complex Systems, ICS-7: Biomechanics, Forschungszentrum Jülich GmbH, Jülich 52425, Germany c Institute of Complex Systems, ICS-8: Bioelectronics, Forschungszentrum Jülich GmbH, Jülich 52425, Germany d Interdisciplinary Centre for Clinical Research (IZKF) Aachen, RWTH Aachen University Medical School, 52074 Aachen, Germany e Institute for Biomedical Technology e Cell Biology, RWTH Aachen University Medical School, Aachen, Germany f Department of Plastic and Reconstructive Surgery, RWTH Aachen University Medical School, 52074 Aachen, Germany
a r t i c l e i n f o
a b s t r a c t
Article history: Received 28 March 2014 Accepted 22 April 2014 Available online xxx
Matrix elasticity guides differentiation of mesenchymal stem cells (MSCs) but it is unclear if these effects are only transient e while the cells reside on the substrate e or if they reflect persistent lineage commitment. In this study, MSCs were continuously culture-expanded in parallel either on tissue culture plastic (TCP) or on polydimethylsiloxane (PDMS) gels of different elasticity to compare impact on replicative senescence, in vitro differentiation, gene expression, and DNA methylation (DNAm) profiles. The maximal number of cumulative population doublings was not affected by matrix elasticity. Differentiation towards adipogenic and osteogenic lineage was increased on soft and rigid biomaterials, respectively e but this propensity was no more evident if cells were transferred to TCP. Global gene expression profiles and DNAm profiles revealed relatively few differences in MSCs cultured on soft or rigid matrices. Furthermore, only moderate DNAm changes were observed upon culture on very soft hydrogels of human platelet lysate. Our results support the notion that matrix elasticity influences cellular behavior while the cells reside on the substrate, but it does not have major impact on cellintrinsic lineage determination, replicative senescence or DNAm patterns. Ó 2014 Elsevier Ltd. All rights reserved.
Keywords: Mesenchymal stem cells Elasticity Long-term culture Platelet lysate DNA-methylation Epigenetic
1. Introduction Cells are normally embedded in a complex microenvironment of specific topography and matrix elasticity [1e4]. The elastic modulus of extracellular matrix (ECM), also known as Young’s modulus, ranges from 0.1 kPa in brain, some few kPa in fat tissue, to several hundred MPa in calcified bone [5,6]. In contrast, in vitro cell culture is usually performed on very rigid polystyrene surfaces of several GPa. Various studies have indicated that substrate elasticity is an important regulator for cellular function and these were mostly performed on polyacrylamide hydrogels [5,7,8]. However,
* Corresponding author. Helmholtz-Institute for Biomedical Engineering, Stem Cell Biology and Cellular Engineering, RWTH Aachen University Medical School, Pauwelsstrasse 20, 52074 Aachen, Germany. Tel.: þ49 241 8088611. E-mail address: [email protected] (W. Wagner).
changes in topography upon surface creasing of polyacrylamide substrates can significantly influence cell behavior [9]. Alternatively, elastomers of cross-linked polydimethylsiloxane (PDMS), a viscoelastic organic polymer, display rather smooth surfaces [10]. PDMS is widely used in biomedical engineering and cell culture due to good biocompatibility, optical transparency and tunable stiffness e it is therefore a suitable substrate for mechanotransduction experiments [11]. Effects of matrix elasticity were often addressed in mesenchymal stem cells (MSCs). These cells comprise a multipotent subset capable of differentiation potential towards osteocytes, chondrocytes and adipocytes and they are concurrently tested in a wide variety of clinical trials [12,13]. It has been suggested that matrix elasticity alone can direct MSC faith: soft (0.1e1 kPa), intermediate (8e17 kPa), and rigid (>34 kPa) polyacrylamide hydrogels supported differentiation towards neurogenic, myogenic and osteogenic lineages, respectively [5]. MSCs reveal directed
http://dx.doi.org/10.1016/j.biomaterials.2014.04.079 0142-9612/Ó 2014 Elsevier Ltd. All rights reserved.
Please cite this article in press as: Schellenberg A, et al., Matrix elasticity, replicative senescence and DNA methylation patterns of mesenchymal stem cells, Biomaterials (2014), http://dx.doi.org/10.1016/j.biomaterials.2014.04.079
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migration towards stiffer substrates, called durotaxis [14]. Cell adhesion, cell spreading, and proliferation rates are significantly increased on rigid biomaterials [8,15,16]. It has even been suggested that stiffness gradients, rather than stiffness alone, might be crucial for regulation of MSC behavior [14]. This is of particular relevance for tissue engineering approaches with MSCs embedded in matrix or scaffolds of specific elasticity. If matrix elasticity also entails persistent lineage conversion it would be relevant for generation of tailored cell products, too. Cellular differentiation is reflected by the epigenetic makeup. DNA methylation (DNAm) is the best characterized epigenetic modification: cytosine guanine dinucleotides (CpGs) can be methylated at cytosine moieties and this governs chromatin structure and gene regulation [17]. It is yet unclear if matrix elasticity impacts on DNAm profiles e which would be expected if matrix elasticity induces persistent cell-intrinsic modifications. All primary cells enter replicative senescence after a certain number of cell divisions, the so called Hayflick-Limit [18]. This process is accompanied by large-spread morphology, loss of differentiation potential, and unequivocal proliferation arrest [19e 22]. We have demonstrated that long-term culture of MSCs is associated with highly reproducible modifications in the DNAm pattern, particularly in developmental genes [23,24]. Notably, these senescence-associated DNAm changes are reversed upon reprogramming of MSCs into induced pluripotent stem cells (iPSCs) e and iPSCs do not reveal replicative senescence while in pluripotent state [25]. Recently, it has been reported that tissue stiffness and stress increase lamin-A levels, which stabilizes the nucleus [26]. Furthermore, hypomethylation upon long-term culture is enriched in lamin-associated domains [27] (Hänzelmann S. et al., manuscript in preparation). Hence it might be speculated that substrate stiffness, which alters cellular and nuclear morphology, contributes to cellular senescence. In this study, we have continuously culture expanded MSCs on PDMS of different Young’s modulus until they reached replicative senescence. Thereby, we wanted to determine if matrix elasticity affects cellular aging, whether it induces persistent lineage commitment, and if this is also reflected on DNAm level. 2. Methods 2.1. Fabrication of elastomeric substrates Polydimethylsiloxan substrates (Sylgard 184 Silicone Elastomer Kit, Dow Corning, MI, USA) were generated by mixing base and cross linker at ratios of 70:1 (1.5 kPa), 60:1 (6.5 kPa), 50:1 (15 kPa), 40:1 (50 kPa) and 30:1 (100 kPa). The mixture was then degassed and 2 ml were added in wells of 6-well plates. Cross-linking was performed at 60 C for 16 h. Characterization of elastomer material properties were performed by stretching cylindrical test pieces and macroscopic indentation tests as described previously [28,29]. For generation of human platelet lysate (hPL) gels 5 platelet units were pooled as described before [30]. Culture medium with 10% hPL gelatinized in the absence of anticoagulants due to the plasma components. The cells were then seeded on these hPL-gels in the same culture medium with 10% hPL supplemented with heparin (1 IU/ml) [31]. For passaging, MSCs were harvested together with the hPL-gel by pipetting, diluted with culture medium, and then plated onto new plates with hPLgel as described previously [30]. To determine cell numbers, fractions of the cell suspension were plated on TCP and conventionally counted upon adhesion to TCP (within 1 day). 2.2. Isolation and culture of MSCs Mesenchymal stem cells were isolated from lipoaspirates after patient’s written consent using guidelines approved by the Ethic Committee of the RWTH Aachen University Medical School (Permit Number EK163/07). Lipoaspirates were digested with 2 g/L collagenase type I (PAA, Pasching, Austria) [32] and seeded in culture medium consisting of Dulbecco’s Modified Eagles Medium-Low Glucose (DMEM-LG; PAA) with 2 mM L-glutamine (Sigma Aldrich, St. Louis, MO, USA), 100 U/mL penicillin/ streptomycin (pen/strep; Lonza, Basel, Switzerland) and 10% hPL [33]. Prior to cell seeding PDMS substrates were coated with 2.5 mg/cm2 fibronectin (Biochrom Berlin, Germany) for 45 min at 37 C. Cultures were maintained at 37 C in a humidified atmosphere containing 5% CO2 with medium changes twice per week. Fibroblastoid
colony-forming unit (CFU-f)-frequencies were determined 14 days after seeding of MSCs using crystal violet staining [32]. Upon 80% confluent growth MSCs were harvested from the substrates (either TCP or PDMS) using trypsin, counted with a Neubauer chamber (Brand, Wertheim, Germany), and re-seeded at a density of 10,000 cells/cm2 in 6-well-plates (Nunc Thermo Fisher Scientific, Langenselbold, Germany). Cell population doublings were analyzed for each passage and cumulative population doublings (cPD) were calculated from the first passage onward as described before [32]. 2.3. Characterization of MSCs Surface marker expression was analyzed on a FACSCanto II (Beckton Dickinson, BD) upon staining with the following antibodies as described before [32]: CD14allophycocyanin (APC, clone M5E2, BD), CD29-phycoerythrin (PE, clone MAR4, BD), CD31-PE (clone WM59, BD), CD34-APC (clone 8G12, BD), CD45-APC (clone HI30, BD), CD73-PE (clone AD2, BD), CD90-APC (clone 5E10, BD), CD105-fluorescein isothiocyanate (FITC, clone MEM-226 Immuno Tools). Osteogenic and adipogenic differentiation potential of MSCs was assessed in parallel on the different substrates as described before [22,34]. Alternatively, some of the cells were harvested from the PDMS substrates and cultured on TCP for two passages before induction of differentiation. Osteogenic differentiation was estimated by Alizarin Red staining and quantified with a Tecan infinite M200 platereader (405 nm) [35]. Adipogenic differentiation was estimated by counting cells with lipid-droplets in phase contrast images from five randomly chosen areas (Lipophilic dyes could not be used on PDMS) with counter staining with DAPI (40 ,6Diamidin-2-phenylindol; both Molecular Probes, Eugene, Oregon, USA) [32,36]. Images were captured using a Leica DM IL LED microscope (Leica, Wetzlar, Germany). 2.4. Analysis of senescence-associated b-galactosidase Expression of pH-dependent senescence associated b-galactosidase (SA-b-gal) activity was analyzed by flow cytometry as described before [37]: MSCs were pretreated with Bafilomycin A1 (Sigma, St Louis, MO, USA) and 5dodecanoylaminofluorescein di-b-D-galactopyranoside (C12FDG, Invitrogen, Eugene, OR, USA) was subsequently used as fluorogenic substrate for bgalactosidase. 2.5. Apoptosis assay Apoptosis was estimated by flow cytometry using Annexin V and propidium iodid (PI) double staining. In brief, MSCs were harvested from the substrates, washed twice in phosphate buffered saline (PBS), and 1 105 cells were stained in 5 ml Annexin V with 100 ml binding buffer (Becton Dickinson [BD], San Jose, USA) for 15 min at RT in the dark. Subsequently, 400 ml Annexin V binding puffer (BD) and 2 ml PI were added prior to analysis on a FACSCanto II (BD). 2.6. Immunofluorescence and scanning electron microscopy MSCs were grown on glass or PDMS substrates for 8 days, fixed using 3.7% formaldehyde (Merck, Germany) and stained for vinculin and actin expression as previously described [38]. For scanning electron microscopy of cells on PDMS, 5000 cells were grown on cross-linked PDMS substrates exhibiting elasticities of 1.5 and 100 kPa, respectively. After 24 h of growth cells were treated as described before [39]. Substrates were sputtered with a thin layer of gold and analyzed by a LEO-1550 scanning electron microscope (Zeiss, Germany). Scanning electron microscopy of hPL gels was performed as described before [30]. 2.7. Gene expression analysis Gene expression profiles were analyzed in three MSC preparations (always at passage 4) upon continuous culture-expansion on PDMS (1.5 kPa or 50 kPa) or TCP. RNA was isolated from 106 cells using the NucleoSpin RNA Isolation Kit (Macherey Nagel, Düren, Germany) and quality control was carried out using the Agilent 2100 Bioanalyzer (Agilent Technologies, Inc., Santa Clara, CA, USA). RNA was amplified and hybridized to the GeneChip Human Gene 1.0 ST Arrays (Affymetrix, Santa Clara, CA, USA) according to the manufacturer’s instructions. Raw data were normalized by RMA (Affymetrix Power Tools). The microarray data are accessible in NCBIs Gene Expression Omnibus (GEO, http://www.ncbi.nlm.nih.gov/geo/) through accession number GSE55867. Pairwise limma-T-test analysis was calculated in R to select for differentially expressed genes (adjusted p-value < 0.05). Quantitative real time PCR (RT-qPCR) of selected genes was performed using the High Capacity cDNA Reverse Transcription Kit and a StepOnePlus instrument (both Applied Biosystems, Carlsbad, CA, USA) in a MicroAmp optical 96-well reaction plate. TaqMan Gene Expression Assays (Applied Biosystems) used in this study are listed in Supplemental Table 1: Gene expression levels were normalized to GAPDH expression. 2.8. DNA-methylation profiling DNA methylation profiles were analyzed in four MSC preparations (always at passage 4) upon continuous culture-expansion on PDMS (1.5 kPa or 50 kPa) or TCP.
Please cite this article in press as: Schellenberg A, et al., Matrix elasticity, replicative senescence and DNA methylation patterns of mesenchymal stem cells, Biomaterials (2014), http://dx.doi.org/10.1016/j.biomaterials.2014.04.079
A. Schellenberg et al. / Biomaterials xxx (2014) 1e8 Furthermore, we compared DNAm profiles of 4 MSC preparations (passage 4) cultured either on TCP or hPL-gel. Genomic DNA was isolated from 106 cell using the QIAamp DNA Blood Midi Kit (Qiagen, Hilden, Germany) and subsequently bisulfiteconverted (EZ DNA MethylationTM Kit, Zymo, Irvine, USA). DNAm profiles were analyzed using the HumanMethylation450 BeadChip (Illumina, San Diego, USA) as described before [40]. In brief, about 200 ng of bisulfite converted DNA was hybridized according to the manufacturer’s instructions. After single-base extension using DNP- and Biotin-labeled ddNTPs, the array was fluorescently stained, scanned, and the intensities of non-methylated and methylated bead types measured. Hybridization and initial data analysis with the BeadStudio Methylation Module was performed at the DKFZ Gene Core Facility (Heidelberg, Germany). The complete dataset is accessible through GEO Series accession number GSE55888. Hierarchical clustering of DNAm profiles was performed using the MultiExperiment Viewer (MeV, TM4; Euclidian distance). Pairwise limma-T-test analysis was calculated in R to select for CpGs with significant different DNAm levels (adjusted p-value < 0.05). Under these parameters no CpGs reached the significance threshold and therefore we alternatively selected CpGs with at least 20% differential DNAm in MSCs isolated on different substrates. Genes associated with the differentially methylated CpG sites were classified by Gene Ontology analysis using DAVID Bioinformatics Resources 6.7 (http://david.abcc.ncifcrf.gov/home.jsp). 2.9. Statistics Results are expressed as mean standard deviation (SD) of at least three independent experiments. To estimate the probability of differences we have adopted the paired two-sided Student’s T-test.
3. Results 3.1. Elasticity and replicative senescence of MSCs Mesenchymal stem cells were isolated from adipose tissue and continuously cultured in parallel either on TCP or on PDMS of different stiffness (1.5 kPa, 6.5 kPa, 15 kPa, 50 kPa and 100 kPa). Scanning electron microscopy revealed that PDMS substrates were
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evenly flat and cells attached on this substrate (Fig. 1A). Initial colony formation was higher on TCP than on PDMS (Fig. 1B). After 7 days (passage 0) all cell preparations displayed the typical immunophenotypic pattern of MSCs (CD14-, CD29þ, CD31, CD34þ/-, CD45, CD73þ, CD90þ, CD105þ and CD140; Supplemental Fig. 1), regardless of the substrate they were isolated on. Proliferation rate was slightly higher on TCP and there was no significant difference in the long-term growth curves (Fig. 1C): MSCs continuously expanded in parallel on TCP, PDMS 1.5 kPa, or PDMS 50 kPa entered replicative senescence within 90 days with a similar maximal number of cumulative population doublings (cPDs; Fig. 1C). However, senescent cells on elastic substrates did not reveal typical large “fried-egg” morphology and remained overall smaller (Fig 1D; Supplemental Fig. 2). Furthermore, neither flow cytometric measurement of senescence associated b-galactosidase (SA-b-gal) activity (Fig. 1E, F), nor apoptosis rates (Fig. 1G) revealed significant differences upon expansion on the various substrates. Thus, matrix elasticity affects initial outgrowth and morphology of MSCs but it does not impact on replicative senescence. 3.2. Impact of elasticity on in vitro differentiation MSCs culture expanded on the different substrate elasticities were differentiated towards osteogenic or adipogenic lineage: osteogenic differentiation, reflected by calcium phosphate depositions in Alizarin Red staining after 14 days, markedly increased with substrate stiffness (Fig. 2A); whereas adipogenic differentiation, estimated by the percentage of cells with lipid droplets, was more pronounced on softer PDMS substrates (Fig. 2B). These tendencies have also been described in previous studies [5,8,15,41].
Fig. 1. Long-term culture of MSCs on elastic PDMS substrates. (A) Scanning electron microscopy of MSCs cultured on PDMS 1.5 kPa (scale bar 20 mm). (B) Initial fibroblastoid colony forming unit (CFU-f) frequency of adipose tissue derived MSCs was higher on tissue culture plastic (TCP) than on PDMS substrates (n ¼ 5; *p < 0.05). (C) MSCs of 3 donors were continuously culture expanded on soft PDMS (1.5 kPa), stiffer PDMS (50 kPa) and rigid TCP until replicative senescence. Cumulative population doublings (cPDs) were calculated from the first passage onward (transient inclination of proliferation after 30 d might be due to change of hPL supplements). (D) No significant differences were observed in the maximum number of cPDs. (E) Cells of late passages on elastic substrates did not show the typical “fried egg morphology” of senescent cells (scale bar 100 mm). (F, G) However, flow cytometric analysis of senescence-associated b-galactosidase by C12FDG was not affected by the biomaterials (n ¼ 3). (H) Analysis of apoptotic cells by flow cytometric analysis of Annexin V did not reveal any differences, too (n ¼ 3).
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Fig. 2. Transient effect of substrate elasticity on differentiation of MSCs. (A) Osteogenic differentiation was induced for two weeks in MSCs cultured on either tissue culture plastic (TCP) or on cross-linked PDMS substrates with indicated Young’s moduli. Semiquantitative analysis of calcium phosphate precipitates with a plate reader upon Alizarin red staining revealed increased differentiation with substrate stiffness (*p < 0.05). (B) In contrast, adipogenic differentiation, determined by the percentage of cells containing lipid-droplets, was increased on softer matrices (*p < 0.05). (C) Staining for actin (green) and vinculin (red) in MSCs on PDMS substrates and glass. If indicated, cells were additionally differentiated towards osteogenic and adipogenic lineages for two weeks (size bar ¼ 50 mm). (D, F) MSCs which were continuously cultured on the substrates as indicated were re-seeded on TCP before induction of in vitro differentiation e thereby the above mentioned propensity of lineage-specific differentiation was lost. (For interpretation of the references to color in this figure legend, the reader is referred to the web version of this article.)
Immunostaining of vinculin and actin revealed heterogeneous cell morphologies on all elasticities. Upon osteogenic differentiation the cells enlarged with more stress fibers which could be visualized on all substrates. In contrast adipogenic differentiation resulted in rather round cellular morphology with less pronounced actin structures and accumulation of lipid droplets was particularly observed on soft substrates (Fig. 2C). Subsequently, MSCs were harvested from the different substrates for two additional passages on TCP. Now the initially observed propensities for in vitro differentiation e osteogenic differentiation on stiff and adipogenic differentiation on soft matrices e were not evident any more (Fig. 2D, F). Thus matrix elasticity appears to affect in vitro differentiation only while the cells are on the matrix, but it does not result in cellintrinsic fate conversion which remains upon passaging on other substrate elasticities. 3.3. Effects of elasticity on gene expression profiles Global gene expression profiles were compared in MSCs continuously cultured on different substrates (TCP, PDMS 1.5 kPa, PDMS 50 kPa; all passage 4). Hierarchical cluster analysis revealed close relationship of MSC preparations derived from the same donor (Fig. 3A). However, gene expression profiles of MSCs cultured on PDMS 1.5 kPa versus PDMS 50 kPa did not reveal any significant differences (adjusted limma paired-T-test < 0.05). Only nine genes were differentially expressed in MSCs expanded on PDMS 1.5 kPa versus TCP (Fig 3B): sulfatase 1 (SULF1), fibronectin type III domain containing 1 (FNDC1), retinoic acid receptor
responder 2 (RARRES2), myosin heavy chain 11 (MYH11), and vascular cell adhesion protein 1 (VCAM1) were higher expressed on TCP, whereas ATP-binding cassette C3 (ABCC3), chemokine C-X-C motif ligand 1 (CXCL1), ribosomal protein SA pseudogene 52 (RPSAP52), and interleukin 8 (IL-8) were higher expressed on PDMS 1.5 kPa. Furthermore, only seven genes were differential expressed in MSCs expanded on PDMS 50 kPa versus TCP (Fig. 3C): SULF1, FNDC1, RARRES2, MYH11, lymphoid enhancer-binding factor-1 (LEF1), and thymocyte expressed, positive selection associated 1 (KIAA0748) are higher expressed on TCP, whereas ABCC3 was higher expressed on PDMS 50 kPa. Differential expression was confirmed by RT-qPCR (Fig. 3D, E). These results were somewhat unexpected, as previous studies indicated substrate elasticity has major impact on gene expression profiles [5]. We have subsequently focused on the described lineage-specific gene expression changes but the effects of matrix elasticity were not recapitulated in our data (Supplemental Fig. 3). 3.4. Effects of elasticity on DNA-methylation profiles Subsequently, we analyzed DNAm profiles of MSCs expanded on different substrates (TCP, PDMS 1.5 kPa, or PDMS 50 kPa) using the Illumina HumanMethylation450 platform which assays more than 480,000 CpG sites at single base resolution (covering 99% of RefSeq genes and 96% of CpG islands) [40]. Hierarchical cluster analysis of DNAm profiles demonstrated donor-specific differences as described before [42] (Fig. 4A). MSCs did not cluster according to the substrate elasticities and this was also observed in principal
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Fig. 3. Gene expression profiles of MSCs on different substrates. (A) Gene expression profiles of MSCs cultured for 4 passages in parallel on PDMS substrates (1.5 kPa or 50 kPa) or tissue culture plastic (TCP) were analyzed on Affymetrix 1.0 ST microarrays. The three individual donor samples clustered closely together. (B) Nine genes with differential gene expression in MSCs on PDMS 1.5 kPa versus TCP, (C) and seven genes with differential expression on PDMS 50 kPa versus TCP are depicted (Limma paired t-test, adjusted pvalue < 0.05). (D, E) Differential expression of these genes was validated by RT-qPCR.
component analysis (PCA, not shown). Significance Analysis of Microarray (SAM) or limma paired T-test did not reveal any CpG with differential DNAm between the growth conditions (adjusted p-value < 0.05). Alternatively, we focused on the four CpGs with more than 20% difference in DNAm level. Culture of MSCs on stiffer substrates resulted in hypermethylation of two CpGs associated with the homeobox genes HOXA5 and HOXC4 (Fig. 4BeD). However, these differences were neither significant, nor were they correlated
with differential gene expression. Alternatively, we have focused on CpGs which reflect senescence-associated DNAm changes in MSCs (1702 CpGs with >20% hypermethylated; 2116 CpGs with >20% hypomethylation in MSCs of late passage [25]), but there was no association with DNAm patterns of MSCs expanded on different substrate elasticities (Supplemental Fig. 4). Overall, elasticity therefore does not evoke significant differences in global DNAm profiles.
Fig. 4. DNA methylation profiles of MSCs on elastic substrates. (A) DNAm profiles of MSCs cultured in parallel on PDMS 1.5 kPa, PDMS 50 kPa and TCP (passage 4) were compared by HumanMethylation450 BeadChips. Hierachical clustering revealed separation according to the four donor samples. (B, C, D) Scatterplots depict CpGs with more than 20% differential methylation between MSCs cultured on the different substrates and the corresponding gene IDs are highlighted. None of these CpGs revealed significant DNAm changes (adjusted pvalue < 0.05).
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3.5. DNA-methylation patterns on platelet lysate-hydrogel To further verify the effects of substrate elasticity on DNAm we performed additional experiments with MSCs cultured on hPL-gel. These hydrogels consist of a fibrin-network which forms in the absence of anticoagulants and possess an extremely low Young’s modulus of about 30 Pa [30] (Fig. 5A). Notably, hPL-gel consists of the same components as the over-layered culture medium, apart from heparin, and thus the cells are embedded in a matrix without immediate contact to artificial biomaterials (Fig. 5B, C). DNAm profiles of cells expanded for 4 passages on hPL-gel did not reveal significant differences in comparison to those expanded on TCP. Alternatively, we used the cutoff of 20% differential methylation: 149 CpG sites were more than 20% hypermethylated and 106 CpG sites were more than 20% hypomethylated on hPL-gel (Fig. 5D; Supplemental Table 2) demonstrating that there was more variation than on PDMS. Differentially methylated CpGs were particularly associated with homeobox genes such as HOXB3, HOXB6, HOXB7, HOXC10 and HOXD3. In fact, the HOX clusters reflected some coherent DNAm changes in neighboring CpGs upon culture on hPLgel versus TCP indicating that this pattern might be regulated (Supplemental Fig. 5). Gene ontology analysis of genes with differentially methylated CpGs revealed significant enrichment in GO terms for cartilage development and skeletal system morphogenesis (Fig. 5E). Furthermore, senescence-associated DNAm changes were slightly enhanced in MSCs cultured on hPL-gel as compared to TCP (Fig. 5F). This moderate effect can be attributed to increased proliferation and more population doublings on hPL-gel.
Overall, even on very soft hPL-gel the DNAm profiles of MSCs were hardly different to those cultured on TCP - despite the different culture methods and growth patterns throughout cultureexpansion. 4. Discussion It is commonly accepted that matrix elasticity has major impact on cellular function. So far, this has been particularly addressed by seeding of cells on different substrates and functional comparison whilst on the substrate [5]. Here, we describe that elasticity of PDMS has no sustained effect on replicative senescence, cellintrinsic lineage commitment, or DNAm profiles of cells which are continuously cultured on these substrates. Tissue culture plastic is biofunctionalized by plasma treatment to facilitate better cell attachment and this may contribute to the higher CFU-f frequency in comparison to PDMS. Plasma treatment of PDMS would vitrify the surface and distort the elasticity moduli and therefore hydrophobic PDMS substrates were pre-coated with fibronectin [8]. Furthermore, adsorption of serum proteins e such as fibrin e notoriously takes place during cell culture. Different response of such matrix proteins to the different PDMS substrates may contribute to differences in cell adhesion [43]. We did not observe sharp folds in the gel surface, as described for polyacrylamide substrates after swelling [9]. Absence of topographical features on PDMS may therefore contribute to discrepancies to previous described studies on polyacrylamide gels [5,15].
Fig. 5. DNA methylation profiles on MSCs on human platelet lysate gel. (A) Histological sections of MSCs cultured for 4 days on hPL-gel (10% hPL) reveal multilayered growth at the interface to the culture medium (also with 10% hPL; size bar ¼ 100 mm). (B) Scanning electron microscopy (SEM) of the reticular fibrin scaffold in hPL-gel (size bar ¼ 2 mm). (C) SEM image of MSCs cultured on hPL-gel (size bar ¼ 20 mm). (D) DNAm profiles were compared in four individual MSC preparations which were cultured in parallel either on tissue culture plastic or hPL-gel (up to passage 4). Scatterplot analysis reveals mean DNAm levels: 149 CpGs are >20% hypermethylated on hPL-gel (red); 106 CpGs are > 20% hypomethylated on hPL-gel (green). (E) The corresponding genes to these CpGs were classified by Gene ontology categories. (F) Alternatively, we have highlighted CpGs with either senescence-associated hypermethylation (red), or hypomethylation (green) upon long-term culture in MSCs [25]. Overall, there is only a very moderate effect of the substrates on senescence-associated DNAm which might be attributed to higher proliferation on hPL-gel. (For interpretation of the references to color in this figure legend, the reader is referred to the web version of this article.)
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We demonstrate that substrate elasticity does not affect replicative senescence. Other authors have demonstrated that decreased hydrophobic properties of poly(ethylene-co-vinyl alcohol) impair long-term culture and induces typical senescence-associated phenotypic changes in fibroblasts but this effect was compensated by absorption of ECM proteins to the biomaterials [44]. Furthermore, proliferation of adherent cells increases with higher substrate stiffness and may even recruit quiescent cells into cell cycle [8,15,16]. We observed a moderate growth-stimulatory effect on rigid PDMS substrates whereas the maximal number of cumulative population doublings did not differ in MSCs continuously culture expanded on the different substrate elasticities. Notably, senescent cells were smaller and did not reveal the typical “fried egg” morphology on soft PDMS. This is in line with previous reports that cells grown on soft matrix have smaller cell size [45] and reduced spreading [15,45,46]. Either way, the very high stiffness of TCP, which may entail alterations of nuclear morphology [26], is not the driving force of replicative senescence. Differentiation of MSCs is influenced by the elasticity of the substrate they reside on [5,7,8,41]. In analogy to previous studies we observed that adipogenic differentiation was increased on soft substrates corresponding to the elastic modulus of adipose tissue [47]. In contrast, osteogenic differentiation was more pronounced on stiffer substrates mimicking the elastic modulus of bone [48] and it was associated with increased stress fiber formation [49,50]. Higher stiffness may provide better anchorage for these stress fibers and thereby support osteogenic differentiation. However, the propensity for differentiation towards adipogenic or osteogenic lineage was not maintained when reseeded on TCP. These results indicate that matrix elasticity supports differentiation only while the cells are in contact to the substrate. Biomaterials have impact on gene expression of MSCs [51e54] and it has been suggested that matrix elasticity up-regulates particularly lineages-specific genes [5]. These effects were not recapitulated in our study. In fact, no significant differences were observed in MSCs cultured on softer versus stiffer PDMS. Furthermore, only few genes were significantly differentially expressed on TCP as compared to PDMS e they might therefore rather be evoked by the different biomaterial than specifically by substrate elasticity. Many of the previous studies did not involve biological replica for sophisticated bioinformatics analysis of gene expression changes on different substrate elasticities. The discrepancy may also be due to creasing of polyacrylamide substrates [9]. Our results do not reveal up-regulation of lineage-specific gene expression by substrate elasticity and conversely this aspect deserves further analysis in other studies. Substrate properties can influence the epigenetic makeup: micropatterns, microgrooves and mechanical strain on substrate can modulate nuclear shape, HDAC activity, and histone acetylation [55]. Recently, Downing et al. reported that micro-grooved PDMS substrates induce pronounced changes in histone acetylation and methylation patterns in fibroblasts [56]. So far, effects of only matrix elasticity on the epigenetic makeup have not been analyzed. In this study, we demonstrate that DNAm profiles are not significantly influenced by the Young’s modulus of PDMS. Alternatively, we analyzed DNAm profiles of MSCs cultured on hPL-hydrogels. These hydrogels are very soft and facilitate multilayered cell growth at the interface of the fibrin network. Notably, even on hPL-gel none of the CpGs revealed significant DNAm changes. Several CpGs reached a threshold of at least 20% differential DNAm and these modifications can partially be attributed to senescence-associated DNAm changes e due to higher proliferation rates on hPL-gel. The high enrichment of DNAm changes in homeobox genes may suggest further impact on cellular differentiation [57-59] even though reproducibility was not high enough to reach statistical significance. Taken together,
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matrix elasticity did hardly induce reproducible DNAm changes in MSCs e other epigenetic modifications, such as the histone code, need to be further analyzed in future studies. 5. Conclusions Mesenchymal stem cells which are continuously cultured on the TCP or PDMS of different elasticity reveal very similar cell-intrinsic differentiation potential, gene expression profiles, and DNAm profiles. Furthermore, we demonstrate that substrate elasticity does not affect replicative senescence and it has relatively little impact on the DNAm profiles even on the very soft hPL-gel. This indicates that neither specific subpopulations are selected by the different biomaterials e which might be anticipated given to the heterogeneous composition of MSCs [60,61] e nor the cells are primed for specific lineages by matrix elasticity. In this regard, matrix elasticity needs to be taken into account for tissue engineering approaches where cells directly interact with biomaterials, but it appears to be is less relevant for cellular therapy with cells harvested from their substrates. Acknowledgments The authors would like to thank Sonja Hänzelmann (Computational Biology Lab, IZKF, RWTH Aachen University) for support in graphical presentation of gene expression results. This work was supported by the German Research Foundation (DFG; WA 1706/32), within the Boost-Fund Project “MechCell” of the excellence initiative of RWTH Aachen University, by the Else Kröner-Fresenius Stiftung and by the Stem Cell Network North Rhine Westphalia. Appendix A. Supplementary data Supplementary data related to this article can be found at http:// dx.doi.org/10.1016/j.biomaterials.2014.04.079. References [1] Hynes RO. The extracellular matrix: not just pretty fibrils. Science 2009;326: 1216e9. [2] Chai C, Leong KW. Biomaterials approach to expand and direct differentiation of stem cells. Mol Ther 2007;15:467e80. [3] Lim JY, Donahue HJ. Cell sensing and response to micro- and nanostructured surfaces produced by chemical and topographic patterning. Tissue Eng 2007;13:1879e91. [4] Hinz B. Matrix mechanics and regulation of the fibroblast phenotype. Periodontol 2000 2013;63:14e28. [5] Engler AJ, Sen S, Sweeney HL, Discher DE. Matrix elasticity directs stem cell lineage specification. Cell 2006;126:677e89. [6] Samani A, Bishop J, Luginbuhl C, Plewes DB. Measuring the elastic modulus of ex vivo small tissue samples. Phys Med Biol 2003;48:2183e98. [7] Sharma RI, Snedeker JG. Biochemical and biomechanical gradients for directed bone marrow stromal cell differentiation toward tendon and bone. Biomaterials 2010;31:7695e704. [8] Rowlands AS, George PA, Cooper-White JJ. Directing osteogenic and myogenic differentiation of MSCs: interplay of stiffness and adhesive ligand presentation. Am J Physiol Cell Physiol 2008;295:C1037e44. [9] Saha K, Kim J, Irwin E, Yoon J, Momin F, Trujillo V, et al. Surface creasing instability of soft polyacrylamide cell culture substrates. Biophys J 2010;99: L94e6. [10] Palchesko RN, Zhang L, Sun Y, Feinberg AW. Development of polydimethylsiloxane substrates with tunable elastic modulus to study cell mechanobiology in muscle and nerve. PLoS ONE 2012;7:e51499. [11] Rosenzweig DH, Matmati M, Khayat G, Chaudhry S, Hinz B, Quinn TM. Culture of primary bovine chondrocytes on a continuously expanding surface inhibits dedifferentiation. Tissue Eng Part A 2012;18:2466e76. [12] Dominici M, Le Blanc K, Mueller I, Slaper-Cortenbach I, Marini F, Krause D, et al. Minimal criteria for defining multipotent mesenchymal stromal cells. The international society for cellular therapy position statement. Cytotherapy 2006;8:315e7. [13] Horwitz EM, Le Blanc K, Dominici M, Mueller I, Slaper-Cortenbach I, Marini FC, et al. Clarification of the nomenclature for MSC: the international society for cellular therapy position statement. Cytotherapy 2005;7:393e5.
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Please cite this article in press as: Schellenberg A, et al., Matrix elasticity, replicative senescence and DNA methylation patterns of mesenchymal stem cells, Biomaterials (2014), http://dx.doi.org/10.1016/j.biomaterials.2014.04.079