A scaffold-free in vitro model for osteogenesis of human mesenchymal stem cells

Tissue and Cell 43 (2011) 91–100

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A scaffold-free in vitro model for osteogenesis of human mesenchymal stem cells Cornelia Hildebrandt, Heiko Büth, Hagen Thielecke ∗ Department of Biohybrid Systems, Fraunhofer IBMT, Ensheimerstrasse 48, 66386 St. Ingbert, Germany

a r t i c l e

i n f o

Article history: Received 24 March 2010 Received in revised form 21 December 2010 Accepted 27 December 2010 Available online 16 February 2011 Keywords: Mesenchymal stem cells Osteogenic differentiation 3D Spheroid

a b s t r a c t For studying cellular processes three-dimensional (3D) in vitro models are of a high importance. For tissue engineering approaches osseous differentiation is performed on 3D scaffolds, but material depending influences promote cellular processes like adhesion, proliferation and differentiation. To investigate developmental processes of mesenchymal stem cells without cell–substrate interactions, self-contained in vitro models mimicking physiological condition are required. However, with respect to scientific investigations and pharmaceutical tests, it is essential that these tissue models are well characterised and are of a high reproducibility. In order to establish an appropriate in vitro model for bone formation, different protocols are compared and optimised regarding their aggregate formation efficiency, homogeneity of the aggregates, the viability and their ability to induce differentiation into the osteogenic lineage. The protocols for the generation of 3D cell models are based on rotation culture, hanging drop technique, and the cultivation in non adhesive culture vessels (single vessels as well as 96 well plates). To conclude, the cultivation of hMSCs in 96 well non adhesive plates facilitates an easy way to cultivate homogenous cellular aggregates with high performance efficiency in parallel. The size can be controlled by the initial cell density per well and within this spheroids, bone formation has been induced. © 2011 Elsevier Ltd. All rights reserved.

1. Introduction In vivo hMSCs play an important role in tissue repair and regeneration processes of the adult organism. Since MSCs can be easily obtained from the bone marrow, and due to their selfrenewing capacity expanded in vitro (Owen and Friedenstein, 1988; Digirolamo et al., 1999; Colter et al., 2001), they are promising for cell-based therapies, tissue engineering approaches as well as for pharmaceutical research. hMSCs have the plasticity to differentiate multipotently into osteoblasts, chondrocytes, adipocytes, stromal cells (Pittenger et al., 1999), tenoblasts (Young et al., 1998) and the myogenic lineage (Wakitani et al., 1995; Dezawa et al., 2005). However, the exploitation of the potential of MSCs as an attractive cell source for cell-based technologies will depend on the ability to understand the factors that influence cell fate decisions to direct the development into a specific cell type. Unlike conventional monolayer or suspension cultures three-dimensional (3D) cell aggregates, or so called spheroids, comprise the complexity of tissues and mimic physiological conditions like cell–cell and cell–matrix interactions (Rossi et al., 2005; Wang et al., 2009). Such cell aggregates represent a complex 3D network displaying and controlling biological mechanisms such as proliferation, differentiation and apoptosis and having the potential of providing an insight

∗ Corresponding author. E-mail address: [email protected] (H. Thielecke). 0040-8166/$ – see front matter © 2011 Elsevier Ltd. All rights reserved. doi:10.1016/j.tice.2010.12.004

into the regulating signal gradients induced by specific bioactive substances and factors (Berrier and Yamada, 2007). For that purpose 3D cell-based aggregates as an improved and refined in vitro model representing a link between conventional cell models and whole organs became more important over the last decades. For example, cell-based aggregates are widely used in cancer research to screen the efficiency of radiation therapies (Wheldon, 1994) or the effects of various drugs (Kobayashi et al., 1993). Moreover, organotypic models were established based on hepatocytes (Miranda et al., 2009), pituitary gland cells for research on hormone release (Bael et al., 1995), embryonic chicken cardiomyocytes as a contractive heart muscle model (Bartholomä et al., 2005) and 3D in vitro models were established to investigate neural differentiation and degeneration (Chatterjee and Nöldner, 1994; Layer et al., 1992). To direct the differentiation of hMSCs into the osteogenic and chondrogenic lineages research is focused on the development of biocompatible and biodegradable scaffolds that act as shape and guidance templates for in vitro and in vivo tissue development (Ohgushi and Caplan, 1999; Risbud and Sittinger, 2002). Polymeric gels, for example hyaluronic acid (Kujawa and Caplan, 1986), collagen (Mueller and Glowacki, 2001), agarose (Huang et al., 2004), gelatin (Ponticiello et al., 2000), alginate and chitosan capsules (Pound et al., 2007) as well as poly(alpha-hydroxy esters) like PLGA (polylactic acid), PGA (polyglycolic acid) and copolymer PLGA (Banu et al., 2005; Zwingmann et al., 2007) have been used extensively to maintain the chondrogenic phenotype.

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However, for the generation of artificial bone substitutes for tissue engineering approaches scaffold containing collagen or phosphorous calcium substrates such as tricalcium phosphate and hydroxyapatite (Ohgushi et al., 1990), mineralized collagen (Niemeyer et al., 2004) or partially demineralized bone (Mauney et al., 2004) have shown osteoinductive properties (Sawyer et al., 2005). The support of the osteogenic development is not only constricted on osseous matrix compounds, but also macro and micro structured glass ceramics containing SiO2 , Na2 O, CaO, P2 O2 , Al2 O3 or titanium devices are common scaffolds to mediate bone formation in vitro and in vivo (Ohgushi et al., 2003; Ohgushi and Caplan, 1999). Beside this, a scaffold-free 3D culture technique, a so called micromass or pellet culture, for the differentiation of hMSCs into the chondrogenic lineage was previously described (Sekiya et al., 2002). Culturing hMSC after artificial condensation by treatment with chondrogenic factors leads to the generation of hyaline cartilage (Johnstone et al., 1998). Recent work has shown that it is possible to direct the osteogenic differentiation in such a scaffoldfree culture system (Muraglia et al., 2003). Differentiation in scaffold-free 3D aggregates supports not only the investigation of cellular organization and accompanying changes of gene expression patterns, but also provides new insights into specific cell physiological and pathological processes influenced by various biomolecules. Even if several protocols are known to induce the differentiation of bone marrow MSCs successfully, there is still a lack of knowledge of the signal transduction pathways releasing functional cellular reactions to specific factors or drugs. The major aim of this study consisted in the development of a 3D in vitro model system for the osteogenic differentiation of hMSCs free from non-physiological matrix compounds. Traditional techniques for the generation of cell-based spheroids, for example the suspension culture in spinner flask, the hanging drop technique, the cultivation on non adhesive surfaces or the encapsulation in hydro gel polymers, have drawbacks like a heterogeneous size of the aggregates and a low spheroid formation efficiency. Moreover, an additional challenge was the cultivation of vital spheroids over a long time period needed for the osteogenic differentiation. Within this investigation, several techniques for spheroid formation and cultivation have been optimized in order to establish a 3D in vitro model for osteoblast development of hMSCs. Here we present a rather straightforward, mild technique to generate homogenous hMSC aggregates with a high spheroid formation efficiency accomplishing the osteogenic differentiation successfully. 2. Materials and methods 2.1. Material All chemicals, fetal bovine serum (FBS), the leukocyte alkaline phosphatase staining kit (#86), pluronic (#P2443), methylcellulose (#274429) and poly-2-hydrxyethylmethlacrylate (poly-HEMA,

#P3932) were obtained from Sigma–Aldrich (Hamburg, Germany). The cell culture materials were all purchased from Greiner Bio-one GmbH (Frickenhausen, Germany). Phosphate buffered saline (PBS), alpha MEM (without desoxyribonucleotides or ribonucleotides, with GlutarMAX), penicillin/streptomycin, trypsin/EDTA solution and goat serum were obtained from Gibco (Invitrogen, Karlsruhe, Germany). The primary antibody anti-Collagen Type 1, clone Col-1, mouse IgG1 was purchased from Sigma–Aldrich (#C2456, Hamburg, Germany) and the secondary antibody Alexa Fluor dye 568 from Invitrogen (goat anti mouse, #A11031, Karlsruhe, Germany). HistoBond positive charged slides were obtained from Marienfelder (Mergentheim, Germany) and Vectashield mounting medium from Vector (Burlington, Canada). Cell proliferation reagent WST-1 was purchased from Roche (Mannheim, Germany). The RNeasy plus mini kit, the QiaShredder columns, the QuantiTect Transcription kit and SYBR Green PCR kit were obtained from Qiagen (Hilden, Germany). 2.2. Culture of hMSCs hMSCs were obtained from biopsy material after total hip replacement of femoral heads after informed consent of the donors. The age of the 7 donors was ranging from 51 to 81 years; cells were used for experiments from passage 3 to 10. hMSCs were cultured with alpha MEM, 15% FBS, 100 U/ml penicillin and 100 ␮g/ml streptomycin at 37 ◦ C, 5% CO2 in a humidified atmosphere. The medium exchange was performed twice a week. Prior experiments the MSCs were characterized according to the standard criteria for MSCs (Dominici et al., 2006). The cells demonstrated the expression of the MSC-associated positive markers CD73, CD90 and CD105 and lacked the expression of CD34, CD45 and HLA-DR. Further, all hMSC-isolations exhibited the typical fibroblast-like morphology and had been tested for their osteogenic capacity in monolayer culture by alkaline phosphatase staining and von Kossa staining. For experiments, the cells were selected randomly. The cells were harvested with 0.05% trypsin/EDTA solution for 3 min at 37 ◦ C and recovered cells were seeded out at the desired density for spheroid formation. 2.3. Spheroid formation systems and osteogenic induction The different culture techniques tested for the formation of multicellular aggregates are illustrated in Fig. 1. The aggregate cultures were assessed by the spheroid formation efficiency, homogeneity of size and the vitality of the aggregates during long term cultivation. The formation efficiency is defined as the ratio of obtained aggregates to the initially number of seeded aggregates, e.g. number of hanging drops or culture tubes or wells, as percentage. The osteogenic differentiation was started after spheroid formation. Culture medium was replaced by osteogenic medium with 3 × 10−4 M l-ascorbic acid phosphate, 1 × 10−7 M dexamethasone and 5 × 10−3 ␤-glycerophosphate whereas control spheroids were

Fig. 1. Culture techniques. (A) Spheroid formation and cultivation by liquid suspension culture on a rotation platform. (B) Hanging drop (HD) technique for initially spheroid formation. For long term cultivation HD spheroids were transferred into (A) or (C), a semi solid gel culture. (D) Cultivation and aggregation of a cell suspension in 96 well non adhesive culture plates culture plates and (E) in a polypropylene tube culture.

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kept in complete culture medium. The medium exchange was performed every 2–3 days. 2.3.1. Liquid suspension culture hMSCs were seeded at a density of 6.5 × 104 cells/ml in 35 mm non adhesive bacterial grade Petri dishes (Greiner, #627102) at a total volume of 3 ml and cultivated on a rotating platform (Heidolph Rotamax 120, 20 mm orbit) at a starting range of 75 rpm. After spheroid formation on day 2 the rotation range was raised up (85–95 rpm) to minimize the aggregation of multi-spheroids. 2.3.2. Combined hanging drop (HD) and suspension culture Hanging droplets of 20 ␮l cell suspension containing 5 × 104 hMSCs were prepared in a lid of a 12 cm quadratic culture dish by utilizing the surface tension. The lid was inverted without disturbing the drops and placed over the bottom filled with PBS to prevent a drying out. The cells were allowed to aggregate in the hanging drops for 2 days before transferring the spheroids by washing in a liquid suspension culture or in a semi solid culture for further cultivation. A semi solid culture was prepared by adding 1% methylcelluloses (MC) to the culture medium. 2.3.3. Polypropylene tube culture Conical polypropylene tubes were used for the generation of spheroids by aggregation of hMSCs. A cell suspension with the desired number of hMSCs was incubated in polypropylene tubes on a rotation platform (65 rpm) for 2 days to generate more spherical aggregates. A cell suspension containing 2 × 105 hMSCs was adequate for cultivation in a total volume of 2 ml culture medium in 15 ml tubes, whereas a number of cells of 2 × 104 were cultivated in a total volume of 0.5 ␮l in 1.5 ml tubes.

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to the primary antibody anti-Collagen Type 1 in a dilution of 1:100 for 1 h in a humidified chamber at room temperature. After rinsing three times with PBS, slides were incubated with the secondary antibody goat anti mouse IgG1 (Alexa Fluor dye 568) in a dilution of 1:200 for 1 h at room temperature or alternatively over night at 4 ◦ C. Then, sections were washed three times in PBS and then in dH2 O once. Afterwards sections were embedded in Vectashield mounting medium and analyzed with fluorescent light microscopy (Axiophot, Carl Zeiss GmbH, Oberkochen, Germany). 2.5.3. Alkaline phosphatase staining The detection of the alkaline phosphatase (ALP) on the sections was conducted using the leukocyte alkaline phosphatase staining kit. The sections were washed with dH2 O and incubated with the staining solution prepared according to the manufacturer’s instructions at room temperature for 20 min in the dark. The colorimetric reaction was stopped by washing with dH2 O and allowed to air dry, and then sections were analyzed by bright field microscopy (IX 81, Olympus GmbH, Hamburg, Germany). 2.5.4. von Kossa staining The mineral deposition of the bone specific extra cellular matrix was determined by von Kossa staining for calcium. Briefly, slides were washed two times with dH2 O and incubated with 5% silver nitrate solution for 1 h in the dark. The cryosections were washed thoroughly three times for 5 min with deionised water and developed with 2.5% carbonate buffered formalin for 3 min maximum. After washing with dH2 O three times the sections were embedded in Vectashield mounting medium and analyzed by bright field microscopy (IX 81 from Olympus GmbH). 2.6. WST-1 assay of hMSC aggregates

2.3.4. 96 well non adhesive culture plates For spheroid formation in 96 well, U-bottom, non adhesive plates (Greiner, #650185) a cell suspension at concentrations of 1 × 104 to 2 × 104 hMSCs per well was cultured in a total volume of 150 ␮m on a rotation platform at 75 rpm. To prevent cell adhesion the culture plates were coated with 10% pluronic in dH2 O or 2% poly-HEMA in acetone/ethanol (1:1). After two days of cellular aggregation on a rotation platform, a static cultivation was possible without cell attachment on the substrate surface. 2.4. Live and dead staining with FDA/PI Spheroids were collected and washed with PBS. Then spheroids were incubated with a solution of 25 ␮g/ml fluorescein diacetate (FDA) and 40 ␮g/ml propidium iodide (PI) in PBS for 1 min. After washing two times with PBS the spheroids were analyzed with fluorescent light microscopy (Axiophot, Carl Zeiss GmbH, Oberkochen, Germany). 2.5. Histological investigation 2.5.1. Cryosections Spheroids were collected, washed with PBS and fixed with ice cold methanol/acetone (7:3) for 30 min. After washing two times with PBS spheroids were transferred into 25% sucrose and cryosections (7 ␮m) were prepared using a cryostat (Leica Microsystems, Wetzlar, Germany) and placed onto HistoBond positive charged slides. Sections were stored at −20 ◦ C. Before used for further analyses the sections were thawed and allowed to air dry for 1 h at 37 ◦ C. 2.5.2. Immunhistochemistry Sections were washed three times with PBS and blocked with 10% goat serum, 2% BSA in PBS for 30 min. Sections were exposed

The metabolic activity of comparable spheroids was determined by cell proliferation and cell viability reagent WST-1. Spheroids in a volume of 20 ␮l were transferred into a 96 well cell culture plate. To prepare the WST-1 assay 80 ␮l fresh medium and 10 ␮l reagent WST-1 were added per well. For background control 100 ␮l fresh medium and 10 ␮l WST-1 were used. The spheroids were incubated for 24 h, 37 ◦ C, 5% CO2 at a humidified atmosphere, then 100 ␮l of the reaction mix was transferred into a 96 well culture plate for measurement of the absorbance at 405 nm (SLT Rainbow, Tecan, Crailsheim, Germany). 2.7. Quantitative real time PCR Total RNA was obtained from 24 pooled spheroids per sample using the RNeasy plus mini kit according to the manufacturer’s instructions. For homogenization of the samples QiaShredder columns were used additionally. The amount of RNA was determined spectrometrically (BioPhotometer, Eppendorf, Hamburg, Germany) in 10 mM Tris–HCl, pH 7.5. 200 ␮g total RNA was used as a template for cDNA synthesis using QuantiTect Reverse Transcription kit with integrated removal of genomic DNA. For amplification 2 ␮l aliquots of the first strand reaction were performed using Quanti Tect Primer Assays (Qiagen) for collagen-1 type A1 (NM000088), collagen-3 type A1 (NM000090), BMP2 (NM001200), osteopontin (OPN; NM000582) and ␤-Tubulin (NM000478). Quantitative real-time PCR analyses were performed using Quanti Fast SYBR Green PCR kit in an iCycler (BioRad; München, Germany). Following a 5 min Taq Polymerase activating step at 95 ◦ C, 40 amplification cycles were performed by denaturing at 95 ◦ C for 30 s and annealing and elongation for 30 s at 60 ◦ C. To check genomic DNA contamination a PCR reaction mixture produced without the addition of the reverse transcriptase was analysed for each primer. Specificity of the PCR products

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was verified by performing a melting curve analysis at a rate of 0.2 ◦ C/s. Levels of target gene CT -values were standardized against ␤-Tubulin expression, and then induction of osteoblast related genes in osteogenic spheroids was normalized to controls. Every target gene was tested in triplicates. 2.8. Data acquisition of the size of the spheroids The cross-sectional diameter of the aggregates was determined by transmitted light microscopy using the Analysis pro software (Olympus GmbH, Hamburg, Germany). The data are presented as mean values ± standard deviation. 2.9. Statistical intervention If not stated otherwise, all experiments were performed on 8 spheroids per group and the data are presented as mean values ± Gaussian error distribution. The OriginPro software package was used for statistical analyses (Origin-Lab Corp., Northampton, USA). Levels of significance were calculated by one-way ANOVA. Differences were considered significant at p < 0.05. 3. Results

85–95 rpm, whereas higher rotation led to disintegration of the tightly packed spheroids. No spheroid formation was obtained by static cultivation of a cell suspension in bacterial grade dishes. In this case, hMSCs adhered to the bottom of the bacterial grade dishes and formed a monolayer (data not shown). The tendency to adhere was also shown after spheroid formation on a rotation platform when the aggregates were cultivated without further shaking (data not shown). 3.2. HD/liquid suspension culture The spheroids were generated in HD with initial cell densities of 5 × 103 cells per drop. After two days tightly packed spheroids had formed, but the final spheroid formation efficiency of 46% was rather low. By replacing the lid on the culture dish some droplets were disturbed and a loss of spheroids appeared by transferring them into a Petri dish for further cultivation on a rotation platform (Fig. 2B2). The initially generated spheroids were homogeneous, but after 3 days of cultivation on a rotation platform the formation of multi-aggregates started. At this time point the small aggregates of 2–3 spheroids were still vital as shown by PI/FDA staining (Fig. 2B2), but further cultivation led to irregularly shaped multi-aggregates with a reduced vitality.

3.1. Liquid suspension culture 3.3. HD/semi solid gel Cultivation of a cell suspension of 5 × 104 cells per ml with bacterial grade dishes on a rotation platform generated many randomly sized spheroids after 2 days (Fig. 2A1). The size of the individual spheroids and the size distribution in different cultures varied extremely and multi-aggregation was observed leading large irregular shaped masses during the first week of cultivation. The vitality estimated using PI/FDA double staining of hMSCs in these multi aggregates was very low (Fig. 2A2). The aggregation was slightly reduced by scaling-up the initial shaking speed from 65 rpm to

388 spheroids were generated by HD technique with a density of 5 × 103 hMSCs per spheroid. After transferring the spheroids into a semi solid culture only 165 spheroids were observed, leading to an effective spheroid formation efficiency of 45% (Table 1). The semi solid media inhibited the movement of the individual spheroids and their aggregation. As shown by PI/FDA staining, vital spheroids were cultured over a time period of 17 days (Fig. 2C) with a final diameter of 171 ␮m ± 28 ␮m for spheroids cultured with differen-

Fig. 2. Spheroid formation maintained by different culture techniques and the corresponding PI/FDA staining. Vital cells were marked green, non vital cell were red. (A1) Heterogeneous cellular aggregation maintained on a rotation culture at day 2. (A2) During long term cultivation these aggregates formed multi aggregates with a low vitality. (B1) Spheroids generated by hanging drop technique and subsequent transferred into a rotation culture. After transfer, single aggregates were observed, which start to form multi-aggregates soon after 3 days (B2). Initially, the vitality was still high, but multi aggregation proceeds with a reduction in vitality (see A2). (C) Spheroids generated in a hanging drop culture and transferred into a semi solid gel kept vital, compact and homogenous even after 17 days of cultivation. (D) Spheroid formation in 1.5 ml conical polypropylene tubes. After 29 days of cultivation the vitality of the spheroids was high. (E and F) Spheroids performed in 96 well non adhesive culture plates on a rotation platform. (E) Spheroid formation in pluronic 10% coated plates was incomplete whereas (F) poly-HEMA coating prevented cell adhesion at the bottom of the culture vessel. Obtained spheroids were vital after a culture period of 29 days.

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Table 1 The table shows the efficiency of spheroid formation and the cross-sectional diameter generated by different culturing techniques. The spheroid formation efficiency was shown as percentages of the number of seeded aggregates (n). The values for HD/suspension and HD/semi solid were obtained after transfer in a liquid suspension culture or a semi solid gel. Growth depending differences affected by cultivation with standard culture medium (control) and osteogenic medium are compared in the last two columns. n Liquid suspension culture HD/suspension HD/semi solid 1.5 ml polypropylene tube 15 ml polypropylene tube 96 well non adhesive plates

– 160 388 20 6 96

Spheroid formation efficiency 46% 45% 90% 83% 95%

Cells per spheroid

Cross-sectional diameter of control

Cross-sectional diameter of differentiation

Statistic 5,000 5,000 20,000 200,000 11,000

Multi aggregation Multi aggregation 159 ␮m ± 19 ␮m (±16.4%) 323 ␮m ± 29 ␮m (±12%) Non specified 200 ␮m ± 18 ␮m (±9%)

Multi aggregation Multi aggregation 171 ␮m ± 28 ␮m (±12%) 360 ␮m ± 56 ␮m (±15 5%) Non specified 231 ␮m ± 26 ␮m (±11%)

tiation medium and 159 ␮m ± 15 ␮m for standard culture medium. Spheroids cultured with osteogenic medium for 17 days were positive for ALP staining while control spheroids showed no expression (Fig. 3A and B).

3.4. Polypropylene tube culture By cultivation of a cell suspension in 15 ml polypropylene tubes aggregation of hMSCs could be performed and it was possible to

Fig. 3. Osteogenic differentiation in hMSC aggregates. (A) Alkaline phosphatase staining of aggregates maintained by 5 × 103 hanging drop and transferred in a semi solid culture after osteogenic induction for 17 days and (B) aggregates cultured without osteogenic factors as control. Osteogenesis of hMSC aggregates generated in polypropylene tube cultures. (C) Strong mineralization after 35 days of osteogenic induction inside big aggregates of 2 × 105 cells maintained by 15 ml tube cultures and (D) controls. (E and F) Combined von Kossa and ALP staining in small aggregates of 2 × 104 cells cultured in 1.5 ml polypropylene tubes. (E) After 29 days of osteogenic stimulation ALP expression was observed, but no mineralization. (F) Aggregates for control were negative for ALP and mineralization. (A and B) bar = 50 ␮m and (C–F) bar = 200 ␮m.

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Fig. 4. Osteogenesis in hMSCs spheroids generated in 96 well non adhesive culture plates. (A) Appearance of ALP positive cells close to the surface after 21 days of osteogenic treatment. (B) Control spheroids cultured with standard culture medium were negative for ALP. (C) Deposition of mineralized ECM after 28 days of osteogenic induction while controls (D) were negative for mineralization. (E) Expression of collagen-1 throughout the osteogenic spheroids and (F) control. (G) Quantitative RT-PCR analysis of the n-fold expression of osteoblast-associated genes compared to the values of controls. The regulation of collagen-1 (Col1), collagen-3 (Col3), osteopontin (OPN) and bone morphogenic protein 2 (BMP-2) are demonstrated after 24 days of osteogenic induction. Bar = 50 ␮m.

direct the osteogenic differentiated as shown by von Kossa staining after 35 days of cultivation (Fig. 3C and D). In order to scale down this technique a cell suspension of 2 × 104 hMSCs was cultured in 1.5 ml polypropylene tubes with a formation efficiency of 90% (Table 1). Data of PI/FDA staining indicated that the spheroids kept vital for a culture period of 29 days (Fig. 2D) and the differentiation into the osteogenic lineage was shown by ALP staining (Fig. 3E), while controls were negative (Fig. 3F). Whereas the spheroids held in differentiation medium had a diameter of 360 ␮m ± 60 ␮m, spheroids in standard culture medium had a smaller diameter of 323 ␮m ± 29 ␮m and the shape was slightly irregular (Table 1). 3.5. 96 well non adhesive culture plates The formation of tightly packed, round spheroids was observed after 2 days of cultivation in 96 well non adhesive culture plates. Under these conditions the spheroid formation in 96 wells was over 95% (Table 1), whereas cultivation without additionally poly-HEMA coating led to adhesion to the bottom of the culture vessel. hMSCs differentiated into osteoblasts as shown by ALP staining on day 21 of osteogenic treatment, von Kossa staining for calcium deposition of ECM on day 28 as well as anti-collagen-1 immunostaining (Fig. 4). Quantitative real time RT-PCR was used to examine the regulation of osteoblast-associated genes compared to control spheroids on day 24 of 16 pooled spheroids. Compared to control spheroids an increase of the marker genes BMP-2 (15-fold), collagen-1 (6fold), collagen-3 (4-fold) and OPN (30-fold) was expressed in the osteogenic spheroids. The data of PI/FDA staining (Fig. 2F) showed a predominant green color indicating the vitality. Further, the cellular viability in spheroids was quantified by WST-1 assay according to the cultivation time. The absolute extinction was shown as percentages of the viability at day 1 (Fig. 5A). There was a clear tendency toward a decreased viability of the control spheroids as well as the osteogenic spheroids found in these data, but the viability of the osteogenic spheroids at day 21 was significantly higher than controls. For the control spheroids there was a decrease in viabil-

ity found from 100% ± 6% at culture day 1 to 40% ± 3% at day 21 and in case of the osteogenic spheroids from 100% ± 4% at day 1 to 60% ± 4% at day 21. In accordance, the corresponding cross-sectional diameter of the spheroids (Fig. 5B) decreased significantly (p < 0.0001) from 632 ␮m ± 35 ␮m at day 1 to 353 ␮m ± 28 ␮m at day 21 in case of controls and from 630 ␮m ± 21 ␮m at day 1 to 349 ␮m ± 16 ␮m at day 21 in case of the osteogenic spheroids. At day 21 there was no significant difference in size found between both groups. Based on this data the viability per diameter of the spheroids was calculated and normalized against the values of day 1 (Fig. 5C). The viability per diameter was demonstrated to increase from the initial level of 1 ± 0.06 for control and 1 ± 0.05 for osteogenic spheroids to 1.2 ± 0.03 for control and 1.3 ± 0.03 at day 5. During further cultivation time a slightly decrease in the viability per diameter was found for the osteogenic spheroids, the final viability per diameter was 1.1 ± 0.08 conforming the initial level. Compared to the osteogenic spheroids there was a significant loss in viability for the control spheroids down to 0.7 ± 0.06. Depending on the initial cell density per well, the diameter of the performed spheroids could be controlled as shown by the mean of the cross-sectional diameter of control and osteogenic spheroids at a total cultivation time of 21 days (Table 2). A cellular density of 1 × 104 cells led to spheroids with a mean cross-sectional diameter of 267 ␮m ± 13 ␮m for control and 304 ␮m ± 45 ␮m for the osteogenic samples. A cellular density of 1.5 × 104 cells let to aggregates with a diameter of 383 ␮m ± 23 ␮m for controls and 397 ␮m ± 24 ␮m for the osteogenic spheroids, and 2 × 104 cells to spheroids of 435 ␮m ± 13 ␮m for controls and 409 ␮m ± 27 ␮m for the osteogenic spheroids. There was no significant difference (p > 0.05) in size found between osteogenic and control spheroids for all seeded cell numbers. 4. Discussion In this study a systematic analysis of different strategies for the generation of a 3D scaffold-free in vitro model for osteogenesis of hMSCs was conducted. In comparison to monolayer cultures, 3D

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Fig. 5. Examination of the metabolic activity and size development of hMSCs aggregates generated in 96 well, U-bottom, non adhesive culture plates according to the cultivation time. Data were shown as mean ± standard deviation (n = 8). The cellular aggregation was performed on 1.8 × 104 cells per spheroid and then, aggregates were cultivated with standard culture medium as control and osteogenic differentiation medium. (A) Metabolic activity of aggregates measured by WST-1 assay shown as percentages of day 1. (B) Corresponding diameter of these spheroids. (C) Metabolic activity per diameter. The absolute extinction of WST-1 was shown per diameter in relation to day 1.

cell culture systems provide a well controlled environment that corresponds to the physiological conditions in vivo where the cells are surrounded by other cells or fibrous layers (Berrier and Yamada, 2007). The 3D microenvironment is similar as observed in the stem cell niche in vitro and mediated the cell behavior (Watt and Hogan, 2000; Wang et al., 2009). It is reported that integrin specific engagement influences adhesion, proliferation and differentiation processes in MSCs (Martino et al., 2009; Kale et al., 2000). Cultivation of hMSCs in scaffold free 3D cell aggregates provides better intercellular as well as ECM interactions and tissue specific capacities without any scaffold depending influences (Wang et al., 2009). Hence, these culture techniques are displaying as a link between conventional monolayer or 3D cultures and whole organs (Rossi et al., 2005). During osteogenesis in vitro the development of mature osteoblasts is associated with a cellular condensation, which proceeds during matrix mineralization (Dunlop and Hall, 1995; Kale et al., 2000). However, in vitro there is also strong evidence that a 3D microenvironment is essential for bone formation. For example, the appearance of so called bone nodules associated with osteoblast-related ECM expression and mineralization in MSC monolayer cultures (Beresford et al., 1993; Maniatopoulos et al., 1988) indicates the importance of 3D cellular interaction for this developmental process. Nonetheless, there is less information about osteogenic differentiation of hMSCs in scaffold free aggregates. The expression of mineralized ECM of hBMSCs (bone

marrow stromal cells) in pellet cultures differentiated into chondroblast first by TGF-␤1 treatment and then sequentially exposed to osteogenic conditions (Muraglia et al., 2003) had described. Moreover, direct evidence of osteogenic differentiation of hMSCs in spheroid cultures was reported already (Wang et al., 2009) and discussed to be mediated by specific integrin interactions in the high density cultures (Kale et al., 2000). Previous studies have demonstrated cell scaffold-free aggregation of ES cells (Karp et al., 2006; Doetschman et al., 1985), when cultivated in bacterial grade Petri dishes. Within this investigation this culture technique was adapted and led to the formation of hMSCs aggregates. Despite of the hydrophobic features of these culture dishes, the rotation was necessary to avoid cell and spheroid attachment at the bottom of the Petri dish. The aggregation of hMSCs by liquid suspension culture on a rotation platform occurred spontaneously resulting in heterogeneous spheroids in size as well as in number. Consequently, it was not possible to determine the average of size and the spheroid formation efficiency. Moreover, since the aggregates tend to form larger amorphous masses with a low vitality after 1 week, the method was not proper for the long term cultivation needed for the process of osteogenic differentiation. To avoid multi-aggregation in a liquid suspension culture on a rotation platform, the aggregation step was performed in HD, a widely used technique for spheroid formation (Takahashi et al., 2003). The cellular aggregation occurs in single droplets, but a

Table 2 Cross-sectional diameter of hMSCs spheroids (n = 20) performed in 96 well non adhesive culture plates after 21 days of cultivation depending on varying initial cell numbers. Data were presented as mean value ± standard deviation. Initial cell number per 96 well Diameter of control spheroids Diameter of osteogenic treated spheroids

10,000 267 ␮m ± 13 ␮m (±6%) 305 ␮m ± 50 ␮m (±16%)

15,000 382 ␮m ± 24 ␮m (±6%) 397 ␮m ± 26 ␮m (±7%)

20,000 435 ␮m ± 14 ␮m (±4%) 409 ␮m ± 2 ␮m (±7%)

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medium exchange is not possible. Consequently the spheroids were transferred for long term cultivation and differentiation into a liquid suspension culture or alternatively into a semi solid gel. Even if the aggregation of hMSCs in the HD culture was well controlled and led to homogenous spheroids, the fusion of multi-aggregates with a low vitality was observed again after transfer in a liquid suspension culture for long term cultivation as a feature of this cultivation methods. To overcome this disadvantage, a semi solid gel culture was introduced for long term cultivation, a technique previously described for embryoid body formation (Dang et al., 2002) as well as for colony forming assays, e.g. for bone marrow derived progenitor cells (Rennick et al., 1987). Under these culture conditions the spheroids kept vital during the whole cultivation period (Fig. 2C) and osteogenesis of hMSCs was induced successfully as shown by ALP staining after 17 days (Fig. 3A). The gel-like texture prevented multi-aggregation as well as attachment on the bottom of the culture vessel, but the final formation efficiency was only 45% (Table 1) and the viscous properties hindered the medium exchange. Therefore, all techniques including a formation step in a HD culture are problematic. The droplet were disturbed easily by inverting the lid and in consequence of the transferring step there was an additional lose of aggregates. Another strategy investigated to maintain aggregation of hMSCs and induce osseous differentiation successfully was the cultivation of a cell suspension in 15 ml polypropylene tubes. This cultivation technique has been previously described for the chondrogenic differentiation in pellet or micromass cultures (Sekiya et al., 2002; Mackay et al., 1998), and the importance of strong cell–cell interaction in 3D cell cultures on the chondrogenic differentiation in combination with specific differentiation factors is discussed already (Johnstone et al., 1998; Bosnakovski et al., 2006). Besides this, self-induced chondrogenic differentiation of bovine MSCs was demonstrated in micromass cultures (Bosnakovski et al., 2004). However, this culture system was appropriate for the osteogenic differentiation of hMSCs as shown by the deposition of mineralized ECM after 35 days of osteogenic treatment (Fig. 3C), but due to the large amount of cells the cultivation in 15 ml tubes has its limitations and consequently this technique was transferred into smaller polypropylene vessels for cultivation in parallel. Even if the formation efficiency in the small tubes was found to be high as 90%, the generated aggregates had a slightly irregular geometry and no mineralization was found after 29 days of osteogenic treatment (Fig. 3E). The distinction in mineralization found in 15 ml polypropylene tube cultures but not in the 1.5 ml tubes could be related to inter donor variability, cellular senescence in culture (Digirolamo et al., 1999; Baxter et al., 2004; D’Ippolito et al., 1999; Wagner et al., 2009) or reduced oxygen transfer causes by the closed lids. The overall most effective and convenient technique to perform tightly packed spherical aggregates was the cultivation of a cell suspension in 96 well non adhesive culture plates. This technique has been previously described for example for embryonic body formation from mouse embryonic stem cells by statically cultivation (Kurosawa et al., 2003) as well as for aggregation of human embryonic stem cells in low attachment plates by centrifugation (Ng et al., 2005). The spheroids generated by cultivation of a cell suspension in 96 well non adhesive culture plates were round and tightly packed with a constant range of size reflected by the mean of the cross-sectional diameter which differs approximately 9% for control spheroids and 11% (Table 1) for osseous spheroids. However, this technique offered an easy way to generate spheroids with a formation efficiency of over 95%, the highes value of the tested methods, and the advantage of size control of the spheroids. In comparison, techniques starting with an initial formation step in a hanging drop culture (HD/liquid suspension culture

and HD/semi solid culture) had a low effective spheroid formation efficiency of approximately 50%. Although the formation efficiency in polypropylene tubes was as high as 90%, the handling of the single tubes was not suitable for high parallelism and the amount of culture media was higher compared to cultivation in non adhesive, 96 well plates. Especially, when expenses arising for specific differentiation factors could be very high a cost saving technique is required for the generation of higher numbers of spheroids. In 96 well non adhesive culture plates the osteogenic differentiation could be induced successfully by soluble factors added to the culture medium as demonstrated by the expression of ALP at day 18 as well as the expression of collagen-1 and the deposition of mineralized ECM and at day 21. These histological findings could be confirmed by the data of real time PCR, at which an upregulation in the gene expression of collagen-1, collagen-3, BMP-2 and osteopontin was shown after 25 days of osteogenic treatment (Fig. 4). On the basis of experimental data, it is not intended to associate the expression of specific osteogenic markers to a specific time point or cultivation method, because of the inter-donor variability and the ranging passage numbers. Nonetheless, these results provide ample evidence to verify the osteogenesis in scaffold-free 3D aggregates. With respect to our findings, we conclude that the cultivation and differentiation of hMSCs in scaffold-free aggregates facilliates a tissue specific in vitro model for the investigation of developmental processes as well as for drug testing. Very interesting was the decrease in size of the spheroids during long-time cultivation (Fig. 5B). This tendency was determined in several experiments, whereas after approximately 2 weeks this size effect stopped (data not shown). Thus, this seems to reflect a process of strong condensation of the spheroids. This proceeding condensation of the spheroids might be reflected in the decreasing values of WST-1 assay (Fig. 5A), which not only interfered the passive diffusion of the reagent and its metabolite, but also be addressed by cellular organization and inhibition of proliferate activities. After normalization of the viability against the crosssectional diameter the viability per diameter was found to be relatively constant for the osteogenic treated spheroids, while controls had a reduced final viability. However, the investigation of the vitality of these aggregates is hardly to determine. For scaffold based 3D cultures assays to estimate metabolic activities are previously described (Ghods et al., 2007), but these cultures are more permeable than the strong condensate spheroid cultures and assays like the life and dead PI/FDA staining (Carpenedo et al., 2007) only detects the outer cells. Nonetheless, the condensation of the aggregates was proportional to the initial cell seeding, so it was possible to control the final size of the spheroids (Table 2). At all, on the basis of our data the size of the spheroids was not affected significant by the differentiation process in 96 well non adhesive culture plates, in contrast to previously reported differences in size associated with chondrogenic development in pellet cultures (Mackay et al., 1998; Sekiya et al., 2001) solely, and in case of an change from chondrogenic to osteogenic conditions (Muraglia et al., 2003).

5. Conclusion In this study we have compared and optimised different culture techniques for the generation of a 3D scaffold-free in vitro model for osteogenesis of hMSCs. Based on non adhesive 96 well culture plates a protocol for efficient generation of spherical aggregates with low variation of relevant properties was established. The formation of homogenous spherical aggregates occurred in a high efficiency and the size was well controlled in dependency of the cell number seeded per well initially.

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Using this culturing method it was demonstrated that the spherical cell aggregates can be differentiated into the osteogenic direction. Thereby an important basis for studying cell and molecular biological processes of osteogenic mesenchymal stem cell differentiation in scaffold-free 3D in vitro models was provided. However, in contrast to the use of individual polypropylene tubes, non-adhesive, 96 well culture plates were more appropriate for the generation of cell aggregates in high numbers, since the handling was easier and less working steps were required. In case of hanging drop techniques the final outcome of cell aggregates was low. This technique did not support a medium exchange, hence the aggregtes had to be transfered to another culture method. Acknowledgements This work was funded by EU under the framework VI project OsteoCord. The authors are solely responsible for its content, it does not represent the opinion of the European Community and the Community is not responsible for any use that might be made of the information contained therein. References Bael, A.V., Proesmans, M., Tilemans, D., Denef, C., 1995. Interaction of LHRH with growth hormone-releasing factor-dependent and -independent postnatal development of somatotrophs in rat pituitary cell aggregates. J. Mol. Endocrinol. 14, 91–100. Banu, N., Banu, Y., Sakai, M., Mashino, T., Tsuchiya, T., 2005. Biodegradable polymers in chondrogenesis of human articular chondrocytes. J. Artif. Organs 8, 184–191. Bartholomä, P., Gorjup, E., Monz, D., Reininger-Mack, A., Thielecke, H., Robitzki, A., 2005. Three-dimensional in vitro reaggregates of embryonic cardiomyocytes: a potential model system for monitoring effects of bioactive agents. J. Biomol. Screen. 10, 814–822. Baxter, M.A., Wynn, R.F., Jowitt, S.N., Wraith, J.E., Fairbairn, L.J., Bellantuono, I., 2004. Study of telomere length reveals rapid aging of human marrow stromal cells following in vitro expansion. Stem Cells 22, 675–682. Beresford, J.N., Graves, S.E., Smoothy, C.A., 1993. Formation of mineralized nodules by bone derived cells in vitro: a model of bone formation? Am. J. Med. Genet. 45, 163–178. Berrier, A.L., Yamada, K.M., 2007. Cell–matrix adhesion. J. Cell Physiol. 213 (3), 565–573. Bosnakovski, D., Mizuno, M., Kim, G., Ishiguro, T., Okumura, M., Iwanaga, T., Kadosawa, T., Fujinaga, T., 2004. Chondrogenic differentiation of bovine bone marrow mesenchymal stem cells in pellet cultural system. Exp. Hematol. 32, 502–509. Bosnakovski, D., Mizuno, M., Kim, G., Takagi, S., Okumura, M., Fujinaga, T., 2006. Chondrogenic differentiation of bovine bone marrow mesenchymal stem cells (MSCs) in different hydrogels: influence of collagen type II extracellular matrix on MSC chondrogenesis. Biotechnol. Bioeng. 93, 1152–1163. Carpenedo, R.L., Sargent, C.Y., McDevitt, T.C., 2007. Rotary suspension culture enhances the efficiency, yield, and homogeneity of embryoid body differentiation. Stem Cells 25, 2224–2234. Chatterjee, S.S., Nöldner, M., 1994. An aggregate brain cell culture model for studying neuronal degeneration and regeneration. J. Neural Transm. Suppl. 44, 47–60. Colter, D.C., Sekiya, I., Prockop, D.J., 2001. Identification of a subpopulation of rapidly self-renewing and multipotential adult stem cells in colonies of human marrow stromal cells. Proc. Natl. Acad. Sci. U.S.A. 98, 7841–7845. D’Ippolito, G., Schiller, P.C., Ricordi, C., Roos, B.A., Howard, G.A., 1999. Age-related osteogenic potential of mesenchymal stromal stem cells from human vertebral bone marrow. J. Bone Miner. Res. 14, 1115–1122. Dang, S.M., Kyba, M., Perlingeiro, R., Daley, G.Q., Zandstra, P.W., 2002. Efficiency of embryoid body formation and hematopoietic development from embryonic stem cells in different culture systems. Biotechnol. Bioeng. 78, 442–453. Dezawa, M., Ishikawa, H., Itokazu, Y., Yoshihara, T., Hoshino, M., Takeda, S., Ide, C., Nabeshima, Y., 2005. Bone marrow stromal cells generate muscle cells and repair muscle degeneration. Science 309, 314–317. Digirolamo, C.M., Stokes, D., Colter, D., Phinney, D.G., Class, R., Prockop, D.J., 1999. Propagation and senescence of human marrow stromal cells in culture: a simple colony-forming assay identifies samples with the greatest potential to propagate and differentiate. Br. J. Haematol. 107, 275–281. Doetschman, T.C., Eistetter, H., Katz, M., Schmidt, W., Kemler, R., 1985. The in vitro development of blastocyst-derived embryonic stem cell lines: formation of visceral yolk sac, blood islands and myocardium. J. Embryol. Exp. Morphol. 87, 27–45. Dominici, M., Blanc, K.L., Mueller, I., Slaper-Cortenbach, I., Marini, F., Krause, D., Deans, R., Keating, A., Prockop, D., Horwitz, E., 2006. Minimal criteria for defining multipotent mesenchymal stromal cells. The International Society for Cellular Therapy position statement. Cytotherapy 8 (4), 315–317.

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