Jellyfish collagen scaffolds for cartilage tissue engineering.

Acta Biomaterialia xxx (2013) xxx–xxx

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Acta Biomaterialia journal homepage: www.elsevier.com/locate/actabiomat

Jellyfish collagen scaffolds for cartilage tissue engineering Birgit Hoyer a,⇑, Anne Bernhardt a, Anja Lode a, Sascha Heinemann b, Judith Sewing c,d, Matthias Klinger e, Holger Notbohm c, Michael Gelinsky a a

University Hospital and Medical Faculty, Technische Universität Dresden, Centre for Translational Bone, Joint and Soft Tissue Research, Fetscher Str. 74, D-01307 Dresden, Germany Max Bergmann Center of Biomaterials and Institute for Materials Science, Technische Universität Dresden, Budapester Str. 27, 01069 Dresden, Germany c Institute of Virology and Cell Biology, University of Lübeck, Ratzeburger Allee 160, 23562 Lübeck, Germany d CRM Coastal Research & Management GmbH, Tiessenkai 12, 24159 Kiel, Germany e Institute of Anatomy, University of Lübeck, Ratzeburger Allee 160, 23562 Lübeck, Germany b

a r t i c l e

i n f o

Article history: Received 27 May 2013 Received in revised form 28 August 2013 Accepted 22 October 2013 Available online xxxx Keywords: Marine collagen Fibril formation Jellyfish Mesenchymal stem cells Chondrogenic differentiation

a b s t r a c t Porous scaffolds were engineered from refibrillized collagen of the jellyfish Rhopilema esculentum for potential application in cartilage regeneration. The influence of collagen concentration, salinity and temperature on fibril formation was evaluated by turbidity measurements and quantification of fibrillized collagen. The formation of collagen fibrils with a typical banding pattern was confirmed by atomic force microscopy and transmission electron microscopy analysis. Porous scaffolds from jellyfish collagen, refibrillized under optimized conditions, were fabricated by freeze-drying and subsequent chemical cross-linking. Scaffolds possessed an open porosity of 98.2%. The samples were stable under cyclic compression and displayed an elastic behavior. Cytotoxicity tests with human mesenchymal stem cells (hMSCs) did not reveal any cytotoxic effects of the material. Chondrogenic markers SOX9, collagen II and aggrecan were upregulated in direct cultures of hMSCs upon chondrogenic stimulation. The formation of typical extracellular matrix components was further confirmed by quantification of sulfated glycosaminoglycans. Ó 2013 Acta Materialia Inc. Published by Elsevier Ltd. All rights reserved.

1. Introduction Collagen is the prevailing component of extracellular matrices in connective tissues. Due to its biocompatibility, biodegradability, low immunogenicity and cell-adhesive properties, collagen is one of the most frequently utilized materials in the field of tissue engineering [1]. Collagen can be extracted from a variety of organisms. Preferential sources of collagen for tissue engineering applications are bovine skin and tendon as well as porcine skin. However, collagen of bovine origin involves the risk of infection with diseases such as bovine spongiform encephalopathy. Furthermore, mammalian collagens, especially of porcine origin, are increasingly rejected for religious reasons [2]. Marine organisms are an alternative natural source of collagen and, presumably, are safer compared to mammals. Recent publications focus mainly on the isolation and characterization of collagen from different fish species, such as salmon [3], shark [4] or deep sea redfish [5] and marine sponges [6]. Another attractive marine source for the extraction of collagen is jellyfish. The global increase in jellyfish population causes major problems in the ecological environment, and their potential use in tissue engineering, next to food industry ⇑ Corresponding author. Tel.: +49 351 458 6694; fax: +49 351 458 7210. E-mail address: [email protected] (B. Hoyer).

and medicine, may help to reduce their further expansion [7]. With a collagen content of more than 60% [8], jellyfish has the potential to become a significant source of collagen in biomedical applications [9]. Isolation and molecular characterization of jellyfish collagen derived from Stomophulus nomurai has been reported decades ago [10,11]. More recent investigations are concerned with collagen from other jellyfish species, e.g. Rhopilema asamushi, Stomolophus meleagris, Catostylus tagi and Rhizostoma pulmo [7– 9,12]. High collagen recovery rates have consistently been reported. Amino acid analyses revealed a composition similar to vertebrate collagen with, however, a lower content of hydroxyproline, which leads to relatively low denaturation temperatures between 26 and 29.9 °C. Differences in the subunit composition of collagens from different species are detectable in their respective electrophoretic pattern. Hence, it may be stated that collagens of different jellyfish species show similarities to different vertebrate collagen types. Some jellyfish collagens are comparable to vertebrate collagen IV or V [10,11], others seem to resemble vertebrate collagen I [7,12,13] and some show a unique structure with a fourth a-chain [9]. Hsieh [14] postulates that jellyfish collagen of S. meleagris is similar to vertebrate collagen type II according to the molecular mobility, salting-out concentration, high content of hydroxylysine, solubility properties, absence of disulfide bonds and highly hygroscopic nature. Similar findings are described by

1742-7061/$ - see front matter Ó 2013 Acta Materialia Inc. Published by Elsevier Ltd. All rights reserved. http://dx.doi.org/10.1016/j.actbio.2013.10.022

Please cite this article in press as: Hoyer B et al. Jellyfish collagen scaffolds for cartilage tissue engineering. Acta Biomater (2013), http://dx.doi.org/10.1016/ j.actbio.2013.10.022

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B. Hoyer et al. / Acta Biomaterialia xxx (2013) xxx–xxx

Bermueller et al. for jellyfish collagen of Rhopilema esculentum, which consists of only one type of a-chain and shows a degree of glycosylation similar to that of vertebrate collagen type II [15]. Since collagen II, as the main component of cartilage extracellular matrix, has been shown to support the chondrogenic differentiation and maintenance of chondrogenic phenotype to a higher extent compared to collagen I [16], jellyfish collagen derived from R. esculentum might be a predominant prospective material for the preparation of scaffolds in cartilage tissue engineering. So far only a few publications have reported on the preparation of jellyfish collagen-based scaffolds for tissue engineering applications. Song et al. [17] generated porous scaffolds by freeze-drying and subsequent chemical cross-linking of acidsolubilized jellyfish collagen. Biocompatibility investigations comprised the attachment of human fibroblasts as well as the immune response after implantation of the scaffolds in vivo, being similar to other collagen sources. Another type of porous scaffold was established by the same group, combining jellyfish collagen and hyaluronic acid [18]. Tubular porous scaffolds from jellyfish collagen reinforced with poly(lactic-co-glycolic) acid fibers were developed by freeze-drying and electrospinning techniques [19,20]. Seeded with osteosarcoma cells, these constructs were cultivated to test the influence of electrospinning parameters on cell adhesion and proliferation, or, when seeded with endothelial cells and smooth muscle cells in a perfusion system, to generate vascular grafts. There are three different types of cartilage: hyaline, fibrocartilaginous and elastic. All cartilage types consist of chondrocytes and an extracellular matrix with collagens, proteoglycans and water, differing in protein types and proportions [21]. Joint surfaces are covered by hyaline cartilage, which is predominantly collagen type II (90–95%), followed by collagen XI and IX [22,23]. On the contrary, in fibrocartilage (in knee joint menisci and intervertebral discs), which has properties of both connective tissues and hyaline cartilage, collagen type I and fewer proteoglycans are present [24]. Analogous to hyaline cartilage, elastic cartilage is mainly composed of collagen type II. The main difference to the other two types is elastin and a higher number of cells [21]. Cartilage repair is very challenging since all three types of cartilage tissue are nonvascularized and have therefore a poor intrinsic regeneration capacity. One approach to the regeneration of cartilage defects is the implantation of a tissue-engineered scaffold colonized with cells. In clinical practice autologous chondrocytes are used for this purpose. There are, however, limitations to this procedure due to the induction of morbidity at the donor site and instability in monolayer cultivation [25]. Mesenchymal stem cells from various tissues, such as bone marrow or adipose tissue [25], represent an alternative source of superior availability and are for that reason the object of research in cartilage tissue engineering [26,27]. Materials for scaffolds in cartilage tissue engineering consist mainly of natural or synthetic polymers [26], both in the shape of either hydrogels [26,28] or porous matrices [27,29,30]. Natural polymers include agarose, collagen, silk, alginate or chitosan [28,30–32]. In the clinical environment, mammalian collagen is already used in the form of membranes, e.g. CaReS (Arthro Kinetics AG, Germany), Chondro-GideÒ (Geistlich Pharma AG, Switzerland), CartimaixÒ (Matricel GmbH, Germany), NovocartÒ 3D (TETECÒ Tissue Engineering Technologies AG, Germany) or hydrogels, e.g. ChondroFillerÒ (Amedrix, Germany). The aim of the present study was to characterize fibril formation parameters of jellyfish collagen to find optimal parameters for the fabrication of stable porous scaffolds from refibrillized jellyfish collagen. Furthermore we wanted to evaluate the suitability of the scaffolds for potential usage in cartilage tissue engineering.

2. Materials and methods 2.1. Collagen Collagen was extracted by pepsin digestion from cured jellyfish R. esculentum (LiroyBV, Rotterdam, The Netherlands) as described before [33]. Briefly, salted jellyfish was cut into pieces and extensively rinsed with cold water until salinity was 60.01. After equilibration in 0.5 M acetic acid for at least 30 min, pieces of jellyfish were homogenized. Following 60 h of pepsin digestion at 4 °C, the solution was centrifuged for cleaning. Collagen was then precipitated from the supernatant by adding NaH2PO4 and KCl at pH 7 for 12 h. After another centrifugation step the collected collagen was redissolved in 0.05% acetic acid and dialyzed against 0.05% acetic acid. It was stored at 20 °C and lyophilized only when required to avoid unwanted cross-linking. Prior to use, a stock solution was generated under constant stirring at 4 °C with a concentration of 5 mg ml1 lyophilized collagen dissolved in 0.01 M HCl. 2.2. Fibril formation For reassembly analysis, 500 ll collagen stock solution in graded concentrations of 5, 4, 3, 2 and 1 mg ml1 were thoroughly mixed with 500 ll of 50 mM tris-(hydroxymethyl)-aminomethane (Tris) buffer containing graded sodium chloride concentrations of 20, 40, 60, 80 and 100 mM; final pH was adjusted to 7.4. Turbidity was measured at 313 nm over 60 min at 4 °C or 25 °C using a UV– Vis spectrophotometer Cary 50 Bio (Varian, Germany). Fibril formation was quantified indirectly by measuring the collagen concentration in the supernantant using the modified Bradford assay, as described previously [34,35]. In brief, upon completion of the fibril formation process over 4 h at 4 or 25 °C, the suspension was centrifuged for 15 min at 10,000g, at 4 °C. 5 ll of the supernantant were mixed with 250 ll Bradford reagent (Sigma–Aldrich, USA) containing 0.035 mg ml1 SDS (Sigma–Aldrich). Absorbance was measured after 15 min at 590 nm using a microplate reader Infinite M200Pro (Tecan, Switzerland). For calibration, a graded series of jellyfish collagen stock solution was used. Finally, the ratio of fibrillized to total initial collagen was calculated, revealing the degree of fibril formation. The graphs show the mean ± standard deviation (n = 6). 2.3. Scaffold preparation Based on the results of the fibril formation studies we developed the following protocol for scaffold preparation. A 5 mg ml1 stock solution of jellyfish collagen was merged 1:1 with 50 mM Tris buffer, pH 8. This preparation with a resulting pH 7.4 was stirred for 12 h at 4 °C. Upon centrifugation at 5000g and 4 °C, the pellet was resuspended in a small amount of supernantant and transferred to 96-well cell culture dishes. Three-dimensional (3D) sponge-like, porous scaffolds were obtained after freezing at a speed of 1 K min1 and subsequent freeze-drying (Alpha 1–2, Christ, Germany) for 24 h. The scaffolds were chemically crosslinked in a 1 wt.% solution of N-(3-dimethylaminopropyl)-N0 -ethylcarbodiimide hydrochloride (EDC, Fluka, Germany) in 80 vol.% ethanol for 2 h. After careful rinsing in deionized water, in 1 wt.% glycine solution, and once more in deionized water, a final freeze-drying step concluded the procedure. 2.4. Atomic force microscopy (AFM) Droplets of refibrillized jellyfish collagen suspended in 50 mM Tris buffer, pH 8, were transferred onto the surface of mica discs.



The collagenous material was allowed to adsorb for 30 min followed by rinsing with deionized water and drying in air. AFM imaging was performed in tapping mode in air using a NanoscopeÒ IIIa Bioscope (Digital Instruments/Veeco, USA) and aluminum reflex coated silicon tips (force constant 40 N m1). Both deflection and height images were captured simultaneously at imaging speeds of 1.2 Hz, scanning 512 lines. Measurements were taken from height images using the Nanoscope software. 2.5. Transmission electron microscopy (TEM) 5 ll graded dilutions of refibrillized jellyfish collagen were applied to formvar-coated copper grids (SF 162-3, Plano GmbH, Germany), negatively stained with 1% phosphotungstic acid in distilled water, and allowed to dry. Grids were examined with a Philips TEM 400 at 60 kV. Fibril diameter was measured semiquantitatively from two pictures with Axio Vision 3.1 software (Zeiss, Germany), in which several sections of the widest fibrils were measured. 2.6. Scanning electron microscopy (SEM) Cell-seeded scaffolds were fixed with 2.5% glutaraldehyde in phosphate-buffered saline (PBS), followed by dehydration in graded series of ethanol and critical point drying (BAL-TEC CPD 30, Liechtenstein). All samples were fixed on carbon pads and sputter-coated with gold. A Philips XL 30/ESEM with field emission gun operated in SEM mode was used. 2.7. Confocal laser scanning microscopy (cLSM) Cell-seeded scaffolds were fixed with 3.7% formaldehyde in PBS. After treatment with 0.2% Triton X-100 in PBS for 5 min, samples were rinsed five times with PBS, followed by blocking of autofluorescence with 3% bovine serum albumin (Sigma–Aldrich) in PBS. The cytoskeleton of the cells was stained using Alexa Fluor 488Ò phalloidin (Invitrogen, USA) and nuclei with DAPI (Sigma–Aldrich). Samples were imaged using a Zeiss cLSM 510. 2.8. Porosity The real volume of freeze-dried jellyfish collagen sponges was evaluated in a helium pycnometer (Ultrapyc1200e, Quantachrome Instruments, USA). Height, diameter and weight were measured to calculate porosity. 2.9. Differential scanning calorimetry (DSC) After thoroughly rinsing in deionized water, samples were sealed in 50 ll melting pans of aluminum (BO14-3003 and BO 14-3017, Perkin Elmer, USA) and finally measured in a Pyris 6 DSC, software version 9.1 (Perkin Elmer), with a heating rate of 2 K min1. An empty melting pan was used as reference sample.

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2.11. Cell culture Human mesenchymal stem cells (hMSCs) (kindly provided by Professor Martin Bornhäuser and co-workers, Medical Clinic I, Dresden University Hospital Carl Gustav Carus) were isolated from bone marrow aspirate of two healthy male donors (age 30 and 32) after providing written informed consent. The application of hMSCs for in vitro experiments was approved by the ethics committee of the Medical Faculty of Technische Universität Dresden. Cells were expanded in Dulbecco’s modified Eagle’s medium (DMEM) low glucose (Biochrom, Germany), containing 10% fetal calf serum, 100 U ml1 penicillin and 100 lg ml1 streptomycin (Biochrom) at 37 °C in a humidified, 5% CO2/95% air incubator. Prior to cell seeding, cylindrical, gamma-sterilized jellyfish collagen scaffolds (diameter: 6 mm; height: 3 mm) were incubated in cell culture expansion medium for 24 h. Upon removal of excess liquid, each scaffold was seeded with 1.2 106 cells in expansion medium. After 24 h the culture medium was replaced with chondrogenic medium containing 100 U ml1 penicillin and 100 lg ml1 streptomycin, 10 lg ml1 insulin, 5.5 lg ml1 transferrin, 6.7 ng ml1 selenium, 0.2 lg ml1 ethanolamine (ITS-X mix, Gibco, Germany), 107 M dexamethasone (Sigma–Aldrich), 0.2 mM ascorbic acid-2-phosphate (Sigma–Aldrich), 10 ng ml1 TGF-b3 (Milteny Biotec, Germany) and 35 lM proline (Sigma–Aldrich). The first medium change was 1 day after seeding and later twice a week over 21 days. Samples for biochemical analysis (n = 3) were washed twice with PBS and frozen at 80 °C in 2 ml Nalgene tubes containing six ceramic beads (Peqlab, Germany). For SEM investigations, samples were washed with PBS and fixed in 2.5% glutaraldehyde in PBS. Samples for gene expression analysis (n = 3) were taken after 1 and 21 days of cultivation, washed with PBS and immediately subjected to RNA isolation as described below.

2.12. Cytotoxicity test Cytotoxicity of jellyfish collagen scaffolds was tested by indirect cultivation of cells with scaffold extracts. Jellyfish collagen scaffolds (diameter: 6 mm; height: 3 mm) were incubated in 1 ml cell culture medium (DMEM containing 10% fetal calf serum, 100 U ml1 penicillin and 100 lg ml1 streptomycin) at 37 °C. 8 103 hMSCs were seeded in 48-well tissue culture plates and cultivated for 1, 7 or 14 days, respectively, either with scaffoldconditioned medium or normal medium. On completing the respective cultivation period, samples were washed twice in PBS and frozen at 80 °C for later analysis. After thawing, samples were lysed with 1% Triton X-100 in PBS for 50 min on ice. During lysis, samples were incubated for 10 min in an ice-cooled ultrasonic bath. The CytoTox 96Ò non-radioactive cytotoxicity assay (Promega, USA) was used for determination of the lactate dehydrogenase (LDH) activity according to the manufacturer’s instructions. Absorbance was read at 492 nm in a microplate reader. LDH activity of the samples was correlated with the number of cells using a calibration line of defined cell numbers. Graphs show mean ± standard deviation (n = 3).

2.10. Mechanical measurements 2.13. MTT staining Uniaxial compressive tests were conducted using an Instron 5566 testing machine (Instron GmbH, Germany). Ten cylindrical samples (diameter 10.3 ± 0.3 mm, height 10.7 ± 0.2 mm) were incubated in modified simulated body fluid (SBF) [36] for 24 h prior to measurements. Wet samples were compressed to 50% of their initial height. For static tests, a velocity of 0.1 mm s1 and for cyclic tests (50 cycles) a velocity of 0.3 mm s1 was used.

After 21 days of cultivation, a cell-seeded and a non-seeded scaffold were incubated in culture medium with 1.2 mM 3-(4,5dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT; Sigma–Aldrich) at 37 °C for 4 h. Intracellular conversion of MTT into a dark blue formazan derivative was macroscopically evaluated.


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2.14. Determination of sulfated glycosaminoglycan (sGAG) and DNA content 1000 ll papain digestion solution (125 lg ml1 papain, 100 mM Na2HPO4, 5 mM ethylenediaminetetraacetic acid (EDTA), 5 mM cystein in deionized water) was added to the frozen cell-seeded scaffolds. The samples were homogenized (2 10 s at 5900 rpm) using a PrecellysÒ 24 apparatus (Peqlab) and were then incubated at 60 °C for 24 h. Subsequently, 50 ll of the samples were applied for quantification using the sGAG assay (Kamiya Biomedical Company, USA) according to the manufacturer’s instructions. Absorbance was assessed at 610 nm in a microplate reader. Another aliquot of the cell lysate was mixed with the Quant-iT™ PicoGreenÒ dsDNA reagent (Molecular Probes, USA), diluted 1:800 in TE buffer (10 mM TRIS, 1 mM EDTA) and incubated for 5 min in the dark. The intensity of fluorescence was measured at excitation/emission wavelengths of 485/535 nm, respectively. Relative fluorescence units were correlated to the amount of DNA using a calf thymus DNA calibration line. Graphs of sGAG concentration in relation to DNA content show mean ± standard deviation (n = 6). 2.15. Reverse transcriptase PCR (RT-PCR) RNA was extracted from cell-seeded scaffolds using the peqGOLD Micro Spin Total RNA Kit (Peqlab) according to the manufacturer’s instructions. During the RNA isolation procedure, cell lysates of three samples cultured under identical conditions were pooled. 100 ng of total RNA were reverse transcribed into cDNA in a 20 ll reaction mixture containing 200 U of Superscript II Reverse Transcriptase (Invitrogen, Germany), 0.5 mM dNTPs (Invitrogen), 12.5 ng ll1 random hexamers (Eurofins MWG Operon, Germany) and 40 U of RNase inhibitor RNase OUT (Invitrogen). 1 ll cDNA in 20 ll reaction mixtures containing specific primer pairs were used for amplification in PCR analysis to detect transcripts of collagen I, collagen IIa, collagen X, aggrecan, SOX9, RUNX2 and b-actin, respectively. Primer sequences (Eurofins MWG Operon), annealing temperatures and amplicon sizes for each gene are summarized in Table 1. The PCR experiments were carried out in a Thermocycler (Vapo-Protect Mastercycler Pro, Eppendorf, Germany) and the resulting PCR products were visualized using the FlashGel™ Dock system (Cambrex Bio Science, USA). 2.16. Statistics Mean and standard deviation are shown in the figures. The difference in the proportion of reassembled collagen within each parameter was statistically evaluated with two-way ANOVA and post hoc Tukey test via Origin 8.6.

3. Results 3.1. Influence of collagen concentration, ionic strength and temperature on fibril formation 3.1.1. Influence of final collagen concentration The influence of the final collagen concentration on fibril formation was investigated in the range of 0.5–2.5 mg ml1 by monitoring the change of turbidity at 313 nm (Fig 1A and B) as well as by indirect evaluation of the degree of fibril formation via modified Bradford assay (Fig. 1C). Turbidity started to rise shortly after mixing the solutions and was too fast to be monitored in the spectrometer. Nevertheless, a clear difference in the turbidity as a function of the collagen concentration could be observed in the plateau phases. The degree of fibril formation was not significantly influenced by the collagen concentration. 3.1.2. Influence of NaCl concentration With an increasing amount of sodium chloride from 0 to 100 mM (Fig. 2A and B) in the suspension, the turbidity plateau decreased. Sodium chloride had a significant influence (p < 0.001) on the degree of fibril formation (Fig. 2C). Increase of the salinity over 20 mM reduced the amount of jellyfish collagen fibrils. 3.1.3. Influence of temperature Comparing the collagen concentrations of 2.5 mg ml1 at 4 and at 25 °C (Fig. 3A) revealed slightly lower turbidity plateaus at 25 °C. A similar tendency was observed with different TRIS buffer concentration (data not shown). At 25 °C the amount of reassembled collagen was significantly (p < 0.05) diminished (Fig. 3B). Investigations of varying sodium chloride concentration did not show such a difference between the two tested temperatures. 3.2. Morphology of jellyfish collagen fibrils The morphology of reassembled jellyfish collagen fibrils was visualized by AFM. As shown in Fig. 4A, branched fibrils, varying in thickness and length, were observed. The small fibrils measured 32 ± 5 nm in width, 1.5 ± 0.3 nm in height and 469 ± 103 nm in length. On the other hand, large fibrils exhibited a width of 49 ± 6 nm, a height of 3.1 ± 0.5 nm and a length of 4871 ± 683 nm. Higher magnification (Fig. 4B and C) exhibited irregular banding patterns of the fibrils with 30–40 nm gaps between overlap regions occurring most frequently. TEM analysis of fibrils from jellyfish collagen (Fig. 5) revealed also a banding pattern; however, here a period of 67 nm was observed. Thick representatives of jellyfish collagen fibrils had a width of 53.4 ± 8.6 nm.

Table 1 Primers for RT-PCR. Marker Collagen I Collagen IIa Collagen X Aggrecan SOX9 b-Actin RUNX2

Primer sequences 0

Tannealing (°C) 0

For:5 -CCAGAAGAACTGGTACATCA-3 rev: 50 -CCGCCATACTCGAACTGGAA-30 For: 50 -GAACATCACCTACCACTGCAAG-30 rev: 50 -GCAGAGTCCTAGAGTGACTGAG-30 For: 50 -GCCCACTACCCAACACCAAGAC-30 rev: 50 -CCTGGCAACCCTGGCTCTC-30 For: 50 -ACCACCGAGCCAGAAAACCAGAC-30 rev: 50 -CCTCTGAGGGGAACAGCTCCAC-30 For: 50 -GTGCTCAAAGGCTACGACTG-30 rev: 50 -CGTTCTTCACCGACTTCCTC-30 For: 50 -GGACTTCGAGCAAGAGATGG-30 rev: 50 -AGCACTGTGTTGGCGTACAG-30 For: 50 -GGTAACGATGAAAATTATTCTGCTG-30 rev: 50 -CCGAGGTCCATCTACTGTAAC-30

Amplicon size (bp)

60

96

60

488

50

196

64

268

62

316

55

234

55

201



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Fig. 1. Influence of the final collagen concentration on fibril reassembly of jellyfish collagen: turbidity measurement at 4 °C (A) and 25 °C (B), and degree of fibril formation measured by Bradford assay (C).

Fig. 2. Influence of the NaCl concentration on fibril reassembly of jellyfish collagen: turbidity measurement at 4 °C (A) and 25 °C (B), and degree of fibril formation measured by Bradford assay (C).

Fig. 3. Influence of the reaction temperature on the fibril reassembly of jellyfish collagen exemplary for a final collagen concentration of 2.5 mg ml1: turbidity measurement (A) and degree of fibril formation measured by Bradford assay (B).

3.3. Characterization of porous jellyfish collagen scaffolds Porous 3-D jellyfish collagen scaffolds were engineered in different sizes (Fig. 6A) by means of lyophilization. SEM analysis (Fig. 6B) revealed an open, interconnected pore structure. A porosity of 98.2 ± 0.4% was assessed by measuring the pore volume with a helium pycnometer. DSC (Fig. 7) was used to determine the effect of chemical crosslinking on the denaturation temperature of jellyfish collagen. An increase of 12 K of the peak temperature was achieved by chemical cross-linking of jellyfish collagen with 1% EDC. Mechanical properties were evaluated under wet conditions, with scaffolds incubated in SBF. In static tests (Fig. 8A), a compressive modulus of 9.98 ± 0.93 kPa for 3-D jellyfish collagen scaffolds was measured. The compressive stress at 20% compression was 1.79 ± 0.20 kPa. Cyclic tests (Fig. 8B) revealed a highly elastic behavior of the jellyfish collagen scaffolds.

3.4. Viability and chondrogenic differentiation of hMSC in scaffolds from jellyfish collagen Initially, the cytotoxicity of jellyfish collagen was tested. Cells on polystyrene were cultivated with either scaffold extract medium or fresh medium (Fig. 9). There was no significant difference in the number of cells in both examined media, showing that jellyfish collagen did not release cytotoxic substances. Human mesenchymal stem cells were seeded on porous jellyfish collagen scaffolds and cultivated under chondrogenic stimulation for 21 days. Cell distribution was evaluated macroscopically by MTT staining (Fig. 10). Only viable cells metabolize the yellowish MTT into a dark blue formazan derivative. The staining showed viable cells throughout the whole scaffold. Gene expression of b-actin, collagen I, collagen IIa, collagen X, aggrecan, RUNX2 and SOX9 was investigated (Fig. 11A). The housekeeping gene b-actin and collagen I were expressed at similar levels on


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Fig. 4. AFM images (left: height image, right: amplitude image) of fibrillized jellyfish collagen. Insets in A and B represent the height scale. Panel C is a magnification of the marked section in B and was used for the section analysis.

According to SEM and cLSM images after 21 days of cultivation (Fig. 12A–C), a large number of cells were found on the seeding side of the scaffolds, which spread over the pores, similar to fibroblast phenotype. Yet in between, cells of a more spherical shape (indicated by arrows) were discovered which resemble the chondrogenic phenotype. Fig. 12D shows the opposing seeding side of the jellyfish collagen scaffold. Here only spherical cells are found, arranged in small clusters. 4. Discussion

Fig. 5. TEM image of stained jellyfish collagen fibrils. Bar shows 200 nm.

day 1 and day 21. Genes coding for collagen II, collagen X, aggrecan, RUNX2 and SOX9 were upregulated. Furthermore, sGAG concentration was biochemically quantified, showing a significant increase (p < 0.001) from day 1 to day 21 (Fig. 11B).

Sufficient regeneration of cartilage defects is still one of the major challenges in orthopedics. One strategy to meet this challenge is tissue engineering. The development of suitable matrices for hosting chondrogenically stimulated cells is a prerequisite for this purpose. In this study, we describe the manufacture of fibrillized jellyfish collagen to be used as a biomimetic matrix for cartilage tissue engineering. Collagen is a well-investigated biomaterial that has been studied since the beginning of the last century. Although fibril formation of collagen has been extensively examined, it is still not common to use fibrillized collagen for scaffold fabrication. Porous



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Fig. 6. (A) Scaffolds of different sizes and (B) SEM image of a 3-D jellyfish collagen scaffold.

Fig. 7. DSC measurement of jellyfish collagen without and with 1% EDC crosslinking.

collagen-based scaffolds are more frequently prepared by freezedrying solubilized collagen monomers. There are a few reports on fibrillized collagen matrices in the form of lyophilized sponges [37], hydrogels [28,38], films [39] or as coatings [40,41]. In the field of bone tissue engineering, some studies focus on mineralization of fibrillized collagen to produce a biomaterial which mimics the mineralized bone matrix [35,42–44]. Scaffolds made of fibrillized marine collagens derived from salmon skin have been reported only by a Japanese research group [45] and recently by us [34]. In contrast to structures of freeze-dried monomeric collagen, structures of fibrillized collagen mimic the natural extracellular matrix and can provide additional mechanical strength [46]. A challenge in the handling of marine collagens is their lower denaturation temperature compared to collagens from mammalian sources. For collagen derived from R. esculentum we determined a denaturation temperature of 32 °C by means of circular dichroism (data not shown). Consequently, we conducted the fibril formation studies at 4 and 25 °C, respectively, both temperatures being below the denaturation temperature of jellyfish collagen. The reaction temperature had a considerable influence on the

Fig. 9. Viability of human mesenchymal stem cells cultured with scaffold extract (scaffolds) or fresh medium (control). Number of viable cells was calculated from LDH activity after total lysis.

extent of fibril formation (Fig. 3), which was significantly increased (p < 0.05) at 4 °C compared to 25 °C. Wood and Keech [47] reported a similar temperature effect on the fibril formation of collagen from calf skin caused by different fibril widths of 500 and 1000 nm, respectively. Changes in the collagen concentration (Fig. 1) in the range of 0.5–2.5 mg ml1 did not significantly influence the percentage of fibrillized collagen, while the absorbance values of the turbidity plateaus increased with increasing collagen concentration. Increasing turbidity values of the plateau phase were also documented for salmon collagen type I in the range of 0.2–2.5 mg ml1 [34] and bovine collagen type II in the range of 0.2–0.5 mg ml1 [48]. Fibril formation of jellyfish collagen was further influenced by ionic strength (Fig. 2). With increasing sodium chloride concentration (0–100 mM) the absorbance values of the turbidity plateaus decreased, and consequently the amount of fibrillized collagen was reduced. This was a surprising result since sea water contains 455 mM sodium chloride [49], four times higher than the highest concentration investigated in this study. Based on those findings a

Fig. 8. Compressive stress–strain curve of 3-D jellyfish collagen scaffolds: static compression (A) and cyclic compression (B) of wet samples.


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Fig. 10. MTT staining of viable human mesenchymal stem cells cultured with chondrogenic supplements after 21 days of cultivation. Jellyfish collagen scaffolds without (left) and with (right) cells (A), and cross-section of a cell-seeded scaffold (B). Bars show 3 mm.

Fig. 11. Analysis of unstimulated hMSCs on jellyfish collagen scaffolds after 1 day of cultivation and chondrogenically stimulated hMSCs on jellyfish collagen scaffolds after 21 days of cultivation. RT-PCR expression analysis of different genes (A) and biochemical quantification of sulfated glycosaminoglycan concentration (B).

protocol was established for fibril formation of jellyfish collagen at a temperature of 4 °C: 5 mg ml1 collagen dissolved in hydrochloric acid was mixed 1:1 with 50 mM Tris buffer at pH 8, yielding a solution with the resultant pH of 7.4. Fibril formation was verified with AFM (Fig. 4) and TEM (Fig. 5) images showing the typical banding of collagen fibrils. It is well known that the glycosylation grade is inversely related to the fibril diameter [50]. This was already reported for jellyfish collagen with a glycosylation grade of 23–25 residues/1000 amino acids, leading to fibrils with a diameter of 10–30 nm [11]. Collagen from R. esculentum showed a lower glycosylation grade of 9 residues/1000 amino acids, which could explain the slightly thicker fibrils with diameters between 30 and 60 nm, which are in the range of native mammalian cartilage collagen type II fibrils (5–120 nm) [51]. The D-periodic banding pattern of 67 nm could not be clearly identified in AFM for jellyfish collagen. This is a well-known phenomenon of collagen fibrils with diameters below 30 nm [51]. For example, Tsai et al. [52] demonstrated a banding pattern of 48.6 nm in porcine cartilage collagen. In TEM, however, stained jellyfish collagen fibrils showed a periodic banding of 67 nm, which is typical for collagens of type I and II [53,54]. Nevertheless, banding of jellyfish collagen fibrils was less distinct, indicating that fibril association was slightly disordered. To sum up, jellyfish collagen is a fibrilforming collagen with a distinct banding pattern.

3-D porous scaffolds (Fig. 6) made of fibrillized jellyfish collagen were prepared by a freeze-drying technique, resulting in 98% porosity comparable to bovine collagen scaffolds produced by Dawson et al. [55]. The issue of the denaturation temperature being below 37 °C was solved by cross-linking with a carbodiimid derivative verified with DSC measurements (Fig. 7). Compressive modulus (Fig. 8) of jellyfish collagen scaffolds was increased 13-fold compared to salmon collagen scaffolds [34]. However, collagen scaffolds generally have, compared to other materials, limited mechanical properties, owing to the high porosity of 98% and the use of a natural protein. Cytotoxicity of jellyfish collagen (Fig. 9) was indirectly assessed by cultivation of cells with scaffold extracts. No significant difference in the proliferation rate of hMSCs cultivated with scaffold extracts or fresh medium was observed, indicating the absence of cytotoxic effects of the scaffold extracts. Similar non-cytotoxic effects on human fibroblasts and smooth muscle cells incubated with extracts of collagen of Stomolophus nomurai meleagris were obtained by Song et al. [17]. Direct cultivation of hMSCs under chondrogenic stimulation revealed viable cells (Fig. 10) and upregulation of chondrogenic markers on mRNA level (Fig. 11A). Other studies have demonstrated chondrogenic differentiation of MSCs in the 3-D environment of scaffolds. Collagen-based matrices were applied in particular by Bosnakovski et al. [28], who cultivated bovine MSCs in collagen type I and II gels, and by Chen and co-workers [56], who studied the chondrogenic differentiation of rabbit MSCs in porous scaffolds from bovine collagen type II. Both research groups reported in accordance a substantial increase of collagen type II and aggrecan mRNA after 3 weeks of cultivation. Additionally, we found collagen type X to be upregulated, which is also synthesized in hypertrophic chondrocytes and is therefore associated with hypertrophic induction [57]. However, some researchers express doubts as to whether an increased collagen type X transcript level in chondrogenically stimulated MSC cultures is really associated with hypertrophy. Mwale et al. [58] detected an increase of collagen type X mRNA earlier than collagen II mRNA in chondrogenically stimulated pellet cultures of hMSCs, suggesting that collagen type X is no reliable hypertrophy marker in stem cell differentiation. Similar results were reported by Jakobsen et al. [59], who observed a simultaneous upregulation of the chondrogenic markers SOX9 and aggrecan on the one hand and collagen type X on the

Fig. 12. SEM images (A, C, D) and cLSM image (B) of chondrogenically stimulated hMSCs cultivated on jellyfish collagen scaffolds for 21 days. (A–C) Cell-seeding side and (D) opposite side of scaffold. Bars show 50 lm.



other hand when cultivating chondrogenically stimulated hMSCs in hyaluronic acid scaffolds. In our experiments a tendency to hypertrophic development is furthermore indicated by the upregulation of RUNX2. An upregulated expression of RUNX2 in later chondrocytes regulates hypertrophy [60]. On the contrary, RUNX2 is expressed in bone-marrow-derived MSCs even without osteogenic or chondrogenic differentiation, as confirmed by immunofluorescent staining and gene expression analysis [61]. This is in accordance with our study, where RUNX2 was already expressed after seeding of the scaffolds. Redifferentiation of chondrocytes is accompanied by upregulation of collagen type II and downregulation of collagen type I expression. In our study chondrogenic differentiation of hMSCs led to an increased collagen type II expression; however, the mRNA level of collagen type I did not decrease. Other groups reported likewise no downregulation of collagen type I expression during chondrogenic differentiation of MSCs [59,62]. The chondrogenic induction was probably not sufficient to completely suppress the fibroblast-like MSC phenotype, which could be observed in SEM and cLSM (Fig. 12A–C). Chondrogenic differentiation of hMSCs was further indicated by a 23-fold increase of deposited sGAG in jellyfish collagen scaffolds after 21 days of cultivation (Fig. 11B). An increase of sGAG deposition was also observed in chondrocytes on collagen matrices [16,63]. Furthermore SEM images (Fig. 12) revealed a large number of hMSCs with a spherical phenotype, similar to the findings of Pieper et al. [63] for chondrocytes on collagen matrices. Obviously not all cells have differentiated towards the spherical phenotype yet, requiring further adjustments in chondrogenic stimulation and scaffold modification. However, the potential of fibrillized jellyfish collagen to support chondrogenically stimulated hMSCs was shown beyond dispute.

5. Conclusions This is the first report on fibrillized collagen scaffolds based on collagen from jellyfish R. esculentum. This jellyfish species is a prospective collagen source due to the similarity of its collagen to mammalian collagen type II. We analyzed different parameters influencing the fibril formation of jellyfish collagen. The use of collagen fibrils for producing a matrix mimics the cartilage extracellular matrix, which is quite often not regarded in other publications. Porous 3-D jellyfish collagen scaffolds with an interconnected pore structure were manufactured by freeze-drying and subsequent chemical cross-linking. We successfully engineered a cytocompatible matrix with the potential to support and maintain chondrogenic stimulation of human mesenchymal stem cells. Acknowledgements The authors wish to thank the German Federal Ministry for Education and Research (BMBF) for financial support (Regeneration with cell specific matrices, BMBF contract No. 01GN0962) and Prof. Martin Bornhäuser, Medical Clinic I, University Hospital Carl Gustav Carus, Dresden, for providing hMSCs. Special thanks to Dr. Silke Erdmann for her enlightening and encouraging discussions and support. We also thank Kristin Faulwasser and Rudolf Beiermeister for the preparation and fibril formation analysis. Furthermore, we are grateful for the excellent technical assistance of Sophie Brüggemeier and Ortrud Zieschang.

Appendix A. Figures with essential colour discrimination Certain figures in this article, particularly Figs. 4, 10, and 12, are difficult to interpret in black and white. The full colour images can

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be found in the on-line version, at http://dx.doi.org/10.1016/ j.actbio.2013.10.022.

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