Biomimetically Mineralized Salmon Collagen Scaffolds for Application

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Biomimetically Mineralized Salmon Collagen Scaffolds for Application in Bone Tissue Engineering Birgit Hoyer,*,† Anne Bernhardt,† Sascha Heinemann,‡ Ines Stachel,§ Michael Meyer,§ and Michael Gelinsky† †

Centre for Translational Bone, Joint and Soft Tissue Research, University Hospital and Medical Faculty Carl Gustav Carus, Fetscher Str. 74, Technische Universität Dresden, D-01307 Dresden, Germany ‡ Max Bergmann Center of Biomaterials and Institute for Materials Science, Technische Universität Dresden, Budapester Str. 27, D-01069 Dresden, Germany § Research Institute of Leather and Plastic Sheeting (FILK gGmbH), Meissner Ring 1-5, D-09599 Freiberg, Germany S Supporting Information *

ABSTRACT: Biomimetic mineralization of collagen is an advantageous method to obtain resorbable collagen/hydroxyapatite composites for application in bone regeneration. In this report, established procedures for mineralization of bovine collagen were adapted to a new promising source of collagen from salmon skin challenged by the low denaturation temperature. Therefore, in the first instance, variation of temperature, collagen concentration, and ionic strength was performed to reveal optimized parameters for fibrillation and simultaneous mineralization of salmon collagen. Porous scaffolds from mineralized salmon collagen were prepared by controlled freeze-drying and chemical cross-linking. FT-IR analysis demonstrated the mineral phase formed during the preparation process to be hydroxyapatite. The scaffolds exhibited interconnecting porosity, were sufficiently stable under cyclic compression, and showed elastic mechanical properties. Human mesenchymal stem cells were able to adhere to the scaffolds, cell number increased during cultivation, and osteogenic differentiation was demonstrated in terms of alkaline phosphatase activity.

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INTRODUCTION Degradable materials for bone replacement are needed for a variety of medical therapies. A promising approach is mimicking the natural extracellular matrix of bone tissue with composite materials consisting of type I collagen and hydroxyapatite (HA). Commonly, collagens from mammalian sources are used for this purpose, for example, bovine collagen.1−3 However, the clinical application of mammalian collagen bears the risk of infections with pathogens such as bovine spongiform encephalopathy. Another disadvantage that has been discussed recently is the disuse because of religious reasons.4 An alternative and presumably safer source is collagen from marine organisms, such as collagen from marine sponges5,6 or from fish skin.7,8 The potential for disease transmission is reduced because there is a wide evolutionary distance between fish and human beings. Furthermore, the habitats are quite different. Fish skin and bone are available in high amount as industrial byproduct of the fish industry. They are high in protein content and, therefore, a good source for collagen. While mammalian collagens have been comprehensively investigated within the last 20 years marine organisms are an ambitious new source for collagens. Recent publications mainly concentrate on the isolation and characterization of collagen © 2012 American Chemical Society

from various fish species, for example, salmon, shark, deep sea redfish,8−10 or marine sponges.5,11 There is a lot of potential for the preparation of scaffolds made of fish collagen. However, one disadvantage of fish-derived collagen is the low denaturation temperature. Salmon collagen, for example, is only stable below 19 °C,7 whereas shark collagen and collagen of red stingray denaturate above 30 °C.12,13 In either case, this is not sufficient for application at physiological temperature in human beings. However, the thermal stability of fish-derived collagen scaffolds can be improved by chemical cross-linking.14 Only few publications exist that deal with the preparation of scaffolds from marine collagens. In these studies, mainly salmon was used as the collagen source. Nagai and co-workers14,15 evaluated salmon collagen scaffolds for the culture of human periodontal ligament cells. The same group also applied salmon collagen gels to prepare vascular grafts,16,17 salmon collagen films for wound dressings,18 and recently, salmon collagen sponges for osteochondral repair.19 Matsumoto et al.20 prepared sponges made of salmon collagen and elastin as artificial dermis. However, until now there are no reports on Received: December 13, 2011 Revised: February 5, 2012 Published: February 24, 2012 1059

dx.doi.org/10.1021/bm201776r | Biomacromolecules 2012, 13, 1059−1066

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solutions to receive final concentrations of 45 and 50 mM, respectively. The pH was adjusted to 7.4 by addition of TRIS buffer and Sørensen phosphate buffer with final concentrations of 25 and 27 mM. All solutions were precooled to 4 °C before starting the reaction and the mixture was stirred at 4 °C for 12 h. Under these conditions, the collagen fibril reassembly and the formation of nanocrystalline HAP occurred simultaneously. Mineralized salmon collagen formed a cloudy white precipitate and was collected by centrifugation for 20 min at 5000 g and 4 °C. For preparation of porous scaffolds, the mineralized salmon collagen was resuspended in a small amount of the supernatant and stirred for 1 h. This suspension was poured into the cavities of 48-well cell culture dishes. After freezing at a speed of 1 K/ min, lyophilization was conducted for 24 h to achieve sponge-like porous scaffolds. For stabilization, the scaffolds were then cross-linked with a 1% (weight %) solution of N-(3-dimethylaminopropyl)-N′ethylcarbodiimide (EDC, Fluka, Germany) hydrochloride in 80% (vol %) ethanol for 12 h, followed by thorough rinsing in deionized water, 1% glycine solution, once again in water, freezing at a freezing rate of about 1 K/min, and finally, freeze-drying3 (Alpha 1−2, Christ, Germany). Fourier Transform-Infrared (FT-IR) Spectroscopy. Freezedried collagen samples were embedded in KBr and analyzed by FTIR spectroscopy using a Perkin-Elmer FTS 2000. The detected spectra were baseline-corrected and flattened by using the Savitzky-Golay algorithm with nine supporting points. Atomic Force Microscopy (AFM). Suspensions of fibrillized collagen were incubated on mica (Plano) for 30 min, followed by rinsing with deionized water, and drying on air. AFM measurements were carried out in tapping mode on air using a Nanoscope IIIa Bioscope (Digital Instruments/Veeco, U.S.A.) and aluminum reflexcoated silicon tips (force constant 40 N/m; Budget Sensors, Bulgaria) with imaging speeds of 1.2 Hz, scanning 512 lines. Height and amplitude data was captured simultaneously. Imaging fields were usually chosen to be 20 × 20 μm or 5 × 5 μm. A minimum of 50 fibrils were counted for determination of fibril length, width, and height. Only entire fibrils were counted. For fibrils that codeposited in patches, fibril ends were counted. For bent fibrils, the straight length was taken to be the addition of the bent lengths. Height measurements were not taken where fibrils crossed each other, but only where a single fibril was deposited on the mica surface. Scanning Electron Microscopy (SEM). SEM was carried out using a Philips XL 30/ESEM with field emission gun, operated in SEM mode. All samples were fixed on carbon pads and sputter-coated with gold. Cell-seeded scaffolds were previously washed twice with PBS and fixed in 3.7% formaldehyde in PBS. After dehydration in graded series of ethanol the samples were critical point dried (BAL-TEC CPD 30, Liechtenstein). Loss on Ignition. About 100 mg dehydrated mineralized collagen was weighed into a melting pot. Pyrolysis of the organic substance was carried out in a muffle furnace (TC 405/20, Padelttherm, Germany) at air atmosphere and at a temperature of 1000 °C for 1 h. The remaining materials were weighed again and lost percentage as a measure of the organic content was calculated. Three samples of mineralized collagen were subjected to this procedure. Porosity. Mineralized salmon collagen scaffold were measured (height and diameter) and weighed before and after loss on ignition (see above). With this data the density was calculated. Loss on ignition revealed the percentage of collagen and mineral in the scaffold. This ratio was used to calculate the relative density of a solid material using the densities 1.343 g/cm3 for collagen22 and 3.16 g/cm3 for hydroxyapatite.23 The porosity was calculated with the ratio of geometric to relative density. Porosity was verified experimentally by usage of a helium pycnometer (Ultrapyc 1200e, Quantachrome Instruments, U.S.A.). Mechanical Measurements. Uniaxial cyclic loading experiments were carried out using an Instron 5566 testing machine. Five cylindrical scaffolds of salmon collagen (diameter 11 mm, height 8 mm) were incubated in deionized water for 24 h before starting the experiments. Wet samples were compressed to 50% of the initial

composite materials from fish collagen containing an inorganic mineral phase like hydroxyapatite. In the present study, collagen from salmon skin was used for the first time to fabricate porous scaffolds from mineralized collagen for application in bone tissue engineering. The biomimetic mineralization of collagen, which has been developed in our group, is a promising procedure to generate bone-like collagen/hydroxyapatite composites.2,3 In this biomimetic approach, collagen fibril reassembly and precipitation of nanocrystalline hydroxyapatite occur simultaneous to obtain a structure that is very similar to that of natural bone.2 This procedure has been elaborated for bovine collagen3 and had to be adapted to the special requirements of salmon collagen. After finding suitable parameters for salmon collagen fibril reassembly with regard to collagen concentration, temperature, and ionic strength, porous collagen/HA nanocomposite scaffolds were prepared by synchronous mineralization and fibril reassembly, followed by controlled freeze-drying and chemical cross-linking. Scaffolds were characterized in terms of microstructure, collagen-to-mineral ratio, chemical nature of mineral phase, and mechanical stress. First successful cytocompatibility tests were performed using human mesenchymal stem cells (hMSC).

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MATERIALS AND METHODS

Collagen. Soluble type I like collagen was extracted from skin of Atlantic salmon. Freshly obtained salmon skin was intensively washed in icy water prior to manual cutting into small pieces and extraction in 0.1 M acetic acid at 4 °C for 48 h. After filtration, collagen was precipitated by the addition of sodium chloride. The pellet was then collected by centrifugation at 4000 g for 20 min at 4 °C and resuspended in distilled water. After dialysis against deionized water, the collagen was lyophilized and stored at −20 °C until required. The purity of the collagen was investigated according to ASTM F2212 and full conformity was ascertained. Prior to use, the lyophilized collagen was dissolved in 0.01 M HCl under constant stirring at 4 °C for at least 2 h to receive a stock solution with a concentration of 5 mg/mL. Fibril Formation. To analyze the reassembly process of salmon collagen, 500 μL of the collagen stock solution was mixed with 500 μL of aqueous 50 mM tris-(hydroxymethyl)-aminomethane (TRIS, Fluka, Germany) solution of pH 8 containing various amounts of NaCl. After mixing, the final solutions were transferred to PMMA cuvettes and absorbance at 313 nm was monitored up to 40 min using a UV/vis spectrophotometer Cary 50 Bio (Varian, Germany). Due to the use of different salmon collagen batches, total turbidity varied between the experiments. Collagen concentrations were varied from 0.2 to 2.5 mg/ mL (50 mM NaCl). The influence of ionic strength on fibril formation of a 2.5 mg/mL collagen solution was studied by adjusting final concentrations of 25, 50, 75, 100, and 200 mM NaCl. Fibril formation was carried out at temperatures of 4, 10, and 15 °C at a NaCl concentration of 50 mM. After fibrillation, the collagen suspensions were centrifuged (S3K30, Sigma, Germany) for 15 min at 10000 g, and 4 °C. The protein concentration of the supernatant was determined using a modified Bradford assay.21 A total of 5 μL of supernatant were mixed with 250 μL Bradford reagent (Sigma-Aldrich, U.S.A.) containing 0.035 mg/mL SDS (Carl Roth, Germany). After 15 min, the absorption at 590 nm was measured using a microplate reader SpektraFluorPlus (Tecan, Switzerland). For calibration, graded series of ultrapure bovine collagen type I solutions (from tendon and skin; Sigma, U.S.A.) were used. The degree of fibril formation was calculated by the ratio of fibrillized collagen to total initial collagen. The graphs show the mean ± standard deviation (n = 3). Scaffold Preparation. Based on our already established protocol for the biomimetic mineralization and synchronous refibrillation of soluble bovine collagen,2,3 the formation of mineralized salmon collagen was achieved by the following, adapted procedure: Salmon collagen stock solution (5 mg/mL) was mixed with CaCl2 and NaCl 1060


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Figure 1. Effect of collagen concentration on the fibril reassembly of salmon collagen: (A) turbidity measurement and (B) degree of fibril formation measured by Bradford assay.

Figure 2. Effect of temperature on the fibril reassembly of salmon collagen: (A) turbidity measurement and (B) degree of fibril formation measured by Bradford assay. height with a velocity of 0.3 mm/s for 50 cycles.3,24 The results are presented as mean ± standard deviation. Cell Culture. HMSC, isolated from bone marrow aspirate of two voluntary healthy male donors (both age 37), was kindly provided by Professor Martin Bornhäuser and co-workers, Medical Clinic I, Dresden University Hospital Carl Gustav Carus. Expansion of the cells was performed in Dulbecco’s modified Eagle’s medium (DMEM) low glucose (Biochrom, Germany), containing 10% fetal calf serum, 100 U/mL penicillin, and 100 μg/mL streptomycin (Biochrom, Germany) at 37 °C in a humidified, 7% CO2/93% air incubator. Cells in passage 5 were used for the experiments. All procedures were approved by the Ethical Commission of the Medical Faculty of Technische Universität Dresden. For cell culture experiments, cylindrical mineralized salmon collagen scaffolds with 6 mm in diameter and 3 mm in height were used. Prior to cell seeding, the samples were incubated in 70% ethanol and then in cell culture medium for 24 h. After removing the excess liquid with sterile filter paper the samples were placed in 48 well culture dishes and colonized with 1.6 × 105 cells. As a control hMSC (1 × 104) were seeded on 24well polystyrene culture dishes. After 24 h, cell-seeded samples were transferred to fresh culture dishes. After four days of cultivation, cell culture medium of half of the samples was supplemented with 10−7 M dexamethasone, 10 mM βglycerophosphate, and 0.05 mM ascorbic acid 2-phosphate (osteogenic supplements = OS+, all chemicals from Sigma). Medium was changed twice weekly and cell-seeded scaffolds were cultured for up to 21 days. After 1, 7, 14, and 21 days, samples (n = 3) were placed on filter paper, washed twice with PBS, and frozen in 2 mL Nalgene tubes containing six beads of stainless steel (Peqlab, Germany) at −80 °C. Cells on polystyrene culture dishes were taken after 1, 7, 14, and 21 days. The cell layer was washed twice with PBS and the plates were frozen until further analysis. Determination of Alkaline Phosphatase (ALP) Activity and DNA Content. A volume of 450 μL of ice-cold PBS was added to frozen cell-seeded scaffolds, and the samples were homogenized (2 × 10 s at 5900 rpm) using a Precellys24 apparatus (Peqlab, Germany). After homogenization, 50 μL of 10% Triton X-100 in PBS were added,

the mixture was shaken shortly, and then incubated on ice for 50 min. Cells on polystyrene culture dishes were defrosted for 20 min, followed by 50 min of cell lysis with 1% Triton X-100 on ice. During cell lysis samples were sonicated for 10 min in an ultrasonic bath (Bandelin, Sonorex TK 52, Germany). An aliquot of each cell lysate was added to ALP reaction buffer, containing 1 mg/mL p-nitrophenyl phosphate (Sigma, U.S.A.), 0.1 M diethanolamine, 1% Triton X-100 (pH 9.8), and 1 mM MgCl2, and the mixture was incubated at 37 °C for 30 min. Finally, 1 M NaOH was added to stop the enzymatic reaction. After centrifugation at 16000 g for 10 min, the supernatant was transferred to a microtiter plate and the absorbance was measured at 405 nm with the microplate reader. A calibration line was obtained from different dilutions of 1 mM pnitrophenol. Another aliquot of the cell lysate was mixed with Picogreen ds DNA quantification reagent (Molecular Probes, U.S.A.) 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 cell number using a calibration line. Graphs show mean ± standard deviation (n = 3).

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RESULTS

Effect of Collagen Concentration, Temperature, and Ionic Strength on Fibril Reassembly of Monomeric Salmon Collagen. Effect of collagen concentration: Collagen concentrations ranging from 0.2 to 2.5 mg/mL were evaluated and turbidity changes at 313 nm were monitored for 20 min at 4 °C (Figure 1A). A rapid rise of turbidity was observed for collagen concentrations of 1 mg/mL and above, whereas for 0.2 and 0.5 mg/mL collagen concentration the turbidity change is slightly retarded. The average degree of fibril formation (Figure 1B) for collagen concentrations 1.0, 1.5, and 2.5 mg/mL was 60.5 ± 2.5%, whereas collagen concentrations 0.2 and 0.5 mg/ mL yielded a lower degree of fibril formation. 1061


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Figure 3. Effect of NaCl concentrations on the fibril reassembly of salmon collagen at 4 °C: (A) turbidity measurement and (B) degree of fibril formation measured by Bradford assay.

the strong peak at 1541 cm−1 and the weak peaks at 1562 and 1502 cm−1. The peaks at 1281 and 1244 cm−1 were identified as N−H deformation of amide III. The FT-IR spectrum of mineralized salmon collagen showed the bands of both organic and the mineral phase. In addition to the before mentioned amide bands, the phosphate ion bands caused by ν1 P−O symmetric stretching mode (961 cm−1), ν2 O−P−O bending at 471 cm−1, ν3 P−O antisymmetric stretching mode (1027 cm −1 ), and the ν 4 O−P−O antisymmetric and symmetric stretching modes (601 and 561 cm−1, respectively) were recognized.25,26 The FT-IR spectrum of mineralized salmon collagen shows the same phosphate peaks that were also found in mineralized bovine collagen.2 The morphology of the reassembled salmon collagen fibrils was observed by AFM imaging. The fibrils did not join during reassembly and sample preparation but, rather, were separated (Figure 5), allowing additional analysis of length, height, width,

Effect of temperature: Fibril reassembly of 2.5 mg/mL salmon collagen was carried out at temperatures of 4, 10, and 15 °C. Turbidity measurements showed an acceleration of fibril formation with rising temperature (Figure 2A). The final absorbance value after 20 min of fibril reassembly was higher at 10 and 15 °C, compared to 4 °C. Nevertheless, the percentages of fibril formation determined were similar at all examined temperatures (Figure 2B). Effect of ionic strength: Ionic strength was varied during the reassembly procedure by adding NaCl with final concentrations between 25 and 200 mM. With increasing NaCl concentrations, higher values for maximum turbidity were obtained (Figure 3A). Concomitantly, the turbidity raise was delayed with increasing NaCl concentrations. This effect was particularly pronounced in the case of 200 mM NaCl, where fibril reassembly was delayed, so that the final absorbance was not reached even after 40 min of incubation. In contrast to the findings of turbidity measurements, the degree of fibril assembly calculated using the modified Bradford assay showed no significant influence by NaCl in the range between 25 and 100 mM. Only at 200 mM NaCl the degree of fibril formation was slightly lowered. Composition and Structure of Mineralized Salmon Collagen Scaffolds. Figure 4 shows the FT-IR spectra of

Figure 5. AFM height images and fibril profile of reassembled salmon collagen.

and banding pattern. The average length of the fibrils was in the range of 9.2 ± 4.1 μm, whereas the widths and heights in the middle of the hastate fibrils taken from corresponding height images were calculated to be in the range of 150.2 ± 26.9 and 9.1 ± 1.8 nm, respectively. Higher magnification revealed a straightforward banding pattern with predominantly regular sections exhibiting periodicity (red arrows) in the range of 67 nm, similar to that of mammalian collagen type I. Multiperiodicity, as observed for marine sponge collagen, was not noticed.5 The ratio between collagen and mineral was investigated by loss on ignition. On average, a loss of 46.7 ± 0.21% was measured, indicating a scaffold composition of about 47% salmon collagen and 53% mineral phase. Bovine mineralized

Figure 4. FT-IR spectra of pure (upper, gray line) and mineralized salmon collagen (lower, black line).

mineralized and pure fibrillar salmon collagen. The broad band between 3700 and 3100 cm−1 resulted from an overlap of different vibrations of units with hydrogen, such as O−H stretching of adsorbed water or N−H stretching (amide A). Carbon dioxide in the surrounding air caused the weak peaks between 2360 and 2340 cm−1. The FT-IR spectra of pure salmon collagen showed strong absorption peaks at 1660 and 1631 cm−1 that can be assigned to CO stretching of amide I bands. N−H deformations of amide II (1550−1500 cm−1) were 1062


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salmon collagen scaffolds reached values of 70% of the bovine samples. Figure 7B shows the elastic behavior of mineralized salmon collagen scaffolds with a linear elasticity to about 17% compression; the moduli indicate similar elastic properties of mineralized salmon and bovine collagen scaffolds. Adhesion, Proliferation, and Osteogenic Differentiation of hMSC. HMSC attached to the material and spread onto the surface. SEM investigations after 15 days of cultivation revealed a confluent cell layer on the scaffold surface (Figure 8A,B). Figure S1 in the Supporting Information shows cells spreading over pores in the cross-section of the mineralized salmon collagen scaffold. DNA content after lysis of cell-seeded samples and cells cultivated on polystyrene was correlated with cell number. During the first week of cultivation, the cell number on mineralized salmon collagen samples remained on the same level, followed by an increase of cell number during the second week. During the third week of cultivation only the amount of noninduced cells on salmon collagen scaffolds continued to rise (Figure 9A). These findings were confirmed by the determination of total LDH activity (data not shown). Control cells on polystyrene dishes showed steady proliferation over the observed culture period. Cell number on polystyrene increased 5.5-fold for noninduced and 7.5-fold for osteogenically induced cells from d1 to d21 of cultivation. Cell number on mineralized salmon collagen multiplied 3.6-fold and 2.9-fold, respectively, after three weeks of cultivation. The specific ALP activity (ALP activity related to cell number determined from DNA content) of osteogenically induced hMSC on mineralized salmon collagen scaffolds raised from day 7 to day 14 and 21 of cultivation (Figure 9B). After 14 and 21 days of cultivation, specific ALP activity of cells on mineralized salmon collagen was slightly higher compared to cells on polystyrene; however, these differences were not statistically significant. Specific ALP activity of hMSC that was not treated with osteogenic supplements both on polystyrene and mineralized salmon collagen scaffolds stayed almost unchanged during the whole cultivation period.

scaffolds reached a higher mineral content of up to 62 wt % under similar conditions.2 Based on this collagen-to-mineral ratio, a porosity of 96.9 ± 0.29% has been calculated which was verified by helium pycnometer measurement (96.5%). The high porosity is supported by the SEM image (shown in Figure 6),

Figure 6. SEM image of mineralized salmon collagen scaffold surface.

which demonstrates interconnected pores with diameters up to 200 μm. Because of the similarity to mineralized bovine collagen scaffolds,3 no quantitative pore size measurements were conducted. Mean pore size in mineralized bovine collagen scaffolds was calculated to be 180 ± 12 μm using data from microcomputed tomography using synchrotron irradiation and supported by Brunauer−Emmett−Teller (BET) measurements revealing pores in the range of 80−200 μm.3 Mechanical Properties of the Scaffolds. For the mechanical characterization of the scaffolds, cyclic compression tests were carried out using three types of scaffolds (Figure 7A): pure and mineralized salmon collagen, as well as mineralized bovine collagen as a reference.24 Scaffolds from bovine mineralized collagen were prepared as previously described.3 Due to the fact that all three types of scaffolds do not fail in the wet (swollen) state when being compressed, the stress at distinct levels of deformation was determined and compared to each other. The quantitative results (Table 1) indicate a strengthening effect of collagen mineralization. Mineralized salmon collagen reached a 67 times higher value for maximum compressive stress σd,max at 50% deformation than the one measured for the pure collagen. The parameters σd,max and σd,end of mineralized

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DISCUSSION Simultaneous mineralization and fibril reassembly of bovine collagen type I is a well-established method for the preparation of a material, which combines collagen fibrils with nanocrystalline HA in an intimate connection, resembling the composition of natural bone matrix.2,3 The aim of the present study was to

Figure 7. Cyclic measurements (50 cycles uniaxial loading up to 50% deformation): (A) compressive stress over time comparing scaffolds of mineralized bovine (blue line) and salmon collagen (orange line) and pure salmon collagen scaffolds (black line); (B) compressive stress over strain of mineralized salmon collagen. 1063


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Table 1. Results of Cyclic Compression Tests under Wet Conditions sample

σd,20a (kPa)

σd,maxb (kPa)

σd,endc (kPa)

Ed (kPa)

pure salmon collagen mineralized salmon collagen mineralized bovine collagen

0.30 ± 0.36 4.95 ± 0.61 12.59 ± 7.54

2.40 ± 1.99 27.81 ± 2.60 42.07 ± 4.87

1.04 ± 0.61 17.88 ± 1.83 25.37 ± 2.73

0.77 ± 0.56 23.03 ± 3.73 24.37 ± 3.14

σd,20: compression stress at 20% compression. bσd,max: maximum compressive stress in the first cycle at 50% compression. cσd,end: average compressive stress over the last five cycles. dE: Young’s modulus.

a

Figure 8. SEM images of hMSC cultivated on mineralized salmon collagen for fifteen days (magnifications: (A) 150×, (B) 1000×).

Figure 9. (A) Proliferation of hMSC with osteogenic (+) and without osteogenic (−) stimulation determined by DNA quantification. (B) Specific ALP activity of hMSC (PS, polystyrene; SC, mineralized salmon collagen scaffolds).

Rising concentrations of NaCl strongly affected the fibril formation rate as well as final turbidity. A similar effect was described by Delorenzi et al.,28 who studied the effect of monovalent anions on in vitro fibrillogenesis of bovine collagen I. They reported a decrease of fibrillogenesis rate, as well as an increase of final turbidity upon addition of Cl−, and explained this phenomenon by the stabilization of intermediate states in the pathway of soluble collagen to fibrils. Yunoki and coworkers7 also reported a decrease of fibril formation rate at high NaCl concentrations (140 mM), as well as a reduced maximum turbidity for collagen I of pacific chum salmon. In contrast, Bae et al.13 detected only slight reduction in fibril formation rate with increasing NaCl concentration (140 mM) for collagen I of red stingray. Further investigation of salmon collagen using AFM showed typically elongated fibrils after collagen fibrillation. The fibril structures were within the size range of characteristic Dperiodicity of collagen type I with a period of D = 67 nm.29 Based on the results of fibril formation studies, a protocol for the simultaneous mineralization and fibrillation of salmon collagen was developed. This process is well-investigated for bovine collagen.2,3 It has been shown that the synchronous

examine the simultaneous mineralization and fibrillation of skin collagen from Atlantic salmon and to develop porous scaffolds from biomimetically mineralized salmon collagen for perspective bone engineering application. Salmon collagen is an alternative source since the potential for disease transmission is reduced. In view of the fact that fish skin is a byproduct of fish industry availability of salmon collagen is given at large scale. Challenging in this study of mineralized salmon collagen was the lower denaturation temperature compared to collagen from mammalian sources; for example, Yunoki et al.7 reported 18.6 °C for Pacific chum salmon. Therefore, the well-known process chain of bovine collagen mineralization had to be adapted to a temperature below 19 °C. Consequently, the effect of temperature, collagen concentration, and ionic strength on the rate of fibril formation as well as the degree of fibril formation was examined first. Initial collagen concentrations of 0.2 and 0.5 mg/mL lead to a retarded turbidity change as well as a reduced degree of fibril formation. Comparable results were reported by Raspanti et al.27 who detected a remarkable delay of turbidity-time curve during fibrillation of 0.1 mg/mL bovine collagen compared to higher collagen concentrations. 1064


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mal stem cells was observed upon cultivation on mineralized salmon collagen scaffolds. Other studies likewise demonstrated adhesion and proliferation of different cell types to salmon collagen gels, films, and scaffolds.14−16,33 Moreover, Kawase et al.33 reported an increased ALP activity in response to osteoinduction in periostal sheet cultures grown on salmon collagen-coated polymer meshes. Scaffolds from mineralized bovine collagen have been shown to support adhesion, proliferation, and osteogenic mineralization of human mesenchymal stem cells34−36 and fast remodeling in vivo, as demonstrated in a small animal model.37 The present results suggest a similar potential of salmon-derived mineralized collagen scaffolds for the application in bone regeneration.

collagen biomineralization process is initiated by the formation of an amorphous calcium phosphate phase, precipitating only at the emerging collagen fibrils, which act as a template for mineralization. The amorphous mineral phase slowly converts into nanocrystalline hydroxyapatite, tightly connected to the collagen fibrils. Biomineralization of bovine collagen is accomplished at physiological temperature for mammalians at 37 °C. Our fibril formation studies of salmon collagen insisted on performing the biomineralization at 4 °C, since higher temperatures lead to a very fast fibrillation, even lacking growth phase at 15 °C. Higher temperatures were not applied because of the low denaturation temperature of salmon collagen type I. It is anticipated, that the addition of further ions (calcium and phosphate) to the fibrillation reaction will cause changes in the fibrillation kinetics, however, there is no evidence for this given in the present study, because turbidity measurements would be disturbed by the calcium phosphate precipitation process. The optimal parameters were chosen from the fibril formation studies regarding temperature, collagen concentration and NaCl concentration and varied concerning the collagen-tocalcium and phosphate ratio to obtain biomimetically mineralized salmon collagen. The concentration of 45 mM CaCl2 and 27 mM phosphate buffer resulted in stable scaffolds with a fairly high mineral content, not far from that of natural bone matrix. Furthermore, the calcium-to-phosphate ratio of the mixture of those two solutions is 1.67, which is similar to that of stoichiometric hydroxyapatite. Scaffolds from mineralized collagen were characterized by FT-IR to analyze the chemical nature. The relevant amide stretching in collagen type I of a hydroxyapatite-collagen nanocomposite was investigated by Chang et al.25 The observed amide I bands reflect the triple helical structure of collagen, confirming that the secondary structure of salmon collagen is not altered by the mineralization process, which was observed for bovine collagen before.2 For identification of the mineral phase, the phosphate ion vibration modes were investigated. Typical stretching and bending modes caused by phosphate ions in an apatitic environment2,25,26 were found. Therefore, the precipitated mineral phase could be clearly identified as hydroxyapatite with interlayers of octacalcium phosphate, also shown by Bradt et al.2 for mineralized bovine collagen. Those results show that also salmon collagen can be used as a template for biomineralization. Mineral content of the mineralized salmon collagen scaffolds was determined to be 53%. This was slightly different from the composition of natural bone with about 70% inorganic proportion, mainly carbonate-rich hydroxyapatite.30,31 Cyclic mechanical tests revealed that mineralized salmon collagen scaffolds could withstand 50 loading cycles (50% compression) under wet conditions, showing that the handling of the scaffolds is also possible in the wet state. The relative low compressive stress under wet conditions, similar mechanical properties are also seen in porous collagen−silica or collagen− silica−hydroxyapaptite scaffolds,32 is a compromise to the porous structure, with interconnective pores of up to 200 μm in diameter, the high porosity of 97%, and the composition mimicking the natural extracellular matrix of bone. First cytocompatibility tests showed that scaffolds from mineralized salmon collagen are suitable for the cultivation of human mesenchymal stem cells. SEM of cell-seeded scaffolds revealed good cell attachment and cell number, quantified by DNA content, increased during cultivation for three weeks. Furthermore, osteogenic differentiation of human mesenchy-

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CONCLUSIONS This is the first report on the simultaneous mineralization and fibril reassembly of salmon collagen. The adaption of a wellknown biomimetic mineralization process of bovine collagen for marine collagen with a much lower denaturation temperature has succeeded. Porous scaffolds from mineralized salmon collagen were obtained by freeze-drying and chemical crosslinking. It was confirmed by FT-IR that the mineral phase formed during the preparation process was hydroxyapatite. The scaffolds revealed an interconnected porosity and were stable under cyclic compression. Cell culture experiments demonstrated the cytocompatibility of the newly developed composite material. HMSC were able to adhere, proliferate, and differentiate into the osteoblastic lineage.

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ASSOCIATED CONTENT

S Supporting Information *

SEM images of cross sections of cell-seeded mineralized salmon collagen scaffolds (Figure S1) are added. This material is available free of charge via the Internet at http://pubs.acs.org.

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AUTHOR INFORMATION

Corresponding Author

*Phone: +49-351-458-6694. Fax: +49-351-458-7210. E-mail: [email protected]. Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS We are grateful to O. Zieschang and S. Brüggemeier for their support in scaffold preparation and cell culture. The authors would like to thank the German Federal Ministry for Education and Research for funding the project RECEM (Regeneration with cell specific matrices, BMBF Contract No. 01GN0962).

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REFERENCES

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