- Email: [email protected]
Bio-mimetic composite scaffold from mussel shells, squid pen and crab chitosan for bone tissue engineering
International Journal of Biological Macromolecules 80 (2015) 445–454
Contents lists available at ScienceDirect
International Journal of Biological Macromolecules journal homepage: www.elsevier.com/locate/ijbiomac
Bio-mimetic composite scaffold from mussel shells, squid pen and crab chitosan for bone tissue engineering Amin Shavandi a,∗ , Alaa El-Din A. Bekhit a , M. Azam Ali b , Zhifa Sun c a b c
Department of Food Sciences, University of Otago, Dunedin, New Zealand Department of Applied Sciences, University of Otago, Dunedin, New Zealand Department of Physics, University of Otago, Dunedin, New Zealand
a r t i c l e
i n f o
Article history: Received 5 March 2015 Received in revised form 2 July 2015 Accepted 8 July 2015 Available online 15 July 2015 Keywords: Squid pen Chitosan Hydroxyapatite
a b s t r a c t In the present study, chitosan/hydroxyapatite (HA)/-tircalcium phosphate (-TCP) composites were produced using squid pen derived chitosan (CHS) and commercial crab derived chitosan (CHC). CHS was prepared from squid pens by alkaline N-deacetylation. HA and -TCP were extracted from mussel shells using a microwave irradiation method. Two different composites were prepared by incorporating 50% (w/w) HA/(-TCP) in CHS or CHC followed by lyophilization and cross-linking of composites by tripolyphosphate (TPP). The effect of different freezing temperatures of −20, −80 and −196 ◦ C on the physicochemical characteristics of composites was investigated. A simulated body fluid (SBF) solution was used for preliminary in vitro study for 1, 7, 14 and 28 days and the composites were characterized by XRD, FTIR, TGA, SEM, -CT and ICP-MS. Porosity, pore size, water uptake; water retention abilities and in vitro degradations of the prepared composites were evaluated. The CHS composites were found to have higher porosity (62%) compared to the CHC composites (porosity 42%) and better mechanical properties. The results of this study indicated that composites produced at −20 ◦ C had higher mechanical properties and lower degradation rate compared with −80 ◦ C. Chitosan from the squid pen is an excellent biomaterial candidate for bone tissue engineering applications. © 2015 Elsevier B.V. All rights reserved.
1. Introduction In recent years, much attention has been paid to marine byproducts, scoping their cost-effective processing schemes and their potential for production of high-value products. Natural polymers like chitin, chitosan and calcium phosphate (CaP) compounds can be obtained from waste marine products [1]. Because of their intrinsic properties such as biocompatibility, biodegradation and antimicrobial properties, these natural materials have important biomedical applications [2,3]. In the last two decades, many reports have been published on chitin and chitosan applications in drug delivery, tissue engineering, skin and bone grafting [2,4,5]. Chitosan is a biopolymer consisting of (1,4)-2-amino-2-deoxy-d-glucose units that is obtained by N-deacetylation of chitin under alkaline condition. Chitin can be sourced and extracted from a diverse range of natural organisms, including molluscs, fungi, insects, crustaceans and algae [6]. Chitin exists in three different allomorphic forms depending on the sources of the compound. Most chitins, including
∗ Corresponding author. E-mail address: [email protected] (A. Shavandi). http://dx.doi.org/10.1016/j.ijbiomac.2015.07.012 0141-8130/© 2015 Elsevier B.V. All rights reserved.
crustaceans and insect chitin are in alpha (␣) form which has a two chain antiparallel structure. However, squid pen and some diatoms have beta () chitin which has one chain parallel structure and in gamma (␥) chitin, the biomolecular chains are arranged randomly in which two parallel chains and one antiparallel chain form the polymeric structure [7]. Alpha-chitin and chitosan are commercially available products and are produced normally from shrimp or crab shell. Chitin/chitosan from the squid pen has a structure that is low packed and has weak intermolecular hydrogen bonds. These properties makes it chemically more reactive compared to the heavily packed and strong molecular structure of ␣-chitin/chitosan from shrimp and crab shells [8–10]. In addition, -chitin/chitosan can incorporate water molecules in its structure and forms crystalline structure leading to higher ability to uptake and hold water more than the alpha form, which is advantageous in biomedical applications [11,12]. The source of chitosan can affect its purity, molecular weight, chain length, degree of deacetylation, density, viscosity, solubility, water retention capacity and distribution of the amino/acetamide groups. All these characteristics affect the physicochemical properties of chitosan and therefore, its application [13]. A proper biocomposite should be prepared strategically in a way to have a suitable geometry and pore size, have a
446
A. Shavandi et al. / International Journal of Biological Macromolecules 80 (2015) 445–454
required mechanical strength, support cell attachment in vitro, and have tuneable biodegradable properties [14]. In this work, chitosan was isolated from endoskeleton of New Zealand arrow squid pen (Nototodarus sloanii) (CHS). In addition, a commercially available crab chitosan (CHC) was used for comparison with CHS. The HA and -TCP were prepared using waste green mussel shells. Then CHS, CHC and HA, -TCP were processed to produce biocomposites using freezing at different temperatures (−20, −80 and −196 ◦ C) and lyophilization processes. The produced biocomposites were evaluated to scope their potential for bone tissue engineering applications. To improve the mechanical properties of the composites in this study, tripolyphosphate (TPP) was used as an ionic cross-linker and glycerol was used as plasticizing agent. 2. Materials and methods 2.1. Materials Hydroxyapatite (Ca10 (PO4 )6 (OH)2 ) and -TCP (Ca3 (PO4 )2 ) powders were synthesized from waste mussel shells as previously reported [15,16]. Acetic acid, ethanol, and NaOH were obtained as analytical grade from Univar (Ajax Finechem, USA) and Sigma (St. Louis, USA), respectively. The process of squid pen chitosan (CHS) preparation and fabrication of the biocomposites are displayed in Fig. 1. Dried arrow squid pens (Nototodarus sloanii) were used for the preparation of -chitin (obtained from Independent Fisheries Co., Christchurch, New Zealand). Commercial crab chitosan (CHC) purchased from Weseta International (Shanghai, China) was used as received. 2.2. Preparation of chitosan from squid pen Isolation of chitosan from the squid pen was performed following a method developed by Chaussard and Domard [17] with minor modifications (Fig. 1). In brief, dried squid pens were grinded to particles ≤1 mm in diameter using Waring laboratory grinder (Waring Inc., USA), then deproteinization was carried out using 1 M sodium hydroxide (15 ml/g) at 60 ◦ C and constant stirring using a rotary shaker for 24 h. Then, the leachate was removed by vacuum filtration and particles were washed extensively until neutral pH was achieved. The obtained highly moist extracted material was frozen at −80 ◦ C and lyophilized using a freeze drier (Labconco FreeZone 12 Plus). Then the chitin powder was slowly added to a beaker containing 45% NaOH to obtain a solid/solvent ratio of 1:15 (w/v) and soaked at room temperature for 24 h [18]. The temperature of the reaction was maintained at 60 ◦ C and the mixture was stirred for 10 h. 2.3. Degree of N-deacetylation The degree of deacetylation (DDA) of chitosan (CHS) was calculated from data of elemental analysis (Carlo Erba Elemental Analyser EA 1108). DDA was calculated using Eq. (1) proposed by Xu et al. [19].
DDA (%) = 1 −
(C/N − 5.14) 1.72
∗ 100,
(1)
where C/N is the ratio (w/w) of carbon to nitrogen in chitosan. 2.4. Preparation of the composite Squid chitosan production and fabrication of the composites are shown in Fig. 1. Crab chitosan (CHC) solution (2%) was made by dissolving 5 g of chitosan in 250 ml of 1% acetic acid solution [20]. Due to the high hygroscopic nature of squid pen chitosan
Table 1 Composition of HA/-TCP/CH composites. Composite
A B
Compound (%) HA
-TCP
CHS
CHC
30 30
20 20
– 50
50 –
(CHS), the solution was very viscous at 1% and so the preparation of higher concentrations was technically difficult. To prepare 2% CHS solution, a 1% solution of chitosan dissolved in 1% acetic acid was subjected to microwave irradiation to remove excess water and achieving the desired concentration of 2%. The solution was then mixed by an overhead mixer (IKA T25 Ultra Turrax) for 2 min to obtain a transparent gel. The HA and -TCP powders were mixed together based on ratios shown in Table 1. The powders were weighed and made into a homogeneous paste using ethanol (1:10, w/v). The paste was added to the chitosan solutions, homogenized by the overhead mixer. The chitosan solutions were then sonicated (Elmasonic S40 (H)) for 1 h to remove any air bubbles. The air bubble free mixtures were transferred to 15 ml Poly-Cons® plastic container and frozen at −20 ◦ C, −80 ◦ C, or −196 ◦ C (the latter by direct immersion of the plastic container into liquid nitrogen for approximately 10 s). Then the samples were freeze-dried for 48 h (Labconco FreeZone 12 Plus) to form the HA/-TCP/CH (CHC or CHS) composites. The dried HA/-TCP/CH composites were soaked in 2.5% tripolyphosphate (TPP) aqueous solution at 4 ◦ C for 2 h [21]. Then the composites were rinsed with deionised water for 12 h at 4 ◦ C to remove residual TPP and were freeze dried for 24 h at −40 ◦ C. Composites made with crab chitosan and processed at −20 and −80 ◦ C were denoted as A220 and A280 respectively, and those made with squid pen chitosan were denoted as B220 and B280, respectively.
2.5. Characterization of the composites The distribution of HA/-TCP in the chitosan matrix was analysed using an X-Ray Diffractometer (XRD; PANnalytical X’Pert PRO MPD System) in the range 0◦ < 2 < 60◦ with Cu K␣ radiation (k = 0.15418 nm) with a scan speed of 2.63 s [22]. The functional groups of the samples were identified using Fourier Transform Infrared Spectroscopy (FT-IR; Perkin-Elmer #100) in the region 400–4000 cm−1 with 4 cm−1 spectral resolution using the KBr pellet technique [22]. Thermogravimetric analysis of the composites was carried out using a TGA instrument (TGA; Q 500) up to 1000 ◦ C at a 10 ◦ C/min heating rate under a nitrogen flow. Scanning electron microscopy (SEM) (JEOL 6700F FESEM JEOL Ltd, Tokyo, Japan) was used to examine the microscopic details of the composites.
2.6. Mechanical testing The mechanical properties of the HA/-TCP/CH composites were tested according to the guidelines of ASTM D5024-95a (22). The mechanical properties of composites were determined using a TA.XTPlus, Texture Analyzer (Texture Technologies Corp., Stable Micro Systems, Godalming, Surrey, UK). The analysis was carried out on cylindrical samples with dimensions of 25 mm diameter × 12 mm height. A 250 N load cell was operated at a rate of 0.5 mm min−1 until the sample was compressed to 50% of the original height at room temperature. The compression stress–strain curves were recorded, and compression modulus, yield and ultimate strength were calculated using the Exponent software (version V6.1.5.0) using 5 replicates per each of the composite samples [23].
A. Shavandi et al. / International Journal of Biological Macromolecules 80 (2015) 445–454
447
Fig. 1. Diagram for the fabrication of the HA/-TCP/CH composites.
2.7. Micro architectural analysis of composite X-ray microtomographic (-CT) systems have been widely used as a non-destructive technique to study or characterize microstructural morphology of various biomaterials [24,25]. -CT images of the HA/-TCP/CH composites were obtained using a Skyscan 1172 system (Bruker-Micro CT, Kontuch, Belgium) to quantify the 3-D microstructure of the composite samples. The CT-analyzer software v.1.14.4 (Bruker-Micro CT, Kontuch, Belgium) was used to calculate morphometric parameters and the calculations were performed using a segmented image from a rectangular region of
interest (ROI). The exhibited regions with white pixels are considered as solid portions and regions with black pixels, which are surrounded by white pixels, are pores and reported as percentage. 2.8. Water uptake and retention capacities The water uptake and retention ability of the composites was measured using the method described by Thein-Han et al. [26]. Sample with a known weight (Wd ) was immersed in distilled water for 24 h. Then, the sample was gently removed and placed on a wire rack for 1 min. The sample was weighed (Ww ) to determine
448
A. Shavandi et al. / International Journal of Biological Macromolecules 80 (2015) 445–454
the water uptake. To measure the retention ability of water, the wet composite sample was transferred to a falcon tube with a filter paper at the bottom of the tube was centrifuged using a Beckman GPR centrifuge at 500 RPM for 3 min, and measure its ultimate ). The weight of the sample was averaged from tripweighed (Ww licate measurements (n = 3) for each group of samples. The water uptake and retention percentage of the composites were calculated using the following Eq. (2) [23,26]: EA = [(Ww − Wd )/Wd ] × 100 − W )/W ] × 100 ER = [([(Ww d d
(2)
2.9. In vitro degradation of composites A simulated body fluid that considered a physiological buffer solution consisting of ␣-lactate and lactic acid (Lac-SBF) with an ionic concentration similar to human blood plasma was prepared as described by Cuneyt Tas [27]. The degradation ability of the composites was studied by incubating dry and weighed (W0 ) samples in Lac-SBF (pH 7.4 at 37 ◦ C) for up to 28 days. Approximately 25 mg samples (n = 3 for each group) were placed in glass liquid scintillation vials (volume 20 ml). A 10 ml of Lac-SBF solution were added to each of the vials and were then incubated at 37 ◦ C. The samples were removed at time intervals (1, 7, 14, and 28 days), washed five times with distilled water, and then dried at 60 ◦ C for overnight. The degradation was monitored in terms of changes in the weight of the samples over the time period. The degradation ratio (D) was calculated using the following Eq. (3): D = 100 −
W − W t 0 W0
× 100
CHC and CHS, i.e. the ␣-chitin and -chitin structure, respectively. Various DDA values have been reported for CHS in literature, up to 95%, which was dependent on the extraction conditions and methods used [11]. 3.2. Determination of the components by X-ray crystallography The phase of the composites was confirmed by X-ray diffraction (XRD). Samples processed at −196 ◦ C were all failed and collapsed during freeze drying, and so we omit any further characterization of these samples. Despite deacetylation, freezing temperature does not change crystal structure and chemical bonds of the chitosan; therefore, XRD and FTIR analysis were performed on two treatments (A280 and B280), instead of all four treatments. The XRD patterns of the composites made with CHS and CHC are displayed in Fig. 2A. The wide peak at 2 = 20◦ in all patterns was assigned to chitosan and corresponded to its degree of crystallinity [33]. The crystallinity was decreased with the increase in DDA [10]. As shown by the broad peaks of XRD graph at about 2 = 10◦ and 20◦ , CHC had less ordered structure, and lower crystallinity compared to CHS. The sharp peak at around 2 = 32◦ represents the HA/-TCP in the composite, also the peaks at 2 = 33, 34, 40, 47, 48◦ and 50–54◦ are also assigned to HA/-TCP. These results were in agreement with previous reports [34]. It has been reported that HA/-TCP weakens the intramolecular interaction of the chitosan chain [33]. The crystallinity of the HA/-TCP compound was found to be lower than either pure HA or -TCP [15,16] due to the presence of the chitosan matrix. However, the XRD patterns of natural bone also have broad and overlapping peaks [35].
(3)
where W0 represent the original weight and Wt is the weight of the sample at time t. Changes in the pH of Lac-SBF solution was also measured over the testing time using a pH meter (HI 2211, HANNA instruments, Rhode Island, USA) [28]. 2.10. Pore size determination The pore structure of composites was examined using scanning electron micrograph (SEM) and the average pore size was calculated using ImageJ [29]. The mean pore diameter was estimated by measuring about 100 different pores for each composite and three images were tested for each composite. 2.11. Statistical analysis Triplicate experiments were performed for each sample, and results were expressed as the mean of at least three replicates ± SEM. General linear Model (GLM) was performed between different composites, incubation times and/or temperatures using Minitab 16.2.4 Statistical Software and the differences were considered statistically significant at P < 0.05. The graphs were generated using the GraphPad Prism software (version 6.00 for Windows, GraphPad Software, San Diego, CA, USA, www.graphpad.com).
3.3. FTIR FTIR is a typical technique to investigate the structure-function of chitosan biomacromolecule and its interaction with HA/-TCP [36]. The infrared (IR) spectra of the composites made with CHS and CHC are shown in Fig. 2B. The peak at 1647 and 1560 cm−1 can be assigned to intracellular hydrogen bond between the carbonyl groups of amide I and II of chitosan respectively [26,30,37]. The peak at around 3260 cm−1 represents intramolecular hydrogen bonding between the stretching vibration of the N H bond of chitosan, and OH group of HA/-TCP. The sharp peak at 1415 cm−1 is devoted to the symmetrical deformation mode of CH3 . The peaks at 1029 and 1100 cm−1 are assigned to the C O stretching vibration. The typical band of PO4 observed at 500–700 cm−1 . The FTIR spectra of all composites indicated that the characteristic bands of both CaP compounds and chitosan were present in the composites. This FTIR result suggests that the structure of chitosan biomacromolecule provides a matrix for the HA/-TCP particles and also binds them together in the composites [26]. However, comparing the spectra bands of B280 and A280 in Fig. 2B, different intensities are observed in peaks at 1647 and 1560 cm−1 . The C O bands at 1660 cm−1 for A280 is less intense than B280, this is likely due to the deacetylation process that removed the C C bands. 3.4. Thermal analysis
3. Results and discussion 3.1. Degree of deacetylation The degree of deacetylation (DDA) represents the number of glucosamine units in a chitosan biopolymer chain [30]. DDA determines some of the properties of the polymer such as solubility, water uptake, biodegradation behaviour and its crystallinity [31]. The DDA of CHS and CHC was 72.8 ± 2.9 and 86.9 ± 1.7, respectively [32]. These different DDA values appear to be due to different efficiencies of deacetylation resulted from intramolecular structure of
Thermal degradation behaviour of the composites was studied using TGA. The TGA curves of the four composites are shown in Fig. 2(C). The weight loss for all composite samples began at 50 ◦ C, which could be due to water loss and sharpest decrease was observed at 50–100 ◦ C [38]. The second thermogram peak occurs at 250–350 ◦ C, which could be due to degradation of deacetylated molecules and the formation of saccharide molecule structure. This process includes dehydration of the saccharide ring, polymerization and decomposition of the acetylated units [39]. In case of composites A and B, which processed at different freezing
A. Shavandi et al. / International Journal of Biological Macromolecules 80 (2015) 445–454
449
Fig. 2. X-ray diffraction pattern (A), FTIR spectra (B) and thermograph analysis (C) of composite samples manufactured using two different chitosan (crab (A280) and squid (B280)) and processed at different freezing temperature (−20 and −80 ◦ C). A280 = 50% HA/-TCP and 50% crab chitosan (CHC). B280 = 50% HA/-TCP and 50% squid pen chitosan (CHS).
temperatures, composites of A (A220 and A280) showed a similar trend, while in case of B composites, B220 was slightly more thermally stable than B280. This higher stability might be attributed to its denser and robust structure as a result of its better cross linking
compare to B280. It is anticipated that, HA/-TCP provides a thermal barrier and hinders the degradation of composites [40]. This heat-resistant behaviour of chitosan can be assigned to bonds that formed between hydroxyl (OH) and amino (NH2 ) groups [38]. From
450
A. Shavandi et al. / International Journal of Biological Macromolecules 80 (2015) 445–454
Fig. 3. SEM images and photograph of the synthesized biocomposites. (A) Sample processed in liquid nitrogen, (B) Sample processed at −80 ◦ C. (C) A280 and B280 samples that processed at −20 ◦ C. A280 = 50% HA/-TCP and 50% crab chitosan (CHC). B280 = 50% HA/-TCP and 50% squid pen chitosan (CHS).
600 ◦ C to 1000 ◦ C the curve was stable and about 50% of the samples weight remained for all composites, which reflected the HA/-TCP %. 3.5. Microstructure morphology Scanning electron microscopy (SEM) was used to examine the microstructure of the composites (Fig. 3). The composites processed in liquid nitrogen (−196 ◦ C) illustrated radial and aligned channels. This microstructural formation is likely occurred due to water crystallization in chitosan droplets and subsequent ice sublimation (Fig. 3A). These needle-like channels clearly show the ice crystal shapes [41] since the composites had very compact and almost non porous structure (Fig. 3A), which could be due to its very fast freezing and solidification. The composites prepared at −196 ◦ C were incoherent and collapsed over freeze-drying processes, which possibly as a result of evaporation of water from the compact region and from channelled cavities. In a recent study, Ouyang et al. [41] reported that freezing chitosan beads in liquid nitrogen resulted in width channel size of 10–20 m. However, in that study chitosan drops were immersed in liquid nitrogen and therefore, no structural collapse was observed. Due to disjoint structure and low porosity of −196 ◦ C samples, we were unable to examine the structural morphology of the composites any further. The composite samples
processed at −80 ◦ C exhibited elongated pore structure, which is also showed extended irregular shape (Fig. 3B). However, the composites processed at −20 ◦ C appeared more irregular in shape, the pore structures were layered, and morphological geometry were more collapsed compared to samples processed at −80 ◦ C (Fig. 3C). This phenomenon can be attributed to a weaker structure of composites processed at −20 ◦ C compared to −80 ◦ C, which can be due to its slower freezing rate, bigger water crystals and consequently bigger pores and loser structure of −20 ◦ C samples compared to −80 ◦ C processed samples. Therefore, more intense cross-linking with TPP occurred that caused the breakdown of the internal structure. The presence of HA/-TCP in the composite was also observed on the surface of pore walls. 3.6. Pore size The average diameter of the pores was 122.2 m for the B220 (squid chitosan composite containing 50% HA/-TCP and frozen at −20 ◦ C), 456.2 m for B280 (squid chitosan composite containing 50% HA/-TCP and frozen at −80 ◦ C) composite, 93.1 m for A220 (crab chitosan composite containing 50% HA/TCP and frozen at −20 ◦ C) and 322.13 m for A280 (crab chitosan composite containing 50% HA/TCP and frozen at −80 ◦ C) composites. It was observed that the pore size and pore distribution of
A. Shavandi et al. / International Journal of Biological Macromolecules 80 (2015) 445–454
451
Fig. 4. (A) Pore size of composites fabricated using CHC (A) and CHS (B) that are processed at the freezing temperature −20 and −80 ◦ C, respectively. (B) Two dimensional (2D) cross-section image of composites processed at −20 ◦ C, where composite B220 (left) and composite A220 (right).
452
A. Shavandi et al. / International Journal of Biological Macromolecules 80 (2015) 445–454
Table 2 Micro-architectural parameters for the composites. Type of composite
Degree of anisotropy
A220 A280 B220 B280
1.34 1.31 1.36 1.09
± ± ± ±
0.25 0.23 0.26 0.08
Porosity (%) 45.43 79.10 62.79 66.40
± ± ± ±
2.54 1.62 4.36 5.70
Fig. 5. Porosity of composites fabricated using different chitosan (A) CHC and (B) CHS and freezing temperature (−20 and −80 ◦ C).
composites cross-linked with TPP and processed at −20 ◦ C appeared less homogeneous compared to composites processed at −80 ◦ C (Fig. 4A). It has been reported by Hsieh et al., that temperature of −20 ◦ C generated porous chitosan composites with larger pore-sized [42]. The two dimensional (2D) cross-section images of the A220 and B220 (Fig. 4B) revealed that B220 had a more uniform and regular internal structure compared to A220, suggesting CHS is a good candidate for fabrication of composites with homogeneous microstructure. The composite morphology is an important feature of the composite for considering it for bone-tissue engineering. The structure has to have enough porosity and proper pore size that allow blood vascularization and cell proliferation. Wide range of pore sizes, from 30 to 1000 m, has been reported in literature for adequate bone tissue engineering [43,44]. It has been reported that pore diameter greater than 300 m is essential for vascularization of composites and bone ingrowth [45–48]. The results from our study are in accordance with those previously reported pore sizes for chitosan/hydroxyapatite composites [20,49]. 3.7. Porosity In this study, the fabricated composites showed porosity values from 45 to 79% (Table 2 and Fig. 5). The porosity of the
composites is critical for the use in bone-tissue engineering applications. Porosity facilitates cell migration, blood circulation and vascularization during tissue repairing or regeneration. It has been reported that composite construction via lyophilization technique has demonstrated better pore size compared to sol–gel or precipitation method [50]. As shown in Table 2, composites frozen at −20 ◦ C had lower porosities compared to composites frozen at −80 ◦ C. For example, composite (A220), which was frozen and processed at −20 ◦ C, exhibited lower porosity compared to its counterpart composite (B220). In general, the crab chitosan composites processed at −20 ◦ C exhibited more shrinkage compared to squid chitosan composites frozen at the same temperature. This behaviour can be due to structure loss of composites frozen at −20 ◦ C compared to compact structure of composites at −80 ◦ C. The higher the freezing temperature is the slower the freezing rate of the chitosan solution is, therefore at a slower freezing rate, chitosan phase has more time to grow its grains size and ice crystals were bigger [42]. Therefore, the final dried structure was looser compared to −80 ◦ C. On the other hand, the lower porosity of crab chitosan composites A can be due to its lower initial viscosity compared to squid chitosan composites. Accordingly, TPP appears to be working better in composites processed at −20 ◦ C. At −80 ◦ C, composites A280 showed higher porosity compared to B280; this can be due to unstable and non-uniform structure of A280, which cannot hold all HA/-TCP in its structure and so due to loss of material it has higher porosity. 3.8. Mechanical properties of composites The mechanical properties of composites are shown in Fig. 6. The mechanical behaviour of the composites is a critical factor in their biomedical application and bone healing properties. It is documented that incorporation of CaP into the biomacromolecule matrix enhanced the mechanical properties of the composite [3]. Furthermore, cross-linking of composites with 2.5% TPP caused microstructure changes, which had significant effects on the mechanical properties of the composites. As shown in Fig. 6, composites frozen at −20 ◦ C showed better mechanical properties. In particular, composite B220, which was frozen at −20 ◦ C, recorded highest compression modulus, ultimate strength and yield strength. This can be due to formation of bigger ice crystals during freezing at low freezing temperature (i.e. −20 ◦ C compared to −80 ◦ C); so enabled better TPP cross-linking. Enhanced tensile properties were also reported by Hsieh et al., for composites frozen and processed at −20 ◦ C [42]. In case of the composites that were frozen and processed at −80 ◦ C, there were no significant differences between the composites (A280 and B280) and both showed lower mechanical properties compared to the composites frozen/processed at −20 ◦ C. These findings were in accordance
Fig. 6. Mechanical properties of the composites. (A) Compression modulus and (B) ultimate strength and yield strength of the composites.
A. Shavandi et al. / International Journal of Biological Macromolecules 80 (2015) 445–454
with previously literature by Reys et al., who reported CHS to have better mechanical properties than CHC composites [32]. In addition, when comparing different composites at same temperature of −20 ◦ C, composite A had collapsed internally and was not as strong as composite B (Fig. 4B). 3.9. Water uptake and water retention test The water retention ability of the composite is a critical factor for its efficacy for bone-tissue engineering; in particular, bone grafting. It has been reported that water uptake ability of the composite can significantly effect on cell proliferation and differentiation [51]. In this study, the results of the water uptake and retentions are shown in Fig. 7A and B. All the composites started swelling rapidly in the first day, indicating good water uptake characteristic [26]. From this water uptake data, it can be conferred that the composites
453
can take up and hold water more than its own weight (values are higher than 100%). However, maximum water uptake recorded in this study is almost 1200%, which is lower than the Chitosan/CaP composites (1600% and 2500%) reported in literature [26,52]. This lower water uptake recorded in this study might be due to the cross linking and shrinkage of the composites. As shown in Fig. 7A, composite A280 recorded highest water uptake, which could be due to its loose structural morphology that allows more water uptake and retention. On the other hand, lower water uptake and retention of composite B280 might be due to its degradation from 14 days onward. It is also noteworthy to mention that, the stiffer and rigid structure of composites processed at −20 ◦ C hindered their water uptake and retention ability. 3.10. In vitro degradation of composites Biodegradation properties of composites are an important factor on the long term functionality of the bone regeneration, including, grafting or repairing. The degradation profile of composite samples as a function of weight loss vs soaking time in SBF is presented in Fig. 7C. As shown the weight loss gradually increased by increasing time, which indicates composites degraded with soaking time. It is envisaged that the backbone of the chitosan bio-macromolecule is hydrolytically unstable [53], which enhance its degradation. After 28 days of in vitro degradation, the structural stability of all composite composites seems to be stable except for B280 samples, which degraded by about 40%. This high degradation might be due to two reasons: first the composite processed at −80 ◦ C (B280) did not cross-linked adequately compared to its counterpart B220. This could be due to the loss and weaker structure morphologies of B220 composite. Second, CHS chitosan that used for B composites had smaller DDA which promotes its degradation, as the structural stability of chitosan is inversely related to the DDA [54]. Considering these factors, composites processed at −20 ◦ C and cross-linked afterwards are durable and these composites maintained their structural stability over the tested period. 3.11. Change in pH of physiological solution Chemical stability of the composite is importance. Solubility of composite compartments can lead to change in pH and may affect the cell response [55]. The pH variation of SBF solution of composites monitored over a four-week period (Fig. 8). During the first two week of incubation in SBF, the pH value of all composites increased slightly and then decreased slightly for the following week (Fig. 8). This increase might be due to alkalescent nature of chitosan, which has alkaline groups, and also alkaline ions from degradation of CaP compounds [56,57]. The decrease of pH can be due to consumption of calcium and phosphate ions and their deposition on the composite surface [58]. The pH change pattern of the SBF solution might be
Fig. 7. Water uptake, water retention and degradation properties of HA/-TCP/CH composites. (A) Water uptake and (B) retention ability of the composites. Water uptake for composite B220 was significantly higher than A220 for all the tested times (P < 0.05) and Water uptake of composite A280 was significantly higher than B280 from day 14 onward. There was not significant differences for water retention between the composites. (C) In vitro degradation profile of the composites with various time period (1, 7, 14 and 28 days). Degradation of composite B280 was significantly higher than other composites.
Fig. 8. The pH values of the SBF solution in which samples were incubated.
454
A. Shavandi et al. / International Journal of Biological Macromolecules 80 (2015) 445–454
related to the dissolution characteristic of HA/-TCP and degradation profile of chitosan. Overall, the composites processed at −80 ◦ C showed higher chemical stability than −20 ◦ C composites and their pH of SBF solution only changed slightly and decreased to 7.20 from its original value of 7.45, and remained close to physiological value. 4. Conclusions The squid derived chitosan composites have shown to be more hydrophilic compared to commercial crab chitosan. Additionally, the SEM images and -CT results revealed that squid composites have homogeneous and uniform lamellar structure. The results of this study indicated that composites produced at −20 ◦ C had higher mechanical properties and lower degradation rate compared with −80 ◦ C and in overall, squid chitosan has better mechanical properties. These results indicated that chitosan isolated from the squid pen could be an alternative for existing commercial chitosan to be considered for biomedical applications. Acknowledgements The authors acknowledge the facilities as well as scientific and technical assistance from staff at Otago Centre for Electron Microscopy (OCEM) at the University of Otago. We would also like to thank Mr Damian Wallas for his help and technical support for XRD. The first author acknowledges the PhD scholarship by University of Otago, New Zealand. References [1] T.H. Silva, A. Alves, B.M. Ferreira, J.M. Oliveira, L.L. Reys, R.J.F. Ferreira, R.A. Sousa, S.S. Silva, J.F. Mano, R.L. Reis, Int. Mater. Rev. 57 (2012) 276–306. [2] J. Venkatesan, S.-K. Kim, Mar. Drugs 8 (2010) 2252–2266. [3] W.W. Thein-Han, R.D. Misra, Acta Biomater. 5 (2009) 1182–1197. [4] N.M. Alves, J.F. Mano, Int. J. Biol. Macromol. 43 (2008) 401–414. [5] M.L. Gisela, F.M. João, Biomed. Mater. 7 (2012) 054104. [6] M. Shimojoh, K. Fukushima, K. Kurita, Carbohydr. Polym. 35 (1998) 223–231. [7] I. Aranaz, M. Mengibar, R. Harris, I. Panos, B. Miralles, N. Acosta, G. Galed, A. Heras, Curr. Chem. Biol. 3 (2009) 203–230. [8] R. Minke, J. Blackwell, J. Mol. Biol. 120 (1978) 167–181. [9] N.E. Dweltz, Biochim. Biophys. Acta 51 (1961) 283–294. [10] K. Kurita, K. Tomita, T. Tada, S. Ishii, S.-I. Nishimura, K. Shimoda, J. Polym. Sci. A: Polym. Chem. 31 (1993) 485–491. [11] D.K. Youn, H.K. No, W. Prinyawiwatkul, Int. J. Food Sci. Technol. 48 (2013) 571–577. [12] F.A.A. Sagheer, M.A. Al-Sughayer, S. Muslim, M.Z. Elsabee, Carbohydr. Polym. 77 (2009) 410–419. [13] D. Raafat, H.-G. Sahl, Microb. Biotechnol. 2 (2009) 186–201. [14] E. Khor, in: E. Khor (Ed.), Chitin, 2nd ed., Elsevier, Oxford, 2014, pp. 51–66. [15] A. Shavandi, A.E.-D.A. Bekhit, A. Ali, Z. Sun, J.T. Ratnayake, Powder Technol. 273 (2015) 33–39. [16] A. Shavandi, A.E.-D.A. Bekhit, A. Ali, Z. Sun, Mater. Chem. Phys. 149–150 (2015) 607–616. [17] G. Chaussard, A. Domard, Biomacromolecules 5 (2004) 559–564. [18] K. Kurita, Prog. Polym. Sci. 26 (2001) 1921–1971.
[19] J. Xu, S.P. McCarthy, R.A. Gross, D.L. Kaplan, Macromolecules 29 (1996) 3436–3440. [20] Y. Zhang, M. Zhang, J. Biomed. Mater. Res. 61 (2002) 1–8. [21] S.H. Yu, S.J. Wu, J.Y. Wu, C.K. Peng, F.L. Mi, Molecules (Basel, Switz.) 18 (2012) 27–40. [22] G.S. Kumar, A. Thamizhavel, E.K. Girija, Mater. Lett. 76 (2012) 198–200. [23] M.V. Zugravu, R.A. Smith, B.T. Reves, J.A. Jennings, J.O. Cooper, W.O. Haggard, J.D. Bumgardner, J. Biomater. Appl. 28 (2013) 566–579. [24] A.C. Jones, B. Milthorpe, H. Averdunk, A. Limaye, T.J. Senden, A. Sakellariou, A.P. Sheppard, R.M. Sok, M.A. Knackstedt, A. Brandwood, D. Rohner, D.W. Hutmacher, Biomaterials 25 (2004) 4947–4954. [25] A.S. Lin, T.H. Barrows, S.H. Cartmell, R.E. Guldberg, Biomaterials 24 (2003) 481–489. [26] W.W. Thein-Han, R.D.K. Misra, Acta Biomater. 5 (2009) 1182–1197. [27] A. Cuneyt Tas, J. Non-Cryst. Solids 400 (2014) 27–32. ´ [28] J. Czechowska, A. Zima, Z. Paszkiewicz, J. Lis, A. Slósarczyk, Ceram. Int. 40 (2014) 5523–5532. [29] C.A. Schneider, W.S. Rasband, K.W. Eliceiri, Nat. Mater. 9 (2012) 671–675. [30] K.M. Rudall, W. Kenchington, Biol. Rev. 48 (1973) 597–633. [31] C. Chatelet, O. Damour, A. Domard, Biomaterials 22 (2001) 261–268. [32] L.L. Reys, S.S. Silva, J.M. Oliveira, S.G. Caridade, J.F. Mano, T.H. Silva, R.L. Reis, Biomed. Mater. 8 (2013) 1–11. [33] S.H. Teng, E.J. Lee, B.H. Yoon, D.S. Shin, H.E. Kim, J.S. Oh, J. Biomed. Mater. Res. A 88 (2009) 569–580. [34] Q. Hu, B. Li, M. Wang, J. Shen, Biomaterials 25 (2004) 779–785. [35] C. Meneghini, M.C. Dalconi, S. Nuzzo, S. Mobilio, R.H. Wenk, Biophys. J. 84 (2003) 2021–2029. [36] S.V. Dorozhkin, Biomaterials 1 (2011) 3–56. [37] R. Barabás, M. Czikó, I. Dékány, L. Bizo, E. Bogya, Chem. Pap. 67 (2013) 1414–1423. [38] M.C. Sunny, P. Ramesh, H.K. Varma, J. Mater. Sci.: Mater. Med. 13 (2002) 623–632. [39] A.T. Paulino, J.I. Simionato, J.C. Garcia, J. Nozaki, Carbohydr. Polym. 64 (2006) 98–103. [40] K.S. Katti, D.R. Katti, R. Dash, Biomed. Mater. 3 (2008) 034122. [41] A. Ouyang, Q. Gong, J. Liang, Adv. Eng. Mater. 17 (4) (2015) 460–466. [42] C.-Y. Hsieh, S.-P. Tsai, M.-H. Ho, D.-M. Wang, C.-E. Liu, C.-H. Hsieh, H.-C. Tseng, H.-J. Hsieh, Carbohydr. Polym. 67 (2007) 124–132. [43] K. Tuzlakoglu, C.M. Alves, J.F. Mano, R.L. Reis, Macromol. Biosci. 4 (2004) 811–819. [44] J.M. Oliveira, M.T. Rodrigues, S.S. Silva, P.B. Malafaya, M.E. Gomes, C.A. Viegas, I.R. Dias, J.T. Azevedo, J.F. Mano, R.L. Reis, Biomaterials 27 (2006) 6123–6137. [45] V. Karageorgiou, D. Kaplan, Biomaterials 26 (2005) 5474–5491. [46] Y. Kuboki, Q. Jin, H. Takita, J. Bone Joint Surg. 83 (2001) 105–115. [47] C.M. Murphy, M.G. Haugh, F.J. O’Brien, Biomaterials 31 (2010) 461–466. [48] H.E. Götz, M. Müller, A. Emmel, U. Holzwarth, R.G. Erben, R. Stangl, Biomaterials 25 (2004) 4057–4064. [49] F. Zhao, Y. Yin, W.W. Lu, J.C. Leong, W. Zhang, J. Zhang, M. Zhang, K. Yao, Biomaterials 23 (2002) 3227–3234. [50] J. Venkatesan, R. Pallela, I. Bhatnagar, S.K. Kim, Int. J. Biol. Macromol. 51 (2012) 1033–1042. [51] N. Shanmugasundaram, P. Ravichandran, P.N. Reddy, N. Ramamurty, S. Pal, K.P. Rao, Biomaterials 22 (2001) 1943–1951. [52] A. Sionkowska, J. Kozlowska, Int. J. Biol. Macromol. 52 (2013) 250–259. [53] B.D. Ulery, L.S. Nair, C.T. Laurencin, J. Polym. Sci. B: Polym. Phys. 49 (2011) 832–864. [54] A.M. Martins, R.C. Pereira, I.B. Leonor, H.S. Azevedo, R.L. Reis, Acta Biomater. 5 (2009) 3328–3336. [55] J. Gustavsson, M.P. Ginebra, E. Engel, J. Planell, Acta Biomater. 7 (2011) 4242–4252. [56] X. Wang, G. Song, T. Lou, W. Peng, J. Biomater. Sci. 20 (2009) 1995–2002. [57] X.-m. Pu, Q.-q. Yao, Y. Yang, Z.-z. Sun, Q.-q. Zhang, Int. J. Biol. Macromol. 51 (2012) 868–873. [58] A. Yoshida, T. Miyazaki, E. Ishida, M. Ashizuka, Mater. Trans. 45 (2004) 994–998.
Contents lists available at ScienceDirect
International Journal of Biological Macromolecules journal homepage: www.elsevier.com/locate/ijbiomac
Bio-mimetic composite scaffold from mussel shells, squid pen and crab chitosan for bone tissue engineering Amin Shavandi a,∗ , Alaa El-Din A. Bekhit a , M. Azam Ali b , Zhifa Sun c a b c
Department of Food Sciences, University of Otago, Dunedin, New Zealand Department of Applied Sciences, University of Otago, Dunedin, New Zealand Department of Physics, University of Otago, Dunedin, New Zealand
a r t i c l e
i n f o
Article history: Received 5 March 2015 Received in revised form 2 July 2015 Accepted 8 July 2015 Available online 15 July 2015 Keywords: Squid pen Chitosan Hydroxyapatite
a b s t r a c t In the present study, chitosan/hydroxyapatite (HA)/-tircalcium phosphate (-TCP) composites were produced using squid pen derived chitosan (CHS) and commercial crab derived chitosan (CHC). CHS was prepared from squid pens by alkaline N-deacetylation. HA and -TCP were extracted from mussel shells using a microwave irradiation method. Two different composites were prepared by incorporating 50% (w/w) HA/(-TCP) in CHS or CHC followed by lyophilization and cross-linking of composites by tripolyphosphate (TPP). The effect of different freezing temperatures of −20, −80 and −196 ◦ C on the physicochemical characteristics of composites was investigated. A simulated body fluid (SBF) solution was used for preliminary in vitro study for 1, 7, 14 and 28 days and the composites were characterized by XRD, FTIR, TGA, SEM, -CT and ICP-MS. Porosity, pore size, water uptake; water retention abilities and in vitro degradations of the prepared composites were evaluated. The CHS composites were found to have higher porosity (62%) compared to the CHC composites (porosity 42%) and better mechanical properties. The results of this study indicated that composites produced at −20 ◦ C had higher mechanical properties and lower degradation rate compared with −80 ◦ C. Chitosan from the squid pen is an excellent biomaterial candidate for bone tissue engineering applications. © 2015 Elsevier B.V. All rights reserved.
1. Introduction In recent years, much attention has been paid to marine byproducts, scoping their cost-effective processing schemes and their potential for production of high-value products. Natural polymers like chitin, chitosan and calcium phosphate (CaP) compounds can be obtained from waste marine products [1]. Because of their intrinsic properties such as biocompatibility, biodegradation and antimicrobial properties, these natural materials have important biomedical applications [2,3]. In the last two decades, many reports have been published on chitin and chitosan applications in drug delivery, tissue engineering, skin and bone grafting [2,4,5]. Chitosan is a biopolymer consisting of (1,4)-2-amino-2-deoxy-d-glucose units that is obtained by N-deacetylation of chitin under alkaline condition. Chitin can be sourced and extracted from a diverse range of natural organisms, including molluscs, fungi, insects, crustaceans and algae [6]. Chitin exists in three different allomorphic forms depending on the sources of the compound. Most chitins, including
∗ Corresponding author. E-mail address: [email protected] (A. Shavandi). http://dx.doi.org/10.1016/j.ijbiomac.2015.07.012 0141-8130/© 2015 Elsevier B.V. All rights reserved.
crustaceans and insect chitin are in alpha (␣) form which has a two chain antiparallel structure. However, squid pen and some diatoms have beta () chitin which has one chain parallel structure and in gamma (␥) chitin, the biomolecular chains are arranged randomly in which two parallel chains and one antiparallel chain form the polymeric structure [7]. Alpha-chitin and chitosan are commercially available products and are produced normally from shrimp or crab shell. Chitin/chitosan from the squid pen has a structure that is low packed and has weak intermolecular hydrogen bonds. These properties makes it chemically more reactive compared to the heavily packed and strong molecular structure of ␣-chitin/chitosan from shrimp and crab shells [8–10]. In addition, -chitin/chitosan can incorporate water molecules in its structure and forms crystalline structure leading to higher ability to uptake and hold water more than the alpha form, which is advantageous in biomedical applications [11,12]. The source of chitosan can affect its purity, molecular weight, chain length, degree of deacetylation, density, viscosity, solubility, water retention capacity and distribution of the amino/acetamide groups. All these characteristics affect the physicochemical properties of chitosan and therefore, its application [13]. A proper biocomposite should be prepared strategically in a way to have a suitable geometry and pore size, have a
446
A. Shavandi et al. / International Journal of Biological Macromolecules 80 (2015) 445–454
required mechanical strength, support cell attachment in vitro, and have tuneable biodegradable properties [14]. In this work, chitosan was isolated from endoskeleton of New Zealand arrow squid pen (Nototodarus sloanii) (CHS). In addition, a commercially available crab chitosan (CHC) was used for comparison with CHS. The HA and -TCP were prepared using waste green mussel shells. Then CHS, CHC and HA, -TCP were processed to produce biocomposites using freezing at different temperatures (−20, −80 and −196 ◦ C) and lyophilization processes. The produced biocomposites were evaluated to scope their potential for bone tissue engineering applications. To improve the mechanical properties of the composites in this study, tripolyphosphate (TPP) was used as an ionic cross-linker and glycerol was used as plasticizing agent. 2. Materials and methods 2.1. Materials Hydroxyapatite (Ca10 (PO4 )6 (OH)2 ) and -TCP (Ca3 (PO4 )2 ) powders were synthesized from waste mussel shells as previously reported [15,16]. Acetic acid, ethanol, and NaOH were obtained as analytical grade from Univar (Ajax Finechem, USA) and Sigma (St. Louis, USA), respectively. The process of squid pen chitosan (CHS) preparation and fabrication of the biocomposites are displayed in Fig. 1. Dried arrow squid pens (Nototodarus sloanii) were used for the preparation of -chitin (obtained from Independent Fisheries Co., Christchurch, New Zealand). Commercial crab chitosan (CHC) purchased from Weseta International (Shanghai, China) was used as received. 2.2. Preparation of chitosan from squid pen Isolation of chitosan from the squid pen was performed following a method developed by Chaussard and Domard [17] with minor modifications (Fig. 1). In brief, dried squid pens were grinded to particles ≤1 mm in diameter using Waring laboratory grinder (Waring Inc., USA), then deproteinization was carried out using 1 M sodium hydroxide (15 ml/g) at 60 ◦ C and constant stirring using a rotary shaker for 24 h. Then, the leachate was removed by vacuum filtration and particles were washed extensively until neutral pH was achieved. The obtained highly moist extracted material was frozen at −80 ◦ C and lyophilized using a freeze drier (Labconco FreeZone 12 Plus). Then the chitin powder was slowly added to a beaker containing 45% NaOH to obtain a solid/solvent ratio of 1:15 (w/v) and soaked at room temperature for 24 h [18]. The temperature of the reaction was maintained at 60 ◦ C and the mixture was stirred for 10 h. 2.3. Degree of N-deacetylation The degree of deacetylation (DDA) of chitosan (CHS) was calculated from data of elemental analysis (Carlo Erba Elemental Analyser EA 1108). DDA was calculated using Eq. (1) proposed by Xu et al. [19].
DDA (%) = 1 −
(C/N − 5.14) 1.72
∗ 100,
(1)
where C/N is the ratio (w/w) of carbon to nitrogen in chitosan. 2.4. Preparation of the composite Squid chitosan production and fabrication of the composites are shown in Fig. 1. Crab chitosan (CHC) solution (2%) was made by dissolving 5 g of chitosan in 250 ml of 1% acetic acid solution [20]. Due to the high hygroscopic nature of squid pen chitosan
Table 1 Composition of HA/-TCP/CH composites. Composite
A B
Compound (%) HA
-TCP
CHS
CHC
30 30
20 20
– 50
50 –
(CHS), the solution was very viscous at 1% and so the preparation of higher concentrations was technically difficult. To prepare 2% CHS solution, a 1% solution of chitosan dissolved in 1% acetic acid was subjected to microwave irradiation to remove excess water and achieving the desired concentration of 2%. The solution was then mixed by an overhead mixer (IKA T25 Ultra Turrax) for 2 min to obtain a transparent gel. The HA and -TCP powders were mixed together based on ratios shown in Table 1. The powders were weighed and made into a homogeneous paste using ethanol (1:10, w/v). The paste was added to the chitosan solutions, homogenized by the overhead mixer. The chitosan solutions were then sonicated (Elmasonic S40 (H)) for 1 h to remove any air bubbles. The air bubble free mixtures were transferred to 15 ml Poly-Cons® plastic container and frozen at −20 ◦ C, −80 ◦ C, or −196 ◦ C (the latter by direct immersion of the plastic container into liquid nitrogen for approximately 10 s). Then the samples were freeze-dried for 48 h (Labconco FreeZone 12 Plus) to form the HA/-TCP/CH (CHC or CHS) composites. The dried HA/-TCP/CH composites were soaked in 2.5% tripolyphosphate (TPP) aqueous solution at 4 ◦ C for 2 h [21]. Then the composites were rinsed with deionised water for 12 h at 4 ◦ C to remove residual TPP and were freeze dried for 24 h at −40 ◦ C. Composites made with crab chitosan and processed at −20 and −80 ◦ C were denoted as A220 and A280 respectively, and those made with squid pen chitosan were denoted as B220 and B280, respectively.
2.5. Characterization of the composites The distribution of HA/-TCP in the chitosan matrix was analysed using an X-Ray Diffractometer (XRD; PANnalytical X’Pert PRO MPD System) in the range 0◦ < 2 < 60◦ with Cu K␣ radiation (k = 0.15418 nm) with a scan speed of 2.63 s [22]. The functional groups of the samples were identified using Fourier Transform Infrared Spectroscopy (FT-IR; Perkin-Elmer #100) in the region 400–4000 cm−1 with 4 cm−1 spectral resolution using the KBr pellet technique [22]. Thermogravimetric analysis of the composites was carried out using a TGA instrument (TGA; Q 500) up to 1000 ◦ C at a 10 ◦ C/min heating rate under a nitrogen flow. Scanning electron microscopy (SEM) (JEOL 6700F FESEM JEOL Ltd, Tokyo, Japan) was used to examine the microscopic details of the composites.
2.6. Mechanical testing The mechanical properties of the HA/-TCP/CH composites were tested according to the guidelines of ASTM D5024-95a (22). The mechanical properties of composites were determined using a TA.XTPlus, Texture Analyzer (Texture Technologies Corp., Stable Micro Systems, Godalming, Surrey, UK). The analysis was carried out on cylindrical samples with dimensions of 25 mm diameter × 12 mm height. A 250 N load cell was operated at a rate of 0.5 mm min−1 until the sample was compressed to 50% of the original height at room temperature. The compression stress–strain curves were recorded, and compression modulus, yield and ultimate strength were calculated using the Exponent software (version V6.1.5.0) using 5 replicates per each of the composite samples [23].
A. Shavandi et al. / International Journal of Biological Macromolecules 80 (2015) 445–454
447
Fig. 1. Diagram for the fabrication of the HA/-TCP/CH composites.
2.7. Micro architectural analysis of composite X-ray microtomographic (-CT) systems have been widely used as a non-destructive technique to study or characterize microstructural morphology of various biomaterials [24,25]. -CT images of the HA/-TCP/CH composites were obtained using a Skyscan 1172 system (Bruker-Micro CT, Kontuch, Belgium) to quantify the 3-D microstructure of the composite samples. The CT-analyzer software v.1.14.4 (Bruker-Micro CT, Kontuch, Belgium) was used to calculate morphometric parameters and the calculations were performed using a segmented image from a rectangular region of
interest (ROI). The exhibited regions with white pixels are considered as solid portions and regions with black pixels, which are surrounded by white pixels, are pores and reported as percentage. 2.8. Water uptake and retention capacities The water uptake and retention ability of the composites was measured using the method described by Thein-Han et al. [26]. Sample with a known weight (Wd ) was immersed in distilled water for 24 h. Then, the sample was gently removed and placed on a wire rack for 1 min. The sample was weighed (Ww ) to determine
448
A. Shavandi et al. / International Journal of Biological Macromolecules 80 (2015) 445–454
the water uptake. To measure the retention ability of water, the wet composite sample was transferred to a falcon tube with a filter paper at the bottom of the tube was centrifuged using a Beckman GPR centrifuge at 500 RPM for 3 min, and measure its ultimate ). The weight of the sample was averaged from tripweighed (Ww licate measurements (n = 3) for each group of samples. The water uptake and retention percentage of the composites were calculated using the following Eq. (2) [23,26]: EA = [(Ww − Wd )/Wd ] × 100 − W )/W ] × 100 ER = [([(Ww d d
(2)
2.9. In vitro degradation of composites A simulated body fluid that considered a physiological buffer solution consisting of ␣-lactate and lactic acid (Lac-SBF) with an ionic concentration similar to human blood plasma was prepared as described by Cuneyt Tas [27]. The degradation ability of the composites was studied by incubating dry and weighed (W0 ) samples in Lac-SBF (pH 7.4 at 37 ◦ C) for up to 28 days. Approximately 25 mg samples (n = 3 for each group) were placed in glass liquid scintillation vials (volume 20 ml). A 10 ml of Lac-SBF solution were added to each of the vials and were then incubated at 37 ◦ C. The samples were removed at time intervals (1, 7, 14, and 28 days), washed five times with distilled water, and then dried at 60 ◦ C for overnight. The degradation was monitored in terms of changes in the weight of the samples over the time period. The degradation ratio (D) was calculated using the following Eq. (3): D = 100 −
W − W t 0 W0
× 100
CHC and CHS, i.e. the ␣-chitin and -chitin structure, respectively. Various DDA values have been reported for CHS in literature, up to 95%, which was dependent on the extraction conditions and methods used [11]. 3.2. Determination of the components by X-ray crystallography The phase of the composites was confirmed by X-ray diffraction (XRD). Samples processed at −196 ◦ C were all failed and collapsed during freeze drying, and so we omit any further characterization of these samples. Despite deacetylation, freezing temperature does not change crystal structure and chemical bonds of the chitosan; therefore, XRD and FTIR analysis were performed on two treatments (A280 and B280), instead of all four treatments. The XRD patterns of the composites made with CHS and CHC are displayed in Fig. 2A. The wide peak at 2 = 20◦ in all patterns was assigned to chitosan and corresponded to its degree of crystallinity [33]. The crystallinity was decreased with the increase in DDA [10]. As shown by the broad peaks of XRD graph at about 2 = 10◦ and 20◦ , CHC had less ordered structure, and lower crystallinity compared to CHS. The sharp peak at around 2 = 32◦ represents the HA/-TCP in the composite, also the peaks at 2 = 33, 34, 40, 47, 48◦ and 50–54◦ are also assigned to HA/-TCP. These results were in agreement with previous reports [34]. It has been reported that HA/-TCP weakens the intramolecular interaction of the chitosan chain [33]. The crystallinity of the HA/-TCP compound was found to be lower than either pure HA or -TCP [15,16] due to the presence of the chitosan matrix. However, the XRD patterns of natural bone also have broad and overlapping peaks [35].
(3)
where W0 represent the original weight and Wt is the weight of the sample at time t. Changes in the pH of Lac-SBF solution was also measured over the testing time using a pH meter (HI 2211, HANNA instruments, Rhode Island, USA) [28]. 2.10. Pore size determination The pore structure of composites was examined using scanning electron micrograph (SEM) and the average pore size was calculated using ImageJ [29]. The mean pore diameter was estimated by measuring about 100 different pores for each composite and three images were tested for each composite. 2.11. Statistical analysis Triplicate experiments were performed for each sample, and results were expressed as the mean of at least three replicates ± SEM. General linear Model (GLM) was performed between different composites, incubation times and/or temperatures using Minitab 16.2.4 Statistical Software and the differences were considered statistically significant at P < 0.05. The graphs were generated using the GraphPad Prism software (version 6.00 for Windows, GraphPad Software, San Diego, CA, USA, www.graphpad.com).
3.3. FTIR FTIR is a typical technique to investigate the structure-function of chitosan biomacromolecule and its interaction with HA/-TCP [36]. The infrared (IR) spectra of the composites made with CHS and CHC are shown in Fig. 2B. The peak at 1647 and 1560 cm−1 can be assigned to intracellular hydrogen bond between the carbonyl groups of amide I and II of chitosan respectively [26,30,37]. The peak at around 3260 cm−1 represents intramolecular hydrogen bonding between the stretching vibration of the N H bond of chitosan, and OH group of HA/-TCP. The sharp peak at 1415 cm−1 is devoted to the symmetrical deformation mode of CH3 . The peaks at 1029 and 1100 cm−1 are assigned to the C O stretching vibration. The typical band of PO4 observed at 500–700 cm−1 . The FTIR spectra of all composites indicated that the characteristic bands of both CaP compounds and chitosan were present in the composites. This FTIR result suggests that the structure of chitosan biomacromolecule provides a matrix for the HA/-TCP particles and also binds them together in the composites [26]. However, comparing the spectra bands of B280 and A280 in Fig. 2B, different intensities are observed in peaks at 1647 and 1560 cm−1 . The C O bands at 1660 cm−1 for A280 is less intense than B280, this is likely due to the deacetylation process that removed the C C bands. 3.4. Thermal analysis
3. Results and discussion 3.1. Degree of deacetylation The degree of deacetylation (DDA) represents the number of glucosamine units in a chitosan biopolymer chain [30]. DDA determines some of the properties of the polymer such as solubility, water uptake, biodegradation behaviour and its crystallinity [31]. The DDA of CHS and CHC was 72.8 ± 2.9 and 86.9 ± 1.7, respectively [32]. These different DDA values appear to be due to different efficiencies of deacetylation resulted from intramolecular structure of
Thermal degradation behaviour of the composites was studied using TGA. The TGA curves of the four composites are shown in Fig. 2(C). The weight loss for all composite samples began at 50 ◦ C, which could be due to water loss and sharpest decrease was observed at 50–100 ◦ C [38]. The second thermogram peak occurs at 250–350 ◦ C, which could be due to degradation of deacetylated molecules and the formation of saccharide molecule structure. This process includes dehydration of the saccharide ring, polymerization and decomposition of the acetylated units [39]. In case of composites A and B, which processed at different freezing
A. Shavandi et al. / International Journal of Biological Macromolecules 80 (2015) 445–454
449
Fig. 2. X-ray diffraction pattern (A), FTIR spectra (B) and thermograph analysis (C) of composite samples manufactured using two different chitosan (crab (A280) and squid (B280)) and processed at different freezing temperature (−20 and −80 ◦ C). A280 = 50% HA/-TCP and 50% crab chitosan (CHC). B280 = 50% HA/-TCP and 50% squid pen chitosan (CHS).
temperatures, composites of A (A220 and A280) showed a similar trend, while in case of B composites, B220 was slightly more thermally stable than B280. This higher stability might be attributed to its denser and robust structure as a result of its better cross linking
compare to B280. It is anticipated that, HA/-TCP provides a thermal barrier and hinders the degradation of composites [40]. This heat-resistant behaviour of chitosan can be assigned to bonds that formed between hydroxyl (OH) and amino (NH2 ) groups [38]. From
450
A. Shavandi et al. / International Journal of Biological Macromolecules 80 (2015) 445–454
Fig. 3. SEM images and photograph of the synthesized biocomposites. (A) Sample processed in liquid nitrogen, (B) Sample processed at −80 ◦ C. (C) A280 and B280 samples that processed at −20 ◦ C. A280 = 50% HA/-TCP and 50% crab chitosan (CHC). B280 = 50% HA/-TCP and 50% squid pen chitosan (CHS).
600 ◦ C to 1000 ◦ C the curve was stable and about 50% of the samples weight remained for all composites, which reflected the HA/-TCP %. 3.5. Microstructure morphology Scanning electron microscopy (SEM) was used to examine the microstructure of the composites (Fig. 3). The composites processed in liquid nitrogen (−196 ◦ C) illustrated radial and aligned channels. This microstructural formation is likely occurred due to water crystallization in chitosan droplets and subsequent ice sublimation (Fig. 3A). These needle-like channels clearly show the ice crystal shapes [41] since the composites had very compact and almost non porous structure (Fig. 3A), which could be due to its very fast freezing and solidification. The composites prepared at −196 ◦ C were incoherent and collapsed over freeze-drying processes, which possibly as a result of evaporation of water from the compact region and from channelled cavities. In a recent study, Ouyang et al. [41] reported that freezing chitosan beads in liquid nitrogen resulted in width channel size of 10–20 m. However, in that study chitosan drops were immersed in liquid nitrogen and therefore, no structural collapse was observed. Due to disjoint structure and low porosity of −196 ◦ C samples, we were unable to examine the structural morphology of the composites any further. The composite samples
processed at −80 ◦ C exhibited elongated pore structure, which is also showed extended irregular shape (Fig. 3B). However, the composites processed at −20 ◦ C appeared more irregular in shape, the pore structures were layered, and morphological geometry were more collapsed compared to samples processed at −80 ◦ C (Fig. 3C). This phenomenon can be attributed to a weaker structure of composites processed at −20 ◦ C compared to −80 ◦ C, which can be due to its slower freezing rate, bigger water crystals and consequently bigger pores and loser structure of −20 ◦ C samples compared to −80 ◦ C processed samples. Therefore, more intense cross-linking with TPP occurred that caused the breakdown of the internal structure. The presence of HA/-TCP in the composite was also observed on the surface of pore walls. 3.6. Pore size The average diameter of the pores was 122.2 m for the B220 (squid chitosan composite containing 50% HA/-TCP and frozen at −20 ◦ C), 456.2 m for B280 (squid chitosan composite containing 50% HA/-TCP and frozen at −80 ◦ C) composite, 93.1 m for A220 (crab chitosan composite containing 50% HA/TCP and frozen at −20 ◦ C) and 322.13 m for A280 (crab chitosan composite containing 50% HA/TCP and frozen at −80 ◦ C) composites. It was observed that the pore size and pore distribution of
A. Shavandi et al. / International Journal of Biological Macromolecules 80 (2015) 445–454
451
Fig. 4. (A) Pore size of composites fabricated using CHC (A) and CHS (B) that are processed at the freezing temperature −20 and −80 ◦ C, respectively. (B) Two dimensional (2D) cross-section image of composites processed at −20 ◦ C, where composite B220 (left) and composite A220 (right).
452
A. Shavandi et al. / International Journal of Biological Macromolecules 80 (2015) 445–454
Table 2 Micro-architectural parameters for the composites. Type of composite
Degree of anisotropy
A220 A280 B220 B280
1.34 1.31 1.36 1.09
± ± ± ±
0.25 0.23 0.26 0.08
Porosity (%) 45.43 79.10 62.79 66.40
± ± ± ±
2.54 1.62 4.36 5.70
Fig. 5. Porosity of composites fabricated using different chitosan (A) CHC and (B) CHS and freezing temperature (−20 and −80 ◦ C).
composites cross-linked with TPP and processed at −20 ◦ C appeared less homogeneous compared to composites processed at −80 ◦ C (Fig. 4A). It has been reported by Hsieh et al., that temperature of −20 ◦ C generated porous chitosan composites with larger pore-sized [42]. The two dimensional (2D) cross-section images of the A220 and B220 (Fig. 4B) revealed that B220 had a more uniform and regular internal structure compared to A220, suggesting CHS is a good candidate for fabrication of composites with homogeneous microstructure. The composite morphology is an important feature of the composite for considering it for bone-tissue engineering. The structure has to have enough porosity and proper pore size that allow blood vascularization and cell proliferation. Wide range of pore sizes, from 30 to 1000 m, has been reported in literature for adequate bone tissue engineering [43,44]. It has been reported that pore diameter greater than 300 m is essential for vascularization of composites and bone ingrowth [45–48]. The results from our study are in accordance with those previously reported pore sizes for chitosan/hydroxyapatite composites [20,49]. 3.7. Porosity In this study, the fabricated composites showed porosity values from 45 to 79% (Table 2 and Fig. 5). The porosity of the
composites is critical for the use in bone-tissue engineering applications. Porosity facilitates cell migration, blood circulation and vascularization during tissue repairing or regeneration. It has been reported that composite construction via lyophilization technique has demonstrated better pore size compared to sol–gel or precipitation method [50]. As shown in Table 2, composites frozen at −20 ◦ C had lower porosities compared to composites frozen at −80 ◦ C. For example, composite (A220), which was frozen and processed at −20 ◦ C, exhibited lower porosity compared to its counterpart composite (B220). In general, the crab chitosan composites processed at −20 ◦ C exhibited more shrinkage compared to squid chitosan composites frozen at the same temperature. This behaviour can be due to structure loss of composites frozen at −20 ◦ C compared to compact structure of composites at −80 ◦ C. The higher the freezing temperature is the slower the freezing rate of the chitosan solution is, therefore at a slower freezing rate, chitosan phase has more time to grow its grains size and ice crystals were bigger [42]. Therefore, the final dried structure was looser compared to −80 ◦ C. On the other hand, the lower porosity of crab chitosan composites A can be due to its lower initial viscosity compared to squid chitosan composites. Accordingly, TPP appears to be working better in composites processed at −20 ◦ C. At −80 ◦ C, composites A280 showed higher porosity compared to B280; this can be due to unstable and non-uniform structure of A280, which cannot hold all HA/-TCP in its structure and so due to loss of material it has higher porosity. 3.8. Mechanical properties of composites The mechanical properties of composites are shown in Fig. 6. The mechanical behaviour of the composites is a critical factor in their biomedical application and bone healing properties. It is documented that incorporation of CaP into the biomacromolecule matrix enhanced the mechanical properties of the composite [3]. Furthermore, cross-linking of composites with 2.5% TPP caused microstructure changes, which had significant effects on the mechanical properties of the composites. As shown in Fig. 6, composites frozen at −20 ◦ C showed better mechanical properties. In particular, composite B220, which was frozen at −20 ◦ C, recorded highest compression modulus, ultimate strength and yield strength. This can be due to formation of bigger ice crystals during freezing at low freezing temperature (i.e. −20 ◦ C compared to −80 ◦ C); so enabled better TPP cross-linking. Enhanced tensile properties were also reported by Hsieh et al., for composites frozen and processed at −20 ◦ C [42]. In case of the composites that were frozen and processed at −80 ◦ C, there were no significant differences between the composites (A280 and B280) and both showed lower mechanical properties compared to the composites frozen/processed at −20 ◦ C. These findings were in accordance
Fig. 6. Mechanical properties of the composites. (A) Compression modulus and (B) ultimate strength and yield strength of the composites.
A. Shavandi et al. / International Journal of Biological Macromolecules 80 (2015) 445–454
with previously literature by Reys et al., who reported CHS to have better mechanical properties than CHC composites [32]. In addition, when comparing different composites at same temperature of −20 ◦ C, composite A had collapsed internally and was not as strong as composite B (Fig. 4B). 3.9. Water uptake and water retention test The water retention ability of the composite is a critical factor for its efficacy for bone-tissue engineering; in particular, bone grafting. It has been reported that water uptake ability of the composite can significantly effect on cell proliferation and differentiation [51]. In this study, the results of the water uptake and retentions are shown in Fig. 7A and B. All the composites started swelling rapidly in the first day, indicating good water uptake characteristic [26]. From this water uptake data, it can be conferred that the composites
453
can take up and hold water more than its own weight (values are higher than 100%). However, maximum water uptake recorded in this study is almost 1200%, which is lower than the Chitosan/CaP composites (1600% and 2500%) reported in literature [26,52]. This lower water uptake recorded in this study might be due to the cross linking and shrinkage of the composites. As shown in Fig. 7A, composite A280 recorded highest water uptake, which could be due to its loose structural morphology that allows more water uptake and retention. On the other hand, lower water uptake and retention of composite B280 might be due to its degradation from 14 days onward. It is also noteworthy to mention that, the stiffer and rigid structure of composites processed at −20 ◦ C hindered their water uptake and retention ability. 3.10. In vitro degradation of composites Biodegradation properties of composites are an important factor on the long term functionality of the bone regeneration, including, grafting or repairing. The degradation profile of composite samples as a function of weight loss vs soaking time in SBF is presented in Fig. 7C. As shown the weight loss gradually increased by increasing time, which indicates composites degraded with soaking time. It is envisaged that the backbone of the chitosan bio-macromolecule is hydrolytically unstable [53], which enhance its degradation. After 28 days of in vitro degradation, the structural stability of all composite composites seems to be stable except for B280 samples, which degraded by about 40%. This high degradation might be due to two reasons: first the composite processed at −80 ◦ C (B280) did not cross-linked adequately compared to its counterpart B220. This could be due to the loss and weaker structure morphologies of B220 composite. Second, CHS chitosan that used for B composites had smaller DDA which promotes its degradation, as the structural stability of chitosan is inversely related to the DDA [54]. Considering these factors, composites processed at −20 ◦ C and cross-linked afterwards are durable and these composites maintained their structural stability over the tested period. 3.11. Change in pH of physiological solution Chemical stability of the composite is importance. Solubility of composite compartments can lead to change in pH and may affect the cell response [55]. The pH variation of SBF solution of composites monitored over a four-week period (Fig. 8). During the first two week of incubation in SBF, the pH value of all composites increased slightly and then decreased slightly for the following week (Fig. 8). This increase might be due to alkalescent nature of chitosan, which has alkaline groups, and also alkaline ions from degradation of CaP compounds [56,57]. The decrease of pH can be due to consumption of calcium and phosphate ions and their deposition on the composite surface [58]. The pH change pattern of the SBF solution might be
Fig. 7. Water uptake, water retention and degradation properties of HA/-TCP/CH composites. (A) Water uptake and (B) retention ability of the composites. Water uptake for composite B220 was significantly higher than A220 for all the tested times (P < 0.05) and Water uptake of composite A280 was significantly higher than B280 from day 14 onward. There was not significant differences for water retention between the composites. (C) In vitro degradation profile of the composites with various time period (1, 7, 14 and 28 days). Degradation of composite B280 was significantly higher than other composites.
Fig. 8. The pH values of the SBF solution in which samples were incubated.
454
A. Shavandi et al. / International Journal of Biological Macromolecules 80 (2015) 445–454
related to the dissolution characteristic of HA/-TCP and degradation profile of chitosan. Overall, the composites processed at −80 ◦ C showed higher chemical stability than −20 ◦ C composites and their pH of SBF solution only changed slightly and decreased to 7.20 from its original value of 7.45, and remained close to physiological value. 4. Conclusions The squid derived chitosan composites have shown to be more hydrophilic compared to commercial crab chitosan. Additionally, the SEM images and -CT results revealed that squid composites have homogeneous and uniform lamellar structure. The results of this study indicated that composites produced at −20 ◦ C had higher mechanical properties and lower degradation rate compared with −80 ◦ C and in overall, squid chitosan has better mechanical properties. These results indicated that chitosan isolated from the squid pen could be an alternative for existing commercial chitosan to be considered for biomedical applications. Acknowledgements The authors acknowledge the facilities as well as scientific and technical assistance from staff at Otago Centre for Electron Microscopy (OCEM) at the University of Otago. We would also like to thank Mr Damian Wallas for his help and technical support for XRD. The first author acknowledges the PhD scholarship by University of Otago, New Zealand. References [1] T.H. Silva, A. Alves, B.M. Ferreira, J.M. Oliveira, L.L. Reys, R.J.F. Ferreira, R.A. Sousa, S.S. Silva, J.F. Mano, R.L. Reis, Int. Mater. Rev. 57 (2012) 276–306. [2] J. Venkatesan, S.-K. Kim, Mar. Drugs 8 (2010) 2252–2266. [3] W.W. Thein-Han, R.D. Misra, Acta Biomater. 5 (2009) 1182–1197. [4] N.M. Alves, J.F. Mano, Int. J. Biol. Macromol. 43 (2008) 401–414. [5] M.L. Gisela, F.M. João, Biomed. Mater. 7 (2012) 054104. [6] M. Shimojoh, K. Fukushima, K. Kurita, Carbohydr. Polym. 35 (1998) 223–231. [7] I. Aranaz, M. Mengibar, R. Harris, I. Panos, B. Miralles, N. Acosta, G. Galed, A. Heras, Curr. Chem. Biol. 3 (2009) 203–230. [8] R. Minke, J. Blackwell, J. Mol. Biol. 120 (1978) 167–181. [9] N.E. Dweltz, Biochim. Biophys. Acta 51 (1961) 283–294. [10] K. Kurita, K. Tomita, T. Tada, S. Ishii, S.-I. Nishimura, K. Shimoda, J. Polym. Sci. A: Polym. Chem. 31 (1993) 485–491. [11] D.K. Youn, H.K. No, W. Prinyawiwatkul, Int. J. Food Sci. Technol. 48 (2013) 571–577. [12] F.A.A. Sagheer, M.A. Al-Sughayer, S. Muslim, M.Z. Elsabee, Carbohydr. Polym. 77 (2009) 410–419. [13] D. Raafat, H.-G. Sahl, Microb. Biotechnol. 2 (2009) 186–201. [14] E. Khor, in: E. Khor (Ed.), Chitin, 2nd ed., Elsevier, Oxford, 2014, pp. 51–66. [15] A. Shavandi, A.E.-D.A. Bekhit, A. Ali, Z. Sun, J.T. Ratnayake, Powder Technol. 273 (2015) 33–39. [16] A. Shavandi, A.E.-D.A. Bekhit, A. Ali, Z. Sun, Mater. Chem. Phys. 149–150 (2015) 607–616. [17] G. Chaussard, A. Domard, Biomacromolecules 5 (2004) 559–564. [18] K. Kurita, Prog. Polym. Sci. 26 (2001) 1921–1971.
[19] J. Xu, S.P. McCarthy, R.A. Gross, D.L. Kaplan, Macromolecules 29 (1996) 3436–3440. [20] Y. Zhang, M. Zhang, J. Biomed. Mater. Res. 61 (2002) 1–8. [21] S.H. Yu, S.J. Wu, J.Y. Wu, C.K. Peng, F.L. Mi, Molecules (Basel, Switz.) 18 (2012) 27–40. [22] G.S. Kumar, A. Thamizhavel, E.K. Girija, Mater. Lett. 76 (2012) 198–200. [23] M.V. Zugravu, R.A. Smith, B.T. Reves, J.A. Jennings, J.O. Cooper, W.O. Haggard, J.D. Bumgardner, J. Biomater. Appl. 28 (2013) 566–579. [24] A.C. Jones, B. Milthorpe, H. Averdunk, A. Limaye, T.J. Senden, A. Sakellariou, A.P. Sheppard, R.M. Sok, M.A. Knackstedt, A. Brandwood, D. Rohner, D.W. Hutmacher, Biomaterials 25 (2004) 4947–4954. [25] A.S. Lin, T.H. Barrows, S.H. Cartmell, R.E. Guldberg, Biomaterials 24 (2003) 481–489. [26] W.W. Thein-Han, R.D.K. Misra, Acta Biomater. 5 (2009) 1182–1197. [27] A. Cuneyt Tas, J. Non-Cryst. Solids 400 (2014) 27–32. ´ [28] J. Czechowska, A. Zima, Z. Paszkiewicz, J. Lis, A. Slósarczyk, Ceram. Int. 40 (2014) 5523–5532. [29] C.A. Schneider, W.S. Rasband, K.W. Eliceiri, Nat. Mater. 9 (2012) 671–675. [30] K.M. Rudall, W. Kenchington, Biol. Rev. 48 (1973) 597–633. [31] C. Chatelet, O. Damour, A. Domard, Biomaterials 22 (2001) 261–268. [32] L.L. Reys, S.S. Silva, J.M. Oliveira, S.G. Caridade, J.F. Mano, T.H. Silva, R.L. Reis, Biomed. Mater. 8 (2013) 1–11. [33] S.H. Teng, E.J. Lee, B.H. Yoon, D.S. Shin, H.E. Kim, J.S. Oh, J. Biomed. Mater. Res. A 88 (2009) 569–580. [34] Q. Hu, B. Li, M. Wang, J. Shen, Biomaterials 25 (2004) 779–785. [35] C. Meneghini, M.C. Dalconi, S. Nuzzo, S. Mobilio, R.H. Wenk, Biophys. J. 84 (2003) 2021–2029. [36] S.V. Dorozhkin, Biomaterials 1 (2011) 3–56. [37] R. Barabás, M. Czikó, I. Dékány, L. Bizo, E. Bogya, Chem. Pap. 67 (2013) 1414–1423. [38] M.C. Sunny, P. Ramesh, H.K. Varma, J. Mater. Sci.: Mater. Med. 13 (2002) 623–632. [39] A.T. Paulino, J.I. Simionato, J.C. Garcia, J. Nozaki, Carbohydr. Polym. 64 (2006) 98–103. [40] K.S. Katti, D.R. Katti, R. Dash, Biomed. Mater. 3 (2008) 034122. [41] A. Ouyang, Q. Gong, J. Liang, Adv. Eng. Mater. 17 (4) (2015) 460–466. [42] C.-Y. Hsieh, S.-P. Tsai, M.-H. Ho, D.-M. Wang, C.-E. Liu, C.-H. Hsieh, H.-C. Tseng, H.-J. Hsieh, Carbohydr. Polym. 67 (2007) 124–132. [43] K. Tuzlakoglu, C.M. Alves, J.F. Mano, R.L. Reis, Macromol. Biosci. 4 (2004) 811–819. [44] J.M. Oliveira, M.T. Rodrigues, S.S. Silva, P.B. Malafaya, M.E. Gomes, C.A. Viegas, I.R. Dias, J.T. Azevedo, J.F. Mano, R.L. Reis, Biomaterials 27 (2006) 6123–6137. [45] V. Karageorgiou, D. Kaplan, Biomaterials 26 (2005) 5474–5491. [46] Y. Kuboki, Q. Jin, H. Takita, J. Bone Joint Surg. 83 (2001) 105–115. [47] C.M. Murphy, M.G. Haugh, F.J. O’Brien, Biomaterials 31 (2010) 461–466. [48] H.E. Götz, M. Müller, A. Emmel, U. Holzwarth, R.G. Erben, R. Stangl, Biomaterials 25 (2004) 4057–4064. [49] F. Zhao, Y. Yin, W.W. Lu, J.C. Leong, W. Zhang, J. Zhang, M. Zhang, K. Yao, Biomaterials 23 (2002) 3227–3234. [50] J. Venkatesan, R. Pallela, I. Bhatnagar, S.K. Kim, Int. J. Biol. Macromol. 51 (2012) 1033–1042. [51] N. Shanmugasundaram, P. Ravichandran, P.N. Reddy, N. Ramamurty, S. Pal, K.P. Rao, Biomaterials 22 (2001) 1943–1951. [52] A. Sionkowska, J. Kozlowska, Int. J. Biol. Macromol. 52 (2013) 250–259. [53] B.D. Ulery, L.S. Nair, C.T. Laurencin, J. Polym. Sci. B: Polym. Phys. 49 (2011) 832–864. [54] A.M. Martins, R.C. Pereira, I.B. Leonor, H.S. Azevedo, R.L. Reis, Acta Biomater. 5 (2009) 3328–3336. [55] J. Gustavsson, M.P. Ginebra, E. Engel, J. Planell, Acta Biomater. 7 (2011) 4242–4252. [56] X. Wang, G. Song, T. Lou, W. Peng, J. Biomater. Sci. 20 (2009) 1995–2002. [57] X.-m. Pu, Q.-q. Yao, Y. Yang, Z.-z. Sun, Q.-q. Zhang, Int. J. Biol. Macromol. 51 (2012) 868–873. [58] A. Yoshida, T. Miyazaki, E. Ishida, M. Ashizuka, Mater. Trans. 45 (2004) 994–998.