- Email: [email protected]
Preparation and characterization of bioactive composite scaffolds from polycaprolactone nanofibers-chitosan-oxidized starch for bone regeneration
Accepted Manuscript Title: Preparation and characterization of bioactive composite scaffolds from polycaprolactone nanofibers-chitosan-oxidized starch for bone regeneration Author: Jhamak Nourmohammadi Azadeh Ghaee Samira Hosseini Liavali PII: DOI: Reference:
S0144-8617(15)01145-5 http://dx.doi.org/doi:10.1016/j.carbpol.2015.11.055 CARP 10574
To appear in: Received date: Revised date: Accepted date:
3-9-2015 19-10-2015 23-11-2015
Please cite this article as: Nourmohammadi, J., Ghaee, A., and Liavali, S. H.,Preparation and characterization of bioactive composite scaffolds from polycaprolactone nanofibers-chitosan-oxidized starch for bone regeneration, Carbohydrate Polymers (2015), http://dx.doi.org/10.1016/j.carbpol.2015.11.055 This is a PDF file of an unedited manuscript that has been accepted for publication. As a service to our customers we are providing this early version of the manuscript. The manuscript will undergo copyediting, typesetting, and review of the resulting proof before it is published in its final form. Please note that during the production process errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain.
Preparation and characterization of bioactive composite scaffolds from polycaprolactone nanofibers-chitosan-oxidized starch for bone regeneration
1- Jhamak Nourmohammadi,
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Department of Life science Engineering, Faculty of New Sciences and Technologies University of Tehran, P.O. Box 14395-1561, Tehran, Iran. E-mail: [email protected]
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2- Azadeh Ghaee
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Department of Life science Engineering, Faculty of New Sciences and Technologies University of Tehran, P.O. Box 14395-1561, Tehran, Iran. E-mail: [email protected]
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3- Samira Hosseini Liavali
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Department of Life science Engineering, Faculty of New Sciences and Technologies University of Tehran, P.O. Box 14395-1561, Tehran, Iran.
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E-mail: [email protected]
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*Corresponding author: Jhamak Nourmohammadi; PhD. Faculty of New Sciences and Technologies, University of Tehran, P.O. Box: 143951561,Tehran, Iran. Tel: +98-21-66118560 Email: [email protected]
Abstract The objective of this study was to fabricate and investigate the characteristics of a suitable scaffold for bone regeneration. Therefore, chitosan was combined with various amounts of oxidized starch through reductive alkylation process. Afterwards, chopped CaP-coated PCL nanofibers were added into the chitosan-starch composite scaffolds in order to obtain bioactivity and mimic bone extracellular matrix structure. Scanning electron microscopy confirmed that all scaffolds had well-interconnected porous structure. The mean pore size, porosity, and water uptake of the composite scaffolds increased by incorporation of higher amounts of starch, while this trend was opposite for compressive modulus and strength. Osteoblast-like cells (MG63) culturing on the scaffolds demonstrated that higher starch content could improve cell viability. Moreover, the cells spread and anchored well on the scaffolds, on which the surface was covered with a monolayer of cells.
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Keywords: Chitosan; Oxidized starch; Polycaprolactone; Calcium phosphate, Bone; Scaffold.
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1. Introduction Large bone defects are believed to be one of the major issues caused by trauma and infection. Despite several efforts such as using bone grafts and implants, there are still limitations and drawbacks in bone regeneration. Over the past few decades, bone tissue engineering (BTE) has emerged as a promising alternative to classical therapies (Bose, Roy, & Bandyopadhyay, 2012; Shrivats, Dermott, & Hollinger, 2014). One of the most important aspects of BTE is designing a suitable scaffold that could modulate bone healing and mimic the extracellular matrix (ECM) role in bone tissue. Thus, ideal scaffolds should consist of biodegradable and biocompatible materials, which possess appropriate pore size, porosity, mechanical properties, and osteo-conductivity (Guarino et al., 2013). However, single-phase scaffolds do not provide proper structural and mechanical features required for bone regeneration. Recently, using composite materials has gained much attention to improve the biodegradability and bioactivity (Hutmacher, Schantz, Lam, Tan, & Lim, 2007). Meanwhile, the mechanical and biological properties of the composite scaffolds can be manipulated in order to design a suitable structure similar to the features of the bone tissue (Hutmacher, Schantz, Lam, Tan, & Lim, 2007; Lee, & Shin, 2007). Various natural and synthetic biodegradable polymers have been studied as scaffolds for BTE (Puppi, Chiellini, Piras, & Chiellini, 2010). In this regard, natural hydrogels have gained much popularity because of their similarity to the extracellular matrix (ECM) and higher regeneration rate (Peppas, Hilt, Khademhosseini, & Langer, 2006). Chitosan is a linear, amino polysaccharide derived from chitin through deacetylation process. It is widely used in various fields of biomedical engineering due to its biocompatibility, accessibility, biodegradability, and antibacterial activities (Croisier, & Jérôme, 2013). Chitosan is barley used alone in BTE as it has poor mechanical properties, and high water sensitivity. Reports indicated that the functional properties of chitosan are generally improved by combining it with other natural biopolymers such as alginate (Baysal, Aroguz, Adiguzel, & Baysal, 2013), cellulose (Wu et al., 2004]), and gelatin (Cheng et al., 2010). Among different kinds of biopolymers, starch is one of the most promising material in the field of biomedical engineering, since it is biocompatible, cost-effective, and accessible (Hanafi, Irani, & Zulkifli, 2013; Xie, Pollet, Halley, & Avérous, 2013). Bourtoom, & Chinnan (2008) proposed that the water vapor permeability of rice starchchitosan blend films is lower than the films prepared only by chitosan. Martins et al. (2008) found that the water uptake and biological activity of chitosan hydrogel improved when combining with starch, while its compression strength decreased. As mentioned above, blending of chitosan with starch can improve some properties of chitosan-based hydrogels. However, there are still some limitations regarding their application as BTE scaffolds, including insufficient mechanical properties and bioactivity (Shakir, Jolly, Khan, Iram, & Khan, 2015). Over the past few years, the researchers have incorporated nano-hydroxyapatite into hydrogels to enhance such properties (Chesnutt et al., 2009; Lai, Shalumon, & Chen,2015; Barbani et al., 2011). However, the weak interfacial bonding and inhomogeneous distribution of ceramic fillers throughout the hydrogels were reported in such approaches. More recent studies suggested that hydrogel
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reinforcement with electrospun nanofibers improved its mechanical properties and biological activity (Hadisi, Nourmohammadi, & Mohammadi, 2015; Kai, Prabhakaran, Stahl, Eblenkamp, Wintermantel, & Ramakrishna, 2012). This study introduces a set of new bioactive nanocomposite scaffolds using chitosan, starch, and polycaprolactone (PCL) for BTE applications. Therefore, chitosan was combined with various amounts of oxidized starch through the reductive alkylation process. Afterwards, chopped calcium phosphate-coated PCL nanofibers were added into the fabricated chitosan-starch composite in order to achieve bioactivity and mimic bone ECM. Eventually, the morphology/structure, swelling, mechanical properties and biological response of MG 63 osteoblast-like cells cultivated on the surface of the scaffolds were investigated. 2. Materials and methods 2.1. Materials Chitosan (medium molecular weight, deacetylation degree 85%), acetic acid, formic acid, and polycaprolactone (Mn= 80000 Da) were purchased from Sigma (St. Louis, USA). Sodium periodate (NaIO4), calcium chloride (CaCl2), Tris (Hydroxy-methylamino-methane) ((CH2OH)3CNH2), hydrochloric acid (HCl), sodium hydrogen phosphate (Na2HPO4), and sodium hydroxide (NaOH) were obtained from Merck (Germany). The soluble potato starch was supplied by BioBasic Inc. (Canada). All the chemicals used in this study were of analytical grade. 2.2. Electrospun PCL nanofibers preparation In this study low toxic solvents of acetic acid and formic acid were chosen to produce elctrospun PCL nanofibers. The appropriate amount of PCL was dissolved in the mixture of formic acid and acetic acid with a ratio of 3:1 (v/v) in order to prepare 13 wt% solution. Afterwards, the resulting solution was filled in a 1 ml syringe with a 22 G blunted stainless steel needle. Then, electrospinning (Fanavaran Nano-Meghyas, Iran) was carried out at 11 kV with a constant flow rate of 0.4 ml.h-1. The rotating mandrel (50 m/min) was chosen as a collector and the needle tip distance from the collector was 10 cm. 2.3. Calcium phosphate (CaP) deposition on/in the electrospun PCL mats The surface of the PCL mat was modified using a 2 M NaOH solution in a shaker incubator (T=37oC). After 3 h, the sample was removed from the NaOH solution, carefully washed with deionized water and dried at room temperature. For CaP deposition, the NaOH-treated sample was dipped in calcium and phosphate rich solutions, alternately (Taguchi, Kishida, & Akashi, 1999). Briefly, the fabricated mat was soaked into 10 ml of 0.5 M CaCl2 (pH=7.4) at 37oC for 30 min, washed rapidly with deionized water, and immediately soaked into 10 ml of 0.3 M Na2HPO4 (pH=8.5) at 37 o C for another 30 min. After repeating each cycle 3 more times, the mat was rinsed thoroughly with deionized water and then dried at room temperature. 2.4. Preparation of the oxidized starch Oxidized starch was initially prepared through the procedure described by Hermanson (1996). Briefly, 1.25 ml of sodium iodate (NaIO4) solution (10 mg/ml) and 2% (w/v) starch solution were mixed thoroughly in a light-protected glass vessel at room temperature. After 30 min, 1.125 ml of glycerin was added to the reaction vessel and stirred for another 10 minutes. 2.5. Preparation of composite scaffolds
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Chitosan powder (1 g) was added to the 1% (v/v) acetic acid and stirred overnight to have 1% (w/v) clear solution. Subsequently, different amounts of oxidized starch and chitosan solutions were mixed thoroughly at room temperature for about 1 h. Meanwhile, 1% (w/v) of chopped CaP-coated PCL nanofibers were added into the solution. After stirring for 2 h, the mixture was poured into 24-well tissue culture polystyrene plates, frozen at -20 oC overnight and then freeze-dried. The code and composition of the fabricated scaffolds are presented in Table 1. 3. Characterization 3.1. Characterization of electrospun PCL mat The morphologies of electrospun fibers before and after CaP deposition were observed using a Scanning electron microscopy (SEM, Lecia Cambridge S360). The diameter and distribution of the electrospun fibers were measured by Image J software (National institutes of Health, Bethesda, Maryland, USA). Approximately 100 random fibers of different SEM images were analyzed. The presence of calcium and phosphorus elements on the deposited layer was also evaluated by Electron dispersive spectroscopy (EDS). The changes in the chemical structure of electrospun PCL nanofibers after CaP deposition were studied by Fourier transform infrared spectrometer (FTIR; BRUKER IFS 48) in the range of 400–4000 cm-1. Moreover, the phase compositions of electrospun PCL nanofibers were determined by X-ray diffractometry (XRD; X’Pert Pro MPD) using Cukα radiation. Metrohm pH meter 827 determined the changes in pH value during alternate soaking process. 3.2. Characterization of fabricated composite scaffolds 3.2.1. IR spectroscopy The structural analysis of pure starch, oxidized starch, pure chitosan, and composite scaffolds were examined through Fourier transform infrared spectroscopy in the attenuated total reflectance mode (FTIR-ATR). 3.2.2. Microstructure of the composite scaffolds SEM was performed to examine the internal morphology of the fabricated scaffolds. The pore size of each sample, the total area of the pores in each cross section (AP), and the total area of each cross section (AT) were calculated using Image J software. The measurements were based on the five SEM images of each sample (n=5). Next, the porosity percentages of the fabricated scaffolds were calculated using the following equation (Annabi, Fathi, Mithieux, Weiss, & Dehghani, 2011): Eq. (1) 3.2.3. Water uptake properties To determine the amount of water uptake, the pre-weighted scaffolds (Wo) were immersed in deionized water (T=37oC) and allowed to completely swell. After that, the samples were removed; the excess water was wiped off with filter paper and weighted (Ws). All measurements were carried out for three samples (n=3). The amount of water uptake was calculated using the following equation (Nasri-Nasrabadi et al., 2014): Water uptake (g/g) Eq. (2) 3.2.4. Mechanical properties The mechanical properties of the scaffolds (d=6 mm, h=12 mm) were determined using Zwick/Roell Z050, equipped with a 0.1 KN load cell with a crosshead speed of 10
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mm/min. The compressive modulus was calculated by the linear section of the stressstrain curve. Moreover, the compressive strength was measured by drawing a line, starting at 1% strain. The intersection of this line with the stress-strain curve was defined as the compressive strength (Kim, Park, Kim, Wada, & Kaplan, 2005).
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3.3. Cytocompatibility assessment 3.3.1. Cell culture Human osteoblast-like cells, MG63, from National Cell Bank of Iran (NCBI; Pasteur institute) were maintained in Dulbecco’s Modification of Eagles Medium (DMEM; GIBCO, Scotland) supplemented with 10% Fetal Bovine Serum (FBS; Gibco, Renfrewshire, Scotland), 100 U/mL penicillin (Sigma, Saint Louis, USA), and 100 µg/mL streptomycin (Sigma, Saint Louis, USA) at 37 °C in a humidified incubator with 5% CO2. Triplicate specimens (n=3) of each specimen were put into petri dishes, disinfected via interval immersion in ethanol and phosphate buffered saline (PBS) for 5 times, and then washed twice with culture medium before cell seeding. 3.3.2. Cell viability characterization Indirect 3-(4, 5-dimethylthiazol-2-yl)-2, 5-diphenyltetrazolium-bromide (MTT, Sigma, Saint Louis, USA) assay based on extraction method, was used to assess cell viability (Mirahmadi, Tafazzoli-Shadpour, Shokrgozar, & Bonakdar, 2013). Briefly, the 3 and 7day samples’ extractions were added to a 96-well plate containing MG 63 cells for 24 h. Next, the culture medium was removed, replaced by 100 µl of 3 and 7-day extracts and then kept in an incubator. After 24 h, 100 µl MTT solution (0.5 mg/ml) in PBS was added to each cell containing well and incubated for about 4 h at 37 oC. After that, the formed purple formazan crystals were dissolved in 50 µl DMSO (D2650, Sigma, Saint Louis, USA) for 10 min and then absorbance was read at 545 nm using an ELISA Reader (Stat Fax-2100; GMI, Inc., Miami, FL, USA). The serum free culture medium without scaffolds was chosen to be the control group. 3.3.3. Cell morphology observation The morphology of the 5-day cultured MG 63 cells (5x 104 cells/ cm2) was examined using SEM. After 5 days of culture, the cells were fixed using 4% (v/v) glutaraldehyde solution in PBS at 4oC for 30 minutes. Afterwards, the samples were washed thoroughly with deionized water, dehydrated by graded alcohol solution (10% ethanol increments; each step 10 minutes) and finally dried at room temperature. In the end, the dried samples were mounted on an aluminum stub and sputter coated with gold prior to imaging. 3.4. Statistical analysis All data was expressed as mean ± standard deviation (SD). A one-way analysis of variance (ANOVA) was performed to compare any significant difference using StatPlus software. A p< 0.05 was considered statistically meaningful. 4. Results and discussion 4.1. Structure and morphology of electrospun PCL mat Fig. 1a-c shows the SEM image, fiber diameter distribution, and EDS spectrum of pristine electrospun PCL nanofibers. The SEM image consisted of beadless fibers with an average diameter of 149.7 ± 31 nm, presenting a diameter size between 65 nm and 250 nm (Fig.1 a, b). As shown in Fig. 1c, the corresponding EDS spectrum consists of carbon and oxygen. However, new spherical and plate-like morphologies appeared on the surface of electrospun PCL mat after soaking in calcium and phosphate rich solutions
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(Fig. 1d). As shown by EDS analysis, the precipitated layer consists of calcium and phosphate elements (Fig. 1e). Fig. 2 depicts FTIR spectra and XRD patterns of the NaOH-treated PCL nanofibers before and after alternate soaking process. As illustrated in Fig. 2a, the NaOH-treated PCL spectrum has characteristic peaks of 1166 cm-1 (COC symmetric stretching), 1186 cm-1 (OC–O stretching), 1242 cm-1 (COC asymmetric stretching) 1722 cm-1 (C=O stretching), 2871 cm-1 (C-H symmetric stretching), and 2950 cm-1 (C-H asymmetric stretching) (Elzein, Nasser-Eddine, Delaite, Bistac, & Dumas, 2004). However, after the alternate soaking process, new peaks appeared at 550–650 cm-1 (PO43- bending vibration), 876 cm-1 (P-O(H) stretching) and 1000-1100 cm-1 (PO43- asymmetric stretching) (Nourmohammadi, Sadrnezhaad, & Ghader, 2008; Verma, Katti, & Katti, 2006), suggesting that CaP successfully deposited throughout the electrospun PCL mat. The XRD pattern of the resulting NaOH-treated PCL nanofibers (Fig. 2b) exhibited two distinct peaks at about 2θ= 21o and 23o, which relate to the (110) and (200) planes in crystalline PCL (Baji, Wong, Liu, Li, & Srivatsan, 2007). However, after CaP deposition, some changes in the XRD pattern of PCL nanofibers were observed. As indicated from Fig. 2b, the appearance of new peaks at 11.61o, 20.81o, and 29.18o are related to the formation of brushite crystals (Dicalcium phosphate dihydrate; DCPD) on the PCL mat (Mandel, & Tas, 2010 ). Moreover, apatitic CaP is indicated by the appearance of weak peaks at 26o and 32o (Nourmohammadi, Sadrnezhaad, & Ghader, 2008). The XRD results support the SEM and EDS data. The formation of CaP layer throughout the PCL electrospun mat is due to the presence of carboxyl groups. In other words, during NaOH treatment, PCL was partially hydrolyzed (Park et al., 2007), which resulted in creation of carboxylate groups on the surface of PCL mat. These negatively charged carboxylate groups could serve as an inductive site for CaP nucleation (Kawashita et al., 2003; Zheng, Xiong, & Zhang, 2014)]. Therefore, heterogeneous nucleation began through bonding of calcium ions with these negatively charged carboxylate groups after the initial soaking in the calcium rich solution (pH=7.4). Subsequent soaking in the phosphate rich solution resulted in CaP nuclei formation, followed by growth throughout the electrospun mats during soaking time. As indicated by XRD data (Fig. 2b), both brushite and apatitic CaP deposited on the electrospun PCL mats. Moreover, the pH value of CaCl2 solution decreased from 7.4 to 5.95 at the first soaking cycle. This finding is presumably due to the formation of acidic by-product during the hydrolysis reaction of PCL nanofibers, which accelerates after NaOH treatment. Therefore, the reduction in pH should be the reason for brushite precipitation; this is in agreement with previous studies indicating that brushite crystals start to precipitate on the materials in acidic pH ranging from 2 to 6.5 (Mandel, & Tas, 2010). In addition, brushite is a primary phase that can convert to thermodynamically stable apatitic CaP (Mavis, Demirtaş, Gümüşderelioğlu, Gündüz, & Colak, (2009). 4.2. Characterization of fabricated composite scaffolds 4.2.1. IR spectroscopy The IR spectra of pure chitosan and starch as well as oxidized starch and composite scaffolds are shown in Fig. 3. Chitosan and starch present a peak at around 1161 cm−1 due to CO stretching in glycosidic bonds (Silva, 2012; Sevenou, Hill, Farhat, & Mitchell, 2002). In comparison with the starch spectrum, a new peak at 1720 cm-1 appeared after starch oxidation process, which is related to the C=O stretching in aldehyde groups
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(Socrates, 2004). This suggests that aldehyde groups were successfully generated in starch structure without breaking its glycosidic bands. In composite samples, the imine conjugation between aldehyde groups of starch and NH2 of chitosan were observed at 1645 cm-1. In addition, amide II linkage between free carboxyl groups in PCL and NH2 groups in chitosan was also detected at 1532 cm-1. Importantly, the peaks related to the free aldehyde groups at 1720-1740 cm-1 (Socrates, 2004) were not observed in the IR spectra, suggesting that all of the aldehyde groups in starch were covalently cross-linked with chitosan. 4.2.2. Microstructure of the composite scaffolds The SEM images of the fabricated composite scaffolds and their pore size distribution are shown in Fig. 4. In addition, average pore size and the porosity percentage of each scaffold are summarized in Table 2. In order to characterize the presence and distribution of CaP-coated PCL nanofibers in the porous structure, an EDS elemental mapping of Ca and P were performed. As shown in Fig. S1-4 (supplementary files), both Ca and P were uniformly distributed throughout the fabricated scaffolds. This suggests that the CaPcoated PCL nanofibers were dispersed in the hydrogel matrix. In the sample S1, the pore size varied from 34.30 ± 5.33 µm to 148.67 ± 3.43 µm with the average size of 73.40 ± 28.19 µm. However, the pore size distribution became broader as starch content increased (Fig. S5; supplementary files). Furthermore, the average size of the pores rose to 115.32 ± 37.62 µm in S2, 122.068 ± 51.92 µm in S3, and 135.34 ± 63.03 µm in S4 (Table 2). Thus, increasing starch content resulted in larger pore formation, which makes such scaffolds suitable for BTE applications. It has been reported that the optimal mean pore size needed for BTE scaffolds is in the range of 100-135 µm (Murphy, Haugh, & O'Brien, 2010). As shown in Table 2, the porosity percentage also increased with the increase in starch content. It can be speculated that the rise in the pore size and porosity is due to the formation of imine conjugation (shiff base) between the aldehyde in oxidized starch and amino groups of chitosan. It is generally held that a water molecule was displaced during imine formation (Clayden, Greeves, & Warren, 2012). Therefore, elimination of the formed water molecules during freeze-drying may increase both pore size and porosity values. 4.2.3. Water uptake properties The changes in the equilibrium water uptake of the scaffolds containing various amounts of starch are represented in Table 2. The lowest water uptake occurred in the scaffold consisting of 3% starch (S1; 11.35 ± 0.34 g/g), while this amount increased to 11.47 ± 0.63 (g/g) in S2, 12.96 ± 0.97 (g/g) in S3, and 13.84 ± 1.32 (g/g) in S4. These results are probably due to the presence of more hydroxyl groups and larger pore size in samples having more starch. Previous studies also confirmed that the pore size of the scaffold has effect on the water uptake (Hadisi, Nourmohammadi, & Mohammadi, 2015; Mirahmadi, Tafazzoli-Shadpour, Shokrgozar, & Bonakdar, 2013). 4.2.4. Mechanical properties Determining the compressive properties is another key factor in designing bone tissue engineering scaffolds. The compressive modulus and strength of the fabricated hybrid composites are represented in Fig. 5. It can be seen that the highest compressive modulus was observed for S1 (2.64 ± 0.19 MPa). The modulus decreased with increasing starch content, starting at 2.43 ± 0.13 MPa in S2, 1.6 ± 0.25 MPa in S3, and reaching 0.8 ± 0.15 MPa in S4. A similar trend was observed for compressive strength of the scaffolds. The
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compressive strength of sample S1 (1.2 ± 0.05 MPa) decreased to 0.89 ± 0.025 MPa in S2, 0.54 ± 0.09 MPa in S3, and 0.44 ± 0.025 MPa in S4. It should be noted that the compressive properties of materials are highly affected by their solid content (Gibson, & Ashby, 1988). The rise in the pore size and porosity percentage that occurred at higher starch concentration may cause this reduction. This is in agreement with previous reports that increasing pore sizes and porosity led to a decrease in the compressive strength and modulus of scaffolds (Mirahmadi, Tafazzoli-Shadpour, Shokrgozar, & Bonakdar, 2013; Gibson, & Ashby, 1988). 4.2.5. Cytocompatibility assay The results of the cell viability from indirect MTT assay of all samples in comparison with the control group for days 3 and 7 are illustrated in Fig. 6a. As shown in Fig. 6a, all samples have cellular viability more than 90%, which suggests that the fabricated scaffolds are biocompatible and did not show any cytotoxic effects on cells. There is no significant difference between MTT values for different samples after exposed to 3 days extracts. After 7 days, the cell viability increased as the amount of oxidized starch rose (p < 0.05). This can be related to the formation of high-crosslinked structure by increasing oxidized starch amounts, which led to the release of less toxic agents in the culture medium and consequently increased cellular viability. The morphology of MG 63 cells cultured on the scaffolds after 5 days is shown in Fig. 6b. The cells spread on the surface of all scaffolds by their filopodia and they had penetrated inside the pores. It was observed that the cells attached well on the matrix and were either flat or round in morphology. Here, the flat cells attached tightly on the surface by their lamellopodia, and the confluent monolayers of the cells were observed through the pores’ walls. Moreover, the round MG 63 cells attached to the surface by their filopodia. Both SEM images and MTT results suggest that the composites are biocompatible and can be considered as suitable scaffolds for BTE applications. Martins et.al (2008) confirmed that the adhesion, spreading, and growth of both osteoblast and fibroblast cells are enhanced by blending chitosan with starch, which is in agreement with the result of this study. Conclusion This study offers a new bioactive composite scaffold based on chitosan-oxidized starch for potential bone tissue regeneration. At the outset, oxidized starch prepared by periodate oxidation was blended with chitosan through the reductive alkylation process. Meanwhile, CaP-coated electrospun PCL nanofibers were added into the oxidized starchchitosan solution, followed by freeze-drying. The average pore size, porosity, swelling ratio, and viability of osteoblast-like cells (MG 63) increased by adding more starch, while the compressive modulus and strength were reported to decline. In addition, the cells were spread well and anchored, on which the scaffold was covered with a monolayer of cells. References: Park, J. S., Kim, J. M., Jun, L. S., Geun, L. S., Young-Keun, J., Kim, S. E., & Lee, S. C. (2007). Surface Hydrolysis of Fibrous Poly(ε-caprolactone) Scaffolds for Enhanced Osteoblast Adhesion and Proliferation. Macromolecular Research , 15 (5), 424-429. Mavis, B., Demirtaş, T. T., Gümüşderelioğlu, M., Gündüz, G., & Colak, U. (2009). Synthesis, characterization and osteoblastic activity of polycaprolactone nanofibers coated with biomimetic calcium phosphate. Acta biomaterialia , 5 (8), 3098-3111.
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Gibson, L. J. A., & Ashby, M. F. (1988). Cellular Solids: Structure and Properties. Oxford: Pergamon Press. Hutmacher, D. W., Schantz, J. T., Lam, C. X., Tan, K. C., & Lim, T. C. (2007). State of the art and future directions of scaffold-based bone engineering from a biomaterials perspective. Journal Of Tissue Engineering And Regenerativ , 4 (1), 245-260. Hadisi, Z., Nourmohammadi, J., & Mohammadi, J. (2015). Composite of porous starchsilk fibroin nanofiber-calcium phosphate for bone regeneration. Ceramics International , 41, 10745-10754. Hanafi, I., Irani, M., & Zulkifli, A. (2013). Starch-Based Hydrogels: Present Status and Applications. International Journal of Polymeric Materials and Polymeric Biomaterials , 62, 411-420. Hermanson, G. T. (1996). Bioconjugate Techniques. Academic Press. Kawashita, M., Nakao, M., Minoda, M., Kim, H. M., Beppu, T., Miyamoto, T., Nakamura, T. (2003). Apatite-forming ability of carboxyl group-containing polymer gels in a simulated body fluid. Biomaterials , 24 (14), 2477-2484. Kai, D., Prabhakaran, M. P., Stahl, B., Eblenkamp, M., Wintermantel, E., & Ramakrishna, S. (2012). Mechanical properties and in vitro behavior of nanofiberhydrogel composites for tissue engineering applications. Nanotechnology , 23 (9). Kim, U. J., Park, J., Kim, H. J., Wada, M., & Kaplan, D. L. (2005). Three-dimensional aqueous-derived biomaterial scaffolds from silk fibroin. Biomaterials , 26, 2775-2785. Lai, G. J., Shalumon, K. T., & Chen, J. P. (2015). Response of human mesenchymal stem cells to intrafibrillar nanohydroxyapatite content and extrafibrillar nanohydroxyapatite in biomimetic chitosan/silk fibroin/nanohydroxyapatite nanofibrous membrane scaffolds. International Journal of Nanomedicine , 10, 567-584. Lee, S. H., & Shin, H. (2007). Matrices and scaffolds for delivery of bioactive molecules in bone and cartilage tissue engineering. Advanced Drug Delivery Reviews , 59, 339-359. Nasri-Nasrabadi, B., Mehrasa, M., Rafienia, M., Bonakdar, S., Behzad, T., & Gavanji, S. (2014). Porous starch/cellulose nanofibers composite prepared by salt leaching technique for tissue engineering. Carbohydrate Polymers , 108, 232-238. Nourmohammadi, J., Sadrnezhaad, S., & Ghader, A. B. (2008). Bone-like apatite layer formation on the new resin-modified glass-ionomer cement. Journal of Materials Science: Materials in Medicine , 19, 3507-3514. Murphy, C. M., Haugh, M. G., & O'Brien, F. J. (2010). The effect of mean pore size on cell attachment, proliferation and migration in collagen–glycosaminoglycan scaffolds for bone tissue engineering. Biomaterials , 31 (3), 461-466. Mandel, S., & Tas, A. C. (2010). Brushite (CaHPO4·2H2O) to octacalcium phosphate (Ca8(HPO4)2(PO4)4·5H2O) transformation in DMEM solutions at 36.5 °C. Materials Science and Engineering: C , 30 (2), 245-254. Martins, A. M., Santos, M. I., Azevedo, H. S., Malafaya, P. B. , & Reis, R. L. (2008). Natural origin scaffolds with in situ pore forming capability for bone tissue engineering applications. Acta Biomaterialia 4 (6), 1637-1645. Mirahmadi, F., Tafazzoli-Shadpour, M., Shokrgozar, M. A., & Bonakdar, S. (2013). Enhanced mechanical properties of thermosensitive chitosan hydrogel by silk fibers for cartilage tissue engineering. Materials Science and Engineering: C 33 (2013) 4786-4794. , C33, 4786-4794.
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Puppi, D., Chiellini, F., Piras, A. M., & Chiellini, E. (2010). Polymeric materials for bone and cartilage repair. Polymer Science , 35 (4), 403–440. Peppas, N. A. ,Hilt, J. Z. , Khademhosseini, A. , & Langer, R. (2006). Hydrogels in biology and medicine: from molecular principles to bionanotechnology. Advanced Materials , 18, 1345-1360. Sevenou, O., Hill, S., Farhat, I., & Mitchell, J. (2002). Organisation of the external region of the starch granule as determined by infrared spectroscopy. International Journal of Biological Macromolecules , 31, 79-85. Silva, S. M. L., Braga, C. R. C., Fook, M. V. L., Raposo, C. M. O., Carvalho, L.H., & Canedo, E. L. (2012). Application of Infrared Spectroscopy to Analysis of Chitosan/Clay Nanocomposites. In T. Theophanides (Ed.), Infrared Spectroscopy - Materials Science, Engineering and Technology (pp. 43-63). InTech. Shakir, M., Jolly, R., Khan, M. S., Iram, N. E., & Khan, H. M. (2015). Nanohydroxyapatite/chitosan–starch nanocomposite as a novel bone construct: Synthesis and in vitro studies. International Journal of Biological Macromolecules , 80, 282-292. Shrivats, A. R., Dermott, M. C., & Hollinger, J. O. (2014). Bone tissue engineering: state of the union. Drug discovery today , 19, 781-786. Socrates, G. (2004). in: Infrared and Raman Characteristic Group Frequencies: Tables andCharts. John Wiley & Sons. Taguchi, T., Kishida, A., & Akashi, M. . (1999). Apatite formation on/in hydrogel matrices using an alternate soaking process: II. Effect of swelling ratios of poly (vinyl alcohol) hydrogel matrices on apatite formation. Journal of Biomaterials Science, Polymer Edition , 10, 331-339.
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Fig.1. (a) SEM image of electrospun PCL fibers (b) fiber diameter distribution (c) EDS spectrum of pristine PCL fibers (d) SEM image of CaP-coated PCL nanofibers, and (e) EDS spectrum of CaP-coated PCL nanofibers. Fig.2. Changes in the (a) IR spectra and (b) XRD patterns of PCL nanofibers before and after CaP deposition. Fig.3. The IR spectra of chitosan, starch, oxidized starch, and composite scaffolds. Fig.4. The SEM images of composite scaffolds (a) S1, (b) S2, (c) S3, and (d) S4. Fig.5. The (a) compressive modulus and (b) compressive strength of each scaffold (*** p < 0.001).
Fig.6. (a) The viability of MG 63 cells after exposed to scaffolds' extract at different days. (b) the morphology of MG 63 cells culture on the scaffolds after 5 days (*p < 0.05) . Table 1 The code and composition of each fabricated composite scaffolds
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Chitosan
Oxidized starch
S1 S2 S3 S4
97 93 90 85
3 7 10 15
CaP-coated PCL electrospun fibers (w/v %) 1 1 1 1
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Table 2 The average pore size, porosity, and equilibrium water uptake of each scaffold Porosity (%) 54.263 ± 3.27 60.58 ± 4.1 66.97 ± 5.76 71.91 ± 2.32
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Equilibrium water uptake (g/g) 11.35 ± 0.34 11.47 ± 0.63 12.96 ± 0.97 13.84 ± 1.32
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S1 S2 S3 S4
Average pore size (µm) 73.40 ± 28.19 115.32 ± 37.62 122.068 ± 51.92 135.34 ± 63.03
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Oxidized starch prepared by periodate oxidation was blended with chitosan through the reductive alkylation process. The PCl nanofibers were prepared by electrospinning using solvent mixture formic acid/acetic acid. The calcium phosphate (CaP) was deposited throughout the electrospun PCl mat using alternate soaking process. The composite scaffolds from CaP-coated PCl nanofibers-chitosan-oxidized starch were prepared via freeze-drying. Adhesion and viability of osteoblast-like cells (MG 63) were investigated on the scaffolds.
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S0144-8617(15)01145-5 http://dx.doi.org/doi:10.1016/j.carbpol.2015.11.055 CARP 10574
To appear in: Received date: Revised date: Accepted date:
3-9-2015 19-10-2015 23-11-2015
Please cite this article as: Nourmohammadi, J., Ghaee, A., and Liavali, S. H.,Preparation and characterization of bioactive composite scaffolds from polycaprolactone nanofibers-chitosan-oxidized starch for bone regeneration, Carbohydrate Polymers (2015), http://dx.doi.org/10.1016/j.carbpol.2015.11.055 This is a PDF file of an unedited manuscript that has been accepted for publication. As a service to our customers we are providing this early version of the manuscript. The manuscript will undergo copyediting, typesetting, and review of the resulting proof before it is published in its final form. Please note that during the production process errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain.
Preparation and characterization of bioactive composite scaffolds from polycaprolactone nanofibers-chitosan-oxidized starch for bone regeneration
1- Jhamak Nourmohammadi,
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Department of Life science Engineering, Faculty of New Sciences and Technologies University of Tehran, P.O. Box 14395-1561, Tehran, Iran. E-mail: [email protected]
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2- Azadeh Ghaee
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Department of Life science Engineering, Faculty of New Sciences and Technologies University of Tehran, P.O. Box 14395-1561, Tehran, Iran. E-mail: [email protected]
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3- Samira Hosseini Liavali
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Department of Life science Engineering, Faculty of New Sciences and Technologies University of Tehran, P.O. Box 14395-1561, Tehran, Iran.
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E-mail: [email protected]
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*Corresponding author: Jhamak Nourmohammadi; PhD. Faculty of New Sciences and Technologies, University of Tehran, P.O. Box: 143951561,Tehran, Iran. Tel: +98-21-66118560 Email: [email protected]
Abstract The objective of this study was to fabricate and investigate the characteristics of a suitable scaffold for bone regeneration. Therefore, chitosan was combined with various amounts of oxidized starch through reductive alkylation process. Afterwards, chopped CaP-coated PCL nanofibers were added into the chitosan-starch composite scaffolds in order to obtain bioactivity and mimic bone extracellular matrix structure. Scanning electron microscopy confirmed that all scaffolds had well-interconnected porous structure. The mean pore size, porosity, and water uptake of the composite scaffolds increased by incorporation of higher amounts of starch, while this trend was opposite for compressive modulus and strength. Osteoblast-like cells (MG63) culturing on the scaffolds demonstrated that higher starch content could improve cell viability. Moreover, the cells spread and anchored well on the scaffolds, on which the surface was covered with a monolayer of cells.
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Keywords: Chitosan; Oxidized starch; Polycaprolactone; Calcium phosphate, Bone; Scaffold.
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1. Introduction Large bone defects are believed to be one of the major issues caused by trauma and infection. Despite several efforts such as using bone grafts and implants, there are still limitations and drawbacks in bone regeneration. Over the past few decades, bone tissue engineering (BTE) has emerged as a promising alternative to classical therapies (Bose, Roy, & Bandyopadhyay, 2012; Shrivats, Dermott, & Hollinger, 2014). One of the most important aspects of BTE is designing a suitable scaffold that could modulate bone healing and mimic the extracellular matrix (ECM) role in bone tissue. Thus, ideal scaffolds should consist of biodegradable and biocompatible materials, which possess appropriate pore size, porosity, mechanical properties, and osteo-conductivity (Guarino et al., 2013). However, single-phase scaffolds do not provide proper structural and mechanical features required for bone regeneration. Recently, using composite materials has gained much attention to improve the biodegradability and bioactivity (Hutmacher, Schantz, Lam, Tan, & Lim, 2007). Meanwhile, the mechanical and biological properties of the composite scaffolds can be manipulated in order to design a suitable structure similar to the features of the bone tissue (Hutmacher, Schantz, Lam, Tan, & Lim, 2007; Lee, & Shin, 2007). Various natural and synthetic biodegradable polymers have been studied as scaffolds for BTE (Puppi, Chiellini, Piras, & Chiellini, 2010). In this regard, natural hydrogels have gained much popularity because of their similarity to the extracellular matrix (ECM) and higher regeneration rate (Peppas, Hilt, Khademhosseini, & Langer, 2006). Chitosan is a linear, amino polysaccharide derived from chitin through deacetylation process. It is widely used in various fields of biomedical engineering due to its biocompatibility, accessibility, biodegradability, and antibacterial activities (Croisier, & Jérôme, 2013). Chitosan is barley used alone in BTE as it has poor mechanical properties, and high water sensitivity. Reports indicated that the functional properties of chitosan are generally improved by combining it with other natural biopolymers such as alginate (Baysal, Aroguz, Adiguzel, & Baysal, 2013), cellulose (Wu et al., 2004]), and gelatin (Cheng et al., 2010). Among different kinds of biopolymers, starch is one of the most promising material in the field of biomedical engineering, since it is biocompatible, cost-effective, and accessible (Hanafi, Irani, & Zulkifli, 2013; Xie, Pollet, Halley, & Avérous, 2013). Bourtoom, & Chinnan (2008) proposed that the water vapor permeability of rice starchchitosan blend films is lower than the films prepared only by chitosan. Martins et al. (2008) found that the water uptake and biological activity of chitosan hydrogel improved when combining with starch, while its compression strength decreased. As mentioned above, blending of chitosan with starch can improve some properties of chitosan-based hydrogels. However, there are still some limitations regarding their application as BTE scaffolds, including insufficient mechanical properties and bioactivity (Shakir, Jolly, Khan, Iram, & Khan, 2015). Over the past few years, the researchers have incorporated nano-hydroxyapatite into hydrogels to enhance such properties (Chesnutt et al., 2009; Lai, Shalumon, & Chen,2015; Barbani et al., 2011). However, the weak interfacial bonding and inhomogeneous distribution of ceramic fillers throughout the hydrogels were reported in such approaches. More recent studies suggested that hydrogel
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reinforcement with electrospun nanofibers improved its mechanical properties and biological activity (Hadisi, Nourmohammadi, & Mohammadi, 2015; Kai, Prabhakaran, Stahl, Eblenkamp, Wintermantel, & Ramakrishna, 2012). This study introduces a set of new bioactive nanocomposite scaffolds using chitosan, starch, and polycaprolactone (PCL) for BTE applications. Therefore, chitosan was combined with various amounts of oxidized starch through the reductive alkylation process. Afterwards, chopped calcium phosphate-coated PCL nanofibers were added into the fabricated chitosan-starch composite in order to achieve bioactivity and mimic bone ECM. Eventually, the morphology/structure, swelling, mechanical properties and biological response of MG 63 osteoblast-like cells cultivated on the surface of the scaffolds were investigated. 2. Materials and methods 2.1. Materials Chitosan (medium molecular weight, deacetylation degree 85%), acetic acid, formic acid, and polycaprolactone (Mn= 80000 Da) were purchased from Sigma (St. Louis, USA). Sodium periodate (NaIO4), calcium chloride (CaCl2), Tris (Hydroxy-methylamino-methane) ((CH2OH)3CNH2), hydrochloric acid (HCl), sodium hydrogen phosphate (Na2HPO4), and sodium hydroxide (NaOH) were obtained from Merck (Germany). The soluble potato starch was supplied by BioBasic Inc. (Canada). All the chemicals used in this study were of analytical grade. 2.2. Electrospun PCL nanofibers preparation In this study low toxic solvents of acetic acid and formic acid were chosen to produce elctrospun PCL nanofibers. The appropriate amount of PCL was dissolved in the mixture of formic acid and acetic acid with a ratio of 3:1 (v/v) in order to prepare 13 wt% solution. Afterwards, the resulting solution was filled in a 1 ml syringe with a 22 G blunted stainless steel needle. Then, electrospinning (Fanavaran Nano-Meghyas, Iran) was carried out at 11 kV with a constant flow rate of 0.4 ml.h-1. The rotating mandrel (50 m/min) was chosen as a collector and the needle tip distance from the collector was 10 cm. 2.3. Calcium phosphate (CaP) deposition on/in the electrospun PCL mats The surface of the PCL mat was modified using a 2 M NaOH solution in a shaker incubator (T=37oC). After 3 h, the sample was removed from the NaOH solution, carefully washed with deionized water and dried at room temperature. For CaP deposition, the NaOH-treated sample was dipped in calcium and phosphate rich solutions, alternately (Taguchi, Kishida, & Akashi, 1999). Briefly, the fabricated mat was soaked into 10 ml of 0.5 M CaCl2 (pH=7.4) at 37oC for 30 min, washed rapidly with deionized water, and immediately soaked into 10 ml of 0.3 M Na2HPO4 (pH=8.5) at 37 o C for another 30 min. After repeating each cycle 3 more times, the mat was rinsed thoroughly with deionized water and then dried at room temperature. 2.4. Preparation of the oxidized starch Oxidized starch was initially prepared through the procedure described by Hermanson (1996). Briefly, 1.25 ml of sodium iodate (NaIO4) solution (10 mg/ml) and 2% (w/v) starch solution were mixed thoroughly in a light-protected glass vessel at room temperature. After 30 min, 1.125 ml of glycerin was added to the reaction vessel and stirred for another 10 minutes. 2.5. Preparation of composite scaffolds
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Chitosan powder (1 g) was added to the 1% (v/v) acetic acid and stirred overnight to have 1% (w/v) clear solution. Subsequently, different amounts of oxidized starch and chitosan solutions were mixed thoroughly at room temperature for about 1 h. Meanwhile, 1% (w/v) of chopped CaP-coated PCL nanofibers were added into the solution. After stirring for 2 h, the mixture was poured into 24-well tissue culture polystyrene plates, frozen at -20 oC overnight and then freeze-dried. The code and composition of the fabricated scaffolds are presented in Table 1. 3. Characterization 3.1. Characterization of electrospun PCL mat The morphologies of electrospun fibers before and after CaP deposition were observed using a Scanning electron microscopy (SEM, Lecia Cambridge S360). The diameter and distribution of the electrospun fibers were measured by Image J software (National institutes of Health, Bethesda, Maryland, USA). Approximately 100 random fibers of different SEM images were analyzed. The presence of calcium and phosphorus elements on the deposited layer was also evaluated by Electron dispersive spectroscopy (EDS). The changes in the chemical structure of electrospun PCL nanofibers after CaP deposition were studied by Fourier transform infrared spectrometer (FTIR; BRUKER IFS 48) in the range of 400–4000 cm-1. Moreover, the phase compositions of electrospun PCL nanofibers were determined by X-ray diffractometry (XRD; X’Pert Pro MPD) using Cukα radiation. Metrohm pH meter 827 determined the changes in pH value during alternate soaking process. 3.2. Characterization of fabricated composite scaffolds 3.2.1. IR spectroscopy The structural analysis of pure starch, oxidized starch, pure chitosan, and composite scaffolds were examined through Fourier transform infrared spectroscopy in the attenuated total reflectance mode (FTIR-ATR). 3.2.2. Microstructure of the composite scaffolds SEM was performed to examine the internal morphology of the fabricated scaffolds. The pore size of each sample, the total area of the pores in each cross section (AP), and the total area of each cross section (AT) were calculated using Image J software. The measurements were based on the five SEM images of each sample (n=5). Next, the porosity percentages of the fabricated scaffolds were calculated using the following equation (Annabi, Fathi, Mithieux, Weiss, & Dehghani, 2011): Eq. (1) 3.2.3. Water uptake properties To determine the amount of water uptake, the pre-weighted scaffolds (Wo) were immersed in deionized water (T=37oC) and allowed to completely swell. After that, the samples were removed; the excess water was wiped off with filter paper and weighted (Ws). All measurements were carried out for three samples (n=3). The amount of water uptake was calculated using the following equation (Nasri-Nasrabadi et al., 2014): Water uptake (g/g) Eq. (2) 3.2.4. Mechanical properties The mechanical properties of the scaffolds (d=6 mm, h=12 mm) were determined using Zwick/Roell Z050, equipped with a 0.1 KN load cell with a crosshead speed of 10
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mm/min. The compressive modulus was calculated by the linear section of the stressstrain curve. Moreover, the compressive strength was measured by drawing a line, starting at 1% strain. The intersection of this line with the stress-strain curve was defined as the compressive strength (Kim, Park, Kim, Wada, & Kaplan, 2005).
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3.3. Cytocompatibility assessment 3.3.1. Cell culture Human osteoblast-like cells, MG63, from National Cell Bank of Iran (NCBI; Pasteur institute) were maintained in Dulbecco’s Modification of Eagles Medium (DMEM; GIBCO, Scotland) supplemented with 10% Fetal Bovine Serum (FBS; Gibco, Renfrewshire, Scotland), 100 U/mL penicillin (Sigma, Saint Louis, USA), and 100 µg/mL streptomycin (Sigma, Saint Louis, USA) at 37 °C in a humidified incubator with 5% CO2. Triplicate specimens (n=3) of each specimen were put into petri dishes, disinfected via interval immersion in ethanol and phosphate buffered saline (PBS) for 5 times, and then washed twice with culture medium before cell seeding. 3.3.2. Cell viability characterization Indirect 3-(4, 5-dimethylthiazol-2-yl)-2, 5-diphenyltetrazolium-bromide (MTT, Sigma, Saint Louis, USA) assay based on extraction method, was used to assess cell viability (Mirahmadi, Tafazzoli-Shadpour, Shokrgozar, & Bonakdar, 2013). Briefly, the 3 and 7day samples’ extractions were added to a 96-well plate containing MG 63 cells for 24 h. Next, the culture medium was removed, replaced by 100 µl of 3 and 7-day extracts and then kept in an incubator. After 24 h, 100 µl MTT solution (0.5 mg/ml) in PBS was added to each cell containing well and incubated for about 4 h at 37 oC. After that, the formed purple formazan crystals were dissolved in 50 µl DMSO (D2650, Sigma, Saint Louis, USA) for 10 min and then absorbance was read at 545 nm using an ELISA Reader (Stat Fax-2100; GMI, Inc., Miami, FL, USA). The serum free culture medium without scaffolds was chosen to be the control group. 3.3.3. Cell morphology observation The morphology of the 5-day cultured MG 63 cells (5x 104 cells/ cm2) was examined using SEM. After 5 days of culture, the cells were fixed using 4% (v/v) glutaraldehyde solution in PBS at 4oC for 30 minutes. Afterwards, the samples were washed thoroughly with deionized water, dehydrated by graded alcohol solution (10% ethanol increments; each step 10 minutes) and finally dried at room temperature. In the end, the dried samples were mounted on an aluminum stub and sputter coated with gold prior to imaging. 3.4. Statistical analysis All data was expressed as mean ± standard deviation (SD). A one-way analysis of variance (ANOVA) was performed to compare any significant difference using StatPlus software. A p< 0.05 was considered statistically meaningful. 4. Results and discussion 4.1. Structure and morphology of electrospun PCL mat Fig. 1a-c shows the SEM image, fiber diameter distribution, and EDS spectrum of pristine electrospun PCL nanofibers. The SEM image consisted of beadless fibers with an average diameter of 149.7 ± 31 nm, presenting a diameter size between 65 nm and 250 nm (Fig.1 a, b). As shown in Fig. 1c, the corresponding EDS spectrum consists of carbon and oxygen. However, new spherical and plate-like morphologies appeared on the surface of electrospun PCL mat after soaking in calcium and phosphate rich solutions
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(Fig. 1d). As shown by EDS analysis, the precipitated layer consists of calcium and phosphate elements (Fig. 1e). Fig. 2 depicts FTIR spectra and XRD patterns of the NaOH-treated PCL nanofibers before and after alternate soaking process. As illustrated in Fig. 2a, the NaOH-treated PCL spectrum has characteristic peaks of 1166 cm-1 (COC symmetric stretching), 1186 cm-1 (OC–O stretching), 1242 cm-1 (COC asymmetric stretching) 1722 cm-1 (C=O stretching), 2871 cm-1 (C-H symmetric stretching), and 2950 cm-1 (C-H asymmetric stretching) (Elzein, Nasser-Eddine, Delaite, Bistac, & Dumas, 2004). However, after the alternate soaking process, new peaks appeared at 550–650 cm-1 (PO43- bending vibration), 876 cm-1 (P-O(H) stretching) and 1000-1100 cm-1 (PO43- asymmetric stretching) (Nourmohammadi, Sadrnezhaad, & Ghader, 2008; Verma, Katti, & Katti, 2006), suggesting that CaP successfully deposited throughout the electrospun PCL mat. The XRD pattern of the resulting NaOH-treated PCL nanofibers (Fig. 2b) exhibited two distinct peaks at about 2θ= 21o and 23o, which relate to the (110) and (200) planes in crystalline PCL (Baji, Wong, Liu, Li, & Srivatsan, 2007). However, after CaP deposition, some changes in the XRD pattern of PCL nanofibers were observed. As indicated from Fig. 2b, the appearance of new peaks at 11.61o, 20.81o, and 29.18o are related to the formation of brushite crystals (Dicalcium phosphate dihydrate; DCPD) on the PCL mat (Mandel, & Tas, 2010 ). Moreover, apatitic CaP is indicated by the appearance of weak peaks at 26o and 32o (Nourmohammadi, Sadrnezhaad, & Ghader, 2008). The XRD results support the SEM and EDS data. The formation of CaP layer throughout the PCL electrospun mat is due to the presence of carboxyl groups. In other words, during NaOH treatment, PCL was partially hydrolyzed (Park et al., 2007), which resulted in creation of carboxylate groups on the surface of PCL mat. These negatively charged carboxylate groups could serve as an inductive site for CaP nucleation (Kawashita et al., 2003; Zheng, Xiong, & Zhang, 2014)]. Therefore, heterogeneous nucleation began through bonding of calcium ions with these negatively charged carboxylate groups after the initial soaking in the calcium rich solution (pH=7.4). Subsequent soaking in the phosphate rich solution resulted in CaP nuclei formation, followed by growth throughout the electrospun mats during soaking time. As indicated by XRD data (Fig. 2b), both brushite and apatitic CaP deposited on the electrospun PCL mats. Moreover, the pH value of CaCl2 solution decreased from 7.4 to 5.95 at the first soaking cycle. This finding is presumably due to the formation of acidic by-product during the hydrolysis reaction of PCL nanofibers, which accelerates after NaOH treatment. Therefore, the reduction in pH should be the reason for brushite precipitation; this is in agreement with previous studies indicating that brushite crystals start to precipitate on the materials in acidic pH ranging from 2 to 6.5 (Mandel, & Tas, 2010). In addition, brushite is a primary phase that can convert to thermodynamically stable apatitic CaP (Mavis, Demirtaş, Gümüşderelioğlu, Gündüz, & Colak, (2009). 4.2. Characterization of fabricated composite scaffolds 4.2.1. IR spectroscopy The IR spectra of pure chitosan and starch as well as oxidized starch and composite scaffolds are shown in Fig. 3. Chitosan and starch present a peak at around 1161 cm−1 due to CO stretching in glycosidic bonds (Silva, 2012; Sevenou, Hill, Farhat, & Mitchell, 2002). In comparison with the starch spectrum, a new peak at 1720 cm-1 appeared after starch oxidation process, which is related to the C=O stretching in aldehyde groups
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(Socrates, 2004). This suggests that aldehyde groups were successfully generated in starch structure without breaking its glycosidic bands. In composite samples, the imine conjugation between aldehyde groups of starch and NH2 of chitosan were observed at 1645 cm-1. In addition, amide II linkage between free carboxyl groups in PCL and NH2 groups in chitosan was also detected at 1532 cm-1. Importantly, the peaks related to the free aldehyde groups at 1720-1740 cm-1 (Socrates, 2004) were not observed in the IR spectra, suggesting that all of the aldehyde groups in starch were covalently cross-linked with chitosan. 4.2.2. Microstructure of the composite scaffolds The SEM images of the fabricated composite scaffolds and their pore size distribution are shown in Fig. 4. In addition, average pore size and the porosity percentage of each scaffold are summarized in Table 2. In order to characterize the presence and distribution of CaP-coated PCL nanofibers in the porous structure, an EDS elemental mapping of Ca and P were performed. As shown in Fig. S1-4 (supplementary files), both Ca and P were uniformly distributed throughout the fabricated scaffolds. This suggests that the CaPcoated PCL nanofibers were dispersed in the hydrogel matrix. In the sample S1, the pore size varied from 34.30 ± 5.33 µm to 148.67 ± 3.43 µm with the average size of 73.40 ± 28.19 µm. However, the pore size distribution became broader as starch content increased (Fig. S5; supplementary files). Furthermore, the average size of the pores rose to 115.32 ± 37.62 µm in S2, 122.068 ± 51.92 µm in S3, and 135.34 ± 63.03 µm in S4 (Table 2). Thus, increasing starch content resulted in larger pore formation, which makes such scaffolds suitable for BTE applications. It has been reported that the optimal mean pore size needed for BTE scaffolds is in the range of 100-135 µm (Murphy, Haugh, & O'Brien, 2010). As shown in Table 2, the porosity percentage also increased with the increase in starch content. It can be speculated that the rise in the pore size and porosity is due to the formation of imine conjugation (shiff base) between the aldehyde in oxidized starch and amino groups of chitosan. It is generally held that a water molecule was displaced during imine formation (Clayden, Greeves, & Warren, 2012). Therefore, elimination of the formed water molecules during freeze-drying may increase both pore size and porosity values. 4.2.3. Water uptake properties The changes in the equilibrium water uptake of the scaffolds containing various amounts of starch are represented in Table 2. The lowest water uptake occurred in the scaffold consisting of 3% starch (S1; 11.35 ± 0.34 g/g), while this amount increased to 11.47 ± 0.63 (g/g) in S2, 12.96 ± 0.97 (g/g) in S3, and 13.84 ± 1.32 (g/g) in S4. These results are probably due to the presence of more hydroxyl groups and larger pore size in samples having more starch. Previous studies also confirmed that the pore size of the scaffold has effect on the water uptake (Hadisi, Nourmohammadi, & Mohammadi, 2015; Mirahmadi, Tafazzoli-Shadpour, Shokrgozar, & Bonakdar, 2013). 4.2.4. Mechanical properties Determining the compressive properties is another key factor in designing bone tissue engineering scaffolds. The compressive modulus and strength of the fabricated hybrid composites are represented in Fig. 5. It can be seen that the highest compressive modulus was observed for S1 (2.64 ± 0.19 MPa). The modulus decreased with increasing starch content, starting at 2.43 ± 0.13 MPa in S2, 1.6 ± 0.25 MPa in S3, and reaching 0.8 ± 0.15 MPa in S4. A similar trend was observed for compressive strength of the scaffolds. The
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compressive strength of sample S1 (1.2 ± 0.05 MPa) decreased to 0.89 ± 0.025 MPa in S2, 0.54 ± 0.09 MPa in S3, and 0.44 ± 0.025 MPa in S4. It should be noted that the compressive properties of materials are highly affected by their solid content (Gibson, & Ashby, 1988). The rise in the pore size and porosity percentage that occurred at higher starch concentration may cause this reduction. This is in agreement with previous reports that increasing pore sizes and porosity led to a decrease in the compressive strength and modulus of scaffolds (Mirahmadi, Tafazzoli-Shadpour, Shokrgozar, & Bonakdar, 2013; Gibson, & Ashby, 1988). 4.2.5. Cytocompatibility assay The results of the cell viability from indirect MTT assay of all samples in comparison with the control group for days 3 and 7 are illustrated in Fig. 6a. As shown in Fig. 6a, all samples have cellular viability more than 90%, which suggests that the fabricated scaffolds are biocompatible and did not show any cytotoxic effects on cells. There is no significant difference between MTT values for different samples after exposed to 3 days extracts. After 7 days, the cell viability increased as the amount of oxidized starch rose (p < 0.05). This can be related to the formation of high-crosslinked structure by increasing oxidized starch amounts, which led to the release of less toxic agents in the culture medium and consequently increased cellular viability. The morphology of MG 63 cells cultured on the scaffolds after 5 days is shown in Fig. 6b. The cells spread on the surface of all scaffolds by their filopodia and they had penetrated inside the pores. It was observed that the cells attached well on the matrix and were either flat or round in morphology. Here, the flat cells attached tightly on the surface by their lamellopodia, and the confluent monolayers of the cells were observed through the pores’ walls. Moreover, the round MG 63 cells attached to the surface by their filopodia. Both SEM images and MTT results suggest that the composites are biocompatible and can be considered as suitable scaffolds for BTE applications. Martins et.al (2008) confirmed that the adhesion, spreading, and growth of both osteoblast and fibroblast cells are enhanced by blending chitosan with starch, which is in agreement with the result of this study. Conclusion This study offers a new bioactive composite scaffold based on chitosan-oxidized starch for potential bone tissue regeneration. At the outset, oxidized starch prepared by periodate oxidation was blended with chitosan through the reductive alkylation process. Meanwhile, CaP-coated electrospun PCL nanofibers were added into the oxidized starchchitosan solution, followed by freeze-drying. The average pore size, porosity, swelling ratio, and viability of osteoblast-like cells (MG 63) increased by adding more starch, while the compressive modulus and strength were reported to decline. In addition, the cells were spread well and anchored, on which the scaffold was covered with a monolayer of cells. References: Park, J. S., Kim, J. M., Jun, L. S., Geun, L. S., Young-Keun, J., Kim, S. E., & Lee, S. C. (2007). Surface Hydrolysis of Fibrous Poly(ε-caprolactone) Scaffolds for Enhanced Osteoblast Adhesion and Proliferation. Macromolecular Research , 15 (5), 424-429. Mavis, B., Demirtaş, T. T., Gümüşderelioğlu, M., Gündüz, G., & Colak, U. (2009). Synthesis, characterization and osteoblastic activity of polycaprolactone nanofibers coated with biomimetic calcium phosphate. Acta biomaterialia , 5 (8), 3098-3111.
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Zheng, Y., Xiong, C., & Zhang, L. (2014). Formation of bone-like apatite on plasmacarboxylated poly(etheretherketone) surface. Materials Letters , 126 (1), 147-150. Wu, Y. B., Yu, S. H., Mi, F. L., Wu, C. W., Shyu, S. S., Peng, C. K., & Chao, A. C. (2004). Preparation and characterization on mechanical and antibacterial properties of chitsoan/cellulose blends. Carbohydrate Polymers , 57 (4), 435-440. Xie, F., Pollet, E., Halley, P. J. , & Avérous, L. (2013). Starch-based nano-biocomposites. Progress in Polymer Science , 38 (10-11), 1590-1628. Verma, D., Katti, K., & Katti, D. (2006). Experimental investigation of interfaces in hydroxyapatite/polyacrylic acid/polycaprolactone composites using photoacoustic FTIR spectroscopy. Journal of Biomedical Materials Research Part A , 77 (1), 59-66. Annabi, N., Fathi, A., Mithieux, S. M. ,Weiss, A. S., & Dehghani, F. (2011). Fabrication of porous PCL/elastin composite scaffolds for tissue engineering applications. The Journal of Supercritical Fluids , 59, 157-167. Baysal, K., Aroguz, A. Z., Adiguzel, Z., & Baysal, B. M. (2013). Chitosan/alginate crosslinked hydrogels: Preparation, characterization and application for cell growth purposes. International Journal of Biological Macromolecules , 59, 342-348. Baji, A., Wong, S. C., Liu, T., Li, T., & Srivatsan, T. S. (2007). Morphological and X-ray diffraction studies of crystalline hydroxyapatite-reinforced polycaprolactone. Journal of Biomedical Materials Research Part B: Applied Biomaterials , 81 (2), 343-350. Barbani, N., Rosellini, E., Cristallini, C., Guerra, G. D., Krajewski, A., & Mazzocchi, M. (2011). Hydroxyapatite-collagen composites. Part I: can the decrease of the interactions between the two components be a physicochemical component of osteoporosis in aged bone? Journal of Materials Science: Materials in Medicine , 22 (3), 637–646. Bourtoom, T., & Chinnan, M. S. (2008). Preparation and properties of rice starchchitosan blend biodegradable film. LWT - Food Science and Technology , 41, 1633-1641. Bose, S., Roy, M., & Bandyopadhyay, A. (2012). Recent advances in bone tissue engineering scaffolds. Trends in Biotechnology , 30 (10), 546-554. Cheng, Y. H., Yang, S. H., Su, W. Y., Chen, Y. C., Yang, K. C., Cheng, W. T. K., Lin, F.H. (2010). Thermosensitive Chitosan–Gelatin–Glycerol Phosphate Hydrogels as a Cell Carrier for Nucleus Pulposus Regeneration: An In Vitro Study. Tissue Engineering Part A , 16 (2), 695-703. Chesnutt, B.M., Viano, A. M., Yuan, Y., Yang, Y., Guda, T., Appleford, M. R., Bumgardner, J. D. (2009). Design and characterization of a novel chitosan/nanocrystalline calcium phosphate composite scaffold for bone regeneration. Journal of Biomedical Materials Research Part A , 88 (2), 491-502. Clayden, J., Greeves, N., & Warren, S. (2012). Organic chemistry (second ed.). New York: Oxford University Press. Croisier, F., & Jérôme, C. (2013). Chitosan-based biomaterials for tissue engineering. European Polymer Journal , 49, 780-792. Elzein, T., Nasser-Eddine, M., Delaite, C., Bistac, S., & Dumas, P. (2004). FTIR study of polycaprolactone chain organization at interfaces. Journal of Colloid and Interface Science , 273, 381-387. Guarino, V., Raucci, M. G., Alvarez-Perez, M. A., Cirillo, V., Ronca, A., & Ambrosio, L. (2013). Scaffold Design for Bone Tissue Engineering: From Micrometric to Nanometric Level. In I. Antoniac (Ed.), Biologically Responsive Biomaterials for Tissue Engineering (pp. 1-16). Springer Series in Biomaterials Science and Engineering.
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Gibson, L. J. A., & Ashby, M. F. (1988). Cellular Solids: Structure and Properties. Oxford: Pergamon Press. Hutmacher, D. W., Schantz, J. T., Lam, C. X., Tan, K. C., & Lim, T. C. (2007). State of the art and future directions of scaffold-based bone engineering from a biomaterials perspective. Journal Of Tissue Engineering And Regenerativ , 4 (1), 245-260. Hadisi, Z., Nourmohammadi, J., & Mohammadi, J. (2015). Composite of porous starchsilk fibroin nanofiber-calcium phosphate for bone regeneration. Ceramics International , 41, 10745-10754. Hanafi, I., Irani, M., & Zulkifli, A. (2013). Starch-Based Hydrogels: Present Status and Applications. International Journal of Polymeric Materials and Polymeric Biomaterials , 62, 411-420. Hermanson, G. T. (1996). Bioconjugate Techniques. Academic Press. Kawashita, M., Nakao, M., Minoda, M., Kim, H. M., Beppu, T., Miyamoto, T., Nakamura, T. (2003). Apatite-forming ability of carboxyl group-containing polymer gels in a simulated body fluid. Biomaterials , 24 (14), 2477-2484. Kai, D., Prabhakaran, M. P., Stahl, B., Eblenkamp, M., Wintermantel, E., & Ramakrishna, S. (2012). Mechanical properties and in vitro behavior of nanofiberhydrogel composites for tissue engineering applications. Nanotechnology , 23 (9). Kim, U. J., Park, J., Kim, H. J., Wada, M., & Kaplan, D. L. (2005). Three-dimensional aqueous-derived biomaterial scaffolds from silk fibroin. Biomaterials , 26, 2775-2785. Lai, G. J., Shalumon, K. T., & Chen, J. P. (2015). Response of human mesenchymal stem cells to intrafibrillar nanohydroxyapatite content and extrafibrillar nanohydroxyapatite in biomimetic chitosan/silk fibroin/nanohydroxyapatite nanofibrous membrane scaffolds. International Journal of Nanomedicine , 10, 567-584. Lee, S. H., & Shin, H. (2007). Matrices and scaffolds for delivery of bioactive molecules in bone and cartilage tissue engineering. Advanced Drug Delivery Reviews , 59, 339-359. Nasri-Nasrabadi, B., Mehrasa, M., Rafienia, M., Bonakdar, S., Behzad, T., & Gavanji, S. (2014). Porous starch/cellulose nanofibers composite prepared by salt leaching technique for tissue engineering. Carbohydrate Polymers , 108, 232-238. Nourmohammadi, J., Sadrnezhaad, S., & Ghader, A. B. (2008). Bone-like apatite layer formation on the new resin-modified glass-ionomer cement. Journal of Materials Science: Materials in Medicine , 19, 3507-3514. Murphy, C. M., Haugh, M. G., & O'Brien, F. J. (2010). The effect of mean pore size on cell attachment, proliferation and migration in collagen–glycosaminoglycan scaffolds for bone tissue engineering. Biomaterials , 31 (3), 461-466. Mandel, S., & Tas, A. C. (2010). Brushite (CaHPO4·2H2O) to octacalcium phosphate (Ca8(HPO4)2(PO4)4·5H2O) transformation in DMEM solutions at 36.5 °C. Materials Science and Engineering: C , 30 (2), 245-254. Martins, A. M., Santos, M. I., Azevedo, H. S., Malafaya, P. B. , & Reis, R. L. (2008). Natural origin scaffolds with in situ pore forming capability for bone tissue engineering applications. Acta Biomaterialia 4 (6), 1637-1645. Mirahmadi, F., Tafazzoli-Shadpour, M., Shokrgozar, M. A., & Bonakdar, S. (2013). Enhanced mechanical properties of thermosensitive chitosan hydrogel by silk fibers for cartilage tissue engineering. Materials Science and Engineering: C 33 (2013) 4786-4794. , C33, 4786-4794.
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Puppi, D., Chiellini, F., Piras, A. M., & Chiellini, E. (2010). Polymeric materials for bone and cartilage repair. Polymer Science , 35 (4), 403–440. Peppas, N. A. ,Hilt, J. Z. , Khademhosseini, A. , & Langer, R. (2006). Hydrogels in biology and medicine: from molecular principles to bionanotechnology. Advanced Materials , 18, 1345-1360. Sevenou, O., Hill, S., Farhat, I., & Mitchell, J. (2002). Organisation of the external region of the starch granule as determined by infrared spectroscopy. International Journal of Biological Macromolecules , 31, 79-85. Silva, S. M. L., Braga, C. R. C., Fook, M. V. L., Raposo, C. M. O., Carvalho, L.H., & Canedo, E. L. (2012). Application of Infrared Spectroscopy to Analysis of Chitosan/Clay Nanocomposites. In T. Theophanides (Ed.), Infrared Spectroscopy - Materials Science, Engineering and Technology (pp. 43-63). InTech. Shakir, M., Jolly, R., Khan, M. S., Iram, N. E., & Khan, H. M. (2015). Nanohydroxyapatite/chitosan–starch nanocomposite as a novel bone construct: Synthesis and in vitro studies. International Journal of Biological Macromolecules , 80, 282-292. Shrivats, A. R., Dermott, M. C., & Hollinger, J. O. (2014). Bone tissue engineering: state of the union. Drug discovery today , 19, 781-786. Socrates, G. (2004). in: Infrared and Raman Characteristic Group Frequencies: Tables andCharts. John Wiley & Sons. Taguchi, T., Kishida, A., & Akashi, M. . (1999). Apatite formation on/in hydrogel matrices using an alternate soaking process: II. Effect of swelling ratios of poly (vinyl alcohol) hydrogel matrices on apatite formation. Journal of Biomaterials Science, Polymer Edition , 10, 331-339.
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Fig.1. (a) SEM image of electrospun PCL fibers (b) fiber diameter distribution (c) EDS spectrum of pristine PCL fibers (d) SEM image of CaP-coated PCL nanofibers, and (e) EDS spectrum of CaP-coated PCL nanofibers. Fig.2. Changes in the (a) IR spectra and (b) XRD patterns of PCL nanofibers before and after CaP deposition. Fig.3. The IR spectra of chitosan, starch, oxidized starch, and composite scaffolds. Fig.4. The SEM images of composite scaffolds (a) S1, (b) S2, (c) S3, and (d) S4. Fig.5. The (a) compressive modulus and (b) compressive strength of each scaffold (*** p < 0.001).
Fig.6. (a) The viability of MG 63 cells after exposed to scaffolds' extract at different days. (b) the morphology of MG 63 cells culture on the scaffolds after 5 days (*p < 0.05) . Table 1 The code and composition of each fabricated composite scaffolds
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Chitosan
Oxidized starch
S1 S2 S3 S4
97 93 90 85
3 7 10 15
CaP-coated PCL electrospun fibers (w/v %) 1 1 1 1
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Samples
Table 2 The average pore size, porosity, and equilibrium water uptake of each scaffold Porosity (%) 54.263 ± 3.27 60.58 ± 4.1 66.97 ± 5.76 71.91 ± 2.32
an
Highlights
Equilibrium water uptake (g/g) 11.35 ± 0.34 11.47 ± 0.63 12.96 ± 0.97 13.84 ± 1.32
cr
S1 S2 S3 S4
Average pore size (µm) 73.40 ± 28.19 115.32 ± 37.62 122.068 ± 51.92 135.34 ± 63.03
us
Sample
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Oxidized starch prepared by periodate oxidation was blended with chitosan through the reductive alkylation process. The PCl nanofibers were prepared by electrospinning using solvent mixture formic acid/acetic acid. The calcium phosphate (CaP) was deposited throughout the electrospun PCl mat using alternate soaking process. The composite scaffolds from CaP-coated PCl nanofibers-chitosan-oxidized starch were prepared via freeze-drying. Adhesion and viability of osteoblast-like cells (MG 63) were investigated on the scaffolds.
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