Journal of Biomaterials Science, Polymer Edition, 2014 Vol. 25, No. 18, 2080–2093, http://dx.doi.org/10.1080/09205063.2014.970604
Effects of preparation methods on the bone formation potential of apatite-coated chitosan microspheres Fei Xua,b, Huifen Dinga, Fangfang Songa and Jiawei Wanga* a
Hubei-MOST KLOS & KLOBM, School and Hospital of Stomatology, Wuhan University, Wuhan 430079, P.R. China; bDepartment of Stomatology, Xiangya Hospital, Central South University, Changsha 410008, P.R. China (Received 30 June 2014; accepted 25 September 2014) To investigate the effects of preparation methods on the bone formation potential of apatite-coated chitosan microspheres, coacervate precipitation method and emulsion cross-linking method were chosen to prepare chitosan microspheres, and then apatite coatings were deposited using simulated body fluid. Rat bone marrow-derived mesenchymal stem cells (BMSCs) were seeded on these microspheres. Cell adhesion, proliferation, and differentiation potential were monitored. For in vivo analysis, some cell/microsphere constructs were implanted in the subcutaneous pockets of male Wistar rats. After 3, 6, 12 weeks, the samples were retrieved and stained with hematoxylin and eosin (HE). Some cell/microsphere constructs were implanted in the calvarial defects of rats. Micro-CT and HE analysis were performed to analyze the new bone formation. It was found that BMSCs on apatite-coated emulsion cross-linked microspheres (EM1) exhibited better proliferation and differentiation than cells on apatite-coated coacervate-precipitated microspheres. The in vivo results showed that no bone was observed in ectopic areas. While in calvarial defects, both histological slices and Micro-CT images demonstrated that a substantial amount of new bone was formed in the EM1/BMSCs construct. These data suggest that preparation methods do exert great influence on the in vitro cell behaviors and in vivo orthotopic bone regeneration of apatite-coated chitosan microspheres. Appropriate method should be considered when preparing chitosan microspheres for bone tissue engineering scaffold. Keywords: chitosan microspheres; preparation methods; BMSCs; bone regeneration
1. Introduction The use of microspheres as scaffold is of great interest for bone tissue engineering, which exhibits several advantages over pre-shaped bulk scaffolds. First of all, the spherical nature of microspheres allow them to fill irregularly shaped bone defects through minimally invasive procedures, thus avoiding surgical trauma.[1] Second, microspheres are considered as useful cell delivery vehicles, and the cell/microsphere constructs can be implanted directly in vivo which is able to simplify the procedures and improve cell viability.[2] Third, it is meaningful to conduct surface modifications for microspheres to increase cell–substrate interaction, since they own large specific surface area.[3]
*Corresponding author. Email: [email protected] © 2014 Taylor & Francis
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Chitosan, a linear polysaccharide composed of glucosamine and N-acetyl glucosamine, is usually obtained by partial deacetylation of chitin.[4] Owing to its biocompatibility, enzyme-regulated degradation potential, and similarity to extracellular matrix, chitosan is suitable for constructing microspheres.[5] Various preparation methods have been proposed.[6] Among those, the coacervate precipitation method and the emulsion cross-linking method are mostly chosen. However, with regard to the complexity of bone tissue, a pure chitosan scaffold lacks bioactivity to induce bone formation. To enhance the osteogenic potential of polymers, two methods are proposed. One method is to mix polymer with hydroxyapatite so as to form a polymer-apatite slurry. Nevertheless, the mechanical blend of polymer with hydroxyapatite generally lacks strong mineral/polymer integration and interaction,[7] and the hydroxyapatite particles are generally embedded deeply inside the polymer matrix and distributed unevenly. Another way is to prepare apatite coating on the surface of polymer.[8–10] These apatite-coated scaffolds have some unique advantages. For instance, since apatite coating directly contacts with the tissue, better attachment, proliferation, and differentiation of cells are expected.[11] Additionally, the apatite coating is deposited in simulated body fluid (SBF) without using special equipment or extremely high processing temperature, which mimics the biomimetic process.[12] In our previous paper, apatite-coated chitosan microspheres fabricated by different methods have been evaluated for their physicochemical and in vitro biological properties.[13] However, whether these microspheres can promote in vivo bone formation is unknown. To further learn the effect of preparation methods, in this paper, bone formation potential of apatite-coated chitosan microspheres is evaluated through ectopic and cranial bone defect models. 2. Materials and methods 2.1. Preparation of chitosan microspheres Chitosan microspheres were fabricated according to our previous study,[13] but with some modifications. Briefly, chitosan powder (85% deacetylation degree, Aldrich) was dissolved in 2% (v/v) acetic acid to give a 3% (w/v) chitosan solution. Genipin powder (Wako Chemicals) was dissolved in 60% (v/v) ethanol. The ratio of genipin/chitosan was kept at 10% (w/w). The microspheres obtained from coacervate precipitation method were made by dropping chitosan solution via a syringe into 2% (m/v) sodium hydroxide solution in 80% ethanol (v/v). After stirring for 1 h, the microspheres were rinsed with deionized water and were cross-linked in genipin solution for 24 h, followed by rinsing and drying (CM0). Microspheres obtained from emulsion crosslinking method were fabricated with a water/oil technique. Briefly, 25 mL chitosan solution was slowly added into 100 mL paraffin oil containing 1 mL span 80. Under constant stirring, genipin solution was added. 24 h later, the cross-linked microspheres were collected and rinsed (EM0). Apatite coating was prepared in a two-step procedure as previously described.[14] Briefly, both microspheres were soaked into five times SBF solution for two days. The resulting microspheres were then rinsed gently with deionized water and dried. They were abbreviated as apatite-coated coacervate-precipitated microspheres (CM1) and emulsion cross-linked microspheres (EM1), respectively.
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2.2. Physicochemical properties of chitosan microspheres Since the preparation process was modified, some physicochemical properties of microspheres were assessed again.[13] To determine the cross-linking degree, ninhydrin assay was performed.[15] Briefly, the test samples were heated within the ninhydrin solution for 20 min at 100 °C. Then the absorbance of the solution at 570 nm was recorded by a spectrophotometer (BIO-TEK ELX808, USA) using known concentrations of glycine as standard. The degree of cross-linking was defined as the ratio of the consumed amino groups in the cross-linked samples to the free amino groups in the corresponding uncross-linked samples.[16] The morphology of microspheres was examined by highresolution field scanning electron microscopy (SEM, FEI Sirion 200, USA). The chemical composition of the coating was characterized by X-ray diffractometer (XRD, Bruker D8 Advance, Germany) and Fourier transform infrared-attenuated total reflectance spectroscopy (FTIR-ATR, Nicolet 5700, USA). 2.3. In vitro biocompatibility using rat bone marrow-derived mesenchymal stem cells as the seeding cells 2.3.1. Cell culture Rat bone marrow-derived mesenchymal stem cells (BMSCs) were isolated and cultured according to the protocol reported previously.[17] Cells at passage 2–4 were used. Before the cell seeding, microspheres were sterilized by ultraviolet light and incubated with culture medium for 24 h. 2.3.2. Cell adhesion To evaluate cell adhesion and morphology, BMSCs (3.2 × 104 cells/well) were seeded on 10 mg microspheres in 96 culture plate. After culturing for 1 or 3 days, the cell/ microsphere constructs were observed by SEM. 2.3.3. Cell proliferation To assess cell proliferation, BMSCs were seeded on microspheres, in the same manner as the adhesion test. Twelve hours later, the cell/microsphere constructs were carefully transferred into a new plate. After 1, 3, 5, and 7 days, CCK-8 solution (Dojindo Laboratory, Kumamoto, Japan) was added to each well. Three hours later, the optical density of supernatant solution was measured at a wavelength of 450 nm (n = 6). 2.3.4. Cell differentiation To measure alkaline phosphatase (ALP) activity and the gene expression of osteogenic markers, cells (1 × 106 cells/well) were seeded on 40 mg microspheres in 24 culture plates. Twelve hours later, the cell/microsphere constructs were transferred into a new plate and were cultured in osteogenic medium, including 10 mM β-glycerophosphate, 50 μg/mL ascorbic acid, and 10−8 M dexamethasone. On day 7 and day 14, the samples were rinsed with PBS, lysed in 0.2% Triton X-100, followed by sonication and centrifugation. The ALP activity and total amount of protein were determined by p-nitrophenyl phosphate method (n = 6 for each time point).[18] On day 14, mRNA expression levels of ALP, collagen type I (COL I), osteocalcin (OCN), and glyceraldehyde 3-phosphate
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Primer sequences for RT-PCR.
Gene
Forward primer (5′-3′)
Reverse primer (5′-3′)
Alkaline phosphatase (ALP) Collagen type I (COL I) Osteocalcin (OCN) GADPH
CAATTAACGGCTGACACTGC TCTGCGACACAAGGAGTCTG CAAGTCCCACACAGCAACTC AACGACCCCTTCATTGAC
TTTCAGGGCATTTTTCAAGG GGGACCATCAACACCATCTC GGCTCCAAGTCCATTGTTGA TCCACGACATACTCAGCAC
dehydrogenase were also detected (Table 1). Total RNA was extracted using Trizol (Sigma–Aldrich, St. Louis, MO, USA). RNA (1 μg) was reverse transcribed using First Strand cDNA Synthesis Kit (Fermentas, Maryland, USA). Then polymerase chain reaction (PCR) was performed using 2 × PCR mix (Tiangen Biotech, Beijing, China). Finally, the PCR products were electrophoresed on a 2% agarose gel and were analyzed using Image J software. 2.4 In vivo transplantation 2.4.1. Ethics statement and preparation of BMSCs/microsphere constructs All animal experiments were approved by the Ethics Committee of School and Hospital of Stomatology, Wuhan University. Male Wistar rats with average weight at 200 g ± 15 g were obtained from the Experimental Animal Centre of Hubei Province. To prepare BMSCs/microsphere constructs, BMSCs (3.2 × 105 cells/well) were seeded on 15 mg microspheres in 96 culture plate and cultured in osteogenic medium for 1 week before transplantation. Moreover, in order to fix the microspheres transplanted in cranial defects, chitosan films were prepared. Briefly, 3% chitosan solution was cast on culture dishes and dried at 50 °C. Then, the formed thin films were neutralized with NaOH (0.1 M), washed with deionized water, and sterilized by ultraviolet light before transplantation. 2.4.2. Ectopic bone regeneration To assess ectopic bone regeneration, four dorsal subcutaneous pockets were created for each rat. The cell/microsphere constructs were randomly transplanted and the skin was carefully sutured. After 3, 6, and 12 weeks, three rats were sacrificed at each time point. The transplanted samples were removed, fixed, decalcified, serials dehydrated, and embedded in paraffin (n = 6). Serial sections (5 μm thick) were stained with hematoxylin and eosin (HE). 2.4.3. Orthotopic bone regeneration For cranial defect transplantation, a sagittal incision of approximately 15 mm was made in the dorsal part of the rat cranium. The subcutaneous tissue, muscle, and periosteum were dissected. For each rat, two full-thickness bone defects (5 mm in diameter) were created using a trephine bur under constant irrigation with sterile saline. Thirty cranial defects were randomly repaired as follows: (1) CM0/BMSCs group, the defects were filled with CM0 seeded with BMSCs (n = 6); (2) EM0/BMSCs group, the defects were filled with EM0 seeded with BMSCs (n = 6); (3) CM1/BMSCs group, the defects
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were filled with CM1 seeded with BMSCs (n = 6); (4) EM1/BMSCs group, the defects were filled with EM1 seeded with BMSCs (n = 6); (5) Black group, the defects were filled with nothing (n = 6). After that, the chitosan films were covered on the defects and the wound was closed in layers. Eight weeks later, all rats were sacrificed. The defect site and surrounding bone were carefully dissected. All specimens were placed in a custom-made holder with 4% paraformaldehyde and scanned by a Scanco μCT50 imaging system (Scanco Medical, Bassersdorf, Switzerland). The scanning was performed at 70 kVp and 112 mA with a 10 μm pixel size, 1024 reconstruction matrix, and 250 ms integration time. A Gaussian filter (sigma = 0.8 and support = 1) and a fixed threshold (value = 220) were used to properly segment the scaffolds and the mineralized bone from the background. After three-dimensional (3-D) reconstruction, the relative maturity of bone was analyzed using a protocol provided by the manufacturer of the Micro-CT scanner and distinguished by colors. The red color meant that the bone was relatively mature, while the green color meant that the bone was relatively immature. Since it was difficult to distinguish new bone from apatite coating through separating the μCT signal using global thresholds, a quantitative analysis of bone volume could not be obtained. After the μCT scanning, all samples were decalcified, dehydrated, embedded in paraffin, and sectioned into 5 μm thick sections for HE analysis. Histometric measurements were analyzed by Image-Pro-Plus 5.0 (Media Cybernetic, USA), according to the previous study.[19] New bone area ratio was calculated as the new bone area in the defect divided by the entire defect area. Defect closure was identified by the new bone ingrowth (mm) in the defect divided by the original defect width (mm). 2.5. Statistical analysis All data were presented as the means ± standard deviation. Statistical significance was assessed using one-way ANOVA with the Tukey Post Hoc test. The value of p < 0.05 was considered as statistically significant. 3. Results 3.1. Physicochemical properties of chitosan microspheres The cross-link degree of both microspheres is shown in Figure 1. EM0 had significant lower value than CM0 (40% vs. 80%, p < 0.05). The surface morphology is shown in Figure 2. EM0 demonstrated smooth surface (Figure 2(B) and (D)), whereas CM0 presented rough surfaces (Figure 2(A) and (C)). After being immersed in five times SBF,
Figure 1.
Cross-linking degree of microspheres. *p < 0.05.
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Figure 2. SEM, XRD, and FTIR spectra of chitosan microspheres. (A, C) Coacervate-precipitated microspheres before coating (CM0); (B, D) emulsion cross-linked microspheres before coating (EM0); (E, G, I) coacervate-precipitated microspheres after coating (CM1); (F, H, J) emulsion cross-linked microspheres after coating (EM1); (K) XRD patterns; (L) FTIR spectra. CaP: carbonate apatite. ■: Characteristic peak of carbonate apatite; *: Characteristic peak of chitosan.
the surface of EM1 was uniformly covered by a continuous coating (Figure 2(F) and (H)), whereas the coating on CM1 was not continuous, and the substrate of CM1 could be seen in some places (Figure 2(E) and (G)). Under higher magnification, the minerals formed on EM1 (Figure 2(J)) were denser than the minerals formed on CM1 (Figure 2(I)). The composition of the coatings was characterized by XRD and FTIR. CM1 showed both characteristic peaks of chitosan (2θ = 20°) and characteristic peaks of carbonate apatite (2θ = 25.8°, 28.8°, 31.8°, and 46.7°). These peaks corresponded to the (0 0 2), (2 1 0), (2 1 1), and (2 2 2) planes of apatite, respectively.[20], while the XRD patterns of EM1 only showed the characteristic peaks of apatite (Figure 2(K)). The FTIR spectra of CM0 and EM0 exhibited characteristic bands of peaks around 2900 cm−1 corresponding to –CH2, while 1654 cm−1 was the characteristic peak of amide I. The sharp peaks at 1153, 1070, and 1031 cm−1 were assigned to the C–O stretching vibrations. The peak at 895 cm−1 was assigned to the C–N bond. After coating, new spectra of apatite at 563, 601, 1021, 875, and 1413 cm−1 appeared in CM1 and EM1, in which 563, 601, and 1021 cm−1 were assigned to PO4 bond, while 875 and 1413 cm−1 were assigned to CO3 band.[8,21–24] It could be further observed that the bands of chitosan at 1654 and 895 cm−1 with weak intensity appeared in CM1 but not EM1 (Figure 2(L)). Based on the data of SEM, XRD, and FTIR, we concluded that the surface of EM1 was completely coated by an apatite layer, while the surface of CM1 was partly or thinly coated with an apatite layer.
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3.2. In vitro biocompatibility To assess the cell attachment, we cultured BMSCs on the microspheres for 1 day and 3 days, and evaluated the cell morphology with SEM. On day 1, cells were spherical on CM0 (Figure 3(A)) and EM0 (Figure 3(B)). In contrast, cells were flat and star-like with numerous pseudopodia spread around the cell body on CM1 (Figure 3(C)) and EM1 (Figure 3(D)). On day 3, although the cell number increased on CM0 (Figure 3(E)) and EM0 (Figure 3(F)), it was obviously lower than the cell population on CM1 (Figure 3(G)) and EM1 (Figure 3(H)). Also, the cells on EM1 had grown to sheet-like form and covered the whole surface area (Figure 3(H)). CCK-8 assay showed that after 1 day, there were more cells on CM1 and EM1 than the cells on CM0 and EM0, while there was no significant difference between CM1 and EM1. Five days later, the amount of cells on EM1 was higher than on CM1 (Figure 3(I), p < 0.05). We also found that cells on EM1 had higher ALP activity than cells on CM1 on both day 7 and day 14 (Figure 3(J), p < 0.05). To further monitor the cell differentiation status, the expression of osteogenetic genes was analyzed (Figure 3(K) and (L)). The ALP, COL I, and OCN expression levels were significantly increased in cells cultured on EM1 compared with cells cultured on CM1 (p < 0.05). 3.3. In vivo transplantation 3.3.1. Ectopic transplantation As shown in Figure 4, after 3 and 6 weeks, microspheres in both EM1/BMSCs group and CM1/BMSCs group were capsulized by fibrous tissue without evidence of acute
Figure 3. BMSCs adherence, proliferation, and osteogenic differentiation. (A, E) SEM images of BMSCs grown on CM0; (B, F) SEM images of BMSCs grown on EM0; (C, G) SEM images of BMSCs grown on CM1; (D, H) SEM images of BMSCs grown on EM1; (I) CCK-8 analysis; (J) Quantitative ALP activity; (K) Electrophoresis of PCR product for ALP, COL I, OCN, and GADPH; (K) The ratio of ALP, COL I, and OCN to GADPH mRNA expression. *p < 0.05.
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Figure 4. HE staining of sections from ectopic samples. The arrows indicate the microspheres which have undergone degradation. Bar = 200 μm.
inflammation and bone formation. The EM1/BMSCs group began to degrade with irregular border line after 6 weeks, and after 12 weeks, significant degradation was found. In contrast, the morphology of CM1 did not change much. There was still no bone formation in both groups after 12 weeks. 3.3.2. Cranial transplantation The 3D μCT images of the bone defect were shown in Figure 5. EM1/BMSCs group had the greatest potential to promote bone formation among the groups. As we found that mineralized tissue almost filled the whole defect region (Figure 5(E)).
Figure 5. Micro-CT evaluation of the repaired calvarial bone defects at eight weeks after implantation. (A, F) Blank group; (B, G) CM0/BMSCs group; (C, H) EM0/BMSCs group; (D, I) CM1/BMSCs group; (E, J) EM1/BMSCs group; (A–E) 3-D reconstructed images; (F–J) Analysis of the relative bone maturity. The yellow circle indicates the dimension of 5 mm bone defect. Bar = 1 mm. (Please see the online article for the colour version of this figure: http://dx.doi.org/ 10.1080/09205063.2014.970604.)
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Furthermore, the maturity of the new bone was similar to that of the native bone around the defect, both of which was indicated by red color (Figure 5(J)). So the formed new bone was considered to be relatively mature. CM1/BMSCs group (Figure 5(D)) had less potential to promote bone formation than EM1/BMSCs group (Figure 5(E)), but it had greater potential than the control groups, including Black group (Figure 5(A)), CM0/BMSCs group (Figure 5(B)), and EM0/BMSCs group (Figure 5(C)). Part of the formed new bone in the CM1/BMSCs group was indicated by green color which meant that the new bone was relatively immature (Figure 5(I)). Histological data further supported the Micro-CT finding. No obvious new bone was found in the Blank group (Figure 6(A)) and EM0/BMSCs group (Figure 6(B)). A small amount of bone was observed in the CM0/BMSCs group (Figure 6(C)), whereas obvious new bone grew along the edges of the defects in the CM1/BMSCs group (Figure 6(D)). In the EM1/BMSCs group, the formed new bone was thick and almost covered the whole defect (Figure 6(E)). At higher magnification, active new bone formation could be observed in the CM1/BMSCs group (Figure 6(I)) and EM1/BMSCs group (Figure 6(J)), as osteocytes and blood vessels were found in the inside and osteoblasts were lined at the outside of new bone, whereas in the control groups, only isolated bone islands (Figure 6(H)) or loose connective tissues (Figure 6(G) and (F)) were found. Histomorphometric analysis is shown in Figure 6(K) and (L). The percentages of defect closure and new bone area ratio in the EM1/BMSCs group were significantly higher than those in the CM1/BMSCs group (p < 0.05). 4. Discussion Chitosan microspheres have been widely used in tissue engineering to provide a structural template for cell seeding and extracellular matrix formation. According to our previous study, preparation methods could cause great effect on microspheres’ properties, which further influenced the growth and differentiation of MC3T3-E1 cells in vitro. However, whether preparation methods could affect the regeneration of bone in vivo was still unknown. For this reason, BMSCs, one of the mostly used seeding cells in bone tissue engineering, were thus investigated in place of MC3T3-E1 cell in this study. At the same time, some procedures were modified to optimize the microsphere preparation. For instance, previously, after dropping the chitosan solution into the alkali solution, the stabilized microspheres were firstly dried and then cross-linked. During this process, the drying before cross-linking was time-taking and did affect the morphology of microspheres. So, in this study, the microspheres were directly cross-linked in the solution without first drying. The process was thus simplified and the microspheres were really more spherical than before (data not shown). However, unexpectedly, the cross-linking degree of CM0 increased much, which reached 80% and was significantly higher than that of EM0’s. We speculated that the highly hydrated microspheres in this study might be more vulnerable to the cross-link reaction than the dried microspheres in our previous study. Oppositely, SEM observation, XRD patterns, and FTIR spectra demonstrated that the apatite coating was more favorable to form on EM1 than on CM1. Studies have shown that amino groups could act as the nucleation site during the coating process.[10,25] Therefore, the calcium-binding capacity of chitosan was strictly controlled by the number of free amino groups.[25] Since EM0 had lower cross-linking degree, they naturally possessed more free amino groups on the surface. As a result, apatite could be formed more efficiently on EM1 than on CM1. Toshiki Miyazaki et al. also
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Figure 6. Histological analysis of bone formation in calvarial bone defects stained with HE. (A, F) Blank group; (B, G) EM0/BMSCs; (C, H) CM0/BMSCs; (D, I) CM1/BMSCs; (E, J) EM1/ BMSCs; (K) histomorphometirc analysis of the defect closure; (L) histomorphometirc analysis of the new bone area ratio, (A–E) Bar = 400 μm; (F–J) Bar = 50 μm. The thick black arrow: newly formed bone; the thin black arrow: new vessels; the thin white arrow: osteoblasts; the thin blue arrow: osteocytes. *p < 0.05. (Please see the online article for the colour version of this figure: http://dx.doi.org/10.1080/09205063.2014.970604.)
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found that PGA hydrogels with low cross-linking degree had high apatite-forming ability.[26] However, in our previous study, apatite coating was more favorable to form on CM1 than on EM1.[13] The reason might be that, the difference of cross-linking degree between CM0 and EM0 in the previous study was much smaller than that in the present study (10% vs 40%), so the effect of cross-linking degree on the apatite formation was minimized. Furthermore, the surface of CM0 in the previous study was irregular and shriveled (data not shown), which might favor the physical deposition of apatite. Turning to the biocompatibility, the CCK-8 assay was chosen to evaluate the cell proliferation, since it could monitor the change of cell numbers dynamically. We found that rat BMSCs presented higher proliferation and ALP activity on EM1 than those on CM1. Osteogenic gene expression of BMSCs on the microspheres was also assessed. ALP is an early marker and one of the most frequently used indicators to demonstrate osteogenic differentiation.[27] COL I constitutes around 90% of the organic matrix, which plays an important role in the formation of inorganic matrix of bone.[28] OCN is a later marker of differentiation marker, which is exclusively secreted by mature osteoblasts and present in mineralized matrix.[29] We found that the gene expressions of ALP, COL 1, and OCN were increased on EM1 in contrast to CM1. The reason might be that EM1 had better apatite coatings and apatite could promote the osteogenic differentiation. We further evaluated the in vivo bone regeneration of microspheres using ectopic and orthotopic models. Firstly, CM1/BMSCs and EM1/BMSCs were implanted subcutaneously in rats. After 3, 6, and 12 weeks, it was found that there was no obvious bone formation. Ectopic bone formation model was quite different from the complex biological environment of bone.[30,31] Some researchers reported that apatite-containing biomaterials could successfully promote ectopic bone formation, while others held a totally different view.[32,33] The reason might be that many factors could affect the in vivo behaviors of apatite-containing biomaterials, such as the physicochemical and structural properties of the apatite crystals and the animal models. It was also found that the degradation of EM1 was faster than that of CM1, which might stem from the lower cross-linking degree of EM1. Additional, both groups demonstrated no obvious inflammation indicating that both CM1 and EM1 were safe to the peri-tissue. Secondly, microspheres were implanted in the bone defects. Different from subcutaneous implantation, bone regeneration in cranial defects was favorable. We found that EM1/BMSCs group demonstrated better performance than CM1/BMSCs group. The reasons were as follows: (1) Apatite significantly enhanced the bone repair of chitosan microspheres. It has been reported that ion dissolution products containing Ca and P ions that were released from apatite could stimulate cell attachment, proliferation, and differentiation. In addition, Ca and P ions could activate the bone morphogenetic protein-signaling pathway, which played an important role in an autocrine/paracrine osteoinduction loop and enhanced bone formation in defects.[34] It has also been demonstrated that an apatite layer could preferentially adsorb a high quantity of protein from the serum such as fibronectin, which were then utilized by the osteoblasts to form a mineralized extracellular matrix.[35] (2) The difference in the amount of apatite coatings might affect the cell behaviors and bone formation of scaffolds. In this study, we found that EM1 had the higher degree of apatite formation, since the surface of EM1 was completely coated, while the surface of CM1 was partly or thinly coated by an apatite layer. It has been reported that high apatite content was more effective in inducing new bone formation than low apatite content scaffold.[36] He and coworkers
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further clarified that hydroxyapatite could act as nucleation sites for mineralization and promote further deposition of apatite crystals from cells, thus enhancing the formation of a mineral structure.[37] Several methods, including mass change method, scratch method, and alizarin red-based assay have been tried out to quantify the mineral contents. However, we were not successful since the microspheres were small, spherical, and the content of mineral was low. An effective and accurate method to quantify the amount of apatite coating might be taken in the further research. 5. Conclusions Findings from the present study demonstrate that preparation methods do exert great influence on the in vitro cell behaviors and in vivo orthotopic bone regeneration of apatite-coated chitosan microspheres. Appropriate method should be considered when preparing chitosan microspheres for bone tissue engineering scaffold. Funding This work was supported by the National Natural Science Foundation of China [grant number 81200812], [grant number 81271179]; the Provincial Natural Science Foundation of Hubei [grant number 2011CDB470].
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[13] Shen S, Fu D, Xu F, Long T, Hong F, Wang J. The design and features of apatite-coated chitosan microspheres as injectable scaffold for bone tissue engineering. Biomed. Mater. 2013;8:025007. [14] Chou YF, Chiou WA, Xu Y, Dunn JCY, Wu BM. The effect of pH on the structural evolution of accelerated biomimetic apatite. Biomaterials. 2004;25:5323–5331. [15] Aldana AA, González A, Strumia MC, Martinelli M. Preparation and characterization of chitosan/genipin/poly(N-vinyl-2-pyrrolidone) films for controlled release drugs. Mater. Chem. Phys. 2012;134:317–324. [16] Yan LP, Wang YJ, Ren L, Wu G, Caridade SG, Fan JB, Wang LY, Ji PH, Oliveira JM, Oliveira JT. Genipin-cross-linked collagen/chitosan biomimetic scaffolds for articular cartilage tissue engineering applications. J. Biomed. Mater. Res. A. 2010;95A:465–475. [17] Xu L, Lv K, Zhang W, Zhang X, Jiang X, Zhang F. The healing of critical-size calvarial bone defects in rat with rhPDGF-BB, BMSCs, and β-TCP scaffolds. J. Mater. Sci. Mater. Med. 2012;23:1073–1084. [18] Tan F, Xu X, Deng T, Yin M, Zhang X, Wang J. Fabrication of positively charged poly(ethylene glycol)-diacrylate hydrogel as a bone tissue engineering scaffold. Biomed. Mater. 2012;7:055009. [19] Sohn JY, Park JC, Um YJ, Jung UW, Kim CS, Cho KS, Choi SH. Spontaneous healing capacity of rabbit cranial defects of various sizes. J. Periodontal Implant Sci. 2010;40:180– 187. [20] Chang C, Peng N, He M, Teramoto Y, Nishio Y, Zhang L. Fabrication and properties of chitin/hydroxyapatite hybrid hydrogels as scaffold nano-materials. Carbohydr. Polym. 2013;91:7–13. [21] Chang C, Peng N, He M, Teramoto Y, Nishio Y, Zhang L. Fabrication and properties of chitin/hydroxyapatite hybrid hydrogels as scaffold nano-materials. Carbohydr. Polym. 2013;91:7–13. [22] Thein-Han W, Misra R. Biomimetic chitosan–nanohydroxyapatite composite scaffolds for bone tissue engineering. Acta Biomater. 2009;5:1182–1197. [23] Manjubala I, Scheler S, Bössert J, Jandt KD. Mineralisation of chitosan scaffolds with nanoapatite formation by double diffusion technique. Acta Biomater. 2006;2:75–84. [24] Bagheri-Khoulenjani S, Mirzadeh H, Etrati-Khosroshahi M, Ali Shokrgozar M. Particle size modeling and morphology study of chitosan/gelatin/nanohydroxyapatite nanocomposite microspheres for bone tissue engineering. J. Biomed. Mater. Res. A. 2013;101A:1758–1767. [25] Li B, Wang Y, Jia D, Zhou Y. Gradient structural bone-like apatite induced by chitosan hydrogel via ion assembly. J. Biomater. Sci., Polym. Ed. 2011;22:505–517. [26] Miyazaki T, Mukai J, Ishida E, Ohtsuki C. Apatite mineralization behavior on polyglutamic acid hydrogels in aqueous condition: effects of molecular weight. Biomed. Mater. Eng. 2013;23:339–347. [27] Rodrigues MT, Lee B-K, Lee SJ, Gomes ME, Reis RL, Atala A, Yoo JJ. The effect of differentiation stage of amniotic fluid stem cells on bone regeneration. Biomaterials. 2012;33:6069–6078. [28] Mathews S, Gupta PK, Bhonde R, Totey S. Chitosan enhances mineralization during osteoblast differentiation of human bone marrow-derived mesenchymal stem cells, by upregulating the associated genes. Cell Prolif. 2011;44:537–549. [29] Sato M, Yasui N, Nakase T, Kawahata H, Sugimoto M, Hirota S, Kitamura Y, Nomura S, Ochi T. Expression of bone matrix proteins mRNA during distraction osteogenesis. J. Bone Miner. Res. 1998;13:1221–1231. [30] Sadr N, Pippenger BE, Scherberich A, Wendt D, Mantero S, Martin I, Papadimitropoulos A. Enhancing the biological performance of synthetic polymeric materials by decoration with engineered, decellularized extracellular matrix. Biomaterials. 2012;33:5085–5093. [31] Scott MA, Levi B, Askarinam A, Nguyen A, Rackohn T, Ting K, Soo C, James AW. Brief review of models of ectopic bone formation. Stem Cells Dev. 2012;21:655–667. [32] Levi B, Nelson ER, Li S, James AW, Hyun JS, Montoro DT, Lee M, Glotzbach JP, Commons GW, Longaker MT. Dura mater stimulates human adipose-derived stromal cells to undergo bone formation in mouse calvarial defects. Stem Cells. 2011;29:1241–1255. [33] Dragoo JL, Lieberman JR, Lee RS, Deugarte DA, Lee Y, Zuk PA, Hedrick MH, Benhaim P. Tissue-engineered bone from BMP-2 transduced stem cells derived from human fat. Plast. Reconstr. Surg. 2005;115:1665–1673.
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[34] Chai YC, Roberts SJ, Schrooten J, Luyten FP. Probing the osteoinductive effect of calcium phosphate by using an in vitro biomimetic model. Tissue Eng. Part A. 2011;17:1083–1097. [35] Nishio K, Neo M, Akiyama H, Nishiguchi S, Kim HM, Kokubo T, Nakamura T. The effect of alkali- and heat-treated titanium and apatite-formed titanium on osteoblastic differentiation of bone marrow cells. J. Biomed. Mater. Res. 2000;52:652–661. [36] Xia Z, Yu X, Jiang X, Brody HD, Rowe DW, Wei M. Fabrication and characterization of biomimetic collagen-apatite scaffolds with tunable structures for bone tissue engineering. Acta Biomater. 2013;9:7308–7319. [37] He P, Sahoo S, Ng KS, Chen K, Toh SL, Goh JC. Enhanced osteoinductivity and osteoconductivity through hydroxyapatite coating of silk-based tissue-engineered ligament scaffold. J. Biomed. Mater. Res. A. 2013;101A:555–566.
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Effects of preparation methods on the bone formation potential of apatite-coated chitosan microspheres Fei Xua,b, Huifen Dinga, Fangfang Songa and Jiawei Wanga* a
Hubei-MOST KLOS & KLOBM, School and Hospital of Stomatology, Wuhan University, Wuhan 430079, P.R. China; bDepartment of Stomatology, Xiangya Hospital, Central South University, Changsha 410008, P.R. China (Received 30 June 2014; accepted 25 September 2014) To investigate the effects of preparation methods on the bone formation potential of apatite-coated chitosan microspheres, coacervate precipitation method and emulsion cross-linking method were chosen to prepare chitosan microspheres, and then apatite coatings were deposited using simulated body fluid. Rat bone marrow-derived mesenchymal stem cells (BMSCs) were seeded on these microspheres. Cell adhesion, proliferation, and differentiation potential were monitored. For in vivo analysis, some cell/microsphere constructs were implanted in the subcutaneous pockets of male Wistar rats. After 3, 6, 12 weeks, the samples were retrieved and stained with hematoxylin and eosin (HE). Some cell/microsphere constructs were implanted in the calvarial defects of rats. Micro-CT and HE analysis were performed to analyze the new bone formation. It was found that BMSCs on apatite-coated emulsion cross-linked microspheres (EM1) exhibited better proliferation and differentiation than cells on apatite-coated coacervate-precipitated microspheres. The in vivo results showed that no bone was observed in ectopic areas. While in calvarial defects, both histological slices and Micro-CT images demonstrated that a substantial amount of new bone was formed in the EM1/BMSCs construct. These data suggest that preparation methods do exert great influence on the in vitro cell behaviors and in vivo orthotopic bone regeneration of apatite-coated chitosan microspheres. Appropriate method should be considered when preparing chitosan microspheres for bone tissue engineering scaffold. Keywords: chitosan microspheres; preparation methods; BMSCs; bone regeneration
1. Introduction The use of microspheres as scaffold is of great interest for bone tissue engineering, which exhibits several advantages over pre-shaped bulk scaffolds. First of all, the spherical nature of microspheres allow them to fill irregularly shaped bone defects through minimally invasive procedures, thus avoiding surgical trauma.[1] Second, microspheres are considered as useful cell delivery vehicles, and the cell/microsphere constructs can be implanted directly in vivo which is able to simplify the procedures and improve cell viability.[2] Third, it is meaningful to conduct surface modifications for microspheres to increase cell–substrate interaction, since they own large specific surface area.[3]
*Corresponding author. Email: [email protected] © 2014 Taylor & Francis
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Chitosan, a linear polysaccharide composed of glucosamine and N-acetyl glucosamine, is usually obtained by partial deacetylation of chitin.[4] Owing to its biocompatibility, enzyme-regulated degradation potential, and similarity to extracellular matrix, chitosan is suitable for constructing microspheres.[5] Various preparation methods have been proposed.[6] Among those, the coacervate precipitation method and the emulsion cross-linking method are mostly chosen. However, with regard to the complexity of bone tissue, a pure chitosan scaffold lacks bioactivity to induce bone formation. To enhance the osteogenic potential of polymers, two methods are proposed. One method is to mix polymer with hydroxyapatite so as to form a polymer-apatite slurry. Nevertheless, the mechanical blend of polymer with hydroxyapatite generally lacks strong mineral/polymer integration and interaction,[7] and the hydroxyapatite particles are generally embedded deeply inside the polymer matrix and distributed unevenly. Another way is to prepare apatite coating on the surface of polymer.[8–10] These apatite-coated scaffolds have some unique advantages. For instance, since apatite coating directly contacts with the tissue, better attachment, proliferation, and differentiation of cells are expected.[11] Additionally, the apatite coating is deposited in simulated body fluid (SBF) without using special equipment or extremely high processing temperature, which mimics the biomimetic process.[12] In our previous paper, apatite-coated chitosan microspheres fabricated by different methods have been evaluated for their physicochemical and in vitro biological properties.[13] However, whether these microspheres can promote in vivo bone formation is unknown. To further learn the effect of preparation methods, in this paper, bone formation potential of apatite-coated chitosan microspheres is evaluated through ectopic and cranial bone defect models. 2. Materials and methods 2.1. Preparation of chitosan microspheres Chitosan microspheres were fabricated according to our previous study,[13] but with some modifications. Briefly, chitosan powder (85% deacetylation degree, Aldrich) was dissolved in 2% (v/v) acetic acid to give a 3% (w/v) chitosan solution. Genipin powder (Wako Chemicals) was dissolved in 60% (v/v) ethanol. The ratio of genipin/chitosan was kept at 10% (w/w). The microspheres obtained from coacervate precipitation method were made by dropping chitosan solution via a syringe into 2% (m/v) sodium hydroxide solution in 80% ethanol (v/v). After stirring for 1 h, the microspheres were rinsed with deionized water and were cross-linked in genipin solution for 24 h, followed by rinsing and drying (CM0). Microspheres obtained from emulsion crosslinking method were fabricated with a water/oil technique. Briefly, 25 mL chitosan solution was slowly added into 100 mL paraffin oil containing 1 mL span 80. Under constant stirring, genipin solution was added. 24 h later, the cross-linked microspheres were collected and rinsed (EM0). Apatite coating was prepared in a two-step procedure as previously described.[14] Briefly, both microspheres were soaked into five times SBF solution for two days. The resulting microspheres were then rinsed gently with deionized water and dried. They were abbreviated as apatite-coated coacervate-precipitated microspheres (CM1) and emulsion cross-linked microspheres (EM1), respectively.
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2.2. Physicochemical properties of chitosan microspheres Since the preparation process was modified, some physicochemical properties of microspheres were assessed again.[13] To determine the cross-linking degree, ninhydrin assay was performed.[15] Briefly, the test samples were heated within the ninhydrin solution for 20 min at 100 °C. Then the absorbance of the solution at 570 nm was recorded by a spectrophotometer (BIO-TEK ELX808, USA) using known concentrations of glycine as standard. The degree of cross-linking was defined as the ratio of the consumed amino groups in the cross-linked samples to the free amino groups in the corresponding uncross-linked samples.[16] The morphology of microspheres was examined by highresolution field scanning electron microscopy (SEM, FEI Sirion 200, USA). The chemical composition of the coating was characterized by X-ray diffractometer (XRD, Bruker D8 Advance, Germany) and Fourier transform infrared-attenuated total reflectance spectroscopy (FTIR-ATR, Nicolet 5700, USA). 2.3. In vitro biocompatibility using rat bone marrow-derived mesenchymal stem cells as the seeding cells 2.3.1. Cell culture Rat bone marrow-derived mesenchymal stem cells (BMSCs) were isolated and cultured according to the protocol reported previously.[17] Cells at passage 2–4 were used. Before the cell seeding, microspheres were sterilized by ultraviolet light and incubated with culture medium for 24 h. 2.3.2. Cell adhesion To evaluate cell adhesion and morphology, BMSCs (3.2 × 104 cells/well) were seeded on 10 mg microspheres in 96 culture plate. After culturing for 1 or 3 days, the cell/ microsphere constructs were observed by SEM. 2.3.3. Cell proliferation To assess cell proliferation, BMSCs were seeded on microspheres, in the same manner as the adhesion test. Twelve hours later, the cell/microsphere constructs were carefully transferred into a new plate. After 1, 3, 5, and 7 days, CCK-8 solution (Dojindo Laboratory, Kumamoto, Japan) was added to each well. Three hours later, the optical density of supernatant solution was measured at a wavelength of 450 nm (n = 6). 2.3.4. Cell differentiation To measure alkaline phosphatase (ALP) activity and the gene expression of osteogenic markers, cells (1 × 106 cells/well) were seeded on 40 mg microspheres in 24 culture plates. Twelve hours later, the cell/microsphere constructs were transferred into a new plate and were cultured in osteogenic medium, including 10 mM β-glycerophosphate, 50 μg/mL ascorbic acid, and 10−8 M dexamethasone. On day 7 and day 14, the samples were rinsed with PBS, lysed in 0.2% Triton X-100, followed by sonication and centrifugation. The ALP activity and total amount of protein were determined by p-nitrophenyl phosphate method (n = 6 for each time point).[18] On day 14, mRNA expression levels of ALP, collagen type I (COL I), osteocalcin (OCN), and glyceraldehyde 3-phosphate
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Primer sequences for RT-PCR.
Gene
Forward primer (5′-3′)
Reverse primer (5′-3′)
Alkaline phosphatase (ALP) Collagen type I (COL I) Osteocalcin (OCN) GADPH
CAATTAACGGCTGACACTGC TCTGCGACACAAGGAGTCTG CAAGTCCCACACAGCAACTC AACGACCCCTTCATTGAC
TTTCAGGGCATTTTTCAAGG GGGACCATCAACACCATCTC GGCTCCAAGTCCATTGTTGA TCCACGACATACTCAGCAC
dehydrogenase were also detected (Table 1). Total RNA was extracted using Trizol (Sigma–Aldrich, St. Louis, MO, USA). RNA (1 μg) was reverse transcribed using First Strand cDNA Synthesis Kit (Fermentas, Maryland, USA). Then polymerase chain reaction (PCR) was performed using 2 × PCR mix (Tiangen Biotech, Beijing, China). Finally, the PCR products were electrophoresed on a 2% agarose gel and were analyzed using Image J software. 2.4 In vivo transplantation 2.4.1. Ethics statement and preparation of BMSCs/microsphere constructs All animal experiments were approved by the Ethics Committee of School and Hospital of Stomatology, Wuhan University. Male Wistar rats with average weight at 200 g ± 15 g were obtained from the Experimental Animal Centre of Hubei Province. To prepare BMSCs/microsphere constructs, BMSCs (3.2 × 105 cells/well) were seeded on 15 mg microspheres in 96 culture plate and cultured in osteogenic medium for 1 week before transplantation. Moreover, in order to fix the microspheres transplanted in cranial defects, chitosan films were prepared. Briefly, 3% chitosan solution was cast on culture dishes and dried at 50 °C. Then, the formed thin films were neutralized with NaOH (0.1 M), washed with deionized water, and sterilized by ultraviolet light before transplantation. 2.4.2. Ectopic bone regeneration To assess ectopic bone regeneration, four dorsal subcutaneous pockets were created for each rat. The cell/microsphere constructs were randomly transplanted and the skin was carefully sutured. After 3, 6, and 12 weeks, three rats were sacrificed at each time point. The transplanted samples were removed, fixed, decalcified, serials dehydrated, and embedded in paraffin (n = 6). Serial sections (5 μm thick) were stained with hematoxylin and eosin (HE). 2.4.3. Orthotopic bone regeneration For cranial defect transplantation, a sagittal incision of approximately 15 mm was made in the dorsal part of the rat cranium. The subcutaneous tissue, muscle, and periosteum were dissected. For each rat, two full-thickness bone defects (5 mm in diameter) were created using a trephine bur under constant irrigation with sterile saline. Thirty cranial defects were randomly repaired as follows: (1) CM0/BMSCs group, the defects were filled with CM0 seeded with BMSCs (n = 6); (2) EM0/BMSCs group, the defects were filled with EM0 seeded with BMSCs (n = 6); (3) CM1/BMSCs group, the defects
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were filled with CM1 seeded with BMSCs (n = 6); (4) EM1/BMSCs group, the defects were filled with EM1 seeded with BMSCs (n = 6); (5) Black group, the defects were filled with nothing (n = 6). After that, the chitosan films were covered on the defects and the wound was closed in layers. Eight weeks later, all rats were sacrificed. The defect site and surrounding bone were carefully dissected. All specimens were placed in a custom-made holder with 4% paraformaldehyde and scanned by a Scanco μCT50 imaging system (Scanco Medical, Bassersdorf, Switzerland). The scanning was performed at 70 kVp and 112 mA with a 10 μm pixel size, 1024 reconstruction matrix, and 250 ms integration time. A Gaussian filter (sigma = 0.8 and support = 1) and a fixed threshold (value = 220) were used to properly segment the scaffolds and the mineralized bone from the background. After three-dimensional (3-D) reconstruction, the relative maturity of bone was analyzed using a protocol provided by the manufacturer of the Micro-CT scanner and distinguished by colors. The red color meant that the bone was relatively mature, while the green color meant that the bone was relatively immature. Since it was difficult to distinguish new bone from apatite coating through separating the μCT signal using global thresholds, a quantitative analysis of bone volume could not be obtained. After the μCT scanning, all samples were decalcified, dehydrated, embedded in paraffin, and sectioned into 5 μm thick sections for HE analysis. Histometric measurements were analyzed by Image-Pro-Plus 5.0 (Media Cybernetic, USA), according to the previous study.[19] New bone area ratio was calculated as the new bone area in the defect divided by the entire defect area. Defect closure was identified by the new bone ingrowth (mm) in the defect divided by the original defect width (mm). 2.5. Statistical analysis All data were presented as the means ± standard deviation. Statistical significance was assessed using one-way ANOVA with the Tukey Post Hoc test. The value of p < 0.05 was considered as statistically significant. 3. Results 3.1. Physicochemical properties of chitosan microspheres The cross-link degree of both microspheres is shown in Figure 1. EM0 had significant lower value than CM0 (40% vs. 80%, p < 0.05). The surface morphology is shown in Figure 2. EM0 demonstrated smooth surface (Figure 2(B) and (D)), whereas CM0 presented rough surfaces (Figure 2(A) and (C)). After being immersed in five times SBF,
Figure 1.
Cross-linking degree of microspheres. *p < 0.05.
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Figure 2. SEM, XRD, and FTIR spectra of chitosan microspheres. (A, C) Coacervate-precipitated microspheres before coating (CM0); (B, D) emulsion cross-linked microspheres before coating (EM0); (E, G, I) coacervate-precipitated microspheres after coating (CM1); (F, H, J) emulsion cross-linked microspheres after coating (EM1); (K) XRD patterns; (L) FTIR spectra. CaP: carbonate apatite. ■: Characteristic peak of carbonate apatite; *: Characteristic peak of chitosan.
the surface of EM1 was uniformly covered by a continuous coating (Figure 2(F) and (H)), whereas the coating on CM1 was not continuous, and the substrate of CM1 could be seen in some places (Figure 2(E) and (G)). Under higher magnification, the minerals formed on EM1 (Figure 2(J)) were denser than the minerals formed on CM1 (Figure 2(I)). The composition of the coatings was characterized by XRD and FTIR. CM1 showed both characteristic peaks of chitosan (2θ = 20°) and characteristic peaks of carbonate apatite (2θ = 25.8°, 28.8°, 31.8°, and 46.7°). These peaks corresponded to the (0 0 2), (2 1 0), (2 1 1), and (2 2 2) planes of apatite, respectively.[20], while the XRD patterns of EM1 only showed the characteristic peaks of apatite (Figure 2(K)). The FTIR spectra of CM0 and EM0 exhibited characteristic bands of peaks around 2900 cm−1 corresponding to –CH2, while 1654 cm−1 was the characteristic peak of amide I. The sharp peaks at 1153, 1070, and 1031 cm−1 were assigned to the C–O stretching vibrations. The peak at 895 cm−1 was assigned to the C–N bond. After coating, new spectra of apatite at 563, 601, 1021, 875, and 1413 cm−1 appeared in CM1 and EM1, in which 563, 601, and 1021 cm−1 were assigned to PO4 bond, while 875 and 1413 cm−1 were assigned to CO3 band.[8,21–24] It could be further observed that the bands of chitosan at 1654 and 895 cm−1 with weak intensity appeared in CM1 but not EM1 (Figure 2(L)). Based on the data of SEM, XRD, and FTIR, we concluded that the surface of EM1 was completely coated by an apatite layer, while the surface of CM1 was partly or thinly coated with an apatite layer.
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3.2. In vitro biocompatibility To assess the cell attachment, we cultured BMSCs on the microspheres for 1 day and 3 days, and evaluated the cell morphology with SEM. On day 1, cells were spherical on CM0 (Figure 3(A)) and EM0 (Figure 3(B)). In contrast, cells were flat and star-like with numerous pseudopodia spread around the cell body on CM1 (Figure 3(C)) and EM1 (Figure 3(D)). On day 3, although the cell number increased on CM0 (Figure 3(E)) and EM0 (Figure 3(F)), it was obviously lower than the cell population on CM1 (Figure 3(G)) and EM1 (Figure 3(H)). Also, the cells on EM1 had grown to sheet-like form and covered the whole surface area (Figure 3(H)). CCK-8 assay showed that after 1 day, there were more cells on CM1 and EM1 than the cells on CM0 and EM0, while there was no significant difference between CM1 and EM1. Five days later, the amount of cells on EM1 was higher than on CM1 (Figure 3(I), p < 0.05). We also found that cells on EM1 had higher ALP activity than cells on CM1 on both day 7 and day 14 (Figure 3(J), p < 0.05). To further monitor the cell differentiation status, the expression of osteogenetic genes was analyzed (Figure 3(K) and (L)). The ALP, COL I, and OCN expression levels were significantly increased in cells cultured on EM1 compared with cells cultured on CM1 (p < 0.05). 3.3. In vivo transplantation 3.3.1. Ectopic transplantation As shown in Figure 4, after 3 and 6 weeks, microspheres in both EM1/BMSCs group and CM1/BMSCs group were capsulized by fibrous tissue without evidence of acute
Figure 3. BMSCs adherence, proliferation, and osteogenic differentiation. (A, E) SEM images of BMSCs grown on CM0; (B, F) SEM images of BMSCs grown on EM0; (C, G) SEM images of BMSCs grown on CM1; (D, H) SEM images of BMSCs grown on EM1; (I) CCK-8 analysis; (J) Quantitative ALP activity; (K) Electrophoresis of PCR product for ALP, COL I, OCN, and GADPH; (K) The ratio of ALP, COL I, and OCN to GADPH mRNA expression. *p < 0.05.
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Figure 4. HE staining of sections from ectopic samples. The arrows indicate the microspheres which have undergone degradation. Bar = 200 μm.
inflammation and bone formation. The EM1/BMSCs group began to degrade with irregular border line after 6 weeks, and after 12 weeks, significant degradation was found. In contrast, the morphology of CM1 did not change much. There was still no bone formation in both groups after 12 weeks. 3.3.2. Cranial transplantation The 3D μCT images of the bone defect were shown in Figure 5. EM1/BMSCs group had the greatest potential to promote bone formation among the groups. As we found that mineralized tissue almost filled the whole defect region (Figure 5(E)).
Figure 5. Micro-CT evaluation of the repaired calvarial bone defects at eight weeks after implantation. (A, F) Blank group; (B, G) CM0/BMSCs group; (C, H) EM0/BMSCs group; (D, I) CM1/BMSCs group; (E, J) EM1/BMSCs group; (A–E) 3-D reconstructed images; (F–J) Analysis of the relative bone maturity. The yellow circle indicates the dimension of 5 mm bone defect. Bar = 1 mm. (Please see the online article for the colour version of this figure: http://dx.doi.org/ 10.1080/09205063.2014.970604.)
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Furthermore, the maturity of the new bone was similar to that of the native bone around the defect, both of which was indicated by red color (Figure 5(J)). So the formed new bone was considered to be relatively mature. CM1/BMSCs group (Figure 5(D)) had less potential to promote bone formation than EM1/BMSCs group (Figure 5(E)), but it had greater potential than the control groups, including Black group (Figure 5(A)), CM0/BMSCs group (Figure 5(B)), and EM0/BMSCs group (Figure 5(C)). Part of the formed new bone in the CM1/BMSCs group was indicated by green color which meant that the new bone was relatively immature (Figure 5(I)). Histological data further supported the Micro-CT finding. No obvious new bone was found in the Blank group (Figure 6(A)) and EM0/BMSCs group (Figure 6(B)). A small amount of bone was observed in the CM0/BMSCs group (Figure 6(C)), whereas obvious new bone grew along the edges of the defects in the CM1/BMSCs group (Figure 6(D)). In the EM1/BMSCs group, the formed new bone was thick and almost covered the whole defect (Figure 6(E)). At higher magnification, active new bone formation could be observed in the CM1/BMSCs group (Figure 6(I)) and EM1/BMSCs group (Figure 6(J)), as osteocytes and blood vessels were found in the inside and osteoblasts were lined at the outside of new bone, whereas in the control groups, only isolated bone islands (Figure 6(H)) or loose connective tissues (Figure 6(G) and (F)) were found. Histomorphometric analysis is shown in Figure 6(K) and (L). The percentages of defect closure and new bone area ratio in the EM1/BMSCs group were significantly higher than those in the CM1/BMSCs group (p < 0.05). 4. Discussion Chitosan microspheres have been widely used in tissue engineering to provide a structural template for cell seeding and extracellular matrix formation. According to our previous study, preparation methods could cause great effect on microspheres’ properties, which further influenced the growth and differentiation of MC3T3-E1 cells in vitro. However, whether preparation methods could affect the regeneration of bone in vivo was still unknown. For this reason, BMSCs, one of the mostly used seeding cells in bone tissue engineering, were thus investigated in place of MC3T3-E1 cell in this study. At the same time, some procedures were modified to optimize the microsphere preparation. For instance, previously, after dropping the chitosan solution into the alkali solution, the stabilized microspheres were firstly dried and then cross-linked. During this process, the drying before cross-linking was time-taking and did affect the morphology of microspheres. So, in this study, the microspheres were directly cross-linked in the solution without first drying. The process was thus simplified and the microspheres were really more spherical than before (data not shown). However, unexpectedly, the cross-linking degree of CM0 increased much, which reached 80% and was significantly higher than that of EM0’s. We speculated that the highly hydrated microspheres in this study might be more vulnerable to the cross-link reaction than the dried microspheres in our previous study. Oppositely, SEM observation, XRD patterns, and FTIR spectra demonstrated that the apatite coating was more favorable to form on EM1 than on CM1. Studies have shown that amino groups could act as the nucleation site during the coating process.[10,25] Therefore, the calcium-binding capacity of chitosan was strictly controlled by the number of free amino groups.[25] Since EM0 had lower cross-linking degree, they naturally possessed more free amino groups on the surface. As a result, apatite could be formed more efficiently on EM1 than on CM1. Toshiki Miyazaki et al. also
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Figure 6. Histological analysis of bone formation in calvarial bone defects stained with HE. (A, F) Blank group; (B, G) EM0/BMSCs; (C, H) CM0/BMSCs; (D, I) CM1/BMSCs; (E, J) EM1/ BMSCs; (K) histomorphometirc analysis of the defect closure; (L) histomorphometirc analysis of the new bone area ratio, (A–E) Bar = 400 μm; (F–J) Bar = 50 μm. The thick black arrow: newly formed bone; the thin black arrow: new vessels; the thin white arrow: osteoblasts; the thin blue arrow: osteocytes. *p < 0.05. (Please see the online article for the colour version of this figure: http://dx.doi.org/10.1080/09205063.2014.970604.)
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found that PGA hydrogels with low cross-linking degree had high apatite-forming ability.[26] However, in our previous study, apatite coating was more favorable to form on CM1 than on EM1.[13] The reason might be that, the difference of cross-linking degree between CM0 and EM0 in the previous study was much smaller than that in the present study (10% vs 40%), so the effect of cross-linking degree on the apatite formation was minimized. Furthermore, the surface of CM0 in the previous study was irregular and shriveled (data not shown), which might favor the physical deposition of apatite. Turning to the biocompatibility, the CCK-8 assay was chosen to evaluate the cell proliferation, since it could monitor the change of cell numbers dynamically. We found that rat BMSCs presented higher proliferation and ALP activity on EM1 than those on CM1. Osteogenic gene expression of BMSCs on the microspheres was also assessed. ALP is an early marker and one of the most frequently used indicators to demonstrate osteogenic differentiation.[27] COL I constitutes around 90% of the organic matrix, which plays an important role in the formation of inorganic matrix of bone.[28] OCN is a later marker of differentiation marker, which is exclusively secreted by mature osteoblasts and present in mineralized matrix.[29] We found that the gene expressions of ALP, COL 1, and OCN were increased on EM1 in contrast to CM1. The reason might be that EM1 had better apatite coatings and apatite could promote the osteogenic differentiation. We further evaluated the in vivo bone regeneration of microspheres using ectopic and orthotopic models. Firstly, CM1/BMSCs and EM1/BMSCs were implanted subcutaneously in rats. After 3, 6, and 12 weeks, it was found that there was no obvious bone formation. Ectopic bone formation model was quite different from the complex biological environment of bone.[30,31] Some researchers reported that apatite-containing biomaterials could successfully promote ectopic bone formation, while others held a totally different view.[32,33] The reason might be that many factors could affect the in vivo behaviors of apatite-containing biomaterials, such as the physicochemical and structural properties of the apatite crystals and the animal models. It was also found that the degradation of EM1 was faster than that of CM1, which might stem from the lower cross-linking degree of EM1. Additional, both groups demonstrated no obvious inflammation indicating that both CM1 and EM1 were safe to the peri-tissue. Secondly, microspheres were implanted in the bone defects. Different from subcutaneous implantation, bone regeneration in cranial defects was favorable. We found that EM1/BMSCs group demonstrated better performance than CM1/BMSCs group. The reasons were as follows: (1) Apatite significantly enhanced the bone repair of chitosan microspheres. It has been reported that ion dissolution products containing Ca and P ions that were released from apatite could stimulate cell attachment, proliferation, and differentiation. In addition, Ca and P ions could activate the bone morphogenetic protein-signaling pathway, which played an important role in an autocrine/paracrine osteoinduction loop and enhanced bone formation in defects.[34] It has also been demonstrated that an apatite layer could preferentially adsorb a high quantity of protein from the serum such as fibronectin, which were then utilized by the osteoblasts to form a mineralized extracellular matrix.[35] (2) The difference in the amount of apatite coatings might affect the cell behaviors and bone formation of scaffolds. In this study, we found that EM1 had the higher degree of apatite formation, since the surface of EM1 was completely coated, while the surface of CM1 was partly or thinly coated by an apatite layer. It has been reported that high apatite content was more effective in inducing new bone formation than low apatite content scaffold.[36] He and coworkers
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further clarified that hydroxyapatite could act as nucleation sites for mineralization and promote further deposition of apatite crystals from cells, thus enhancing the formation of a mineral structure.[37] Several methods, including mass change method, scratch method, and alizarin red-based assay have been tried out to quantify the mineral contents. However, we were not successful since the microspheres were small, spherical, and the content of mineral was low. An effective and accurate method to quantify the amount of apatite coating might be taken in the further research. 5. Conclusions Findings from the present study demonstrate that preparation methods do exert great influence on the in vitro cell behaviors and in vivo orthotopic bone regeneration of apatite-coated chitosan microspheres. Appropriate method should be considered when preparing chitosan microspheres for bone tissue engineering scaffold. Funding This work was supported by the National Natural Science Foundation of China [grant number 81200812], [grant number 81271179]; the Provincial Natural Science Foundation of Hubei [grant number 2011CDB470].
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