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Preparation of porous bioactive ceramic microspheres and in vitro osteoblastic culturing for tissue engineering application
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Acta Biomaterialia 5 (2009) 1725–1731 www.elsevier.com/locate/actabiomat
Preparation of porous bioactive ceramic microspheres and in vitro osteoblastic culturing for tissue engineering application S.-J. Hong a, H.-S. Yu a, H.-W. Kim a,b,* a
Department of Biomaterials Science, School of Dentistry, Dankook University, Cheonan, 330-714, South Korea Institute of Tissue Regeneration Engineering (ITREN), Dankook University, Cheonan, 330-714, South Korea
b
Received 4 June 2008; received in revised form 11 November 2008; accepted 5 December 2008 Available online 25 December 2008
Abstract Microparticulates are useful for directly filling defective tissues as well as for delivering cells and bioactive molecules in regenerative medicine. This paper reports on the production of bioactive ceramic microspheres with an interconnected macropore structure. The sol–gel derived calcium silicate powder was homogenized with an oligomeric Camphene melt, which was used as a novel porogen, and sphericalshaped microparticulates were obtained by an oil-in-water emulsion method. A porous structure was generated through the sublimation of Camphene within the calcium silicate–Camphene solidified blend under ambient conditions. The microspheres retained the crystalline phase of apatite and wollastonite during heat treatment and induced calcium phosphate precipitation under a body-simulating medium, showing the characteristics of bone-bioactive materials. Osteoblastic cells were observed to anchor to and spread well over the surface of the porous microspheres, and further to proliferate actively with culturing time. The bioactive and porous microspheres developed are considered potentially useful in the regeneration of hard tissues as a matrix for tissue engineering as well as a direct filling material. Ó 2008 Acta Materialia Inc. Published by Elsevier Ltd. All rights reserved. Keywords: Bioactive ceramics; Microspheres; Porous structure; Osteoblast culturing; Tissue engineering
1. Introduction Skeletal defects caused by trauma, damage and developmental recession are generally treated by a variety of therapeutic approaches and with the aid of medical materials [1,2]. Some types of bioceramics, such as calcium phosphates and silica-based glasses, are already used clinically to regenerate defective bone and tooth structures [2–5]. However, their granular forms ranging in size from tens to hundreds of micrometers have been applied mainly to recover the small-sized and/or non-load-bearing defects, owing to mechanical weakness [6]. Granular bioceramics introduced to fill the defective sites act as a scaffold for the cellular population and matrix synthesis, which ulti* Corresponding author. Address: Department of Biomaterials Science, School of Dentistry, Dankook University, Cheonan, 330-714, South Korea. Tel.: +82 41 550 1926/1928. E-mail address: [email protected] (H.-W. Kim).
mately form neo-bones and regenerate the structure through a remodeling process [6–8]. Recent studies on bioceramic granules have suggested their advanced use as bioactive fillers in conjunction with polymeric matrices as an injectable device [9,10]. Moreover, these materials might find future applications in the stem-cell-based tissue engineering of bioactive carriers [11,12]. The present study produced porous spherical microparticulates of a bioactive ceramic in an attempt to gain optimal performance of such applications in regenerative medicine. Compared with conventionally used irregular granules with a dense form, the current spherical-shaped microparticles contained highly interconnected pore channels, which are believed to play effective roles in the recruitment of cellular reaction and population and neo-tissue formation. Based on studies on polymeric or composite microparticles, special emphasis should be paid to control over the macro-structure and composition of the substrate to
1742-7061/$ - see front matter Ó 2008 Acta Materialia Inc. Published by Elsevier Ltd. All rights reserved. doi:10.1016/j.actbio.2008.12.006
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improve the cell and tissue responses [10–12]. The incorporation of a bioactive inorganic composition, such as bioactive glass and hydroxyapatite within polymeric microspheres resulted in an enhanced osteogenic potential [10,13,14]. Moreover, the introduction of an open space within the polymeric microparticulate was suggested to hold and populate more cells [14,15]. With this in mind, the present study developed porous bioactive ceramic microparticulates, where a novel porogen ‘‘Camphene” was used to create open channels. This paper describes the processing tools used to produce porous bioceramic microspheres, along with their in vitro biological performance. 2. Materials and methods 2.1. Preparation of porous microspheres Calcium silicate powders were first synthesized as a precursor for bioactive ceramic microspheres. Briefly, Ca(NO3)2 4H2O and tetraethyl orthosilicate (TEOS) were mixed in ethanol at a molar ratio of 1:1, using a basic catalyst (NaOH). After vigorous stirring, the solution was left to stand at 40 °C until it became a gel, and was then dried at 70 °C. The dried powder was crushed and calcined at 500 °C for 2 h to obtain fine powder. The calcined powder was mixed with Camphene (C10H16, Sigma–Aldrich) which was used as a porogen at 1:8 by weight at a temperature of 50 °C by ball milling, and the slurry mixture was added with a surfactant KD4 (oligomeric hypermer from Uniqema) at 0.75 wt.%. Poly vinyl butyral (PVB, Sigma–Aldrich) dissolved at 10 wt.% in dichloromethane was added at 15 vol.% to the slurry mixture. The use of Camphene in combination with KD4 was based on a previous report [16], and PVB was used as the binder. The mixture was homogenized by ball milling further for 24 h. The slurry mixture was added dropwise into a distilled water pool containing 2% PVA, with continuous stirring at 750 rpm. The slurry mixture was solidified for 30 min, and washed with icecooled distilled water through a filter paper. The filtered product was stored at 20 °C for 10 min, gathered onto an alumina crucible and dried overnight under ambient conditions. The weight change of the microspheres due to the sublimation of Camphene during the drying process was measured. The dried product was heat treated as follows: ramping to a temperature of 1400 °C at a heating rate of 2 °C min 1, holding for 3 h, and air cooling in a furnace. The experimental procedures used to prepare the porous bioactive microspheres are shown schematically in Fig. 1. 2.2. Characterization The morphology and microstructure of the microspheres were evaluated by scanning electron microscopy (SEM; Hitachi) after the Pt coating. The size distribution
of the microspheres was analyzed from the images using optical microscopy. The phase of the powders either heat-treated or not was analyzed by X-ray diffraction ˚ ) at a scan(XRD) using Cu Ka1 radiation (k = 1.54056 A 1 ning rate of 2 min . The in vitro bioactivity of the microspheres was assessed by incubating the microspheres in a simulated body fluid (SBF; containing ions of Na+ 142.0 mM, K+ 5.0 mM, Ca2+ 2.5 mM, Mg2+ 1.5 mM, Cl 147.8 mM, HCO3 4.2 mM, HPO42 1.0 mM, and SO42 0.5 mM) [17]. In particular, 100 mg of the microspheres contained in the polyethylene tube was immersed in 10 ml of SBF and incubated at 37 °C for different periods, with continuous stirring at 120 rpm. After incubation, the samples were removed, washed with distilled water and ethanol, dried in the oven, and the change in weight was recorded. The surface morphology was examined by SEM to observe precipitates produced on the microsphere surface, and the elements of the precipitates were analyzed by energy dispersive spectroscopy (EDS). 2.3. Cell growth test Preliminary cellular tests on the porous microspheres were carried out using undifferentiated murine calvarial cells (MC3T3-E1). Before the cells were seeded, the microspheres were washed twice with serum-free medium and loaded into the individual wells of a 96-well plate. Ten milligrams of the microspheres was placed into each well of a 96-well plate, and then a 75 ll aliquot of the cell suspension prepared at a density of either 6 104 cells ml 1 (low density) or 5 105 cells ml 1 (high density) was dropped onto each well containing the microspheres. After culturing for 6 h in an incubator humidified with 95% CO2, 125 ll aliquot of a culturing medium supplemented with 50 lg ml 1 sodium ascorbate, 10 mM sodium b-glycerol phosphate and 10 nM dexamethasone was added to each well in order to allow osteoblastic differentiation. After culturing for pre-determined periods, the morphology of the cells grown on the microspheres was observed by SEM after being fixed with glutaraldehyde, dehydrated with a graded series of ethanol (50%, 70%, 90%, 95% and 100%), treated twice with a hexamethyldisilazane (HMDS) solution, and coated with Pt. The level of cell growth was measured using an MTS (3(4,5-dimethylthiazol-2-yl)-5(3-carboxymethonyphenol)-2(4-sulfophenyl)-2H-tetrazolium) assay. At each culturing period (1, 5 and 15 day), the culture medium was decanted, and the CellTitero 96 AQueous One Solution Reagent (Promega, Madison) was added to each sample and reacted at 37 °C for 2 h. The absorbance at 490 nm was read using an Elisa Plate Reader. Five replicate samples were tested for each condition, and the data were expressed as means ± SD. Statistical analysis was carried out by one-way analysis of variance (ANOVA) followed by Bonferroni correction. The P value was considered significant at a level <0.05.
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Fig. 1. Schematic showing the experimental procedures used to prepare bioactive and porous ceramic microspheres. The mixture slurry of ceramic powder and Camphene porogen mediated by surfactant and binder is formulated into spheres within a poly(vinyl alcohol) (PVA)–water pool. During sphere formation, Camphene is solidified by a dendritic growth and subsequently sublimed under ambient conditions to generate interconnected pore channels. The microspheres produced are further heat treated to consolidate and provide structural stability and bioactivity.
3. Results and discussion 3.1. Production of porous microspheres Fig. 1 schematically depicts the procedure to fabricate macroporous bioactive ceramic microspheres. As illustrated, the mixture of calcium silicate powder and hydrophobic Camphene requires a surfactant, and the oligomeric hypermer KD4 used herein was observed to be effective in dispersing the ceramic nano-powders within the Camphene oil melt. Because of the low melting point of the Camphene, the powder slurry could be easily prepared within a molten Camphene without the use of other solvents. Within the slurry, the PVB binder should also be added to prevent the bulk-shaped microspheres from disin-
tegrating during the drying and heat treatment processes. In order to shape the mixture into a spherical form, PVA was added to the water pool to mediate the interface of oil droplets and water. Above all, the role of Camphene should be underscored in the sense that the powder–Camphene mixture was solidified easily in the course of the microsphere formulation because of the freezing point of Camphene (37 °C). Moreover, under atmospheric condition, Camphene easily sublimed to produce interconnected pore channels. These characteristics of Camphene allow the production of pores within the microspheres under ambient conditions. In practice, recent studies have used Camphene to fabricate ceramic porous scaffolds [16,18]. Specifically, Camphene solidifies with a dendritic growth [19] to form an interconnected network,
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which was separated from the ceramic phase. After sublimation of the solidified Camphene phase, microspheres with totally interconnected pores could thus be generated. Fig. 2 shows the typical morphology of the microspheres obtained after the Camphene sublimation and subsequent
Fig. 2. (A and B) SEM image of the microspheres heat treated at 1400 °C at different magnifications, and (C) size distribution of the microspheres. Interconnected pore channels can be seen, and the surface of the microspheres is microporous. Average diameter of the microspheres is 241 lm.
heat treatment at 1400 °C. Pores 20–50 lm in size were well generated throughout the spheres, while the surface was observed to be microporous (Fig. 2A and B). The mean diameter of the microspheres was measured to be 241 lm (Fig. 2C). The size of the microspheres was reduced considerably (30%) after heat treatment, which was attributed to the sintering of powders and the removal of binder. The pore size, which was less than 100 lm, might be increased by adjusting the solidification temperature of Camphene. It was reported that, in the alumina porous body, a pore size over 100 lm could be obtained by means of increasing the solidification temperature up to 35 °C, i.e., decreasing the driving force (temperature difference from melt to freezing) for the solidification. Practically, it is of benefit to increase the pore size because the microsphere system herein is intended to be used as a scaffolding matrix for the cells and tissue regeneration; which highlights the need for further study.
Fig. 3. (A) Weight change of the powder–Camphene blend microspheres showing almost complete sublimation of Camphene within 6 h. (B) XRD pattern of the Camphene-sublimed calcium silicate microspheres after heat treatment at different temperatures. New crystalline phases of apatite and wollastonite are observed after the heat treatment (CS, CaSiO3; A, apatite; W, wollastonite).
S.-J. Hong et al. / Acta Biomaterialia 5 (2009) 1725–1731
As shown in Fig. 3A, the Camphene sublimation occurred quite rapidly, finishing almost within 6 h. The heat treatment temperature over 1200 °C was needed to provide a level of mechanical stability associated with densification to the porous microspheres. As shown by the XRD pattern (Fig. 3B), the initial calcium silicate phase was observed to change into hydroxyapatite (denoted by A) and wollastonite (denoted by W) after heat treatment [20]. The crystalline peaks became more apparent with increasing heat treatment temperature. The apatite–wollastonite (A–W) dual phase is the major crystalline phase gen-
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erally observed in an A–W glass ceramic. Moreover, it has been reported to be bone-bioactive, forming a bone mineral-like phase on the surface both in vitro and in vivo, which consequently regulates the osteogenic responses and results in direct bonding to hard tissues [20,21]. 3.2. Bioactivity and osteoblastic culturing The in vitro bone bioactivity was investigated by immersing the microspheres in a SBF and observing the precipitation of calcium phosphate on the surface.
Fig. 4. In vitro apatite formation on the bioactive ceramic microspheres in SBF: (A–C) SEM images after incubation for 1 day (A) and 7 days (B and C). Some tiny crystallites start to precipitate on the surface at day 1 (A). At day 7, although there is no observable change at low magnification (C), significant formation of apatitic crystallites is noticed at high magnification (B). The EDS analysis of the SBF sample immersed for 7 days shown in (D) reveals the existence of P and the appearance of higher intensity of Ca relative to that of Si, with a Ca/P ratio of 1.53, which is in direct contrast to that of the initial sample (E), suggesting that the calcium phosphate mineral formed is similar to the calcium-deficient biomimetic apatite. (F) Weight change of the microspheres during SBF immersion, showing an increase initially and saturation at a later stage, which is associated with ionic dissolution and precipitation of calcium phosphate.
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Fig. 5. Osteoblastic cell growth behavior upon the microspheres: (A–C) electron micrograph of cells grown for (A) 5 and (B) 15 days at high cell-seeding density, and (C) for 15 days at low cell-seeding density. The cells adhere and grow actively on the microspheres, initially bridging the gaps (A), and later covering the surface almost completely at high cell-seeding density (B), while a relatively small number of cells are observed at low cell-seeding density (C). The cell viability with two different seeding densities is assessed using an MTS method at days 1, 5 and 15 (D), showing that the microspheres play an efficient role as a three-dimensional supporting matrix for cell growth. Significant difference between the groups seeded at different densities is noted at all culturing periods (ANOVA, P < 0.05).
Fig. 4 shows the SEM morphology of the microspheres after immersion in SBF for 1 and 7 days. Tiny precipitates were noticed on most parts of the microspheres after immersion for 1 day (Fig. 4A). At 7 days, nanocrystallites larger than those at 1 day were found to cover the whole surface of the microspheres (Fig. 4B). At the macroscopic level, there appears to be no significant change in the pore structure and microsphere shape (Fig. 4C at 7 days). The EDS spectrum in Fig. 4D shows the atomic composition of the precipitates after 7 days of immersion. The existence of P and a higher intensity of Ca relative to Si suggest the precipitation of calcium phosphate (compare with
Fig. 4E). The Ca/P ratio was measured to be 1.53, which is similar but slightly lower than that of stoichiometric hydroxyapatite (1.67), suggesting the formation of calcium-deficient hydroxyapatite. The weight of the microspheres was observed to increase with increasing immersion time, indicating the precipitation of calcium phosphate (Fig. 4F). Although ionic dissolution (such as calcium and silicon) from the microspheres occurs simultaneously, the precipitation process is considered to dominate, at least during the period tested in the present study. The result suggests the microspheres containing A–W phase are bioactive, at least in vitro, to form cal-
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cium phosphate crystals in SBF, and are considered useful for hard tissue regeneration. The tissue cell responses to the porous bioactive ceramic microspheres were briefly examined in terms of the initial cell adhesion and viability with culturing time, as shown in Fig. 5. Pre-osteoblastic MC3T3-E1 cells were seeded onto the microspheres at two different cell densities (low 6 104 ml 1 and high 5 105 ml 1), and cultured statically in an osteogenic medium for up to 15 days. When seeded at high density, the cells adhered well to the microsphere surface, bridging the surface gaps during the initial culturing period (for 5 days) (Fig. 5A). Moreover, the cells proliferated actively with prolonged culturing (for 15 days), covering the surface almost completely (Fig. 5B). Although the cells seeded at low density adhered well and multiplied with increasing culturing time, they could not effectively cover the entire surface of the microspheres even after culturing for 15 days (Fig. 5C). The cell viability, as measured by the MTS assay (Fig. 5D), exhibited an ongoing increase with culturing time upon the microspheres at both cell densities. However, the difference was significant (P < 0.05) at all culturing periods, reflecting the electron images. The initial difference at the cell adhesion stage (day 1) was shown to maintain up to the following cell proliferative phase (5 and 15 days). Based on the cell growth rate, an extended culturing time may be required to gain full coverage of the surface of the microspheres at the low seeding density. Data suggest that the initial cell-seeding density should be carefully adjusted when using the microspheres as a cell delivery vehicle and tissue engineering matrix. The population density and confluence time of the cells upon the three-dimensional substrate are of special importance for gaining optimal conditions for regulating osteogenic differentiation and the nature of extracellular matrices. Based on this preliminary cell culturing study, the porous bioactive microspheres showed favorable responses to osteoblastic cells, providing appropriate scaffold conditions for the initial adhesion and growth of cells. However, more in-depth studies on the osteogenic potential and tissue engineering feasibility of these microspheres are needed. 4. Conclusion The present study developed novel porous microspheres constituted by an A–W bioactive ceramic phase for the applications in hard tissue regeneration. Based on the in vitro bone bioactivity and cell population behavior, the microspheres are considered to be useful as a three-dimensional matrix for the bone tissue engineering and cell delivery.
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References [1] Hench LL, Polak JM. Third-generation biomedical materials. Science 2002;295:1014. [2] Ikada Y. Challenges in tissue engineering. J R Soc Interface 2006;22:589–601. [3] Sepulveda P, Jones JR, Hench LL. Bioactive sol–gel foams for tissue repair. J Biomed Mater Res A 2002;59A:340–8. [4] Kokubo T. Bioactive glass ceramics: properties and applications. Biomaterials 1991;12:155. [5] Jarcho M. Calcium phosphate ceramics as hard tissue prosthesis. Clin Orthop 1981;157:259–78. [6] Al Ruhaimi KA. Bone graft substitutes: a comparative qualitative histologic review of current osteoconductive grafting materials. Int J Oral Maxillofac Implants 2001;16(1):105–14. [7] Damien CJ, Parsons JR. Bone graft and bone graft substitutes: a review of current technology and applications. J App Biomat 1991;2:187–208. [8] Behairy Y, Jasty M. Bone grafts and bone substitutes in hip and knee surgery. Orthop Clin North Am 1999;30(4):661–7. [9] Kim HW, Yoon BH, Kim HE. Microsphere of apatite–gelatin nanocomposite as bone regenerative filler. J Mater Sci Mater Med 2005;16(12):1105–9. [10] Gu HJ, Lee HH, Kim HW. Microspheres of collagen–apatite nanocomposites with osteogenic potential for tissue engineering. Tissue Eng 2007;13(5):965–72. [11] Borden M, Attawia M, Khan Y, Laurencin CT. Tissue engineered microsphere-based matrices for bone repair: design and evaluation. Biomaterials 2002;23:551–9. [12] Rokstad AM, Holtan S, Strand B, Steinkjer B, Ryan L, Kulseng B, Skjak-Braek G, Espevik T. Microencapsulation of cells producing therapeutic proteins: optimizing cell growth and secretion. Cell Transplant 2002;11(4):313–24. [13] Qiu QQ, Ducheyne P, Ayyaswamy PS. New bioactive, degradable composite microspheres as tissue engineering substrates. J Biomed Mater Res 2000;52:66. [14] Maeda H, Kasuga T. Preparation of poly(lactic acid) composite hollow spheres containing calcium carbonates. Acta Biomater 2006;2(4):403–8. [15] Lee HH, Hong SJ, Kim CH, Kim EC, Jang JH, Shin HI, et al. Preparation of hydroxyapatite spheres with an internal cavity as a scaffold for hard tissue regeneration. J Mater Sci Mater Med 2008;19:3029–34. [16] Koh YH, Lee EJ, Yoon BH, Song JH, Kim HE, Kim HW. Effect of polystyrene addition on freeze casting of ceramic/camphene slurry for ultra-high porosity ceramics with aligned pore channels. J Am Ceram Soc 2006;89:3646–53. [17] Kokubo T. Bioactive glass ceramics: properties and applications. Biomaterials 1991;12:155–63. [18] Lee EJ, Koh YH, Yoon BH, Kim HE, Kim HW. Highly porous hydroxyapatite bioceramics with interconnected pore channels using camphene-based freeze casting. Mater Lett 2007;61:2270–3. [19] Araki K, Halloran JW. New freeze casting technique for ceramics with sublimable vehicle. J Am Ceram Soc 2004;87:1859–63. [20] Yamamuro T. A/W glass–ceramic: clinical applications. In: Hench LL, Wilson J, editors. An introduction to bioceramics, advanced series in ceramics, vol. 1. Singapore: World Scientific; 1993. p. 89–103. [21] Nakamura T, Yamamuro T, Higashi S, Kokubo T, Ito S. A new glass–ceramic for bone replacement: evaluation of its bonding to bone tissue. J Biomed Mater Res 1985;19:685–9.
Acta Biomaterialia 5 (2009) 1725–1731 www.elsevier.com/locate/actabiomat
Preparation of porous bioactive ceramic microspheres and in vitro osteoblastic culturing for tissue engineering application S.-J. Hong a, H.-S. Yu a, H.-W. Kim a,b,* a
Department of Biomaterials Science, School of Dentistry, Dankook University, Cheonan, 330-714, South Korea Institute of Tissue Regeneration Engineering (ITREN), Dankook University, Cheonan, 330-714, South Korea
b
Received 4 June 2008; received in revised form 11 November 2008; accepted 5 December 2008 Available online 25 December 2008
Abstract Microparticulates are useful for directly filling defective tissues as well as for delivering cells and bioactive molecules in regenerative medicine. This paper reports on the production of bioactive ceramic microspheres with an interconnected macropore structure. The sol–gel derived calcium silicate powder was homogenized with an oligomeric Camphene melt, which was used as a novel porogen, and sphericalshaped microparticulates were obtained by an oil-in-water emulsion method. A porous structure was generated through the sublimation of Camphene within the calcium silicate–Camphene solidified blend under ambient conditions. The microspheres retained the crystalline phase of apatite and wollastonite during heat treatment and induced calcium phosphate precipitation under a body-simulating medium, showing the characteristics of bone-bioactive materials. Osteoblastic cells were observed to anchor to and spread well over the surface of the porous microspheres, and further to proliferate actively with culturing time. The bioactive and porous microspheres developed are considered potentially useful in the regeneration of hard tissues as a matrix for tissue engineering as well as a direct filling material. Ó 2008 Acta Materialia Inc. Published by Elsevier Ltd. All rights reserved. Keywords: Bioactive ceramics; Microspheres; Porous structure; Osteoblast culturing; Tissue engineering
1. Introduction Skeletal defects caused by trauma, damage and developmental recession are generally treated by a variety of therapeutic approaches and with the aid of medical materials [1,2]. Some types of bioceramics, such as calcium phosphates and silica-based glasses, are already used clinically to regenerate defective bone and tooth structures [2–5]. However, their granular forms ranging in size from tens to hundreds of micrometers have been applied mainly to recover the small-sized and/or non-load-bearing defects, owing to mechanical weakness [6]. Granular bioceramics introduced to fill the defective sites act as a scaffold for the cellular population and matrix synthesis, which ulti* Corresponding author. Address: Department of Biomaterials Science, School of Dentistry, Dankook University, Cheonan, 330-714, South Korea. Tel.: +82 41 550 1926/1928. E-mail address: [email protected] (H.-W. Kim).
mately form neo-bones and regenerate the structure through a remodeling process [6–8]. Recent studies on bioceramic granules have suggested their advanced use as bioactive fillers in conjunction with polymeric matrices as an injectable device [9,10]. Moreover, these materials might find future applications in the stem-cell-based tissue engineering of bioactive carriers [11,12]. The present study produced porous spherical microparticulates of a bioactive ceramic in an attempt to gain optimal performance of such applications in regenerative medicine. Compared with conventionally used irregular granules with a dense form, the current spherical-shaped microparticles contained highly interconnected pore channels, which are believed to play effective roles in the recruitment of cellular reaction and population and neo-tissue formation. Based on studies on polymeric or composite microparticles, special emphasis should be paid to control over the macro-structure and composition of the substrate to
1742-7061/$ - see front matter Ó 2008 Acta Materialia Inc. Published by Elsevier Ltd. All rights reserved. doi:10.1016/j.actbio.2008.12.006
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improve the cell and tissue responses [10–12]. The incorporation of a bioactive inorganic composition, such as bioactive glass and hydroxyapatite within polymeric microspheres resulted in an enhanced osteogenic potential [10,13,14]. Moreover, the introduction of an open space within the polymeric microparticulate was suggested to hold and populate more cells [14,15]. With this in mind, the present study developed porous bioactive ceramic microparticulates, where a novel porogen ‘‘Camphene” was used to create open channels. This paper describes the processing tools used to produce porous bioceramic microspheres, along with their in vitro biological performance. 2. Materials and methods 2.1. Preparation of porous microspheres Calcium silicate powders were first synthesized as a precursor for bioactive ceramic microspheres. Briefly, Ca(NO3)2 4H2O and tetraethyl orthosilicate (TEOS) were mixed in ethanol at a molar ratio of 1:1, using a basic catalyst (NaOH). After vigorous stirring, the solution was left to stand at 40 °C until it became a gel, and was then dried at 70 °C. The dried powder was crushed and calcined at 500 °C for 2 h to obtain fine powder. The calcined powder was mixed with Camphene (C10H16, Sigma–Aldrich) which was used as a porogen at 1:8 by weight at a temperature of 50 °C by ball milling, and the slurry mixture was added with a surfactant KD4 (oligomeric hypermer from Uniqema) at 0.75 wt.%. Poly vinyl butyral (PVB, Sigma–Aldrich) dissolved at 10 wt.% in dichloromethane was added at 15 vol.% to the slurry mixture. The use of Camphene in combination with KD4 was based on a previous report [16], and PVB was used as the binder. The mixture was homogenized by ball milling further for 24 h. The slurry mixture was added dropwise into a distilled water pool containing 2% PVA, with continuous stirring at 750 rpm. The slurry mixture was solidified for 30 min, and washed with icecooled distilled water through a filter paper. The filtered product was stored at 20 °C for 10 min, gathered onto an alumina crucible and dried overnight under ambient conditions. The weight change of the microspheres due to the sublimation of Camphene during the drying process was measured. The dried product was heat treated as follows: ramping to a temperature of 1400 °C at a heating rate of 2 °C min 1, holding for 3 h, and air cooling in a furnace. The experimental procedures used to prepare the porous bioactive microspheres are shown schematically in Fig. 1. 2.2. Characterization The morphology and microstructure of the microspheres were evaluated by scanning electron microscopy (SEM; Hitachi) after the Pt coating. The size distribution
of the microspheres was analyzed from the images using optical microscopy. The phase of the powders either heat-treated or not was analyzed by X-ray diffraction ˚ ) at a scan(XRD) using Cu Ka1 radiation (k = 1.54056 A 1 ning rate of 2 min . The in vitro bioactivity of the microspheres was assessed by incubating the microspheres in a simulated body fluid (SBF; containing ions of Na+ 142.0 mM, K+ 5.0 mM, Ca2+ 2.5 mM, Mg2+ 1.5 mM, Cl 147.8 mM, HCO3 4.2 mM, HPO42 1.0 mM, and SO42 0.5 mM) [17]. In particular, 100 mg of the microspheres contained in the polyethylene tube was immersed in 10 ml of SBF and incubated at 37 °C for different periods, with continuous stirring at 120 rpm. After incubation, the samples were removed, washed with distilled water and ethanol, dried in the oven, and the change in weight was recorded. The surface morphology was examined by SEM to observe precipitates produced on the microsphere surface, and the elements of the precipitates were analyzed by energy dispersive spectroscopy (EDS). 2.3. Cell growth test Preliminary cellular tests on the porous microspheres were carried out using undifferentiated murine calvarial cells (MC3T3-E1). Before the cells were seeded, the microspheres were washed twice with serum-free medium and loaded into the individual wells of a 96-well plate. Ten milligrams of the microspheres was placed into each well of a 96-well plate, and then a 75 ll aliquot of the cell suspension prepared at a density of either 6 104 cells ml 1 (low density) or 5 105 cells ml 1 (high density) was dropped onto each well containing the microspheres. After culturing for 6 h in an incubator humidified with 95% CO2, 125 ll aliquot of a culturing medium supplemented with 50 lg ml 1 sodium ascorbate, 10 mM sodium b-glycerol phosphate and 10 nM dexamethasone was added to each well in order to allow osteoblastic differentiation. After culturing for pre-determined periods, the morphology of the cells grown on the microspheres was observed by SEM after being fixed with glutaraldehyde, dehydrated with a graded series of ethanol (50%, 70%, 90%, 95% and 100%), treated twice with a hexamethyldisilazane (HMDS) solution, and coated with Pt. The level of cell growth was measured using an MTS (3(4,5-dimethylthiazol-2-yl)-5(3-carboxymethonyphenol)-2(4-sulfophenyl)-2H-tetrazolium) assay. At each culturing period (1, 5 and 15 day), the culture medium was decanted, and the CellTitero 96 AQueous One Solution Reagent (Promega, Madison) was added to each sample and reacted at 37 °C for 2 h. The absorbance at 490 nm was read using an Elisa Plate Reader. Five replicate samples were tested for each condition, and the data were expressed as means ± SD. Statistical analysis was carried out by one-way analysis of variance (ANOVA) followed by Bonferroni correction. The P value was considered significant at a level <0.05.
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Fig. 1. Schematic showing the experimental procedures used to prepare bioactive and porous ceramic microspheres. The mixture slurry of ceramic powder and Camphene porogen mediated by surfactant and binder is formulated into spheres within a poly(vinyl alcohol) (PVA)–water pool. During sphere formation, Camphene is solidified by a dendritic growth and subsequently sublimed under ambient conditions to generate interconnected pore channels. The microspheres produced are further heat treated to consolidate and provide structural stability and bioactivity.
3. Results and discussion 3.1. Production of porous microspheres Fig. 1 schematically depicts the procedure to fabricate macroporous bioactive ceramic microspheres. As illustrated, the mixture of calcium silicate powder and hydrophobic Camphene requires a surfactant, and the oligomeric hypermer KD4 used herein was observed to be effective in dispersing the ceramic nano-powders within the Camphene oil melt. Because of the low melting point of the Camphene, the powder slurry could be easily prepared within a molten Camphene without the use of other solvents. Within the slurry, the PVB binder should also be added to prevent the bulk-shaped microspheres from disin-
tegrating during the drying and heat treatment processes. In order to shape the mixture into a spherical form, PVA was added to the water pool to mediate the interface of oil droplets and water. Above all, the role of Camphene should be underscored in the sense that the powder–Camphene mixture was solidified easily in the course of the microsphere formulation because of the freezing point of Camphene (37 °C). Moreover, under atmospheric condition, Camphene easily sublimed to produce interconnected pore channels. These characteristics of Camphene allow the production of pores within the microspheres under ambient conditions. In practice, recent studies have used Camphene to fabricate ceramic porous scaffolds [16,18]. Specifically, Camphene solidifies with a dendritic growth [19] to form an interconnected network,
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which was separated from the ceramic phase. After sublimation of the solidified Camphene phase, microspheres with totally interconnected pores could thus be generated. Fig. 2 shows the typical morphology of the microspheres obtained after the Camphene sublimation and subsequent
Fig. 2. (A and B) SEM image of the microspheres heat treated at 1400 °C at different magnifications, and (C) size distribution of the microspheres. Interconnected pore channels can be seen, and the surface of the microspheres is microporous. Average diameter of the microspheres is 241 lm.
heat treatment at 1400 °C. Pores 20–50 lm in size were well generated throughout the spheres, while the surface was observed to be microporous (Fig. 2A and B). The mean diameter of the microspheres was measured to be 241 lm (Fig. 2C). The size of the microspheres was reduced considerably (30%) after heat treatment, which was attributed to the sintering of powders and the removal of binder. The pore size, which was less than 100 lm, might be increased by adjusting the solidification temperature of Camphene. It was reported that, in the alumina porous body, a pore size over 100 lm could be obtained by means of increasing the solidification temperature up to 35 °C, i.e., decreasing the driving force (temperature difference from melt to freezing) for the solidification. Practically, it is of benefit to increase the pore size because the microsphere system herein is intended to be used as a scaffolding matrix for the cells and tissue regeneration; which highlights the need for further study.
Fig. 3. (A) Weight change of the powder–Camphene blend microspheres showing almost complete sublimation of Camphene within 6 h. (B) XRD pattern of the Camphene-sublimed calcium silicate microspheres after heat treatment at different temperatures. New crystalline phases of apatite and wollastonite are observed after the heat treatment (CS, CaSiO3; A, apatite; W, wollastonite).
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As shown in Fig. 3A, the Camphene sublimation occurred quite rapidly, finishing almost within 6 h. The heat treatment temperature over 1200 °C was needed to provide a level of mechanical stability associated with densification to the porous microspheres. As shown by the XRD pattern (Fig. 3B), the initial calcium silicate phase was observed to change into hydroxyapatite (denoted by A) and wollastonite (denoted by W) after heat treatment [20]. The crystalline peaks became more apparent with increasing heat treatment temperature. The apatite–wollastonite (A–W) dual phase is the major crystalline phase gen-
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erally observed in an A–W glass ceramic. Moreover, it has been reported to be bone-bioactive, forming a bone mineral-like phase on the surface both in vitro and in vivo, which consequently regulates the osteogenic responses and results in direct bonding to hard tissues [20,21]. 3.2. Bioactivity and osteoblastic culturing The in vitro bone bioactivity was investigated by immersing the microspheres in a SBF and observing the precipitation of calcium phosphate on the surface.
Fig. 4. In vitro apatite formation on the bioactive ceramic microspheres in SBF: (A–C) SEM images after incubation for 1 day (A) and 7 days (B and C). Some tiny crystallites start to precipitate on the surface at day 1 (A). At day 7, although there is no observable change at low magnification (C), significant formation of apatitic crystallites is noticed at high magnification (B). The EDS analysis of the SBF sample immersed for 7 days shown in (D) reveals the existence of P and the appearance of higher intensity of Ca relative to that of Si, with a Ca/P ratio of 1.53, which is in direct contrast to that of the initial sample (E), suggesting that the calcium phosphate mineral formed is similar to the calcium-deficient biomimetic apatite. (F) Weight change of the microspheres during SBF immersion, showing an increase initially and saturation at a later stage, which is associated with ionic dissolution and precipitation of calcium phosphate.
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Fig. 5. Osteoblastic cell growth behavior upon the microspheres: (A–C) electron micrograph of cells grown for (A) 5 and (B) 15 days at high cell-seeding density, and (C) for 15 days at low cell-seeding density. The cells adhere and grow actively on the microspheres, initially bridging the gaps (A), and later covering the surface almost completely at high cell-seeding density (B), while a relatively small number of cells are observed at low cell-seeding density (C). The cell viability with two different seeding densities is assessed using an MTS method at days 1, 5 and 15 (D), showing that the microspheres play an efficient role as a three-dimensional supporting matrix for cell growth. Significant difference between the groups seeded at different densities is noted at all culturing periods (ANOVA, P < 0.05).
Fig. 4 shows the SEM morphology of the microspheres after immersion in SBF for 1 and 7 days. Tiny precipitates were noticed on most parts of the microspheres after immersion for 1 day (Fig. 4A). At 7 days, nanocrystallites larger than those at 1 day were found to cover the whole surface of the microspheres (Fig. 4B). At the macroscopic level, there appears to be no significant change in the pore structure and microsphere shape (Fig. 4C at 7 days). The EDS spectrum in Fig. 4D shows the atomic composition of the precipitates after 7 days of immersion. The existence of P and a higher intensity of Ca relative to Si suggest the precipitation of calcium phosphate (compare with
Fig. 4E). The Ca/P ratio was measured to be 1.53, which is similar but slightly lower than that of stoichiometric hydroxyapatite (1.67), suggesting the formation of calcium-deficient hydroxyapatite. The weight of the microspheres was observed to increase with increasing immersion time, indicating the precipitation of calcium phosphate (Fig. 4F). Although ionic dissolution (such as calcium and silicon) from the microspheres occurs simultaneously, the precipitation process is considered to dominate, at least during the period tested in the present study. The result suggests the microspheres containing A–W phase are bioactive, at least in vitro, to form cal-
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cium phosphate crystals in SBF, and are considered useful for hard tissue regeneration. The tissue cell responses to the porous bioactive ceramic microspheres were briefly examined in terms of the initial cell adhesion and viability with culturing time, as shown in Fig. 5. Pre-osteoblastic MC3T3-E1 cells were seeded onto the microspheres at two different cell densities (low 6 104 ml 1 and high 5 105 ml 1), and cultured statically in an osteogenic medium for up to 15 days. When seeded at high density, the cells adhered well to the microsphere surface, bridging the surface gaps during the initial culturing period (for 5 days) (Fig. 5A). Moreover, the cells proliferated actively with prolonged culturing (for 15 days), covering the surface almost completely (Fig. 5B). Although the cells seeded at low density adhered well and multiplied with increasing culturing time, they could not effectively cover the entire surface of the microspheres even after culturing for 15 days (Fig. 5C). The cell viability, as measured by the MTS assay (Fig. 5D), exhibited an ongoing increase with culturing time upon the microspheres at both cell densities. However, the difference was significant (P < 0.05) at all culturing periods, reflecting the electron images. The initial difference at the cell adhesion stage (day 1) was shown to maintain up to the following cell proliferative phase (5 and 15 days). Based on the cell growth rate, an extended culturing time may be required to gain full coverage of the surface of the microspheres at the low seeding density. Data suggest that the initial cell-seeding density should be carefully adjusted when using the microspheres as a cell delivery vehicle and tissue engineering matrix. The population density and confluence time of the cells upon the three-dimensional substrate are of special importance for gaining optimal conditions for regulating osteogenic differentiation and the nature of extracellular matrices. Based on this preliminary cell culturing study, the porous bioactive microspheres showed favorable responses to osteoblastic cells, providing appropriate scaffold conditions for the initial adhesion and growth of cells. However, more in-depth studies on the osteogenic potential and tissue engineering feasibility of these microspheres are needed. 4. Conclusion The present study developed novel porous microspheres constituted by an A–W bioactive ceramic phase for the applications in hard tissue regeneration. Based on the in vitro bone bioactivity and cell population behavior, the microspheres are considered to be useful as a three-dimensional matrix for the bone tissue engineering and cell delivery.
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