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Shear-mediated crystallization from amorphous calcium phosphate to bone apatite
journal of the mechanical behavior of biomedical materials 54 (2016) 131–140
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Research paper
Shear-mediated crystallization from amorphous calcium phosphate to bone apatite Xufeng Niua,b,c,n, Liyang Wanga, Feng Tiana, Lizhen Wanga, Ping Lia, Qingling Fengd, Yubo Fana,nn a Key Laboratory for Biomechanics and Mechanobiology of Ministry of Education, School of Biological Science and Medical Engineering, Beihang University, Beijing 100191, China b BUAA Research Institute, Guangzhou 510530, China c Research Institute of Beihang University in Shenzhen, Shenzhen 518057, China d State Key Laboratory of New Ceramic and Fine Processing, Tsinghua University, Beijing 100084, China
art i cle i nfo
ab st rac t
Article history:
The contribution of fluid shear stress (FSS) on the conversion of amorphous calcium
Received 31 May 2015
phosphate (ACP) to bone apatite is investigated. The ACP precursors are prepared by using
Received in revised form
a wet-chemistry method and further exposed to the constant FSS environment with values
13 September 2015
of 0.5, 1.0, 1.5, and 2.0 Pa. At the designated time points, the apatites are characterized by
Accepted 21 September 2015
transmission electron microscopy, X-ray diffraction, and inductively coupled plasma-mass
Available online 30 September 2015
spectroscopy. The results show that, the low FSS ( r1.0 Pa) has positive effects on the
Keywords:
transition of ACP, characterized by the accelerated crystallization velocity and the well-
Fluid shear stress
organized calcium-deficient hydroxyapatite (CDHA) structure, whereas the high FSS
Amorphous calcium phosphate
(41.0 Pa) has negative effects on this conversion process, characterized by the poor CDHA
Apatite
crystal morphologies and the destroyed structures. The bioactivity evaluations further
Crystallization
reveal that, compared with the FSS-free group, the CDHA prepared under 1.0 Pa FSS for 9 h
Cone-and-plate viscometer
presents the more biocompatible features with pre-osteoblast cells. These results are helpful for understanding the mechanism of apatite deposition in natural bone tissue. & 2015 Elsevier Ltd. All rights reserved.
1.
Introduction
dynamic tissue capable of mechanical load-induced structural and functional adaptation through the process of bone remodel-
Beginning with the pioneering work of Julius Wolff in the 19th
ing (Knothe Tate et al., 2011; Martínez-Reina et al., 2014; Weiner
century, it is well-recognized that the external mechanical
and Wagner, 1998). At cellular and molecular levels, mounting
stimulus plays a dominant role in bone formation (Chen et al.,
evidences suggest that both bone cells and interstitial fluid flow
2010; Frost, 2000). According to the present knowledge, bone is a
within the lacunar–canalicular system are able to sense and
n Corresponding author at: Key Laboratory for Biomechanics and Mechanobiology of Ministry of Education, School of Biological Science and Medical Engineering, Beihang University, Beijing 100191, China. Tel.: þ86 10 82338755; fax: þ86 10 82315554. nn Corresponding author. Tel./fax: þ86 10 82339428. E-mail addresses: [email protected] (X. Niu), [email protected] (Y. Fan).
http://dx.doi.org/10.1016/j.jmbbm.2015.09.024 1751-6161/& 2015 Elsevier Ltd. All rights reserved.
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journal of the mechanical behavior of biomedical materials 54 (2016) 131 –140
respond to the external mechanical load (Ciani et al., 2009; Han et al., 2004; Huo et al., 2010; Ponik et al., 2007). Bone cells, including osteocytes, osteoblasts, and osteoclasts, are the broadly investigated cells as mechanosensors during the process of bone remodeling (Khosla et al., 2008). These cells compose basic multicellular unit (BMU) within the bone remodeling compartment, where osteocytes are imprisoned in bone matrix whereas osteoclasts and osteoblasts are located in the surface (Eriksen et al., 2007; Hauge et al., 2001). In each BMU, osteocytes are the primary mechanosensor which can sense mechanical strain and trigger bone remodeling (Seeman and Delmas, 2006). Osteoclasts and osteoblasts are the performers of bone anabolism, in which the former can induce bone absorption while the latter can enhance bone formation (Ducy et al., 2000; Teitelbaum, 2000). Signals generated by osteocytes deep within the BMU reach the surface and elicit remodeling events by osteoclasts and osteoblasts. To respond to the external mechanical stimuli timely and accurately, these cells achieve the crosstalk through cell–cell contacts or through signal molecules (Canalis et al., 2013; Haugh et al., 2015; Malone et al., 2007). Besides bone cells, bone matrix is another structural component in natural bone tissue. Apatite is the main inorganic phase in bone matrix, which endows bone with superior mechanical properties. Bone apatite is composed of calcium phosphate-based minerals, predominantly by hydroxyapatite (HA) (Huang et al., 2014; Niu et al., 2009; Zhou et al., 2015). Previously, many researches were focused on the crystallization behavior during calcium phosphate formation (Massera et al., 2015; Nganga et al., 2012). One common view was that bone apatite came from amorphous calcium phosphate (ACP) (Boonrungsiman et al., 2012; Liu et al., 2011; Mahamid et al., 2010). To obtain the desired HA structure in vitro, the conversion of ACP was simulated in aqueous media and various influencing factors were investigated, including pH, temperature, and presence of foreign ions (Combes and Rey, 2010; Yoder et al., 2012a; Mccubbin et al., 2008). The results revealed that the presence of carbonate (Pasteris et al., 2012; Mason et al., 2009) and molecular H2O (Pasteris et al., 2014; Yoder et al., 2012b) groups played a critical role in stabilizing the structure of apatite. However, thinking that bone has the mechanical adaptation and interstitial fluid flow can also act on bone matrix, it is rational to deduce that the external biomechanical stimulus may have influence on the conversion of ACP. Regrettably, it was always neglected in the previous study. Among the complicated mechanical forces generated by external loading, fluid shear stress (FSS) is regarded as the predominant stimulus to bone cells and matrix under the physiological conditions and at the micro-level (Fritton and Weinbaum, 2009; Li et al., 2012). Moreover, in most conversion reactions of ACP occurring at physiological pH, the obtained apatite is calcium-deficient hydroxyapatite (CDHA). Therefore, in this paper, ACP precursor was prepared and further exposed to constant FSS environment. The evolution of ACP to CDHA was investigated by transmission electron microscopy (TEM) and X-ray diffraction (XRD), with the purpose of illuminating the effects of FSS in this conversion process.
2.
Materials and methods
2.1.
Preparation of ACP precursor
A wet chemistry method was used to get the ACP precursor. Two kinds of solution, 12.5 mmol of calcium nitrate tetrahydrate and 7.5 mmol of ammonium phosphate, were prepared individually with volume of 20 mL in distilled water. Calcium ions solution was then gradually dropped into orthophosphate ions solution to form a mixture, which contained calcium and orthophosphate ions with Ca/P mole ratio of 1.67 (corresponding to Ca/P mole ratio in HA (Dorozhkin, 2007; Zhang et al., 2003)). Aqua ammonia was used to adjust the pH to 7.4 during the dropping process. The obtained ACP precipitation was brought to the next procedure right away.
2.2.
Exposure of shear stress
Cone-and-plate viscometer was used to provide constant FSS environment in this research. The ACP precursor was shifted to Brookfield R/S-CPSþ rheometer (USA) and exposed individually to FSS with values of 0.5, 1.0, 1.5, 2.0 Pa for aging 3, 6, and 9 h, respectively. The obtained precipitation was then washed 3 times with distilled water and further lyophilized using the SP Scientific VirTis Advantage XL-70 freeze dryers (USA) to gain CDHA powders. The control, FSS-free CDHA sample, was prepared using the same procedure except biomechanical exposure.
2.3.
Characterization
To get the morphological properties, the CDHA powders were observed by JEOL JEM-2100F TEM (Japan). The samples were dispersed in ethanol and ultrasonicated for 30 min. Then, they were picked up with carbon coated copper grid (200 mesh) for TEM characterization. To acquire the information about crystalline structure, selected area electron diffraction (SAED) was used and XRD patterns of samples were also carried out at room temperature on Rigaku D/Max X-ray diffractometer (Japan) with Cu Kα radiation source (wavelength¼ 1.54 Å). The supplied voltage and current were set to 40 kV and 120 mA, respectively. The samples were exposed at a scan rate of 101/min from 31 to 701. To identify the composition of CDHA, the powders were further dissolved in 8% nitric acid and the Ca/P mole ratios were measured by Varian Vista MPX inductively coupled plasma mass spectroscopy (ICP-MS, USA).
2.4.
In vitro bioactivity assay
2.4.1.
Cell culture and CDHA sterilization
The thawed mouse calvaria-derived preosteoblastic cells MC3T3-E1 (American type culture collection) were incubated in α-MEM supplemented with 10% fetal bovine serum (FBS), 100 U/mL penicillin, and 100 μg/mL streptomycin in a humidified incubator at 37 1C with 5% CO2 (Thermo Fisher Scientific, China). The medium was changed every other day. Two types of CDHA powders, prepared under 1.0 Pa FSS or FSS-free environment for 9 h, were sterilized by exposing in ultraviolet for 1 h, and further used for cell seeding and evaluation.
journal of the mechanical behavior of biomedical materials 54 (2016) 131 –140
2.4.
2 Proliferation assay
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10% FBS. The CDHA powders were added at a density of
The MC3T3-E1 cells were seeded in 96-well tissue culture
300 μg/mL based on α-MEM solution. Cell counting kit-8 (CCK-
plate (2.5 103 cells/well) containing α-MEM medium plus
8) assay was carried on days 1, 4, and 7 after cell seeding. Cell
Fig. 1 – Schematic representation of FSS loading by cone-and-plate viscometer during the process of CDHA preparation. ACP precursor was synthesized using the wet chemistry method and further exposed to various FSS from 0.5 to 2.0 Pa. The obtained CDHA was separated at designated aging time points.
Fig. 2 – Morphologies and crystal characteristics of ACP to CDHA conversion under FSS-free environment. (a–e) TEM characterization after aging for 0, 3, 6, 9 and 24 h, respectively, with SAED pattern in the top right corner of each image. (f) XRD patterns after aging for 0, 3, 6, 9 and 24 h.
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journal of the mechanical behavior of biomedical materials 54 (2016) 131 –140
proliferation was measured following the CCK-8 assay kit (Dojindo, Japan) instructions. Briefly, 10 μL CCK-8 kit was added to each well and then the plate was incubated at 37 1C for 1.5 h. The absorbance was read at 450 nm in Varioskan Flash (Thermo Scientific, USA). All experiments were repeated in triplicate and each group had three replicates.
2.4.3.
Alkaline phosphatase (ALP) activity assay
For ALP assay, the MC3T3-E1 cells were co-cultured with 300 μg/mL CDHA powders in α-MEM medium plus 10% FBS for 7 and 14 d, respectively. Afterwards, the medium was removed. ; The cells with number of 1 105 were transferred to microcentrifuge tube and treated with RIPA (Applygen, China) at 4 1C for 10 min. The lysates were clarified by centrifugation. The supernatant was collected for ALP assay by measuring the release of p-nitrophenol at 405 nm using the ALP activity kit (Zhongsheng, China). The amount of ALP in the cells was calculated according to the formula provided by the kit.
2.4.4.
Statistical analysis
All experiments were conducted at least three times. The data were expressed as mean7standard deviation. Statistical analysis was performed using one-way analysis of variance. The difference was regarded as statistical significance when po0.05.
3.
Results and discussion
3.1.
Preparation of CDHA under FSS environment
Bone tissue is exposed to the complicated biomechanical environment. Previous theoretical studies have demonstrated that interstitial fluid in bone tissue can induce shear stress on the order of 0.8–3.0 Pa in vivo (Weinbaum et al., 1994). According to these results, FSS used in this research was set as 0.5, 1.0, 1.5 and 2.0 Pa. Moreover, just as shown in Fig. 1, cone-and-plate viscometer was introduced to perform FSS loading, which could supply homogeneous biomechanical environment towards the conversion of ACP to CDHA during the whole experimental procedure.
3.2. Crystallization of ACP to CDHA under different FSS environment Fig. 2 shows the conversion of ACP to CDHA at various aging times under FSS-free environment, with morphological transformations and SAED patterns shown at the top right corner in Fig. 2a–e and XRD patterns shown in Fig. 2f. Fig. 2a shows the spherical precursor, which could be defined as ACP, since this unstable calcium phosphate status had a typical diffraction
Fig. 3 – Morphologies and crystal characteristics of ACP to CDHA conversion under 0.5 Pa FSS environment. (a–c) TEM characterization after aging for 3, 6, and 9 h, respectively, with SAED pattern in the top right corner of each image. (d) HRTEM result after aging for 9 h. (e) XRD patterns after aging for 0, 3, 6, and 9 h.
journal of the mechanical behavior of biomedical materials 54 (2016) 131 –140
135
pattern of amorphous halo ring (Dorozhkin, 2010). With aging for
was almost accomplished. Such results were coincident with
3 h, the precursor became curly but still did not crystallize well (Fig. 2b). After aging for 6 h, the tiny sticks appeared with the length of 50–150 nm roughly, under which might be the platelets of octacalcium phosphate or not well crystallized HA (Fig. 2c). At the same time, the diffraction pattern changed from diffused
the previous reports (Dorozhkin, 2009; Kim et al., 2005). Therefore, to fully investigate the influence of biomechanical exposure on crystal formation and growth, the maximum FSS duration was chosen to 9 h in the following study. Fig. 3 shows the conversion of ACP to CDHA after exposing
rings to polycrystalline rings. Further extending aging time to 9 h led to the formation of needle-like or stick-like crystallites (Fig. 2d). Besides, the crystal morphologies in aging for 24 h group were similar to that in aging for 9 h group, and the series
to 0.5 Pa FSS for various aging times up to 9 h, with morphological transformations and SAED patterns shown at the top right corner in Fig. 3a–c and XRD patterns shown in Fig. 3e. Compared to the corresponding FSS-free groups, some distinct features could be distinguished. With exposing for 3 h
of rings in SAED pattern resulted in the almost accomplished crystallization (Fig. 2e). In XRD patterns (Fig. 2f), the curve was broad and flat for ACP group, which was the symbol of amorphous status. Then some main peaks of CDHA appeared after aging for 3 h, such as (002)
(Fig. 3a), the spherical ACP precursors elongated and amorphous halo rings disappeared. After exposing for 6 h (Fig. 3b), some needle-like or stick-like crystallites formed, which looked like thinner and longer compared to Fig. 2c. The
near 25.81, (211) near 31.81. The peak (100) at 10.81 and three peaks of (222) at 46.71, (213) at 49.51 and (004) at 53.21 arose gradually after aging for 6 and 9 h, which were the typical XRD pattern of CDHA and all these diffraction peaks could be
diffraction rings corresponding to (002) and (004) suggested that the crystallites already possessed the typical structure of CDHA. Further extending time to 9 h revealed some arcs consisted of rings (Fig. 3c), which were the character of
assigned to the standard card. Further prolonging time to 24 h did not bring about extra variations of XRD pattern. Based on these results, it was rational to infer that the crystallization of ACP to CDHA in the wet chemistry method occurred mostly within 9 h. After that, the crystal transition
crystalline structure having certain orientation. In XRD patterns (Fig. 3e), the typical peaks of CDHA already emerged after aging for 3 h, which included (002), (211), (222), (213) and (004). Further prolonging aging time to 6 and 9 h just increased peak intensities. These results meant
Fig. 4 – Morphologies and crystal characteristics of ACP to CDHA conversion under 1.0 Pa FSS environment. (a–c) TEM characterization after aging for 3, 6, and 9 h, with SAED pattern in the top right corner of each image. (d) HRTEM result after aging for 9 h. (e) XRD patterns after aging for 0, 3, 6, and 9 h.
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that 0.5 Pa FSS could promote transition of ACP towards CDHA. High resolution TEM (HRTEM) was another evidence of well-organized CDHA structure after exposing to 0.5 Pa FSS, since the d-spacing of 0.25 nm measured from Fig. 3d was coincident with one of d-spacings of HA (Olszta et al., 2007). Fig. 4 shows the conversion of ACP to CDHA after exposing to 1.0 Pa FSS for various aging times up to 9 h, which shows the similar results as shown in Fig. 3. Both TEM and XRD characterizations revealed the accelerated conversion of ACP precursors towards CDHA in comparison with FSS-free group. The formed CDHA crystallites had the trends to be thinner and longer. They also presented the well-organized crystal structure and homogenous orientation, characterized by crystal lattices arranged in order with the distance of 0.23 nm for (310) and 0.33 nm for (002) (Fig. 4d), both were d-spacings of HA (Olszta et al., 2007). All these features demonstrated the positive effects of 1.0 Pa FSS on the crystallization of ACP. Fig. 5 shows the conversion of ACP to CDHA after exposing to 1.5 Pa FSS for various aging times up to 9 h. When exposing for 3 h, both TEM and XRD characterizations indicated the accelerated crystallization process. As a result, Fig. 5a presents the typical thin and straight needle-like CDHA crystallites. However, after exposing for 6 and even 9 h (Fig. 5b and c), TEM characterization revealed the poorer crystal morphologies as compared with the corresponding groups in FSS-free
environment (Fig. 2c and d). HRTEM also illustrated the different d-spacings at various districts in an individual crystallite (Fig. 5d), which was different from the results shown in Figs. 3d and 4d and meant that the inner structure of CDHA crystallites might not be benefitted from the increased FSS. Fig. 6 further shows the conversion of ACP to CDHA after exposing to 2.0 Pa FSS for 3 and 6 h. Further extending aging time to 9 h led to the absence of any CDHA crystals (data not shown). The imperfect SAED patterns shown in Fig. 6a and b suggested the slowed crystallizing speed. XRD patterns also indicated that the crystallization process was greatly decreased, since the representative peaks of CDHA appeared only after exposing for 6 h (Fig. 6d). Another distinguishable difference in morphology was the reduced crystal lengths compared to FSS-free group. Besides, HRTEM characterization revealed the various crystal orientations (Fig. 6c). All these evidences exhibited the prevention of 2.0 Pa FSS on the crystallization of ACP.
3.3. Mechanism of FSS-mediated conversion of ACP to CDHA The amorphous-to-crystalline transformation of ACP might proceed along several pathways, including dissolution and reprecipitation (Somrani et al., 2005), internal structural rearrangements (Onuma, 2006), or formation of crystalline phases directly within
Fig. 5 – Morphologies and crystal characteristics of ACP to CDHA conversion under 1.5 Pa FSS environment. (a–c) TEM characterization after aging for 3, 6, and 9 h, respectively, with SAED pattern in the top right corner of each image. (d) HRTEM result after aging for 9 h. (e) XRD patterns after aging for 0, 3, 6, and 9 h.
journal of the mechanical behavior of biomedical materials 54 (2016) 131 –140
137
Fig. 6 – Morphologies and crystal characteristics of ACP to CDHA conversion under 2.0 Pa FSS environment. (a, b) TEM characterization after aging for 3 and 6 h, with SAED pattern in the top right corner of each image. (c) HRTEM result after aging for 6 h. (d) XRD patterns after aging for 0, 3, and 6 h.
Fig. 7 – Schematic illustration of a hypothesized mechanism for the crystallization of ACP to CDHA under FSS environment. A moderate external FSS could accelerate local diffusion and rearrangement of calcium and orthophosphate ions in an ACP system, which made the formed CDHA crystallites presented the well-organized structure and elongated morphology with certain orientation.
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journal of the mechanical behavior of biomedical materials 54 (2016) 131 –140
Table 1 – Ca/P mole ratio analysis of CDHA prepared under different conditions by ICP-MS characterization. FSS (Pa)
Exposing time (h)
Ca/P mole ratio
0 0.5 0.5 1.0 1.5 2.0 0
0 3 6 6 6 6 6
1.48 1.53 1.63 1.66 1.65 1.67 1.60
Fig. 8 – Biocompatibility of CDHA prepared under FSS and FSS-free environments. (a) The proliferation of MC3T3-E1 cells after co-culturing for 1, 4 and 7 d with CDHA prepared under 1.0 Pa FSS for 9 h and FSS-free environments. (b) The ALP activity of MC3T3-E1 cells after co-culturing for 7 and 14 d with CDHA prepared under 1.0 Pa FSS for 9 h and FSS-free environments.
ACP (Tao et al., 2009; Wang et al., 2009). Although the roles of FSS in this process remains not been well elucidated, Fig. 7 schematically illustrates a hypothesized mechanism for the transformation of ACP to CDHA under FSS environment based on the previous and the present researches. In Fig. 7a, calcium and orthophosphate ions formed pairs and clusters and further composed initial solid phase, which was originally amorphous in nature. At multiple sites within a primary particle, crystalline domains gradually developed. Since the growth of crystalline domains consumed the surrounding ion pairs and clusters, the mechanical strength decreased in the inter-domain regions. Under the action of external FSS, the primary particles collapsed and released many separated crystallites. These liberated crystallites could trigger the rapid ACP crystallization to form elongated needle-like crystals following the direction of FSS. Fig. 7b further illustrates the details of particle collapse and ions rearrangement when they were exposed in FSS. During the stage of crystalline domains development, the primary particle took up calcium ions and released protons. Once it was collapsed, various calcium and orthophosphate ions need to be consumed for the following ions rearrangement and nucleation. A moderate FSS could increase the local diffusivity of calcium and orthophosphate ions in the system, hence having a positive influence on this process. This inference could also be confirmed from Ca/P mole ratio analysis. In Table 1, Ca/P ratio increased from 1.48 to 1.63 with extending aging time from 0 to 6 h under 0.5 Pa FSS. Similarly, under the same aging time (6 h), all 4 FSS groups
exhibited the higher Ca/P mole ratios than the FSS-free group. With enrichment of calcium ions, the crystallites presented well-organized structure and orientation. On the other hand, although a high FSS could also accelerate ionic spread and suppress appearance of large clusters, it might destroy crystalline nuclei as well and thereby resulted in the defective crystalline structure.
3.4.
Biocompatibility evaluation
The biocompatibility of CDHA prepared under FSS environment was evaluated by using pre-osteoblast MC3T3-E1 cells. Fig. 8a shows the proliferation behaviors of MC3T3-E1 cells co-cultured with CDHA prepared under 1.0 Pa FSS for 9 h and FSS-free mediums. The cell numbers were increased in both groups with the culturing time being prolonged from 1 to 4 d. Further extending time to 7 d, the cell numbers decreased in both groups, which revealed that the MC3T3-E1 cells began to differentiation. More importantly, during 7 d of culture, the FSS group always presented the higher cell number than the FSS-free group (po0.05). The ALP contents of cells in FSS group were also significantly higher than that in FSS-free group (po0.05) after 7 and 14 d of culture (Fig. 8b), confirming that the well-organized crystal structure of CDHA prepared under FSS environment enhances osteoblastic differentiation of the cells.
journal of the mechanical behavior of biomedical materials 54 (2016) 131 –140
4.
Conclusions
FSS influences the conversion of ACP precursor and the formed crystal structure of CDHA. Under the low FSS ( r1.0 Pa), it has positive effects on the crystallization of ACP, characterized by the accelerated speed and wellorganized CDHA structure. Under the high FSS (41.0 Pa), it has negative effects on this conversion process, characterized by the poor CDHA crystal morphologies and destroyed structures. The better CDHA structure prepared under low FSS environment further presents the more biocompatible features with pre-osteoblast cells and provides a new candidate for bone regeneration.
Acknowledgments This research was financially supported by the National Natural Science Foundation of China (Nos. 11272038, 31470915 and 11421202), the National Science and Technology Pillar Program of China (Nos. 2012BAI18B01, 2014BAI 11B02 and 2014BAI11B03), the Fundamental Research Funds for the Central Universities (No. YWF-15-YG-021), Pearl River S&T Nova Program of Guangzhou (No. 2013J2200010), State Key Laboratory of New Ceramic and Fine Processing Tsinghua University, the 111 Project (No. B13003), and the International Joint Research Center of Aerospace Biotechnology and Medical Engineering, Ministry of Science and Technology of China.
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Nganga, S., Zhang, D., Moritz, N., Vallittu, P.K., Hupa, L., 2012. Multi-layer porous fiber-reinforced composites for implants: in vitro calcium phosphate formation in the presence of bioactive glass. Dent. Mater. 28, 34–45. Niu, X., Feng, Q., Wang, M., Guo, X., Zheng, Q., 2009. Porous nanoHA/collagen/ PLLA scaffold containing chitosan microspheres for controlled delivery of synthetic peptide derived from BMP2. J. Control. Release 134, 111–117. Olszta, M.J., Cheng, X.G., Jee, S.S., Kumar, R., Kim, Y.Y., Kaufman, M.J., Douglas, E.P., Gower, L.B., 2007. Bone structure and formation: a new perspective. Mater. Sci. Eng. R 58, 77–116. Onuma, K., 2006. Recent research on pseudobiological hydroxyapatite crystal growth and phase transition mechanisms. Prog. Cryst. Growth Charact. Mater. 52, 223–245. Pasteris, J.D., Yoder, C.H., Sternlieb, M.P., Liu, S., 2012. Effect of carbonate incorporation on the hydroxyl content of hydroxylapatite. Mineral. Mag. 76, 2741–2759. Pasteris, J.D., Yoder, C.H., Wopenka, B., 2014. Molecular water in nominally unhydrated carbonated hydroxylapatite: the key to a better understanding of bone mineral. Am. Mineral. 99, 16–27. Ponik, S.M., Triplett, J.W., Pavalko, F.M., 2007. Osteoblasts and osteocytes respond differently to oscillatory and unidirectional fluid flow profiles. J. Cell. Biochem. 100, 794–807. Seeman, E., Delmas, P.D., 2006. Bone quality-the material and structural basis of bone strength and fragility. N. Engl. J. Med. 354, 2250–2261. Somrani, S., Banu, M., Jemal, M., Rey, C., 2005. Physico-chemical and thermochemical studies of the hydrolytic conversion of amorphous tricalcium phosphate into apatite. J. Solid State Chem. 178, 1337–1348.
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Research paper
Shear-mediated crystallization from amorphous calcium phosphate to bone apatite Xufeng Niua,b,c,n, Liyang Wanga, Feng Tiana, Lizhen Wanga, Ping Lia, Qingling Fengd, Yubo Fana,nn a Key Laboratory for Biomechanics and Mechanobiology of Ministry of Education, School of Biological Science and Medical Engineering, Beihang University, Beijing 100191, China b BUAA Research Institute, Guangzhou 510530, China c Research Institute of Beihang University in Shenzhen, Shenzhen 518057, China d State Key Laboratory of New Ceramic and Fine Processing, Tsinghua University, Beijing 100084, China
art i cle i nfo
ab st rac t
Article history:
The contribution of fluid shear stress (FSS) on the conversion of amorphous calcium
Received 31 May 2015
phosphate (ACP) to bone apatite is investigated. The ACP precursors are prepared by using
Received in revised form
a wet-chemistry method and further exposed to the constant FSS environment with values
13 September 2015
of 0.5, 1.0, 1.5, and 2.0 Pa. At the designated time points, the apatites are characterized by
Accepted 21 September 2015
transmission electron microscopy, X-ray diffraction, and inductively coupled plasma-mass
Available online 30 September 2015
spectroscopy. The results show that, the low FSS ( r1.0 Pa) has positive effects on the
Keywords:
transition of ACP, characterized by the accelerated crystallization velocity and the well-
Fluid shear stress
organized calcium-deficient hydroxyapatite (CDHA) structure, whereas the high FSS
Amorphous calcium phosphate
(41.0 Pa) has negative effects on this conversion process, characterized by the poor CDHA
Apatite
crystal morphologies and the destroyed structures. The bioactivity evaluations further
Crystallization
reveal that, compared with the FSS-free group, the CDHA prepared under 1.0 Pa FSS for 9 h
Cone-and-plate viscometer
presents the more biocompatible features with pre-osteoblast cells. These results are helpful for understanding the mechanism of apatite deposition in natural bone tissue. & 2015 Elsevier Ltd. All rights reserved.
1.
Introduction
dynamic tissue capable of mechanical load-induced structural and functional adaptation through the process of bone remodel-
Beginning with the pioneering work of Julius Wolff in the 19th
ing (Knothe Tate et al., 2011; Martínez-Reina et al., 2014; Weiner
century, it is well-recognized that the external mechanical
and Wagner, 1998). At cellular and molecular levels, mounting
stimulus plays a dominant role in bone formation (Chen et al.,
evidences suggest that both bone cells and interstitial fluid flow
2010; Frost, 2000). According to the present knowledge, bone is a
within the lacunar–canalicular system are able to sense and
n Corresponding author at: Key Laboratory for Biomechanics and Mechanobiology of Ministry of Education, School of Biological Science and Medical Engineering, Beihang University, Beijing 100191, China. Tel.: þ86 10 82338755; fax: þ86 10 82315554. nn Corresponding author. Tel./fax: þ86 10 82339428. E-mail addresses: [email protected] (X. Niu), [email protected] (Y. Fan).
http://dx.doi.org/10.1016/j.jmbbm.2015.09.024 1751-6161/& 2015 Elsevier Ltd. All rights reserved.
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journal of the mechanical behavior of biomedical materials 54 (2016) 131 –140
respond to the external mechanical load (Ciani et al., 2009; Han et al., 2004; Huo et al., 2010; Ponik et al., 2007). Bone cells, including osteocytes, osteoblasts, and osteoclasts, are the broadly investigated cells as mechanosensors during the process of bone remodeling (Khosla et al., 2008). These cells compose basic multicellular unit (BMU) within the bone remodeling compartment, where osteocytes are imprisoned in bone matrix whereas osteoclasts and osteoblasts are located in the surface (Eriksen et al., 2007; Hauge et al., 2001). In each BMU, osteocytes are the primary mechanosensor which can sense mechanical strain and trigger bone remodeling (Seeman and Delmas, 2006). Osteoclasts and osteoblasts are the performers of bone anabolism, in which the former can induce bone absorption while the latter can enhance bone formation (Ducy et al., 2000; Teitelbaum, 2000). Signals generated by osteocytes deep within the BMU reach the surface and elicit remodeling events by osteoclasts and osteoblasts. To respond to the external mechanical stimuli timely and accurately, these cells achieve the crosstalk through cell–cell contacts or through signal molecules (Canalis et al., 2013; Haugh et al., 2015; Malone et al., 2007). Besides bone cells, bone matrix is another structural component in natural bone tissue. Apatite is the main inorganic phase in bone matrix, which endows bone with superior mechanical properties. Bone apatite is composed of calcium phosphate-based minerals, predominantly by hydroxyapatite (HA) (Huang et al., 2014; Niu et al., 2009; Zhou et al., 2015). Previously, many researches were focused on the crystallization behavior during calcium phosphate formation (Massera et al., 2015; Nganga et al., 2012). One common view was that bone apatite came from amorphous calcium phosphate (ACP) (Boonrungsiman et al., 2012; Liu et al., 2011; Mahamid et al., 2010). To obtain the desired HA structure in vitro, the conversion of ACP was simulated in aqueous media and various influencing factors were investigated, including pH, temperature, and presence of foreign ions (Combes and Rey, 2010; Yoder et al., 2012a; Mccubbin et al., 2008). The results revealed that the presence of carbonate (Pasteris et al., 2012; Mason et al., 2009) and molecular H2O (Pasteris et al., 2014; Yoder et al., 2012b) groups played a critical role in stabilizing the structure of apatite. However, thinking that bone has the mechanical adaptation and interstitial fluid flow can also act on bone matrix, it is rational to deduce that the external biomechanical stimulus may have influence on the conversion of ACP. Regrettably, it was always neglected in the previous study. Among the complicated mechanical forces generated by external loading, fluid shear stress (FSS) is regarded as the predominant stimulus to bone cells and matrix under the physiological conditions and at the micro-level (Fritton and Weinbaum, 2009; Li et al., 2012). Moreover, in most conversion reactions of ACP occurring at physiological pH, the obtained apatite is calcium-deficient hydroxyapatite (CDHA). Therefore, in this paper, ACP precursor was prepared and further exposed to constant FSS environment. The evolution of ACP to CDHA was investigated by transmission electron microscopy (TEM) and X-ray diffraction (XRD), with the purpose of illuminating the effects of FSS in this conversion process.
2.
Materials and methods
2.1.
Preparation of ACP precursor
A wet chemistry method was used to get the ACP precursor. Two kinds of solution, 12.5 mmol of calcium nitrate tetrahydrate and 7.5 mmol of ammonium phosphate, were prepared individually with volume of 20 mL in distilled water. Calcium ions solution was then gradually dropped into orthophosphate ions solution to form a mixture, which contained calcium and orthophosphate ions with Ca/P mole ratio of 1.67 (corresponding to Ca/P mole ratio in HA (Dorozhkin, 2007; Zhang et al., 2003)). Aqua ammonia was used to adjust the pH to 7.4 during the dropping process. The obtained ACP precipitation was brought to the next procedure right away.
2.2.
Exposure of shear stress
Cone-and-plate viscometer was used to provide constant FSS environment in this research. The ACP precursor was shifted to Brookfield R/S-CPSþ rheometer (USA) and exposed individually to FSS with values of 0.5, 1.0, 1.5, 2.0 Pa for aging 3, 6, and 9 h, respectively. The obtained precipitation was then washed 3 times with distilled water and further lyophilized using the SP Scientific VirTis Advantage XL-70 freeze dryers (USA) to gain CDHA powders. The control, FSS-free CDHA sample, was prepared using the same procedure except biomechanical exposure.
2.3.
Characterization
To get the morphological properties, the CDHA powders were observed by JEOL JEM-2100F TEM (Japan). The samples were dispersed in ethanol and ultrasonicated for 30 min. Then, they were picked up with carbon coated copper grid (200 mesh) for TEM characterization. To acquire the information about crystalline structure, selected area electron diffraction (SAED) was used and XRD patterns of samples were also carried out at room temperature on Rigaku D/Max X-ray diffractometer (Japan) with Cu Kα radiation source (wavelength¼ 1.54 Å). The supplied voltage and current were set to 40 kV and 120 mA, respectively. The samples were exposed at a scan rate of 101/min from 31 to 701. To identify the composition of CDHA, the powders were further dissolved in 8% nitric acid and the Ca/P mole ratios were measured by Varian Vista MPX inductively coupled plasma mass spectroscopy (ICP-MS, USA).
2.4.
In vitro bioactivity assay
2.4.1.
Cell culture and CDHA sterilization
The thawed mouse calvaria-derived preosteoblastic cells MC3T3-E1 (American type culture collection) were incubated in α-MEM supplemented with 10% fetal bovine serum (FBS), 100 U/mL penicillin, and 100 μg/mL streptomycin in a humidified incubator at 37 1C with 5% CO2 (Thermo Fisher Scientific, China). The medium was changed every other day. Two types of CDHA powders, prepared under 1.0 Pa FSS or FSS-free environment for 9 h, were sterilized by exposing in ultraviolet for 1 h, and further used for cell seeding and evaluation.
journal of the mechanical behavior of biomedical materials 54 (2016) 131 –140
2.4.
2 Proliferation assay
133
10% FBS. The CDHA powders were added at a density of
The MC3T3-E1 cells were seeded in 96-well tissue culture
300 μg/mL based on α-MEM solution. Cell counting kit-8 (CCK-
plate (2.5 103 cells/well) containing α-MEM medium plus
8) assay was carried on days 1, 4, and 7 after cell seeding. Cell
Fig. 1 – Schematic representation of FSS loading by cone-and-plate viscometer during the process of CDHA preparation. ACP precursor was synthesized using the wet chemistry method and further exposed to various FSS from 0.5 to 2.0 Pa. The obtained CDHA was separated at designated aging time points.
Fig. 2 – Morphologies and crystal characteristics of ACP to CDHA conversion under FSS-free environment. (a–e) TEM characterization after aging for 0, 3, 6, 9 and 24 h, respectively, with SAED pattern in the top right corner of each image. (f) XRD patterns after aging for 0, 3, 6, 9 and 24 h.
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journal of the mechanical behavior of biomedical materials 54 (2016) 131 –140
proliferation was measured following the CCK-8 assay kit (Dojindo, Japan) instructions. Briefly, 10 μL CCK-8 kit was added to each well and then the plate was incubated at 37 1C for 1.5 h. The absorbance was read at 450 nm in Varioskan Flash (Thermo Scientific, USA). All experiments were repeated in triplicate and each group had three replicates.
2.4.3.
Alkaline phosphatase (ALP) activity assay
For ALP assay, the MC3T3-E1 cells were co-cultured with 300 μg/mL CDHA powders in α-MEM medium plus 10% FBS for 7 and 14 d, respectively. Afterwards, the medium was removed. ; The cells with number of 1 105 were transferred to microcentrifuge tube and treated with RIPA (Applygen, China) at 4 1C for 10 min. The lysates were clarified by centrifugation. The supernatant was collected for ALP assay by measuring the release of p-nitrophenol at 405 nm using the ALP activity kit (Zhongsheng, China). The amount of ALP in the cells was calculated according to the formula provided by the kit.
2.4.4.
Statistical analysis
All experiments were conducted at least three times. The data were expressed as mean7standard deviation. Statistical analysis was performed using one-way analysis of variance. The difference was regarded as statistical significance when po0.05.
3.
Results and discussion
3.1.
Preparation of CDHA under FSS environment
Bone tissue is exposed to the complicated biomechanical environment. Previous theoretical studies have demonstrated that interstitial fluid in bone tissue can induce shear stress on the order of 0.8–3.0 Pa in vivo (Weinbaum et al., 1994). According to these results, FSS used in this research was set as 0.5, 1.0, 1.5 and 2.0 Pa. Moreover, just as shown in Fig. 1, cone-and-plate viscometer was introduced to perform FSS loading, which could supply homogeneous biomechanical environment towards the conversion of ACP to CDHA during the whole experimental procedure.
3.2. Crystallization of ACP to CDHA under different FSS environment Fig. 2 shows the conversion of ACP to CDHA at various aging times under FSS-free environment, with morphological transformations and SAED patterns shown at the top right corner in Fig. 2a–e and XRD patterns shown in Fig. 2f. Fig. 2a shows the spherical precursor, which could be defined as ACP, since this unstable calcium phosphate status had a typical diffraction
Fig. 3 – Morphologies and crystal characteristics of ACP to CDHA conversion under 0.5 Pa FSS environment. (a–c) TEM characterization after aging for 3, 6, and 9 h, respectively, with SAED pattern in the top right corner of each image. (d) HRTEM result after aging for 9 h. (e) XRD patterns after aging for 0, 3, 6, and 9 h.
journal of the mechanical behavior of biomedical materials 54 (2016) 131 –140
135
pattern of amorphous halo ring (Dorozhkin, 2010). With aging for
was almost accomplished. Such results were coincident with
3 h, the precursor became curly but still did not crystallize well (Fig. 2b). After aging for 6 h, the tiny sticks appeared with the length of 50–150 nm roughly, under which might be the platelets of octacalcium phosphate or not well crystallized HA (Fig. 2c). At the same time, the diffraction pattern changed from diffused
the previous reports (Dorozhkin, 2009; Kim et al., 2005). Therefore, to fully investigate the influence of biomechanical exposure on crystal formation and growth, the maximum FSS duration was chosen to 9 h in the following study. Fig. 3 shows the conversion of ACP to CDHA after exposing
rings to polycrystalline rings. Further extending aging time to 9 h led to the formation of needle-like or stick-like crystallites (Fig. 2d). Besides, the crystal morphologies in aging for 24 h group were similar to that in aging for 9 h group, and the series
to 0.5 Pa FSS for various aging times up to 9 h, with morphological transformations and SAED patterns shown at the top right corner in Fig. 3a–c and XRD patterns shown in Fig. 3e. Compared to the corresponding FSS-free groups, some distinct features could be distinguished. With exposing for 3 h
of rings in SAED pattern resulted in the almost accomplished crystallization (Fig. 2e). In XRD patterns (Fig. 2f), the curve was broad and flat for ACP group, which was the symbol of amorphous status. Then some main peaks of CDHA appeared after aging for 3 h, such as (002)
(Fig. 3a), the spherical ACP precursors elongated and amorphous halo rings disappeared. After exposing for 6 h (Fig. 3b), some needle-like or stick-like crystallites formed, which looked like thinner and longer compared to Fig. 2c. The
near 25.81, (211) near 31.81. The peak (100) at 10.81 and three peaks of (222) at 46.71, (213) at 49.51 and (004) at 53.21 arose gradually after aging for 6 and 9 h, which were the typical XRD pattern of CDHA and all these diffraction peaks could be
diffraction rings corresponding to (002) and (004) suggested that the crystallites already possessed the typical structure of CDHA. Further extending time to 9 h revealed some arcs consisted of rings (Fig. 3c), which were the character of
assigned to the standard card. Further prolonging time to 24 h did not bring about extra variations of XRD pattern. Based on these results, it was rational to infer that the crystallization of ACP to CDHA in the wet chemistry method occurred mostly within 9 h. After that, the crystal transition
crystalline structure having certain orientation. In XRD patterns (Fig. 3e), the typical peaks of CDHA already emerged after aging for 3 h, which included (002), (211), (222), (213) and (004). Further prolonging aging time to 6 and 9 h just increased peak intensities. These results meant
Fig. 4 – Morphologies and crystal characteristics of ACP to CDHA conversion under 1.0 Pa FSS environment. (a–c) TEM characterization after aging for 3, 6, and 9 h, with SAED pattern in the top right corner of each image. (d) HRTEM result after aging for 9 h. (e) XRD patterns after aging for 0, 3, 6, and 9 h.
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that 0.5 Pa FSS could promote transition of ACP towards CDHA. High resolution TEM (HRTEM) was another evidence of well-organized CDHA structure after exposing to 0.5 Pa FSS, since the d-spacing of 0.25 nm measured from Fig. 3d was coincident with one of d-spacings of HA (Olszta et al., 2007). Fig. 4 shows the conversion of ACP to CDHA after exposing to 1.0 Pa FSS for various aging times up to 9 h, which shows the similar results as shown in Fig. 3. Both TEM and XRD characterizations revealed the accelerated conversion of ACP precursors towards CDHA in comparison with FSS-free group. The formed CDHA crystallites had the trends to be thinner and longer. They also presented the well-organized crystal structure and homogenous orientation, characterized by crystal lattices arranged in order with the distance of 0.23 nm for (310) and 0.33 nm for (002) (Fig. 4d), both were d-spacings of HA (Olszta et al., 2007). All these features demonstrated the positive effects of 1.0 Pa FSS on the crystallization of ACP. Fig. 5 shows the conversion of ACP to CDHA after exposing to 1.5 Pa FSS for various aging times up to 9 h. When exposing for 3 h, both TEM and XRD characterizations indicated the accelerated crystallization process. As a result, Fig. 5a presents the typical thin and straight needle-like CDHA crystallites. However, after exposing for 6 and even 9 h (Fig. 5b and c), TEM characterization revealed the poorer crystal morphologies as compared with the corresponding groups in FSS-free
environment (Fig. 2c and d). HRTEM also illustrated the different d-spacings at various districts in an individual crystallite (Fig. 5d), which was different from the results shown in Figs. 3d and 4d and meant that the inner structure of CDHA crystallites might not be benefitted from the increased FSS. Fig. 6 further shows the conversion of ACP to CDHA after exposing to 2.0 Pa FSS for 3 and 6 h. Further extending aging time to 9 h led to the absence of any CDHA crystals (data not shown). The imperfect SAED patterns shown in Fig. 6a and b suggested the slowed crystallizing speed. XRD patterns also indicated that the crystallization process was greatly decreased, since the representative peaks of CDHA appeared only after exposing for 6 h (Fig. 6d). Another distinguishable difference in morphology was the reduced crystal lengths compared to FSS-free group. Besides, HRTEM characterization revealed the various crystal orientations (Fig. 6c). All these evidences exhibited the prevention of 2.0 Pa FSS on the crystallization of ACP.
3.3. Mechanism of FSS-mediated conversion of ACP to CDHA The amorphous-to-crystalline transformation of ACP might proceed along several pathways, including dissolution and reprecipitation (Somrani et al., 2005), internal structural rearrangements (Onuma, 2006), or formation of crystalline phases directly within
Fig. 5 – Morphologies and crystal characteristics of ACP to CDHA conversion under 1.5 Pa FSS environment. (a–c) TEM characterization after aging for 3, 6, and 9 h, respectively, with SAED pattern in the top right corner of each image. (d) HRTEM result after aging for 9 h. (e) XRD patterns after aging for 0, 3, 6, and 9 h.
journal of the mechanical behavior of biomedical materials 54 (2016) 131 –140
137
Fig. 6 – Morphologies and crystal characteristics of ACP to CDHA conversion under 2.0 Pa FSS environment. (a, b) TEM characterization after aging for 3 and 6 h, with SAED pattern in the top right corner of each image. (c) HRTEM result after aging for 6 h. (d) XRD patterns after aging for 0, 3, and 6 h.
Fig. 7 – Schematic illustration of a hypothesized mechanism for the crystallization of ACP to CDHA under FSS environment. A moderate external FSS could accelerate local diffusion and rearrangement of calcium and orthophosphate ions in an ACP system, which made the formed CDHA crystallites presented the well-organized structure and elongated morphology with certain orientation.
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Table 1 – Ca/P mole ratio analysis of CDHA prepared under different conditions by ICP-MS characterization. FSS (Pa)
Exposing time (h)
Ca/P mole ratio
0 0.5 0.5 1.0 1.5 2.0 0
0 3 6 6 6 6 6
1.48 1.53 1.63 1.66 1.65 1.67 1.60
Fig. 8 – Biocompatibility of CDHA prepared under FSS and FSS-free environments. (a) The proliferation of MC3T3-E1 cells after co-culturing for 1, 4 and 7 d with CDHA prepared under 1.0 Pa FSS for 9 h and FSS-free environments. (b) The ALP activity of MC3T3-E1 cells after co-culturing for 7 and 14 d with CDHA prepared under 1.0 Pa FSS for 9 h and FSS-free environments.
ACP (Tao et al., 2009; Wang et al., 2009). Although the roles of FSS in this process remains not been well elucidated, Fig. 7 schematically illustrates a hypothesized mechanism for the transformation of ACP to CDHA under FSS environment based on the previous and the present researches. In Fig. 7a, calcium and orthophosphate ions formed pairs and clusters and further composed initial solid phase, which was originally amorphous in nature. At multiple sites within a primary particle, crystalline domains gradually developed. Since the growth of crystalline domains consumed the surrounding ion pairs and clusters, the mechanical strength decreased in the inter-domain regions. Under the action of external FSS, the primary particles collapsed and released many separated crystallites. These liberated crystallites could trigger the rapid ACP crystallization to form elongated needle-like crystals following the direction of FSS. Fig. 7b further illustrates the details of particle collapse and ions rearrangement when they were exposed in FSS. During the stage of crystalline domains development, the primary particle took up calcium ions and released protons. Once it was collapsed, various calcium and orthophosphate ions need to be consumed for the following ions rearrangement and nucleation. A moderate FSS could increase the local diffusivity of calcium and orthophosphate ions in the system, hence having a positive influence on this process. This inference could also be confirmed from Ca/P mole ratio analysis. In Table 1, Ca/P ratio increased from 1.48 to 1.63 with extending aging time from 0 to 6 h under 0.5 Pa FSS. Similarly, under the same aging time (6 h), all 4 FSS groups
exhibited the higher Ca/P mole ratios than the FSS-free group. With enrichment of calcium ions, the crystallites presented well-organized structure and orientation. On the other hand, although a high FSS could also accelerate ionic spread and suppress appearance of large clusters, it might destroy crystalline nuclei as well and thereby resulted in the defective crystalline structure.
3.4.
Biocompatibility evaluation
The biocompatibility of CDHA prepared under FSS environment was evaluated by using pre-osteoblast MC3T3-E1 cells. Fig. 8a shows the proliferation behaviors of MC3T3-E1 cells co-cultured with CDHA prepared under 1.0 Pa FSS for 9 h and FSS-free mediums. The cell numbers were increased in both groups with the culturing time being prolonged from 1 to 4 d. Further extending time to 7 d, the cell numbers decreased in both groups, which revealed that the MC3T3-E1 cells began to differentiation. More importantly, during 7 d of culture, the FSS group always presented the higher cell number than the FSS-free group (po0.05). The ALP contents of cells in FSS group were also significantly higher than that in FSS-free group (po0.05) after 7 and 14 d of culture (Fig. 8b), confirming that the well-organized crystal structure of CDHA prepared under FSS environment enhances osteoblastic differentiation of the cells.
journal of the mechanical behavior of biomedical materials 54 (2016) 131 –140
4.
Conclusions
FSS influences the conversion of ACP precursor and the formed crystal structure of CDHA. Under the low FSS ( r1.0 Pa), it has positive effects on the crystallization of ACP, characterized by the accelerated speed and wellorganized CDHA structure. Under the high FSS (41.0 Pa), it has negative effects on this conversion process, characterized by the poor CDHA crystal morphologies and destroyed structures. The better CDHA structure prepared under low FSS environment further presents the more biocompatible features with pre-osteoblast cells and provides a new candidate for bone regeneration.
Acknowledgments This research was financially supported by the National Natural Science Foundation of China (Nos. 11272038, 31470915 and 11421202), the National Science and Technology Pillar Program of China (Nos. 2012BAI18B01, 2014BAI 11B02 and 2014BAI11B03), the Fundamental Research Funds for the Central Universities (No. YWF-15-YG-021), Pearl River S&T Nova Program of Guangzhou (No. 2013J2200010), State Key Laboratory of New Ceramic and Fine Processing Tsinghua University, the 111 Project (No. B13003), and the International Joint Research Center of Aerospace Biotechnology and Medical Engineering, Ministry of Science and Technology of China.
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