Effects of Three-Dimensional Culture of Mouse Calvaria-Derived

Oct 2, 2017 - †Graduate School of Life Science, ‡Faculty of Advanced Life Science, and §Global Station for Soft Matter, Global Institution for Co...
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Effects of Three-Dimensional Culture of Mouse Calvaria-Derived Osteoblastic Cells in a Collagen Gel with a Multichannel Structure on the Morphogenesis Behaviors of Engineered Bone Tissues Saki Yahata,† Kazuya Furusawa,*,‡,§ Kei Nagao,∥ Masahiro Nakajima,⊥ and Toshio Fukuda#,△ †

Graduate School of Life Science, ‡Faculty of Advanced Life Science, and §Global Station for Soft Matter, Global Institution for Collaborative Research and Education, Hokkaido University, Kita 10 Nishi 8, Kita-ku, Sapporo 060-0810, Japan ∥ Department of Micro-Nano Systems Engineering and ⊥Center for Micro-nano Mechatronics, Nagoya University, Furo-cho, Chikusa-ku, Nagoya 464-8603, Japan # Department of Mechatronics Engineering, Meijo University, 1-501, Shiogamaguchi, Tempaku-ku, Nagoya 468-8502, Japan △ Intelligent Robotics Institute, Beijing Institute of Technology, 5 South Zhongguancun Street, Haidian District, Beijing 100081, China S Supporting Information *

ABSTRACT: Bone has a complex hierarchical structure that contributes to its superior mechanical properties. Therefore, reproducing the complex hierarchical structure of bone tissue is a promising strategy to construct functional engineered bone tissues. In this study, we aimed to reproduce this complex hierarchical structure by developing a method for the threedimensional culture of MC3T3-E1 osteoblastic cells in a collagen gel with a multichannel structure (MCCG), which mimics the parallel arrangement of Haversian canals in bone tissue. MCCG was homogeneously calcified via the biomineralization properties of MC3T3-E1s. Confocal laser scanning microscopy revealed that MCCG could support the growth and proliferation of MC3T3-E1 cells in the deeper parts of the engineered bone tissue and that the cells formed a toroidal structure on the channel surface and a network-like structure in the gel matrix region. Furthermore, quasi-quantitative measurement of osteocalcin and dentin matrix protein 1 expression indicated the coexistence of two types of cells with different morphologies and differentiation phenotypes. Thus, three-dimensional culture of MC3T3-E1 cells in MCCG yielded engineered tissues mimicking the hierarchical structures of bone tissues. Engineered bone tissues with a biomimetic hierarchical structure could be used as a model system for investigating bone metabolism and evaluating the efficacy of novel drugs. KEYWORDS: tissue engineering, bone hierarchical structure, Haversian canal, collagen gel, multichannel structure, morphogenesis of engineered tissue

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INTRODUCTION

tissue formation during the past century may be helpful for reproducing the complex hierarchical structure of bone tissues in vitro. Tissue engineering techniques comprise promising methodologies for constructing biomimetic engineered bone tissues and various technologies to reconstruct the engineered bone tissues with complex hierarchical structures in vitro have been developed.5−10 Many reports have suggested that bone cells should be cultured in a three-dimensional (3D) environment to construct tissues mimicking bone tissues in vitro. Therefore, various cellular scaffolds have been developed for the 3D culture of bone cells. For example, Cunniffe et al. developed collagen-nHA biocomposite scaffolds and collagen sponges as

Bone supports the weight of our bodies to combat the effects of gravity. The superior mechanical properties of bone are attributed to its complex hierarchical structure.1 Long bones, such as the femur and tibia, consist of compact bone tissues with high bone density and cancellous bone tissues with low bone density. The compact bone tissues consist of many osteons, i.e., metabolic units of bone, which contain concentric multilayered bone matrices and a central lumen, termed the Haversian canal. Osteoblasts and osteoclasts adhere on the interior surface of the Haversian canals, and osteocytes are embedded into the lacunae.2 These bone cells make up the complex hierarchical structure of compact bone tissues during the development process and maintain the mechanical properties of the bone in response to mechanical stimulation.3,4 Such details arising from the extensive investigations into the molecular biological and biochemical mechanisms of bone © XXXX American Chemical Society

Received: March 27, 2017 Accepted: October 2, 2017 Published: October 2, 2017 A

DOI: 10.1021/acsbiomaterials.7b00190 ACS Biomater. Sci. Eng. XXXX, XXX, XXX−XXX

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ACS Biomaterials Science & Engineering

Figure 1. Schematic illustration of the preparation of collagen gel using a dialysis chamber.

scaffolds with suitable mechanical and biological properties.6 Murphy et al. fabricated collagen scaffolds with suitable pore sizes for cells.7 In addition, many researchers have focused on constructing 3D-engineered bone tissues by culturing bone cells on 3D scaffolds. In particular, cellular scaffolds consisting of biodegradable polymers, such as polycaprolactone (PCL) and poly(D,L-lactide-co-glycolide) (PLGA), have been used for constructing 3D-engineered bone tissues.9,10 Although various techniques have been established for the culture of bone cells in 3D environments, these techniques have not been able to reproduce the complex hierarchical structure of bone tissue. In particular, the construction of osteon-like structures has been challenging.11−13 Although the morphologies of the constructed tissues mimic or resemble the osteon, they still differ from the original osteon and do not have the ability to reproduce the hierarchical structures of compact bone tissue. Recently, a direct construction method of the anatomic bone model reproducing macroscopic bone morphology using 3D-printing technology has been reported.14 The anatomic bone model is designed to reproduce the mechano-mimetic structures of bone tissue. Therefore, the biomaterial could be used as a template cellular scaffold for constructing biomimetic engineered bone tissues. However, it is difficult to embed the bone cells into the matrix region of the biomaterial during the 3D-printing process. Moreover, many of these constructed tissues and biomaterials consist of nonbone matrix polymers, such as synthetic polymers and other biopolymers. Biomimetic cellular scaffolds that can be used to three-dimensionally culture the bone cells are required to resolve this issue. The materials that mimic the complex hierarchical structure of bone tissue play an important role in the fields of regenerative medicine and tissue engineering. In particular, hydrogels with unidirectionally aligned transport channels (multichannel hydrogels [MCGs]) may be promising candidate cellular scaffolds for constructing engineered bone tissues with native hierarchical structures. MCGs have been prepared by dialysis-induced anisotropic gelation.15−17 Alginate MCGs prepared in this manner15 and composites of alginate MCGs with hydroxyl apatite have been used as template scaffolds for constructing engineered bone tissues18 and blood vessel-like structures.19 However, the cells adhere on the surface of the channel but are not embedded in the gel matrix region; these techniques therefore do not provide the essential 3D culture conditions required for appropriate bone tissue formation. In our previous studies, we have found that dialysis of a collagen aqueous solution against phosphate-buffer solution (PBg) results in collagen hydrogels with multichannel structures (multichannel collagen gels [MCCGs]).17 The multichannel structure mimics the parallel arrangements of Haversian canals in compact bone. Furthermore, the collagen fibers, which constitute the building blocks of collagen

hydrogels, align parallel to the surface of the MCCG channels. Therefore, MCCG mimics the complex hierarchical structure of compact bone tissues from the level of the collagen molecule to the parallel arrangements of Haversian canals. By using this biomimetic multichannel structure, we have constructed engineered bone tissues by seeding MC3T3-E1 cells on the surface of MCCGs.20 In our previous work, we found that MC3T3-E1 cells form a toroidal-like structure on the channel surface of MCCGs after being seeded on the top surface of MCCGs for 21 days. However, our previous findings showed that almost all cells preferentially invaded into the inner channel region, whereas there were no cells in the gel matrix region of MCCGs. Recently, we have established a method to culture cells by embedding cells directly into the collagen gel matrix region of MCCGs.21 Using the method, we could perform 3D culture of osteoblasts in the gel matrix regions of MCCGs, allowing us to construct engineered bone tissues that were more biomimetic than those constructed in previous studies. Accordingly, in the present study, we aimed to elucidate the effects of 3D culture of osteoblasts in MCCGs on the morphogenesis behaviors of the engineered bone tissue and to validate the utility of this 3D culture method in MCCGs as a methodology for constructing engineered bone tissues with biomimetic hierarchical structures.

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MATERIALS AND METHODS

Materials. In this study, we used an “atelo-collagen solution”, which was purchased from KOKEN Co. Ltd. (Tokyo, Japan), for preparing MCCGs. For this collagen solution, atelo-collagen lacking telopeptide at both terminal regions of native collagen was prepared at 5 mg/mL in 1 mM HCl. To induce the formation of MCCGs, we used PBg containing 13 mM KH2PO4 and 20 mM Na2HPO4 (Wako Pure Chemical Co., Ltd., Osaka, Japan). To reduce the difference between the osmotic pressure of PBg and cells, we added 5% D-(+)-glucose (Sigma Chemicals, St. Louis, MO, USA) or 194 mM glycerol to the PBg. The pH of PBg was 7.0. Dulbecco’s phosphate-buffered saline [PBS(−)], consisting of 2.3 mM KCl, 1.5 mM KH2PO4, 8.4 mM Na2HPO4, and 137 mM NaCl (pH 7.4), was used for washing the cells and tissues. The growth medium was minimum essential medium alpha (MEMα; Wako) supplemented with 10% fetal bovine serum replacement (FBR; Equitech-Bio, Inc., TX, USA) and 1% penicillinstreptomycin (Sigma Chemicals). The differential medium was growth medium supplemented with 10 mM glycerophosphate disodium salt pentahydrate (MP Biomedicals, Irvine, CA, USA) and 50 μg/mL Lascorbic acid phosphate magnesium salt n-hydrate (Wako). Preparation Methods of MCCG and Homogeneous Collagen gel. We obtained 2.8 M glycerol solution (Wako) by dissolving glycerol in 1 mM HCl (Wako). The atelocollagen solution was mixed with the glycerol solution at a mixing ratio of glycerol solution:atelocollagen solution = 1:9. The osmotic pressure of the collagen solution was balanced to that of the cells in order to reduce cellular damage. In this case, we obtained collagen solution containing 4.5 mg/ mL atelo-collagen and 0.28 M (≈ 280 mOsm) glycerol. The collagen B

DOI: 10.1021/acsbiomaterials.7b00190 ACS Biomater. Sci. Eng. XXXX, XXX, XXX−XXX

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ACS Biomaterials Science & Engineering solution was dispensed into a 1.5 mL sterilized microtube (Watson) and centrifuged with a desktop centrifuge for 10 min to degas the solution. To prepare the collagen gels, we used a dialysis chamber (Figure 1). Collagen solution (120 μL) was injected into the bore (8 mm diameter) of the bottom silicone rubber sheet (1 mm thickness), and a dialysis membrane was then placed on the top of the collagen solution. The top silicone rubber sheet was placed on the dialysis membrane. To induce MCCG formation, PBg was poured over the silicon rubber well. To prepare a collagen hydrogel without multichannel structures (Col), we used growth medium as an extradialytic solution instead of PBg. The samples were incubated at room temperature for 30 min and the PBg was then exchanged with the growth medium. The samples were incubated for 3 h at 37 °C in an atmosphere containing 5% CO2. After the incubation, the top silicone rubber sheet and the dialysis membrane were carefully removed to avoid damaging the samples. Finally, we obtained cylindrical collagen hydrogels both with and without multichannel structures in the dialysis chamber. The dimensions of the hydrogels were 8 mm diameter and 1 mm thickness. Preparation of MC3T3-E1 Cells. A murine osteoblastic precursor cell line derived from calvaria (MC3T3-E1 cells) was provided by RIKEN BRC through the National Bio-Resource Project of MEXT (Japan). The medium for MC3T3-E1 cell preculture was the growth medium. MC3T3-E1 cells were seeded in 100 mm dishes and cultured in growth medium at 37 °C in an incubator with an atmosphere containing 5% CO2. The medium was exchanged every 3 days. The cells were cultured up to 80−90% confluence and then used for the experiments described below. Construction of Engineered Bone Tissues. The harvested cells were resuspended in 4.5 mg/mL collagen solution with 0.28 M glycerol at a density of 4.0 × 106 cells/mL. To embed MC3T3-E1 cells into the collagen gel matrix of MCCG, we dialyzed the cell suspension against PBg by using the dialysis chamber. In addition, the cell suspension was dialyzed against the growth medium to embed MC3T3-E1 cells into Col. We also seeded MC3T3-E1 cells onto the top surface of MCCGs, the homogeneous collagen gel, or 35 mm glass bottom dishes. The samples were cultured in growth medium for 1 day and the growth medium was then exchanged with differentiation medium for samples cultured under differentiation conditions. All samples were cultured in an incubator at 37 °C in an atmosphere containing 5% CO2 with the medium replaced every 3 days. Samples cultured for 7, 14, 21, and 28 days were washed with PBS(−) three times and then fixed with 4% paraformaldehyde for 2 h at 4 °C. Samples were then washed with PBS(−) three times. We also prepared engineered bone tissues cultured in the growth medium and collagen hydrogels immersed in the differentiation medium for the same culture periods. The 3D engineered bone tissues with and without the multichannel structures cultured in the differentiation medium were termed as MCCG/Dif+/3D and Col/Dif+/3D, respectively. The samples where the cells were seeded on the top surface of hydrogels and cultured in the differentiation medium were termed as MCCG/Dif+/2D and Col/ Dif+/2D. For the samples cultured in the growth medium, “Dif−” was substituted in the name, giving MCCG/Dif−/3D and Col/Dif−/3D. The collagen gels with and without the multichannel structure immersed in the differentiation medium were termed as MCCG/Dif+ and Col/Dif+, respectively. The abbreviations for the samples are given in Table 1. Alizarin Red S Staining. To elucidate the effects of differences in hierarchical structures and seeding methods on the calcified morphologies for the engineered bone tissue, we visualized the deposited calcium ions on the sample by staining with Alizarin red S. The samples were immersed in 2% (w/v) Alizarin red S (Sigma Chemicals)/PBS(−) solution. After incubation for 1 h at 37 °C, the specimens were washed with Milli-Q water at least four times. Photographs of the stained samples were taken using a digital camera (Nikon D600, lens AF-S Micro NIKKOR 60 mm f/2.8G ED) under same exposure conditions. The mineralization area was measured by using ImageJ, available as a public domain NIH image program (http://rsb.info.nih.gov/nih-image). We first changed the type of

Table 1. Sample Abbreviations cultured in the differentiation medium

cultured in the growth medium

cells seeded on MCCG

MCCG/Dif+/2D

cells embedded in the collagen matrix of MCCG cells seeded on the collagen gel without multichannel structure cells embedded in the collagen matrix of collagen gel without multichannel structure MCCG without MC3T3-E1 cells Col without MC3T3-E1 cells

MCCG/Dif+/3D Col/Dif+/2D

MCCG/ Dif−/2D MCCG/ Dif−/3D Col/Dif−/2D

Col/Dif+/3D

Col/Dif−/3D

sample

MCCG/Dif+ Col/Dif+

photograph from 8-bit color to 8-bit grayscale and then the grayscale photographs were converted to the binary images with a same threshold gray value (= 128 a.u.). We measured the area of the dark region that corresponded to the area of calcified region for the binary image and the area of the cross-section of the whole sample using the original photographs. The ratio of the mineralization area to the whole sample was calculated and compared for each sample (n = 3). Wide-Angle X-ray Diffraction. To determine the main component of deposited mineral, we measured the wide-angle X-ray diffraction (WAXD) profiles for the samples. Samples cultured for 21 days were placed on 60 mm dishes and immersed in 50% ethanol (Junsei Chemical Co., Ltd., Tokyo, Japan) for 30 min, 70% ethanol for 30 min, and 100% ethanol overnight to remove the water from the samples. We obtained 2D WAXD patterns for the samples using an Xray diffractometer (Micro Max 007 HF; Rigaku, Tokyo Japan). The wavelength of the X-rays was 1.54 Å. The incident beam was focused on the samples using total reflection mirrors. The diffraction image was recorded on an R-Axis 4++ (Rigaku) imaging plate area detector. The distance from the samples to the imaging plate was 140 mm. The 2D WAXD patterns were recorded using Crystalclear (Rigaku) and converted to one-dimensional (1D) WAXD profiles by FIT2D (Andy Hammersley). Confocal Scanning Laser Microscopy. The blocking solution consisted of 1% bovine serum albumin (Sigma Chemicals) in PBS(−). Goat antitype I collagen antibodies (SC-25974; Santa Cruz Biotechnology, Inc., Dallas, TX, USA) and mouse antiosteocalcin antibodies (SC-376835; Santa Cruz Biotechnology, Inc.) were diluted 1:200 in blocking solution. Rabbit antidentin matrix protein 1 antibodies (M176; TaKaRa Bio, Inc., Shiga, Japan) were diluted 1:100 in blocking solution. Alexa Fluor 633-conjugated rabbit antigoat IgG, Alexa Fluor 633-conjugated goat antirabbit IgG, and Alexa Fluor 488-conjugated chicken antimouse IgG (all from Molecular Probes, Life Technologies Corporation, Carlsbad, CA, USA) solutions diluted 1:1000 in blocking solution were used as secondary antibodies for fluorescently labeling the primary antibodies. For F-actin staining, 10 U/mL Alexa Fluor 488-conjugated phalloidin solution was diluted 1:10 in blocking solution. For nuclear staining, 5 mM SYTOX Blue solution was diluted 1:1000 in blocking solution. The samples were placed in 2 mL sterilized microtubes (Watson). For the primary antibody treatment to label osteocalcin and dentin matrix proteins, the samples were decalcified by immersion in decalcification reagent containing 0.5 M ethylenediaminetetraacetic acid (EDTA) in PBS (Wako Chemical Co. Ltd.) overnight. To permeabilize the cell membrane, the samples were immersed in 0.5% Triton X-100 (Wako) in PBS(−) for 30 min at 4 °C. The samples were washed with PBS(−) three times and then incubated in blocking solution overnight. After blocking, the samples were incubated with the primary antibody solution in an incubator at 37 °C overnight and then washed with PBS(−) three times. After washing, the samples were incubated in a mixture of secondary antibodies, phalloidin solution, and SYTOX Blue solution in an incubator at 37 °C for 3 h. After washing with PBS(−) three times, the samples were mounted on a coverslip and observed using a confocal laser scanning microscope C

DOI: 10.1021/acsbiomaterials.7b00190 ACS Biomater. Sci. Eng. XXXX, XXX, XXX−XXX

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Figure 2. (A) Photographs of the samples stained with Alizarin red S. (B) Culture time courses of the calcified area normalized by the cross-sectional area of the sample (Sc). The error bars show standard deviations (n = 3). Asterisks indicate significant differences: p < 0.05. (CLSM; Leica, TCS SP5). The magnification of the objective lens was 10×, and the zoom factor was 1.7, 4, or 8. We obtained images from the top surface to a depth of 100 μm and images from the bottom surface to a depth of 100 μm. The images were collected using Leica Application Suite Advanced Fluorescence software (LAS AF, Leica). Statistical Analysis. All experiments were performed at least three times. The statistical comparisons for multiple groups were performed using one-way ANOVA, and then we compared each pair by using the Tukey method. The statistical analysis was performed with SigmaPlot 11.0 software (Systat Software, Inc., San Jose, CA, USA).

Dif+/2D, Sc increased on day 21 and then further increased on day 28, irrespective of the scaffolds and seeding methods. In particular, Sc for MCCG/Dif+/3D was significantly higher than those for other samples. The results showed that 3D culture of MC3T3-E1 cells in MCCGs resulted in the production of efficiently and homogeneously calcified engineered tissues. In addition, we observed the calcified morphologies for MCCG/ Dif+/3D on day 7 and day 28 by using scanning electron microscope-computed tomography (SEM-CT) imaging (Figure S2A, B). To quantitatively analyze the difference in the degree of calcification of the samples, we obtained CT-intensity profiles. Figure S2C, D show the CT-intensity profiles for the samples at day 7 and day 28, respectively. We observed the minimum CT-intensity near the mid portion (0.4 ≤ z̃ ≤ 0.6) of the sample at day 7 (Figure 2C), whereas a plateau was observed near the mid portion of the sample at day 28 (Figure 2D). The average CT-intensities at the mid portion (Imid) of the samples on day 28 were significantly higher than those on day 7 (Figure S2E). The results showed that the deeper parts of MCCG/Dif+/3D were also calcified in the depth direction. Composition of Minerals Deposited in the Engineered Bone Tissues. To further elucidate the effects of 3D culture of MC3T3-E1 cells in MCCGs on the calcified behaviors, we investigated the microscopic structures of samples using WAXD analysis and CLSM. WAXD profiles for the samples cultured in the differentiation medium are shown in Figure 3A. We observed two broad and intense peaks at 2θ = 26° and 2θ = 32° for samples cultured in differentiation medium. The peak positions of the two diffractions agreed with the (002) and (211) diffractions of hydroxyapatite (HAp).22,23 Therefore, the results showed that the minerals deposited in the samples mainly consisted of HAp. For the WAXD profiles of MCCG/Dif+/3D and Col/Dif+/3D, WAXD peaks that were not assigned to HAp were also observed, indicating the existence of additional mineral components. In particular, the peak near the 2θ = 25° might consist two peaks, as the peak has a shoulder. A major peak for monetite (CaHPO4) was observed near the 2θ = 25°.23 Therefore, the deposited minerals for MCCG/Dif+/3D and Col/Dif+/3D were likely predominantly

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RESULTS AND DISCUSSION Macroscopic Morphologies of the Engineered Bone Tissues. Photographs of samples stained with Alizarin red S are shown in Figure 2A. As calcium ions deposited in the samples were stained with Alizarin red S, the macroscopic morphologies of calcified samples could be observed in the photographs. MCCG/Dif−/3D, MCCG/Dif−/2D, Col/Dif−/3D, and Col/ Dif−/2D were not calcified (Figure S1A). In addition, samples without cells (MCCG/Dif+ and Col/Dif+) showed no deposition of Ca2+ ions (Figure S1B). Conversely, the samples with MC3T3-E1 cells cultured in differentiation medium were calcified on day 21, and further calcification of the samples occurred on day 28. In the photographs of MCCG/Dif+/3D at day 21 and day 28, the calcium was deposited entirely onto the samples. We defined the entirely calcified morphology as homogeneous calcification. In contrast, a spot-like and partial calcium deposition was observed for MCCG/Dif+/2D, Col/Dif +/3D, and Col/Dif+/2D at day 21. Such calcified morphologies were defined as inhomogeneous calcification. At day 21, MCCG/Dif+/3D was homogeneously calcified, whereas calcified morphologies for other samples were inhomogeneous. The calcified morphologies for all samples at day 28 were more homogeneous than those at day 21. To quantitatively analyze the homogeneity of the calcified samples, we measured the area of the calcified region normalized to the total area of the cross-section of the samples (Sc). Culture time courses of Sc are shown in Figure 2B. For MCCG/Dif+/3D, MCCG/Dif+/2D, Col/Dif+/3D, and Col/ D

DOI: 10.1021/acsbiomaterials.7b00190 ACS Biomater. Sci. Eng. XXXX, XXX, XXX−XXX

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of HAp crystallite.1,24 In the present study, we used the collagen gels as cellular scaffolds. Although the hierarchical structures of the collagen gels differed (Figure S3), the building blocks, i.e., the collagen fibers, were the same. Because the microenvironments surrounding MC3T3-E1 cells were not dependent on differences in the macroscopic structures of collagen gels, there were no significant differences in the microscopic structures and compositions of minerals deposited in the samples. Microscopic Morphologies of the Engineered Bone Tissues. The CLSM images at low magnification for MCCG/ Dif+/3D, MCCG/Dif+/2D, Col/Dif+/3D, and Col/Dif+/2D obtained near the top surface region on day 28 are shown in Figure 4A. In the ZX and YZ plane of the CLSM images, MC3T3-E1 cells covered the top surfaces of the samples, irrespective of the construction method of tissues. In contrast, in the deeper parts of the samples, the number of cells (Nc) decreased as the depth from the top surface of the sample (Z) increased. To quantitatively analyze changes in Nc with Z, we measured Nc by counting cell nuclei at different Z. Figure 5A shows the dependence of Nc on Z. For all samples, the maximum cell number was observed near the surface of samples, with Nc then decreasing as the Z increased further. Because nutrients and oxygen mainly diffuse from the top surface of the samples, the cells were well proliferated on the top surface or migrated from the deeper parts of samples to the top surface. Nc at the shallow part (Z = 30 μm) for MCCG/Dif +/3D and MCCG/Dif+/2D were significantly higher than those for Col/Dif+/3D and Col/Dif+/2D (Figure 5B). In contrast, Nc at a deeper part (Z = 60, and 90 μm) for MCCG/ Dif+/3D was significantly larger than those for other samples (Figure 5C-D). Therefore, the cell density in the deeper parts of MCCG/Dif+/3D was highest in the constructed engineered bone tissues. Because the collagen fiber density in the deep channel region was less than those of the gel matrix regions of MCCGs and the homogeneous collagen gel, the multichannel structure could function as transport ducts for oxygen and nutrients. Furthermore, the cells were seeded in the 3D gel matrix region of MCCGs. Therefore, MC3T3-E1 cells efficiently proliferated in MCCG/Dif+/3D. Although the cells were initially embedded in the gel matrix region as shown in Figure S4, the morphology for MCCG/Dif +/3D at day 28 shows that many cells were found on the surface of the channel. We speculate that the cells on the channel surface migrated from the collagen gel matrix or proliferated near the channel. In native bone tissue, osteoblasts adhere on the surface of the Harversian canals and form toroidlike structures.25 For MCCG/Dif+/3D, the MC3T3-E1 cells on the channel surface elongated along the channel surface and also formed a toroid-like structure. In comparison, osteocytes in native bone tissue are found in a bone matrix and form a network structure by means of numerous processes.26 For MCCG/Dif+/3D, MC3T3-E1 cells embedded in the collagen matrix region also formed a network-like structure. A CLSM image for MCCG/Dif+/3D at high magnification (Figure 4C) shows that the network-like structure consisted of cells linked with process-like protrusions and further that it was also connected with the cells forming the toroid-like structures on the channel surface (Figure 4D). These results indicate that MCCG/Dif+/3D reproduces the morphology of native bone tissue. In the CLSM image of MCCG/Dif+/2D, toroid-like structures were also observed on the channel surface of MCCG, whereas the number of cells in the collagen matrix region was much less than that for MCCG/Dif+/3D. Several studies have

Figure 3. Wide-angle X-ray diffraction profiles for samples cultured in (A) differentiation medium or (B) growth medium.

HAp and partially monetite. Monetite constitutes the alternate crystalline state of calcium phosphate that is formed under lower pH conditions. We found that the color of the medium easily changed from red to orange during the long culture period, indicating that the pH of medium decreased. Therefore, the presence of monetite might be attributed to partially dissolved HAp. As shown below, the number of cells in the deeper portion for MCCG/Dif+/3D and Col/Dif+/3D were higher than that for MCCG/Dif+/2D and Col/Dif+/2D. The consumption of nutrients and oxygen increased with increasing number of cells in the samples. However, the oxygen and nutrients are only provided from the surface of the samples. In the deeper part of the samples, therefore, the amounts of nutrients and oxygen might not be sufficient to maintain the cells, with insufficient nutrients and oxygen potentially inducing cell necrosis. This in turn would decrease the pH of medium. This further supports that the HAp was partially dissolved by decreasing pH of the medium during culture. In contrast, no diffraction peaks were observed for samples cultured in growth medium, as shown in Figure 3B. These results showed that calcification of the samples with the cells could be attributed to the osteogenic behaviors of MC3T3-E1 cells. Osteoblasts produce osteoids, which consist of bone matrix proteins, such as type I collagen and osteocalcin, and then they mineralize the osteoid through deposition of HAp.3 During the biomineralization process, collagen fibers play a role in template E

DOI: 10.1021/acsbiomaterials.7b00190 ACS Biomater. Sci. Eng. XXXX, XXX, XXX−XXX

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Figure 4. Confocal scanning laser microscopic (CLSM) images of the engineered bone tissues constructed using different methods. (A) CLSM images observed from the top surface of the samples. The images were obtained at Z = 30 μm. Arrowheads indicate the toroid-like structure. (B) CLSM images observed from the bottom surface of the samples. The images were obtained at Z = 20 μm. The images for the ZX and YZ planes of MCCG/Dif+/3D (upper left) are shown in reverse orientation; i.e., the upper side is the inner part and the lower side is the bottom part. (C, D) High-magnification CLSM images obtained near the top surface of MCCG/Dif+/3D on day 28. C. Network-like structure formed in the collage matrix of MCCG. (D) Toroid-like structures formed on the channel surface. (E, F): High-magnification CLSM images obtained near the bottom of MCCG/Dif+/3D on day 28. (E) Network-like structure formed in the collage matrix of MCCG. F. Toroid-like structures formed on the channel surface. Nuclei, F-actin, and type I collagen are colored in blue, green, and red, respectively.

channel surface. The preferential invasion of MC3T3-E1 cells into the inner channel region may be attributed to the efficient transport of oxygen and nutrients through the multichannel

shown that osteoblasts or osteoblastic cells seeded on the surface of a collagen gel can invade into the deep part of the collagen gel.27,28 However, almost all cells were found on the F

DOI: 10.1021/acsbiomaterials.7b00190 ACS Biomater. Sci. Eng. XXXX, XXX, XXX−XXX

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Figure 5. A: Numbers of cells in the engineered bone tissues (Nc) measured as a function of the depth from the top surface of the samples (Z). (B, D) Statistical comparisons of Nc at each Z. The error bars show standard deviations (n = 3). Asterisks indicate significant differences: p < 0.05.

Figure 6. (A) Number of cells in the engineered bone tissues (Nc) measured as a function of the depth from the bottom surface of the samples (Z′). (B−D) Statistical comparisons of Nc at each Z′. Error bars show standard deviations (n = 3). Asterisks indicate significant differences: p < 0.05.

G

DOI: 10.1021/acsbiomaterials.7b00190 ACS Biomater. Sci. Eng. XXXX, XXX, XXX−XXX

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ACS Biomaterials Science & Engineering

solution, which is limited by the rate of fibrillogenesis of collagen fibers, decreases as the ionic strength increases.30,31 Because the ionic strength of the growth medium was much higher than that for the PBg, the gelation rate of the homogeneous collagen gel could be slower than that for MCCG. Therefore, the number of MC3T3-E1 cells that settled to the bottom region during the formation process for Col/Dif +/3D was higher than that for MCCG/Dif+/3D. As shown in Figure 4, we found that MC3T3-E1 cells in MCCG/Dif+/3D were adhered on the surface of the channel and were also embedded in the gel matrix region of MCCG. To elucidate the differences between the cells on the channel surface and in the gel matrix, we observed the expression of osteocalcin (OC) and dentin matrix protein-1 (DMP1). Figure 7 shows CLSM images for MC3T3-E1 immunofluorescently

structure. In the CLSM image of Col/Dif+/3D, the cells were also embedded in the collagen gel matrix and formed networklike structures. In addition, clusters consisted of cells were observed in the collagen matrix as well. The CLSM image indicates that the network-like structure in Col/Dif+/3D was formed by connecting the cell clusters with the elongated cells, as for MCCG/Dif+/3D. However, no toroid-like structures were observed because there were no multichannel structures in the deeper parts of the homogeneous collagen gel. In contrast, in the CLSM image of Col/Dif+/2D, almost no cells were observed in the collage matrix of Col. These results could be attributed to the lack of transport ducts for nutrients and oxygen in the deeper parts of the homogeneous collagen gel. Figures 4B and 6 show the tissue morphology near the bottom of samples and Nc characterized as a function of the depth from the bottom surface (Z′), respectively. In a previous study, we showed that the channel diameter of MCCG increased with increasing Z, whereas the channel number decreased with increasing Z.29 For MCCG/Dif+/3D and MCCG/Dif+/2D, therefore, the bottom channel diameter was larger than that near the top surface, whereas the bottom channel number was lower than that near the top surface. The CLSM image obtained at the bottom region of MCCG/Dif +/3D showed that MC3T3-E1 cells were completely surrounded by the collagen gel matrix. Because the thickness of the gel matrix region between the channels was larger than the size of the cells, MC3T3-E1 cells could be embedded in the collagen gel matrix. A high magnification CLSM image for MCCG/Dif+/3D (Figure 4E) clearly shows that the cells embedded in the collagen matrix exhibit stretched process-like protrusions that were interconnected. Therefore, the cells embedded in the collagen matrix also formed network-like structures. Figure 4F shows that some of the cells also elongated along the channel surface and formed toroid-like structures. The cells forming the network-like structure in the collagen matrix were also connected with the cells forming the toroid-like structure in the inner channel region. As shown in Figure 4, we found that there were two types of cells cultured under different conditions. The cells embedded in the collagen matrix were subjected to 3D culture, whereas the cells adhered on the channel surface were subjected to 2D culture. The differences among these cells are discussed below. Although the number of cells was small, the cellular morphology for Col/Dif+/3D was almost the same as that near the top surface. The cells embedded themselves into the collagen matrix and formed a stretched shape. However, they did not form a network-like structure, because number of cells was insufficient. The number of cells near the bottom surfaces of MCCG/Dif+/2D and Col/Dif+/2D were much lower than those for MCCG/Dif+/3D and Col/Dif+/3D (Figure 6). Because there were no cells in the bottom region for MCCG/ Dif+/2D and Col/Dif+/2D just after seeding cells on the top surface, these results were easily expected. Nc for MCCG/Dif+/3D above Z′ = 60 μm were significantly higher than those for Col/Dif+/3D and Col/Dif+/2D (Figure 6D, E). The average value of Nc for MCCG/Dif+/3D was higher than that for MCCG/Dif+/2D from Z′ = 10 to 100 μm, although the difference was not statistically significant (Figure 6A). In contrast, Nc for Col/Dif+/3D below Z′ = 30 μm were significantly higher than those for other samples (Figure 6B). When we constructed Col/Dif+/3D, we dialyzed the collagen solution containing MC3T3-E1 against the growth medium. Under isothermal conditions, the gelation rate of the collagen

Figure 7. Expression patterns of osteocalcin (OC) and dentin matrix protein-1 (DMP1). Arrowheads indicate examples of the cells selected for measuring the fluorescence intensities of OC and DMP1. We selected single cells in single CLSM images and scanned the whole cell body of each cell.

labeled with OC and DMP1 on a glass-bottomed dish and in the gel matrix region of MCCG on day 1. Although the cells were cultured in growth medium for only 1 day after seeding or embedding, they expressed OC and DMP1. Figure 7 also shows the expression patterns of OC and DMP1 in the cells that adhered on the channel surface and embedded in the gel matrix region. MC3T3-E1 cells also expressed OC and DMP1, irrespective of their positions. Notably, we identified several H

DOI: 10.1021/acsbiomaterials.7b00190 ACS Biomater. Sci. Eng. XXXX, XXX, XXX−XXX

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ACS Biomaterials Science & Engineering

Figure 8. (A, B) Fluorescence intensities of OC and DMP1 for cells under different microenvironmental conditions. (C) Ratios of IOC to IDMP1 (RI) for cells under different microenvironmental conditions. Error bars show standard deviations (n = 9). Asterisks indicate significant differences: p < 0.05.

an osteocyte marker.28,32 RI for the cells in the gel matrix were significantly lower than those for the cells on the channel surface. These results indicated that the cells in the gel matrix had an osteocyte-like phenotype, whereas the cells on the channel surface had an osteoblast-like phenotype. To support this suggestion, we observed the expression levels of Sclerostin (ISclerostin) that is another osteocyte marker, and OC (IOC) for the cells under different conditions in MCCG (Figure S5). As shown in Figure S6, R′I, which is defined as an intensity ratio of IOC to ISclerostin (R′I = IOC/ISclerostin), for the cells in the collagen matrix on day 28 was significantly lower than that for the cells on the channel surface at the same day of culture. This result supports our speculation. We believe that the coexistence of osteocyte-like and osteoblast-like cells reproduced the tissue structure of compact bone. To precisely determine the differentiation stage of MC3T3-E1 cells, we should measure the expression levels of osteoblast and osteocyte makers for the cells under different culture conditions with more quantitative methods, such as quantitative PCR and quantitative Western blotting. However, it is difficult to separately evaluate the expression levels for the cells in MCCG. Therefore, the development of selective methods for measuring the expression levels of the differentiation makers for the cells in the collagen matrix and on the channel surface are required. Effects of Cell Culture Conditions on the Calcification Behaviors of Engineered Bone Tissues. For MCCG/Dif +/3D, the samples were homogeneously calcified by MC3T3E1. CLSM images showed that MC3T3-E1 cells were homogeneously embedded in the gel matrix of MCCGs and were adhered on the channel surface. Therefore, the contact area between the cells and the collagen matrix was wider than that for other samples. Furthermore, the cell density at the deep part for MCCG/Dif+/3D was significantly higher than that for the other samples. Thus, the efficient and homogeneous calcification of MCCG/Dif+/3D could be attributed to the large contact area between the cells and the matrix and the high cell density, suggesting that MCCG may be suitable for constructing engineered tissues with homogeneous calcification and a high cell density. Reproduction of the Structures of Native Bone Tissues. In native bone tissues, osteoblasts adhere on the interior surface of Haversian canals, and osteocytes are embedded in the bone matrix.1 The MCCG/Dif+/3D constructed in this study had various biomimetic features of

cells with high DMP1 and low OC expression in the gel matrix region on day 21 and day 28. Therefore, to quantitatively compare the expression levels of DMP1 and OC, we measured the averaged fluorescence intensities scaled based on the cell area. Figure 8A and B show the average fluorescence intensities of DMP1 (IDMP1) and OC (IOC). Figure 8A shows that no significant change in IDMP1 for the cells in the collagen matrix between day 21 and day 28 was observed, whereas IDMP1 for the cells on the channel was significantly decreased on day 28. In addition, IDMP1 for the cells on the channel on day 21 was significantly larger than that for the cells in the collagen matrix. Figure 8B shows that there were no significant differences in IOC for the cells in the collagen matrix on day 21 and day 28, whereas IOC for the cells on the channel significantly decreased on day 28. Furthermore, IOC for the cells on the channel surface was significantly larger than that for the cells in the collagen matrix. For mouse calvarial osteoblasts cultured on a plastic cell culture plate, which provides a 2D cell culture condition, it was reported that the maximum gene expression level for DMP1 was observed on day 21 and then it was restricted on day 28.32 As the cells on the channel surface are essentially cultured under 2D conditions, IDMP for the cells on the channel surface was also restricted at day 28. In contrast, Uchihashi et al. reported that the gene expression level of DMP1 for MC3T3E1 cultured three-dimensionally in a type I collagen gel increases at the late stage of osteoblastic differentiation.28 Although our result did not show a significant increase in IDMP1 for the cells in the collagen matrix on day 28, it was still maintained. These results may be attributed to the 3D culture in the collagen matrix of MCCG. In addition, the expression of OC increases through the late stages of osteogenesis in osteoblasts and then further progression of osteogenesis results in decreased expression of OC.3,33 Therefore, the observed changes in IOC and IDMP1 could be attributed to the progression of osteogenesis. To discuss differences in the differentiation behaviors of cells on the channel surface and in the collagen matrix, we evaluated the fluorescence intensity ratios of IOC to IDMP (RI = IOC/IDMP). Figure 8C shows the RI for the cells cultured under different conditions in MCCG. The RI values for the cells in the collagen matrix were significantly lower than those for the cells on the channel surface on day 21 and day 28. OC and DMP1 are expressed in osteoblasts during the late stages of osteogenesis and in osteocytes.3,28,32−35 Several reports have used OC as an osteoblast marker and DMP1 as I

DOI: 10.1021/acsbiomaterials.7b00190 ACS Biomater. Sci. Eng. XXXX, XXX, XXX−XXX

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ACS Biomaterials Science & Engineering native bone tissue. For example, the extracellular matrix mainly consisted of type I collagen and the multichannel structure could reproduce the parallel array of Haversian canals. The MCCG/Dif+/3D contained cells embedded in the bone matrix and cells adhered on the channel surface, reproducing the locations of osteocytes and osteoblasts in the native bone tissue. However, although MCCG/Dif+/3D exhibited various biomimetic structures, its mechanical properties were still soft and fragile. Therefore, further developments are required for reproducing the mechanical properties of bone tissues. Among the various technologies used to reconstruct osteonlike structures, biomaterials constructed using 3D-printing technology excellently reproduces the osteon structures of compact bone tissue in the macroscopic scale.14 Because the biomaterial is designed for mimicking the mechanical properties, it might be used as an anatomic bone model and mechanomimetic implant. However, the method is not designed for embedding the cells in the matrix region of the biomaterial, and it is difficult to reproduce the microscopic structure resulting from collage fiber alignment. Therefore, this technique is not currently adequate for constructing model systems to investigate the cellular behaviors in bone tissues. In contrast, MCCG mimics the complex hierarchical structures of compact bone tissues. Furthermore, in the present study, we have shown that MCCG can be used as a cellular scaffold for threedimensionally culturing the MC3T3-E1 cells. The 3D engineered bone tissues constructed by using MCCG are more suitable for investigating various cellular behaviors in the bone tissues and therefore provide a model system to study bone metabolism and to test the efficacy of drugs.

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AUTHOR INFORMATION

Corresponding Author

*E-mail: [email protected]. ORCID

Kazuya Furusawa: 0000-0002-4465-3711 Author Contributions

K.F., M.N., and T.F. designed the research. S.Y. and K.F. constructed the engineered bone tissues. S.Y., K.F., K.N., and M.N. performed experiments and analyzed the results. K.N., M.N., and T.F. contributed to obtaining SEM-CT images of the samples. S.Y. and K.F. wrote the paper. Notes

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The authors declare no competing financial interest.

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CONCLUSION In this study, we investigated the effects of the structure of the collagen gel and the dimensionality of cell culture on the morphogenic behaviors of engineered bone tissues. The engineered bone tissues constructed by embedding the cells in the gel matrix of MCCGs were highly and homogeneously calcified and had a high cell density. Therefore, 3D culture in MCCGs provided appropriate conditions for constructing homogeneously calcified engineered bone tissues with a high cell density. Furthermore, the observation of tissue morphology and the comparisons of the expression levels of differentiation makers showed that 3D culture of MC3T3-E1 cells in MCCGs resulted in the coexistence of two types of cells with different morphologies and differentiation stages. In a previous study, we developed a method for constructing transport ducts, including blood vessels and kidney tubules, in a multichannel structure.29 By combining the present method for constructing 3D engineered bone tissues with the construction methods of transport ducts, we could further reproduce the hierarchical structure of bone tissues. If engineered bone tissues reproducing the hierarchical structure of bone tissues could be constructed, they could be used to investigate the morphogenic mechanism of bone tissues and for other basic bone science studies, such as studies on effects of mechanical and electrical stimuli on the remodeling behaviors of bone tissues.

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Photographs for the engineered bone tissues cultured in the growth medium stained and for MCCG and Col immersed in the differentiation medium stained with Alizarin red S (Figure S1); SEM-CT analysis for MCCG/Dif+/3D on day 7 and day 28 (Figure S2). photographs for MCCG and Col obtained under the natural light and polarized light (Figure S3); initial microscopic morphology for an engineered bone tissue where MC3T3-E1 cells embedded into the collagen gel matrix of MCCG (Figure S4); measurement of expression levels of Sclerostin and Osteocalcin for cells in the collagen matrix and cells on the channel surface (Figure S5); fluorescence intensities of Sclerostin (ISclerostin) and OC (IOC), and ratios of IOC to ISclerostin (R′I) for cells under different microenvironmental conditions (Figure S6) (PDF)

ACKNOWLEDGMENTS This work was supported by a Grant-in-Aid for Scientific Research on Innovative Areas, “Hyper Bio Assembler for 3D Cellular Innovation” (23106006 and. 26106703), from the Ministry of Education, Culture, Sports, Science, and Technology (MEXT) and by a Grant-in-Aid for Young Scientists (B) (no. 15K16330), from the Japan Society for the Promotion of Science (JSPS).

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DOI: 10.1021/acsbiomaterials.7b00190 ACS Biomater. Sci. Eng. XXXX, XXX, XXX−XXX