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3D Printing of Nano Hydroxyapatite/Poly(ester urea) Composite Scaffolds with Enhanced Bioactivity Jiayi Yu, Yanyi Xu, Shan Li, Gabrielle V Seifert, and Matthew L. Becker Biomacromolecules, Just Accepted Manuscript • DOI: 10.1021/acs.biomac.7b01222 • Publication Date (Web): 11 Oct 2017 Downloaded from http://pubs.acs.org on October 12, 2017

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3D Printing of Nano Hydroxyapatite/Poly(ester urea) Composite Scaffolds with Enhanced Bioactivity

Jiayi Yu, † Yanyi Xu, † Shan Li, † Gabrielle V. Seifert, † and Matthew L. Becker*,†,∥

†

Department of Polymer Science, The University of Akron, Akron, Ohio 44325, United States

∥

Department of Biomedical Engineering, The University of Akron, Akron, Ohio 44325, United States

KEYWORDS: Hydroxyapatite, composite, osteogenic differentiation, porous scaffolds, 3D printing

ACS Paragon Plus Environment

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ABSTRACT Polymer-bioceramic composites incorporate the desirable properties of each material while mitigating the limiting characteristics of each component. 1,6-hexanediol L-phenylalanine-based poly(ester urea) (PEU) blended with hydroxyapatite (HA) nanocrystals were 3D-printed into porous scaffolds (75% porosity) via fused deposition modeling (FDM) technique and seeded with MC3T3-E1 pre-osteoblast cells in vitro to examine their bioactivity. The resulting 3Dprinted scaffolds exhibited a compressive modulus of ~50 MPa after 1-week incubation in PBS at 37 °C, a cell viability of >95%, and a composition dependent enhancement of radio-contrast. The influence of HA on MC3T3-E1 proliferation and differentiation was measured using quantitative real-time polymerase chain reaction, immunohistochemistry and biochemical assays. After 4 weeks, alkaline phosphatase (ALP) activity increased significantly for the composite with 30% HA with values reaching 2.5-fold greater compared to the control. Bone sialoprotein (BSP) showed an approximately 880-fold higher expression and 15-fold higher expression of osteocalcin (OCN) on the 30% HA composite compared to the control. Calcium quantification results demonstrated a 185-fold increase of calcium concentration in mineralized extracellular matrix deposition after 4 weeks of cell culture in samples with higher HA content. 3D-printed HA-containing polymer composites provide an efficient and customized method to promote bone regeneration and have the potential to be used in orthopedic applications.


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INTRODUCTION Bone defects continue to pose significant problems for surgeons. More than 6 million patients in the United States suffer from bone defects or injuries each year.1,2 Regenerating osseous tissue using cells and a synthetic extracellular matrix is a new approach that aims to replace or supplement the transplantation of harvested auto-/allo- grafts.3-8 Using the synthetic approach, a highly porous three-dimensional scaffold is needed as a substrate to facilitate cell attachment, migration, proliferation, differentiation and ultimately tissue regeneration. Several synthetic polyesters derived from lactides, glycolides, lactones and their respective copolymers have been shown to be non-toxic and biodegradable in a number of applications in the literature. Many of these materials have been used in tissue engineering scaffolds and have been included in Federal Food and Drug Administration approved clinical applications including surgical sutures and implantable devices.5,9-12 One disadvantage of these materials is that the degradation products are acidic, reduce the local pH when the constructs are large which may in turn accelerate the degradation rate and induce local and systemic inflammation.13-15 Among numbers of other biodegradable polymers used clinically, α-amino acid-based poly(ester urea)s (PEUs) are promising materials for biomedical applications. They have highly tunable mechanical and degradation properties, nontoxic hydrolysis byproducts and can be functionalized easily via “click” chemistry.16-23 Specifically, 1,6-hexanediol L-phenylalanine-based PEU (poly(1-PHE-6)) has shown a high elastic modulus (~3.2 GPa) and no evidence of local acidification-induced inflammatory response has been observed in vivo, which can be explained by the presence of acid-neutralizing urea groups at each repeat unit. The structural, mechanical and chemical properties of poly(1PHE-6) make it a translationally relevant candidate for bone tissue regeneration applications.


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While synthetic materials have advantages, they are not osteoinductive or osteoconductive and can not drive osteoblast differentiation nor enhance the extent and rate of bone regeneration without additives. Bioceramics, especially CaP-based ceramics, have been used commonly to treat bone defects due to their low density and compositional similarities to natural bone.24-26 Hydroxyapatite (HA, Ca10(PO4)6(OH)2) has been used in a number of bone tissue-engineering applications including bone cement for craniofacial defect repair and coatings for femoral components of hip replacements due to its strong mechanical properties and osteoconductivity.2729

In vivo studies have previously shown that bone has a greater affinity for implants containing

higher quantities of HA over those with only trace amounts. However, the inherent brittleness of HA makes the manufacturing process challenging and inefficient.30 A solution to potential solution to the problem is a polymer-bioceramic composite material that incorporates the desirable properties of the ceramic and polymeric materials while mitigating the limiting characteristics of each component. Polymer-bioceramic composites mimic natural bone which is a hierarchical composite of organic compounds (mainly collagen) and inorganic ceramic (mainly nano-sized HA). The organization of calcium phosphate crystals and collagen fibers renders bone both bioactive and strong.31 In addition, Webster et al. have shown significant increases in protein adsorption and osteoblast adhesion on the nano-sized HA.32 Significant effort has attempted to create porous polymer-bioceramic composites from traditional fabrication methods including salt-leaching, phase separation and gas foaming. These methods generally result in heterogeneous pores and suboptimal distribution of cells. Additive manufacturing technologies are well suited to create well-defined porosity and patient-specific biomimetic structures.33-43 Fused deposition modeling (FDM) is a rapid prototyping technique that yields a three-dimensional architecture from a computer-aided design/manufacturing


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(CAD/CAM) model. FDM has been used to fabricate a number of biomedical implants including highly porous polyurethane human ear scaffolds and artificial porous polybutylene terephthalate canine cancellous bone scaffolds.44,45 FDM uses a sequential layer-by-layer fabrication process that results in a three-dimensional structure that is able to adapt the outer edges of an implant to a patient specific defect while controlling the inner pore architecture.46 It also enables the ability to fabricate bone components that mimic the organic/inorganic chemical composition and the architecture of natural bone.47,48 Herein, we utilize FDM to fabricate a series of three-dimensional porous poly(ester urea)/nano-hydroxyapatite composite scaffolds with various amounts of HA. The influence of HA concentration on the scaffold fabrication, mechanical properties, in vitro cytotoxicity, osteoinductivity and osteoconductivity are thoroughly investigated. The results showed that the composite scaffolds possess sufficient mechanical strength to be used in segmental bone defect applications and the scaffolds with higher HA contents promote cell proliferation, osteogenic differentiation and mineralization of MC3T3-E1 cells. We feel these constructs can be translated to more advanced preclinical studies which are ongoing.

EXPERIMENTAL SECTION

Materials. All chemicals and reagents were purchased from Sigma or Fisher Scientific and used as received unless noted otherwise.

Chloroform was distilled after drying overnight with

calcium hydride.

Synthesis and Characterization of Poly(ester urea) Monomer and Polymer.


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Synthesis of Di-p-toluene Sulfonic Acid Salts of Bis-L-phenylalanine-hexane-1,6-diester Monomer (PEU-M). Di-p-toluene sulfonic acid salts of bis-L-phenylalanine-diol-diester monomer was prepared using procedures published previously, as shown in Scheme 1(a).17,18 (1H NMR, 13C NMR, ESI mass spectrum in Figure S1, Figure S2 and Figure S3) 1

H NMR (500 MHz, DMSO-d6): 1.04-1.13 (m, 4H, -COOCH2CH2CH2-) 1.38-1.44 (m, 4H, -

COOCH2CH2CH2-) 2.27 (s, 6H, CH3Ar-) 2.50 (DMSO) 2.98-3.19 (m, 4H, -CHCH2-Ar-) 3.894.03 (m, 4H, -COOCH2CH2-) 4.25-4.32 (m, 2H, +NH3CHCOO-) 7.09-7.13 (d, 4 H, aromatic H) 7.21-7.34 (m, 10H, aromatic H) 7.47-7.50 (d, 4H, aromatic H) 8.36 (s, 6H, +NH3-).

13

C-NMR

(125 MHz, DMSO- d6): 20.75, 24.66, 27.62, 35.97, 38.80-40.28 (DMSO-d6), 53.07, 65.46, 125.39, 127.14, 127.95, 128.49, 129.30, 134.69, 137.78, 145.33, 169.03. Synthesis of 1,6-hexanediol L-phenylalanine based Poly(ester urea) (PEU). The polymer was synthesized by interfacial polymerization as shown in Figure 1(a). (1H NMR in Figure 1(b). 13C NMR and IR in Figure S2 and Figure S6) 1

H NMR (500MHz, DMSO-d6): 1.15 (m, 4H, -COOCH2CH2CH2-), 1.43 (m, 4H, -

COOCH2CH2CH2-), 2.5 (DMSO), 2.87-2.94 (m, 4H, -CHCH2Ar-), 3.94 (m, 4H, CHCOOCH2CH2-), 4.34-4.40 (m, 2H, -NHCHCOO-), 6.47-6.5 (m, 2 H, -NH-), 7.14-7.17 (m, 4H, aromatic), 7.19-7.28 (d, 6H, aromatic). 13C NMR (75MHz, DMSO-d6): 25.33, 28.38, 38.10, 39.40-40.56 (DMSO), 54.49, 64.65, 127.04, 128.78, 129.65, 137.34, 157.07, 172.89. Characterization of PEU-M and PEU. 500MHz 1H NMR and 125MHz

13

C NMR spectra were recorded using a Varian NMR

Spectrophotometer. All chemical shifts were reported in ppm (δ), and referenced to the chemical shifts of the residual solvent resonances (1H NMR DMSO-d6 2.50 ppm;

13

C NMR DMSO- d6

39.50 ppm). The following abbreviations were used to explain the multiplicities: s=singlet,


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d=doublet, t=triplet, br=broad singlet, m=multiplet. Infrared spectra of all compounds were recorded using MIRACLE 10, Shimadzu Corp. ATR-FTIR spectrometer with 4 cm-1 resolution. Electrospray ionization (ESI) of PEU-M was performed using a HCT Ultra II quadrupole ion trap mass spectrometer (Bruker Daltonics, Billerica, MA) equipped with an electrospray ionization source. Number-average molecular mass (Mn), weight-average molecular mass (Mw) and post-precipitation molecular mass distribution (ÐM) were determined by size exclusion chromatography (SEC). The SEC analyses were performed using a TOSOH HLC-8320 gel permeation chromatograph instrument. The experiments were carried out with a flow rate of 1mL/min using HPLC grade N,N-dimethylformamide (DMF) with 25 mM LiBr as the eluent at 323K with a refractive index (RI) detector and molecular mass values were determined relative to polystyrene standards.

Synthesis and Characterization of Hydroxyapatite (HA). The hydroxyapatite was synthesized by hydrothermal method. 0.75 M calcium solution (pH 10.5) was mixed with 0.25 M phosphate solution (pH 10.5) with a Ca/P ratio of 1.71 at 100 ºC for 2 hours and kept stirring at room temperature for another 20 hours under nitrogen. White precipitant was collected, washed with ammonium hydroxide solution (pH 10.5) and characterized. The crystalline structure of HA was measured by wide angle X-ray diffraction measurement (WAXD) (Bruker AXS diffractometer with a two-dimensional detector) at room temperature. The generator was operated at 40 kV and 40 mA with a beam monochromatized to Cu Kα radiation. A typical exposure time was 5 min. The air scattering was subtracted. One-dimensional WAXD curve was integrated from the two-dimensional image. A survey scan of X-ray


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photoelectron spectroscopy (XPS) measurement was performed on a Kratos AXIS Ultra DLD spectrometer to calculate the Ca/P ratio. The X-ray source was monochromated Al Kα, scanning over a binding-energy range of (0 to 700) eV with a dwell time of 100 ms. Each spectrum was collected over a 300 × 700 µm sample area. The Ca/P ratio was also measured by inductively coupled plasma-atomic emission spectrometry (ICP-AES) (Agilent Technologies 700 series, Santa Clara, CA, USA). HA was dissolved in concentrated HCl and the concentration of Ca and P ions was measured. The morphology of HA was characterized by scanning electron microscope (SEM) (JSM-7401F, JEOL, Peabody, MA) and transmission electron microscope (TEM) (JEOL1230 TEM, Peabody, MA).

Fabrication and Characterization of Composite Scaffolds. Poly(ester urea) and HA were mixed in hexafluoroisopropanol (HFIP) with a weight ratio of 100:0, 90:10, 80:20, 70:30,60:40 respectively. The solutions were sonicated at room temperature for 2 hours and freeze-dried to remove the solvent. After getting the composite, the filament with a diameter of 1.8 cm was obtained from capillary rheometer (Dynisco LCR 7000) at 165 ºC and fed into the heated nozzle (Cartesio 3D printer). The material was extruded through a 0.3 mm nozzle and printed layer by layer in the x and y plane with a printing speed of 2 mm/s. The stage moved up and down in the z axis. Resulting geometry is a serious of rods layered perpendicularly. The thermal stability of the polymer composites was measured using thermogravimetric analysis (TGA, TA Q500) across a temperature range of 0ºC to 600ºC at a scanning rate of 20ºC under nitrogen. The experimental determined HA contents in the scaffolds were obtained from the weight% at 550 ºC in TGA. The thermal characteristics of the polymer composites were


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determined using differential scanning calorimetry (DSC, TA Q2000) from 0ºC to 220ºC at a scanning rate of 20 ºC/min. The resulting values of the thermal properties were determined from three individual measurements. The glass transition temperature was determined from the midpoint in the second heating cycle of DSC. The structure of the filament and 3D printed porous scaffolds was characterized nondestructively using X-ray micro-computed tomography (µ-CT, Skyscan 1172). 3D scanning of scaffolds was carried out under the following parameters: 60 kV voltage, medium camera, no filter, 30 ms camera exposure preset time, 0.5 µm resolution for the scans of filaments and 10.0 µm resolution for the scans of scaffolds. The mechanical properties of the composite scaffolds were studied by compression tests (Instron 5543 Universal Testing Machine) with a Hencky strain of 0.1 min-1 at three different conditions: tested at room temperature, physiological temperature and tested after 1 week of incubation in PBS. The dimensions of the scaffolds were 7.7 mm in diameter and 1.5 mm in thickness. The compressive moduli were calculated using the slope of linear fitting in the linear regime. The reported results are average values from three individual measurements.

in vitro Cell Culture on 3D printed scaffolds. Sterilization of Scaffolds. All 3D printed scaffolds were sterilized by 12 h cycle of ethylene oxide sterilization and 48 h of degas. Then scaffolds were submerged in 1 mL of cell medium for 5 h prior to cell seeding. Cell Culture. MC3T3-E1 Subclone 4 mouse preosteoblast cells (ATCC, Manassas, VA) were expanded and cultured in α-MEM media with ribonucleosides, deoxyribonucleosides, 2 mM Lglutamine and 1 mM sodium pyruvate, but without ascorbic acid (GIBCO, Life Technologies,


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Grand Island, NY) supplemented with 10% fetal bovine serum (FBS) (Invitrogen, Grand Island, NY), 100 units/mL penicillin (Invitrogen, Grand Island, NY), and 100 µg/mL streptomycin (Invitrogen, Grand Island, NY) at 37°C in a 5% CO2 humidified atmosphere. For cell seeding, cells were rinsed with DPBS and detached from the bottom of the flask using 0.05% trypsin/ethylenediaminetetraacetic acid tetrasodium salt (EDTA) solution at 37 °C, 95% humidity, 5% CO2 for 5 min. Detached cells were collected into a conical tube containing equal parts media to trypsin. Cells were centrifuged into a pellet at 3000 rpm, 4 °C for 1 min. The media/trypsin was aspirated and resuspended in fresh media. Then cells (passage 5-6) were counted using a hemocytometer with trypan blue exclusion and seeded on the 3D printed scaffolds at a density of 2.5×105 cells/scaffold. All cells were cultured at 37 °C, 95% humidity, 5% CO2 for up to 4 weeks and fed every 3 days. Viability Assay. Viability was evaluated after 1 day of cell culture using a Live/Dead viability/cytotoxicity kit (Invitrogen, UK). 20 µL of the 1 mM Calcein-AM stock solution and 10 µL of the 2 mM ethidium homodimer-1 (EtmD-1) stock solution were added to 10 mL of DPBS to prepare the Live/Dead staining solution. After aspiration of the old medium, samples were rinsed once with DPBS, and then 2 mL of the working solution was added to each well. Samples were incubated for 10 min at 37 °C before imaging at 4× magnification using CellSENS imaging software with an IX81fluorescence microscope (Olympus, Center Valley, PA) equipped with a Hamamatsu Orca R2 CCD camera and a filter cube containing 494 nm (green, Calcein) and 528 nm (red, EthD-1) emission filters. Images were analyzed for live/dead cell counts using ImageJ (NIH) software with a cell counter plugin. For quantitative analysis, a total of 200 cells were counted from each sample over 25 randomly chosen areas and the viable and non-viable cells counts were recorded. Cells stained green were counted as live and cells that stained red were


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counted as dead. Live and dead cell counts for all images per sample were totaled to calculate % viability for each sample Immunohistochemical Staining of Cytoskeletal Actin and Vinculin. Cell spreading was evaluated after 3 days of cell culture by immunohistochemical staining of actin and vinculin proteins. Cells on the scaffolds were fixed in pre-warmed 0.8 mL cell culture media and 1.2 mL 3.7% paraformaldehyde (PFA) in cytoskeletal stabilization (CS) buffer for 5 minutes at 37°C, and then fixed in 3.7 % PFA solution in PBS buffer at 37°C for 5 minutes. After washing with 1×PBS 3 times, cells were permeabilized using 2 mL of Triton X-100 in CS buffer (0.5% v/v) for 10 minutes at 37°C. The scaffolds were rinsed with 1×PBS 3 times at room temperature. Aldehyde fluorescence was then quenched using freshly made 0.1 wt% NaBH4 in 1×PBS for 10 minutes and non-specific binding was blocked using 5% donkey serum for 20 minutes at room temperature. After aspiration, the samples were incubated in vinculin primary antibody Mouse (Life Technology, monoclonal anti-vinculin antibody produced in mouse, county of origin Israel) in 1×PBS (v/v 1:200) at 4 °C overnight. After washing with 1% donkey serum 3 times, the samples were stained in a solution of rhodamine phalloidin (v/v 1:40) (Invitrogen, Eugene, OR) and Alexa Flour 488 secondary antibody Mouse (v/v 1:200) (Invitrogen, Eugene, OR) for 1 hour at room temperature avoiding light. After washing with 1×PBS 3 times, the nuclei were stained with DAPI (Invitrogen, Eugene, OR) in 1×PBS (6 µL/ 10 mL) for 20 minutes at room temperature in the dark. After washing with 1×PBS 3 times to remove excess staining, the samples were mounted and viewed under an IX81 fluorescence microscope equipped with a Hamamatsu Orca R2 CCD camera and filters of FITC, TRITC and DAPI. Images were analyzed for cell aspect ratio and cell area using Image J software. Cell aspect ratio was quantified using the cells greatest length divided by the diameter of the cell across the center of the nucleus. A


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total of 50 cells over 10 randomly chosen areas were used to calculate the average cell aspect ratio as well as cell area for each group. Cell Proliferation Assay. MC3T3-E1 cell proliferation on day 1, day 7 and day 14 was evaluated by CyQUANT cell proliferation assay kit (Invitrogen, UK). To generate a standard curve, a CyQUANT GR/cell-lysis buffer working solution was prepared by adding CyQUANT GR dye solution (Component A, v/v 1:400) and cell-lysis buffer (Component B, v/v 1:20) in nucleasefree distilled water. Serially diluted 200 µL of bacteriophage λ DNA with concentrations from 50 pg/mL to 1.0 µg/mL using CyQUANT GR/cell-lysis buffer working solution were added in the wells of a plastic 96-well microplate with a corresponding blank well. The fluorescence intensity by excitation at 480 nm and emission at 520 nm was measured and the blank absorbance value was subtracted from each reading. The standard curve was fitted with a linear relationship by plotting fluorescence intensity versus DNA concentration and the coefficient of determination (R2) was above 0.99. To determine the cell numbers, the scaffolds were taken out from the old medium, put in 1 mL of 1×cell-lysis buffer (Component B, v/v 1:20) and stored at -80 ºC at each time point (n=3 for each group at each time point). After thawing at room temperature, 200 µL of the thawed solution and 0.5 µL of CyQUANT GR dye solution (Component A, v/v 1:400) were added to each sample well in triplicate. After mixing gently under dark, the samples’ fluorescence was measured by plate reader (Synergy Biomax) by excitement at 480 nm and emission at 520 nm. The cell numbers were calculated from the observed fluorescence using the DNA standard curve and normalized to the cell numbers on Day 1. Alkaline Phosphatase (ALP) Activity Assay. ALP activity on week 2 and week 4 was measured by SensoLyte pNPP ALP assay kit (AnaSpec Inc, San Jose, CA, USA) following the provided protocol. The scaffolds were taken out from the old medium, put in 1 mL of lysis buffer (20 µL


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Triton X-100 in 10 mL 1× Assay Buffer) and stored at -80 ºC at each time point (n=3 for each group at each time point). After thawing at room temperature, the solution was homogenized by vortex and centrifuged at 2500×g for 10 min at 4°C. After centrifuge, the supernatant was collected for ALP and DNA quantification. To quantify the ALP activity, a standard curve was measured with an ALP solution at concentrations of 0, 3.1, 6.2, 12.5, 25, 50, 100 and 200 ng/mL. 50 µL of serially diluted ALP standard solution or 50 µL of supernatant was mixed with 50 µL pNPP solution in each well of a 96-well plate. The solution was mixed by gently shaking for 30 sec. After incubation under dark at room temperature for 1 h, 50 µL of stop solution (Component C) was added and the absorbance at 405 nm was measured by plate reader after shaking the 96-well plate for 1 min. Three replicates were measured for each sample. The standard curve was fitted with a linear relationship by plotting absorbance versus ALP concentration with a coefficient of determination (R2) above 0.98. The ALP activity of cell lysate from each sample was calculated with obtained equation. To account for the difference in cell numbers, the ALP activity was normalized with total amount of DNA which was quantified by CyQUANT cell proliferation assay kit (Invitrogen, UK). A standard curve was measure with a serially diluted 200 µL of bacteriophage λ DNA with concentrations from 50 pg/mL to 1.0 µg/mL using CyQUANT GR/cell-lysis buffer working solution. The fluorescence intensity by excitation at 480 nm and emission at 520 nm was measured and the blank absorbance value was subtracted from each reading. The standard curve was fitted with a linear relationship by plotting fluorescence intensity versus DNA concentration and the coefficient of determination (R2) was above 0.99. 200 µL of the supernatant and 0.5 µL of CyQUANT GR dye solution were added to each sample well in triplicate. After mixing gently


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under dark, the samples’ fluorescence was measured by plate reader (Synergy Biomax) by excitement at 480 nm and emission at 520 nm. The total amount of DNA was calculated from the observed fluorescence using the DNA standard curve. Quantitative Real Time Reverse Transcription Polymerase Chain Reaction (RT-qPCR). Total RNA was isolated from the MC3T3-E1 cell-seeded scaffolds on week 2 and week 4 using the RNeasy Mini Kit (Qiagen, Valencia, CA). In brief, scaffolds were taken out from the old medium, put in RNA protect cell reagent and stored at -80 ºC until used. After thawing at room temperature, the scaffolds were taken out and put into a micro-centrifuge with 350 µL of RLT Plus buffer (10 µL 2-mercaptoethanol in 1 mL RLT buffer). The scaffolds were torn up into small pieces with filaments exposed to the buffer solution. The lysate was homogenized by vortex and mixed well with 350 µL of 70% ethanol by pipetting. The lysate was transferred to the RNeasy spin column for total RNA isolation following the manufacturer’s protocol. DNase digestion was performed during RNA isolation using the Qiagen RNase free DNase set (Qiagen, Valencia, CA). Total RNA was quantified using a Take3 Multi Volume Plate and Synergy MX Microplate Reader (BioTek, Winooski, VT) at 260 nm. The 260 nm/280 nm ratio was used to determine purity of RNA with wells showing values of above 1.7 being used for RT-qPCR. RNA was stored at -80 ºC until used for reverse transcription. The TaqMan Reverse Transcription Reagents kit (Applied Biosystems, Life Technologies) was used for reverse transcription of mRNA into complementary DNA (cDNA) using 1000 ng total RNA as the template in a 100 µL reaction following the manufacturer’s protocol. The reverse transcription thermal protocol was conducted on a 2720 Thermal Cycler (Applied Biosystems) and consisted of incubation at 25 ºC for 10 min, reverse transcription at 48 ºC for 30 min, and inactivation at 95 ºC for 5 min. cDNA was stored at -20 ºC until qRT-PCR was performed. The qRT-PCR was performed on a 7500 Real Time PCR


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System (Life technologies, Grand Island, NY) using SYBR Green PCR Master Mix (Applied Biosystems, Thermal Fisher Scientific) and oligonucleotide primers (Integrated DNA Technologies). In brief, 50 µL reactions were prepared in PCR plates with 2× SYBR Green Master Mix, 209.4 nM forward primer, 209.4 nM reverse primer (Table S1) and 10 ng of cDNA. The remaining reaction volume was filled with DNase/RNase free H2O. For all genes, the thermal protocol consisted of reverse transcription at 50 ºC for 30 min, activation at 95 ºC for 15 min, and 50 amplification cycles of denaturing for 30 s at 95 ºC, annealing for 1.5 min at 58 ºC, and extension for 2 min at 72 ºC. Following amplification, a melt curve analysis was performed at 1ºC increments from 50 ºC to 95 ºC to analyze the purity of the product generated. qRT-PCR results were used for relative quantification of bone sialoprotein (BSP) and osteocalcin (OCN) using glyceraldehyde-3-phosphate dehydrogenase (GAPDH) as the endogenous reference gene (housekeeping gene). Relative expression of BSP and OCN was reported as a normalized gene expression value, 2-∆∆Ct, which was calculated using the following equation. ∆CT-cell control = CT-cell control, targeted gene – CT-cell control, GAPDH ∆CT-scaffold = CT-scaffold, targeted gene - CT-scaffold, GAPDH ∆∆C = ∆CT- scaffold - ∆CT-cell control Fold difference = 2- ∆∆C where CT-cell

control, targeted gene

is the average threshold value for the gene of interest on

undifferentiated cell control; CT

-cell control, GAPDH

is the average threshold value for the

housekeeping gene on undifferentiated cell control; CT-scaffold, targeted gene is the average threshold value for the gene of interest on differentiated cells on the scaffold; CT-scaffold, GAPDH is the average threshold value for the housekeeping gene on differentiated cells on the scaffold. All experiments were conducted at four replicates (n=4). All quantitative data are presented as the average ±


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standard deviation. Immunohistochemical Staining of Bone Sialoprotein (BSP) and Osteocalcin (OCN). Osteogenic differentiation was evaluated after 2 weeks and 4 weeks of cell seeding by immunohistochemical staining of bone sialoprotein (BSP) and osteocalcin (OCN). Cell-seeded scaffolds were fixed in pre-warmed 2 mL 3.7% paraformaldehyde (PFA) in PBS buffer for 2 hours at room temperature. After washing with 1×PBS 3 times, scaffolds were soaked in 70% EtOH 2 hours to dehydrate, processed into wax overnight using a tissue processor (Leica ASP300S, Leica Biosystems) and embedded in paraffin wax (Richard-Allan Scientific Type 9) for sectioning. Blocks were removed from -20 ºC, cut into 5 µm thick sections by microtome (Leica RM2255, Leica Biosystems). Sections were mounted on coated glass slides and dried at 40 ºC overnight before staining. Samples were put into 60 ºC oven for 1 h to obtain better attachment between sections and slides, and then rehydrated through a series of washes: xylenes (2×2min), 50% xylenes/50% EtOH (1×2 min), 100% EtOH (2×2 min), 95% EtOH (1×2 min), 70% EtOH (1×2 min), 50% EtOH (1×2 min) and deionized H2O (3×2 min). Immunohistochemistry samples were incubated in 0.5% pepsin reagent for 15 min at room temperature for antigen retrieval. After washing with 1×TBS buffer 3 times, slides were incubated in blocking buffer (10% donkey serum in 1×PBS) for 1 h at room temperature to block the non-specific binding. After washing with 1×TBS buffer 3 times, samples were incubated in bone sialoprotein (BSP) primary antibody Mouse (v/v 1:400) and osteocalcin (OCN) primary antibody Goat in 1×PBS (v/v 1:100) overnight at 4°C. After washing with 1×TBS buffer 3 times, samples were incubated in corresponding secondary antibodies conjugated to Alexa Fluor 488 (v/v 1:300 in 1×PBS) (Invitrogen, Eugene, OR) and Alexa Fluor 546 (v/v 1:300 in 1×PBS) (Invitrogen, Eugene, OR) for 1 h at room temperature in the dark. After washing with 1×TBS buffer 3 times, samples were incubated in DAPI solution (6


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µg /10 mL in 1×PBS) (Invitrogen, Eugene, OR) for 15 minutes at room temperature to stain the nuclei. After washing with 1×TBS buffer 3 times, samples were mounted and imaged using an IX81microscope (Olympus) equipped with Hamamatsu Orca R2 CCD camera and DAPI, FITC and TRITC fluorescence filters. Alizarin Red S. staining. The extents of mineralized extracellular matrix after 4 weeks were examined by Alizarin Red S. Staining. After rehydration of the sectioned slides, histological samples were incubated in freshly made Alizarin Red S. solution (2 g in 100 mL dd H2O, pH adjusted to 4.2) for 10 min, then washed and dehydrated through a series of solvents: deionized H2O (2×2min), 70% EtOH (1×2 min), 95% EtOH (1×2 min), 100% EtOH (1×2 min), and xylenes (2×2min), mounted by coverslip and imaged using an IX81microscope (Olympus) equipped with QImaging Micropublisher 3 camera under bright field. Calcium quantification. The scaffolds after 4 weeks of cell seeding were taken out from the old medium, washed with DPBS buffer (Mg2+, Ca2+ free) once, and put in 2 mL of 1×cell-lysis buffer. After 3 cycles of freeze-thaw to destroy the cell membrane, 400 µL of the supernatant was collected to quantify the total amount of DNA by CyQUANT cell proliferation assay kit (Invitrogen, UK) following the protocol. 4 mL of 1 M HCl was added to the 1.6 mL residue supernatant and the samples were agitated at room temperature for 4 hours. The concentration of calcium ions in the supernatant was measured using inductively coupled plasma optical emission spectrometry (ICP-OES) (Agilent Technologies 700 series, Santa Clara, CA, USA). The emission wavelength was set at 393.366 nm to quantify Ca2+. A standard curve was measured with solutions of Ca2+ concentration of 0.125, 0.25, 0.5, 1, and 2 ppm. Triplicate measurements were carried out for each sample. The calcium amount of each sample was normalized with total amount of DNA.


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Statistics. A one-way analysis of variance (ANOVA) with Tukey analysis was used to assess significance in the cell viability, cell spreading, cell proliferation data, ALP activity, real time RT-qPCR data and Ca2+ quantification to compare each sample type at all different time points. A significance value of p ≤ 0.05 with a 95% confidence interval or p ≤ 0.01 with a 99% confidence interval was set for statistical analysis (* indicates p value < 0.05, and ** indicates p value < 0.01). All quantitative data are presented as the average ± standard deviation.

RESULTS AND DISCUSSION Synthesis and Characterization of Composites. The degradable poly(ester urea) (PEU) polymer was synthesized in 2 steps from Lphenylalanine amino acid, 1,6-hexanediol and triphosgene using an interfacial polymerization as described in Figure 1(a).17,18 The chemical structures of the monomer and polymer were confirmed by 1H nuclear magnetic resonance (NMR) (Figure 1(b), Figure S1), 13C NMR (Figure S2), electrospray ionization (ESI) mass spectrometry (Figure S3) and fourier transform infrared spectroscopy (FT-IR) (Figure S6). The ester peak at 3.94 ppm, urea peak at 6.47-6.5 ppm in the 1

H NMR and carbonyl (C=O ester) stretching at 1735-1750 cm-1, N-H urea stretching at 3350-

3500 cm-1 in the FT-IR spectra demonstrated the successful formation of ester bonds and urea groups. The molecular mass, molecular mass distribution and thermal properties of the polymer were measured by size exclusion chromatography (SEC), thermogravimetric analysis (TGA) and differential scanning calorimetry (DSC) respectively. High molecular mass polymer (Mw 100 kDa) was synthesized through a step growth polymerization and a narrower molecular distribution (ƉM 1.79) was obtained than the theoretical value of ƉM 2.0 due to a postprecipitation and fractionation during precipitation. The TGA results showed the high thermal


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stability of the polymer. The degradation temperature (Td, 286 ºC) is significantly higher than the glass transition temperature (Tg, 54 ºC) which allows for high temperature processing techniques with limited degradation. Furthermore, the polymer is soluble in polar organic solvents, including N,N-dimethylformamide (DMF), dimethyl sulfoxide (DMSO), N-methyl-2-pyrrolidone (NMP) and hexafluoroisopropanol (HFIP). Hydroxyapatite (HA) was synthesized from mixing calcium and phosphate solutions using a hydrothermal method as described in Figure 1(d).50,51 The hexagonal crystalline structure was confirmed by wide angle X-ray diffraction measurement (XRD) as shown in Figure 1(c). The pattern showed diffraction peaks corresponding to the characteristic peaks of synthetic standard HA.52 The peaks occurred at 26º, 32º, 40º, 47º, 50º, 53º and 64º referring respectively to (002), (211), (130), (222), (213), (004) and (304) faces of the HA crystals. The Ca/P ratio was measured by a survey scan of X-ray photoelectron spectroscopy (XPS) and inductively coupled plasma atomic emission spectrometry (ICP-AES) (Figure S4). The XPS elemental analysis results showed phosphorous peaks at 132 eV (P 2p), 190 eV (P 2s) and calcium peaks at 346 eV (Ca 2p), 438 eV (Ca 2s). The integration of the area under the peaks gave a Ca/P ratio of 1.6 which is close to the ideal Ca/P ratio of 1.67 in stoichiometric HA. The needle-like morphology of the synthesized HA was measured using scanning electron microscope (SEM) (Figure S4) and transmission electron microscope (TEM) (Figure 1(e)) with dimensions of ~200 nm in length and ~10 nm in width. A series of PEU and HA composites (PEU-0% HA, PEU-10% HA, PEU-20% HA, PEU-30% HA and PEU-40% HA) were fabricated by solution mixing in HFIP with a weight ratio of 100:0, 90:10, 80:20, 70:30,60:40 respectively. The increased intensity of P-O stretching at 1030 cm-1 in the IR spectra (Figure S6) and enhanced crystalline signal of HA in the XRD patterns (Figure 7)


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indicated the presence of HA in the composites. The HA content present in the composites was quantified by the residual weight% at 550 ºC from TGA (Figure S5) as HA is the only component that does not decompose at that temperature. The experimental determined HA content shown from TGA are close to the feed ratio, indicating that the amount of HA added into the composite can be controlled precisely. The DSC (Figure S8, Table 1) results showed that the glass transition temperatures of the composites are higher than 37 ºC even though there is a slight decrease as the HA content increases. The high thermal stability of the composites (Td > 280 ºC) and glass transition temperatures suggest the composites are suitable for scaffold fabrication via 3D printing.

Fabrication and Characterization of Composite Scaffolds One of the major design challenges for 3D printed scaffolds is to mimic the structure and biological functions of the natural ECM. The ideal scaffolds are thought to be three-dimensional and porous with an interconnected pore network that facilitates the transport of nutrients and metabolic waste and affords tissue integration and vascularization.8 In this work, highly porous composite scaffolds were fabricated by fused deposition modeling (FDM). FDM is a rapid prototyping technology used to fabricate three-dimensional scaffolds by extruding melt materials layer by layer. In the setup shown in Figure 2(a), the filament of each composite was fabricated using extrusion via capillary rheometer at 165 ºC with a diameter of 1.75 mm which can be fed into the printing nozzle. The composite material was deposited through a 0.3 mm nozzle and printed layer by layer in the X and Y plane with a printing speed of 2 mm/s. The stage moved down in the Z axis. The resulting geometry of the scaffolds was a series of rods layered perpendicularly with a porosity of 75%. The dimensions of the scaffolds were ~7.7 mm in


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diameter and ~1.5 mm in thickness (Figure 2(c)). The morphology and microstructural formability of the filament and 3D printed porous scaffolds were characterized nondestructively using X-ray micro-computed tomography (µCT). The µCT images (Figure 2(b), Figure S11) and SEM images (Figure S9) of the filaments showed that the HA was homogeneously distributed within the filament. There was no significant aggregation and an enhancement in scattering length density was observed when increasing the HA content which could help tracking the scaffolds when implanted in vivo. The µCT images of the scaffolds (Figure 2(d), Figure S10) demonstrated that these composite scaffolds were homogeneous with highly reproducible spatial arrangement of pores and channels. The pore size and diameter of the microfilament were ~320 µm and ~300 µm respectively which were consistent with the printing parameters of the original design and nozzle size. Previous studies have shown that larger pore sizes (200-400 µm) are required for bone regeneration to facilitate cell penetration into the scaffold.20,53 Smaller pores (75-100 µm) have been shown to result in unmineralized osteoid tissue.54,55 Therefore, the large void volume of these 3D printed composite scaffolds could support for proper osteogenic differentiation as well as cell penetration. The amount of porosity was around 75% as calculated from 3D reconstructed models which was consistent with the printing design. The 3D models obtained from the µCT images (Figure S10) revealed the complete interconnectivity and integrity of the three-dimensional porous scaffolds which facilitates the flow transport of nutrients and metabolic wastes as well as migrations of cells through the entire scaffold construct. These microstructural characteristics and incorporation of HA also impart hydrophilicity to the scaffolds. Additionally, the 3D printing speed is sufficient to generate anatomically scaled and patient-specific grafts, sterilize and prepare for use in hours. The linear print speeds could be achieved up to 25 cm/s (instrument limitation) with no more post printing process required


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before getting the final product. This linear deposition rate equates to volume deposition rates as high as 80 cm3/hour from one single 0.3 mm nozzle or total object volume of 320 cm3/hour when the architecture is 75% porous. The production time that it takes to fabricate scaffolds could be dramatically decreased by automation. Before further investigation, it is necessary to determine the actual content of HA within the composite scaffolds. During the filament fabrication or 3D printing process, it is possible that some amounts of HA particles could be lost due to the nozzle size or rheological effects. The mass% at 550 ºC in the TGA is considered to be the actual amount of HA within the composite scaffolds. For the HA/PEU scaffolds that were prepared with the initial HA content of 0, 10, 20, 30 and 40% w/w, the actual HA contents (Table 1) were determined to be 0.5, 10.3, 18.9, 27.1 and 38.2% w/w, respectively in the filament; 0.6, 10.9, 19.1, 29.9 and 38.9% w/w, respectively in the scaffold; 1.5, 11.9, 21.3, 30.3 and 40.1% w/w, respectively in the decellularized scaffolds after 4 weeks of cell culture. The TGA results validated that the HA weight percentage in the filament, in the 3D printed scaffolds and in the scaffolds following 4 weeks of cell culture remained identical indicating there was no loss of HA during processing and cell culture. One of the major challenges in developing load-bearing scaffolds for bone tissue engineering is the conflict between material porosity and mechanical strength. A highly porous structure (porosity > 70%) is necessary for cell growth and proliferation but results in a decrease of mechanical strength. Although there is no clearly defined criteria between porosity and mechanical strength required for bone tissue engineering, it is generally regarded that the scaffold should have mechanical properties similar to that of natural bones.56,57 The mechanical properties of these composite scaffolds were measured via uniaxial compression tests using Instron with a Hencky strain of 0.1 min-1 at three different conditions: tested at room


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temperature, physiological temperature and tested after 1 week of incubation in PBS. When the scaffolds were compressed in the Z direction of 3D printing process (Figure 3), the filament junctions of adjacent layers primarily supported the applied load. In that process, an initial linear elastic deformation was observed and significant shear deformation of the filament joints was also involved. As the compression strain increased, the linear elastic regime was truncated by the sliding/stretching of filament layers which results in a decrease of slope in the stress-strain curve (Figure S12). On further compression, the scaffolds underwent densification and an increase of slope was obtained. The mechanical properties of the scaffolds would get closer to the property of the bulk materials until failure. The compressive moduli were determined using the slope of linear fitting in the beginning linear regime and reported in Figure 3 with the average values from three individual measurements. The 3D printed scaffolds showed an averaged compressive modulus of 65-85 MPa when tested at room temperature. The HA content does not appear to significantly influence the mechanical properties during compression. This is because of the balance between the reinforcement effect and unoptimized filler/matrix interaction. HA is very stiff which could reinforce the composite and enhance the mechanical properties of the material. However, it is also possible to observe a decrease of modulus with an increase of HA content especially if the polymer/particle interface is not optimized. A higher effect of stress concentration induced by the presence of HA particles could become more relevant when increasing the HA concentration. When the testing temperatures increased from room temperature to physiological temperature, the compressive moduli dropped slightly (49-65 MPa) as the temperature approached to the glass transition temperatures of the composites. When the tests were repeated at physiological temperature after 1 week of phosphate buffered saline solution incubation, lower values of compressive moduli were observed (38-54 MPa). This can


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be explained by the weakening of the interface and the plasticization effect due to the presence of water. Even though there was a slight decrease of moduli when the scaffolds were wet, these 3D printed porous scaffolds approached the stiffness of the natural cancellous bone (50-100 MPa)58,59 and exhibited a significantly higher compressive modulus compared to the salt-leached 80% porous PEU scaffolds (1-3 MPa); 3D printed 72% porous poly(Ɛ-caprolactone) (PCL) scaffolds (~27 MPa)60; 3D printed 75% porous poly(lactic-co-glycolic acid) (PLGA)/HA (10:90 w/w) (~4 MPa)61, as the structure and perfect fusion between filaments at the joints strengthened the mechanical properties of the scaffolds. Even though the solid volume fraction was only 25% of the total construct, the 3D printed scaffolds can hold more than 6000 N (~500 MPa) without yield and fracture which is much greater mechanical loads than the load bearing capacity of human cortical bone (~190 MPa).62 This is significant as the highly porous scaffolds show sufficiently high strength which lower the chance of implant failure due to mechanical property mismatch and are translationally relevant to craniofacial and other load bearing applications.

in vitro Culture of MC3T3-E1 Cells on Composite Scaffolds. In vitro experiments were performed to assess the ability of 3D printed PEU-0% HA, PEU10% HA, PEU-20% HA, PEU-30% HA and PEU-40% HA scaffolds to support MC3T3-E1 preosteoblast cell adhesion, proliferation and induce osteogenic differentiation and mineralization in the absence of osteogenic growth factors and peptides. MC3T3-E1 pre-osteoblast cells, which have been frequently used for elucidating the responses of bone cells to biomaterials63,64, were seeded on the composite materials with a cell density of 3750 cells/cm2. After 24 h of cell culture, the dominant green fluorescence from live cells using live/dead assay (Figure 4(a)) demonstrated the materials are nontoxic. The MC3T3-E1 cell


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viabilities on these composites (Figure 4(c)) are higher than 95% and no significant difference was observed when increasing the HA content. After 3 days of cell culture, the MC3T3-E1 cells were fluorescently stained to visualize actin (green) and vinculin (red) to assess the organization of cytoskeleton and spatial distribution of focal adhesion contacts. The cells were attached on the substrate and the focal adhesion contacts between cells and substrates formed as shown in Figure 4(b). Quantification of cell spread area and aspect ratio (Figure 4(d)) revealed that there is no significant statistic difference in cell area and aspect ratio after 3 days of cell culture indicating that HA has no effects in the adhesion of MC3T3-E1 cells onto scaffolds in 3 days of cultivation. Also, cells spread comparably with those cultured on glass (Figure S13) which means cells put down attachments and stretched out on the composite materials as they normally would on glass substrates. The growth of MC3T3-E1 cells on the three-dimensional porous scaffolds was studied by CyQUANT proliferation assay on day 1, 7 and 14 of cell culture (Figure 5(a)). 2.5 ×105 MC3TCE1 cells were seeded on each scaffold and the change of cell numbers was recorded. From day 1 to day 7, the cells were mostly attaching to the surface, adjusting to the new environment and starting to proliferate, therefore, the cell number increased ~4-fold at similar proliferation rates on day 7. There is no significant difference in cell numbers among scaffolds with different HA contents. This similar ascendant tendency of the cell population demonstrated that the HA scaffolds do not have significant influence on cell growth in the short term, which is consistent with the cell spreading data on day 3. From day 7 to day 14, the MC3TC-E1 cells continued to proliferate. All of the samples supported cell proliferation without exhibiting toxic effects through 14 days of in vitro culture which demonstrate that the architecture of the scaffold is suitable for osteoblast seeding and growth. The cells on PEU-30% HA scaffolds showed the


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highest cell number compared with others indicating these scaffolds are more conductive to MC3T3-E1 cell proliferation (p < 0.05). This is likely due to increased roughness or the fact that HA may contribute to the integrin-mediated osteoblast adhesion from day 7 to day 14. Integrins connect the cell’s cytoskeleton and surrounding ECM and initiate intracellular signaling pathways to nucleate signaling proteins including focal adhesion kinase.65 HA also functions as an inductive substrate for osteoblast adhesion, promotes cell surface integrin presentation and clustering, modulate formation of focal contacts which serve to promote cell growth.66 HA recruits cell growth factors via adsorption processes such as bone morphogenic protein (BMP)like factors from complex serum to yield an osteoinductive and osteoconductive microenvironment.67 When implanted in vivo the protein/growth factors in the body fluid and circulation adsorb on the HA surface and promote adhesion of osteoblasts. The adsorbed factors leads to the sequential expression of integrins, FAK and ERK genes required for cell attachment that is followed by the expected proliferative and cell differentiation gene expression stages. 68 Following cell proliferation, MC3TC-E1 cells enter early stage of differentiation. In that process, the extracellular matrix is deposited and become matured. Early differentiation of MC3T3-E1 cells is marked by the expression of alkaline phosphatase (ALP). ALP is an essential enzyme for cell mineralization that hydrolyzes the calcification inhibitor.

In this study, a

standard colorimetric assay was performed at week 2 and week 4 to quantify the ALP activity and the ALP activity values were normalized with total amount of DNA to account for the difference in cell numbers. As shown in Figure 5(b), there was an increase of ALP activity from 2 weeks to 4 weeks due to maturation of osteoblasts suggesting the cells have approached or passed the peak time as the expression of ALP would be down-regulated after mineralization starts.69. In addition, the HA at higher concentrations exhibited an enhancement effect on the


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ALP activity. This up-regulation of ALP activity indicated that the HA preserves its ability to stimulate the dephosphorylation which is an essential activity involved in the mineralization process. By 2 weeks, the PEU-30% HA scaffold showed a 1.5-fold higher ALP expression of MC3T3-E1 cells compared to PEU-0% HA scaffold (p < 0.01). By 4 weeks, the ALP activity of MC3T3-E1 cells was significantly increased in PEU-30% HA scaffold with values reaching 2.5fold greater compared to the PEU-0% HA (p < 0.05). The resultant higher proliferation level on the PEU-30% HA scaffold potentially provided faster cell-to-cell interaction which could promote subsequent osteogenic differentiation.70 This explains the shift to a more differentiated state of cells and better bone forming ability in the PEU-30% HA scaffolds. The osteogenic differentiation was characterized by tracking the temporal expression of osteogenic markers: bone sialoprotein (BSP) and osteocalcin (OCN). BSP is a multifunctional protein that promotes cell adhesion and induces mineralization. The gene expression of BSP in the seeded cells on the composite scaffolds was analyzed at both 2 weeks and 4 weeks by real time quantitative reverse transcription polymerase chain reaction (RT-qPCR). The results in Figure 6(a) indicate that the expression of BSP mRNA in the cells cultured on the PEU-0% HA, PEU-10% HA, PEU-20% HA, PEU-30% HA and PEU-40% HA scaffolds was upregulated by 130-fold, 6-fold, 4-fold, 2-fold and 1.3-fold at week 4, respectively compared to that at week 2. In addition, the expression levels of BSP mRNAs significantly increased in cells cultured on scaffolds with higher HA contents than those in cells cultured on PEU-0% HA scaffolds at both 2 weeks and 4 weeks, which showed the advantages of the osteoinductivity imparted by HA. By week 2, there was a 6900-fold, 22,000-fold, 69,000-fold and 61,000-fold greater expression of BSP in MC3T3-E1 cells on the PEU-10% HA, PEU-20% HA, PEU-30% HA and PEU-40% HA scaffolds, respectively, compared to that on the PEU-0% HA scaffolds (p < 0.01). By week 4,


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there was a 320-fold, 690-fold, 880-fold and 620-fold higher expression of BSP in the cells on the PEU-10% HA, PEU-20% HA, PEU-30% HA and PEU-40% HA scaffolds, respectively, compared to that on the PEU-0% HA scaffolds (p < 0.01). In Figure 6(c), the immunohistochemistry staining has shown the cells on the scaffolds secreted abundant amounts of BSP proteins at both 2 weeks and 4 weeks as indicated by the strong green fluorescence. There was more expression of BSP on the scaffolds cultured for 4 weeks than 2 weeks and higher expression on the scaffolds at higher concentration of HA. This suggests the HA upregulates the osteogenic differentiation of MC3T3-E1 cells at protein levels as well as confirms our results from RT-qPCR. Peter Ma and his co-workers also reported the higher cell numbers and greater expression of BSP mRNA of MC3T3-E1 cells on 89% porous poly(L-lactic acid) (PLLA)/HA (1:1 w/w) composite scaffold compared to pure 92% porous PLLA scaffolds up to 6 weeks.71 BSP has been shown to bind to HA through its polyglutamic acid residues and mediates cell attachment through an RGD sequence (Arg-Gly-Asp), nucleate new HA crystal formation, promote pre-osteoblast cells differentiate into mature osteoblast and ultimately stimulating bone mineralization.72 Therefore, the increased expression of BSP affords cell attachment to the surface of the scaffold and enhanced osteogenic differentiation. OCN is a late stage marker expressed when cells have reached mature osteoblast stage. It is secreted solely by osteoblasts so it is a very specific protein for osteoblast differentiation and mineralization. The RT-qPCR results in Figure 6(b) have shown the expression of OCN mRNA in the cells cultured on the PEU-0% HA scaffolds was upregulated by 260-fold at week 4 compared to that at week 2, while lower expression of OCN on the PEU-10% HA, PEU-20% HA, PEU-30% HA and PEU-40% HA scaffolds was observed compared to that at week 2, indicating the cells have reached mature osteoblast stage within the first 2 weeks. In addition,


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there was a significant greater expression of OCN in cells on the scaffolds with higher HA contents. By week 2, there was a 16,000-fold, 9200-fold, 12400-fold and 11,900-fold greater expression of OCN in MC3T3-E1 cells on the PEU-10% HA PEU-20% HA PEU-30% HA and PEU-40% HA scaffolds, respectively, compared to that on the PEU-0% HA scaffolds (p < 0.01). By week 4, there was a 4.5-fold, 7.3-fold, 15.6-fold and 11.5-fold higher expression of OCN in the cells on the PEU-10% HA, PEU-20% HA, PEU-30% HA and PEU-40% HA scaffolds compared to that on the PEU-0% HA scaffolds (p < 0.01). In Figure 4(c), the immunohistochemistry staining has shown the expression of OCN proteins in the cytoplasm at both 2 weeks and 4 weeks as indicated by the strong red fluorescence. There was more expression of OCN on the scaffolds with higher HA content. Studies have indicated that integrinextracellular matrix interactions may play a key role in the regulation of the osteocalcin genes and HA could promote cell surface integrin presentation.66,73 This explains the upregulation of OCN on the PEU-30% HA scaffold by week 4. Calcium deposition occurs during the final stages of osteogenic differentiation. The extent of mineralized extracellular matrix after 4 weeks of cell culture was examined by Alizarin Red S. staining, a red dye that forms chemical complex with the calcium ions. As shown in Figure 7(a) and 7(b), there was pigmentation of each microfilament with red color due to the presence of HA and calcium deposition on the surface of each microfilament with dark red color indicating the MC3T3-E1 cells differentiated to matured osteoblast and secreted mineralized extracellular matrix. The calcium content in the mineralized extracellular matrix on the surface of the scaffolds was quantified by ICP-AES. The concentration of Ca2+ ion in the cell cultured scaffolds substrates the concentration of HA in the scaffolds, divided by the total amount of DNA to account for the difference in cell numbers. The results listed in Figure 7(c) have shown that the


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PEU-10% HA, PEI-20% HA, PEU-30% HA and PEU-40% HA scaffolds exhibited 5.8-fold, 21.9-fold, 60.3-fold and 185-fold higher concentration of calcium ions, respectively, compared to that on the PEU-0% HA scaffolds (p < 0.01). Therefore, the scaffolds with higher HA content could promote the mineralization of MC3T3-E1 cells. results are consistent with previous work that reports pre-incorporation of HA in a nanofiber scaffolds promotes the initial rate of apatite deposition.74

4. CONCLUSION Three-dimensional porous scaffolds were fabricated using FDM techniques with custom nanoHA PEU filaments without further chemical modification of their surface. These highly porous HA/PEU composite scaffolds are strong and possess architectures (~75% porosity and ~300 µm pore size) suitable for MC3T3-E1 cell growth, differentiation and maturation. The mechanical properties of the 3D printed composite scaffolds (65-85 MPa) are significantly higher than those of polymer scaffolds fabricated using other techniques such as salt leaching and those of other 3D printed pure polymer scaffolds. The long-term cell survival, cell growth, highly differentiated state by improving ALP activity and expression of osteoblast genes and proteins as well as enhanced mineralization in the scaffolds with higher HA contents demonstrated the HA impart osteoinductivity and osteoconductivity to the scaffold. This is a viable approach for designing and fabricating novel biomaterial formulations that enhance cell proliferation and differentiation without growth factors or peptides. The combination of synthesis and processing with tailored structure and optimal property to direct subsequent functions of cells make these polymer/HA composite scaffolds superior to other polymer scaffolds for osseous tissue engineering. More advanced preclinical studies on the composite scaffold degradation and in vivo tissue formation


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are ongoing to understand explore the translational potentials of these materials for clinical application.

ASSOCIATED INFORMATION Supporting Information The supporting Information is available free of charge on the ACS Publications website at DOI: xxx. 1

H NMR (DMSO-d6),

13

C NMR (DMSO-d6), ESI of di-p-toluene sulfonic acid salts of bis-L-

phenylalanine-hexane-1,6-diester monomer and L-phenylalanine based poly(ester urea); XPS and SEM of HA; TGA, IR, XRD, DSC of PEU/HA composites; dispersion of HA in PEU; exemplar µCT images of PEU-20% HA scaffold; exemplar 2D XRD of PEU-40% HA microfilament; exemplar stress vs strain curve of PEU-20% HA scaffold; cell spreading of MC3T3-E1 cells on composite scaffolds and RT-qPCR primers.

AUTHOR INFORMATION Corresponding Author *E-mail: [email protected] ORCID Jiayi Yu: 0000-0003-3697-9137 Matthew L. Becker: 0000-0003-4089-6916 Notes The authors declare no competing financial interest.

ACKNOWLEDGEMENTS


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The authors are grateful to Thomas J. Quick for the help in ICP-AES tests, Shichen Yuan for the help in the XRD measurements and Dr. Nikolov Zhorro for the help with XPS characterization. This work was supported by The United States Department of Defense USAMRMC (Project ID, W81XWH-15-1-0718). MLB acknowledges support from the W. Gerald Austen Professor in Polymer Science and Polymer Engineering endowed by the Knight Foundation.

REFERENCES

(1) Patel, Z. S.; Young, S.; Tabata, Y.; Jansen, J. A.; Wong, M. E. K.; Mikos, A. G. Bone 2008, 43, 931. (2) Bucholz, R. W. Clinical Orthopaedics and Related Research 2002, 395, 44. (3) Cancedda, R.; Giannoni, P.; Mastrogiacomo, M. Biomaterials 2007, 28, 4240. (4) Kneser, U.; Schaefer, D. J.; Polykandriotis, E.; Horch, R. E. Journal of Cellular and Molecular Medicine 2006, 10, 7. (5) Puppi, D.; Chiellini, F.; Piras, A. M.; Chiellini, E. Progress in Polymer Science 2010, 35, 403. (6) Finkemeier, C. G. JBJS 2002, 84, 454. (7) Gazdag, A. R.; Lane, J. M.; Glaser, D.; Forster, R. A. JAAOS - Journal of the American Academy of Orthopaedic Surgeons 1995, 3, 1. (8) Burg, K. J. L.; Porter, S.; Kellam, J. F. Biomaterials 2000, 21, 2347. (9) Zhang, Q.; Mochalin, V. N.; Neitzel, I.; Knoke, I. Y.; Han, J.; Klug, C. A.; Zhou, J. G.; Lelkes, P. I.; Gogotsi, Y. Biomaterials 2011, 32, 87. (10) Prabhakaran, M. P.; Venugopal, J.; Ramakrishna, S. Acta Biomaterialia 2009, 5, 2884. (11) Zheng, J.; Hua, G.; Yu, J.; Lin, F.; Wade, M. B.; Reneker, D. H.; Becker, M. L. ACS Macro Letters 2015, 4, 207. (12) Lin, F.; Zheng, J.; Yu, J.; Zhou, J.; Becker, M. L. Biomacromolecules 2013, 14, 2857. (13) Hakkarainen, M.; Höglund, A.; Odelius, K.; Albertsson, A.-C. Journal of the American Chemical Society 2007, 129, 6308. (14) Ignatius, A. A.; Claes, L. E. Biomaterials 1996, 17, 831. (15) Athanasiou, K. A.; Niederauer, G. G.; Agrawal, C. M. Biomaterials 1996, 17, 93. (16) Lin, F.; Yu, J.; Tang, W.; Zheng, J.; Xie, S.; Becker, M. L. Macromolecules 2013, 46, 9515. (17) Stakleff, K. S.; Lin, F.; Smith Callahan, L. A.; Wade, M. B.; Esterle, A.; Miller, J.; Graham, M.; Becker, M. L. Acta Biomaterialia 2013, 9, 5132. (18) Yu, J.; Lin, F.; Lin, P.; Gao, Y.; Becker, M. L. Macromolecules 2014, 47, 121. (19) Li, S.; Yu, J.; Wade, M. B.; Policastro, G. M.; Becker, M. L. Biomacromolecules 2015, 16, 615. (20) Policastro, G. M.; Lin, F.; Smith Callahan, L. A.; Esterle, A.; Graham, M.; Sloan Stakleff, K.; Becker, M. L. Biomacromolecules 2015, 16, 1358.


Page 33 of 42

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Biomacromolecules

(21) Yu, J.; Lin, F.; Becker, M. L. Macromolecules 2015, 48, 2916. (22) Chen, K.; Yu, J.; Guzman, G.; Es-haghi, S. S.; Becker, M. L.; Cakmak, M. Macromolecules 2017, 50, 1075. (23) Li, S.; Xu, Y.; Yu, J.; Becker, M. L. Biomaterials 2017, 141, 176. (24) Dorozhkin, S. V. Biomaterials 2010, 31, 1465. (25) LeGeros, R. Z. Clinical Orthopaedics and Related Research 2002, 395, 81. (26) Ducheyne, P.; Qiu, Q. Biomaterials 1999, 20, 2287. (27) Weiner, S.; Wagner, H. D. Annual Review of Materials Science 1998, 28, 271. (28) Du, C.; Cui, F. Z.; Zhu, X. D.; de Groot, K. Journal of Biomedical Materials Research 1999, 44, 407. (29) Du, C.; Cui, F. Z.; Feng, Q. L.; Zhu, X. D.; de Groot, K. Journal of Biomedical Materials Research 1998, 42, 540. (30) Cooke, F. W. Clinical Orthopaedics and Related Research 1992, 276, 135. (31) Rho, J.-Y.; Kuhn-Spearing, L.; Zioupos, P. Medical Engineering & Physics 1998, 20, 92. (32) Webster, T. J.; Siegel, R. W.; Bizios, R. Biomaterials 1999, 20, 1221. (33) Adam, E. J.; Alexandra, L. R.; Ramille, N. S. Biomedical Materials 2016, 11, 014102. (34) Grasman, J. M.; Zayas, M. J.; Page, R. L.; Pins, G. D. Acta Biomaterialia 2015, 25, 2. (35) Chen, W.; Zhou, H.; Tang, M.; Weir, M. D.; Bao, C.; Xu, H. H. K. Tissue Engineering. Part A 2012, 18, 816. (36) Deville, S.; Saiz, E.; Tomsia, A. P. Biomaterials 2006, 27, 5480. (37) Thomson, R. C.; Yaszemski, M. J.; Powers, J. M.; Mikos, A. G. Biomaterials 1998, 19, 1935. (38) Travitzky, N.; Bonet, A.; Dermeik, B.; Fey, T.; Filbert-Demut, I.; Schlier, L.; Schlordt, T.; Greil, P. Advanced Engineering Materials 2014, 16, 729. (39) Heinemann, S.; Rössler, S.; Lemm, M.; Ruhnow, M.; Nies, B. Acta Biomaterialia 2013, 9, 6199. (40) Akkineni, A. R.; Luo, Y.; Schumacher, M.; Nies, B.; Lode, A.; Gelinsky, M. Acta Biomaterialia 2015, 27, 264. (41) Zein, I.; Hutmacher, D. W.; Tan, K. C.; Teoh, S. H. Biomaterials 2002, 23, 1169. (42) Childers, E. P.; Wang, M. O.; Becker, M. L.; Fisher, J. P.; Dean, D. MRS Bulletin 2015, 40, 119. (43) Wang, M. O.; Vorwald, C. E.; Dreher, M. L.; Mott, E. J.; Cheng, M.-H.; Cinar, A.; Mehdizadeh, H.; Somo, S.; Dean, D.; Brey, E. M.; Fisher, J. P. Advanced Materials 2015, 27, 138. (44) Wen, Z.; Feng, L.; Tingchun, S.; Renji, Z.; Yongyan, N.; Jie, R.; Tianrui, Z. Rapid Prototyping Journal 2008, 14, 280. (45) Tellis, B. C.; Szivek, J. A.; Bliss, C. L.; Margolis, D. S.; Vaidyanathan, R. K.; Calvert, P. Materials Science and Engineering: C 2008, 28, 171. (46) Meakin, J. R.; Shepherd, D. E. T.; Hukins, D. W. L. The British Journal of Radiology 2004, 77, 504. (47) Shor, L.; Güçeri, S.; Wen, X.; Gandhi, M.; Sun, W. Biomaterials 2007, 28, 5291. (48) Vorndran, E.; Klammert, U.; Ewald, A.; Barralet, J. E.; Gbureck, U. Advanced Functional Materials 2010, 20, 1585. (49) Pang, X.; Chu, C.-C. Biomaterials 2010, 31, 3745. (50) Koutsopoulos, S. Journal of Biomedical Materials Research 2002, 62, 600. (51) Eanes, E. D.; Gillessen, I. H.; Posner, A. S. Nature 1965, 208, 365.


Biomacromolecules

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

(52) Lu, H. B.; Campbell, C. T.; Graham, D. J.; Ratner, B. D. Analytical Chemistry 2000, 72, 2886. (53) Kasugai, S.; Nagata, T.; Sodek, J. Journal of Cellular Physiology 1992, 152, 467. (54) Boyan, B. D.; Hummert, T. W.; Dean, D. D.; Schwartz, Z. Biomaterials 1996, 17, 137. (55) Leong, K. F.; Cheah, C. M.; Chua, C. K. Biomaterials 2003, 24, 2363. (56) Ma, P. X. Materials Today 2004, 7, 30. (57) Hutmacher, D. W. Biomaterials 2000, 21, 2529. (58) Yaszemski, M. J.; Payne, R. G.; Hayes, W. C.; Langer, R.; Mikos, A. G. Biomaterials 1996, 17, 175. (59) Gerhart, T. N.; Renshaw, A. A.; Miller, R. L.; Noecker, R. J.; Hayes, W. C. Journal of Biomedical Materials Research 1989, 23, 1. (60) Hoque, M. E.; San, W. Y.; Wei, F.; Li, S.; Huang, M.-H.; Vert, M.; Hutmacher, D. W. Tissue Engineering Part A 2009, 15, 3013. (61) Jakus, A. E.; Rutz, A. L.; Jordan, S. W.; Kannan, A.; Mitchell, S. M.; Yun, C.; Koube, K. D.; Yoo, S. C.; Whiteley, H. E.; Richter, C.-P.; Galiano, R. D.; Hsu, W. K.; Stock, S. R.; Hsu, E. L.; Shah, R. N. Science Translational Medicine 2016, 8, 358ra127. (62) Zioupos, P.; Currey, J. D. Bone 1998, 22, 57. (63) Quarles, L. D.; Yohay, D. A.; Lever, L. W.; Caton, R.; Wenstrup, R. J. Journal of Bone and Mineral Research 1992, 7, 683. (64) Beck, G. R.; Sullivan, E. C.; Moran, E.; Zerler, B. Journal of Cellular Biochemistry 1998, 68, 269. (65) Gronthos, S.; Simmons, P. J.; Graves, S. E.; G. Robey, P. Bone 2001, 28, 174. (66) Gajjeraman, S.; He, G.; Narayanan, K.; George, A. Advanced Functional Materials 2008, 18, 3972. (67) Kodaira, K.; Imada, M.; Goto, M.; Tomoyasu, A.; Fukuda, T.; Kamijo, R.; Suda, T.; Higashio, K.; Katagiri, T. Biochemical and Biophysical Research Communications 2006, 345, 1224. (68) Rouahi, M.; Champion, E.; Hardouin, P.; Anselme, K. Biomaterials 2006, 27, 2829. (69) Golub, E. E.; Boesze-Battaglia, K. Current Opinion in Orthopaedics 2007, 18, 444. (70) Aubin, J. E.; Liu, F.; Malaval, L.; Gupta, A. K. Bone 1995, 17, S77. (71) Ma, P. X.; Zhang, R.; Xiao, G.; Franceschi, R. Journal of Biomedical Materials Research 2001, 54, 284. (72) Kirkham, G.; Cartmell, S. Ashammakhi N, Reis R, Chiellini E, editores. Topics in Tissue Engineering 2007, 3. (73) Kirkham, G. R.; Cartmell, S. H. In Topics in Tissue Engineering.; Ashammakhi, N., Reis, R., Chiellini, E., Eds. 2007; Vol. 3. (74) Wei, G.; Ma, P. X. Journal of Biomedical Materials Research Part A 2006, 78A, 306.


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Figure 1. Synthesis and characterization of poly(ester urea) (PEU) and hydroxyapatite (HA). (a) A twostep general synthetic route of L-phenylalanine based PEU using interfacial polymerization. (b) 1

H NMR (DMSO-d6) of L-phenylalanine based PEU. The ester peak at 3.94 ppm and urea peak at 6.47-6.5 ppm showed the successful formation of ester bonds and urea groups. (d) The synthetic route of HA. (c) The X-ray diffraction pattern of HA showed the highly crystalline hexagonal structure of HA. (e) TEM images showed the needle-like morphology of the synthesized HA with around 200 nm in length and 10 nm in width.


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Figure 2. Fabrication and characterization of 3D printed porous composite scaffolds. (a) The composites with different HA contents were made by solution mixing. The filaments of the composite around 1.8 cm were obtained from capillary rheometer and fed into the heated nozzle. The material was extruded and printed layer by layer in the X and Y plane. The stage moved down in the Z axis. The resulting geometry of the scaffolds was a serious of rods layered perpendicularly. (b) The µCT images of the filaments proved that the HA were homogeneously distributed within the filament. There was no significant aggregation but an enhancement of radio contrast when increasing the HA content. (c-d) The µCT images of the scaffolds showed the porous structure with a porosity of 75%. The pore size was ~ 320 µm and the diameter of the strut ~ 300 µm.


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Figure 3. The mechanical properties of the 3D printed scaffolds were measured by compression tests at three different conditions: room temperature, physiological temperature and tested after 1 week of incubation. The 3D printed scaffolds exhibited a high compressive modulus compared to the porous scaffolds fabricated by traditional methods such as salt leaching. The 3D printed scaffolds can hold more than 6000 N without yield and fracture.

Table 1. Characterization summary of the composites HA weight% at 550 ºC from TGA a Tg / filament 3D printed Decellularized 3D printed scaffold Td/ b c ºC ºC scaffold after 4 w cell culture PEU0.5 0.6 1.5 308 54 0%HA PEU10.3 10.9 11.9 288 53 10%HA PEU18.9 19.1 21.3 293 49 20%HA PEU27.1 29.9 30.3 290 48 30%HA PEU38.2 38.9 40.1 285 46 40%HA a HA weight% at 550 ºC from TGA in the filament, 3D printed scaffolds and decellularized 3D printed scaffolds after 4 weeks of cell culture. b Onset degradation temperatures of composite scaffolds from TGA under nitrogen. c Glass transition temperatures of composite scaffolds from the 2nd heating cycle of DSC.


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Figure 4. Cell viability and spreading of MC3TC-E1 cells on the composite scaffolds after 1 day and 3 days of cell culture. (a) in vitro cell viability of MC3T3-E1 cells after 1 day of cell culture on the composite scaffolds was studied by live-dead assay (scale bar 500 µm). Live cells were stained green and dead cells were stained red. (c) The cell viability was calculated from a total of 10 images and 3 replicates of each sample. (b) in vitro cell attachment and spreading after 3 days on the composite scaffolds was studied by immunohistochemistry (scale bar 100 µm). Actin filaments was stained red. Vinculin proteins were stained green. Nucleus were stained blue. The cell area and aspect ratios are quantified by Image J (d). 95% of cells survived and no significant difference in cell viability and spreading was observed among different composite samples (p > 0.05).


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Figure 5. Cell proliferation and alkaline phosphatase (ALP) activity of MC3TC-E1 cells on the composite scaffolds after 1 day, 1 week, 2 weeks and 4 weeks of cell culture. (a) MC3TC-E1 cell proliferation within the first 2 weeks was quantified by CyQUANT cell proliferation assay. Cell numbers were normalized to day 1. (b) ALP activity of MC3TC-E1 cells was measured using a standard colorimetric assay. The values were normalized with total amount of DNA to account for the difference in cell numbers. * p < 0.05 and ** p < 0.01


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Figure 6. Osteogenic differentiation of MC3TC-E1 cells within the composite scaffolds after 2 weeks and 4 weeks of cell culture. (a-b) mRNA levels of transcription factor genes of bone sialoprotein (BSP) and osteocalcin (OCN) were measured by real-time RT-qPCR. Data represents fold increase to the levels of control (undifferentiated MC3TC cells, set as 1). Mean values and standard deviations were calculated from four samples. * p < 0.05 and ** p < 0.01 (c) BSP and OCN proteins were secreted by MC3T3-E1 cells as demonstrated by the immunohistochemical staining (scale bar 100 µm). OCN proteins was stained red. BSP proteins were stained green. Nucleus were stained blue.


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Figure 7. Mineralization of MC3TC-E1 cells on the composite scaffolds after 4 weeks of cell culture. (a) Ca2+ deposition in the mineralized ECM on the surface of the filament in the scaffolds were visualized by Alizarin Red S. Staining. The dark red layers on the surface of the filaments in the bright field microscope images indicated the promoted mineralization resulted from higher HA contents in the scaffolds. (b-c) The contents were quantified with ICP-AES by subtracting the Ca2+ concentration of HA in the scaffolds from the Ca2+ concentration of the cell seeded scaffolds after 4 weeks of cell culture and normalized to the total amount of DNA (CyQUANT cell proliferation assay) to account for the difference in cell numbers. * p < 0.05 and ** p < 0.01


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