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An innovative cryopreservation process using a modified core/ shell cell-printing with microfluidic system for cell-laden scaffolds Jae Yoon Lee, YoungWon Koo, and GeunHyung Kim ACS Appl. Mater. Interfaces, Just Accepted Manuscript • DOI: 10.1021/acsami.7b18360 • Publication Date (Web): 23 Feb 2018 Downloaded from http://pubs.acs.org on February 23, 2018

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An innovative cryopreservation process using a modified core/shell cell-printing with microfluidic system for cell-laden scaffolds Jae Yoon Lee1, YoungWon Koo1, GeunHyung Kim*

Department of Biomechatronic Eng., College of Biotechnology and Bioengineering, Sungkyunkwan University (SKKU), Suwon, South Korea 1

These authors contributed equally to this study.

*

Author to whom any correspondence should be addressed.

*

Corresponding author

GeunHyung Kim, Ph.D Associate Professor Department of Biomechatronic Engineering, College of Biotechnology and Bioengineering, Sungkyunkwan University (SKKU), Suwon, South Korea. Email: [email protected], Tel.: +82-31-290-7828.

Keywords: cryopreservation, organ banking, cell-printing, cell-laden scaffold

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Abstract This work investigated the printability and applicability of a core/shell cell-printed scaffold for medium-term (for up to 20 days) cryopreservation and subsequent cultivation with acceptable cellular activities including cell viability. We developed an innovative cell printing process supplemented with a microfluidic channel, a core/shell nozzle, and low temperature working stage to obtain a cell-laden 3D porous collagen scaffold for cryopreservation. The 3D porous biomedical scaffold consisted of core/shell struts with a cell-laden collagen-based bioink/dimethyl sulfoxide mixture in the core-region, and an alginate/poly(ethylene oxide) mixture in the shell-region. Following 2 weeks of cryopreservation, the cells (osteoblast-like-cells or human adipose stem cells) in the scaffold showed good viability (over 90%), steady growth, and similar mineralization to that of a control scaffold fabricated using a conventional cell-printing process without cryopreservation. We believe these results are attributable to the optimized fabrication processes the cells underwent, including safe freezing/thawing processes. Based on these results, this fabrication process has great potential for obtaining core/shell cell-laden collagen scaffolds for cryopreservation, which have various tissue engineering applications.

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Introduction Organ banking, the storage of natural and bioartificial organs with live cells for transplant operations, and tissue engineering have been gaining attention as ideal methodologies to treat damaged organs or tissues. Several researchers feel the need to build a tissue-banking system in preparation for the urgent requests of specific tissues or organs.1 However, the preservation process to successfully store the tissue grafts or organs has several significant issues. One of the issues is related to the long-term preservation and stable storage of ‘living’ biomaterials.2 To overcome this challenge, several investigators have attempted to develop a system for the long-term storage of living tissues or organs. These systems have focused on storage at a very low temperature at which all biological, chemical, and physical processes can be ceased, called cryopreservation.1 Cryopreservation has been used to store, transport, and distribute cells, tissues, organs, and other biological constructs under very low temperatures. Cell cryopreservation needs cell preservation solutions like DMSO (dimethyl sulfoxide) to prevent physicochemical damage to cells during freezing, such as ice crystal formation inside and outside of cells.1,3 By understanding the physicochemical effects of freezing on cells, cell freezing methods for numerous types of cells have been successfully established.1 Likewise, researchers trying to establish tissue banks also desire to find a successful cryopreservation method for living tissues or organs.1,2 However, cryopreservation of large volumes of tissues or organs has some problems that must be overcome. For example, nonuniform temperature gradients of tissues and organs during freezing and thawing processes can damage the biostructures. Recently, Manuchehrabadi et al. developed a nanowarming process, which uses magnetic particles called msIONPs to overcome this problem.4 Through the nanowarming process, they were able to uniformly thaw larger tissues (up to 50 mL; porcine arteries and aortic heart valve leaflet tissues) with over 80% cell viability after warming. However, they had to perform an additional washout step to remove remaining nanoparticles in the tissues. This may be critical, since the cryopreserved tissues have to go through complex post-thaw procedures, which may render the cells unsuitable for transplant operations.5 Another current limitation of the cryopreservation of tissues and organs is a more fundamental problem, shortage of tissue or organ supply. To counter the difficulty of obtaining native tissues or organs in sufficient quantities, cryopreservation has been applied to tissue engineering technologies by many researchers. Tissue engineering is an actively progressing field of research which includes biofabrication or 3D cell/tissue printing techniques.6-8 Tissue engineering has great promise as it can reproducibly fabricate and manufacture three-dimensional (3D) biostructures with living cells for tissue regeneration and repair.2,4 Costa et al. investigated the possibility of applying cryopreservation to tissue engineering by analyzing the effect of cryopreservation on cell-seeded porous and non-porous tissue-engineered constructs.9 They made non-porous disc-shaped scaffolds and porous fiber mesh scaffolds consisting

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of a starch and poly(ε-caprolactone), and seeded both with goat bone marrow stem cells. Both biostructures maintained their functionalities such as surface roughness, porosity, and mechanical properties, while cell survival percentages after cryopreservation were 20% and 54% for discs and fiber mesh, respectively. This low cell viability after cryopreservation has been commonly shown in most studies of the cryopreservation of cell-seeded biostructures.9-11 Liu and McGrath determined the optimal preservation method and vitrification solution for osteoblast cells, which they applied to the cryopreservation of a cell-seeded hydroxyapatite scaffold. For suspended osteoblasts, the cell viability after cryopreservation was over 90%, while the viability of seeded cells on the scaffold was only 43% after the freezing process. They concluded that the reason for the low viability was not the cryopreservation method, but was more likely to be the physicochemical destructive forces caused by the complex shapes of attached cells in the scaffold.10,12 Katsen-Globa et al. found that the shape of cells can influence the effectiveness of the freezing process.11 They analyzed the effects of different pre-cultivation times (0.5, 2, and 24 h) before freezing on the post-cryopreservation viability of human mesenchymal stem cells within alginategelatin scaffold. Shorter cultivation periods yielded higher cell viability and cell activity after cryopreservation, due to the round shape of seeded cells frozen within 2 hours, which can offer cells and their membranes higher stability to osmotic shrinkage during freezing process.11,13-16 The viability of the cells in the scaffold with 2 hours of pre-cultivation showed over 99% (normalized cell viability to control) directly after thawing, much higher than that with 24 hours of pre-culture. However, the initial viability of over 99% had fallen by around 78% at 24 hours after thawing, requiring another 24 hours to recover the original viability. Based on the previous works, we can determine that the cryopreservation process is a critical factor for the successfully preservation of cells or tissues. In particular, the majority of previous studies did not determine the appropriate conditions for various cryopreservation parameters such as temperature gradient and DMSO mixing time. In this study, we developed an innovative fabrication system for a cell-containing core/shell scaffold, which was supplemented with a 3D printing and microfluidic mixing system, for cryopreservation to overcome the shortcomings of the conventional methods. Our process uses a direct cell-printing system supplemented with core/shell nozzle and various temperature controllers, which yield high viability (over 85% after cryopreservation) of the cells loaded in the scaffold by maintaining a round cell shape and including protective vitrification solutions and natural polymers in the bioink. The scaffolds survived over 2 weeks of cryopreservation with high cell viability. Based on these results, we believe that the proposed cryopreservation system using a cell-laden scaffold could be a promising step in the development of manufacturing processes for ready-to-use scaffolds for long-term organ or tissue banking.

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Experimental Materials Osteoblast-like cells (MG63; ATCC, USA) and human adipose-derived stem cells (hASCs; Anterogen Corp., South Korea) were used in the cell-printing processes. Low-viscosity, high-Gcontent LF10/60 alginate (FMC BioPolymer, Norway), poly(ethylene oxide) (PEO; Mv = 900,000, Sigma-Aldrich, USA), and type I collagen from porcine tendon (Bioland, South Korea) were used in the cell-printing processes. Dimethyl sulphoxide (DMSO) (Sigma-Aldrich, USA) was prepared for cryopreservation of the cell-laden scaffold. Calcium chloride (CaCl2) (Sigma–Aldrich, USA) was used as a crosslinking agent to crosslink the alginate.

Preparation of alginate/PEO bioink for conventional low temperature direct printing process (LTDP) To prepare the bioink used in the conventional LTDP scaffold, the alginate, PEO, and DMSO were dissolved in minimum essential medium (MEM). Briefly, the alginate and PEO were dissolved in MEM by stirring the solution the solution at 4°C overnight. The DMSO was added to this solution and stirred for 1 h. The final concentrations were alginate (4 wt%), PEO (4 wt%), and DMSO (10% (v/v)). Before printing, the cells were mixed with the bioink using a three-way stopcock at a density of 1 × 106 cells mL-1 for 10 min.

Preparation of collagen bioink and alginate/PEO solution for core/shell LTDP process In the modified LTDP process supplemented with core (microfluidic channel)/shell nozzle, collagen (0.5 wt%), DMSO (10% (v/v)), and cells (1 × 106 cells mL-1) were mixed through the custom-built microfluidic channel by dispensing the collagen (0.5 wt%)/DMSO (20%) mixture and the collagen (0.5 wt%)/cell (2 × 106 cells mL-1) mixture through the two different inlets of the microchannel. Two types of bioink in the core region were prepared for the different cell types (MG63 cells and hASCs). For MG63 cells, 0.5 wt% collagen in MEM was mixed with DMSO (10% v/v). For hASCs, 0.5 wt% collagen in fetal bovine serum (FBS) was mixed with DMSO. For the alginate/PEO solution in the shell region, the alginate (4 wt%) and PEO (4 wt%) were dissolved in phosphate-buffered saline (PBS) in an autoclave at 120°C for 15 min and cooled to room temperature.

Fabrication of cell-laden scaffold by conventional LTDP process A three-axis robot system (DTR3-2210-T-SG; DASA Robot, South Korea) was used with a dispenser connected to a single nozzle to obtain multilayered cell-laden mash structures. The prepared bioink was printed with an air pressure of 200 kPa on the low-temperature (-15oC) working stage to fabricate stable 3D mesh scaffold by rapidly freezing the hydrogel structure. The temperature of the working stage was set at -15oC according to the previous studies of LTDP.17,18 In the previous studies, hydrogels with/without cells had been printed on the low-temperature (under -15oC) plate and had

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formed a fine 3D mesh structure due to the instant congelation of the hydrogel, showing highly precise printing resolution.17,18,21

Modified LTDP supplemented with microchannel and core/shell nozzle The same model of the three-axis robot system used in the conventional LTDP was supplemented with a microfluidic channel and a core/shell nozzle. The microfluidic channel with 300 µm diameter was used to mix the prepared collagen bioink with DMSO and was connected to the core nozzle (400 µm inner diameter and 700 µm outer diameter). The microfluidic channel was prepared by a company (Illuminaid Inc., Seongnam-si, South Korea). The prepared alginate/PEO solution was then printed through the shell nozzle (1.26 mm inner diameter and 1.65 mm outer diameter). The flow rates of the core and shell regions were set as 4 and 32 mL h-1, respectively. The core/shell strut was printed on the low-temperature (-15oC) working stage as in the conventional LTDP system. Control scaffold was fabricated with the core/shell nozzle and an in-situ aerosol printing process (10 wt% of CaCl2 and 0.93 ± 0.12 mL min-1).19

Cryopreservation and thawing process of cell-laden core/shell scaffold After printing the scaffolds, the frozen samples were first placed in a precooled (-20oC) isopropanol (IPA) container. The container was then stored in a deep freezer (-80oC) overnight for gradual ice formation. The samples were then taken from the container and stored in the deep freezer for up to 14 days before thawing. For assessment of cellular activities of the samples, the cell-laden core/shell scaffolds were crosslinked in CaCl2 solution (2 wt% in cell culture media) for 2 min and washed two times with PBS. During the crosslinking process and PBS washing, the temperature was maintained at 36oC for rapid thawing and removal of DMSO.3

Mechanical analysis of cell-laden core/shell scaffold with/without cryopreservation The mechanical properties of core/shell scaffolds with and without cryopreservation were measured at room temperature in a compression mode using a universal test instrument (Top-tech; Chemilab, Seoul, South Korea). The stress-strain curves of the scaffolds (5 × 5 × 2 mm3) were recorded at a compression rate of 0.05 mm s−1. The scaffolds were measured in wet state. The compressive modulus was obtained as the slope of linear region of stress-strain curves.

Viability assay The cell viability of our samples with/without cryopreservation was examined by counting live/dead cells in fluorescent images at 1, 3, and 7 days of culture. The samples were treated by 50 mM Ethylenediaminetetraacetic acid (EDTA) solution to dissolve shell region alginate for better fluorescent staining. The samples were then treated with 0.15 mM calcein AM and 2 mM ethidium

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homodimer-1 for 30 min in an incubator without light exposure. The stained samples were observed and analyzed by fluorescence microscopy. On the fluorescence images, live and dead cells were presented in green and red, respectively. The numbers of live and dead cells were counted using Image-J software and the ratio of the number of live cells to the total number of cells (including live and dead cells) was calculated as cell viability. The value was normalized by the initial cell viability of the bioink before cell-printing, which was determined using trypan blue (Mediatech, Herndon, VA, USA).

DAPI and phalloidin staining Cell-laden core/shell scaffolds at 7 or 14 days of culture were treated with 50 mM EDTA solution to dissolve the shell region for better fluorescent staining and then stained with diamidino-2phenylindole (DAPI; 1:100, Invitrogen, Carlsbad, CA, USA) staining to analyze the cell nuclei, and Alexa Fluor 568 linked phalloidin (1:100, Invitrogen) to analyze the f-actin cytoskeleton of the cells. The staining was performed for one hour in a darkened room. The stained cell samples were observed and evaluated using a confocal fluorescence microscope. Cell number was counted by counting the cell nuclei stained with DAPI presented in blue and the f-actin area per cell was calculated by dividing total f-actin area presented in red in the fluorescence images with cell nuclei number.

Alkaline phosphatase (ALP) activity ALP activity staining was assessed by determining the released amount of p-nitrophenol (pNP). The cell-laden core/shell scaffolds were washed two times in PBS and transferred into Tris-buffer saline (TBS; 10 mM, pH 7.5) containing 0.1% Triton X-100 for 10 min at 36oC. 100 µL of the aliquot was then added into each well of 96-well culture plates containing 100 µL of pNP solution prepared using an ALP kit (Sigma-Aldrich). ALP activity was measured spectrophotometrically using a microplate reader (Spectra III; SLT Lab Instruments, Salzburg, Austria) with a spectrum at 405 nm.

Osteocalcin (OCN) analysis for osteogenic differentiation potential tests for hASCs The hASC-laden scaffolds with/without cryopreservation were cultured in an osteogenic differentiation medium (DMEM with 10% FBS, 0.1 µM dexamethasone, 50 µM ascorbic acid, and 10 mM b-glycerol phosphate). A MicroVue Osteocalcin EIA Kit (Quidel, San Diego, CA, USA) was used to measure the OCN levels. The samples were gently rinsed three times with 1X wash buffer and 125 µL of anti-osteocalcin was added to each sample followed by incubation at 25°C for 2 h. The samples were washed three times with 1X wash buffer and 150 µL of reconstituted enzyme conjugate was added before incubating for another 1 h at 25°C. A 150 µL aliquot of substrate solution was added, and the wells were incubated for 40 min at 25°C. Without rinsing a 50 µL of 0.5 N NaOH was added to stop the reaction. Then the OD values were obtained by reading at 405 nm using a microplate

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reader.

Statistical analysis All data were presented as the mean ± standard deviation. Statistical analyses were performed using SPSS (SPSS, Chicago, IL, USA) and consisted of single-factor analyses of variance (ANOVAs). In all analyses, P < 0.05 was taken to indicate statistical significance.

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Results and discussion The aim of this study was to develop a fabrication system of cryopreservable cell-laden core/shell scaffolds for medium-term (up to 20 days) storage with reasonable cell viability. This would enable the longer-term storage and possible stockpiling of regenerative medicines with living cells for tissue repair and other medical purpose, as shown in Figure 1. In this way, delay or interruption of the supply of regenerative medicines can be solved, as the prepared scaffolds would be available at any time. However, we had to overcome the limitations of the conventional method, such as ineffective cryopreservation process for the cell-laden scaffold resulting in low cell viability.

Limitation of conventional low temperature direct printing (LTDP) for cell-laden scaffolds DMSO has been widely used in cell freezing procedures for various cell types. However, it can induce critical damage to the cells in some conditions, such as long-term treatment and treatment at specific temperatures.3,20 Figure 2(a, b) shows the effect of DMSO on the cell viability in the cellladen bioink (the mixture of 0.5 wt% of collagen, MG63 cells at a density of 1 × 106 mL-1, and 10% DMSO) for a range of time periods (1 ~ 120 min) at two temperatures (10 and 25oC). As shown in the images (live and dead cells are shown in green and red respectively), with increasing time, the number of dead cells observed clearly increased. Particularly, the cell viability decreased after 10 min. In addition, the cell viability decreased as the temperature increased from 10oC to 25oC, while it did not seem to be affected by the temperature change at shorter time periods than 10 min. This suggests that the DMSO has negligible cytotoxicity in treatments shorter than 10 min for some specific temperature. For this reason, the cell-laden bioink should be in contact with DMSO for a minimum period of time (10 min), to reduce cell death due to the cytotoxicity of the DMSO. Figure 2(c) shows how the cellladen bioink (a mixture of alginate 4 wt%, PEO 4 wt%, MG63 cells at 1 × 106 cells mL-1, and DMSO 10%) prepared at room temperature (25oC) was printed via the conventional low temperature direct printing (LTDP) process, which uses a low working plate temperature (usually around -15oC) to fabricate a 3D mesh structure.17,18,21 The LTDP process was used to obtain relatively the high cellviability of the scaffold because it avoids an abrupt temperature change between the fabricating process and cryopreservation.21 The scaffold was fabricated and cryopreserved at -80oC during one day. Then the scaffold was crosslinked in 2 wt% of CaCl2 solution and washed 2 times in PBS before culturing. Figure 2(d) shows the optical and scanning electron microscope (SEM) images of the crosslinked scaffold. Although the mesh structure of the scaffold was well formed, the cell viability after 1 day of culture after cryopreservation for 1 day was only about 37.1 ± 0.5%, as shown in the live (green)/dead (red) fluorescence images in Figure 2(e). The viability of the cells was extremely low since the DMSO treatment to the cells in the bioink occurred at room temperature (25oC) for over 60 min of processing time (bioink and DMSO mixing followed by printing time to fabricate the cellladen scaffold) and due to thermal shock, which occurred during the processing. This result

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corresponds to the cytotoxicity test of DMSO in Figure 2(a). Based on the results, it was required to develop an instant mixing system for DMSO and cell-containing bioink right before printing at an appropriate low temperature to lower the harmful effect of DMSO and thermal shock.

Development of a microfluidic channel-supplemented printing system As the instant mixing system assisted with temperature controller was required above, we developed the printing system supplemented with a microfluidic channel (channel diameter: 300 µm) to instantly mix the bioink (collagen 0.5%, DMSO 10%, and MG63 cells) thoroughly directly prior to printing (Figure 3(a)). To simply observe the mixing ability of this channel, we ran FITC and rhodamine solution with the ratio 1:1 at a flow rate of 2 mL h-1 for each solution. As shown in Figure 3(b), in the initial contact region of FITC (green) and rhodamine (red) at the starting point of the mixing area (Bregion of Figure 3(b)), the solution was a divided flow-region due to the laminar flow of the solution, but the solutions were well mixed in the end region of the mixing area of the channel. Although this simple analysis cannot provide the exact mixing ability of the channel due to the different viscosities between the experimental solution (0.5 wt% of collagen/cells and DMSO) and FITC/rhodamine solutions, we predict that the experimental solution will be well mixed due to the low viscosity of the experimental solution. The mixing process in the microchannel occurred for 144, 72, 36, and 18 ± 1 second with flow rates of 1, 2, 4, and 8 mL h-1 for the mixed solution, respectively (Figure 3(c)). DMSO should not affect the cell viability at any temperatures in such short times (less than 5 min) according to the results of the cytotoxicity test of DMSO in Figure 2(a). However, since the working plate temperature was around -15oC as required for the low temperature printing,17,18,21 the large difference in temperature between the bioink extruded from the microfluidic channel and the working plate may have affected the viability of the printed cells. Figure 3(d, e) shows the live/dead images and cell viability of the bioink printed using the microfluidic channel cooled to various temperatures (-5, 10, and 25oC) and cultured for 1 day after cryopreservation (1 day) shown in Figure 3(a). During the printing process, the working plate was maintained at -15oC. As shown in the cell viability assay, the cell viability in the bioink printed with the microfluidic channel at 25oC was extremely low (37%), while at a microfluidic channel temperature of -5oC, the cell viability of the bioink increased dramatically to about 87%. The low cell viability of the bioink printed through the microchannel at room temperature may be due to the relatively high temperature difference between the bioink (25oC) and the working plate stage (-15oC), resulting in high freezing rate and relatively high damage to the cells. For this reason, we set the temperature of the microfluidic channel at -5oC and which was controlled by the Peltier CPU cooling system. To efficiently perform the cooling of the bioink, the region (‘A’ in the Figure 3(b)) of the microfluidic channel was used to accelerate the cooling of the DMSO and collagen solution. However, there was a limitation for the low viscosity of the bioink mixture (0.5 wt% collagen + cells)

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due to the mixing effectiveness of the microfluidic channel, since the fluids were required to be low viscosity to flow and mix appropriately within the microchannel. The bioink with 0.5 wt% of collagen had adequate viscosity for good flow in the channel (data not shown). However, the bioink (0.5 wt% of collagen/cells and DMSO) had too low viscosity to fabricate 3D biomedical scaffolds through LTDP in the temperature range of -25oC ~ -5oC, as shown in Figure 3(f). According to several researchers,17,18,22 a low-temperature working plate can be used to fabricate biomaterials including hydrogels from low viscosity (low mechanical strength) materials. However, in our case, the formation of a 3D biomedical scaffold was impossible due to the low viscosity bioink and DMSO, even with the extremely low temperature (-25oC). In addition, the cell viability of the bioink was meaningfully lowered when the temperature was lower than -15oC, due to the overly harsh conditions of the working plate (see Figure 3(g, h)).

Application of core/shell nozzle for the microchannel cell-printing It is well accepted that cell-laden structures should have a mesh structure with appropriate pores, which are required for transferring oxygen or metabolic wastes between the encapsulated cells and the neighboring environment.23,24 To fabricate the mechanically stable 3D porous mesh structure of the collagen-based bioink, we designed a cell-laden collagen scaffold consisting of core/shell struts. Here, we accommodated two bioactive polymers, alginate and poly(ethylene oxide) (PEO), which were used in the shell region to mechanically reinforce the low mechanical properties of the collagen bioink. Alginate is a well-known anionic linear polysaccharide and it has been applied extensively as a tissue engineering material since the alginate can speed up epithelialization as well as easily encapsulate cells because of relatively fast gelation with calcium ions.25 We also used the PEO as a sacrificial component to produce pores on the strut surface, which can induce effectual mass-transfer between the embedded cells and the culture medium, and efficiently enable the release of loaded cells.26 The mixture of alginate (4 wt%)/PEO (4 wt%) was printed through the shell region to enable the formation of the mesh-structured scaffold by enfolding the low viscosity core region containing cellladen collagen/DMSO (see Figure 4(a)). In this experiment, we set the printing temperature of the shell region at a temperature of 5oC because the relatively high viscosity of the shell region can firmly cover the core region, resulting in the stable formation of a core/shell structure. Printing the shell region at room temperature (25oC) caused unstable core/shell structure formation, shown in the optical images of Figure 4(b). The freezing rate of the alginate (4 wt%)/PEO (4 wt%) at 5oC and 25oC is presented in Figure 4(c) and the lower temperature (5oC) showed quicker saturation in the grey scale values, meaning faster freezing, resulting in more stable formation of the core/shell structure (see the cross-sectional view in Figure 4(c)). Figure 4(d) shows the storage modulus of the mixture of alginate (4 wt%)/PEO (4 wt%) in the shell region for temperature sweep. By using the core/shell nozzle and the processing temperature in the shell region (5oC), stable formation of the cell-laden

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mesh scaffold was possible at the working plate temperature of -15oC, compared to higher temperatures (-5oC and -10oC) (Figure 4(e)). In addition, high cell viability in the core region (collagen 0.5 wt%, DMSO 10%, and MG63 cells) was maintained around 85 ~ 90% at these processing temperatures (working plate temperature: -15oC and shell region temperature: 5oC), as shown in Figure 4(e). The overall printability analysis for the printing conditions (working plate temperature, nozzle moving speed, and flow rate of core region) is presented in the Figure 4(f). Based on these results, we set the working plate temperature and moving speed of printing nozzle as -15oC and 10 mm s-1, respectively, to obtain a 3D cell-laden biomedical core/shell scaffold for cryopreservation.

Flow rate analysis for stable formation of core/shell structure In general, the flow rate in the nozzle-based system can cause high wall shear stress (cell-damage) and also it can significantly affect the core/shell structure formation. Because of these aspects, the controllable flow rate of the shell and core region should be considered. Figure 5(a-d) shows the assessment of printability (the ability of core/shell mesh structure formation) and cell viability at a range of core flow rates under a fixed shell flow rate (32 mL h-1). In Figure 5(a, b), the strut size of the printed structure remained constant around 1500 µm at all core flow rates, while the core region (red color with rhodamine) enlarged from around 300 to 800 µm with increasing core flow rate. This phenomenon was because the low viscose cell-laden collagen/DMSO (core solution) was easily diffused in the shell region. In addition, both printability and cell viability (1 day) after cryopreservation (1 day) were stable until the core flow rate reached 4 mL h-1, while the printability became unstable with core flow rate of 8 mL h-1 and the cell viability also decreased, probably due to the relatively higher wall shear stress of the bioink generated by the higher flow rate (Figure 5(c, d)). Based on this, we selected the core flow rate of 4 mL h-1 for the maximum cell-density in the printed struts. In addition, Figure 5(e-h) shows the assessment of the printability and cell viability with varying shell flow rate. The printability was stable with the shell flow rate of 32 mL h-1, while flow rates greater or less than this could cause unstable structure formation (Figure 5(e)). In addition, the cell viability for each shell flow rate (11, 18, 32, and 51 mL h-1) was around 90% and showed no significant difference. As a result, the shell flow rate of 32 mL h-1 showed the most acceptable shapeability for the mesh structure with the former conditions (stage temperature: -15oC, core temperature: -5oC, shell temperature: 5oC, nozzle moving speed: 10 mm s-1, and core flow rate: 4 mL h-1), as shown in Table 1.

Characterization of the cryopreserved core/shell scaffold The final scaffold was fabricated with the optimal conditions selected in the previous sections and

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characterized as shown in Figure 6. In Figure 6(a), the whole structure and the core/shell structure of the scaffold are shown in optical images (live cells in green spots). In addition, the alginate in the shell region was dissolved with 50 mM ethylenediaminetetraacetic acid (EDTA) solution to observe the cells and collagen in the core region (see Figure 6(b)). The fabricated scaffold maintained high cell viability over 90% at 1 day after thawing after 1, 7, 14, and 20 days of cryopreservation as shown in Figure 6(c, d). In addition, the mechanical property of the cryopreserved core/shell scaffold was measured and compared with the scaffold without cryopreservation (see Figure 6(e, f)). These results indicate that the medium-term (for up to 20 days) cryopreservation has no effect on the mechanical properties of the core/shell scaffold and the cell viability of the cells in the core region of the scaffold. Although it needs testing for longer periods of time (e.g. over a month), the cell viability may remain acceptably high even after months of cryopreservation since the high cell viability was maintained after 20 days.

Cell activities of the cryopreserved scaffold To observe the cellular activities of the cryopreserved scaffolds, we cultured the scaffold (after cryopreservation for 14 days) for a further 14 days. In addition, as a control, we used a cell-printed core/shell structure using a bioink (shell: 4 wt% of alginate + 4 wt% of PEO and core: 0.5 wt% collagen + cells (1 × 106 mL-1)) and our previous aerosol printing process, which was not treated with the DMSO, and was not subjected to the cryopreservation process.19 Figure 7(a, b) shows the live/dead and nuclei/f-actin images for the control and scaffolds cryopreserved for 1 and 14 days. The cell viability of the control was slightly higher (94.58 ± 0.88% at culturing day 1 and 97.34 ± 0.95% at day 7) than that of the cryopreserved scaffolds (1-day cryopreserved: 88.87 ± 1.24% at culturing day 1 and 92.86 ± 0.60% at day 7, 7-day cryopreserved: 87.87 ± 1.09% at day 1 and 92.25 ± 0.90% at day 7), while the cell proliferation rate for the culture period (1 and 7 days) was similar (Figure 7(c)). This cell viability of the cryopreserved scaffolds (around 90%) is still much higher compared to previous cryopreserving methods for tissue engineered structures (around 54% for 7-day cryopreserved goat bone MSCs in the study of Costa et al.9; around 43% for MC3T3-E1 cells in the study of Liu et al.10; around 41.2 ~ 78.4% for 1-day cryopreserved hMSCs in the study of KatsenGloba et al.11), which have shown very low cell viabilities caused by the physicochemical destructive forces by the complex shapes of their seeded cells.10,12 The high cell viability of our cryopreservable cell-laden core/shell structures have more possibility to be applied to various tissue engineering studies. Likewise, the f-actin area per cell and the relative ALP activity, indicating cell morphology and bone differentiation of the MG63 cells in the scaffold, respectively, did not show any significant difference between the control and cryopreserved scaffolds (Figure 7(d, e)). All scaffolds showed similar

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increases in the cell proliferation indicators (cell viability and f-actin area per cell) and almost doubled in the bone differentiation indicator (ALP activity; Figure 7(e)) from 7 days to 14 days of the cellculture period. These biological results conclude that the scaffold after the cryopreservation (14 days) showed similar cellular activities and functionality to the control scaffold.

Summarized fabrication processes for the cryopreservable core/shell scaffold As the cellular activities have verified that our fabrication procedure of cryopreserved cell-laden core/shell scaffold was acceptable for MG63 cells, we set the printing processes as shown in Figure 8. In summary, the shell region (alginate 4 wt% + PEO 4 wt%) was cooled to 5oC to wrapping the core region for structural stability. The core region was cooled to -5oC, where the bioink (cell-laden collagen and DMSO) mixed gradually through the microchannel, to reduce cell damages caused by DMSO and the rapid temperature change between the bioink and the working stage (-15oC). The stage temperature was set as -15oC for the stable formation of the core/shell and mesh structures. Core and shell flow rate and nozzle moving speed were confirmed as 4 mL h-1, 32 mL h-1, and 10 mm s-1, respectively. After printing, the samples were stored at -80oC for 14 days and crosslinked with 2 wt% of CaCl2 in cell culture media and washed two times in PBS at 36oC for rapid thawing. The osteoblast-like cells in this process showed complete similar cellular activities (cell viability, proliferation, and ALP activities shown in Figure 7(c-e)) to the control scaffold.

Application of the fabrication and cryopreservation process to adipose-derived stem cells (ASCs) The developed process conditions for the cryopreserved scaffold have also been applied to human adipose-derived stem cells (hASCs) for future application to the regeneration of various tissues, since the hASCs are known to be able to differentiate to different into several kinds of tissue cells, such as bone, cartilage, muscle, or skin cells.27,28 In Figure 9(a,b), the cellular activities of hASCs (cell density: 1 × 106 mL-1) in the same composition of the collagen-based bioink are presented with live/dead assay and nuclei (blue)/f-actin (blue) images. The hASC-containing scaffold cryopreserved for 10 days showed similar cell viability compared to the control scaffold containing hASCs, which was fabricated using an aerosol printing process, was not treated with the DMSO, and was not subjected to the cryopreservation process.19 The cell viability data of the cryopreserved cell-laden core/shell scaffold were presented in Figure 9(c) and the cells have well survived after 10 days of cryopreservation. DAPI and phalloidin staining showed a significant increase of the f-actin area per cell from 7 to 14 days of culture in both hASC samples, as presented in Figure 9(d). When the hASC-laden scaffolds were cultured in an osteogenic medium, the osteogenesis of hASCs have been occurred, as in our previous studies.29 To show the multilineage differentiation potential of hASCs cryopreserved in 7 days in this study, the degree of differentiation was also evaluated with ALP activity (Fig. 9(e)) and OCN (Fig. 9(f)) for the cell-laden scaffolds with/without the

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cryopreservation. As shown in the results, after 7 days of cell culture, the hASCs in the cryopreserved scaffold demonstrated similar ALP and OCN gene expressions compared to those of the control. These results confirmed the possibility of the application of this fabrication of the cryopreserved scaffold containing hASCs for various tissue regenerations.

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Conclusion In this study, an innovative cell printing process supplemented with a microfluidic channel, a core/shell nozzle, and low temperature working stage was proposed to obtain a cell-laden collagen scaffold with core/shell structure for cryopreservation. By controlling various processing factors, such as the temperature of microfluidic channel, working plate temperature, flow rates in core and shell region, and nozzle moving speed, a porous cell-laden scaffold (core: cell-laden collagen/DMSO, shell: alginate/PEO) with high cell viability could be successfully fabricated. After cryopreservation for 14 days, the cells (osteoblast-like cells) loaded into the scaffold had good viability and were able to proliferate and mineralize. When comparing the cell-laden scaffold and a control fabricated using the general cell-printing method without DMSO and cryopreservation, the cellular activities were similar in all measurements. To expand the utility of the process to various tissue engineering applications, we fabricated hASC-laden collagen scaffolds using the same processing conditions. These also showed very similar cellular activities to the control, which was fabricated using the same processing conditions without DMSO or cryopreservation. Based on these results, this fabrication process has enormous potential for obtaining hASC-laden collagen scaffolds for cryopreservation, which have various tissue engineering applications.

Acknowledgements This study was supported by a grant from the National Research Foundation of Korea funded by the Ministry of Education, Science, and Technology (MEST) (Grant No. NRF-2015R1A2A1A15055305).

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References (1) Karlsson, J. O.; Toner, M. Long-term storage of tissues by cryopreservation: critical issues. Biomaterials. 1996, 17, 243-256. (2) Lewis, J. K.; Bischof, J. C.; Braslavsky, I.; Brockbank, K. G.; Fahy, G. M.; Fuller, B. J.; Rabin, Y.; Tocchio, A.; Woods, E. J.; Wowk, B. G.; Acker J. P.; Giwa, S. The Grand Challenges of Organ Banking: Proceedings from the first global summit on complex tissue cryopreservation. Cryobiology. 2016, 72, 169-182. (3) Pegg, D. E. Principles of cryopreservation. Cryopreservation and freeze-drying protocols. 2007, 39-57. (4) Manuchehrabadi, N.; Gao, Z.; Zhang, J.; Ring, H. L.; Shao, Q.; Liu, F.; McDermott, M.; Fok, A.; Rabin, Y.; Brockbank, K. G.; Garwood, M.; Haynes, C. L.; Bischof, J. C. Improved tissue cryopreservation using inductive heating of magnetic nanoparticles. Sci. Transl. Med. 2017, 9, eaah4586. (5) Borel Rinkes, I. H.; Mehmet, T.; Sheehan, S. J.; Tompkins, R. G.; Yarmush, M. L. Long-term functional recovery of hepatocytes after cryopreservation in a three-dimensional culture configuration. Cell Transplant. 1992, 1, 281-292. (6) Groll, J.; Bolan, T.; Blunk, T.; Burdick, J. A.; Cho, D. W.; Dalton, P. D.; Derby, B.; Forgacs, G.; Li, Q.; Mironov, V. A.; Moroni, L.; Nakamura, M.; Shu, W.; Takeuchi, S.; Vozzi, G.; Woodfield, T. B. F.; Xu, T.; Yoo, J. J.; Malda, J. Biofabrication: reappraising the definition of an evolving field. Biofabrication. 2016, 8, 013001. (7) Gao, G.; Lee, J. H.; Jang, J.; Lee, D. H.; Kong, J. S.; Kim, B. S.; Choi, Y.; Jang, W. B.; Hong, Y. J.; Kwon, S. M.; Cho, D. W. Tissue engineered bio-blood-vessels constructed using a tissue-specific bioink and 3D coaxial cell printing technique: a novel therapy for ischemic disease. Adv. Funct. Mater. 2017, 27, 1700798. (8) Pati, F.; Ha, D. H.; Jang, J.; Han, H. H.; Rhie, J. W.; Cho, D. W. Biomimetic 3D tissue printing for soft tissue regeneration. Biomaterials. 2015, 62, 164-175. (9) Costa, P. F.; Dias, A. F.; Reis, R. L.; Gomes, M. E. Cryopreservation of cell/scaffold tissueengineered constructs. Tissue Eng. Part C Methods. 2012, 18, 852-858. (10) Liu, B. L.; McGrath, J. Vitrification solutions for the cryopreservation of tissue-engineered bone. Cell Preserv. Technol. 2004, 2, 133-143. (11) Katsen-Globa, A.; Meiser, I.; Petrenko, Y. A.; Ivanov, R. V.; Lozinsky, V. I.; Zimmermann, H.; Petrenko, A. Y. Towards ready-to-use 3-D scaffolds for regenerative medicine: adhesionbased cryopreservation of human mesenchymal stem cells attached and spread within alginate–gelatin cryogel scaffolds. J. Mater. Sci. Mater. Med. 2014, 25, 857-871. (12) Acker, J. P.; Larese, A.; Yang, H.; Petrenko, A.; McGann, L. E. Intracellular ice formation is affected by cell interactions. Cryobiology. 1999, 38, 363-371.

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(13) Guthrie, H. D.; Liu, J.; Critser, J. K. Osmotic tolerance limits and effects of cryoprotectants on motility of bovine spermatozoa. Biol. Reprod. 2002, 67, 1811-1816. (14) Meryman, H. T. Osmotic stress as a mechanism of freezing injury. Cryobiology. 1971, 8, 489-500. (15) Ragoonanan, V.; Hubel, A.; Aksan, A. Response of the cell membrane–cytoskeleton complex to osmotic and freeze/thaw stresses. Cryobiology. 2010, 61, 335-344. (16) Ragoonanan, V.; Less, R.; Aksan, A. Response of the cell membrane–cytoskeleton complex to osmotic and freeze/thaw stresses. Part 2: The link between the state of the membrane– cytoskeleton complex and the cellular damage. Cryobiology. 2013, 66, 96-104. (17) Lee, H. J.; Kim, G. H. Cryogenically fabricated three-dimensional chitosan scaffolds with pore size-controlled structures for biomedical applications. Carbohydr. Polym. 2011, 85, 817823. (18) Lee, H. J.; Kim, G. H. Cryogenically direct-plotted alginate scaffolds consisting of micro/nano-architecture for bone tissue regeneration. RSC Adv. 2012, 2, 7578-7587. (19) Yeo, M. G.; Lee, J.; Chun, W.; Kim, G. H. An innovative collagen-based cell-printing method for obtaining human adipose stem cell-laden structures consisting of core–sheath structures for tissue engineering. Biomacromolecules. 2016, 17, 1365-1375. (20) Elmoazzen, H. Y.; Poovadan, A.; Law, G. K.; Elliott, J. A.; McGann, L. E.; Jomha, N. M. Dimethyl sulfoxide toxicity kinetics in intact articular cartilage. Cell Tissue Bank. 2007, 8, 125-133. (21) Ahn, S. H.; Lee, H. J.; Lee, E. J.; Kim, G. H. A direct cell printing supplemented with lowtemperature processing method for obtaining highly porous three-dimensional cell-laden scaffolds. J. Mater. Chem. B. 2014, 2, 2773-2782. (22) Xu, W.; Wang, X.; Yan, Y.; Zhang, R. Rapid prototyping of polyurethane for the creation of vascular systems. J. Bioact. Compat. Polym. 2008, 23, 103-114. (23) Hwang, C. M.; Sant, S.; Masaeli, M.; Kachouie, N. N.; Zamanian, B.; Lee, S. H.; Khademhosseini, A. Fabrication of three-dimensional porous cell-laden hydrogel for tissue engineering. Biofabrication. 2010, 2, 035003. (24) Park, J. H.; Chung, B. G.; Lee, W. G.; Kim, J.; Brigham, M. D.; Shim, J.; Lee, S.; Hwang, C. M.; Durmus, N. G.; Demirci U.; Khademhosseini, A. Microporous cell laden hydrogels for engineered tissue constructs. Biotechnol. Bioeng. 2010, 106, 138-148. (25) Alsberg, E.; Anderson, K. W.; Albeiruti, A.; Franceschi, R. T.; Mooney, D. J. Cell-interactive alginate hydrogels for bone tissue engineering. J. Dent. 2001, 80, 2025-2029. (26) Lee, H. J.; Kim, G. H. Enhanced cellular activities of polycaprolactone/alginate-based cellladen hierarchical scaffolds for hard tissue engineering applications. J. Colloid Interface Sci. 2014, 430, 315-325.

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(27) Strem, B. M.; Hicok, K. C.; Zhu, M.; Wulur, I.; Alfonso, Z.; Schreiber, R. E.; Fraser, J. K.; Hedrick, M. H. Multipotential differentiation of adipose tissue-derived stem cells. Keio J. Med. 2005, 54, 132-141. (28) Bunnell, B. A.; Flaat, M.; Gagliardi, C.; Patel, B.; Ripoll, C. Adipose-derived stem cells: isolation, expansion and differentiation. Methods. 2008, 45, 115-120. (29) Yeo, M. G.; Kim, G. H. A cell-printing approach for obtaining hASC-laden scaffolds by using a collagen/polyphenol bioink. Biofabrication. 2017, 9, 025004.

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Tables

Table 1. The fabrication conditions for the cryopreservable mesh scaffold consisting of core/shell struts (core: cell-laden collagen/DMSO and shell (alginate/PEO)). Core region:

-5

Shell region:

5

Working plate:

-15

Core region:

4

Shell region:

32

Temperature (oC)

Flow rate (mL h-1) Nozzle moving speed:

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Figures

Figure 1 Application of cryopreservation/cell-printing process to tissue engineering processes.

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Figure 2 Limitations of the conventional low-temperature direct printing process. (a) Cytotoxicity of DMSO assessed by live (green)/dead (red) assay and (b) cell viability of MG63 cells plotted by DMSO treatment time. (c) Schematics of the conventional low-temperature direct printing process (working plate temperature: -15oC) and (d) optical (left) and magnified SEM (right) images of the scaffold fabricated via the conventional method. (e) Cell viability of the scaffold was examined by live (green)/dead (red) assay.

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Figure 3 Development of the microfluidic channel for cell printing with DMSO. (a) Schematics of microchannel-supplemented low-temperature direct printing process. (b) Structure of the microfluidic channel and mixing test using FITC (green) and rhodamine (red). The channel was comprised of the cooling region (A) (30 × 30 mm2) and mixing region (B) (30 × 30 mm2), and the diameter of the channel was about 300 µm. FITC (substitute for collagen 0.5% + DMSO 20%) was cooled in the cooling region and mixed with rhodamine (substitute for collagen 0.5% + MG63 cells). (c) The mixing time in the mixing area is determined by the flow rate of the mixed bioink (collagen 0.5%/DMSO 10%/MG63 cells). (d, e) Cell viability of MG63 at a range of temperatures (-5 ~ 25oC) of the microchannel by live (green)/dead (red) assay. (f) Poor printability (formation of mesh structure) of the mixed bioink, (g) live/dead, and (h) cell viability analysis at different temperatures (-25 ~ -5oC) of working plate. Asterisks (*) indicate P < 0.05, and NS means statistical non-significance.

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Figure 4 (a) Schematics of the modified low-temperature direct printing supplemented with microchannel and core/shell nozzle. (b, c) Assessment of shell (alginate (4 wt%)/PEO (4 wt%) solution) structure formation at two temperatures (5 and 25oC) and (d) the rheological analysis. The ice formation was quantified by grey scale intensity of the optical images. (e) Optical images of printed mesh scaffolds at different plate temperature (-5, 10, and -15oC) and the whole image of scaffold fabricated at -15oC with stable pore and cell viability. (f) Overall printability analysis for the printing conditions (working plate temperature, nozzle moving speed, and flow rate of core region).

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Figure 5 Flow rate analysis for core and shell nozzle for stable formation of mesh structure. (a) Optical images of core/shell flow from the core/shell nozzle and the fabricated scaffold and (b) core/shell strut size analysis at different core region flow rates (1 ~ 8 mL h-1), shell flow rate was fixed at 32 mL h-1. (c, d) Cell viability was assessed by live (green)/dead (red) assay for the different core flow rates. (e) Optical images of core/shell flow from the nozzle and the fabricated scaffold and (f) core/shell strut size analysis at different shell region flow rates (11 ~ 51 mL h-1), core flow rate was fixed at 4 mL h-1. (g, h) Cell viability examination by live/dead assay for the different shell flow rates. Asterisks (*) indicate P < 0.05, and NS means statistical non-significance.

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Figure 6 (a) Optical and SEM images of final scaffold with core/shell mesh structure. (b) Optical images of final scaffold before/after EDTA treatment and fluorescence images of live cells and collagen fiber in core strut after EDTA treatment. (c, d) Cell viability analyzed by live (green)/dead (red) assay of the scaffolds cryopreserved for 1, 7, 14, and 20 days. (e) Compressive stress-strain curves and (f) calculated moduli of core/shell scaffolds with/without cryopreservation. NS means statistical non-significance.

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Figure 7 Cellular activities of MG63 in the cryopreserved scaffolds. (a) Fluorescence images of live (green) and dead (red) cells and (b) nuclei and actin of the cells in control and cryopreserved samples (1 and 14 days). (c) Cell viability and (d) f-actin area per cell were analyzed from the fluorescence images. (e) Bone differentiation of the samples was assessed with relative ALP activity analysis. NS means statistical non-significance.

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Figure 8 The overall schematics of the final fabrication process of the cryopreservable scaffold. In short, bioink (collagen 0.5 wt%/DMSO 10%/MG63 cells) was mixed through the microchannel (-5oC) and the alginate (4 wt%)/PEO (4 wt%) solution was dispensed through the shell nozzle. The core/shell mesh structure was fabricated on the low-temperature plate (-15oC) and cryopreserved in a deep freezer at -80oC for up to 14 days. The scaffold was crosslinked in 2 wt% CaCl2 and washed 2 times with PBS for rapid thawing before use.

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Figure 9 Application of the final fabrication process of cryopreservable scaffold to human adipose-derived stem cells (hASCs). (a) Live/dead assay and (b) DAPI/phalloidin staining of control and cryopreserved samples (10 days) were performed at 1, 3, 7 days and 7, 14 days of culture, respectively, and presented in fluorescence images. (c) Cell viability and (d) F-actin area per cell was analyzed from the fluorescence images. Osteogenic differentiation potential tests for hASCs with (e) ALP and (f) OCN gene expressions. Asterisks (*) indicate P < 0.05, and NS means statistical non-significance.

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Table of contents (TOC)

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