Biocompatible Macroscale Cell

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A Strategy to Achieve Highly Porous/biocompatible Macroscale Cellblocks, Using a Collagen/genipin-bioink and an Optimal 3D Printing Process Yongbok Kim, Hyeongjin Lee, and GeunHyung Kim ACS Appl. Mater. Interfaces, Just Accepted Manuscript • DOI: 10.1021/acsami.6b11669 • Publication Date (Web): 09 Nov 2016 Downloaded from http://pubs.acs.org on November 12, 2016

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ACS Applied Materials & Interfaces

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A Strategy to Achieve Highly Porous/biocompatible Macroscale Cell-blocks, Using a Collagen/genipin-bioink and an Optimal 3D Printing Process Yong Bok Kim, Hyeongjin Lee, and Geun Hyung Kim* Department of Biomechatronic Engineering, College of Biotechnology and Bioengineering, Sungkyunkwan University (SKKU), Suwon, South Korea *

E-mail: [email protected], Tel.: +82-31-290-7828

Abstract Recently, a three-dimensional (3D) bioprinting process for obtaining a cell-laden structure has been widely applied because of its ability to fabricate biomimetic complex structures embedded with and without cells. To successfully obtain a cell-laden porous block, the cell-delivering vehicle, bioink, is one of the significant factors. Until now, various biocompatible hydrogels (synthetic and natural biopolymers) have been utilized in the cell-printing process, but a bioink satisfying both biocompatibility and print-ability requirements to achieve porous structure with reasonable mechanical strength has not been issued. Here, we propose a printing strategy with optimal condition including a safe cross-linking procedure for obtaining a 3D porous cell-block composed of a biocompatible collagen-bioink and genipin, a cross-linking agent. To obtain the optimal processing conditions, we modified the 3D printing machine and selected an optimal cross-linking condition (~1 mM and 1 h) of genipin solution. To show the feasibility of the process, 3D pore-interconnected cellladen constructs were manufactured using osteoblast-like-cells (MG63) and human adipose stem cells (hASCs). Under these processing conditions, a macroscale 3D collagen-based cell-block of 21 × 21 × 12 mm3 and over 95% cell-viability was obtained. In vitro biological testing of the cell-laden 3D porous structure showed that the embedded cells were sufficiently viable, and their proliferation was significantly higher; the cells also exhibited increased osteogenic activities compared to the conventional alginate-based bioink (control). The results indicated the fabrication process using the collagen-bioink would be an innovative platform to design highly biocompatible and mechanically stable cell-blocks.

Keywords: Collagen, Genipin, Bioink, hASC, Porous cell-block, Cell-printing

1 ACS Paragon Plus Environment


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Introduction The cell-printing process, an additive manufacturing method, is an outstanding technology for fabricating three-dimensional (3D) implantable matrices laden with cells because it provides precise controllability of various geometrical internal/external shapes without the loss of cell-viability, using a computer-aided system. In addition, the method using a cell-laden bioink can enable homogeneous cell distribution in the whole matrix and even be used to deposit multiple cell-types in desired regions of the matrix.1,2 The challenges associated with the cell-printing process include achieving good printability without loss of in situ cell viability after the printing procedure, the fabrication of realistic and safe macroscale 3D structures for various cell types, and creation of a microscale internal porous structure to allow vascularization and transport of nutrients and metabolic wastes.3 In bone tissue regeneration, the microscale pore over 300 µm is known to enhance osteogenesis and prevent osteochondral ossification.4 Furthermore, the 3D cell-laden structure should be biocompatible and allow various cellular activities that occur in the biological environment.3,5 To achieve the most appropriate cell-laden structure, the development of a mechanical bio-printing system and biomimetic bioinks is essential. The most versatile method for obtaining a cell-laden porous structure is direct cell-printing, using a microsized core-sheath nozzle with pneumatic pressure3 and piezo-electric transducer,6 laser,7 tentative aerosol-cross-linking, etc.8 However, although these methods have been applied to overcome several shortcomings in fabricating cell-laden porous structures, many issues remain, including low mechanical properties inducing the collapse of internal micropores, in turn reducing pore-interconnectivity, smaller-than-desired structure (below centimeter scale), and failure to incorporate biocompatible/biomimetic bioinks. 3, 6-8 To date, hydrogels have been the most widely used bioinks because of their physical similarity to extracellular matrix (ECM) and comparably low cytotoxicity. 9 One of the typical bioink hydrogels is alginate, and it has been widely used because of its rapid gelation upon exposure to calcium ions, low toxicity, and controllable rheological and mechanical properties allowed for by the manipulation of the cross-linking conditions and its components.3 However, alginate does not have cell-binding or proliferating components to help cell attachment and proliferation, and it is generally modified with the arginine-glycine-aspartic acid (RGD) peptide sequence.3 Since the modification of alginate with RGD peptide requires the use of a chemical agent [1-ethyl-3-(3-dimethylaminopropyl) carbodiimide (EDC)] which is highly toxic, the resulting product is not completely free from chemical toxicity, although several cleaning processes can be performed. One of the representative ECM proteins, collagen, has been widely recognized as the most appropriate biomedical material for regenerating various tissues. For this reason, many researchers have tried to use it as the ‘ideal’ bioink. However, printed collagen-bioink cannot sustain its printed porous structure because of its low viscosity and


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mechanical properties, and the cross-linking procedure for collagen requires toxic chemical agents (EDC or glutaraldehyde). Hence, despite its outstanding biocompatibility, collagen has not been widely used as a bioink.

Here, we introduce a new cell-printing method, using a collagen bioink to attain a 3D cell-laden structure that is highly porous with fully interconnected pores and macroscale size (over centimeters), without loss of initial cell-viability. To achieve this goal, this study starts with a systematic methodology to describe the particular requirements to stably print a cell-laden structure. The ability to print a multi-layered structure with stable pore

geometry and proper mechanical properties, using collagen-bioink, was evaluated by comparing various printing temperatures affecting the unique gelation phenomenon, which was determined based on the rheological properties of collagen. Further, cross-linking ability of the cell-laden collagen structure, using ‘genipin,’ a material obtained from the fruit of Gardenia jasminoides, a natural crosslinking agent which has been widely used for various biomaterials [gelatin microsphere, chitosan-alginate composite, and poly(ethylene)-glycol hydrogel], 10-12 was optimized. Through the evaluation of printability and biocompatibility, we could select a processing window describing a range for obtaining a promising cell-laden porous structure that is mechanically stable and biologically safe. Osteoblast-like-cells (MG63) and human adipose stem cells (hASC) were used in the work. As based on the results, the proposed printing and cross-linking method will provide a new fabricating criteria and tool for the development of successful collagen-based cell-laden structures.

Experimental Materials Osteoblast-like cells (MG63; ATCC, Manassas, VA, USA) and human adipose-derived stem cells (hASCs; Anterogen Corp., South Korea) were used in this work. As an experimental cell carrier material, type-I collagen (Matrixen-PSP; SKBioland, South Korea) derived from porcine tendon, and high-G-content LF10/60 alginate (FMC BioPolymer, Drammen, Norway), as a control material, were used. To make a neutral collagen solution, 10× enriched DMEM solution was mixed with the collagen solution at a volume ratio of 1:1.13 The neutralized weight portions (3, 5, and 7 wt%) of collagen were used in this cell-printing process, and the mixture of cells (1 × 106 cells mL−1 for MG63s and hASCs) and collagen solutions was utilized as a collagen-bioink. For the cross-linking of the printed cell-laden collagen structures, they were incubated in 0.1, 0.5, 1, 3, and 5 mM genipin solution (Challenge Bioproducts, Taiwan) in medium for 1, 6, 24, and 48 h. In addition, a cell-laden alginate [5 wt% in phosphate-buffered saline (PBS)] having the same cell density was used as a control material. Before



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performing the mixing process of alginate/cells solution, the alginate solution was lightly cross-linked in a 0.5 wt% CaCl2 solution at a ratio of 7:3. Rheological testing Various weight fractions of cell-laden collagen (cell density, 1 × 106 mL−1) were used to measure rheological properties (complex viscosity (n*), storage (G′), and tangent delta). A rotational rheometer (Bohlin Gemini HR Nano; Malvern Instruments, Surrey, UK) installed with cone-and-plate geometry (40 mm diameter, 4° cone angle, 150-µm gap) was used to evaluate the properties. A temperature sweep (ranging between 10oC and 50oC, and ramping rate = 5oC min-1) was conducted with 1% strain and frequency = 1 Hz within the range of the linear viscoelastic region. For frequency sweep tests (0.1 Hz ~ 10 Hz), temperature and strain were set to 37oC and 1%, respectively.

Fabrication of cell-laden mesh structure A dispensing system (DTR2–2210T; Dongbu Robot, Bucheon, South Korea) equipped with a printing nozzle (outer diameter of 310 µm) was used to obtain a multi-layered mesh structure. To generate Gcode commands, the standard 3D printing software was used to control printing process, which was provided by Dongbu Robot Co. In the printing system, the temperature of the barrel/nozzle and working stage was precisely controlled, shown in the schematic image of Fig. 1(a). We applied pneumatic pressure in the range of 110–300 kPa in the micro-size nozzle. The speed of the moving nozzle was fixed at 10 mm s-1. A mesh structure (a 0°/90° strut structure) was obtained using a layerby-layer manner. After printing the structure, genipin solution (0.1, 0.5, 1, 3, 5 mM) was used to cross-link for 1, 6, 24, and 48 h. As a control, an alginate-based cell-laden mesh structure (5 wt% of alginate) was fabricated with our previous fabrication procedure.8 The physical structure, including pore size, cell-laden strut size, geometrical size, and cell density were completely identical to the mesh structure of the collagen-based cell-laden structure.


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Figure 1. Scheme of procedure for fabricating cell-laden collagen-mesh-structure. (a) Collagen bioink mixed with culture medium, (b) 3D printing system supplemented with temperature controllers, and (c) cross-linking with genipin and washing. Characterizations of cell-laden structure An optical microscope system (Model BX FM-32; Olympus, Tokyo, Japan) and a scanning electron microscopy (SEM) system (Sirion, Hillsboro, OR, USA) were used for the observation of surface morphology and mesh structure. Also, to evaluate the elemental calcium and phosphorus distribution in the cultured cell-laden structure, energy-dispersive spectroscopy (EDS) analyses were performed. The pore-size of the cell-laden structure was defined as the distance between the parallel struts. The compressive mechanical properties of the mesh structures were assessed in compressive mode using a universal testing instrument (Top-tech 2000; Chemilab, Seoul, South Korea) at 27 oC. The compressive stress-strain curves of the cell-laden mesh structure were evaluated at a compression rate of 0.2 mm s-1 with diameter = 6 mm and thickness = 1.2 mm. Fourier-transform infrared (FTIR) spectroscopy (Model 6700; Nicolet, West Point, PA, USA) was performed to assess the materials used during the fabrication of the bioinks. Infrared (IR) spectra perform the average of 30 scans in the range of 500 cm-1 and 4000 cm-1 (resolution of 10 cm-1).

In vitro cell culture The cell-laden mesh structures containing MG63s or hASCs were cultured in 6-well culture plates using Dulbecco’s modified Eagle’s medium (DMEM) supplemented with 10% fetal bovine serum (FBS) and 1% penicillin-streptomycin (Thermo Fisher Scientific, USA). The mesh structures were preserved in DMEM with 5% CO2 at 37°C, and the medium was changed every 2 days.



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Osteogenic differentiation of hASC-laden mesh structures The growth medium of DMEM containing 10% FBS and 1% penicillin-streptomycin was used to culture hASCs embedded in mesh structures for 7 days. To induce osteogenesis, the media changed to differentiation medium of DMEM added with 10% FBS, 0.1 µM dexamethasone, 50 µM ascorbic acid, and 10 mM β -glycerol phosphate. Then, the media was exchanged every 3 days, and the samples were cultured for 4 weeks.

Live and dead cell assay and cell proliferation After fabrication, the mesh structures were stained with 0.15 × 10-3 M calcein AM and 2 × 10-3 M ethidium homodimer-1 and incubated for 45 min. The stained samples were then analyzed under a microscope (TE2000-S; Nikon, Tokyo, Japan) installed with an epifluorescence attachment and a SPOT RT digital camera (SPOT Imaging Solutions, Sterling Heights, MI). For the evaluation of cell viability at 1 h, 6 h, 1 day, 2 day, 3 day, and 7 day, images were taken and the quantitative evaluation (cell count of green and red cells) was performed using ImageJ software (NIH, Bethesda, MD). The ratio of the quantity of viable cells to the total quantity of cells (viable and dead cells) was then calculated and normalized to the cell viability, which was obtained from trypan blue (TB) stain (Mediatech, Herndon, VA), before using the bioinks. MTT assay (Cell Proliferation Kit I; Boehringer Mannheim) was used to determine the proliferation of viable cells. Cell-laden mesh structures were put in a 0.5-mg mL-1 MTT solution for 4 h in the incubator at 37°C. The volume of the cultured sample for measurement was 5 × 5 × 1.2 mm3. The amount of MTT solution used during the incubation was 400 µL. Only 100 µL of the supernatant around the mesh structure in the MTT solution was selected as a reading sample and moved to a 96well plate. The structures were not recycled, and different samples were prepared for different time points. The absorbency values at wavelength 570 nm were measured using a microplate reader (EL800; Bio-Tek Instruments). Four samples were tested.

DAPI/Phalloidin analysis The cultured cell-laden samples were stained using diamidino-2-phenylindole (DAPI; 1:100, Invitrogen, Carlsbad, CA, USA) staining to analyze the cell nuclei, and to analyze the actin cytoskeleton of the cells, Alexa Fluor 568 phalloidin (1:100, Invitrogen) was used.

ALP activities and alizarin red S staining ALP was assessed based on the measurement of the released p-nitrophenol (pNP). The cultured samples were washed twice with PBS and put in the incubator for 10 min after adding Tris buffer (10 mM, pH 7.5) containing 0.1% Triton X-100. Then, 100 µL of the aliquot was added into each well of


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96-well culture plates containing 100 µL of pNPP solution prepared using an ALP kit (SigmaAldrich). ALP activity was measured spectrophotometrically at 405 nm, using a microplate reader (Spectra III; SLT Lab Instruments, Salzburg, Austria). Calcium mineralization was evaluated by alizarin red S (ARS) staining of MG63 cells in 24-well culture plates. The cultured sample was rinsed three times with PBS, followed by fixation using 70% (v/v) cold ethanol at 4oC for 1 h, and then air-dried. The sample was stained with 40 mM alizarin red S (pH 4.2) for 1 h and rinsed three times with purified water. Lastly, the sample was destained with 10% cetylpyridinium chloride in 10 mM sodium phosphate buffer (pH 7.0) for 15 min. Staining result was shown using an optical microscope, and the absorption (at 562 nm) was measured using a Spectra III UV microplate reader. All data values are defined as means ± SD (n = 5).

Quantitative real-time polymerase chain reaction (qRT-PCR) For quantitative gene expressions, qRT-PCR was used to measure the osteogenic marker levels of hASCs cultured for 28 days. To isolate total RNA from the hASCs cultured on mesh structures, TRI reagent (Sigma-Aldrich, St. Louis, MO) was used followed by the manufacturer’s protocols. Then, the purity and concentration of isolated RNA were examined by spectrophotometer (FLX800T; Biotek, Winooski, VT, USA) using absorbance at 260 nm. The cDNA synthesis from RNase-free DNasetreated total RNA (1 µg) was conducted with a reverse transcription system. Using Fast Universal PCR Master Mix (Applied Biosystems), qRT-PCR was conducted with StepOnePlus Real-Time PCR System (Applied Biosystems, Foster City, CA). The TaqMan Gene Expression assays (Applied Biosystems) were used for osteogenic marker expression (Table 1). For the assay with fast conditions, total 40 cycles of denature (95 °C) for 1 s and annealing (60 °C) for 20 s were performed. The relative gene expression was normalized by endogenous reference transcript (human glyceraldehyde 3phosphate dehydrogenase (GAPDH)) using the comparative threshold cycle (Ct) method. Then, the hASCs cultured on alginate mesh structure were used to normalize and represent relative gene expression.

Table 1. TaqMan osteogenic gene markers GAPDH

Hs02758991_g1

Bone morphogenetic proteins (BMP-2)

Hs00154192_m1

Runt-related transcription factor 2 (Runx2)

Hs00231692_m1

Collagen type 1 (Col-I)

Hs00164099_m1

Osteocalcin(OCN)

Hs01587814_g1



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Statistical analysis All data values are represented as means ± SD. Data were analyzed by Single-factor analysis of variance (ANOVA), and the statistically significance level was defined as p < 0.05 (*). Results and Discussion Rheological behavior of collagen bioink in culture medium The rheological properties of collagen solution, which is liquefied in acetic acid solution, are highly dependent on the temperature. In particular, storage modulus (G′) or complex viscosity (n*) are rapidly decreased at 30~35oC owing to the breakdown of the collagen helices to a randomized coil structure.14,15 Figure 2(a) shows the graph of the storage modulus and tangent delta (tan δ) of the collagen solution (4 wt% in acetic acid solution) for the temperature range. Below 29oC, the modulus only slightly decreased because of the temperature effect, but above 30oC, the modulus decreased rapidly, reflecting the change in collagen structure. Generally, the temperature of the maximum tan δ is called the denaturation temperature.

Figure 2. Rheological analysis of collagen solution. (a) Storage modulus (G′) and tangent delta (tan-δ) of the 4 wt% collagen in acetic acid solution and (b) the 4 wt% collagen solution in the culture medium, DMEM. Recently, collagen gelation by thermal treatment has been researched for obtaining mechanically stable hydrogel biomaterials. The gelation of collagen, using calcium/phosphate-loaded liposomes or cell-culture medium, can be attained in three steps: (1) fibril formation from several triple helices, (2) fibril growth in a linear direction, and (3) a structure networking between linear fibrils. 16,17 Fig. 2(b) shows the rheological results (G′ and tan δ) for the same weight fraction (4 wt%) of collagen in culture medium (α-MEM). As shown in the graph, the gelation significantly affected the modulus of


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the collagen solution near 29oC. However, the G′ of the collagen has a maximum peak at 30~32oC, and at temperatures above 32oC the modulus was dramatically decreased because of the breakdown of the collagen structure. This phenomenon is due to the competitive process between the gelation and breakdown of the collagen structure. To observe the competitive process for 3, 5 and 7 wt% of cell (MG63 density = 1 × 106 cells mL-1)laden collagen, we measured G’ and tan δ (Fig. 3(a-c)). As expected, the G′ of the cell-laden collagen solutions showed maximum values near 30~40oC. Furthermore, the maximum G′ and temperature of the solutions were increased as the weight fraction of collagen increased (Fig. 3(d)). Based on the rheological assessment results, the processing temperature of the cell-laden collagen to attain a mechanically stable cell-laden structure can be observed. However, as previously reported, the high viscosity or modulus of the cell-laden bioink can cause high wall shear stress in the microsized nozzle utilized in extrusion-based cell-printing.6,18,19

Figure 3. Rheological properties of cell (MG63, 1×106 cells mL-1)-laden collagen solution. Storage modulus (G′) and tangent delta (tan-δ) of various concentrations ((a) 3, (b) 5, and (c) 7 wt%) of collagen bioinks in the culture medium. (d) Maximum storage modulus and gelation temperature for the collagen concentrations. (e) Live (green) and dead (red) images for the in situ printed cell-laden collagen mesh structure at 25oC and measured cell-viabilities. (f) Cell-viability for various weight fraction of collagen. Asterisks (*) indicate P < 0.05 and NS means non-significance. Here, two factors inducing cell-damageable high shear stress in the printing system existed: (1) collagen weight fraction of the bioink and (2) the processing temperature of the bioink in the



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extrusion nozzle. To consider the problems, we should select a reasonable weight fraction of collagen in the bioink and a processing temperature in the nozzle. Figure 3(e) shows the viable cells in the printed microsized struts for the three cell-laden bioinks (3, 5, and 7 wt% of collagen), using the processing temperature (25oC), which was selected to avoid the gelation temperature, of the barrel/nozzle region and the similar mass flow rate (1.52 ~1.62 mg/s). As shown in the live/dead results, 7 wt% collagen caused the cell viability to significantly decrease due to the high wall shear stress, while less than 5 wt% collagen resulted in high cell viability (93%) [Fig. 3(f)]. Based on these result, we utilized bioink with 5 wt% collagen in the subsequent experiments.

Printability of collagen bioink In general, pores and pore interconnectivity are imperative factors determining printability because of their effects on various cellular responses (cell viability, growth, and even differentiation).1,20 For this reason, we used pores consisting of printed cell-laden struts to evaluate the level of printability. A test was conducted with the pneumatic pressures (110 ~ 300 kPa) to fabricate a similar strut size. To evaluate the thermal effect of the barrel/nozzle on the printability and cell viability, we printed the bioink (5 wt% collagen) at four different temperatures (10, 15, 20, and 25oC), which were below the gelation temperature (35oC), in the barrel/nozzle region. This prevents wall shear stress from becoming too high and causing severe damage to the cells laden in the structure (in situ cell viability using the gelation temperature (35oC): 71 ± 2%). Figure 4(a-d) shows the printed structures at each temperature of the barrel/nozzle (B/N-10oC, B/N15oC, B/N-20oC, and B/N-25oC). The temperature of the working plate was 25oC. As displayed in the optical images, when we increased the deposition layers, the pores collapsed, but the processing temperature of 10oC resulted in relatively stable formation of the porous structure until 1.2 mm thickness compared to the others due to the slight increase of the viscosity. In addition, we measured the in situ cell viability for the fabricated structures, using the processing temperatures 10oC and 25oC. As shown in Fig. 4(e,f), the cell viability was above 92% for both processing temperatures. For this reason, we utilized a nozzle temperature in the barrel/nozzle of 10oC.


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Figure 4. Printability of the 5 wt% cell-laden collagen bioink for various barrel/nozzle temperatures ((a) 10, (b) 15, (c) 20, and (d) 25oC). Optical images show various heights printed using the collagen bioink, describing the structure printability. The measured pore size of the printed mesh structure can be used to assess the printability of the structure. (e) Live/dead images for temperatures 10oC and 25oC of the barrel/nozzle. (f) Measured cell viability for each processing temperature of the barrel/nozzle. NS means non-significance. Although the selected processing temperature in the barrel/nozzle could be used to help construct the cell-laden porous structure, the resulting thickness was below 1.2 mm. To obtain a more reasonable thickness of the cell-laden structure, the temperature of the printing stage was manipulated to the gelation temperature of the bioink. Through the previous rheometer data, we could observe the gelation temperature (35oC) of the bioink (5 wt% collagen). To assess the effect of the gelation temperature, we used three different temperatures in the working stage (W-S-25, W-S-30, and W-S35oC). Figure 5(a-c) shows the optical images and pore size distributions for various thicknesses. As expected, the most appropriate thickness having homogenous pore size in the cell-laden structure was about 4 mm using the gelation temperature. However, unfortunately, as the height increased the pores in the structure began to collapse because of the limitation of the heat transfer from the working stage to the thickness direction and the accumulated mass of the struts. Figure 5(d) shows the magnified optical images for the cell-laden structures using the processing temperatures of the working stage, 30oC and 35oC, respectively.



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Moreover, as seen in Fig. 5(e), the live/dead images of cells in structures obtained using the temperatures 25 and 35oC indicated high initial cell-viability (above 92%) (Fig. 5(f)).

Figure 5. Printability of the 5 wt% cell-laden collagen bioink for various working stage temperatures ((a) 25, (b) 30, and (c) 35oC). Optical images and pore size distribution show the stable region of the printable structure. (d) Magnified cell-laden collagen mesh structure for the conditions W-S-30oC and W-S-35oC. (e) Live/dead images for processing temperatures W-S-25oC and W-S-35oC. (f) Measured cell viability for the processing temperature of the working stage. NS means non-significance. Optimal concentration of genipin for cross-linking cell-laden collagen structures We obtained a cell-laden collagen structure with high cell viability and homogenous pores up until a thickness of 4 mm by controlling the printing temperature of the barrel/nozzle and working stage. However, although the printed structure had self-supporting stiffness on the working stage, the mechanical properties were not sufficient for handling the cell-laden structure for cell-culture or implantation into a defected region. To overcome this deficiency, we adjusted the cross-linking process of the cell-laden structure using genipin. Figure 5(a) shows the compressive stress-strain curve of the fabricated collagen structure with cells printed with the gelation temperature before cross-linking with a genipin solution. The compressive modulus of the cell-laden collagen structure was about 17 ± 2 kPa, indicating significantly low mechanical properties, limiting its practical use in tissue regeneration. Generally, collagen can be crosslinked with highly toxic synthetic chemicals (EDC and glutaraldehyde etc.), but the cytotoxic chemicals cannot be used in the cell-laden structure because


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they cause significant cell damage. For this reason, we proposed the use of the natural agent (genipin), extracted from the gardenia fruit, for cross-linking the cell-laden structure because it has a comparable rapid cross-linking time and its cytotoxicity is significantly low compared to synthetic crosslinking agents.21 The collagen crosslinking process using the genipin molecule is described in Fig. 6(b). According to Sung et al., the crosslinking process starts with a nucleophilic substitution of the amine group in the collagen structure on the C3 carbon of genipin to obtain the formation of a nitrogeniridoid.22 Next, the oxygen in the genipin can be replaced with the collagen’s tertiary nitrogen, and the cross-links are the connected cyclical structure of nitrogen-exchanged genipin. Finally, two iridoids successively dimerize via a radical reaction. Figure 6(c) showed the storage modulus (G’) for 5 min, 1 h, 3 h, and 6 h of crosslinking, respectively, and various concentrations (0.5, 1, and 3 mM) of genipin. As shown in the result, as increasing the cross-linking time and genipin concentration, the modulus was significantly increased. Furthermore, the crosslinking ability of various genipin concentrations (0.1, 1, and 5 mM) and crosslinking times (1, 6, 24, and 48 h) on the printed cell-laden structure was observed. Figure 6(d) shows the optical images of cross-linked cell-laden collagen-mesh-structures for various genipin solutions and times. As expected, as the genipin concentration and cross-linking time increased, the qualitative stiffness of the mesh structure continuously improved and also the blue color of the crosslinked structure became denser because of the increased fluorescence intensity (emission at 630 nm) induced by genipin crosslinking.23 To quantitatively observe the stiffness of the cross-linked mesh structure, the compressive modulus was measured (Fig. 6(e)). Unsurprisingly, the stiffness was significantly increased with greater concentration of genipin and longer crosslinking period. FT-IR analysis was performed to detect the chemical reaction during the cross-linking process. Figure 6(f) shows the three different spectra describing pure genipin, collagen, and cross-linked collagen with the genipin solution (1 mM and 1 h). In the genipin spectrum, the three main peaks were at 990, 1080, and 1635 cm-1, which can be attributed to the ring C-H out-of-plane bend, ring C-H in-plane bend, and C=C double bond ring stretch modes of the core of the genipin molecule, respectively. The peak at 1104 cm-1, which was obtained from the chemical reaction of genipin with collagen was attributed to the C-N stretching of the tertiary nitrogen with the neighboring aliphatic carbon existing in lysine or arginine residues of the collagen structure.24 The peak at 1104 cm-1 was identified in the IR spectra of the cross-linked collagen, indicating that the collagen was well crosslinked with genipin solution.



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Figure 6. (a) Compressive stress-strain curve for uncross-linked, cell-laden collagen structure printed with the processing condition W-S-35oC. (b) Cross-linking of collagen using genipin. (c) Frequency sweep result (G’) for various cross-linking times and weight fractions of genipin at 37oC. (d) Optical images of cell-laden collagen lattice structures for several cross-linking conditions using genipin


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(concentration and cross-linking time). (e) Compressive modulus of the cross-linked cell-laden collagen mesh structure for various crosslinking conditions. (f) FTIR spectra for pure genipin, collagen, and collagen bioink crosslinked with 1 mM of genipin solution and a crosslinking time of 1 h. (g) Optical and live/dead images of MG63-laden collagen structures after various cross-liking processes. (h) Cell-viability of the MG63-laden collagen structures after various cross-linking processes. (i) Live/dead images of hASC-laden collagen structures after various cross-liking processes. (j) Cell-viability of the hASC-laden collagen structures after various cross-linking processes. Asterisks (*) indicate P < 0.05 and NS means non-significance. Furthermore, Fig. 6(g,h) shows the live/dead images and measured cell viability of the cell-laden structure crosslinked with genipin. The printed cells in the collagen structure were homogeneously distributed, but we found that the cell-viabilities of the collagen structures exposed to the genipin solutions of 0.1, 0.5, and 1 mM for 48 h were relatively high for osteoblast-like-cells (MG63), while 3 and 5 mM genipin concentrations resulted in significant cell damage. The results indicated that in order to successfully obtain a mechanically stable and sufficiently cell-viable cell-laden collagen structure a limited range of genipin concentrations should be utilized (~1 mM genipin concentration for the MG63 cells). To expand the usage of the genipin solution for cross-linking the cell-laden structure embedding human adipose stem cells (hASCs), we repeated the same fabricating and testing procedure. Notably, as shown in the live/dead images and measured cell-viability in Fig. 6(i,j), the ~1 mM genipin concentration was also safe for the hASC-printed collagen structure. Similar results for fibroblasts were also obtained by Sundararaghavan et al.21 They found that for the L929 fibroblasts in collagenbased biomaterial, genipin concentration of ~1 mM and crosslinking time for 24 h was safe, while 5 and 10 mM genipin solutions resulted in critical cell damage.

Cell responses in cell-laden collagen structure In situ cell viability after printing of cells is an important parameter directly affecting cell growth and even differentiation for several culture periods.3,5 In the previous experiments, we identified the optimum genipin crosslinking conditions and to observe the cell growth in the cell (MG63)-laden collagen structure, a cell-laden alginate mesh structure (pore size = 435±32 µm) obtained using the aerosol tentative crosslinking process8 was used as a control. The control structure (cell-laden alginate) had the same cell density (1 × 106 cells/mL) and similar pore structure (pore size = 452 ± 53 µm and the diameter of cell-laden strut = 437 ± 67 µm) compared to that of the cell-laden collagen scaffold (Fig. 7(a)).



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Figure 7. (a) Optical images of 3D mesh structures printed using MG63-laden alginate (control) and MG63-laden collagen bioink. (b) Live/dead images at 1 and 7 days of the porous mesh structure fabricated using the bioinks containing the osteoblast-like-cells (MG63). (c) Cell viability and (d) cell proliferation determined with MTT assay. (e) Proliferation rate of viable cells. Asterisks (*) indicate P < 0.05. Figure 7(b) displayed the live and dead images for the cell-laden mesh structure after 1 and 7 days of cell culture. The results were applied to quantify the cell viability [Fig. 7(c)], and the collagen bioink structure showed significantly higher cell viabilities at day one and longer culture periods than the control. To detect the metabolic activities of the control and collagen-based cell-laden structures, MTT assay was performed at 1, 3, and 7 days and the proliferation rates were calculated [Figure 7(d,e)]. To measure the rate of cell proliferation, the equation OD(t) = ODoert was used, where OD(t) is the optical density at time (t), ODo is the initial optical density, and r is the proliferation rate. The cellladen collagen structure exhibited higher metabolic activity relative to the control. This is because the printing and cross-linking process is safe and the collagen bioink provides an exceptional cellsupporting material to encourage an effective microcellular environment between cells.25 Figure 8(a) shows DAPI and phalloidin images at 14 and 21 days of cell culture. To quantitatively analyze the morphological behavior of the proliferated cells, the nuclei number and F-actin area were


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measured [Fig. 8(b,c)]. As shown, the printed cells were more proficiently distributed/proliferated on the cell-embedded strut surface in the collagen-based bioink than in the control, indicating that the metabolic activities of the printed cells in the collagen mesh structure were more bioactive for the culturing periods than those in the control.

Figure 8. (a) DAPI/phalloidin images for 14 and 21 days of cell culture on the mesh-structure printed with alginate and collagen bioinks. (b) Cell nuclei density and (c) actin fiber area fraction. (d,e) Optical images of ALP and alizarin red staining describing qualitative mineralization and (f,g) quantitative results of ALP and calcium deposition. SEM and EDS results showing the amounts of ‘Ca’ and ‘P’ after 21 days of culture on the scaffolds printed with (h) alginate and (i) collagen bioinks. Asterisks (*) indicate P < 0.05.

ALP and calcium deposition were used to show the osteogenic differentiation of the cell-laden scaffold, which was cultured for 7 and 14 days in the osteogenic media. As shown in the optical and analyzed data [Fig. 8(d-g)], the levels of ALP activity and calcium deposition in the collagen-based



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mesh structure were significantly higher than those in the control. Moreover, from the EDS results for the cultured structures for 21 days, the quantities of calcium (Ca) and phosphorus (P) elements were measured from the cell-laden alginate and collagen structures (Fig. 8 (h,i)). As a result, the Ca and P peaks were significantly increased for the cell-laden collagen structure than those of the control, and the Ca/P ratio of the cell-laden collagen structure was 1.38, while for the control the ratio was 1.04. The results confirmed that the structure using the collagen-based bioink induced more satisfactory mineralization than the control because of the biocompatible collagen-bioink.

Osteogenic differentiation of hASC-laden collagen structure The optical images of hASCs on alginate and collagen mesh structure were obtained after 28 days of cell-culture (Fig. 9(a,b)). The hASCs were grown in growth media for 7 days, and then the media was changed to differentiation media to promote osteogenesis. For quantification of osteogenic effect, osteogenic specific markers were measured from hASCs cultured on alginate and collagen mesh structure using quantitative real-time PCR (Fig. 9 (c-f)). Generally, osteogenic markers like bone morphogenic protein-2 (BMP-2), Runx2, collagen type-I (Col-I), and osteocalcin (OCN) have been known as differentiation markers.26 BMP-2 are involved in osteogenesis, which increases the transcription of the osteogenic marker (Runx2) and bone structural proteins (Col-I and OCN). 27 In addition, Runx2 is a regulatory factor that assists osteogenesis by propagating hASCs into the osteoblast lineage.28 The significant increase in BMP-2 and Runx2 of collagen mesh structure were measured as 3.9-fold and 20.2-fold compared to that of alginate mesh structure. Col-I is one of the most plentiful components of ECM, and is necessary in influencing cellular behaviors.29 In postproliferative period, OCN is expressed when mineralization occurs, and results in the mineralized bone.29 The gene expression of Col-I and OCN of the collagen mesh structure were 1.39-fold and 1.64-fold compared to the alginate mesh structure. From these results, significantly high osteogenesis from hASCs was observed by characteristics of the collagen mesh structure, the similar environment to ECM, and sufficient cell-binding/proliferating sites.30


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Figure 9. Optical images of (a) hASC-laden alginate mesh structure and (b) hASC-laden collagen structure. Expression levels of osteogenic specific genes, (c) BMP-2, (d) Runx2, (e) Col-I, and (f) OCN by qRT-PCR using hASCs on day 28. Gene expression was normalized to that of GAPDH, and then normalized by the value of hASCs cultured on the alginate mesh structure. An asterisk (*) indicates a significant difference. A strategy for obtaining macroscale collagen-based cell-blocks Furthermore, to demonstrate the feasibility of creating a macroscale 3D-shaped cell-laden collagen structure with a highly porous mesh and high cell viability, we used the selected processing and crosslinking conditions with genipin. To achieve the macroscale cell-laden structure, we used a LEGOblock strategy. Figure 10(a) displays a schematic image demonstrating the fabricated LEGU-block structure. By using several steps shown in Fig. 10(b), one protruding structure like a brick unit of LEGO-blocks was attained. Using the brick unit, we could successfully achieve a macroscale rectangular cell-laden structure (21 × 21 × 12 mm3) (Fig. 10(c)).



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Figure 10. (a) Schematic image of a LEGO-block. (b) Optical images demonstrating the fabricating steps and one protruding structure like the LEGO-block for fabricating a macroscale hASCs-block using the collagen bioink (in situ cell viability : 95±2%). (c) Assembling method and an assembled macroscale rectangular cell-laden structure (21 × 21 × 12 mm3). Conclusion Here, a new cell-printing method and crosslinking process using collagen-based bioink is proposed. By manipulating numerous processing parameters (i.e., printing temperatures of the nozzle and working stage) and the concentration and crosslinking time using the cross-linker (genipin), a highly porous (pore size = over 400 µm)/mechanically stable and biocompatible cell-laden collagen-block of 21 × 21 × 12 mm3 was attained. The in vitro assessment of in situ cell viability, live/dead, and DAPI/phalloidin staining outcomes for several culture periods of the 3D porous cell-block showed that the osteoblast-like-cells and hASCs were sufficiently viable without damage during the printing and crosslinking processes. Significant rates of cell proliferation and gene expression levels of various osteogenic markers were observed in the collagen-based cell-block because the collagen is highly biocompatible in comparison with the alginate-based cell-block. Based on the results, the collagenbased cell-blocks described here have great potential for hard tissue regeneration applications.

ACKNOWLEDGMENTS 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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Biocompatible Macroscale Cell

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