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Collagen Microparticle-mediated 3D Cell Organization: A Facile Route to Bottom-Up Engineering of Thick and Porous Tissues Yuya Yajima, Masumi Yamada, Rie Utoh, and Minoru Seki ACS Biomater. Sci. Eng., Just Accepted Manuscript • DOI: 10.1021/acsbiomaterials.7b00131 • Publication Date (Web): 12 Jul 2017 Downloaded from http://pubs.acs.org on July 16, 2017

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ACS Biomaterials Science & Engineering

Collagen Microparticle-mediated 3D Cell Organization: A Facile Route to Bottom-Up Engineering of Thick and Porous Tissues

Yuya Yajima, Masumi Yamada*, Rie Utoh, and Minoru Seki

Department of Applied Chemistry and Biotechnology, Graduate School of Engineering, Chiba University, 1-33 Yayoi-cho, Inage-ku, Chiba 263-8522, Japan *[email protected]

KEYWORDS tissue engineering, collagen, microparticle, membrane emulsification, cytochrome P450, hepatocyte

ACS Paragon Plus Environment

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ABSTRACT

In closely-packed artificial 3D cellular constructs, cells located near the center of the constructs are not functional because of the limited supply of oxygen and nutrition. Here we describe a simple, unique, and highly versatile approach to organizing cells into thick but porous 3D tissues, using cell-sized collagen microparticles as particulate scaffolds. When cells and particles are mixed and seeded in a non-cell-adhesive planar chamber, they gather to form sheetshaped structures with a thickness of 100–150 µm. In the construct, uniformly distributed particles work as a binder between cells, and modulate the strong intercellular contraction. We confirmed that several factors, including the particle/cell ratio and particle size, critically affect the stability and shrink behaviors of porous tissues prepared using mouse embryonic fibroblasts (NIH-3T3 cells).

Cross-sectional observation, together with cell proliferation and viability

assays, revealed that the cells composing the tissues are functional primarily because interior pores between cells/particles worked as a path for efficient molecular transport. Furthermore, we prepared thick cell tissues of a liver model using human hepatocarcinoma cells (HepG2 cells), and confirmed that liver-specific functions were upregulated when composite tissues were formed using collagen microparticles prepared with several different stabilization protocols by glutaraldehyde, genipin, and methyl acetate). The process presented would be highly useful in enabling one-step production of thick cellular constructs in which porosity and morphology are tunable.


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INTRODUCTION With the recent advancements in mammalian cell culture techniques, construction of functional, three-dimensional (3D) tissues from dispersed cells has attracted a great deal of attention in the fields of regenerative medicine, drug assays using organ models, and development of extracorporeal/ implantable bioartificial devices. Techniques to assemble cells into 3D constructs are generally classified into two types: top-down and bottom-up approaches. In the top-down processes, cells are seeded onto porous scaffolds with pre-defined shapes1. In contrast, bottom-up processes are used to organize individual cells into small building blocks, which are then further assembled as unit structures into larger architectures2,3. In the unit structures used in bottom-up processes, close cell-to-cell interactions are reconstituted if hydrogel matrices or solid scaffolds are not used. Many studies have reported that cell functions and survival in 3D tissues are improved compared to conventional 2D cell cultures for several cell types, including hepatocytes and pancreatic islet cells4-8, because of the physiological similarity with the in vivo cellular environment. Various types of bottom-up approaches have been proposed, as represented by the assembly of multicellular aggregates9-11 and cell sheets12-14.

To create centimeter-sized, thick cellular

constructs using cell sheets, one-cell-thick sheets are first prepared, e.g., using a temperatureresponsive cell culture dish12,15 or electrochemical cell detachment techniques16, and they are then stacked using stamps or cell scoopers17,18.

The prepared constructs are suitable for

transplantation therapies because of their large size and high operability. In addition, several types of state-of-the-art bottom-up strategies have been reported, including the accumulation of cells at the interface of aqueous solutions19, stepwise deposition of extracellular matrix (ECM)-


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coated cells20, and forced cell stacking using magnetic fields21 to produce thick cellular constructs. Based on these processes, multilayered muscle cell sheets22,23, cardiovascular networks24, eart valves25, liver tissue models26,27, and neural tube-incorporating tissues28 have been produced. However, most of these techniques are not capable of independently controlling tissue morphology and cell density at the same time. This problem becomes more serious for relatively large-sized tissues; the cellular environment becomes hypoxic, and cells located near the center of the tissue are no longer viable when tissue thickness becomes greater than ~200 µm29,30. Efforts have been made to induce angiogenesis and capillary formation in multilayered tissues24, but labor-intensive and time-consuming manipulations are necessary.

In addition, the

morphological characteristics of tissues are highly dependent on intercellular contraction forces, thereby making it difficult to control cell density. When cells with relatively high contraction forces are used, the formed constructs dramatically shrink during cultivation, and the tissue shape cannot be maintained. On the other hand, constructs become fragile and unstable when cells with low contraction force are employed. A new technique to easily produce stable thick tissues, while enabling control of both construct morphology and internal cell density/porosity, is therefore highly desired. Recently, techniques to incorporate microengineered ECM components in 3D cell culture platforms have been reported31-35. For example, collagen hydrogel microbeads, ~100-µm in diameter, were produced using microfluidic technology and used as cell carriers to enable cell proliferation and formation of large tissues by bottom-up accumulation31. In addition, collagen or gelatin microbeads produced by bulk emulsification have been mixed with cells and used to form composite 3D tissues32,33. These particles enable the formation of cell-ECM interactions in


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3D tissues, which contribute to the maintenance of cell viability and differentiation ability32. More recently, we developed a process to produce highly condensed, single-micrometer-sized collagen particles using droplet microfluidics or membrane emulsification34. In this process, aqueous droplets incorporating collagen molecules are formed in a continuous phase of a polar organic solvent; the water molecules in droplets are rapidly dissolved into the continuous phase, resulting in the significant condensation of collagen molecules and subsequent particle formation. We were able to produce single-cell-sized type I collagen particles with a final collagen concentration of more than 10%. These particles exhibited non-spherical, disc-shaped morphologies, where collagen fibrils with a diameter of 10-20 nm were reconstituted.

In

addition, we have demonstrated that the functions of primary hepatocytes were upregulated when multicellular spheroids incorporating collagen microparticles were formed, because of the positive effects of collagen, a natural ECM component. We expected that the cell density in thick tissues could be controlled while stabilizing the tissue shape if collagen microparticles with appropriate size and amounts were incorporated. In such tissues, relatively solid particles would work as a binder to control the cell-cell and cell-matrix interactions while stabilizing the tissue morphology. In this study, we present a simple, easy, and highly versatile approach to fabricating thick and porous tissues using collagen microparticles as particulate scaffolds. The concept is shown in Fig. 1. Type I collagen microparticles with sizes similar to the cell size are prepared, and then a mixture of the particles and cells is seeded onto a non-cell-adhesive culture chamber. Because the particles and cells do not adhere to the chamber surface, they gather and form sheet-shaped composite tissues with shapes depending on the initial geometry of the culture platform. Due to the incorporated collagen microparticles, both cell-cell and cell-ECM interactions take place. In


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addition, porous tissues would be generated when an appropriate amount of particles are incorporated. Here we used two types of cells, mouse embryonic fibroblasts (NIH-3T3 cells) and human hepatocarcinoma cells (HepG2 cells), which have relatively strong and weak intercellular contraction forces, respectively10,36. We investigated the effects of particle size and the ratio of particles and cells on the formation and shrinkage behaviors of the formed thick and porous tissues. In addition, we examined whether this 3D cell cultivation process could maintain the cell viability, proliferative ability, and cell-specific functions. Moreover, we employed several types of particles prepared by different stabilization processes, and evaluated the effect of particle type on the expression of liver-specific functions of liver tissue models

Fig. 1. Schematic of the fabrication process of a thick tissue using collagen microparticles as a binder. A suspension of cells and collagen microparticles is introduced onto a non-cell adhesive culture chamber. The accumulated cells and collagen particles gather to form a porous sheetshaped tissue.

MATERIALS AND METHODS Preparation of collagen microparticles A membrane emulsification process, with subsequent droplet dehydration, collagen condensation, and chemical stabilization, was employed to produce collagen microparticles34.


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Briefly, an aqueous solution of ~10 mg/mL type I collagen solution (from rat tail; Corning, NY, USA) was diluted with an aqueous solution of 0.02 M acetic acid to adjust the final collagen concentration to 0.05% (0.5 mg/mL). Methyl acetate (MA; purity >98%; Wako Pure Chemical Industries, Osaka, Japan) was used as the continuous phase: 200 mL of MA was poured into a 400-mL beaker and vigorously stirred at 600 rpm with a magnetic stirrer (SW-RS077D; Nissin Rika, Tokyo, Japan). The diluted collagen solution was continuously pumped into MA through a Shirasu Porous Glass (SPG) membrane (Ф: 8 mm, SPG Direct connector; SPG Technology, Miyazaki, Japan) at a flow rate of 40–100 µL/min using a syringe pump (KDS200; KD Scientific, MA, USA). We used three types of SPG membranes with different pore sizes (3, 5, and 10 µm). After introducing ~6 mL of the diluted collagen solution, the particle suspension in MA was collected in plastic tubes, and the particles were concentrated into a ~10 mL suspension via centrifugation. Collagen molecules were then chemically stabilized at room temperature, either by (i) adding 1 mL of 2.5% glutaraldehyde (GA; Wako) solution in Dulbecco’s phosphate buffered saline (PBS; Takara Bio, Shiga, Japan) and incubating for 15 min; (ii) adding 0.1 mL of 400-mM genipin (GP; Challenge Bioproducts, Taiwan) in 60% ethanol (final genipin concentration of 4 mM) and incubating for 24 h; or (iii) just incubating the particles in MA for 48–60 h. After the stabilization reaction was completed, microparticles were washed with Dulbecco’s modified Eagle’s medium (DMEM; Sigma-Aldrich, MO, USA) with 10% fetal bovine serum (FBS; Thermo Fisher Scientific, MA, USA) at least thrice via centrifugation, to completely remove the solvent and/or the crosslinker.

Particle sizes were analyzed from

captured micrographs using Image J software.

Cell culture


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NIH-3T3 cells and HepG2 cells (both provided by Riken BRC, Ibaraki, Japan) were cultured in DMEM with 10% FBS, 100 U/mL penicillin, and 0.1 mg/mL streptomycin (Sigma-Aldrich) at 37 °C in a CO2 incubator (APM-30D; ASTEC, Fukuoka, Japan) with 5% CO2 under a humidified atmosphere. Confluent cells were harvested from cell culture dishes by trypsinEDTA (Sigma-Aldrich) treatment. The cell suspensions were filtered through a cell strainer (BD Biosciences, CA, USA) with a mesh size of 40 µm (NIH-3T3 cells) or 70 µm (HepG2 cells) before use in experiments to remove large cell aggregates.

Fabrication of cell culture chamber First, chamber structures (Ф = 8 mm, depth = 2 mm) were formed on a poly(methyl methacrylate) (PMMA) plate using a drilling machine (Sakai Mini drill-1; Sakai Machine Tool, Osaka, Japan). Polydimethylsiloxane (PDMS; Silpot 184, Dow Corning Toray, Tokyo, Japan) prepolymer, with the mixing ratio of the base polymer and the curing agent of 10:1, was poured onto the PMMA plate and cured, to obtain a PDMS mold. An aqueous solution of 3% agarose I (Dojindo Laboratories, Kumamoto, Japan) with 0.9% NaCl was poured onto the PDMS mold. After incubating at 4 °C for at least 30 min, the gelled agarose hydrogel plate with the culture chamber was peeled off from the mold. The prepared hydrogel chambers were then immersed in the cell culture medium before use in cell culture experiments.

Preparation of porous tissues Collagen microparticles and cells (NIH-3T3 or HepG2) were mixed, and the suspensions of cells/particles were dropped onto the culture chambers and incubated in the CO2 incubator. The number-based ratio of particles and cells, defined as the “particle/cell ratio”, was varied at 0:1,


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1:1, 4:1, 10:1, or 20:1, while the initial cell density was fixed at 1.0 × 106 cells/cm2 (5.0 × 105 cells for a circular 8-mm chamber) unless otherwise noted. During the cell culture period, photographs were taken using a stereo microscope (SZX16, Olympus, Tokyo, Japan) equipped with a CCD camera (DP72, Olympus). Areas of the generated tissues were quantitatively analyzed using Image J software. NIH-3T3 cells were cultured in the cell culture medium with 10% FBS.

On the other hand, HepG2 cells were cultured in a medium with a low FBS

concentration (1%), to modulate the high proliferation speed and enhance the hepatocyte-specific functions. We mainly used particles stabilized by GA with an average diameter of ~ 5 µm, unless otherwise specified. When porous tissues formed from multiple distinct layers (“distinct-layered tissues”), cells were stained in green or red using PKH67 and PKH26 (Sigma-Aldrich) according to the manufacturer’s protocol. Temperature-responsive cell culture dishes (UpCell, 12 well, 10.55mm diameter; CellSeed, Tokyo, Japan) were used. Cells stained with different color dyes were seeded stepwise, together with the particles, 2 or 3 times at intervals of 12 h. The final cell and particle densities were constant (1.0 × 106 cells/cm2 and 4.0 × 106 particles/cm2).

Evaluation of cell proliferation and viability The ability of cells to proliferate to compose the porous tissues was evaluated using Alamar Blue assay (Bio-Rad, CA, USA), following the manufacturer’s protocol. Briefly, the culture medium was replaced with fresh medium containing 10% Alamar Blue reagent. After 12 h of cultivation, 100 µL of the culture supernatant was collected into a black-bottom 96-well plate, and the fluorescence intensity of the collected sample was measured using a microplate reader


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(MTP-810 Lab; Corona Electronic, Tokyo, Japan), with excitation and emission wavelengths of 550 nm and 610 nm, respectively. To test cell viability, the prepared tissues were dispersed into single cells by trypsin/EDTA treatment. After suspended cells were washed twice, live and dead cells were stained with calcein AM and ethidium homodimer-1 (Live/Dead viability kit; Thermo Fisher Scientific), respectively.

Cells were observed using a phase contrast/fluorescence microscope (IX71;

Olympus). At least 200 cells were counted in randomly selected 5−10 fields for each condition (n = 4).

Histological analysis The formed tissues at Day 2 and 14 were washed thrice with PBS, and then immersed in 10% formalin neutral buffer solution (Wako) at room temperature overnight. After transferring the fixed tissues to 10%, 20%, and 30% sucrose solutions in a stepwise manner (1 h of incubation time each), the tissues were embedded in an O.C.T. compound (Sakura Finetek, Tokyo, Japan) and frozen at -80 °C. Sections with a thickness of 5 µm were then prepared using a cryostat (Leica CM1510S; Leica Biosystems, Wetzlar, Germany), followed by staining using Sirius red (for collagen particles; Direct red 80, Sigma-Aldrich or Picrosirius Red Stain Kit, Polysciences, PA, USA) and Fast green (for cells; Sigma-Aldrich). Stained tissue sections were observed by optical microscopy. Immunohistochemical analysis of Ki-67 at Day 2 and 5 was also performed. The prepared sections (thickness of 7 µm) were treated with 10 mM sodium citrate buffer (pH of 6.0) at 98 °C for 10 min. Then the sections were blocked with 3% horse serum in PBS for 30 min, followed by incubation with anti-Ki-67 rabbit monoclonal antibody (Thermo Fisher) at 4 °C overnight.


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Then the sections were treated with 0.3% hydrogen peroxide in methanol for 30 min to block endogenous peroxidase activity, and subsequently incubated with ImmPRESS Reagent, antirabbit Ig (Vector Laboratories, Burlingame, CA) for 30 min.

Finally, the horseradish

peroxidase-conjugated antibodies were visualized with Metal Enhanced DAB substrate kit (Thermo Fisher) and the sections were counterstained with Mayer’s hematoxylin. The Ki-67positive cells were counted from at least 200 cells in randomly selected 5−10 images (n = 4).

Gene expression analysis Multilayered tissues were prepared using HepG2 cells and collagen particles and cultured for 5 days. Total RNA was extracted using TRIzol Reagent (Thermo Fisher) and a PureLink RNA Mini Kit (Thermo Fisher) according to the manufacturer’s protocol. DNase I (Thermo Fisher) was used to digest genomic DNA in the samples. cDNAs were synthesized from 1 µg of total RNA using a SuperScript VILO cDNA Synthesis Kit (Thermo Fisher). Real-time RT-PCR was performed using Taqman Gene Expression Assays (Thermo Fisher) with a StepOne Plus realtime PCR System (Thermo Fisher). The following genes were examined: glyceraldehyde-3phosphate dehydrogenase (GAPDH), albumin (ALB), ornithine transcarbamylase (OTC), apolipoprotein A1 (APOA1), vascular endothelial growth factor (VEGF), cytochrome P450 1A2 (CYP1A2), 2B6 (CYP2B6), 2C19 (CYP2C19), 2E1 (CYP2E1), and 3A4 (CYP3A4). GAPDH was used as the internal control to quantify the relative gene expressions by the comparative CT method. Results were statistically analyzed by one-way analysis of variance (ANOVA) with Tukey's honest significance difference test (HSD) or Bonferroni post hoc test using IBM SPSS Statistics 22 software (IBM Japan, Tokyo, Japan). A value of p less than 0.05 was considered to be statistically significant.


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RESULTS AND DISCUSSION Preparation of collagen microparticles In an attempt to obtain single cell-sized collagen microparticles, we previously proposed a process that used either microfluidics or membrane emulsification34. In the present study, we employed membrane emulsification, which allows for relatively large-scale particle production. Porous membranes with different pore sizes (3, 5, and 10 µm) were used to control the particle size.

In addition, we examined 3 different protocols to chemically stabilize the collagen

molecules. Particles prepared using 5-µm pore membranes with different stabilization processes, washed with and immersed in the cell culture medium, are shown in Fig. 2 (a). Because the initial collagen concentration in the precursor solution was very low (0.05%), single-cell-sized, highly condensed particles were obtained.

These particles were not stable, and gradually

dissolved when they were introduced into the cell culture medium immediately after particle formation without chemical stabilization (data not shown). In contrast, the shape of particles stabilized by one of the chemical treatment protocols was maintained even in the cell culture medium. It was interesting that the particles incubated only in MA for a long time period (> 48 h) were also stable, although we did not use chemical crosslinkers.

This result could be

explained by the weak fixation effect of general organic solvents on protein molecules37,38. The average sizes and size distributions of the collagen particles were not significantly changed by the different stabilization processes employed (Fig. 2 (b)); the average sizes ± standard deviation (SD) of the particles prepared using 5-µm pore membranes were 5.1 ± 1.6 µm, 5.2 ± 1.1 µm, and 5.4 ± 1.3 µm, when stabilized by GA, GP, and MA, respectively. It is well known that the physicochemical stability and biocompatibility of chemically fixed biomaterials are affected by


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type of crosslinker39-41, and hence, it would be possible to choose a suitable stabilization protocol depending on the application. In addition, we were able to control the size of the particles simply by using membranes with different pore sizes (Fig. 2 (c)). When membranes with pore sizes of 3 and 10 µm were used, the average diameters ± SD of the particles were 3.3 ± 1.0 µm and 10.3 ± 4.1 µm, respectively.


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Fig. 2. (a) Microscopic images of collagen microparticles obtained using different crosslinking protocols (left: glutaraldehyde (GA); center: genipin (GP); and right: methyl acetate (MA)). Scale bar, 10 µm. (b, c) Size distributions of collagen microparticles prepared using (b) different crosslinkers, and (c) porous membranes with different pore sizes (3, 5, and 10 µm). On average, ~200 particles were evaluated for each condition.

Fabrication of multilayered thick tissues Using the prepared collagen microparticles, we fabricated composite thick tissues. First, we used NIH-3T3 cells, which have a relatively strong contraction force10,36. As depicted in Fig. 1, a mixture of the collagen particles (average diameter ~5 µm, crosslinked by GA) and cells (NIH3T3) were mixed and seeded onto non-cell-adhesive agarose chambers. Figure 3 shows the formation behaviors of the tissues with and without the collagen microparticles. The initial cell density was 1.0 × 106 cells/cm2, which is 8~10 times higher than cell confluency (~1 × 105 cells/cm2), and the particle/cell ratio was 10:1 (1.0 × 107 particles/cm2). The average diameter ± SD of the NIH-3T3 cells was 11.7 ± 2.0 µm; hence, the volume ratio of the particles to the cells was ~2:1 under this condition. When cells (and particles) were seeded in the chamber, they were precipitated and accumulated on the bottom within several minutes (Fig. 3 (a, d)). Cells and particles did not adhere on the surface, and initially they were uniformly mixed and evenly dispersed. Cells and particles formed a sheet-shaped structure within 1 day of cultivation (Fig. 3 (b, e)). In the case in which collagen particles were used, the circular shape of the tissues, in accordance with the circular morphology of the chamber, was clearly maintained. The tissues shrank due to cell-cell and cell-particle contraction forces, and the tissue diameter decreased


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from 8 mm to ~5 mm (Fig. 3 (c)). In contrast, a cell aggregate was formed when collagen microparticles were not used, but it rapidly and dramatically shrunk, finally forming a highlycondensed small aggregate (Fig. 3 (f)). These results clearly demonstrated the remarkable effect of the collagen microparticles in maintaining the shape of the 3D cellular constructs. We confirmed that the collagen particles, evenly incorporated into the tissues as particulate scaffolds, mediated the cell-matrix interaction while modulating the strong intercellular contraction.

Fig. 3. Photographs showing the formation behaviors of NIH-3T3 cell tissues (a-c) with or (d-f) without using collagen microparticles (average diameter of ~5 µm).

(a, d) Photographs

immediately after seeding; (b, e) after 1 day of cultivation; and (c, f) after 2 days of cultivation and recovery from the chamber. The particle/cell ratio was 10:1. Scale bar, 2 mm.

We next investigated the effects of the particle/cell ratio on the formation behaviors of the tissues. Figure 4 shows photographs of the planar sheet-shaped tissues prepared using 5-µm particles (Fig. 4 (a-c)), and the time-course change of the relative area of the tissues (Fig. 4 (b)),


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when the particle/cell ratio was changed from 0:1 to 20:1. Under all these conditions, cell aggregates were formed in the chambers within 1 day of cultivation, with the relative areas of the tissues less than 50% of the chamber area. It was clear that tissue size was critically dependent on the particle/cell ratio; larger tissues were formed under high particle/cell ratio conditions. The tissues gradually shrank, and the relative areas reached a plateau after 3–5 days of cultivation. When the particle/cell ratio was equal to or higher than 4:1, the circular shape of the tissue was stably maintained for at least 7 days. In particular, smooth, unwrinkled sheet-shaped tissues were obtained and tissue shrinkage was suppressed when the particle/cell ratio was higher than 10:1. For example, the area of the tissue maintained ~40% of the chamber area when the particle/cell ratio was 20:1, with a corresponding decrease of tissue diameter from ~8 to ~5 mm. On the other hand, the tissues shrank significantly into a small aggregate when the ratios were 0:1 (without employing particles) and 1:1. In conventional methods to create small tissues with densely packed cells (e.g., multicellular spheroids), the cell-cell interaction is the primary driving force for cells to form aggregates42,43. With the present cell culture technique, the relative cellcell interaction was reduced with the increase in the amount of incorporated collagen particles, and this is thought to be the primary reason why tissue shrinkage was suppressed under a high particle/cell ratio. These results clearly suggested that the particle/cell ratio is a crucial factor in the controllability of tissue shape. Under a proper particle/cell ratio condition, we were able to produce tissues with various morphologies depending on the chamber geometry. For example, cylindrical tissues with a diameter of ~1.2 mm and a height of 1.8 mm were obtained (Figure S1 in Supporting Information) using cylindrical cambers (diameter of 2.0 mm, depth of 3.0 mm). It is worth noting that uniformly thick tissues were not obtained, and some pores were formed when the initial cell density was less than 5.0 × 105 cells/cm2, even when collagen particles were


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used. The initial cell concentrations higher than 5.0 × 106 cells/cm2 were not also suitable, because the cellular environment may become hypoxic.

Fig. 4. (a-c) Thick porous tissues of NIH-3T3 cells prepared under different particle/cell ratio conditions at day 2. The particle/cell ratios were: (a) 1:1; (b) 4:1; and (c) 20:1. Scale bar, 2 mm. (d) Time-course change of the relative area of the sheet-shaped tissues formed under different particle/cell ratio conditions. The area of the chamber (50 mm2) was standardized to be 1. Collagen particles with an average diameter of ~5 µm, crosslinked with glutaraldehyde (GA), were used.

Effect of particle size


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Size is one of the most critical particle characteristics, so we examined the effects of particle size on the formation behaviors of the tissues. Collagen particles with different sizes (average size of 3 or 10 µm) were employed, and the particle/cell ratio was changed from 1:1 to 20:1. Figure 5 shows the sheet-shaped tissues formed after 2 days of cultivation. When 3-µm particles were used, the tissues that formed shrank dramatically, even though a relatively large number of particles were incorporated (particle/cell ratio of 20:1).

On the other hand, larger 10-µm

particles suppressed tissue deformation and shrinkage even at low particle/cell ratios (Fig. 5 (c, d)). Under the particle/cell ratio condition of 4:1 with 10-µm particles, the relative tissue area to dish area was as large as ~50% after 2 days of cultivation. Even when the volumetric ratios of the particles were similar, the shrink behaviors of the particles were critically different, as may be seen from the comparison between the 4:1 ratio condition for 5-µm particles (Fig. 4 (b)) and the 20:1 ratio condition for 3-µm particles (Fig. 5 (b)). These data clearly indicated that the optimal particle/cell ratio was significantly different depending on the particle size.

We

concluded that larger particles have a higher anti-shrinkage effect than smaller particles when the volume ratios in the tissues are equal.


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Fig. 5. Photographs of prepared tissues of NIH-3T3 cells incorporating collagen particles with different diameters at different particle/cell ratios at day 2. The average particle sizes were (a, b) 3 µm, and (c, d) 10 µm. The number-based particle/cell ratios were (a) 4:1; (b) 20:1; (c) 1:1; and (d) 4:1. Scale bar, 2 mm.

Evaluation of cell proliferation and viability Next, we investigated the proliferative ability and viability of the cells incorporated into the thick tissues. Figure 6 (a) indicates the proliferative characteristics of NIH-3T3 cells in the tissues evaluated by Alamar Blue assay, under different particle/cell ratio conditions (0:1, 4:1, and 10:1). For the tissues formed without using particles (i.e., a ratio of 0:1), the normalized signal intensity, which indicates cell respiratory activity and is correlated to the absolute cell number, gradually decreased with the progress of cultivation. Considering the gradual decrease in the fluorescence intensity, it was suggested that ~50% of the cells composing the tissue were not viable after 5 days of cultivation. It was most likely that the cell density in the tissues rapidly


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increased at the initial stage of aggregate formation, resulting in an insufficient supply of oxygen/nutrition to the cells. On the other hand, when the particle/cell ratio was 4:1, cell activity was maintained and even increased slightly with the progress of cultivation. When the ratio was further increased to 10:1, the cell activity increased continuously; the apparent cell number showed ~40% increase from day 1 to 5. The results of immunohistochemical staining of Ki-67 positive cells, which indicate proliferating cells, are shown in Figure S2 in Supporting Information. The ratio of Ki-67-positive cells in the tissues prepared using collagen particles was significantly higher (55−60%) than that in the control tissue prepared without using particles (15−20%). These results clearly suggested that cells were proliferating in the inter-particle spaces in formed tissues under the high particle/cell ratio condition. In addition, we examined cell viability after 5 days of cultivation (Fig. 6 (b)). Cell viability under the 10:1 ratio condition was ~92%, indicating that most of the cells were viable. Even in the relatively large, millimeterscale cylindrical tissues prepared using cylindrical chambers, ~85% of cells were viable at Day 5 (Figure S1 in Supporting Information). In contrast, cell viability was lower than 80% when particles were not used. These results clarified that a higher particle/cell ratio was suitable not only for stabilizing the tissue shape, but also for maintaining cell viability and proliferative ability.

Compared to the conventional multicellular spheroids, where normal cells do not

substantially proliferate because of contact inhibition, the present approach is advantageous in that it allows cell proliferation through adhesion on the collagen particles. For cells that do not substantially proliferate in vitro (e.g., primary hepatocytes), the low particle/cell ratio might possibly result in insufficient formation of cell-cell interactions, and hence, it would be better to optimize the cell/particle densities depending on the cell type.


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Fig. 6. Evaluation of proliferative property and viability of NIH-3T3 cells in the thick tissues. (a) Time course change of the cell numbers represented by the respiratory activity measured by Alamar Blue assay.

The metabolites of Alamar Blue were quantified by measuring the

fluorescence intensity. (b) Evaluation of the cell viability by live/dead assay after 5 days of cultivation. Each data point is the mean ± SD from 4 Individual samples. *p < 0.05, **p < 0.01 compared to the control group (the particle/cell ratio of 0:1).

Observation of the tissue cross section We next observed the cross-sectional morphology and interior structure of the prepared tissues after 2 days of cultivation. Sections of the recovered tissues were prepared and then stained with


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Sirius red (particles) and Fast green (cells). In this experiment, the particle/cell ratio was either 4:1 or 10:1. The density of NIH-3T3 cells was changed for the 4:1 particle/cell ratio condition (either 1.0 × 106 or 2.0 × 106 cells/cm2). The stained sections are shown in Fig. 7. Cells and particles were almost uniformly distributed throughout the entire region of the cross section. The tissues were not closely packed with cells/particles, forming a porous interior structure (Fig. 7 (ac)).

The cells and particles formed unique, 3D networks in the void spaces between

cells/particles. In addition, the cells were located in the inter-particle spaces, and they adhered and extended on the particle surface, indicating that they functioned as inter-particle bridges to maintain the tissue shape. The surface of the constructs was covered with a one-cell-thick layer, probably because cell proliferation was promoted on the relatively flat surfaces with a greater supply of oxygen/nutrition. Meanwhile, in the control sample without particles, the tissues (small aggregates) were closely packed with cells because of the high intercellular contraction force of NIH-3T3 cells (Fig. 7 (d)). The thicknesses of the sections were in the range of 100-150 µm when particles were used. Under a particle/cell ratio condition of 4:1, most of the cells contacted adjacent cells, covering the particles sufficiently (Fig. 6 (a, b)), and hence, it was naturally deduced that cell proliferation was not promoted because of contact inhibition. On the other hand, when the particle/cell ratio was 10:1, cell density in the tissue was further decreased and larger void spaces were formed (Fig. 6 (c)). Particles were not completely covered with cells, which was likely one of the main reasons why cells continued to grow under this condition. It is worth noting that the thickness of the tissue showed only ~20% increase even though the densities of the particles/cells were doubled (Fig. 6 (a, b)). Considering that the tissue diameters were 4.3 and 5.1 mm for the (a) 1.0 × 106 and (b) 2.0 × 106 cells/cm2 conditions, respectively, the entire volume of the tissues in the latter case was increased ~1.7-fold; hence, cell density was


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increased ~1.3-fold compared to the former condition. These results indicated that not only the appearance but also the tissue porosity, i.e., the density of packed cells, can be controlled simply by adjusting the particle/cell ratio and the density of seeded cells. To produce thick 3D tissues, several bottom-up methods have been previously developed, including the stacking of single-cell-layer cell sheets17,44-46 and accumulation of ECM-coated cells20. The cell-sheet stacking procedure, is advantageous in the sense that intact ECM layers are preserved and aligned between the sheets44,45.

However, there was a limitation in the

thickness of the formed tissues; the thickness was only 20−50 µm even when 5 homotypic cell sheets were stacked46. Compared to these methods, the process presented herein is advantageous in that it allows for the fabrication of thick tissues by a highly simplified process/operation: namely, just seeding cells together with collagen microparticles into non-cell-adhesive chambers. Another benefit is the stability of the shape of produced tissues, even under a floating condition, which facilitates the recovery of the tissues from the cultivation chamber without employing complicated devices or equipment.

Moreover, there were internal void spaces for cell

proliferation, which might have worked as a conduit for an effective supply of nutrition and oxygen even to the cells located at the center of the tissues.


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Fig. 7. Histological observation of the cross sections of the NIH-3T3 cell tissues prepared under different particle/cell ratio conditions; 5-µm particles, stabilized by GA, were used.

The

particle/cell ratios were (a, b) 4:1, and (c) 10:1, and the initial cell densities were (a, c) 1.0 × 106 cells/cm2, and (b) 2.0 × 106 cells/cm2. (d) Control tissue sample prepared without collagen microparticles (the initial cell density was 1.0 × 106 cells/cm2). Cells and particles were stained with green and red, respectively. Scale bar, 50 µm.

Preparation of distinct-layered tissues When cell-adhesive dishes were used to form the particle-incorporating tissues, cells adhered on the surface, thereby making it possible to overlay additional cells/particle layers on the upper


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surface of the formed tissue. Stepwise seeding of cells/particles would therefore enable the formation of hierarchically arranged tissues composed of multiple distinct layers.

To

demonstrate this concept and recover the formed distinct-layered tissues, we employed temperature-responsive cell culture dishes (Ф = 15.5 mm). Collagen microparticles (5 µm) were seeded together with NIH-3T3 cells, with a particle/cell ratio of 4:1. After 12 h of cultivation at 37 °C, cells adhered on the temperature-responsive dish, and by repeating the cell/particle seeding process, distinct-layered tissues were formed (Fig. 8). After 2 days of cultivation, we were able to successfully recover the tissues from the temperature-responsive dish simply by lowering the temperature to 20 °C. Immediately after the tissue detached from the dish, it shrank slightly, and the tissue diameter decreased to ~90% of the dish diameter. Each cell layer was clearly distinguishable from other layers, and they were not mixed at this time point. Compared to the above-mentioned experiments using non-cell-adhesive chambers, tissue shrinkage in the horizontal plane was suppressed, resulting in the formation of thinner tissues (70–80 µm), although the particle/cell densities were the same (Fig. 4 (b), thickness of ~120 µm). This may be explained by the weak cell-cell contraction in the horizontal direction, because some of the cells adhered on the dish surface during cultivation. In this experiment we used only one cell type, but it would be possible to produce tissues composed of hierarchically arranged, multiple layers of heterotypic cells, which would be applicable to the formation of complicated tissue models such as skin equivalents and various types of epithelial tissues.


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Fig. 8. Bright-field and fluorescence micrographs showing the cross sections of the distinctlayered tissues of NIH-3T3 cells; (a, b) double-, and (c, d) triple-layered structures. Tissues were recovered from temperature-responsive dishes after 2 days of cultivation. Scale bar, 100 µm.

Evaluation of liver-specific functions of HepG2 cell tissues In an attempt to demonstrate the applicability of the present approach to the formation of functional tissues, we fabricated a porous 3D liver-tissue model. We employed HepG2, a human hepatoma cell line that is widely used to simulate liver-specific functionalities in vitro4,47-49. It has been well established that the functions of HepG2 cells are upregulated when close cell-cell interactions are formed in a 3D platform4,47. However, cells located near the center of densely packed tissues, e.g., spheroids, are not functional and viable when the tissue size is greater than ~200 µm, mostly because the local cellular environment becomes hypoxic30. Here, we prepared tissues of HepG2 cells using collagen microparticles with an average size of ~5 µm. The particle/cell ratio was fixed at 4:1. We examined three types of particles prepared using different


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stabilization processes (GA, GP, and MA), because it is known that the type of crosslinker used for the solid scaffolds affects cellular adhesion, proliferation, and functions50,51. In addition, promotion of cell proliferation is not necessarily ideal when the prepared tissues are applied to in vitro drug evaluation.

Hence, to enhance functions of HepG2 cells and to suppress cell

proliferation, HepG2 cells were cultured in a medium with a low serum concentration (1%). Figure 9 shows the tissues fabricated of HepG2 cells at Day 5, with and without collagen particles. Unlike in the experiment using NIH-3T3 cells, contiguous tissues of uniform-thickness were formed even when particles were not used; the tissue did not show significant shrinkage, but maintained the circular morphology (Fig. 9 (a)). This result can be attributed to the lower intercellular contraction force between HepG2 cells compared to NIH-3T3 cells10,36. When tissues were prepared using differently stabilized particles, the circular shape of the tissues was also maintained (Fig. 9 (b-d)), but the tissue diameter (5.5-5.7 mm) was smaller than that of the control group (~6.0 mm), for which no particles were used. In addition, small cracks were sometimes observed. The cross sections of the tissues are shown in Fig. 9 (e, f). Both in the tissues with and without collagen particles, porous interior structures were observed. In the case without particles (Fig. 9 (e)), cells were uniformly distributed in the cross section, and the cell shape was mostly spherical. In contrast, in the tissues with collagen particles (Fig. 9 (f)), cells were not uniformly distributed but rather were localized around the particles, and the inner structure became sparser. These results suggest that the cell-particle contraction was greater than the cell-cell contraction for HepG2 cells, and the collagen microparticles enhanced the aggregate formation of the cells. Hence, both cell-cell and cell-ECM interactions were realized in the composite tissues prepared using the cells and particles. These structural characteristics are similar to those found in the in vivo liver tissues, where hepatocytes contact each other at the


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apical side, whereas the basolateral sides are covered with a layer of ECM, the so-called space of Disse.

Fig. 9. (a-d) Photographs of the HepG2 cell tissues after 5 days of cultivation (a) without or (bd) with collagen microparticles. Three types of particles stabilized by different processes were employed: (b) GA, (c) GP, and (d) MA. The particle/cell ratio was 4:1 for (b-d), and the average particle size was ~5 µm. Scale bar, 2 mm. (e, f) Cross-sections of the HepG2 cell tissues (e) without and (f) with collagen particles (stabilized by GA) at day 5. Cells and particles were stained with green and red, respectively. Scale, 50 µm.


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We next evaluated the liver-specific functions of HepG2 cells in the tissues. Figure 10 shows the gene expressions of HepG2 cells after 5 days of cultivation, quantified by real-time RT-PCR. It was clarified that the gene expressions of liver-specific functions (albumin (ALB), apolipoprotein A1 (APOA1), and ornithine transcarbamylase (OTC)) were mostly upregulated when collagen particles were used (Fig. 10 (a-c)), regardless of the types of the particles. There are two factors that could possibly explain the upregulation of these genes: (i) collagen particles enhanced the cell-ECM interactions; and (ii) the void spaces functioned as a path for an effective supply of oxygen and nutrition. The latter is supported by the expression of VEGF (Fig. 10 (d)); this result implies that the cellular environment for the control sample without collagen particles is hypoxic, and the cells produced VEGF to induce angiogenesis, because of the uniform but relatively packed tissue configuration. We also analyzed 5 types of cytochrome P450 family genes, which are involved in drug metabolism. Interestingly, the expression of these genes was affected by the particle type, and some of these genes showed statistically significant differences compared to the control group, especially for the particles stabilized by GP.

Recently,

biomaterials crosslinked with GP were reported to have a positive effect on HepG2 cells41. It is known that the crosslinking process using GP is more biocompatible than those using other crosslinkers, which possibly contributed to the preservation of integrin-mediated cell-supportive functions of the native collagen molecules. Based on these results, we further analyzed HepG2 cell functions after cultivation for a longer period (2 weeks) in the tissues prepared using GP-crosslinked particles. The results are shown in Figure S3 in Supporting Information. We confirmed some of the gene expressions at Day 5 and Day 14 were comparable (e.g., APOA1 and CYP1A2), but others (e.g., ALB and OTC) showed significant increase from Day 5 to Day 14, indicating that tissue maturation is progressing during


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this culture period. Furthermore, we observed the particle stability in the tissues at 2 weeks. The results are shown in Figure S4 in Supporting Information. Unlike the tissues of NIH-3T3 cells tissues, where particles were stably maintained regardless of the crosslinking protocols, collagen particles in the HepG2 cell tissues, stabilized by GP and MA, were significantly degraded. This was probably caused by the matrix metalloproteinases produced by the cancerous HepG2 cells52, indicating that collagen remodeling is taken place by the proteinases. With these experiments, we confirmed the positive effects of the collagen particles on the function of hepatic cells, and showed that the local chemical characteristics surrounding the cells is tunable using different types of particles. It would therefore be possible to apply the obtained tissues to assessment of the metabolism behavior of drug candidates using the porous liver tissue model. In this study, we tested only one cell type. Evaluation of the effect of the particles on other cell types (e.g., primary hepatocytes and other cells surrounded by ECM components in vivo) might be necessary in order to apply the presented approach to various biomedical researches.


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Fig. 10. Gene expression assays of hepatocyte-specific genes of HepG2 tissues at day 5, as evaluated by quantitative RT-PCR. Three different types of collagen particles, stabilized either by GA, GP, or MA were used. (a) ALB, albumin; (b) APOA1, apolipoprotein A1; (c) OTC, ornithine transcarbamylase; (d) VEGF, vascular endothelial growth factor; (e) CYP1A2–CYP3A4, cytochrome p450 1A2–3A4. GAPDH was used as the internal control for relative quantitation. The relative expressions were normalized to that of the control tissue formed without using


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collagen particles. Each data item shows the mean ± SD of 4 samples. *p < 0.05, **p < 0.01 compared to the control group without using particles.

CONCLUSIONS We proposed a simple, easy, versatile, and highly unique bottom-up 3D tissue engineering approach using collagen microparticles as a functional binder for cells. The collagen particles could be uniformly introduced into the formed tissue, which worked as particulate scaffolds to maintain the entire shape of the multilayered tissues. Because the intercellular contraction was modulated by the presence of the particles, cells were not very closely packed in the tissue. The interior pores may have enhanced the supply of oxygen and nutrition, which contributed to the maintenance of cell function and viability, and allowed cells to proliferate in the porous constructs. We investigated several parameters, including the amount of incorporated particles, particle size, and stabilization process on the formation behavior of thick tissues, and clearly showed that these factors critically affected the tissue morphology. Moreover, the functions of liver tissue models were upregulated, especially when particles prepared with specific stabilization protocols were employed. The process presented herein is highly useful because it enables one-step production of thick but porous 3D tissues without necessitating complicated devices or operations. The prepared tissues would be compatible as a unit structure in organson-a-chip devices, as the porous configuration is potentially suitable for medium perfusion and integration of multiple units. Future studies will include the production of tissues of various cell types for drug evaluation and regenerative therapy, and the creation of complicated tissue models for cell biological studies by the stepwise deposition of heterotypic cells.


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ASSOCIATED CONTENT Supporting Information The Supporting Information is available free of charge on the ACS Publications website. Photographs of the cylindrical tissues, immunohistological analysis of Ki-67-positive cells, gene expressions of HepG2 cell tissues at Days 5 and 14, and the tissue cross sections at Day 14 (PDF).

AUTHOR INFORMATION Corresponding Author Dr. Masumi Yamada, Department of Applied Chemistry and Biotechnology, Graduate School of Engineering, Chiba University, 1-33 Yayoi-cho, Inage-ku, Chiba 263-8522, Japan. Tel&Fax: +81-43-290-3398 E-mail: [email protected] ORCID: 0000-0003-2596-3527

Author Contributions All authors contributed to manuscript preparation. All authors have given their approval to the final version of this manuscript.


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Notes The authors declare no competing financial interest.

ACKNOWLEDGMENTS This study was supported in part by Grants-in-Aid for Scientific Research (16H04571, 26350530, and 23106007) and for JSPS Fellows (15J06315 and 16J40041) from the Ministry of Education, Culture, Sports, Science, and Technology of Japan.


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Collagen Microparticle-mediated 3D Cell Organization: A Facile Route to BottomUp Engineering of Thick and Porous Tissues

Yuya Yajima, Masumi Yamada, Rie Utoh, and Minoru Seki


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Collagen Microparticle-Mediated 3D Cell ... - ACS Publications

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