Hydrogels with an embossed surface: An all-in-one platform for mass production and culture of human adipose-derived stem cell spheroids

Biomaterials 188 (2019) 198–212

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Hydrogels with an embossed surface: An all-in-one platform for mass production and culture of human adipose-derived stem cell spheroids

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Se-jeong Kima,b, Jaesung Parka,b, Hayeon Byuna,b, Young-Woo Parka,b, Luke G. Majorc, Dong Yun Leea,b,d, Yu Suk Choic, Heungsoo Shina,b,d,∗ a

Department of Bioengineering, Hanyang University, 222 Wangsimri-ro, Seongdong-gu, Seoul 04763, Republic of Korea BK21 Plus Future Biopharmaceutical Human Resources Training and Research Team, Hanyang University, 222 Wangsimri-ro, Seongdong-gu, Seoul 04763, Republic of Korea c School of Human Science, University of Western Australia, Perth, WA 6009, Australia d Institute of Nano Science & Technology (INST), Hanyang University, 222 Wangsimri-ro, Seongdong-gu, Seoul 04763, Republic of Korea b

ARTICLE INFO

ABSTRACT

Keywords: 3D cell culture Stem cell spheroid 3D printing Surface structure Hydrogel

Stem cell spheroids have been studied extensively in organoid culture and therapeutic transplantation. Herein, hydrogels with an embossed surface (HES) were developed as an all-in-one platform that can enable the rapid formation and culture of a large quantity of size-controllable stem cell spheroids. The embossed structure on the hydrogel was adjustable according to the grit designation of the sandpaper. Human adipose-derived stem cells (hADSCs) were rapidly assembled into spheroids on the hydrogel, with their size distribution precisely controlled from 95 ± 6 μm to 181 ± 15 μm depending on surface roughness. The hADSC spheroids prepared from the HES demonstrated expression of stemness markers and differentiation capacity. In addition, HES-based spheroids showed significantly greater VEGF secretion than spheroids grown on a commercially available low-attachment culture plate. Exploiting those advantages, the HES-based spheroids were used for 3D bioprinting, and the spheroids within the 3D-printed construct showed improved retention and VEGF secretion compared to the same 3D structure containing single cell suspension. Collectively, HES would offer a useful platform for mass fabrication and culture of stem cell spheroids with controlled sizes for a variety of biomedical applications.

1. Introduction Recently, technologies to culture one or more cell types in 3D conditions have attracted a great deal of attention in many research fields, including tumor cell biology, drug development, and regenerative medicine [1–3]. The characteristics of 3D-cultured cells differ from those of cells cultured in 2D in several aspects, such as cellular heterogeneity, morphology, and mass transport [4]. In particular, numerous reports have demonstrated that stem cells cultured in vitro in the form of 3D spheroids have improved viability, self-renewal capacity, and differentiation potential compared to 2D-cultured cells [5–8]. Human adipose-derived stem cells (hADSCs) have been widely used for 3D spheroid culture due to their high extraction yield through a relatively non-invasive process, rapid proliferation, and ability to differentiate into various lineages [9]. Numerous studies have reported that hADSCs cultured as spheroids showed enhanced stemness, proangiogenic factor secretion, and differentiation under defined supplements compared with 2D-cultured hADSCs [9–14]. In addition, in vivo

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transplantation of hADSC spheroids lead to prolonged survival, improved engraftment, and up-regulation of anti-inflammatory and proangiogenic responses [15–19]. Thus, the development of 3D culture platforms that can provide a physiologically in vivo-like environment would be important. Stem cell spheroids can be prepared using various culture techniques, including liquid overlay, hanging drops, and spinner flasks [20,21]. Those methods allow anchorage-dependent stem cells to be exposed to a confined protein adsorption surface or prevented from attaching to the bottom surface of a culture dish via a balanced buoyant force or continuous agitation, respectively [22]. As a result, cell–cell aggregation to promote spontaneous formation of spheroids is maximized, and cell adhesion to the surface through interactions with adsorbed extracellular matrix (ECM) proteins is minimized. Although those techniques generate a desirable environment for stem cell spheroid formation, challenges remain that restrict their wide use in many applications. Hanging drops are labor-intensive, generally requiring a large number of cells and time to induce self-aggregation. In

Corresponding author. Department of Bioengineering, Hanyang University, 222 Wangsimri-ro, Seongdong-gu, Seoul 04763, Republic of Korea. E-mail address: [email protected] (H. Shin).

https://doi.org/10.1016/j.biomaterials.2018.10.025 Received 15 October 2018; Accepted 19 October 2018 Available online 22 October 2018 0142-9612/ © 2018 Elsevier Ltd. All rights reserved.

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addition, the spheroids must be transferred to a second plate for prolonged culture or various assays [23]. Mechanical forces during spinner flask culture can impede a uniform size distribution of the spheroids, affecting the biochemical and physiological functions of the stem cells and potentially interfering with the viability or pro-angiogenic effects of the spheroids through hypoxia, oxygen concentration gradient, or nutrient diffusion [24]. 3D printing technologies have also been introduced for fabrication of spheroids, though most of those techniques are useful for preparing a 3D structure that encapsulates spheroids within crosslinked bioink hydrogels [25,26]. In those conditions, harvesting material-free spheroids is difficult without causing cellular damage. To overcome the drawbacks of existing methods, alternative spheroid culture techniques based on biomaterials with controlled chemical properties and physical structures have been developed [2,3]. For example, chitosan films, either alone or conjugated with hyaluronan, have been used to make a low-adhesive surface for stem cell spheroid culture, with the average diameter of the spheroids controlled by initial seeding density [14,27]. Other studies combined the low adhesive properties of materials with physical patterns on their surfaces to control the diameter of the fabricated spheroids. Fibroblast spheroids have been fabricated on wrinkle patterns induced on poly (2-hydroxyethyla methacrylate) hydrogels, yielding spheroids with a narrow size distribution [28]. In addition, honeycomb structures on poly (dimethylsiloxane) and inverted colloidal crystal scaffolds of poly (acrylamide) hydrogels have been used to fabricate size-controlled mouse embryonic stem cell spheroids and hepatocyte spheroids for functional liver tissue formation, respectively [29,30]. Those studies each provided a culture substrate for producing spheroids in a relatively large quantity with a certain level of size controllability, but an ideal biomaterial system that could accelerate spheroid formation with mass production, controlled size, and long-term culture potential is still highly desired. Given that, we here developed a 3D culture platform based on hydrogels with an embossed surface, enabling mass production of hADSC spheroids and their long-term culture with size control. We hypothesized that the cell-repellant surface of the hydrogel would lead to cohesion of the hADSCs, and that the irregular embossed roughness would rapidly lead to cell aggregation (spheroidization) within the valleys while inhibiting cell adhesion during long-term culture. We also hypothesized that those combined physical and chemical features would improve spheroidization efficiency compared with previous techniques. The main objectives of this study were 1) development of hydrogels with an embossed surface at various micro-scales, 2) investigation of the effect of the embossed surface on the formation of hADSC spheroids of various sizes, 3) investigation of the effect of the spheroids on stemness and multi-lineage differentiation properties of hADSCs, and 4) demonstration of the potentiation of pro-angiogenic properties using size-controlled spheroids in combination with a 3D bioprinting set-up.

CA, USA); EZ-CYTOX, a WST-based cell viability/cytotoxicity assay kit, from DoGenBio (Seoul, Korea); anti-fibronectin antibody from BD Biosciences (Franklin Parks, NJ, USA); anti-Nanog, anti-Oct4, antiSOX2, anti-Collagen IV, and anti-Laminin antibodies from Abcam® (Cambridge, MA, USA); a human VEGF standard ABTS ELISA development kit from PeproTech (Rocky Hill, NJ, USA); SYBR® Premix Ex Taq™ from Takara Bio (Mountain View, CA, USA); Maxime RT premix from Intron Biotechnology (Seongnam, Korea); Hypoxia Probe LOX-1 from Organogenix (Kawasaki, Japan); mounting medium containing 4′,6-diamidino-2-phenylindole (DAPI) from Vectashield® (Burlingame, CA, USA); mounting medium for histology from Richard-Allan Scientific (Kalamazoo, MI, USA); and an RNeasy mini kit from Qiagen (Hilden, Germany). All primers for qPCR were from Cosmogenetech (Seoul, Korea). We purchased NOVAMATRIX® PRONOVA UP LVG sodium alginate from FMC (Industriveien 33 N-1337, Sandvika, Norway); a 27-gauge blunt needle from JUYOUNG Eng (Seoul, Korea); and sodium hydroxide from Duksan Science (Seoul, Korea). 3. Methods 3.1. Preparation of hydrogels with an embossed surface For fabrication of the PDMS master mold, sandpapers with various grit designations (P60, P100, P220, P320, P1000) were placed in a square dish and covered with Silgard silicon elastomer solution (Fig. 1). The elastomer solution was cured for 2 h at 60 °C and then gently detached from the underlying sandpapers. The PDMS master mold was sterilized with 70% EtOH under UV for 2 h before use. For hydrogel formation, the resulting PDMS master mold was placed within a glass mold with a 1-mm gap Teflon spacer. Tetronic®-tyramine (Tet-TA) was synthesized as previously described [31] and dissolved in PBS in a separate tube containing H2O2 or HRP. The prepared solutions were mixed in a 1:1 ratio and directly injected between the glass mold and the PDMS master. The final concentration of each compound was TetTA 12% (w/v), H2O2 0.1% (v/v), and HRP 0.0025 mg/ml. After 10 min of gelation, the hydrogels with an embossed surface (HES) with topographical cues (60, 100, 220, 320, 1000) were separated from the PDMS master. The detached hydrogels were cut into disk shapes and sterilized under UV for 2 h. Before use, the hydrogels were kept in an incubator (37 °C). To analyze the surface embossed structure, the hydrogels were incubated in 1 μg/ml DAPI solution (in PBS) for 10 min. The DAPI-absorbed hydrogels allowed fluorescent 3D reconstruction of the HES. The hydrogels were then placed face up on microscope slides and imaged at 10× magnification using a 405 nm wavelength laser on a confocal laser microscope (C2 confocal microscope, Nikon, Tokyo, Japan). Z-stack images (5 μm steps) of the hydrogels were captured, and volumetric reconstructions were created using NIS-Elements software. The images were color-coded based on z-height to visualize the topographical profile of the HES. Additionally, the surface topography of the sandpaper and the PDMS master were analyzed by scanning electron microscope (NOVA NANO SEM) installed at the Hanyang Center for Research Facilities (Seoul).

2. Materials We purchased Tetronic® 1307 (M.W. 18000) from BASF (Ludwigshafen, Germany); p-nitrophenyl chloroformate, tyramine, hydrogen peroxide (H2O2), horseradish peroxidase (HRP), cytochalasin D, and ultra-low attachment plates from Sigma Aldrich (St. Louis, MO, USA); and AggreWell™400 from STEMCELL Technologies (Vancouver, BC, Canada). A Sylgard® 184 silicone elastomer kit from Dow Corning (W. Salzburg Rd., MI, USA) was used for polydimethylsiloxane (PDMS) master mold fabrication. Phosphate-buffered saline (PBS) and trypsin/ EDTA were purchased from Wisent (St. Bruno, QC, Canada). We bought sandpaper from Starcke (Melle, Germany); rhodamine-phalloidin, Vybrant DiO cell-labeling solution, a LIVE/DEAD® viability/cytotoxicity kit, trypan blue solution, STEMPRO® hADSCs, and MesenPRO RS™ medium from Life Technologies Corp. (Grand Island, NY, USA); low glucose Dulbecco's modified Eagle's medium from Gibco BRL (Carlsbad,

3.2. Formation and characterization of spheroids on HES hADSCs (passage number 3 to 5) were cultured in growth medium (MesenPRO RS™) under standard conditions (37 °C, 5% CO2, 95% humidity). The hADSCs were detached from their tissue culture plates using PBS and trypsin/EDTA. For spheroid formation, HES were placed in 48-well plates. Then, hADSCs were seeded on the HES (seeding density = 1 × 105 cells/cm2) to monitor spheroid formation. Spheroids on HES 60, 100, 220, 320, and 1000 were designated as S-60, S-100, S220, S-320, and S-1000, respectively (Table 1). Phase contrast images were obtained at each time-point using an inverted microscope (CKX41, Olympus, Tokyo, Japan). The diameter and total number of spheroids were measured using NIS-Element AR software (Nikon, Tokyo, Japan). 199

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Fig. 1. Schematic diagram of fabrication of hydrogels with an embossed microstructure (HES) by replica molding. PDMS molds were prepared on commercially available sandpapers with various grit designations. Then, the embossed microstructures were imprinted on the surface of crosslinked hydrogels. Human adiposederived stem cells were spontaneously assembled and formed as spheroids on the embossed microstructure of the hydrogel.

To check spheroid shape, hADSCs were seeded on HES 60, 100, 220, 320, and 1000 (seeding density = 1 × 105 cells/cm2) and cultured for 24 h. The resulting spheroids were fixed using 4% paraformaldehyde. Factin was stained by treatment with rhodamine-phalloidin for 1 h. Subsequently, samples were washed twice with PBS and mounted with mounting medium containing DAPI for nuclear staining. The 3D structure of the spheroids was observed using a confocal laser scanning microscope. For histological analysis, spheroids on HES 60, 100, 220, 320, and 1000 (seeding density = 1 × 105 cells/cm2) were fixed 1 and 3 days after seeding. Fixed spheroids were encapsulated in 3% agarose gel and paraffinized. Paraffinized samples were sliced into 5 μm thick slices. For H&E staining, specimens were deparaffinized and hydrated using xylene and graded EtOH (70–100%) and then treated with hematoxylin and eosin solution. After treatment, specimens were dehydrated and mounted with mounting medium. Stained samples were observed by optical microscopy. For fluorescence staining for Nanog, Oct4, SOX2, fibronectin, collagen IV, and laminin, spheroids were cultured on HES 320 (seeding density = 1 × 105 cells/cm2) for 3 days. The samples were then sliced into 5 μm slices after undergoing the fixation and paraffinization steps described above. After hydration of the specimens, an antigen retrieval step was carried out by treatment with sodium citrate buffer for 10 min at 95 °C (10 mM sodium citrate, 0.05% Tween 20, pH 6.0 in distilled water). Specimens were washed three times with PBS and incubated with primary antibodies for each target for 12 h at 4 °C. Secondary antibody and FITC-conjugated tertiary antibody were added for 1 h at 37 °C. Samples were washed with PBS three times between staining steps. DAPI-containing mounting medium was used to observe the nuclei. Stained samples were observed using a fluorescence microscope. To evaluate the transcriptional factor expression level in spheroids, hADSCs were seeded on HES 320 (seeding density = 1 × 105 cells/cm2) and cultured. As a 2D control, cells were seeded in a 6-well plate (seeding density = 2 × 104 cells/well). After three days, RNA in the spheroids was extracted using an RNeasy mini kit. The total amount of RNA was determined by measuring the absorbance at 260 nm using a NanoDrop™ 2000 spectrophotometer (Thermo Fisher Scientific, Waltham, MA, USA). One microgram of RNA was mixed with Maxime RT premix and put into a thermal cycler (MyCycler™, Bio-Rad, Hercules, CA, USA) for cDNA synthesis according to the manufacturer's instructions. SYBR® Premix Ex Taq™ was used to perform RT-PCR. Relative gene expression levels were measured using a StepOnePlus™

Table 1 Name of each experimental group. Grit designation

Hydrogel name

Spheroid name

Seeding density [cells/ cm2]

None (Control) 1000 320 220 100 60

Flat 1000 320 220 100 60

S-Flat S-1000 S-320 S-220 S-100 S-60

1 × 105

To measure the diameter of the spheroids, we measured the size of all spheroids in images randomly obtained from each experimental group. To investigate the effect of seeding density on spheroid diameter, hADSCs were seeded on HES 320 at 0.5, 1, and 2 × 105 cells/cm2. The size distribution of the spheroids was checked using NIS-Element AR software. To confirm the mechanism of spheroid formation on HES, we used cytochalasin D and Y-27632. Cytochalasin D or Y-27632 was premixed with hADSCs to make a suspension. Then, hADSCs were seeded onto HES 320 (seeding density = 1 × 105 cells/cm2). The final concentration of cytochalasin D and Y-27632 was 1 μM and 100 μM, respectively. The initial spheroid formation process was monitored for 24 h. Phase contrast images were obtained using an inverted microscope (CKX41, Olympus). After 24 h, the viability of each group was checked using a trypan blue exclusion assay. The circularity of the spheroids was measured by image analysis. The projected area and circumcircle area were measured using ImageJ software. Subsequently, the area ratio was calculated. The viability and proliferation of spheroids on HES 60, 100, 220, 320, and 1000 were investigated on days 3 and 7. For the viability assay, a LIVE/DEAD® viability/cytotoxicity kit was used. After the reaction, fluorescence signals were observed using a confocal laser scanning microscope (TCS SP5, Leica, Wetzlar, Germany). Moreover, spheroids were dissociated with trypsin for 5 min, and a trypan blue exclusion assay was carried out to quantify viability. To measure proliferation, the WST-1 assay was used. After treatment with WST-1 solution, absorbance at 440 nm was measured. To confirm the proliferation of spheroids, the absorbance value on day 7 was normalized by the absorbance value on day 3. 200

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density = 5 × 105 cells/ml). The mixture including the spheroids was put into a syringe with a 27-gauge blunt needle and then loaded into the 3D bioprinter (nozzle size: 0.3 mm, printing speed: 12 mm/s, displacement: 60%). After 3D bioprinting, the fabricated 3D gel structure was further incubated with 100 mM calcium chloride solution for 3 min for additional crosslinking. After further crosslinking, the 3D gel structure was washed with culture medium for the following experiments. The printed cells were visualized using a live/dead assay kit. To assess viability, the printed structures were ruptured by gentle pipetting. Spheroids from the printed structures were treated with trypsin for 5 min and completely dissociated. A trypan blue exclusion assay was carried out to quantify spheroid viability after printing. The cell retention rate was confirmed using a DNA assay. Printed structures were cultured in 24-well plates for 7 days. On days 1, 4, and 7, printed structures were collected into microtubes and treated with RIPA lysis buffer. To check the cells exiting the printed structure, the remaining culture medium in the 24-well plate was removed, and RIPA lysis buffer was added to each well. Cell lysates from the printed structures and the 24-well plates were assayed for DNA. The cell retention rate within the printed structures was calculated as the ratio of DNA contents of the cells remaining in the printed structure to DNA contents of the exiting cells. Furthermore, VEGF secretion of the printed structures was monitored for 7 days. The printed structures were cultured in 24-well plates, and culture medium was collected on days 1, 2, 3, 5, and 7 for VEGF ELISA. In addition, stemness (Nanog, SOX2, Oct4), cell junction (ecadherin), and anti-apoptotic (Bcl2L1) gene expression of the spheroids was confirmed using qRT-PCR. The printed structures were cultured for 3 days before RNA extraction.

Real-Time PCR system (Thermo Fisher Scientific, Waltham, MA, USA). Specific gene expression (fibronectin, laminin, e-cadherin, connexin 43, Nanog, Oct4, SOX2) was normalized by GAPDH. 3.3. Differentiation of spheroids on embossed hydrogels To confirm the differentiation capacities of the spheroids, hADSCs were seeded on HES 320 (seeding density = 1 × 105 cells/cm2) and cultured for 14 days under each differentiation condition (adipogenesis, chondrogenesis, and osteogenesis). Culture medium was exchanged to differentiation medium 24 h after seeding. Two-dimensionally cultured cells were used as a control group. To exchange the culture medium, the spheroids on HES were harvested by mild pipetting and pelleted after centrifugation. After aspirating the supernatant, fresh medium was added. The medium-exchanged spheroid suspension was then re-seeded on HES for prolonged culture. Differentiation medium was exchanged every two days for successful differentiation in both the spheroid and control groups. For staining, samples were fixed with 4% paraformaldehyde. Then, spheroids were entrapped within a 3% agarose hydrogel and frozen with sectioning compound. Spheroids were sliced into 10 μm thick specimens and stained with various dyes (Oil red O for adipogenesis, Alcian blue for chondrogenesis, and Alizarin red for osteogenesis). For adipogenic, chondrogenic, and osteogenic marker gene expression analyses, qRT-PCR was carried out at the final time-point. 3.4. Comparison to commercial 3D culture plates Ultra-low attachment plates (LAPs) were used to compare the characteristics of spheroids formed on commercially available 3D culture plates without surface structure to the spheroids formed on the HES. hADSCs were seeded (1 × 105 cells/cm2) on LAPs, flat hydrogels (Flat), and HES 220. The spheroids on each substrate were monitored for 3 days. Phase contrast images were obtained using an inverted microscope (CKX41, Olympus), and the spheroid diameter was measured by image analysis using NIS-Element AR software (Nikon). To sense hypoxia inside the spheroids, LOX-1 was added to the medium 12 h before imaging. Phase contrast and fluorescence images were obtained using a fluorescence microscope (TE 2000, Nikon). Fluorescence images were obtained with the same exposure time (2 s) to normalize fluorescence intensity. To evaluate gene expression levels, all experimental groups were sacrificed for RNA extraction, and qRT-PCR was carried out after 3 days. To quantify VEGF released from the spheroids, the spheroids were harvested every 24 h. Spheroid suspensions were centrifuged to make pellets. Supernatants were used for VEGF ELISA, and the pellets underwent a lysis step for a DNA assay to normalize VEGF secretion with DNA content.

3.6. Statistical analysis All quantitative results are reported as means ± standard deviations. Differences between groups and treatments were evaluated using Student's t-test and analysis of variance, with post hoc comparison by means of Tukey's HSD test (p < 0.05) (n = 4). 4. Results and discussion Fig. 1 illustrates the fabrication of HES using synthetic hydrogels for spontaneous spheroid formation of stem cells. The PDMS master molds were prepared using commercially available sandpapers, and the TetTA hydrogels were chemically crosslinked on the PDMS master, resulting in hydrogels with an embossed surface structure. The embossed structure on the surface of the HES assisted spontaneous hADSC spheroid formation by maximizing cell–cell cohesion and inhibiting cell adhesion on the surface of the hydrogels. Furthermore, the spheroid diameter was controlled by the grit designation of the sandpaper and the seeding density of the hADSCs.

3.5. 3D bioprinting of spheroids

4.1. Surface characterization of HES

To confirm the injectability of the spheroids, hADSCs were seeded (1 × 105 cells/cm2) on HES 60, 100, and 320 and cultured. For the control group, a flat hydrogel was used. After 24 h, the formed spheroids were harvested using a 1 ml syringe with the needle removed and replaced with a 27-gauge needle. Subsequently, the spheroid suspension was injected into a 24-well plate at a speed of 200 μl/s. After injection, the spheroid morphology was observed using an inverted microscope (CKX41, Olympus). Live/dead and trypan blue exclusion assays were carried out after injection to confirm spheroid viability. The fluorescence signals of cells were observed using a fluorescence microscope (TE 2000, Nikon). The 3D structure of the gel was designed using Auto-CAD2016 (Autodesk) and fabricated with an INVIVO® 3D bioprinter (ROKIT, Korea). Briefly, a mixture of 2% (w/v) alginate in 0.9% sodium chloride solution and 0.36% (w/v) collagen was prepared (mixing ratio; alginate : collagen = 1 : 2), and then hADSC spheroids from HES 320 (seeding density = 1 × 105 cells/cm2) were added (final cell

SEM images of sandpaper with diverse grit designations and their corresponding PDMS master molds are shown in Fig. 2(a) and (b), respectively; the surface morphology of the sandpapers showed a coarser structure as the grit designation decreased. Consequently, the topographical features of each sandpaper were engraved on the surface of the PDMS master mold depending on the grit designation. The embossed patterns were then successfully reproduced on the surface of the hydrogels, as shown in Fig. 2(c). More severe light scattering on the HES was observed as the grit designation decreased, and reconstructed 3D images confirmed the presence of protrusions with diverse ranges of depth, duplicating the surface features of each sandpaper. The protrusion size increased as the grit designation decreased. Generally, the average roughness (Ra) values of hydrogels without surface topographical cues are within 1 μm [32–34]. With the master molds tested in this study, we were able to fabricate HES with surface elevation 201

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Fig. 2. SEM images of (a) sandpapers and their corresponding (b) PDMS molds. The value in each image represents corresponding grit designation. (c) Photographs of hydrogels and their surface morphology characterized by confocal laser scanning microscope. (d) Representative line profile results of hydrogels with various embossed structures. (e) Average roughness of hydrogels with embossed structures. “#” indicates statistical significance.

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Fig. 3. (a) Phase contrast microscope images of spheroids formed on the hydrogels with various microstructures depending on grit designation. (b) Diameters of spheroids assembled and cultured on the hydrogels. (c) Size distribution of spheroids formed on HES 320 (grit-sized sandpaper) with various seeding densities. (d) The maximum number of spheroids formed on the hydrogels. (e) Tracking the position of spheroids within the microstructure of hydrogels.

differences up to approximately 400 μm (Fig. 2(d) and Fig. S1). The Ra of the HES from the 1000, 320, 220, 100, and 60 grit designations was 2.1 ± 0.2, 13.7 ± 3.0, 19.3 ± 1.9, 29.3 ± 2.4, and 45.0 ± 6.7 μm, respectively [35,36]. These results suggest that surface roughness can be controlled by the grit designation of the sandpaper. Various types of hydrogel materials have been used for spheroid culture, including polyethylene glycol, alginate, and poly (2-hydroxyethyl methacrylate), because they are non-adhesive to cells, which can promote cell–cell cohesion [8,28,37]. We chose the synthetic Tet-TA hydrogel as a platform material based on our previous reports of its non-adhesive property [38,39]. A non-cell adhesive surface is critical for spheroid formation because it can promote rapid cell–cell cohesion while minimizing cell adhesion onto the surface materials. Furthermore, the

Tet-TA hydrogel is fabricated by chemical crosslinking, which can stabilize the hydrogel surface structure [31]. 4.2. Spheroid formation on HES As shown in Fig. 3(a), successful spheroid formation was observed on every test group after 24 h. It should be noted that the size distribution of the spheroids was highly modulated by the grit designation (Fig. 3(b)). Spheroids on the flat hydrogel (S-Flat) showed an uncontrolled size distribution, which became broader during prolonged culture time from 101 ± 64 μm on day 1–169 ± 70 μm on day 3. In contrast, spheroids on HES maintained their sizes for 3 days, except for the S-1000 group. Interestingly, the size distribution of the spheroids on 203

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HES was highly selective to a specific range depending on the roughness of the hydrogels. For example, the average diameter was 98 ± 8 μm for S-320 (day 3) and 182 ± 12 μm for S-60 (day 3). S-1000 showed a different trend, more similar to S-Flat. Those trends were prolonged for 7 days (Fig. S5). The size of the spheroids can thus be varied by changing the surface roughness of the hydrogel, unlike AggreWell™, which can only form spheroids of 130 ± 10 μm when cells are seeded at a single seeding density (Fig. S2). Additionally, the size of the spheroids was controlled by the seeding density (Fig. 3(c)). Under the same grit designation (320), seeding densities of 0.5, 1, and 2 × 105 cells/cm2 resulted in average spheroid diameters of 60 ± 5, 92 ± 5, and 129 ± 8 μm, respectively. The S-320 group had the greatest density of spheroids (2274 ± 88 spheroids/cm2) (Fig. 3(d)). The number of spheroids decreased as grit designation decreased (S220: 1815 ± 88 spheroids/cm2, S-100: 518 ± 54 spheroids/cm2, S60: 250 ± 50 spheroids/cm2). The S-320 group effectively produced spheroids with a density about 3.6 times greater per unit area than those fabricated by AggreWell™ (Fig. S3). Each spheroid stayed in its original place for more than 3 days (black arrow in Fig. 3(e)) without fusing with other spheroids. These results collectively suggest that HES was highly effective not only for spontaneous assembly of stem cells with a defined size according to surface roughness, but also for culturing spheroids while preventing fusion of multiple spheroids. Surface structure has previously been reported to control the size of spheroids because protrusions on the surface could inhibit unwanted fusion of multiple spheroids. For example, poly (2-hydroxyethyl methacrylate) hydrogels with wrinkle patterns have been used to fabricate fibroblast spheroids [28]. Similarly, in this research, the embossed structures served as a fusion barrier to induce size-controlled spheroid formation. Furthermore, we demonstrated that modulating the surface roughness allowed us to control the size of the spheroids in the range from 100 to 200 μm. The average size of particles on sandpaper is controlled (P1000 = 18 μm, P320 = 46 μm, P220 = 68 μm, P100 = 162 μm, and P60 = 269 μm) by the ISO 6344 standards (the international standards covering abrasives). Our results suggest that both seeding density and embossed features on the hydrogel can determine the size of resulting spheroids. The regular hill-valley-like protruding structures on the surface of the hydrogel apparently caused a similar number of homogeneously distributed hADSCs to rest in each valley, which led to the formation of spheroids with precisely defined sizes. These results also indicate that the number of cells entering the valleys of the HES at the early stage is an important factor in determining the size of the spheroids. It should also be noted that the HES effectively produced massive numbers of spheroids from a relatively small number of cells (i.e., the S-320 spheroids theoretically contain about 44 cells per spheroid). The commonly used pellet culture and hanging drop methods have difficulty producing a large number of spheroids with a small number of cells in each spheroid due to a lack of aggregation, with as few as 1250 cells and as many as 60000 cells per spheroid [24,40]. Furthermore, our results show that HES has better productivity than AggreWell™, which is a commercially available spheroid culture platform. According to the manufacturer's instruction, only 628 and 157 spheroids can be obtained per cm2 using AggreWell™400 and AggreWell™800, respectively, many fewer than we grew on the HES.

the Y-27632 treatment yielded a slightly aggregated structure. The morphological change in the spheroids formed under Y-27632 was traceable only by the ratio of projected area of spheroids to its circumcircle; the circularity was significantly lower in the Y-27632treated group (62.6 ± 10.9%) than in the control group (88.2 ± 2.5%) (Fig. 4(c)). A brief description of how circularity was calculated can be found in the supportive information (Fig. S1). A live/ dead assay after 24 h confirmed that spheroids formed successfully without severe cellular damage (Fig. 4(d) and Fig. S7). A trypan blue exclusion assay on days 3 and 7 quantitatively supported viability over 7 days, as shown in Fig. 4(e), demonstrating no differences compared to spheroids grown on AggreWell™ (Fig. S4). Furthermore, the proliferation of hADSCs within spheroids prepared on HES was significantly higher on day 7 than on day 3 (Fig. 4(f) and Fig. S9); the optical density at 440 nm was significantly increased in all groups on day 7. Reconstructed 3D images showed that cells within the spheroid were assembled with highly organized f-actin stress fibers after 24 h (Fig. 4(g) and Fig. S10). As shown in Fig. 4(h) and Fig. S11, H&E staining revealed that the inside of each spheroid was compactly organized and contained homogeneously distributed cells in all experimental groups on days 1 and 3. During spheroidization of stem cells, cell–cell interactions are anticipated to be preceded by cadherin or integrin, and then actin cytoskeleton modulation through the Rho/ROCK pathway leads to maturation of the spheroids [41]. Y-27632 and CytoD affect stress fiber formation by inhibiting phosphorylation of the myosin light chain and directly inhibiting actin polymerization, respectively. Previously, Tsai et al. reported that treatment with Y-27632 or CytoD inhibits spheroid formation of mesenchymal stem cells on commercially available ultralow attachment plates [42]. Similarly, our results suggest that actin polymerization following initial cell–cell contact seems to be a major factor in spheroid formation on HES. 4.4. Comparison of gene expression level between spheroids and 2Dcultured cells As shown in Fig. 5(a), gene expression of ECM proteins in the spheroids increased significantly compared with 2D-cultured cells, with no differences based on spheroid size; fibronectin expression rose by 4.26 ± 0.60 (S-60), 5.13 ± 0.07 (S-100), and 4.51 ± 0.78 fold (S320) compared to the 2D group. Similarly, the expression of cell junction–related genes was significantly higher in the spheroids; E-cadherin expression was 4.01 ± 0.27 (S-60), 6.26 ± 0.81 (S-100), and 22.62 ± 5.18 fold (S-320) higher than in the 2D group (Fig. 5(b)). Fig. 5(c) shows significantly expanded stemness marker expression in hADSCs within spheroids relative to the control. The increased stemness marker expression of S-320 was highly maintained even after 7 days of culture (Fig. S12). Interestingly, the expression of cell junction–related genes and stemness genes grew as spheroid size decreased. Thus, we selected S-320 as the representative spheroid group for immunofluorescence staining of ECM and the stemness marker. Fig. 5(d) shows fluorescence staining of the stem cell markers in the S-320 spheroids on day 3. Compared to 2D-cultured cells (Fig. 5(e)), the spheroids showed an intensified fluorescence signal indicating Nanog, Oct4, and SOX2 expression by the hADSCs. These results are consistent with many previous reports showing increased expression of embryonic stem cell markers and ECM molecules in adipose-derived stem cells cultured as spheroids relative to those from 2D cultures [13,14,43,44]. The spheroid environment seems to maximize cell–cell interactions with neighboring cells, aiding cell–cell communication. It was quite interesting that the expression of embryonic stem cell markers and cell–cell junction proteins was inverse to the size of the spheroids. Previous works reported that spheroid size is critical for oxygen supply and the diffusion of signaling molecules; in fact, increased hypoxia level inside of spheroids led to both increased VEGF secretion [24,40] and a necrotic core of apoptotic cells. However, the spheroids used in those studies had a large diameter (200 μm–800 μm).

4.3. Characteristics of spheroids prepared on HES Fig. 4(a) shows the spheroid formation process on HES 320 over 24 h; at 6 h, the cells started to form aggregates, and after 24 h, spheroid formation was complete. The viability of S-320 was 96.3 ± 0.7% after 24 h (Fig. 4(b)). To confirm the importance of cytoskeletal stress fiber formation during spheroidization, we used CytoD and Y-27632. Treatment with CytoD and Y-27632 did not alter the viability of the hADSCs (96.4 ± 1.8% and 94.6 ± 0.1%) after 24 h of culture (Fig. 4(b)). However, treating cells with CytoD abolished spheroid formation, and 204

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Fig. 4. (a) Monitoring cellular assembly on the hydrogel with 320 grit size under various conditions. hADSCs were treated with Cytochalasin D or Y-27632. (b) Viability of spheroids after treatment with Cytochalasin D and Y-27632 for 24 h. (c) Circularity of spheroids formed under treatment with Y-27632. Circularity was calculated by measuring the ratio of projected area of spheroids to the corresponding circumcircle area. (d) Representative live/dead images of spheroids formed on the hydrogels after 24 h. (e) Viability of spheroids at each culture time point. (f) Proliferation of hADSCs within spheroids cultured on the hydrogel. (g) Representative three-dimensional confocal microscopic images of spheroids. (h) H&E stained images of spheroids on days 1 and 3.

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Fig. 5. qRT-PCR results for expression of genes associated with (a) extracellular matrix proteins, (b) cell–cell junction proteins, and (c) stem cell markers. All target gene expression was normalized by GAPDH expression. “#”, “&”, and “$” indicate statistical significance compared to the 2D, S-60, and S-100 groups, respectively. Immunofluorescence staining of Nanog, Oct4, SOX2, fibronectin, collagen IV, and laminin in (d) spheroid (S-320) and (e) 2D-cultured cells. Nuclei were stained with DAPI (blue). (For interpretation of the references to color in this figure legend, the reader is referred to the Web version of this article.)

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Fig. 6. Differentiation capacity of spheroids cultured on embossed hydrogels under induction conditions for 14 days. Phase contrast images of (a) the 2D control group and (b) sectioned images of spheroids stained by Oil red O, Alcian blue, and Alizarin red to confirm adipogenic, chondrogenic, and osteogenic differentiation, respectively. qRT-PCR analysis of (c) adipogenic, (d) chondrogenic, and (e) osteogenic marker gene expression in hADSCs from 2D cultures vs. spheroids. “#” indicates statistical significance compared to the 2D group. (For interpretation of the references to color in this figure legend, the reader is referred to the Web version of this article.)

No previous study has shown how spheroids smaller than 200 μm controlled the expression of stemness markers and cell–cell junction proteins. The spheroid-size dependent characteristics could be explained by the difference in the sizes and shapes of the individual cells embedded within each spheroid. The size or shape of single cells has been reported to affect the properties of mesenchymal stem cells [45]. In addition, cells at the periphery of a spheroid are known to be more stretched than those within the core [41]; thus, as the size of the spheroid increases, the proportion of stretched cells also increases. In

any case, further study is warranted on the expression of hypoxia-related genes and its implications for the regulation of stemness marker and cell–cell junction protein expression. Our hydrogel platform might be a great tool for those studies. 4.5. Differentiation of hADSCs within spheroids Because S-320 showed the best results in Fig. 5, we confirmed the differentiation potentials of S-320 after 14 days of culture under various 207

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Fig. 7. Comparison of the effects of surface microstructure on the shape and function of spheroids formed on commercially available ultra-low attachment plates (LAPs), a smooth hydrogel (Flat), and embossed hydrogel with the 220 grit designation. (a) Phase contrast images of spheroids on each substrate after 3 days and (b) fluorescence images of the LOX-1 hypoxia probe inside the spheroids. (c) Size distribution of spheroids prepared from each substrate cultured for 3 days. Relative gene expression levels of (d) ECM proteins (fibronectin, laminin), (e) cell junction protein (E-cadherin), and (f) pro-angiogenic factors (VEGF, PDGF). (g) Secretion of VEGF, measured by ELISA, at various time-points. “#,” “&,” and “$” indicate statistical significance compared to the 2D, LAP, and Flat groups, respectively.

conditions (Fig. 6(a) and (b)). Oil red O staining revealed large lipid droplets inside the spheroids, indicating successful adipogenic differentiation. Alcian blue staining indicated that chondrogenic differentiation was enhanced within the spheroids by positive staining for acidic polysaccharides, including glycosaminoglycan. Higher calcium deposition inside spheroids positively stained for alizarin red was confirmed under osteogenic differentiation conditions. These results collectively indicate that spheroid culture of hADSCs enhanced their differentiation capacities. Using qRT-PCR, we confirmed significantly upregulated gene expression in spheroids compared to 2D cultures (Fig. 6(c) and (d), and 6(e)). PPARγ and LPL expression was 3.76 ± 0.08 and 19.96 ± 4.40 fold higher than in 2D-cultured cells. In addition, chondroadherin, aggrecan, cartilage oligomeric matrix protein (COMP), and SOX9 were 11.69 ± 1.12, 14.47 ± 1.23, 1.35 ± 0.20, and 6.18 ± 0.21 fold upregulated, indicating enhanced chondrogenic differentiation. Similarly, alkaline phosphatase (ALP), osteocalcin (OCN), and Runx2 expression increased 5.09 ± 0.32, 2.46 ± 0.43, and 5.67 ± 0.51 fold. hADSCs can be differentiated into multiple lineages [46]. Therefore, many studies have used them for various therapeutic purposes, including bone and cartilage regeneration [47,48]. However, hADSCs cultured in ex vivo conditions often lose their original stemness and differentiation ability as their passage number increases [49]. Our results are similar to previous studies that demonstrated that mesenchymal stem cells cultured as spheroids maintained their differentiation

capacities to adipogenic (PPARγ, LPL), chondrogenic (aggrecan, collagen II, SOX9), and osteogenic (OCN, OPN, Runx2) lineages for more than 7 days under individual differentiation conditions [13,44,50]. 4.6. Comparison with spheroids prepared from commercially available 3D culture plates As shown in Fig. 7(a), during 3 days of culture, the LAP and Flat spheroids became noncircular, ovoid, and irregular due to uncontrolled fusion of multiple spheroids, whereas the spheroids grown on HES maintained a regular and circular shape. LOX-1 staining as a hypoxia probe showed that the spheroids on HES were less positively stained, whereas those in the LAP and Flat groups had more intense signals, indicating that those spheroids were more hypoxic due to their enlarged size (Fig. 7(b)). The results of our quantitative analysis of the diameters of the spheroids are shown in Fig. 7(c). The LAP and Flat spheroids exhibited broader size distributions than those from HES. In particular, the hydrogel without an embossed structure (Flat) led to spontaneous fusion of multiple spheroids, and no control over the spheroid size was achieved after 3 days. The size of the spheroids affected the gene expression profile, with spheroids on HES having greater expression of ECM and cell junction genes after 3 days than the LAP and Flat spheroids. As shown in Fig. 7(d) and (e), for example, fibronectin expression was 4.34 ± 0.13 and 3.21 ± 0.16 fold greater in the LAP and Flat groups, respectively, relative to 2D culture and was 5.67 ± 0.11 208

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Fig. 8. (a) Morphology and live/dead assay images of spheroids after injection using a 27-gauge needle. (b) Viability of injected spheroids quantified by trypan blue exclusion assay. (c) 3D-printed spheroids and cell suspension. (d) Viability after printing quantified by trypan blue exclusion assay. (e) Cell retention rate of spheroids within the 3D-printed structure. (f) VEGF secretion of 3D-printed spheroids. (g), (h), (i) qRT-PCR analysis of genes associated with stem cell markers, cell junction markers, and anti-apoptotic markers, respectively. “#” indicates statistical significance compared to the cell suspension group. (For interpretation of the references to color in this figure legend, the reader is referred to the Web version of this article.)

fold higher in the HES group. Interestingly, the gene expression of proangiogenic factors (VEGF, PDGF) in spheroids on HES showed the highest level (Fig. 7(f)). Consistently, VEGF secretion measured by protein level (Fig. 7(g)) from the spheroids on HES was also the highest of any group on day 1 (2D: 1.66 ± 0.10, LAP: 0.81 ± 0.36, Flat: 1.02 ± 0.13, and 220: 2.13 ± 0.17 ng/μg DNA). That trend continued on day 3 (2D: 0.56 ± 0.07, LAP: 2.00 ± 0.34, Flat: 2.98 ± 0.90, and 220: 6.02 ± 1.07 ng/μg DNA). Several studies have reported that cell–cell/cell–ECM interactions and secretion of critical chemokines from spheroids can be modulated by the characteristics of the culture substrate, potentially because of

changes in stiffness or surface chemistry [27,51]. Although both the LAP and Flat hydrogel provide a different physical/chemical environment than that on HES during spheroidization, we strongly believe that the size distribution generated by each surface and the corresponding hypoxic conditions within the spheroid are the primary determinant of spheroid function. Hypoxic core formation inside spheroids is inevitable due to diffusion limitations [52,53]. It is known that the proangiogenic factor secretion, proliferation, and differentiation ability of hADSCs are enhanced under hypoxic conditions [54,55]. However, excessive hypoxia might counteract those metabolic processes, triggering cell death and overall protein downregulation [24]. For 209

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Fig. 9. Schematic representing cell distribution on a flat surface and embossed surface. Spheroids with a particular size form spontaneously on the embossed microstructures, with the protruding structure allowing a controlled number of stem cells to be assembled in each valley and inhibiting the fusion and conglomeration of the spheroids.

(g), (h), and (i) show enhanced stemness marker, cell junction, and antiapoptotic gene expression in the spheroid printing group compared to the cell suspension printing group. Both Nanog and Oct4 expression was 1.21 ± 0.03 fold higher in the spheroid printing group. E-cadherin and Bcl2L1 expression was 1.14 ± 0.03 and 1.19 ± 0.06 fold higher, respectively. There are many methods for delivering cells to targets. Among them, injection of cell suspensions has been widely used because it is a simple and easy process, but the viability of large spheroids can be affected by shear stress during injection, depending on extrusion force and needle size [56]. Therefore, larger needles are preferred, but they can be problematic and highly invasive for patients. Aguado et al. simulated shear stress when PBS passed through a 28G needle (inner diameter = 180 μm) under a 1 ml/min flow rate [57] and confirmed that a low shear stress of less than 10 Pa was generated in a region at the center of the needle that was half as wide as the needle's inner diameter. In the case of S-320, the spheroid diameter was controlled to 95 ± 6 μm, which is half of the inner diameter of a 27G needle (210 μm). Thus, spheroid viability was not affected by passing through the needle. By using these characteristics, spheroids on HES were printed into 3D structures while retaining higher viability than the cell suspension printing group. Several studies have used 3D printing techniques to fabricate and culture spheroids after printing a single cell suspension [58,59]. However, it usually took more than 7 days to form spheroids inside the printed structure, which is a relatively long time compared to direct printing of spheroids assisted by our HES platform. Alginate has been widely used as ink in 3D bioprinting applications [60]. However, because alginate does not contain any cell binding motif (such as an RGD sequence), it can inhibit cell adhesion and proliferation inside of the 3D printed structure. Thus, due to lack of adhesion, programmed cell death can occurred in anchorage-dependent cells, which is called anoikis [61]. Moreover, cell–ECM interaction is important to maintain the stemness of stem cells [62]. Thus, because spheroids maximize cell–cell and cell–ECM interactions, we assume that the spheroid printing group maintained its stemness and anti-apoptotic gene expression inside the 3D-printed structure. On the other hand, because hydrogel has physical properties more similar to the in vivo environment than a culture dish, stem cells cultured within the

example, hADSC spheroids with a diameter greater than 700 μm decrease VEGF secretion compared to smaller spheroids due to severe hypoxic conditions and central cellular cohesion loss and cytoplasmic swelling [24]. Therefore, finding appropriate hypoxic conditions is critical to maximizing the therapeutic potential of spheroids without losing cell viability. Despite the simplicity of the system, it seems hard to control the diameter of hADSC spheroids using the commercially available LAP, which showed a rapid increase in the diameter of spheroids as a function of culture time [43,51]. Collectively, our results suggest that narrow control over the size of spheroids on HES rescues them from severe hypoxia through their smaller size while maximizing VEGF secretion. 4.7. Fabrication of injectable spheroids within a 3D-printed structure We then tested the injectability of hADSC spheroids prepared on HES 60, 100, and 320 using a 27-gauge needle (inner diameter = 210 μm). Spheroids on Flat showed uneven and dissociated shapes after injection; relatively smaller spheroids were able to pass through the needle without cellular damage, but larger spheroids seemed to deform into an ellipsoidal morphology, with a significant drop in viability (Fig. 8(a) and (b)). As shown in Fig. 8(b), the viabilities of spheroids after injection were 89.5 ± 3.4% (S-60), 91.2 ± 1.1% (S100), and 91.3 ± 0.4% (S-320), compared to 84.4 ± 2.2% for S-Flat. Subsequently, we used injectable spheroids for 3D bioprinting. We confirmed that the spheroids were encapsulated in the printed structure without structural deformation immediately after printing and were maintained that way for 7 days (Fig. S13). The viability was preserved both in the spheroid printing group and the cell suspension printing group (Fig. 8(c)). However, the spheroid printing group showed higher viability (90.0 ± 0.6%) than the cell suspension printing group (84.2 ± 1.1%) (Fig. 8(d)). Moreover, cell retention was significantly greater in the printed structures when spheroids were used (Fig. 8(e)). Seven days after printing, 82.8 ± 4.5% of cells remained within the spheroid structures, compared to 70.3 ± 1.7% in the suspension printing group. VEGF secretion from the spheroid printing group (2354 ± 5 pg) was significantly higher than that from the cell suspension printing group (1299 ± 260 pg) (Fig. 8(f)) over 7 days. Fig. 8 210

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hydrogel can maintain their pluripotency or growth factor production [63,64]. Therefore, we assume that the gene expression level of the spheroid printing group did not show a dramatic difference from the group printed using a single cell suspension. Nonetheless, the improved retention of the spheroids and the improvement in their VEGF expression are advantages for biofabrication of engineered tissue with a defined structure. Fig. 9 shows a potential spheroid formation mechanism on the HES. On a flat hydrogel surface, the distribution of cells is hard to maintain evenly, whereas the protrusions on the HES can regulate homogenous cell distribution. Moreover, the protrusions can serve as a fusion barrier, inhibiting undesirable fusion of spheroids. Therefore, the spheroids that form on the HES exhibit a controlled size. In addition, the size and spacing between protrusions can be regulated by grit designation of sandpaper, which allowed us to modulate the size of spheroids as we engineered them. Furthermore, these embossed structures can position individual spheroids within each valley, enabling long-term culture without fusion. As a result, our platform is effective, allowing a large number of size-controlled spheroids to be harvested at a desired diameter by controlling seeding density and grit designation.

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5. Conclusions Despite improved therapeutic effects, stem cell spheroids have not been widely used due to the complexity of previous spheroid production methods. Using the HES developed in this study, we were able to harvest a large number of stem cell spheroids with a precisely controlled size. The embossed structure on the hydrogel surface can easily be fabricated using commercially available sandpapers. hADSCs spontaneously formed spheroids on the HES within 24 h, and the sizes of the spheroids were controlled by seeding density and surface roughness from 100 to 200 μm. The hADSC spheroids prepared on the HES maintained higher stemness under general growth conditions and showed increased efficiency toward osteogenic, chondrogenic, and adipogenic differentiation than 2D-cultured stem cells. Furthermore, spheroids on the HES showed higher pro-angiogenic factor secretion than those formed on commercially available LAPs. Exploiting the advantage of being able to control the size of the spheroids, we used spheroids prepared on the HES in 3D bioprinting applications by creating injectable-sized spheroids. Within the 3D-printed construct, the spheroids demonstrated excellent retention and anti-apoptotic and pro-angiogenic properties. Accordingly, spheroid production using the HES could be used in various fields, including cell therapy and ex vivo tissue reconstruction. Acknowledgements This work was supported by a grant from the National Research Foundation of Korea, which is supported by the Korean government (MEST) (NRF-2016R1A2B3009936). The surfaces of the materials fabricated in this work were analyzed using the SEM installed at the Hanyang LINC + Analytical Equipment Center (Seoul). Appendix A. Supplementary data Supplementary data to this article can be found online at https:// doi.org/10.1016/j.biomaterials.2018.10.025. References [1] P. Zorlutuna, N. Annabi, G. Camci-Unal, M. Nikkhah, J.M. Cha, J.W. Nichol, A. Manbachi, H. Bae, S. Chen, A. Khademhosseini, Microfabricated biomaterials for engineering 3D tissues, Adv. Mater. 24 (2012) 1782–1804, https://doi.org/10. 1002/adma.201104631. [2] M. Zanoni, F. Piccinini, C. Arienti, A. Zamagni, S. Santi, R. Polico, A. Bevilacqua, A. Tesei, 3D tumor spheroid models for in vitro therapeutic screening: a systematic approach to enhance the biological relevance of data obtained, Sci. Rep. 6 (2016)

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