Lotus seedpod-inspired hydrogels as an all-in-one platform for culture and delivery of stem cell spheroids

Biomaterials 225 (2019) 119534

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Lotus seedpod-inspired hydrogels as an all-in-one platform for culture and delivery of stem cell spheroids

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Se-jeong Kima,b,1, Jaesung Parka,b,1, Eun Mi Kima,b, Jong-Jin Choic, Ha-Na Kimc, Ian L. Chind, Yu Suk Choid, Sung-Hwan Moonc, Heungsoo Shina,b,∗ 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 Department of Medical Science, School of Medicine, Konkuk University, Seoul, 143-701, Republic of Korea d School of Human Science, University of Western Australia, Perth, WA, 6009, Australia b

A R T I C LE I N FO

A B S T R A C T

Keywords: Nature-inspired materials Stimuli-responsive hydrogel Stem cell spheroid Spheroid delivery 3D cell culture

3D culture of stem cells can improve therapeutic effects. However, there is limited research on how to deliver cultured stem cell spheroids to the desired target. Here, we developed lotus seedpod-inspired hydrogel (LoSH) containing microwells for culture and delivery of stem cell spheroids. Human adipose-derived stem cells (hADSCs) inside the square microwells (200 or 400 μm in width with various depths) spontaneously formed spheroids with high viability (94.08 ± 1.56%), and fibronectins conjugated to the hydrogel successfully gripped the spheroids, similar to the funiculus gripping seeds in the lotus seedpod. The spheroids slightly bound to the LoSH surface at 37 °C were detached by the expansion of LoSH at lower temperature of 4 °C. After spheroid formation, LoSH was placed on the target substrate upside-down, expanded at 4 °C for 10 min, and removed from the target. As a result, the spheroids within the microwell were successfully transferred to the target substrate with high transfer efficiency (93.78 ± 2.30%). A delivery of spheroids from LoSH to full-thickness murine skin wound with chimney model showed significant enhancement of the number of SMA-positive vessels at day 21 compared to the group received the same number of spheroids by injection. Together, our findings demonstrate LoSH as a one-step platform that can culture and deliver spheroids to a large target area, which will be useful for various biomedical applications.

1. Introduction Cell therapy using stem cells has been actively investigated due to the unlimited self-renewal, multi-lineage differentiation, and paracrine signaling of stem cells. Stems cells in a native environment reside within a 3D complex niche. Thus, the culture of stem cells as a selfassembled spherical 3D aggregate (stem cell spheroids) is known to enhance in vitro stemness and the self-renewal ability of stem cells [1–3]. In addition, stem cell spheroids showed increased expression of cytokines associated with neovascularization, anti-inflammation, and anti-apoptosis, enabling greater in vivo tissue engraftment and neovascularization [4–7]. Fabrication of spheroids commonly relies on techniques such as liquid overlay, spinner flasks, hanging drops, and microwell arrays, which are designed to enforce intercellular cohesion of stem cells while minimizing stem cell adhesion to 2D surfaces [2].

Despite the existence of various culture platforms, in vivo transplantation of spheroids is limited to either direct injection using a syringe or use of biomaterial scaffolds [7,8]. However, it is difficult to treat large tissue defects using direct spheroid injection with controlled injection depth [9,10]. The use of biomaterials is still limited due to side effects associated with immune reactions or the toxicity of degraded by-products [11,12]. Therefore, there is an unmet need for all-in-one platforms that enable efficient production of a large number of size-controlled spheroids and their effective delivery to large injuries. A number of devices mimicking the structure and stimuli-responsiveness of natural organisms have been developed in biomedical applications, including biosensors, drug delivery devices, and human tissue analogues [13–16]. For example, the dome-like protuberances inside octopus suckers and the dense fibrillar structure of the gecko foot surface inspired the creation of wound dressings and drug delivery

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Corresponding author. Department of Bioengineering, Hanyang University, 222 Wangsimri-ro, Seongdong-gu, Seoul, 04763, Republic of Korea. E-mail address: [email protected] (H. Shin). 1 These authors contributed equally to this work. https://doi.org/10.1016/j.biomaterials.2019.119534 Received 1 July 2019; Received in revised form 5 September 2019; Accepted 28 September 2019 Available online 30 September 2019 0142-9612/ © 2019 Elsevier Ltd. All rights reserved.

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2.2. Lotus seedpod-inspired hydrogel (LoSH) fabrication and characterization

patches, respectively, which both showed the ability to attach to wet surfaces [15,17,18]. In addition, dynamic twisted or curved movements through swelling or drying of plant seedpods or mimosas were utilized to engineer artificial muscles that can contract or relax due to external stimulation [19–21]. Hydrogels have been explored for those applications due to their biocompatibility and ability to change shape or stiffness in response to external stimuli including pH, temperature, light, and electrical stimulus [22–27]. Among them, poly(N-isopropylacrylamide) or polyethylene oxide (PEO)-based temperature-responsive hydrogels have been used as cell culture and delivery tools because they allow control of expansion and contraction at physiological temperatures [28–30]. Furthermore, using those temperature-responsive properties, it was also possible to culture and harvest spheroids encapsulated within hydrogels or onto hydrogel-based microwell arrays [31–33]. However, those tools are successful for harvest of spheroids, but insufficient to be used as direct spheroid delivery tools for in vivo and in vitro applications. Here, we demonstrate a biologically-inspired, cost-effective, and reproducibly manageable spheroid culture/delivery platform based on a temperature-responsive and cell-interactive hydrogel, enabling rapid, high-throughput production of size-controlled spheroids and their homogeneous transplantation onto a large area open wound. The hydrogel design was inspired by the natural process, in which lotus seeds are loosely bound to the bottom of a seedpod by a funiculus and released by breaking these connections. By mimicking the seed detachment, spheroids (as seeds) were formed on a hydrogel with numerous pockets (as a seedpod), and the attachment and detachment of spheroids were controlled by thermal expansion of the hydrogel. Fibronectin conjugated with the hydrogel was expected to act like the funiculus of a lotus and bind the spheroids on the surface of the hydrogel to precisely control their attachment and detachment. We named this hydrogel LoSH, which is an abbreviation for ‘lotus seedpod-inspired hydrogel.’

The PDMS master mold was fabricated from a micropatterned silicon wafer prepared using a positive photoresist (AZ7220, AZ Electronic Materials Ltd.) for deep reactive ion etching. Each micropattern on the silicon wafer was 200 × 200 × 80 μm (width x length x depth) or 400 × 400 × 80 μm. A silicone elastomer solution was mixed with curing agent at a 10:1 wt ratio and poured on the micropatterned silicon wafer. After 2 h of curing at 60 °C, the PDMS master was separated and sterilized under UV for 24 h. The surface structure of the PDMS master was observed using a scanning electron microscope (Nova NanoSEM 450, FEI) installed at the Hanyang Center for Research Facilities (Seoul). For fabrication of LoSH, Tet-TA polymer was synthesized as previously described [34]. Tet-TA polymers were dissolved in PBS and mixed with fibronectin, HRP, and H2O2. The final concentrations of each compound were 12% (w/v) for Tet-TA, 0.0025 mg/ ml for HRP, and 0.1% (v/v) for H2O2. The concentration of fibronectin was varied as 0, 25, and 100 μg/ml. The mixed solution was injected into the 1 mm gap between the glass and PDMS master. After 10 min of enzymatic crosslinking, the hydrogel was separated from the PDMS and cut into a disk shape (diameter = 10 mm). The hydrogel was sterilized by treatment under UV for 24 h and kept in the incubator prior to use. To measure the depth and width of microwells on the hydrogel, surface and lateral images were acquired using an inverted microscope (CKX41, Olympus) and analyzed using ImageJ software. Before imaging, the hydrogels were incubated at 37 °C or 4 °C for 1 h. To measure fibronectin incorporation efficiency, 1 ml of fibronectin solution (1 mg/ ml in PBS) was mixed with 100 μl of Texas red solution (10 mg/ml) for 1 h on a rotatory shaker. Unreacted Texas red was removed by a desalting column. The hydrogel formed with Texas red-conjugated fibronectin was incubated in 1 ml of PBS for 24 h under orbital shaking at room temperature to extract unreacted fibronectin. The fluorescence intensity of the supernatant was measured using a spectrophotometer (Varioskan LUX, Thermo Fisher Scientific). The conjugation efficiency of fibronectin was then indirectly assessed. The Young's modulus of the hydrogel was measured using an atomic force microscope (AFM; MFP3D, Asylum Research).

2. Materials and methods 2.1. Materials Tetronic® 1307 (MW 18000) was obtained from BASF (Ludwigshafen, Germany). Horseradish peroxidase (HRP), hydrogen peroxide (H2O2), p-nitro-phenylchloroformate (PNC), tyramine, trypan blue solution, and Y-27632 were purchased from Sigma Aldrich (St. Louis, MO, USA). A Sylgard® 184 silicone elastomer kit was purchased from Dow Corning (MI, USA). Fetal bovine serum (FBS), penicillin/ streptomycin (PS), trypsin/EDTA, and phosphate buffered saline (PBS) were obtained from Wisent (St. Bruno, QC, Canada). Rhodaminephalloidin, Vybrant™ DiD/DIO cell-labeling solution, LIVE/DEAD® viability/cytotoxicity kit, STEMPRO® human adipose-derived stem cells (hADSC), and MesenPRO RS™ medium were obtained from Life Technologies Corp. (Grand Island, NY, USA). Human dermal fibroblasts (hDFBs) and high-glucose Dulbecco's modified Eagle's medium (HGDMEM) were purchased from Gibco (Carlsbad, CA, USA). Mouse antifibronectin antibodies and human plasma fibronectin were purchased from BD Biosciences (Franklin Parks, NJ, USA). Rabbit anti-collagen type IV (Col IV) antibody and rabbit anti-laminin antibody were obtained from Abcam® (Cambridge, MA, USA). Mounting medium was purchased from Vectashield® (Burlingame, CA, USA). Zeba™ Spin Desalting Columns and Quant-iT™ PicoGreen™ dsDNA assay kits were purchased from Thermo Fisher Scientific (Waltham, MA, USA). The Human VEGF standard ABTS ELISA development kit was obtained from PeproTech (Rocky Hill, NJ, USA). The RNeasy Mini Kit was purchased from Qiagen (Valencia, CA, USA). All primers for real-time PCR (RTPCR) were obtained from Cosmogenetech (Seoul, Korea). SYBR Premix Ex Taq was purchased from TAKARA (Otsu, Shiga, Japan).

2.3. Spheroid formation using LoSH hADSCs were cultured in growth medium (99% MesenPRO RS™ and 1% PS) under standard conditions (5% CO2 and 37 °C). Cultured hADSCs (passage number 3–5) were harvested using trypsin/EDTA and resuspended at the desired concentration. For harvesting spheroids, LoSH was placed in a 48-well plate, and hADSCs were seeded at various densities (0.5, 1, 2, and 4 × 105 cells/cm2) and centrifuged at 1200 rpm for 5 min. Phase contrast images of spheroid formation were obtained at 0, 2, 6, and 24 h after the process. The diameters of the spheroids at 24 h were measured using NIS elements software (Nikon). The cell attachment area on LoSH depending on fibronectin concentration was measured using ImageJ software at 2 and 6 h after seeding (attachment area (%) = (cell attachment area on LoSH)/(bottom area of microwell on LoSH) x 100). To confirm the effect of contraction force on spheroid formation on LoSH, the hADSC suspension was treated with Y-27632 at 100 μM. Phase contrast images were obtained at 0, 2, 6, and 24 h. To investigate the effect of fibronectin incorporation on spheroids, hADSCs were seeded (2 × 105 cells/cm2) on LoSH prepared with or without fibronectin. After 24 h of spheroid formation, shear force was applied to the spheroids within microwells using a rotatory shaker (250 rpm, 30 s). Phase contrast images were obtained before and after shaking, and the numbers of microwells occupied and unoccupied with spheroids were counted to calculate the binding efficiency of spheroids (binding efficiency (%) = (number of microwells occupied with spheroids after shaking)/(number of total microwell on LoSH) x 100). To confirm the detachment of spheroids by hydrogel expansion, 2

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spheroids was measured using a trypan blue exclusion assay after dissociation of spheroids by treating with trypsin/EDTA for 5 min. To evaluate VEGF secretion, the spheroids within LoSH microwells were cultured for 7 days. Culture medium was collected at each time point (days 1, 4, and 7) for VEGF ELISA. At each time-point, a DNA assay was performed for each sample to normalize VEGF secretion to total amount of DNA.

spheroids within microwells of LoSH were placed at either 37 °C or 4 °C for 10 min, and the same shear stress was applied. The number of microwells with separated spheroids was counted to calculate the detachment ratio (detachment ratio (%) = (number of spheroid separated microwells after shaking)/(number of total microwells) x 100). For adhesion force measurements, atomic force microscope cantilevers (MFP-3D, Asylum Research) were coated with anti-fibronectin antibody (ab2413). Briefly, cantilevers were immersed in 5 M ethanolamine-HCl in dimethylsulfoxide and incubated for 12 h at room temperature after cleaning with chloroform for 30 s. The cantilever was rinsed with PBS and incubated in 25 mM of bis(sulfosuccinimidyl) substrate for 30 min. The cantilever tips were immersed in antibody solution (diluted at a 1:100) for 30 min after PBS rinsing. Surface-coated cantilever tips were dried and stored at 4 °C until use. Hydrogels with various concentrations of fibronectin (0, 25, and 100 μg/ml) were placed in PBS, and the adhesion force was measured in a 3 × 2 array of points spaced ~1 mm apart using the surface-coated cantilever tips at 2 μm/s. A dwell time of 1 s was added between the approach and retraction to promote antibody binding to fibronectin. Adhesion force was analyzed using the Igor Pro software analysis tool (Asylum).

2.5. Transfer of spheroids from LoSH to the target hADSCs spheroids were prepared in a microwell on LoSH and cultured for 24 h. A cover glass (diameter = 10 mm) was coated with fibronectin solution (20 μg/ml) for 2 h as a target for transferred spheroids. The hydrogel was placed on the target cover glass with spheroids facing the target. Subsequently, expansion of LoSH was induced by dropping cold saline (4 °C) onto the hydrogel for 10 min using a pipette (100 μl/min) (Supplementary Fig. 1). The hydrogel was then removed from the glass. Phase contrast images were obtained at each step. The diameter of spheroids after transfer was measured using ImageJ software. The transfer efficiency of spheroids was quantified by image analysis (transfer efficiency (%) = (number of transferred spheroids)/ (number of total spheroids on LoSH before transfer) x 100). We then investigated the effect of microwell depth on spheroid transfer efficiency. The hydrogels (200 μm patterns) containing microwells with various depths (40, 80, 120, and 200 μm) were fabricated by changing the depth of the silicon wafer (and thus the PDMS master). Then, spheroids were prepared on these hydrogels and transferred to the fibronectin-coated cover glass by the previously mentioned process. The transferred spheroids were then fixed in 4% paraformaldehyde solution for 4 h, rinsed with PBS three times, and treated with rhodaminephalloidin solution for 1 h. Nuclei were counterstained by DAPI, 3D images were obtained using a confocal laser microscope (TCS SP5, Leica), and the height of the transferred spheroids was measured using ImageJ software. The viability of transferred spheroids was examined using the previously mentioned trypan blue exclusion assay and the LIVE/DEAD® viability/cytotoxicity kit. Transfer efficiency was calculated as previously described. To measure the mechanical properties of transferred spheroids and hydrogels, AFM probing was performed with an NX10 (Park Systems, South Korea). A 10.8 μm polystyrene bead was glued to a silicon nitride cantilever with a nominal spring constant of 0.08 N/m (NanoAndMore, USA). The individual cantilever tip was calibrated in a culture medium at 37 °C prior to each experiment by measuring the deflection sensitivity on a glass surface, which allowed the cantilever spring constant to be determined using the thermal noise method [35]. Before placement of the sample, the baseline of the force-distance curve was fitted using SmartScan™ software (Park Systems). After calibration of the bead cantilever, transferred spheroids or hydrogels were loaded, and three or four force-distance curves were sequentially acquired from each point to form a two-dimensional array. The approach and retraction velocities were kept constant at 10 μm/s with 5 nN thresholds in a closed zloop to minimize damage to the sample. The Young's modulus of the sample was obtained from the force-distance curves by fitting the Hertzian model (XEI software, Park Systems). More than 10 spheroids and hydrogels were measured for statistically reliable estimation. The maximum duration of the cell-spheroid experiment was restricted to 2 h to minimize any negative impact due to medium evaporation.

2.4. Characterization of spheroids hADSC spheroids were collected by expansion of LoSH at 4 °C for 10 min for structural analysis. Spheroids were then immersed in 4% paraformaldehyde overnight for fixation. For F-actin staining, fixed spheroids were rinsed with PBS, immersed in rhodamine-phalloidin for 1 h, rinsed with PBS, and mounted using mounting medium containing DAPI. The internal structure was observed using a confocal laser microscope (TCS SP5, Leica). For hematoxylin and eosin (H&E) staining, fixed spheroids were rinsed with PBS and encapsulated in 3% agarose hydrogel. The hydrogel was immersed in graded EtOH for dehydration and paraffinized. The hydrogel was then sliced into 3-μm-thick specimens using a sliding microtome, and the specimens were mounted on a slide for H&E staining. For immunostaining, fixed spheroids in 3% agarose hydrogel were frozen overnight with frozen section compound. The frozen samples were sliced into 10-μm-thick specimens and mounted on a slide. Samples were treated with various antibodies (antifibronectin, anti-collagen, and anti-laminin) and incubated at 4 °C overnight. After rinsing with PBS, samples were treated with secondary antibodies at 37 °C for 1 h, followed by washing with PBS. FITC-conjugated tertiary antibodies were incubated with the samples at 37 °C for 1 h, which were then mounted using the same mounting medium. hADSCs were seeded (1 × 104 cells/well) on a 24-well plate as a 2D control group. Images were obtained using a fluorescence microscope (TE2000, Nikon). RNA was extracted from spheroids and 2D control groups using the RNeasy Mini Kit (Qiagen) for analysis of mRNA expression. The total amount of RNA was measured using a NANODROP 2000 (Thermo Fisher Scientific). Then, cDNA was synthesized using a thermal cycler (MyCycler™, Bio-Rad) after mixing with Maxime RT premix. Relative gene expression levels (laminin, collagen, fibronectin, ITGAV, ITGA2, ITGB1, ITGB3 genes) were determined using SYBR® Premix and a StepOnePlus™ Real-Time PCR system (Thermo Fisher Scientific, Waltham, MA, USA). The primer sequences of each target gene are as follows. Laminin (Fw: 5′-AGC GGA TAT GCA GCT CTT GT-3′, Rv: 5′GCC GTC CAC AAG CTC TAG TC-3′), Collagen (Fw: 5′-GAA GAT ACG GAC CAC CTG GA-3′, Rv: 5′-AGG TGG ACC AAA GTG ACT GG-3′), Fibronectin (Fw: 5′-ACC AAC CTA CGG ATG ACT CG-3′, Rv: 5′-GCT CAT CAT CTG GCC ATT TT-3′), ITGAV (Fw: 5′-AAC TCA AGC AAA AGG GAG CA-3′, Rv: 5′-GGG TTG CAA GCC TGT TGT AT-3′), ITGA2 (Fw: 5′GGG CAT TGA AAA CAC TCG AT-3′, Rv: 5′-TCG GAT CCC AAG ATT TTC TG-3′), ITGB1 (Fw: 5′-CAT CTG CGA GTG TGG TGT CT-3′, Rv: 5′GGG GTA ATT TGT CCC GAC TT-3′), and ITGB3 (Fw: 5′-GAC AAG GGC TCT GGA GAC AG-3′, Rv: 5′-ACT GGT GAG CTT TCG CAT CT-3′). The GADPH housekeeping gene was used for normalization. The viability of

2.6. Verification of the therapeutic effect of transferred spheroids using a skin defect model hADSCs were pre-treated with a DiO cell labeling solution for 2 h at 37 °C to observe transplanted spheroids on an excisional wound site. hADSCs were then seeded (2 × 105 cells/cm2) on LoSH (200) and cultured for 24 h to form complete spheroids before transplantation. For animal studies, a chimney animal model was established in mice as 3

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For example, the width and depth changed from 163.48 ± 2.85 μm (37 °C) to 240.33 ± 1.80 μm (4 °C) and from 83.72 ± 3.22 μm (37 °C) to 102.9 ± 5.55 μm in LoSH (200), respectively (Fig. 1d and e). Fibronectin was enzymatically crosslinked during gelation of the endgroup-functionalized Tet-TA polymer, which showed a conjugation yield of 72.05 ± 1.65% (Fig. 1f).

previously described [36]. Briefly, Balb/c nude mice were anesthetized from intraperitoneal injection of 40 μl of a mixture containing rompun (40 mg/kg) and ketamine (10 mg/kg). An excisional wound at the middle of the dorsal surface was formed in each mouse using a biopsy punch (diameter = 8 mm). To compare the effects of spheroid delivery methods on therapeutic efficacy, we used three experimental groups: a control (without treatment of spheroid), spheroid control, and LoSH. For the spheroid control, spheroids were pre-harvested from the hydrogel by treating in cold medium (4 °C) for 10 min. Harvested spheroids were then spread in the chimney structure. For the LoSH group, a hydrogel with spheroids was placed on the wound site upside-down for 10 min to induce cell attachment. The backside of LoSH was then treated with cold saline (4 °C) for 10 min, and the LoSH was removed. In all groups, the chimney was sealed with transparent film (Opsite, Smith & Nephew) to prevent drying and entry of external substrates. To identify transfer of spheroids into the wound site, mice were euthanized after 3 days (n = 5), and the wound tissues were harvested. Specimens were fixed with 4% paraformaldehyde. The whole wound tissues, fixed for one day, were mounted onto the glass slide, and the remaining solution was removed with a lab wiper. The spheroid hADSCs were confirmed using an Eclips-Ti fluorescence microscope (Nikon). For tissue staining, mice were euthanized after 7 and 21 days (n = 5), and the wound tissues were harvested. Specimens were fixed with 4% paraformaldehyde for seven days, dehydrated with a graded ethanol series (50–100%), and embedded in paraffin. Samples of 5 μm thickness were mounted onto the glass slide and stained with hematoxylin and eosin (H&E) to evaluate the inflammatory process during wound repair. In addition, staining with Masson's Trichrome was performed to measure the presence of collagen in the wound repair tissue. Samples were stained using anti-KI67 (Thermo Scientific) and anti-SMA (Abcam) to estimate cell proliferation expression and microvessel density, respectively. The staining signals were visualized, and an EnVision Detection System (Dako) and hematoxylin (Sigma) were used for counterstaining. To quantify the vascularization and proliferation in tissues, images were obtained at 3 random points in each stained sample and analyzed using ImageJ software. All experiments were approved by the Institutional Animal Care and Use Committee of Hanyang University (Approval No. 2017-0189A).

3.2. Stem cell spheroid formation on microwells of LoSH Size-controlled spheroids were then prepared with various seeding densities on the surface of LoSH (200) and LoSH (400). Representative images of the morphologies of the spheroids from each experimental group are shown in Fig. 2a and b. The sizes of the spheroids increased with an increase in seeding density. For example, the spheroid diameters ranged from 59.60 ± 17.05 μm to 184.62 ± 58.98 μm for LoSH (200) at seeding densities from 0.5 to 4 × 105 cells/cm2, respectively. The number of prepared spheroids on LoSH(200) was about 156 spheroids/mm2. The wider size distribution of the 4 × 105 cells/cm2 group may be due to an overflow of cells from the microwell and unwanted fusion of spheroids (Supplementary Fig. 2). As shown in Fig. 2c, centrifugation allowed human adipose-derived stem cells (hADSCs) to be homogeneously distributed within each microwell, and the cells attached to the surface in 2 h. The attached cells then contracted and began to form spheroidal structures within 24 h. Treatment with Y27632 led to impairment of spheroid formation (Fig. 2d). Cell adhesion was regulated by the concentration of fibronectin conjugated to LoSH (Fig. 2e). As shown in Fig. 2f, the projected cell attachment area increased after 2 h of seeding by 48.42 ± 4.32%, 60.74 ± 5.51%, and 71.97 ± 4.36% in the FN0, FN25, and FN100 groups, respectively. 3.3. Characterization of spheroids F-actin staining revealed that stem cells within the spherical structure exhibited a stretched cell morphology in a homogenous distribution with a compact internal arrangement within 24 h (Fig. 3a and b). Factin appears in a well-circumscribed ring of the spheroid rather than across the whole spheroid. It has been reported that the actin stress fibers of the cells surrounding the spheroid surface developed during reorganization and compaction of stem cell spheroids after initial aggregate formation [37]. This may be attributed to intensive F-actin staining in the periphery of the spheroids. The gene expression for ECM proteins such as laminin, collagen, and fibronectin significantly increased by 3.96 ± 1.86-fold, 14.42 ± 6.46-fold, and 55.52 ± 16.23fold, respectively, within spheroids compared with that in 2D cultured cells (Fig. 3c). In addition, expression of the integrin subunits ITGAV, ITGA2, ITGB1, and ITGB3 increased by 12.59 ± 4.38-fold, 19.16 ± 9.85-fold, 6.82 ± 0.28-fold, and 2.74 ± 0.49-fold, respectively, in spheroids relative to 2D culture (Fig. 3d). The spheroids showed proliferation with a 1.57 ± 0.10-fold increase in DNA content over 4 days (Fig. 3e). Viability after formation of spheroids was highly preserved for 94.08 ± 1.76% of spheroids in LoSH (200) and 93.80 ± 1.56% of spheroids in LoSH (400) (Fig. 3f). Immunofluorescence staining for laminin, collagen, and fibronectin also supported ECM-rich spheroid formation (Fig. 3g). Moreover, the amount of vascular endothelial growth factor (VEGF) secreted by the spheroids was 23.02 ± 1.08 ng/μg DNA after 7 days, which was significantly greater than that from 2D-cultured stem cells (2.62 ± 0.16 ng/μg DNA) (Fig. 3h).

2.7. Statistical analysis All quantitative results are presented as mean ± standard deviation. 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 = 5). 3. Results 3.1. Lotus seedpod-inspired hydrogel (LoSH) The engineering principles of the hydrogel mimicking detachment of lotus seeds from the funiculus of the seedpod are illustrated in Fig. 1a. The lotus seedpod holds a seed through its linkage to the funiculus, and external forces such as fluid movement can cleave these mechanical links, enabling seeds to be released from the seedpod. Inspired by this process, we designed a hydrogel (LoSH) in which the spheroids are gripped by cell-adhesive proteins in microwells on the hydrogel surface. As shown in Fig. 1b, polydimethylsiloxane (PDMS) molds with two sizes of square micropatterns (200 and 400 μm) were used for fabrication of microwells on the surface of the hydrogel. The resulting hydrogels (LoSH (200) from the 200 μm PDMS mold and LoSH (400) from the 400 μm PDMS mold) demonstrated homogeneous cuboid microwell structures on the surface with temperature-induced expansion at 4 °C relative to 37 °C (LoSH (200) in Fig. 1c). The overall ratio of shape change of microwells upon reduction of temperature from 37 °C to 4 °C was 1.47 ± 0.03 in width and 1.23 ± 0.08 in depth.

3.4. Attachment of spheroids to the surface of LoSH We hypothesized that (1) fibronectin would serve as the funiculus in the lotus seedpod and sufficiently grip the spheroid within the microwell, and (2) temperature-induced expansion of the hydrogel would break this weak bonding. After application of external shear stress using a rotatory shaker, most spheroids from the microwells of hydrogels 4

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Fig. 1. Characteristics of the lotus seedpod-inspired hydrogel (LoSH). a Spheroid culture and detachment mechanism inspired by the release of seeds from the lotus seedpod. b Fabrication process of LoSH using microfabricated polydimethylsiloxane (PDMS) master mold. c Size change of LoSH and its microwells (depending on temperature change from 37 °C to 4 °C) fabricated from a PDMS master of 200 μm pattern (scale bar: 5 mm (low magnification), 200 μm (high magnification)). d, e Size change of microwells within LoSH depending on temperature. f Conjugation efficiency of fibronectin incorporated into LoSH. In all figures, # indicates statistical significance.

Fig. 2. Spheroid formation on LoSH. a, b Size distribution of spheroids with various seeding densities on LoSH (200) and LoSH (400). Images in the graph show representative spheroids formed in each experimental group. c Spheroid formation of cells seeded on LoSH over time (scale bar: 200 μm). d Morphologies of cells on LoSH after treatment with Rho kinase inhibitor (scale bar: 200 μm). e Cell adhesion on LoSH 2 h after seeding depending on incorporated fibronectin concentration (FN0: 0 μg/ml, FN25: 25 μg/ml, FN100: 100 μg/ml). Projected cell attachment area is highlighted in red (scale bar: 100 μm). f Quantification of projected cell attachment area on LoSH 2 h after cell seeding. In all figures, # indicates statistical significance. (For interpretation of the references to colour in this figure legend, the reader is referred to the Web version of this article.)

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Fig. 3. General characteristics of spheroids formed on LoSH. a Cytoskeletal structures inside the spheroid on LoSH (200). Nuclei were counterstained by DAPI (scale bar: 50 μm). b H&E staining of spheroids on LoSH (200) (scale bar: 50 μm). c, d Relative gene expression level of ECM (laminin, collagen, fibronectin) and integrin (ITGAV, ITGA2, ITGB1, ITGB3) in spheroids on LoSH (200) compared with that from two-dimensionally cultured cells. Gene expression level was normalized by GAPDH expression. e Proliferation of cells inside spheroids on LoSH (200) for 1 or 4 days measured by total DNA content change. f Viability of cells inside the spheroid 24 h after cell seeding on LoSH. g Immunofluorescence staining of laminin, collagen, and fibronectin in spheroids on LoSH (200) and two-dimensionally cultured cells (scale bar: 100 μm). h Secretion of VEGF from spheroids on LoSH (200) and two-dimensionally cultured cells at various time points (days 1, 4, and 7) as measured by ELISA. In all figures, “#” indicates statistical significance.

Fig. 4. Spheroid attachment on the LoSH and separation of spheroids by lowering the temperature. a, b Images of spheroids on LoSH (with different fibronectin concentrations of 0 and 100 μg/ml) before and after shaking (250 rpm, 30 s) (scale bar: 1 mm). c Binding efficiency of spheroids on LoSH depending on incorporated fibronectin concentration (0 and 100 μg/ml). d The measurement of adhesion force between antibody-coated AFM cantilever and LoSH incorporating fibronectin. e Images of LoSH after shaking under the indicated temperatures (scale bar: 200 μm). f Detachment ratio of spheroids after shaking depending on temperature. In all figures, “#” indicates statistical significance.

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Fig. 5. Homogenous delivery of spheroids from LoSH to target. a Schematic diagram and images of spheroid transfer (scale bar: 200 μm). b Diameter of spheroids before and after transfer. c Spheroid transfer efficiency from LoSH to target substrate depending on concentration of incorporated fibronectin (0, 25, and 100 μg/ml). d Spheroid transfer efficiency from LoSH to target substrate depending on temperature (37 and 4 °C).

without fibronectin rapidly detached, while fibronectin-incorporated hydrogels stably held the spheroids within the microwells. The binding efficiency of spheroids increased with fibronectin concentration, from 48.37 ± 2.05% (FN0) to 96.70 ± 2.96% (FN100) and from 13.61 ± 0.64% (FN0) to 95.12 ± 0.19% (FN100) on LoSH (200) and LoSH (400), respectively (Fig. 4a–c). We then measured the adhesive force between antibody-coated AFM cantilever and the hydrogel incorporating various concentrations of fibronectin by using AFM (Fig. 4d). The values were 2.20 ± 0.19 μN and 2.22 ± 0.50 μN for the FN0 and FN25 groups, respectively, which significantly increased to 2.99 ± 0.18 μN for the FN100 group. We next examined the detachment of spheroids from fibronectin-incorporated hydrogel by expansion of LoSH according to temperature (Fig. 4e). Shaking at 37 °C did not result in detachment of the spheroids; however, most spheroids detached from the surface at 4 °C (black arrows) with application of shear stress. The detachment ratio significantly increased from 5.38 ± 2.78% (37 °C) to 80.09 ± 6.15% (4 °C) and from 9.71 ± 4.14% (37 °C) to 81.90 ± 5.91% (4 °C) in LoSH (200) and LoSH (400), respectively (Fig. 4f).

for LoSH (400), as shown in Fig. 5b). The spheroid transfer efficiency was dependent on the concentration of conjugated fibronectin: 10.58 ± 5.30% (FN0), 23.88 ± 3.78% (FN25), and 93.78 ± 2.30% (FN100) (Fig. 5c). Thermal expansion of LoSH was critical for transfer of spheroids, and transfer efficiency was significantly reduced to 32.54 ± 9.01% at 37 °C (Fig. 5d). 3.6. Transfer efficiency of spheroids based on microwell depth on LoSH We then analyzed the effect of depth of microwells on spheroid transfer, with depth varied from 40 to 200 μm. As illustrated in Fig. 6a, excessively shallow or deep microwells may either crush the spheroid or inhibit direct contact of the spheroid with the target surface, respectively. Thus, depth of microwell must be controlled during fabrication by approximating the size of the spheroid. As anticipated, the spheroids from microwells of depths from 40 to 120 μm were homogenously transferred, while the arrangement of transferred spheroids with a depth 200 μm was disturbed (Fig. 6b). Confocal microscopic images confirmed that the spheroids were highly deformed as the depth of the microwell became shallow (Fig. 6c). The transferred spheroids were well-adhered to the target surface (white dotted line in Fig. 6c); however, the spheroids from a depth of 200 μm appeared to have minimal contact with the target surface (white arrows in Fig. 6c). As shown in Fig. 6d and e, the viability of spheroids was highly maintained over 90% even after the transfer process, with slightly decreased viability in the 40-μm group (86.58 ± 3.08%), potentially due to compressive stress during transfer. As shown in Fig. 6f, the transfer efficiency of each group was over 95% except that of the 200-μm group (85.65 ± 3.90%). The height of the transferred spheroids in each group was significantly increased from 42.97 ± 7.71 μm (40-μm group) to 91.74 ± 7.69 μm (200-μm group) (Fig. 6g).

3.5. Transfer of spheroids from hydrogel to targets We then performed proof-of-concept experiments, wherein we transferred spheroids within the microwell to the target substrate by exploiting the temperature-responsive expansion properties of the hydrogel as shown in Fig. 5a. The spheroids were weakly bound to the surface during movement or even inversion of the hydrogel, allowing us to place the LoSH upside-down on the target without loss of spheroids. Cold saline (4 °C) was dropped on the hydrogel surface, and the hydrogel was removed after 10 min. As shown in Fig. 5a, spheroids were successfully delivered and re-attached to the fibronectin-coated glass coverslip with uniform spacing over a large area. Notably, spheroids were successfully transferred from the hydrogel to the target, although some of them were in contact with the walls as well as the bottom of the microwell (Supplementary Fig. 2). The projected diameter of the transferred spheroids slightly increased compared to that before transfer (133.00 ± 7.31 μm for LoSH (200) and 178.36 ± 11.04 μm

3.7. Delivering spheroids on a mouse skin defect model using LoSH The therapeutic effect of spheroids was then examined using a mouse skin defect model. During transplantation of spheroids, the LoSH was inverted on top of an excisional wound formed on mouse skin 7

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Fig. 6. Spheroid transfer depends on depth of the microwell on the LoSH. a Schematic diagram showing the effect of microwell depth (40, 80, 120, and 200 μm) on deformation of spheroids formed on LoSH. b Fluorescence images of spheroids formed on LoSH with various microwell depths and transferred to a large target area. F-actin and nuclei were stained red and blue, respectively (scale bar: 200 μm). c Magnified images of transferred spheroids. Top and side views are shown. White dotted lines indicate the border with target (scale bar: 100 μm). d Representative live/dead images of transferred spheroids (scale bar: 200 μm). e Viability of transferred spheroids as a function of depth of microwells. f Transfer efficiency of spheroids depending on depth of microwells. g Height of transferred spheroids for various microwell depths. (For interpretation of the references to colour in this figure legend, the reader is referred to the Web version of this article.)

of spheroids was used as a spheroid control. After 3 days, transplanted spheroids were observed (white arrows) both in the spheroid control and LoSH groups (Fig. 7d), with a greater level of retention of spheroids in the LoSH group. At 7 and 21 days after transplantation, the transplanted cells remained in the wound (white arrow), and a larger

(Fig. 7a). As shown in Fig. 7b, spheroids (black arrow) were successfully transferred to the wound site and maintained a homogenous distribution. Transferred spheroids were stably localized within the wound site even after insertion of the chimney (Fig. 7c). The control group was a non-treated sham group, and suspended treatment of the same number 8

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Fig. 7. Delivery of spheroids to the skin wound model using LoSH. a Spheroid transfer from the LoSH to the wound site using the chimney model. b, c Transferred spheroid on the surface of an excisional wound (black arrow). Chimney inserted into the excisional wound after spheroid transfer using LoSH. d Remaining spheroids on the wound surface (white arrow) 3 days after transplantation in the indicated experimental groups (scale bar: 500 μm). e Fluorescence signals from transplanted cells (white arrow) within sectioned tissue at 7 and 21 days after spheroid transfer (scale bar: 100 μm).

field at day 21).

number of cells was observed in the LoSH group (Fig. 7e). The transplanted spheroids may have disintegrated into single cells after 7 days.

4. Discussion 3.8. Effect of spheroid delivery on a mouse skin defect model As presented in Fig. 1a, a lotus seedpod has many pockets, each of which can hold a spherical seed linked to the funiculus in the bottom of the ovary well of each pocket. Interestingly, the funiculus does not permanently ‘hold’ and ‘fix’ the seed, but provides ‘temporal weak binding’ to the lotus seed within the pocket. Thus, the seed can be detached by cleavage of the funiculus when an external force is applied. By mimicking this natural structure and process, we devised a hydrogel platform with numerous microwells (similar to pockets in seedpod), in which the spheroids are formed by centrifugation and temporarily adhered to the bottom of the microwell due to the presence of cell-adhesive proteins (like the funiculus). This temporal adhesion can hold spheroids under normal processes at 37 °C, and we were able to break the temporal adhesion and separate the spheroids using thermal expansion of hydrogel at a desired point in time at low temperature. Although various spheroid culture platforms with a microwell structure

Histological analysis revealed that the wound was healed with newly formed vascular structures in the LoSH group at 21 days after transplantation (Fig. 8a). Compared with other groups, regeneration was enhanced in the LoSH group. MT staining also confirmed the regeneration of dense and thick subcutaneous collagen layers in the LoSH group compared with the other groups (Fig. 8b). In particular, inflammatory cells quickly disappeared, and a large number of vascular structures formed. As a result, the number of SMA-positive vessels significantly increased in the LoSH group compared with other groups at day 21 (Fig. 8c), with SMA-positive vessel counts of 4 ± 1 (control), 5 ± 1 (spheroid control), and 9 ± 1 (LoSH), as shown in Fig. 8d. In addition, as shown in Fig. 8e and f, cell proliferation was significantly enhanced in the LoSH group as measured by Ki67 staining (control: 17 ± 5, spheroid control: 27 ± 2, LoSH: 32 ± 3 Ki67-positive cells/ 9

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Fig. 8. Cutaneous regeneration by delivery of spheroids in vivo. a H&E staining of each experimental group after 21 days (scale bar: 250 μm). b Masson's trichrome staining of each experimental group after 21 days (scale bar: 250 μm). c SMA staining of each experimental group after 21 days. Blood vessel structure was stained (black arrow) with antibody (scale bar: 100 μm). d Number of SMA-positive blood vessels in each experimental group at days 7 and 21. e Ki67 staining images of each experimental group after 21 days. Black arrows indicate Ki67-positive area (scale bar: 50 μm). f Number of Ki67-positive cells on each experimental group at days 7 and 21.

dynamic coil-to-globule transition of PEO blocks at a reduced temperature of 4 °C [41,42]. In situ conjugation of proteins or peptides containing a phenol group in the hydrogel has been previously reported [30,34,43]. It should be noted that the balance between cell-cell binding and cell-substrate interaction appears to allow cells to either be attached as a 2D monolayer or to form 3D aggregates, although the surface conveys cell binding signals. For example, our previous study using the same material as in this manuscript demonstrated cell sheet formation on the flat surface of hydrogel [30]. Conversely, in another study, spontaneous spheroid formation was reported on the cell adhesive fibrous scaffolds with a microwell structure when seeding a large number of cells [8]. Thus, despite the presence of fibronectin in the LoSH, stem cells formed spheroids within the microwell structure, in which the seeded cells sank and adhered to the surface of hydrogel within the well in 2 h. They, then, aggregated to form a spheroid over 24 h, as shown in Fig. 2c. This may be due to excessive cell-cell/cellextracellular matrix (ECM) interactions and rearrangement mediated by cytoplasmic stress fiber formation in stem cells, as previously reported [1,37]. Inhibition of spheroid formation by treatment with Y-27632 re-

have been extensively studied, none designed a mechanism to hold a spheroid within each microwell. For these approaches, previous platforms inhibited cell adhesion by utilizing non-adhesive materials while maximizing cell-cell interactions, resulting in spheroid formation [38,39]. In contrast, we controlled the interactions between the spheroids and hydrogel substrate for culture and on-demand release of spheroids. Temperature-responsive hydrogels have been synthesized using many polymers including poly(N-isopropylacrylamide), polyethylene glycol, poly(vinyl methyl ether), and PPO-PEO block copolymer [25,26]. These polymers exhibited transitions from the gel to sol state below a lower critical solution temperature, which can be controlled with polymer composition, functional groups in the polymer main chain, length of hydrophilic segments, and molecular weight [40]. These temperature-responsive properties have been used in biomedical applications including delivery of drugs and cells [23,25]. The Tet-TA hydrogel used in this study was prepared by crosslinking of a terminally-modified PPO-PEO block copolymer. We observed rapid expansion of the hydrogel, reaching the maximum size within 10 min due to the 10

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therapeutic outcomes in previous works [37,57]. However, no report has been available on a biomaterial-based platform to harvest sizecontrollable multi-cellular spheroids and deliver them to a target on a large injured region. Conventional delivery of spheroids relies on syringe-based injection. However, injected spheroids can flow back to the injection port after injection, and the spheroids cannot be directly transplanted onto the open wound surface, such as an excisional wound [6]. Furthermore, it was difficult to confirm a direct therapeutic effect of spheroids injected into the subcutaneous muscular layer [57]. Our results suggest that uniform delivery of spheroids to a large open wound may be achieved using the present novel spheroid delivery platform. No study has yet reported the deformation of spheroids under a compressive stress similar to our transfer process. Therefore, we analyzed the mechanical properties of the hydrogel and spheroids to verify that spheroids can be deformed during transfer without negative effect. As shown in Supplementary Fig. 4, the Young's modulus of fibronectinconjugated LoSH (200) was 13.4 ± 2.6 kPa, which is similar to that of a previously reported fibronectin-conjugated Tet-TA hydrogel [30]. However, this is a much higher value than that of spheroids (2.113 ± 0.577 kPa) (Supplementary Fig. 5), which may lead to spheroid deformation when compressive pressure is applied by the hydrogel. More pressure was applied on the spheroids as the microwell on the LoSH became shallower. It should be noted that successful transfer requires direct contact of the spheroid with the target surface, which seems to accompany severe deformation of spheroids and a decrease in viability when the microwell depth was too shallow. Given that spheroids in a microwell of depth 200 μm showed reduced transfer efficiency, the appropriate depth of microwells is important to maintain the viability of delivered spheroids. Confocal images demonstrated that the actual structures of spheroids were slightly ellipsoidal, as shown in Fig. 6b and c. These results are consistent with previous studies showing that spheroids on hydrogels with a cell adhesive micropattern showed an ellipsoid structure rather than a sphere due to interactions between the spheroid and surface of the hydrogel [58,59]. Skin is one of the most ECM-rich tissues and contains various types of ECM proteins, such as collagen, fibronectin, and laminin, which have abundant cell binding moieties. Transplanted cells on skin through cell spray or cell sheets have been successfully integrated with host tissue [60–63]. Using a cell sheet or cell spray rather than a scaffold on a skin wound has great advantages due to the curved surface structure and low stiffness of skin tissue [64]. In this study, LoSH was able to cover the curved surface of a skin defect, and spheroids on LoSH were successfully attached to skin defects in a homogenous manner. Meanwhile, numerous studies have reported enhanced wound regeneration by transplantation of spheroids, but most studies injected spheroids into peripheral parts of the wound [57,65]. Confirming the direct effect of the implanted spheroid in these cases was difficult. An excisional wound model is an appropriate approach to determine whether spheroids delivered to a wound area show an improved therapeutic effect because of the exposed and large nature of the wound. However, it is difficult to confirm the effect of only the implanted spheroids due to rapid skin contraction in the mouse excisional wound model. Therefore, we used the chimney wound excision model, in which a sectioned tube was inserted around the wound to prevent rapid skin contraction and to confirm the skin regeneration effect of spheroids in human skin-like conditions [66]. hADSC spheroids have been shown to exert excellent therapeutic effects when they are transplanted into an excisional wound due to enhanced secretion of growth factors such as basic fibroblast growth factor (bFGF) and hepatocyte growth factor (HGF) [65]. Hypoxia inducible factor expression was also increased in hADSCs cultured as spheroids, which leads to enhanced neovascularization through VEGF secretion [57]. Meanwhile, the improved therapeutic effect of the LoSH group may be explained by multiple roles of spheroids within the defect. The distance between spheroids is approximately 200 μm, which is an effective distance for paracrine signaling. The homogeneously

confirmed the importance of the contractive force exerted by stress fiber formation within stem cells [44]. Therefore, our results indicate that hADSCs were attached as a monolayer on the surface of the hydrogel within the well, 2–3 h after centrifugation, and the monolayer was subsequently contracted to form a spheroid due to a relatively greater contraction force of cells over cell adhesion to the surface. The cell adhesion was controlled by the number of fibronectin binding sites available on the hydrogel. Meanwhile, following spheroid formation, the presence of fibronectin may serve to weakly hold spheroids through minimal contact, which is described in Supplementary Fig. 3. Stem cells cultured as 3D spheroids enhanced expression of inflammatory cytokines, increased stemness marker expression, and increased differentiation potential due to the abundant cell-cell interactions and cell-ECM communications [3,45]. Consistent with these results, the spheroids from our hydrogel showed over-expression of ECM proteins, integrin, and VEGF [46,47]. The increased VEGF secretion within spheroids may be attributed to several factors. First, upregulation of hypoxia inducible factor mediated by potentially lower oxygen concentration at the core of spheroids may affect the VEGF secretion as previously reported [48]. Secondly, since spheroid has increased cell-cell/cell-ECM interactions compared to two-dimensionally cultured cells, spheroid may have a positive effect on signal transduction, leading to an increase in VEGF secretion [49]. Meanwhile, despite significant progress in the study of stem cell spheroids, many studies have shown conflicting results regarding proliferation of stem cell spheroids [46,50–52]. Among them, proliferation of ADSC spheroids seems to vary depending on culture substrate [46,50]. Some substrates such as chitosan and hyaluronan supported proliferation of ADSCs within the spheroids [50]. We may use other polymers and materials to confirm the proliferation under similar microwells in a future study. Fibronectin conjugated to LoSH is a binding site for cells on the hydrogel. The relatively high density of cells exert high levels of contractile force, which may not be resisted by the limited number of cell binding to fibronectin. Thus, spontaneous spheroid assembly at the center of individual microwells may have occurred. After spheroid formation, some spheroids may touch the walls of the microwell; however, this does not seem to affect transfer efficiency, as shown in Fig. 5. The overall adhesion force between a cell and substrate increases as a function of the number of fibronectin binding to integrin transmembrane receptors [53,54]. Although measurement of the adhesion force between a single cell and substrate has been established using a force microscope, it is still technically challenging to directly measure the force between spheroid and substrate [55]. The contact area by spheroid onto LoSH may be changed according to the size of the spheroid when cultured on the surface of LoSH, and this can affect the adhesion force. In addition, some spheroids may interact with the wall as well as the bottom of the microwell present on the surface of the LoSH, which may affect the adhesion force. Therefore, we decided to indirectly show the interactions between cells (or spheroids) and LoSH conjugated with fibronectin in various ways (Figs. 2f, 4a, b, d). Our results showed that an increase in the number of fibronectin in LoSH with FN100 resulted in increase in overall adhesion force of AFM cantilever to the hydrogel. This concentration did not affect spheroid formation, but was sufficient to grip spheroids formed within each microwell. When the hydrogel expands in response to temperature change, the distance of cell-fibronectin binding sites may become wider, allowing detachment of the spheroids from LoSH. Cell delivery using temperature-responsive materials (e.g., poly(Nisopropylacrylamide)) has been widely used in cell sheet technology [28]. Multi-layered, co-cultured, and vascularized cell sheets have been prepared for transplantation to various targets [29,56]. Despite successful use of cell sheets in regenerative medicine, the cell sheet is mechanically weak as a single layer. This causes problems in handling, and repeated cell seeding is necessary to produce a multi-layered cell sheet. Meanwhile, spheroids have been transplanted for improved 11

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No. NRF-2018M3C1B7021994 and NRF-2019R1A2C2084965). For fabrication of the micropatterned silicon wafer, we are grateful to Jungyul Park and Hyung-kwan Chang (Department of Mechanical Engineering, Sogang University, Seoul, Republic of Korea). We are also grateful to Luke G. Major (School of Human Science, University of Western Australia, Perth, Australia) for assistance with adhesion force measurements using AFM. We would also like to express gratitude to Dae Sung Yoon, Gyudo Lee, and Dongtak Lee (School of Biomedical Engineering, Korea University, Seoul, Republic of Korea) for helping us measure the mechanical properties of materials and spheroids using AFM. The morphology of transferred spheroids in this work was analyzed using the confocal laser microscope installed at the Hanyang LINC + Analytical Equipment Center (Seoul).

distributed spheroids may have secreted signals in a geometrically regulated manner, which may have improved the therapeutic effect. The transplanted spheroids in our delivery platform may be favorable for engraftment into native tissue with improved survival rates compared to randomly distributed spheroids. This is also due to improved signaling by neighboring spheroids. In addition, we applied spheroids to removable medical patches with mild pressure, which may have improved integration and retention of spheroids. As shown in Fig. 7, more spheroids were retained upon transplantation using LoSH after 3 days as compared to control groups. In contrast, spheroid control showed dispersed (not well-organized) distribution of spheroids, which did not seem to be well-integrated into the wound site. The efficiency of engraftment may lead to differences in therapeutic effects. Most previous studies have used non-adhesive materials to maximize cell-cell interactions for spheroid fabrication [67–70]. There are two main technical advances in this study. (1) We developed an all-inone platform for harvest/culture of spheroids and their delivery to tissue defects with a relatively large area in a well-organized manner. All previous studies demonstrated spheroid formation in various conditions; however, transplantation of spheroids was achieved only through syringe-based injection. To our knowledge, no techniques have been reported in which individual spheroids are homogeneously transplanted to the tissue with the 200 μm distance shown in Fig. 7b. Transplanted spheroids may be more effective in paracrine signaling within this distance, producing improved cell survival. (2) We report cell-adhesive hydrogels as an interactive surface to partially grip prepared spheroids, which are then released by breaking these bonds by rapid expansion of the hydrogels in response to temperature change. In addition, the cell adhesive surface within the microwell can support spheroid formation under high density seeding and centrifugation. Therefore, synthetic cell-adhesive substrates can be exploited for applications that require reversible cell attachment/detachment under various physiological environments.

Appendix A. Supplementary data Supplementary data to this article can be found online at https:// doi.org/10.1016/j.biomaterials.2019.119534. Author contributions S-.K. and J.P. designed and performed experiments. E.M.K. collected and analyzed microscopic images. J-.C., H-.K., and S-.M. performed animal experiments and analyzed the results. I.L.C. and Y.S.C. performed physico-chemical analysis of the hydrogel. H.S. managed the project. All authors discussed the results and contributed to the manuscript. References [1] A.-C. Tsai, Y. Liu, X. Yuan, T. Ma, Compaction, fusion, and functional activation of three-dimensional human mesenchymal stem cell aggregate, Tissue Eng. A 21 (2015) 1705–1719, https://doi.org/10.1089/ten.tea.2014.0314. [2] E. Fennema, N. Rivron, J. Rouwkema, C. van Blitterswijk, J. De Boer, Spheroid culture as a tool for creating 3D complex tissues, Trends Biotechnol. 31 (2013) 108–115, https://doi.org/10.1016/j.tibtech.2012.12.003. [3] Z. Cesarz, K. Tamama, Spheroid culture of mesenchymal stem cells, Stem Cell. Int. 2016 (2016) 1–11, https://doi.org/10.1155/2016/9176357. [4] Y. Petrenko, E. Syková, Š. Kubinová, The therapeutic potential of three-dimensional multipotent mesenchymal stromal cell spheroids, Stem Cell Res. Ther. 8 (2017) 94, https://doi.org/10.1186/s13287-017-0558-6. [5] T.J. Bartosh, J.H. Ylöstalo, N. Bazhanov, J. Kuhlman, D.J. Prockop, Dynamic compaction of human mesenchymal stem/precursor cells into spheres self-activates caspase-dependent IL1 signaling to enhance secretion of modulators of inflammation and immunity (PGE2, TSG6, and STC1), Stem Cells 31 (2013) 2443–2456, https://doi.org/10.1002/stem.1499. [6] Y. Xu, T. Shi, A. Xu, L. Zhang, 3D spheroid culture enhances survival and therapeutic capacities of MSCs injected into ischemic kidney, J. Cell Mol. Med. 20 (2016) 1203–1213, https://doi.org/10.1111/jcmm.12651. [7] S.H. Bhang, S. Lee, J.-Y. Shin, T.-J. Lee, B.-S. Kim, Transplantation of cord blood mesenchymal stem cells as spheroids enhances vascularization, Tissue Eng. A 18 (2012) 2138–2147, https://doi.org/10.1089/ten.tea.2011.0640. [8] K. Wang, X. Wang, C. Han, W. Hou, J. Wang, L. Chen, Y. Luo, From micro to macro: the hierarchical design in a micropatterned scaffold for cell assembling and transplantation, Adv. Mater. 29 (2017) 1604600, https://doi.org/10.1002/adma. 201604600. [9] J. Wang, J.V. Jokerst, Stem cell imaging: tools to improve cell delivery and viability, Stem Cell. Int. 2016 (2016) 1–16, https://doi.org/10.1155/2016/9240652. [10] D.J. Mooney, H. Vandenburgh, Cell delivery mechanisms for tissue repair, Cell Stem Cell 2 (2008) 205–213, https://doi.org/10.1016/j.stem.2008.02.005. [11] T. Garg, O. Singh, S. Arora, R.S.R. Murthy, Scaffold: a novel carrier for cell and drug delivery, Crit. Rev. Ther. Drug Carrier Syst. 29 (2012) 1–63, https://doi.org/10. 1615/CritRevTherDrugCarrierSyst.v29.i1.10. [12] F.N. Alaribe, S.L. Manoto, S.C.K.M. Motaung, Scaffolds from biomaterials: advantages and limitations in bone and tissue engineering, Biologia (Bratisl) 71 (2016) 353–366, https://doi.org/10.1515/biolog-2016-0056. [13] A. Vallée-Bélisle, K.W. Plaxco, Structure-switching biosensors: inspired by Nature, Curr. Opin. Struct. Biol. 20 (2010) 518–526, https://doi.org/10.1016/j.sbi.2010. 05.001. [14] J.-W. Yoo, D.J. Irvine, D.E. Discher, S. Mitragotri, Bio-inspired, bioengineered and biomimetic drug delivery carriers, Nat. Rev. Drug Discov. 10 (2011) 521–535, https://doi.org/10.1038/nrd3499. [15] S. Baik, D.W. Kim, Y. Park, T.-J. Lee, S. Ho Bhang, C. Pang, A wet-tolerant adhesive patch inspired by protuberances in suction cups of octopi, Nature 546 (2017) 396–400, https://doi.org/10.1038/nature22382. [16] D.W. Green, B. Ben-Nissan, K.-S. Yoon, B. Milthorpe, H.-S. Jung, Bioinspired

5. Conclusions Despite the demonstrated therapeutic effects, spheroids are not widely used clinically because of the current lack of appropriate delivery methods. Therefore, we developed LoSH to culture a large number of spheroids and successfully deliver them in a uniform manner to the desired target area. The diameter of the spheroids formed on the LoSH surface was adjusted to 100–200 μm according to cell seeding density. The spheroids formed on the LoSH were weakly bound to the surface by conjugated fibronectin (100 μg/ml), and binding was maintained even when shear stress was applied, with spheroids stably attached to the LoSH. However, when the expansion of the hydrogel was induced at 4 °C, spheroids were separated from the surface due to cleavage in binding with fibronectin. Thus, we were able to place the LoSH upside-down on a glass substrate and induce expansion at 4 °C for 10 min. Upon removal of LoSH, we confirmed that the spheroids cultured on LoSH were uniformly transferred to the target with a high transfer efficiency of over 90%, and the high viability of the transferred spheroids was also maintained. We also showed the improved therapeutic efficacy of delivered spheroids in vivo in a mouse full thickness wound model. The spheroids delivered to the surface of the excisional wound through LoSH covered a large area in a homogenous manner. In addition, improved skin regeneration and angiogenesis were observed from spheroids delivered using LoSH. Our results demonstrate that LoSH is a novel platform to culture and deliver spheroids to a large target area and may have potential applications in various fields such as wound, ulcer, and burn treatments. Acknowledgements This research was supported by a National Research Foundation of Korea (NRF) grant funded by the Korean government (MEST) (Grant 12

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