Development of PyramidalMicrowells for Enhanced Cell Spheroid Formation in a Cell-on-Chip Microfluidic System for Cardiac Differentiation of Mouse Embryonic Stem Cells
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Citation: Wongpakham, T.; Chunfong, T.; Jeamsaksiri, W.; Chessadangkul, K.; Bhanpattanakul, S.; Kallayanathum, W.; Tharasanit, T.; Pimpin, A. Development of Pyramidal Microwells for Enhanced Cell Spheroid Formation in a Cell-on-Chip Microfluidic System for Cardiac Differentiation of Mouse Embryonic Stem Cells. Cells 2024,13, 2132. https://doi.org/10.3390/ cells13242132 Academic Editor: Ezequiel Álvarez Received: 18 November 2024 Revised: 13 December 2024 Accepted: 18 December 2024 Published: 23 December 2024 Copyright: © 2024 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (https:// creativecommons.org/licenses/by/ 4.0/). Article Development of Pyramidal Microwells for Enhanced Cell Spheroid Formation in a Cell-on-Chip Microfluidic System for Cardiac Differentiation of Mouse Embryonic Stem Cells Tepparit Wongpakham 1, Thanapat Chunfong 1, Wutthinan Jeamsaksiri 2, Kriengkai Chessadangkul 1, Sudchaya Bhanpattanakul 3,4, Wirakan Kallayanathum 5, Theerawat Tharasanit 4,5,* and Alongkorn Pimpin 1,6,* 1Department of Mechanical Engineering, Faculty of Engineering, Chulalongkorn University, Bangkok 10330, Thailand; [email protected] (T.W.); [email protected] (T.C.); [email protected] (K.C.) 2Thai Microelectronics Center, National Electronics and Computer Technology Center, Chachoengsao 24000, Thailand; [email protected].th 3 Department of Pathology, Faculty of Veterinary Science, Chulalongkorn University, Bangkok 10330, Thailand; [email protected] 4Center of Excellence for Veterinary Clinical Stem Cells and Bioengineering, Chulalongkorn University, Bangkok 10330, Thailand 5Department of Obstetrics, Gynaecology and Reproduction, Faculty of Veterinary Science, Chulalongkorn University, Bangkok 10330, Thailand; [email protected] 6Micro/Nano Electromechanical Integrated Device Research Unit, Faculty of Engineering, Chulalongkorn University, Bangkok 10330, Thailand *Correspondence: [email protected] (T.T.); [email protected] (A.P.) Abstract: Three-dimensional (3D) tissue culture models provide in vivo -like conditions for studying cell physiology. This study aimed to examine the efficiency of pyramidal microwell geometries in microfluidic devices on spheroid formation, cell growth, viability, and differentiation in mouse embryonic stem cells (mESCs). The static culture using the hanging drop (HD) method served as a control. The microfluidic chips were fabricated to have varying pyramidal tip angles, including 66 ◦ , 90 ◦ , and 106 ◦ . From flow simulations, when the tip angle increased, streamline distortion decreased, resulting in more uniform flow and a lower velocity gradient near the spheroids. These findings demonstrate the significant influence of microwell geometry on fluid dynamics. The 90 ◦ microwells provide optimal conditions, including uniform flow and reduced shear stress, while maintaining the ability for waste removal, resulting in superior spheroid growth compared to the HD method and other microwell designs. From the experiments, by Day 3, spheroids in the 90 ◦ microwells reached approximately 400 µ m in diameter which was significantly larger than those in the 66 ◦ microwells, 106 ◦ microwells, and HD cultures. Brachyury gene expression in the 90 ◦ microwells was four times higher than the HD method, indicating enhanced mesodermal differentiation essential for cardiac differentiation. Immunofluorescence staining confirmed cardiomyocyte differentiation. In conclusion, microwell geometry significantly influences 3D cell culture outcomes. The pyramidal microwells with a 90 ◦ tip angle proved most effective in promoting spheroid growth and cardiac differentiation of mESC differentiation, providing insights for optimizing microfluidic systems in tissue engineering and regenerative medicine. Keywords: cell spheroid; embryoid body; microfluidics; microwells; perfusion; pyramidal tip angles 1. Introduction Three-dimensional (3D) tissue culture has become an important bridge between in vitro and in vivo models in various biological research fields [ 1 , 2 ]. Three-dimensional structures provide in vivo -like conditions that are beneficial for studying cell physiology and conducting high-throughput drug discovery and screening [ 3 – 5 ]. In 3D clusters like Cells 2024,13, 2132. https://doi.org/10.3390/cells13242132 https://www.mdpi.com/journal/cells
Cells 2024,13, 2132 2 of 15 spheroids, cells communicate with each other and the extracellular matrix [ 6 , 7 ]. However, static culture systems, while simple and cost-effective, are limited in replicating key cell–environment interactions such as oxygen delivery, nutrient exchange, and waste removal [ 8 , 9 ]. To address these limitations, bioreactors with continuous fluid flow have been developed to improve 3D culture conditions. These bioreactors allow for precise control over fluid dynamics, gas exchange, and mechanical stimuli like shear force [ 10 , 11 ]. This approach has been successful in producing cardiomyocytes in both mouse and human models [ 12 , 13 ]. However, large-volume bioreactors can be costly, especially when significant amounts of chemicals or growth factors are required [14,15]. Microfluidic systems offer a more cost-effective alternative, using small volumes of fluid to control 3D tissue aggregation under precise conditions [ 16 , 17 ]. These systems have been applied to create “heart-on-chip” models for high-throughput drug testing [ 18 ]. Differentiation of mouse embryonic stem cells (mESCs) into cardiomyocytes has been a focal point of these studies [ 19 ]. Traditionally, the gold standard for cardiac differentiation is cell aggregation using the hanging drop (HD) method [ 20 – 23 ]. While effective, this method is labor-intensive and time-consuming [24]. To streamline cell aggregation, automated lab-on-chip technology using microfluidics has been developed. This technology controls small fluid volumes in channels, simulating physiological conditions on a microscale [ 25 , 26 ]. However, several factors influence the effectiveness of spheroid formation in microfluidic systems, including flow shear stress [ 27 ], cell type [ 28 ], and microwell shapes [ 29 ]. For efficient cell aggregation, the microwell platform integrated with microfluidic systems must ensure uniform spheroid size, reproducibility, and the ability to monitor the process [30,31]. The microwell shape significantly impacts cell culture outcomes. Pyramidal microwells have shown promise in reducing cell loss and guiding cells to the bottom, where spheroids are formed [ 32 – 34 ]. Inclined walls help direct cells into aggregation zones, minimizing the removal of unwanted cells during experiments [ 35 ]. Parameters such as the surface area-to-volume (SAV) ratio and tip angle affect how cells aggregate and how nutrients and oxygen are distributed. The pyramidal microwells with acute angles can create stronger flow disturbances, which may aid nutrient transfer but also increase mechanical stress on cells [ 36 ]. Designing effective microwell shapes requires balancing nutrient and oxygen transfer with controlled flow disturbances, such as recirculating flow velocity inside microwells [ 37 ]. Using arrays of pyramidal microwells increases surface area while decreasing SAV, creating better conditions for dynamic cell culture [38,39]. This study investigates how different pyramidal angles of microwells affect flow recirculation, cell spheroid formation, and viability in mESCs. The study focuses on flow characteristics, spheroid morphology, cell density, and viability during culture, aiming to optimize microwell design for improving 3D tissue engineering. 2. Materials and Methods All chemicals were obtained from Sigma Aldrich (St. Louis, MO, USA) unless stated otherwise. 2.1. Fabrication and Design of Microfluidic Device The microfluidic chips were fabricated using a standard soft-lithography process consisting of two parts: the microwell layer and the main chamber layer. The microwell layer was created using a 3D-printed plastic mold, while the main chamber layer was formed with a machined aluminum mold. Poly-dimethylsiloxane (PDMS) mixed with a curing agent at a 10:1 ratio was carefully poured into the molds to avoid air bubble formation. After the PDMS solidified, the microwell layer, main chamber layer, and tube connections were bonded together using oxygen plasma bonding. Figure 1shows the fabrication process of a PDMS microfluidic chip using 3D-printed plastic and aluminum molds. The microwells had varying pyramidal tip angles, including 66◦, 90◦, and 106◦, as shown in Figure 2. The SAV ratio for all cases ranged between 6.67 and 6.70. The microwells
Cells 2024,13, 2132 3 of 15 with 66 ◦ and 90 ◦ angles have a foreplan area of 4.22 mm 2 and 4.00 mm 2 , respectively, while the 106 ◦ microwell has a foreplan area of 3.69 mm 2 . This design enables the investigation of geometry effects on fluid dynamics and cell culture. Cells2025,14,xFORPEERREVIEW3of17 2.1.FabricationandDesignofMicrofluidicDevice Themicrofluidicchipswerefabricatedusingastandardsoft-lithographyprocess consistingoftwoparts:themicrowelllayerandthemainchamberlayer.Themicrowell layerwascreatedusinga3D-printedplasticmold,whilethemainchamberlayerwas formedwithamachinedaluminummold.Poly-dimethylsiloxane(PDMS)mixedwitha curingagentata10:1ratiowascarefullypouredintothemoldstoavoidairbubbleformation.AfterthePDMSsolidified,themicrowelllayer,mainchamberlayer,andtube connectionswerebondedtogetherusingoxygenplasmabonding.Figure1showsthefabricationprocessofaPDMSmicrofluidicchipusing3D-printedplasticandaluminum molds.Themicrowellshadvaryingpyramidaltipangles,including66°,90°,and106°,as showninFigure2.TheSAVratioforallcasesrangedbetween6.67and6.70.Themicrowellswith66°and90°angleshaveaforeplanareaof4.22mm 2 and4.00mm 2 ,respectively,whilethe106°microwellhasaforeplanareaof3.69mm 2 .Thisdesignenablesthe investigationofgeometryeffectsonfluiddynamicsandcellculture. Figure1.FabricationofthePDMSmicrofluidicchipusing3D-printedplasticandaluminummolds. Thefabricateddevice,measuring22mminlengthand12.5mminwidth,featuresanarrayofmicrowellswithvaryingpyramidaltipangles(66°,90°,and106°)and550µmindepth. Figure 1. Fabrication of the PDMS microfluidic chip using 3D-printed plastic and aluminum molds. The fabricated device, measuring 22 mm in length and 12.5 mm in width, features an array of microwells with varying pyramidal tip angles (66◦, 90◦, and 106◦) and 550 µm in depth. Cells2025,14,xFORPEERREVIEW4of17 Figure2.Schematicillustrationofthepyramidalmicrowellswithvaryingtipanglessuchas66°, 90°,and106°.Thedirectionofthemainflowisindicatedbyarrows. 2.2.FlowSimulationandFluidShearStress Acomputationalfluiddynamics(CFD)modelwasdevelopedusingCOMSOLMultiphysics5.3software(COMSOLInc.,Burlington,MA,USA)toanalyzevelocitydistribution,wallshearstress,andshearstressaroundthespheroidsinpyramidalmicrowells. Thesteady-stateNavier–Stokesandconvection–diffusionequationsweresolved,assuminganincompressible,isothermalNewtonianfluid(fluiddensity=998.2kg/m 3 ,viscosity =0.001Pa·s),withno-slipconditionsatthewalls.Themicrofluidicchannelandfourmicrowellsweresimulatedataflowrateof10µL/h(orapproximately1µm/sinletvelocity). Cellspheroidswithfixeddiametersof200µmwereplacedinsidethemicrowells,allowingforadetailedanalysisofshearstressdistribution.Thissetupwaschosentomimic realisticexperimentalconditionsandapproximatethemagnitudeofvelocityandshear stressthatwouldaffectcellviabilityandspheroidfunction. 2.3.CultureofMouseEmbryonicStemCells ThemESCsfromthe129S2/SvPasD3line(ATCC,WashingtonDC,USA)wereculturedonmitomycinC-treatedmouseembryonicfibroblasts(MEFs),whichservedasa feederlayertosupportstemcellmaintenance.ThemESCsweremaintainedinanEScell culturemediumconsistingofhigh-glucoseDulbecco’sModifiedEagleMedium(DMEM) supplementedwith10%ES-gradefetalbovineserum(FBS)(Gibco,Waltham,MA,USA), 1000IU/mLmouseleukemiainhibitoryfactor(LIF)(Millipore,Burlington,MA,USA),0.1 mMβ-mercaptoethanol(β-ME),0.1mMnon-essentialaminoacids(NEAAs),and1%antibiotic–antimycoticsolution(Anti-Anti). 2.4.SpheroidFormationandCardiacDifferentiation TwodifferentapproacheswereemployedforthespheroidformationofmESCssuch asHDandmicrowell(cell-on-chip)methods.ThemESCswereculturedtoapproximately 80%confluenceandthentrypsinizedwith0.25%trypsin–EDTAtoproduceasingle-cell suspension.Thecellswerecountedusingahemocytometeranddilutedtotheappropriate concentrationfordifferentiation.ThedifferentiationmediumconsistedofanESmedium with15%definedFBS(HyClone™,Omaha,NE,USA),excludingLIF.FortheHDmethod, 20µLdropscontaining800dissociatedmESCswereplacedonthelidofabacterialPetri dish,withphosphate-bufferedsaline(PBS)inthebasetopreventevaporation.Cellaggregationoccurredviagravity,leadingtothespontaneousformationofspheroids.The Figure 2. Schematic illustration of the pyramidal microwells with varying tip angles such as 66 ◦ , 90 ◦ , and 106◦. The direction of the main flow is indicated by arrows. 2.2. Flow Simulation and Fluid Shear Stress A computational fluid dynamics (CFD) model was developed using COMSOL Multiphysics 5.3 software (COMSOL Inc., Burlington, MA, USA) to analyze velocity distribution, wall shear stress, and shear stress around the spheroids in pyramidal microwells. The steady-state Navier–Stokes and convection–diffusion equations were solved, assuming an incompressible, isothermal Newtonian fluid (fluid density = 998.2 kg/m 3 , viscosity = 0.001 Pa·s ), with no-slip conditions at the walls. The microfluidic channel and four microwells were simulated at a flow rate of 10 µ L/h (or approximately 1 µ m/s inlet
Cells 2024,13, 2132 4 of 15 velocity). Cell spheroids with fixed diameters of 200 µ m were placed inside the microwells, allowing for a detailed analysis of shear stress distribution. This setup was chosen to mimic realistic experimental conditions and approximate the magnitude of velocity and shear stress that would affect cell viability and spheroid function. 2.3. Culture of Mouse Embryonic Stem Cells The mESCs from the 129S2/SvPas D3 line (ATCC, Washington DC, USA) were cultured on mitomycin C-treated mouse embryonic fibroblasts (MEFs), which served as a feeder layer to support stem cell maintenance. The mESCs were maintained in an ES cell culture medium consisting of high-glucose Dulbecco’s Modified Eagle Medium (DMEM) supplemented with 10% ES-grade fetal bovine serum (FBS) (Gibco, Waltham, MA, USA), 1000 IU/mL mouse leukemia inhibitory factor (LIF) (Millipore, Burlington, MA, USA), 0.1 mM β-mercaptoethanol ( β -ME), 0.1 mM non-essential amino acids (NEAAs), and 1% antibiotic–antimycotic solution (Anti-Anti). 2.4. Spheroid Formation and Cardiac Differentiation Two different approaches were employed for the spheroid formation of mESCs such as HD and microwell (cell-on-chip) methods. The mESCs were cultured to approximately 80% confluence and then trypsinized with 0.25% trypsin–EDTA to produce a single-cell suspension. The cells were counted using a hemocytometer and diluted to the appropriate concentration for differentiation. The differentiation medium consisted of an ES medium with 15% defined FBS (HyClone™, Omaha, NE, USA), excluding LIF. For the HD method, 20 µ L drops containing 800 dissociated mESCs were placed on the lid of a bacterial Petri dish, with phosphate-buffered saline (PBS) in the base to prevent evaporation. Cell aggregation occurred via gravity, leading to the spontaneous formation of spheroids. The formation of spheroids was observed and monitored using a phase contrast microscope (Olympus CKX41, Tokyo, Japan). For cardiac differentiation using the microwells in microfluidic devices, pyramidal microwells were filled with a culture medium by injecting it slowly through the microchannel inlet using a micropipette. A cell suspension of 95 µ L, containing 1 × 10 5 mESCs, was introduced, resulting in an approximated cell density of 800 cells per well. The cells gradually settled into the microwells via gravity. The microfluidic system was then connected to a silicone tube and syringe pump operating in withdrawal mode, as shown in Figure 3, with a fresh culture medium supplied at a continuous flow rate of 10 µ L/h for three days. A fresh culture medium was introduced through the inlet, and fluid was withdrawn through the outlet. All differentiation experiments were conducted in a humidified incubator at 37 ◦C with 5% CO2. Cells2025,14,xFORPEERREVIEW5of17 formationofspheroidswasobservedandmonitoredusingaphasecontrastmicroscope (OlympusCKX41,Tokyo,Japan). Forcardiacdifferentiationusingthemicrowellsinmicrofluidicdevices,pyramidal microwellswerefilledwithaculturemediumbyinjectingitslowlythroughthemicrochannelinletusingamicropipette.Acellsuspensionof95µL,containing1×10 5 mESCs, wasintroduced,resultinginanapproximatedcelldensityof800cellsperwell.Thecells graduallysettledintothemicrowellsviagravity.Themicrofluidicsystemwasthenconnectedtoasiliconetubeandsyringepumpoperatinginwithdrawalmode,asshownin Figure3,withafreshculturemediumsuppliedatacontinuousflowrateof10µL/hfor threedays.Afreshculturemediumwasintroducedthroughtheinlet,andfluidwaswithdrawnthroughtheoutlet.Alldifferentiationexperimentswereconductedinahumidified incubatorat37°Cwith5%CO 2 . Figure3.Schematicoftheexperimentalsetupforspheroidformationusingthemicrofluidicsystem installedinsideaCO 2 incubator. 2.5.SpheroidMorphometryandViabilityAnalysis Toassessspheroidhealthandviabilityundervaryingflowconditionsthroughmorphometricanalysisandlive/deadcellviabilityassays.Spheroidswereimagedusinga brightfieldmicroscope(OlympusCKX41,Tokyo,Japan)tocapturetheirmorphology. SpheroiddiametersandcircularityweremeasuredusingImageJsoftware(ImageJ1.x), ensuringprecisequantificationoftheirstructuralproperties. Fortheviabilityassessment,spheroidswereculturedforthreedaysandthenharvestedfromthemicrowells.Thespheroidsweredissociatedintosinglecellsusingtrypsin–EDTAtreatment,followedbystainingwithCalceinAcetoxymethyl(CalceinAM)and EthidiumHomodimer-1(EthD-1)forlive/deadanalysis.Fluorescentmicroscopy(OlympusBX51,Tokyo,Japan)wasusedtovisualizethecells,withviablecellsshowinggreen fluorescenceinthecytoplasmduetotheretentionofCalceinAM,whiledeadcellsexhibitedredfluorescenceinthenucleusduetoEthD-1stainingofcompromisedmembranes. Fluorescentimageswerecapturedandanalyzedtoquantifytheviabilityofcellswithin eachspheroid. 2.6.ProliferativeActivityEvaluation Cellproliferationwasassessedusingthe3-(4,5-Dimethylthiazol-2-yl)-2,5-DiphenyltetrazoliumBromideassay(MTT)assay.Harvestedspheroidsweretransferredtoa96wellplate,withonespheroidperwell.Eachwellwasfilledwith200µLofMTTsolution (0.5mg/mLinPBS).Thespheroidswereincubatedinthedarkfor30minat37°Cand5% Figure 3. Schematic of the experimental setup for spheroid formation using the microfluidic system installed inside a CO2incubator.
Cells 2024,13, 2132 5 of 15 2.5. Spheroid Morphometry and Viability Analysis To assess spheroid health and viability under varying flow conditions through morphometric analysis and live/dead cell viability assays. Spheroids were imaged using a brightfield microscope (Olympus CKX41, Tokyo, Japan) to capture their morphology. Spheroid diameters and circularity were measured using ImageJ software (ImageJ 1.x), ensuring precise quantification of their structural properties. For the viability assessment, spheroids were cultured for three days and then harvested from the microwells. The spheroids were dissociated into single cells using trypsin–EDTA treatment, followed by staining with Calcein Acetoxymethyl (Calcein AM) and Ethidium Homodimer-1 (EthD-1) for live/dead analysis. Fluorescent microscopy (Olympus BX51, Tokyo, Japan) was used to visualize the cells, with viable cells showing green fluorescence in the cytoplasm due to the retention of Calcein AM, while dead cells exhibited red fluorescence in the nucleus due to EthD-1 staining of compromised membranes. Fluorescent images were captured and analyzed to quantify the viability of cells within each spheroid. 2.6. Proliferative Activity Evaluation Cell proliferation was assessed using the 3-(4,5-Dimethylthiazol-2-yl)-2,5-Diphenyltetrazolium Bromide assay (MTT) assay. Harvested spheroids were transferred to a 96-well plate, with one spheroid per well. Each well was filled with 200 µ L of MTT solution (0.5 mg/mL in PBS). The spheroids were incubated in the dark for 30 min at 37 ◦ C and 5% CO 2 . After incubation, the MTT solution was removed, and the formazan crystals formed in the cells were dissolved with 200 µ L of DMSO per well. Optical density (OD) was measured at 570 nm, with a reference wavelength of 650 nm, using a microplate reader. The resulting absorbance values were compared to a standard curve to indirectly quantify the cell number, serving as a measure of proliferative activity within each spheroid. 2.7. Gene Expression Assessment The expression of the Brachyury gene in spheroids was quantified using a quantitative real-time polymerase chain reaction (qRT-PCR). Total RNA was extracted from EBs using the RNeasy Mini Kit (Qiagen, Venlo, The Netherlands), following the manufacturer’s instructions. The extracted RNA was reverse-transcribed into complementary DNA (cDNA) using SuperScript III Reverse Transcriptase (Invitrogen, Waltham, MA, USA). qRT-PCR was performed using GoTaq ® Green Master Mix (M7122, Madison, Promega, WI, USA) on a CFX96 Connect™ Real-Time PCR System (Bio-Rad, Hercules, CA, USA). The specific primers used for Brachyury were as follows: Forward 5 ′ - TACACCTCTAATGTCCTCCCTTG-3 ′ and Reverse 5 ′ -CCATACAGTTGACTTCCCAACAC3 ′ . The housekeeping gene GAPDH was used as an internal control, with the following primers: Forward 5 ′ -AATGTGTCCGTCGTGGATCT-3 ′ and Reverse 5 ′ -CCTGCTTCACCACCTTCTTG-3 ′ . Relative gene expression was calculated using the ∆∆ Ct method to normalize Brachyury expression levels against GAPDH. 2.8. Assessment of Cardiac Differentiation Efficiency To evaluate cardiac differentiation efficiency, the percentage of beating spheroids was determined. Spheroids harvested from both HD cultures and 90 ◦ microwells (the optimal condition identified in previous experiments) were cultured in a cardiac differentiation medium for fourteen days. The number of beating-cell colonies was counted. The differentiation medium consisted of high-glucose DMEM (Cat. No. SH30022.22, HyClone™, Omaha, NE, USA) supplemented with 15% defined fetal bovine serum (Cat. No. SH30070.03, HyClone™, Omaha, NE, USA), 1% antibiotic–antimycotic, 1% L-glutamine, 1% MEM non-essential amino acids (NEAAs, Cat. No. M7145, Sigma Aldrich, St. Louis, MO, USA), 10 ng/mL Bone Morphogenetic Protein 4 (Recombinant Human BMP-4, Cat. No. 4578, BioVision, Exton, PA, USA), 50 ng/mL Ascorbic Acid (Cat. No. A4544, Sigma Aldrich, St. Louis, MO, USA), and 25.9 mM beta-mercaptoethanol (Cat. No. 21985-023, Gibco, NY, USA).
Cells 2024,13, 2132 6 of 15 2.9. Immunofluorescence Staining Test Mouse cardiomyocytes were fixed in 4% (v/v) paraformaldehyde for 15 min at room temperature, followed by three washes with phosphate-buffered saline (PBS). Permeabilization was performed using 0.1% Triton-X 100 in PBS containing 2% bovine serum albumin (BSA) to block nonspecific binding sites. The cells were then incubated overnight at 4 ◦ C with mouse anti-cardiac Troponin T (cTnT) primary antibodies (1:100; ab33589, Abcam, Cambridge, UK) to specifically label cardiomyocytes. After primary antibody incubation, the cells were washed and incubated with TRITC-conjugated secondary antibodies (1:100; T5393, Sigma Aldrich, St. Louis, MO, USA) for 1 h at 37 ◦ C. The 4 ′ ,6-diamidino2-phenylindole (DAPI) staining was applied for 15 min at room temperature to label the nuclei. The samples were then washed and imaged using a fluorescence microscope (Olympus BX51, Tokyo, Japan), capturing both cardiomyocyte-specific and nuclear staining. 2.10. Statistical Analysis Statistical analysis was performed using SPSS version 29. A normality test was conducted with a 95% confidence level, followed by one-way analysis of variance (ANOVA) and post hoc Bonferroni tests, to determine significant differences between groups. Data are represented as mean ± standard deviation, with differences considered significant at p< 0.05. 3. Results 3.1. Pyramidal Microwells The roughness of the microwell surfaces was measured using a Surface Roughness Tester (Mitutoyo SV-3000, Kanagawa, Japan). The measured roughness was 0.395 ±0.078 µm . The inclined surfaces of the microwells exhibited a stepwise structure with an average width of 90.45 ± 4.56 µ m and a height of 45.58 ± 1.56 µ m. The diameters of the cylindrical openings at the center were 446.96 ± 4.03 µ m, 449.43 ± 5.22 µ m, and 453.15 ±2.80 µm for the 66 ◦ , 90 ◦ , and 106 ◦ microwell configurations, respectively. The microwell arrays consisted of 45 wells for the 66 ◦ tip angle, 42 wells for the 90 ◦ tip angle, and 52 wells for the 106 ◦ tip angle. These geometric and structural variations were critical for analyzing the effects of microwell shape on fluid dynamics and cell behavior within the microfluidic system. 3.2. Velocity Distribution and Fluid Shear Stress Figure 4illustrates the velocity distribution above cell spheroids in microwells with tip angles of 66 ◦ , 90 ◦ , and 106 ◦ . As the tip angle increased, streamline distortion decreased, resulting in more uniform flow and lower velocities near the spheroids. The 106 ◦ microwell exhibited the most uniform flow, indicating a stable environment that could reduce shear stress on cells. The velocity at the center of the 66 ◦ microwell was approximately double that of the 106 ◦ microwell, highlighting the concentrated flow in narrower tip angles. This similar flow pattern was observed at higher flow rates with velocity magnitudes increasing proportionally with higher inlet velocities. Near the spheroids, the velocity was around 1×10−7m/s at a flow rate of 10 µ L/h. These findings demonstrate the significant influence of microwell geometry on fluid dynamics, with wider angles probably promoting more favorable conditions for cell culture by lowering flow disturbance and minimizing fluid shear stress. The flow streamlines through each microwell are depicted in Figure 5. Red streamlines represent flow near the cell spheroids at the microwell bottom, while blue streamlines indicate flows that do not reach the bottom region. In all configurations, only the central flow passes directly above the spheroids. The 106 ◦ microwell shows more uniform flow near the spheroids, whereas narrower angles, like the 66 ◦ microwell, display more confined and distorted flow patterns. These findings emphasize the role of microwell geometry in shaping flow behavior, with wider tip angles facilitating straight and uniformly distributed flows, potentially reducing shear stress on the cells. On the other hand, the ability for waste removal and nutrient transfer might be weakened.
Cells 2024,13, 2132 7 of 15 Cells2025,14,xFORPEERREVIEW8of17 Figure4.Velocitydistributioninmicrowellswithpyramidaltipanglesof66°,90°,and106°:(a)side viewofvelocitycontourdisplayingflowdynamicsaroundembeddedcellspheroidsand(b–d)top viewsofvelocitycontoursfor66°,90°,and106°microwellsataflowrateof10µL/h. TheflowstreamlinesthrougheachmicrowellaredepictedinFigure5.Redstreamlinesrepresentflownearthecellspheroidsatthemicrowellbottom,whilebluestreamlinesindicateflowsthatdonotreachthebottomregion.Inallconfigurations,onlythe centralflowpassesdirectlyabovethespheroids.The106°microwellshowsmoreuniform flownearthespheroids,whereasnarrowerangles,likethe66°microwell,displaymore confinedanddistortedflowpatterns.Thesefindingsemphasizetheroleofmicrowellgeometryinshapingflowbehavior,withwidertipanglesfacilitatingstraightanduniformly distributedflows,potentiallyreducingshearstressonthecells.Ontheotherhand,the abilityforwasteremovalandnutrienttransfermightbeweakened. Figure5.Distributionofstreamlinesinmicrowellswithtipanglesof(a)66°,(b)90°,and(c)106°. Theflowdirectionisindicatedbythearrows. ThedistributionofshearstressonthesurfaceofthecellspheroidisshowninFigure 6.Thisfindingrevealsagradualincreaseinshearstressinboththestreamwiseand Figure 4. Velocity distribution in microwells with pyramidal tip angles of 66 ◦ , 90 ◦ , and 106 ◦ : (a) side view of velocity contour displaying flow dynamics around embedded cell spheroids and (b–d) top views of velocity contours for 66◦, 90◦, and 106◦microwells at a flow rate of 10 µL/h. Cells2025,14,xFORPEERREVIEW8of17 Figure4.Velocitydistributioninmicrowellswithpyramidaltipanglesof66°,90°,and106°:(a)side viewofvelocitycontourdisplayingflowdynamicsaroundembeddedcellspheroidsand(b–d)top viewsofvelocitycontoursfor66°,90°,and106°microwellsataflowrateof10µL/h. TheflowstreamlinesthrougheachmicrowellaredepictedinFigure5.Redstreamlinesrepresentflownearthecellspheroidsatthemicrowellbottom,whilebluestreamlinesindicateflowsthatdonotreachthebottomregion.Inallconfigurations,onlythe centralflowpassesdirectlyabovethespheroids.The106°microwellshowsmoreuniform flownearthespheroids,whereasnarrowerangles,likethe66°microwell,displaymore confinedanddistortedflowpatterns.Thesefindingsemphasizetheroleofmicrowellgeometryinshapingflowbehavior,withwidertipanglesfacilitatingstraightanduniformly distributedflows,potentiallyreducingshearstressonthecells.Ontheotherhand,the abilityforwasteremovalandnutrienttransfermightbeweakened. Figure5.Distributionofstreamlinesinmicrowellswithtipanglesof(a)66°,(b)90°,and(c)106°. Theflowdirectionisindicatedbythearrows. ThedistributionofshearstressonthesurfaceofthecellspheroidisshowninFigure 6.Thisfindingrevealsagradualincreaseinshearstressinboththestreamwiseand Figure 5. Distribution of streamlines in microwells with tip angles of (a) 66 ◦ , (b) 90 ◦ , and (c) 106 ◦ . The flow direction is indicated by the arrows. The distribution of shear stress on the surface of the cell spheroid is shown in Figure 6. This finding reveals a gradual increase in shear stress in both the streamwise and spanwise directions as the angle around the spheroid increases. The maximum shear stress occurs at the top of the spheroid (at 90 ◦ ), then gradually decreases again. Narrower microwell angles (e.g., 66 ◦ ) generate higher shear stress compared to wider angles (e.g., 106 ◦ ), where the shear stress is more evenly distributed. These results show the influence of microwell
Cells 2024,13, 2132 8 of 15 geometry affecting the distribution of shear stress, which is crucial for maintaining the viability and integrity of cell spheroids. Cells2025,14,xFORPEERREVIEW9of17 spanwisedirectionsastheanglearoundthespheroidincreases.Themaximumshear stressoccursatthetopofthespheroid(at90°),thengraduallydecreasesagain.Narrower microwellangles(e.g.,66°)generatehighershearstresscomparedtowiderangles(e.g., 106°),wheretheshearstressismoreevenlydistributed.Theseresultsshowtheinfluence ofmicrowellgeometryaffectingthedistributionofshearstress,whichiscrucialformaintainingtheviabilityandintegrityofcellspheroids. Figure6.Distributionofshearstressonthesurfaceofcellspheroidformicrowellswithtipangles of66°,90°,and106°:(a)shearstressalongthespanwisedirectionand(b)shearstressalongthe streamwisedirection. 3.3.CellGrowthandMorphologicalProgressionofmESC-DerivedSpheroids Aftergentlyseedingcellsuspensions,wesuppliedthemediumataconstantflow ratetofacilitateevendepositionofcellsintothemicrowells.Afterthat,thecellswereallowedtorestfor120mintoenableself-aggregation.Subsequently,weharvestedthecells fromthemicrowellsusingamicropipetteandassessedthecellcountusingtheMTTassay. ThecellnumberswerequantifiedbasedonastandardcurvegeneratedfromtheMTT assay.Wehavecheckedthisprocedurewithothercellswithimagingtechniquesaswell beforeconductingthisresearch,anditwasfoundthatthenumberofcellsineachmicrowellisquiteuniform.Thevariationis5–10%ofthemeanvalue. Inourexperiments,theinitialnumberofmESCsseededonDay0inthe66°,90°,and 106°microwellswas831±89,876±103,and876±103cellsperwell,respectively.Over thethree-dayperiod,allmicrowellconfigurationsshowedanincreaseincellgrowthand viability,withthespheroidsinthe90°microwellsconsistentlyexhibitinglargergrowth comparedtothe66°and106°microwells,aswellasthestaticHDculture.Asshownin Figure7,themorphologyofthemESC-derivedspheroidsvariedwithmicrowellgeometry.OnDay1,spheroidsacrossallmicrowellangleswererelativelyuniforminsize.By Day2,spheroidsinthe90°microwellsbegantoshowmorerapidexpansion,butstillbeing comparabletothoseinthe66°and106°microwells.ByDay3,spheroidsinthe90°microwellsexhibitedthelargestgrowth,reachingdiametersofaround400µm,whichwas significantlylargerthanthoseformedintheothermicrowellconfigurations,asdepicted inFigure8. Itisquitecommontoobserveadarkenedcenterincellaggregates,astheinnermost cellsbecomepackedintoadensemass.Thishighcelldensitycanlimitoxygenandnutrientdiffusionintothecoreoftheaggregate.Additionally,ascellsaggregateintoa3Dstructure,thecenterbecomesmorecompact,increasingcellconcentration.Thisdenserregion Figure 6. Distribution of shear stress on the surface of cell spheroid for microwells with tip angles of 66 ◦ , 90 ◦ , and 106 ◦ : (a) shear stress along the spanwise direction and (b) shear stress along the streamwise direction. 3.3. Cell Growth and Morphological Progression of mESC-Derived Spheroids After gently seeding cell suspensions, we supplied the medium at a constant flow rate to facilitate even deposition of cells into the microwells. After that, the cells were allowed to rest for 120 min to enable self-aggregation. Subsequently, we harvested the cells from the microwells using a micropipette and assessed the cell count using the MTT assay. The cell numbers were quantified based on a standard curve generated from the MTT assay. We have checked this procedure with other cells with imaging techniques as well before conducting this research, and it was found that the number of cells in each microwell is quite uniform. The variation is 5–10% of the mean value. In our experiments, the initial number of mESCs seeded on Day 0 in the 66 ◦ , 90 ◦ , and 106 ◦ microwells was 831 ± 89, 876 ± 103, and 876 ± 103 cells per well, respectively. Over the three-day period, all microwell configurations showed an increase in cell growth and viability, with the spheroids in the 90 ◦ microwells consistently exhibiting larger growth compared to the 66 ◦ and 106 ◦ microwells, as well as the static HD culture. As shown in Figure 7, the morphology of the mESC-derived spheroids varied with microwell geometry. On Day 1, spheroids across all microwell angles were relatively uniform in size. By Day 2, spheroids in the 90 ◦ microwells began to show more rapid expansion, but still being comparable to those in the 66 ◦ and 106 ◦ microwells. By Day 3, spheroids in the 90 ◦ microwells exhibited the largest growth, reaching diameters of around 400 µ m, which was significantly larger than those formed in the other microwell configurations, as depicted in Figure 8. It is quite common to observe a darkened center in cell aggregates, as the innermost cells become packed into a dense mass. This high cell density can limit oxygen and nutrient diffusion into the core of the aggregate. Additionally, as cells aggregate into a 3D structure, the center becomes more compact, increasing cell concentration. This denser region may scatter light differently, contributing to the darker appearance at the spheroid’s center. The darkened center of cell aggregates can influence cardiac differentiation, reflecting processes such as hypoxia, metabolic changes, and apoptosis, which are characteristic of the early stages of cardiac cell development.
Cells 2024,13, 2132 9 of 15 Cells2025,14,xFORPEERREVIEW10of17 mayscatterlightdifferently,contributingtothedarkerappearanceatthespheroid’scenter.Thedarkenedcenterofcellaggregatescaninfluencecardiacdifferentiation,reflecting processessuchashypoxia,metabolicchanges,andapoptosis,whicharecharacteristicof theearlystagesofcardiaccelldevelopment. Theseresultssuggestthatthe90°microwellsprovideoptimalconditionsforspheroid formation.Additionally,nearly100%cellviabilitywasobservedacrossallmicrowellconfigurations,indicatingsuccessfulcultureandgrowthofthespheroids.Thistrendreflects thecorrelationbetweenthemicrowellangleandcellgrowth,withthe90°tipangleprovidingoptimalconditionsforspheroidexpansion. Figure7.MorphologicalprogressionofmESC-derivedspheroidsculturedinpyramidalmicrowells withtipanglesof66°,90°,and106°,comparedtostaticcultureinHD.Imageswerecapturedon Day1,Day2,andDay3.Scalebarsrepresent50µmfortheHDimagesand100µmforthemicrowell images. Figure 7. Morphological progression of mESC-derived spheroids cultured in pyramidal microwells with tip angles of 66 ◦ , 90 ◦ , and 106 ◦ , compared to static culture in HD. Images were captured on Day 1, Day 2, and Day 3. Scale bars represent 50 µ m for the HD images and 100 µ m for the microwell images. Cells2025,14,xFORPEERREVIEW11of17 Figure8.Spheroidsizemeasuredoverthreedaysacrossdifferentculturemethods(n=10per group).Statisticalsignificancebetweengroupsisindicatedbydifferentletters(a–d),withp<0.05. 3.4.ProliferativeActivity TheMTTassaywasusedtoquantifycellproliferationineachspheroid.OnDay1, allmicrowellconfigurationsshowedsimilarcellnumbers,withnosignificantdifferences. ByDay2,allmicrowellsdidnotshowsignificantcellgrowthcomparedtotheHDmethod, whichhadfasterproliferationrates.However,byDay3,cellnumbersincreasedto1500– 2000perwellacrossallmicrowells,withthe90°configurationshowingthemostsignificantgrowthasshowninFigure9.Despitetheseresults,theoverallgrowthratesforthree daysinallmicrowellswereslowerthanthoseintheHDmethod. Compactnesswasfurtherestimatedastheratioofthenumberofcellstothecubeof thespheroiddiameter.OnDay2,compactnesswasmeasuredat0.186±0.038fortheHD, 0.056±0.011forthe66°microwells,0.058±0.012forthe90°microwells,and0.048±0.010 (×10 −3 µm −3 )forthe106°microwells,withanuncertaintyofapproximately20%forall cases.TheseresultsindicatethatconventionaltechniquesastheHDmethodresultedin densercellspheroids.Additionally,thespheroidsinthe106°microwellswererelatively loose,whilethe66°and90°microwellsproducedspheroidswithcomparablecompactness.ByDay3,allspheroidscontinuedgrowing,withcompactnessdecreasingto55–60% ofDay2levels. Figure 8. Spheroid size measured over three days across different culture methods (n= 10 per group). Statistical significance between groups is indicated by different letters (a–d), with p< 0.05.