Generation, High-Throughput Screening, and Biobanking of Human-Induced Pluripotent Stem Cell-Derived Cardiac Spheroids

NOTE: hiPSC-CMs used in this study were generated according to previously described hiPSC culturing and CM differentiation protocols^26,27. Optionally, the hiPSC-CMs can be expanded and cryopreserved as recently published before starting the CS protocol (section 4)¹⁰.

1. Preparation of cell culture media, solutions, and aliquots

1. Prepare basal medium
   1. Equilibrate penicillin-streptomycin and the medium (RPMI 1640) to room temperature (RT) and ensure it has thawed completely. Mix 500 mL of the medium and 5 mL of pen/strep. Store at 4 °C for up to 8 weeks; equilibrate to 37 °C before use.
2. Prepare RPMI + B27
   1. Equilibrate the B27 supplement and the basal medium to RT. Ensure to thaw the supplement completely. Mix 490 mL of the basal medium and 10 mL of the 50x B27 supplement. Store at 4 °C for up to 2 weeks; equilibrate to 37 °C before use.
3. Prepare hiPSC-CM re-plating media
   1. Add Rho-associated, coiled-coil containing protein kinase (ROCK) inhibitor (2 µM final concentration) and 10% knockout serum replacement (KSR) to RPMI + B27 media. Add ROCK inhibitor directly to the RM media as needed. Do not store culture media once supplemented.
4. Prepare CM thawing media
   1. Add a 1:100 concentration of cell survival supplement (e.g., Revitacell) and 20% KSR to RPMI + B27 media and equilibrate to 37 °C before use.
5. Prepare maturation supplement
   1. The previously described maturation medium formula²⁸ consists of: 3 mM glucose, 10 mM L-lactate, 5 mg/mL vitamin B12, 0.82 mM biotin, 5 mM creatine monohydrate, 2 mM taurine, 2 mM L-carnitine, 0.5 mM ascorbic acid, 1x NEAA, 0.5% (w/v) albumax, 1x B27, and 1% KOSR. To prepare one full bottle (500 mL) of maturation supplement, remove 65 mL from a bottle of DMEM without glucose and supplement with 2.7 g of glucose, 5.6 g of L-lactate, 0.025 mg of vitamin B12, 1 mg of Biotin, 3.73 g of creatine monohydrate, 1.25 g of taurine, 1.975 g of L-carnitine, 0.7125 g of ascorbic acid, 50 mL of NEAA, 12.5 g of albumax, and 5 mL of penicillin-streptomycin.
   2. Filter through a sterile disposable filter unit with a 0.22 µm pore Polyethersulfone (PES) membrane.
   3. Aliquot into 45 mL (to prepare 500 mL of maturation medium) or 4.5 mL (to prepare 50 mL of maturation medium). Store at 20 °C for up to 6 months.
6. Prepare maturation media
   1. Equilibrate the B27 supplement, knockout SR, penicillin-streptomycin, the maturation supplement²⁸, and the DMEM no-glucose medium at RT. Ensure to thaw the supplement completely. Mix 435 mL of the DMEM no glucose medium with 10 mL of the 50x B27 supplement, 5 mL of penicillin-streptomycin, 5 mL of knockout SR, and 45 mL of maturation supplement. Store at 4 °C for up to 2 weeks; equilibrate at 37 °C before use.
7. Prepare fluor bright medium
   1. Equilibrate penicillin-streptomycin and DMEM fluorobrite medium at RT. Ensure that the supplement is completely thawed. Mix 500 mL of the DMEM fluorbrite medium with 5 mL of penicillin-streptomycin. Store at 4 °C for up to 1 month; equilibrate at 37 °C before use.
8. Prepare the non-ionic detergent solution
   1. Mix 20% w/v non-ionic detergent powder (e.g., F-127) with PBS. Filter using a 0.22 µm filter and store at 4 °C for up to 6 months; equilibrate at RT before use.
9. Prepare calcium dye medium
   1. Mix the non-ionic detergent solution (final concentration of 0.04% v/v) and 0.1x of the calcium dye (e.g., Cal520 AM) in fluor bright medium. In a 50 mL conical tube, add 10 µL of Cal520 and 20 µL of the non-ionic detergent solution. Mix until fully dissolved. Keep the solution in the dark before adding to the cells.

2. Preparation of buffers

1. Prepare permeabilization and blocking buffer: This buffer contains 10 mL of PBS, 5% wt/v BSA, and 0.3% v/v Triton-X-100.
2. Prepare the flow cytometry buffer: This buffer contains 50 mL of PBS, 1% wt/v BSA, and 0.3% v/v Triton-X-100.
3. Flow cytometry washing buffer: This buffer contains 50 mL of PBS and 1% wt/v BSA.
4. Spheroid washing buffer (SWB): This buffer contains 1 mL of Triton-X-100, 2 mL of 10% (w/v in DPBS) SDS, and 2 g of BSA in 1 L of PBS.
   NOTE: SWB can be stored at 4 °C for up to 2 weeks.
5. Prepare the embedding solution (ES): To prepare 100 mL of the embedding solution, mix 50 mL of glycerol with 9.09 mL of dH₂O, 1 mL of Tris buffer (1 M, pH 8.0), and 200 µL of EDTA (0.5 M). Add 22.7 g of fructose and mix at RT in the dark until dissolved. When clear, add 22.2 g of fructose and mix until dissolved. Then add 15 g of urea and mix until dissolved (store at 4 °C in the dark).
6. Prepare PBT (PBS with Tween-20) buffer. This buffer contains PBS/Tween-20 (0.1% v/v). For 1 L of PBS, add 1 mL of Tween-20.

3. Preparation of small molecules

1. Reconstitute thiazovivin (ROCK inhibitor) powder in 10 mM aliquots of 50 µL in DMSO and store at -20 °C for up to 6 months. Protect from light.
2. Prepare 2.5 mM aliquots of 10 µL each of Cal-520 AM in DMSO and store them at -20 °C for up to 6 months. Protect from light.

4. Cardiac spheroid generation

NOTE: For larger amounts of CSs, seed up to 1 million CMs in a 6-well ultra-low attachment plate with 2 mL of hiPSC-CM re-plating media. This study used a minimum of 2,500 (2.5k CS) up to 20,000 (20k CS) hiPSC-CMs per well of a 96-well plate.

1. For one 96-well plate, prepare a cell culture containing a minimum of 2 x 10⁶ human-induced pluripotent stem cell-derived cardiomyocytes (hiPSC-CMs)¹⁰.
2. When the cultured hiPSC-CMs reach confluence, add 0.1 mL/cm² of sterile cardiac detachment solution (e.g., tryple) to each well. Incubate the plate at 37 °C for 15 min.
3. Using a 5 mL pipet, mechanically dissociate the cells by flushing with 2 mL of warm basal medium to make a single cell suspension. Confirm the detachment with a bright field microscope (4x magnification); cells will look white and have a round shape.
4. Transfer the cell suspension to a 15 mL conical tube and centrifuge for 3 min at 300 x g.
5. Aspirate the supernatant and resuspend the cells in 1 mL of hiPSC-CM re-plating media.
6. Using a 1,000 µL pipet tip, mechanically dissociate the cell pellet. The solution appears homogeneous after three or four mixes. Count the cells. Transfer the appropriate amount of cells in 100 µL of the re-plating medium to each ultra-low attachment round-bottom 96-well well.
7. Place the plate of CSs on an orbital shaker at 70 rpm in the incubator for 24 h. Set the incubator conditions to 37 °C, 5% CO₂, 21% O₂, and 90% humidity.
8. Aspirate 50 µL of medium from each well and add 100 µL of RPMI + B27 medium per well for the first 48 h.
   NOTE: Always keep 50 µL of the medium in the 96-well plate to avoid accidental aspiration and spheroid rupture.
9. Aspirate 100 µL of the medium from each well and add 100 µL of the maturation medium per well. Maintain the cells in the maturation medium and refresh the medium every 2-3 days.

5. Cryopreservation of CSs

NOTE: CSs can be cryopreserved for long-term storage. Cryopreservation can be performed from day 3 after the generation of CSs. CSs can be cryopreserved directly in the wells of a 96-well plate or as a CS suspension in cryovials.

1. Pre-cool the plate by placing the plate on ice for 10 min.
2. Centrifuge the spheroid plate for 3 min at 70 x g.
3. Remove the supernatant till 50 µL remains and add 200 µL of ice-cold hiPSC freezing medium per well.
   NOTE: Keep the CS suspension on ice for the entire duration of the procedure. In the case of a 6-well plate with spheroids, freeze one well in a 500 µL freezing medium cryovial.
4. Seal the plate with plate sealing film.
   NOTE: The 96-well plate needs to be stored in a polystyrene box or, when not available, a silicon mold can be made as described in step 5.5.1.
5. To ensure uniform heat exchange between the well plate and the freezer, place the plate carefully into a polystyrene box or in a silicon mold.
   1. To prepare the silicon mold: Vigorously mix two components of the silicon elastomer kit in a 10:1 ratio. De-bubble the solution using a vacuum pump for 15-20 min. Subsequently, cast the solution inside the bottom part of the well plate and de-bubble using a vacuum pump for 10 min. Place the mold into an oven and cure at 60 °C for 8 h to obtain a semi-flexible elastomer that is peeled off the plate.
6. Freeze the plate at -80 °C for a minimum of 4 h in the polystyrene box or the prepared silicon mold.
7. Transfer the plate to a liquid nitrogen tank or a -150 °C freezer for long-term storage.

6. Thawing of cardiac spheroids

NOTE: Do not thaw more than one plate at a time to ensure a quick thawing process.

1. Prepare 20 mL of 37 °C preheated basal medium in a 50 mL conical tube.
2. Collect the cell plate with CSs from the liquid nitrogen and place it in the incubator for 15 min. Set the incubator conditions to 37 °C, 5% CO₂, 21% O₂, and 90% humidity.
3. Remove the supernatant and the cell pellet remains, and resuspend each well in a warm basal medium. Use 200 µL of medium per well.
4. Centrifuge for 3 min at 70 x g.
5. Repeat steps 6.3 and 6.4.
6. Remove the supernatant until the cell pellet remains and add 200 µL of CM thawing medium in each well.
7. Place the plate of CSs on an orbital shaker at 70 rpm in an incubator for 24 h. Set the incubator conditions to 37 °C, 5% CO₂, 21% O₂, and 90% humidity.
8. Aspirate 50 µL of the medium from each well and add 100 µL of RPMI + B27 medium per well for the first 48 h.
9. Aspirate 100 µL of the medium from each well and add 100 µL of maturation medium per well. Maintain the cells in the maturation media and refresh the medium every 2-3 days.

7. Assessment of intracellular Ca²⁺ transients

NOTE: CSs are in culture for a total of 3 weeks; 2 weeks before freezing, and 1 week after thawing. The 'fresh' controls are age-matched.

1. After 1 week of culture, the thawed CSs are optimal for calcium handling optical imaging. Use a calcium dye (e.g., Cal520AM) to assess the uptake and release of Ca²⁺ from the cells.
2. Treat them with 100 µL of calcium dye medium per well and incubate for 60 min in the incubator. Set the incubator conditions to 37 °C, 5% CO₂, 21% O₂, and 90% humidity.
   NOTE: Cal520AM is light-sensitive. Perform all loading procedures and experiments in the dark.
3. Prepare the calcium acquisition and analysis system.
   1. Power the microscope, ensuring the environmental control option is on.
   2. Adjust the camera and framing aperture dimensions to minimize the background area.
      NOTE: Here, the Leica Thunder DMi8 microscope was used; other microscope systems are applicable as well until they allow a sampling rate above 30 frames/second (fps).
4. Record a video with a consistent stream of 2−10 peaks within 10 s and scan across the 96-well plate, initially moving to the left, then downward in a zig-zag fashion to cover the whole plate. Measure the calcium signal using a 488 nm laser; set the contrast to a black background with a bright green signal during calcium release.
5. After acquiring the Ca²⁺ transients, analyze the data with the fluorescence traces analysis software (e.g., CyteSeer, Vala Sciences) according to the manufacturer's instructions.

8. Flow cytometry analyses of dissociated cardiac spheroids

NOTE: In this study, flow cytometry was used to determine the viability of the CSs before and after the thawing process.

1. Collect the CSs in a 15 mL conical tube using a 5 mL pipet to avoid spheroid damage and centrifuge for 3 min at 70 x g. Aspirate the supernatant and add 1 mL of PBS.
2. Centrifuge for 3 min at 200 x g. Aspirate the supernatant and dissociate the CSs by adding 1 mL of cardiac detachment solution (e.g., tryple). Incubate the tube at 37 °C for 15 min.
3. Using a 5 mL pipet, mechanically dissociate the cells by flushing with 2 mL of basal medium until single cells can be seen when observed under the microscope.
4. Centrifuge for 3 min at 200 x g.
5. Aspirate the supernatant and fix the CMs with 200 µL of 4% paraformaldehyde (PFA) solution in 1x PBS. Incubate for 10 min at RT.
6. Centrifuge for 3 min at 200 x g. Aspirate the supernatant and add 1 mL of PBS.
   NOTE: Pause point: The fixed hiPSC-CMs can be stored at 4 °C for up to 4 weeks.
7. Transfer the cell suspension to a FACS tube and centrifuge for 3 min at 200 x g. Aspirate the supernatant and resuspend 1 x 10⁵ cells in 50 µL of the permeabilization buffer.
8. Incubate the cells for 30 min at 4 °C.
9. For immunofluorescence flow cytometry analysis, perform steps 8.9.1-8.9.4.
   1. Resuspend the cells in the flow cytometry buffer (50 µL) containing the α-actinin antibody at a dilution of 1:300. In another FACS tube, resuspend 1 x 10⁵ cells in the flow cytometry buffer (50 µL) with the respective isotype control (e.g., FITC mouse IgM, κ isotype [1:200 dilution]). Similarly, resuspend 1 x 10⁵ cells in 50 µL of flow cytometry buffer for negative control.
   2. Incubate the cells for 30 min at 4 °C.
   3. Add 2.5 mL of flow cytometry buffer and centrifuge the cells at 200 x g for 3 min at 4 °C; discard the supernatant and repeat this washing step twice.
   4. Resuspend the cells in 50 µL of flow cytometry buffer with the secondary-antibody goat-anti-mouse (1:300 dilution).
      NOTE: Place the tube in the dark since the secondary-antibody solution is light-sensitive.
10. For viability check with propidium iodide (PI), add 150 µL of PI per sample (1:1,000) and incubate for 15 min.
    NOTE: Place the tube in the dark since the PI solution is light-sensitive.
11. Adjust the gates according to the standard gating strategy as shown in Supplementary Figure 1 and analyze the cells with a flow cytometer.

9. Immunofluorescence staining of whole 3D spheroids

NOTE: This protocol is based on the protocol for high-resolution 3D imaging of whole organoids upon immunofluorescent labeling, which was previously published²⁹ and adjusted for cardiac spheroids. During the procedure, all pipet tips and conical tubes can be coated with 1% wt/v BSA-PBS in order to prevent the spheroids from sticking to plastics. To coat the materials, dip into the 1% BSA-PBS. Be careful not to damage the spheroids by using the 5 mL pipet, avoiding mechanical disruption.

1. Collect the CSs in a 15 mL coated tube with a 5 mL pipet. The spheroids are visible to the eye. Collect ~20-50 spheroids per antibody combination. Centrifuge for 3 min at 70 x g and aspirate the supernatant.
2. Carefully resuspend the spheroids in 1 mL of ice-cold 4% paraformaldehyde (PFA) solution in 1x PBS using a coated 1 mL tip.
3. Fix at 4 °C for 45 min. After 20 min, gently resuspend the spheroids using a coated 1 mL tip. This evens fixation among all spheroids.
4. Add 10 mL of ice-cold PBS to the tube and gently mix by inverting the tube. Incubate for 10 min at 4 °C and spin down at 70 x g for 3 min.
   NOTE: From this step onward, coating of tips and conical tubes is generally not needed as CSs do not stick to the tip after fixation.
5. Block the CSs by resuspending the pellet in ice-cold SWB (200 µL of SWB per well) and transfer the spheroids to a 24-well suspension culture plate.
   NOTE: CSs from one large pellet can be split over multiple wells to perform different stainings. Use ~20-50 CSs per antibody combination.
6. Incubate at 4 °C for at least 15 min.
7. Add 200 µL of the SWB in an empty well to serve as a reference well.
   NOTE: For the immunofluorescent staining, 48- or 96-well plates can also be used to reduce antibody usage. However, the staining and washing results can be reduced due to the smaller volume per well.
8. Allow the spheroids to settle at the bottom of the plate, by leaving the plate tilted at a 45° angle for 5 min.
9. Remove the SWB, leaving the CSs in 200 µL of the SWB (use the reference well to estimate the minimal volume of 200 µL).
10. Add 200 µL of the SWB with the primary antibodies 2x concentrated (e.g., ɑ-actinin [1:200] and Troponin T [1:200]) and incubate overnight at 4 °C while rocking/shaking (40 rpm on a horizontal shaker).
11. The next day, add 1 mL of the SWB to each well.
12. Allow the spheroids to settle at the bottom of the plate by leaving the plate at a 45° angle for 5 min.
13. Remove the SWB, leaving 200 µL in the plate. Add 1 mL of SWB and wash for 2 h with slow rocking/shaking.
14. Repeat steps 9.12 and 9.13 twice more.
15. Allow the CSs to settle at the bottom of the plate by leaving the plate tilted at 45° for 5 min. Remove the SWB, leaving 200 µL in each well
16. Add 200 µL of the SWB with secondary antibodies, conjugated antibodies, and dyes 2x concentrated (e.g., DAPI [1 µg/mL], mouse-AF488 [1:500], rabbit-AF568 [1:500]), and incubate overnight at 4 °C in the dark, while slowly rocking/shaking.
17. The next day, repeat steps 9.12 and 9.13 twice more.
18. Carefully transfer the CSs to a 1.5 mL tube and spin down at 70 x g for 3 min.
19. Remove as much of the SWB as possible by pipetting without disrupting the CSs.
20. Add the embedding solution (ES; at least 50 µL, at RT) using a 200 µL tip with the end cut off and resuspend gently to prevent bubble formation and incubate at RT for 20 min.
21. In the meantime, create a square container on a glass slide with either nail polish or silicone sealant.
22. Cut off the end of a 200 µL tip and transfer the CSs in ES to the middle of the square container.
23. Place a square coverslip on top. To reduce air bubbles, place one side of the coverslip first, then slowly lower the coverslip from one side to the other until there is no trapped air under the surface, and then release the coverslip.
24. Gently push on all edges of the coverslip to seal it to the nail polish or silicone sealant.
25. Leave the slide overnight at RT. The next day, the slide is ready for imaging.
    NOTE: Optical clearing by the ES can cause minor tissue shrinkage. However, this cannot affect the overall morphology of the CSs. The staining procedure can be paused here by storing the slides at 4 °C (for at least 1 week) or at -20 °C (for at least 6 months).