Generating Forebrain-Type Cerebral Organoids from Human-Induced Pluripotent Stem Cells

1. Generation of iPSC Aggregates

1. Generation of single-cell monolayer cultures from iPSC colonies
   1. Prepare a basement membrane extract (BME) coated 6-well plate. Thaw BME on the ice at 4 °C for 2-3 h, dilute it with cold Dulbecco's Modified Eagle Medium F12 (DMEM-F12; 1:50 dilution), cover the plate with 1 mL/well of the diluted BME solution, and store the plate overnight at 4 °C.
   2. Aspirate the medium and wash intact iPSC colonies of at least two wells of a 6-well plate with 0.5 mM EDTA in phosphate-buffered saline (PBS) twice before incubating colonies with 0.5 mM ethylenediaminetetraacetic acid (EDTA) in PBS for 4 min at room temperature (RT). Aspirate EDTA solution and gently detach the colonies by washing them off the bottom of the dish with 5 mL of DMEM-F12 medium. Collect them in a 15-mL tube and pellet them by centrifugation (4 min at 1,200 x g at RT).
      NOTE: iPSCs reprogrammed from commercially available fibroblasts have been successfully used.
   3. Aspirate the supernatant and incubate the iPSC colonies with 500 µL of cell-dissociation reagent for 6 min at 37 °C.
   4. Pipet the cell suspension several times gently up and down with a 1 mL pipette to break the remaining cell clusters into single cells.
   5. Add 4 mL of DMEM-F12 to the cell suspension to dilute the cell-dissociation reagent.
   6. Spin the cells down at 1,200 x g for 4 min at RT.
   7. Resuspend the cells in 2 mL of iPSC medium supplemented with 5 µM Y-27632 and seed cells into one well of a BME-coated 6-well plate.
      NOTE: Use the iPSC medium specified in the Table of Materials for a single-cell monolayer iPSC cultures.
   8. On the next day, replace the medium with fresh iPSC medium lacking Y-27632. From this point, continue to culture the cells, changing the medium every day until iPSCs are 100% confluent. Depending on the cell line and the confluency of the starting wells, this will take between 2-4 days.
   9. Once confluent, passage the iPSCs. Aspirate medium and apply 500 µL of cell-dissociation reagent on the cells.
   10. Incubate the cells for 5-10 min at 37 °C. Gently rock the plate to detach the cells.
   11. Wash the cells from the well using 2 mL of DMEM-F12 and collect them in a 15-mL tube. Add DMEM-F12 for a total volume of 5 mL.
   12. Spin down the cells at 1,200 x g for 4 min at RT and aspirate the supernatant.
   13. Seed the cells in iPSC medium supplemented with 5 µM Y-27632 in a 1:2 to 1:4 ratio on a BME coated 6-well plate (2 mL/well of medium).
   14. Continue to culture the cells for 2-5 days and change iPSC medium lacking Y-27632 every day. Once confluent, passage iPSCs (steps 1.1.4-1.1.8).
       NOTE: Culture the iPSCs for at least 2 passages as a monolayer in the iPSC medium specified in the Table of Materials before using them for the generation of iPSC aggregates to allow the cells to adapt to the culture conditions.
2. Use monolayer iPSC cultures when they are 70-90% confluent for the generation of iPSC aggregates.
   ​NOTE: iPSC adapted to the culture conditions as single cells are less prone to stress that leads to cell death during the dissociation and aggregation procedure. Monolayer iPSC cultures need to display typical pluripotent morphology with no evidence of differentiation. Test the cultures on a regular basis for mycoplasma contamination. Work only with mycoplasma-free iPSC cultures.
   1. Aspirate the culture medium from one well of a 6-well plate and apply 500 µL of cell-dissociation reagent on the cells.
   2. Incubate the cells for 5-10 min at 37 °C. Gently rock the plate to detach the cells.
   3. Wash cells from the well using 2 mL of DMEM-F12 and collect them in a 15-mL tube. Add DMEM-F12 for a total volume of 10 mL.
   4. For cell counting, take 25 µL from the cell suspension and mix it with 25 µL of trypan blue to mark dead cells. Count the living cells using a counting chamber.
   5. Collect enough cells (4,500 cells per iPSC aggregate) from the cell suspension in a 15 mL tube.
   6. Spin down the cells at 1,200 x g for 4 min at RT and aspirate the supernatant.
   7. Resuspend the cells in an appropriate volume of iPSC medium supplemented with 50 µM Y-27632 to obtain 4,500 live cells per 150 µL.
      NOTE: Using a high concentration of Y-27632 (50 µM) is critical for the survival of the iPSCs.
   8. Plate 150 µL in each well of a low-attachment 96-well U-bottom plate and place it in the incubator at 37 °C and 5% CO₂.
      NOTE: When generating iPSC aggregates, consider that at least 6 organoids are needed for quality control at day 20 (see Section 5).

2. Induction of Anterior Neuroectoderm

1. Closely monitor morphology changes of the iPSC aggregates every day under the tissue culture microscope using a 4X or 10X power lens. Observe at day 1, cell aggregates with clear borders. Continue to culture iPSC aggregates in the incubator at 37 °C and 5% CO₂.
   NOTE: A certain number of dead cells/well is normal and will not affect organoid generation.
2. Feed the iPSC aggregates starting from day 2 and continuing with every other day by gently aspirating approximately 2/3 of the medium without disturbing the cell aggregates at the bottom. Add an additional 100 µL of iPSC medium lacking Y-27632.
   NOTE: Within 4-6 days the cell aggregates will be 350-450 µm in diameter and exhibit smooth edges.
3. At this stage, pool the cell aggregates with a cut 100 µL pipette tip into a low-attachment 6 cm dish (maximum of 20 aggregates/dish) in cortical induction medium containing DMEM-F12 with N2 supplement (1:200), B27 supplement (1:100), glucose (0.2 mg/mL), cyclic adenosine monophosphate (cAMP; 0.15 µg/mL), 0.5% non-essential amino acids (NEAA), 1% L-alanyl-L-glutamine, heparin (10 µg/mL), and the compounds LDN-193189 (180 nM), A83-01 (500 nM), and inhibitor of Wnt response-1 (IWR-1) (10 µg/mL). Feed the cell aggregates by changing the cortical induction medium 3 days after transferring them to the 6-cm dish.
4. Closely monitor morphological changes during cortical induction under the tissue culture microscope using a 4X power lens.
   NOTE: After 4-5 days in cortical induction medium, edges of the cell aggregates should begin to brighten at the surface, indicating neuroectodermal differentiation. At this stage, radial organization of a pseudostratified epithelium emerges. Proceed to Section 3 to embed the cell aggregates in a BME matrix.

3. Embedding of Neuroectodermal Aggregates in a Matrix Scaffold

1. Thaw BME on ice at 4 °C for 2-3 h. Aliquot undiluted BME in sufficient amounts.
2. Prepare a plastic paraffin film sheet for the embedding procedure. Cut the plastic paraffin film using sterile scissors in 4 cm x 4 cm large pieces, put a piece of the plastic paraffin film over an empty 100 µL tip tray for 100 µL tips, and press with a gloved finger so that small dimples in the plastic paraffin film sheet are created (1 dimple/cell aggregate needed). Clean the plastic paraffin film with 70% ethanol and irradiate it with UV light (power: 15 watts, wavelength: 435 nm) under the closed sterile bench for 30 min.
3. Transfer each cell aggregate to one dimple of the plastic paraffin film sheet using a 100 µL cut tip with 1.5-2 mm opening in diameter. In the case that two cell aggregates are fused, do not separate them but transfer them together to one dimple.
4. Gently aspirate the medium surrounding the cell aggregates using an uncut 100 µL pipette tip.
   NOTE: Be careful not to suck the cell aggregates into the tip, as this will damage the aggregates.
5. Add 40 µL of undiluted BME to each cell aggregate.
6. Position each cell aggregate in the middle of the BME drop using an uncut 100 µL pipette tip.
   NOTE: Be very careful not to harm the developing neuroepithelium with the pipette tip.
7. Carefully transfer the plastic paraffin film sheet using sterile forceps into a 10 cm Petri dish (or another sufficient cell culture dish) and place the dish in the incubator for 15-20 min to allow the BME to solidify.
8. Meanwhile, prepare a low-attachment 6 cm dish containing 5 mL of cortical induction medium.
9. After the polymerization of BME, remove the droplets containing the cell aggregates from the plastic paraffin film sheet. Using sterile forceps, turn the plastic paraffin film over and gently squeeze the cell aggregates into the slightly tilted (about 30 °) low-attachment 6 cm dish until the droplets fall off the plastic paraffin film sheet. Transfer a maximum of 16 cell aggregates into one 6 cm dish.
10. Continue to incubate the cell aggregates at 37 °C.

4. Generation of Forebrain-type Organoids from Neuroectodermal Aggregates

1. One day after embedding in a BME matrix, place organoid culture dishes on a rocking cell culture shaker with a tilting angle of 5 ° and 14 rpm, installed in a cell culture incubator.
2. Monitor organoids every day. Homogeneous neuroepithelial loop-like structures gradually develop after embedding in the BME matrix.
3. When neuroepithelial loop structures are visible, change the medium to organoid differentiation medium containing DMEM-F12 with N2 supplement (1:200), B27 supplement (1:100), glucose (0.2 mg/mL), cAMP (0.15 µg/mL), 0.5% NEAA, 1% L-alanyl-L-glutamine, insulin (2.5 µg/mL)
4. Replace the organoid differentiation medium every 3-4 days until the desired differentiation time point is reached. Then, fix organoids (see Section 5).
   NOTE: Occasionally, the tissue may exhibit buds of optically translucent tissue without loop structures. Although this is not ideal, this is not affecting the development of cortical structures. The organoids can be cultured in organoid differentiation medium for up to 40 days. For extended culture periods (40-100 days) the organoid differentiation medium can be supplemented with 1:50 BME to increase tissue complexity and with BDNF and GDNF to allow neuronal survival and maturation.