An In Vitro Method to Generate Human Brain Organoids

All procedures involving sample collection have been performed in accordance with the institute's IRB guidelines.

1. Generation of Brain Organoids (23 days)

1. Initiation of neuroectoderm (5 days)
   NOTE: The following points should be considered before the start of differentiation. The reprogramming method (lentiviral-, sendai-virus-, episomal-, or microRNA-based etc.) to obtain human iPSCs should ideally be the same for all patient and control iPSC lines. Various reprogramming kits and instructions based on published protocols are available. The quality of the human iPSC lines is the key to accomplishing an optimal differentiation. Monitor colony and cell morphology with a microscope and validate pluripotency by testing the expression of markers such as Oct3/4, Nanog, or TRA-1-60.
   1. Culture human iPSCs under feeder-free conditions in medium A on a dish coated with Engelbreth-Holm-Swarm (EHS) matrix.
      NOTE: Grow human iPSCs feeder-free and serum-free to maintain a defined culture condition and to avoid an additional step for the removal of mouse embryonic feeder cells (MEFs) before the start of differentiation. The optimal passage number for human iPSCs to start differentiation ranges from passage 15 upon reprogramming to passage 70 in total. When passaging, detach hiPSC colonies as cells aggregate using an appropriate cell detachment solution with low mechanical stress and a 2-mL serological pipette to transfer aggregates to a new dish. Avoid dissociation to single cells, as this might induce differentiation and apoptosis in most iPSC lines. It was reported that long-term, single-cell passaging could increase genomic alterations in human iPSCs. Avoid cell detachment procedures, which require a centrifugation step to remove detachment solution, as this reduces the overall viability of iPSCs. Test all cultures for microbial contaminations, particularly mycoplasma, on a regular basis, because this might alter the quality of the iPSCs and their differentiation capacity.
      1. Coat a 60 mm tissue culture dish with EHS matrix as per the manufacturer's instructions.
      2. Thaw an aliquot of 1 x 10⁶ human iPSCs. Seed the iPSCs in a 60-mm tissue culture dish coated in EHS matrix and containing 5 mL of medium A (see the Materials Table). Change the medium daily and passage after 5 to 7 days when the cells reach ~80% confluency.
         NOTE: Passage the thawed iPSCs at 80% confluency using standard methods, such as the enzyme-free detachment of colonies, at least once before starting the differentiation. In brief, remove the iPSC medium, wash the cells once with pre-warmed 37 °C Dulbecco's Modified Eagle's Medium: Nutrient Mixture F-12 (DMEM/F12), and incubate the iPSCs following the manufacturer's instructions using reagent A (see the materials table). Don't exceed the recommended incubation time in order to avoid dissociation to single cells. Human iPSCs should be detached and floating as aggregates, not as single cells. They can then be transferred to new EHS matrix-coated dishes; for example, iPSC aggregates from one 60-mm dish can be distributed to 4 new 60-mm dishes.
         b. Check for mycoplasma with a mycoplasma detection kit according to the manufacturer's instructions. Use only mycoplasma-free iPSCs, as mycoplasma can alter the differentiation capability of iPSCs.
2. Dissociate iPSCs (80% confluent) and prepare a single-cell suspension using reagent B
   1. Wash the iPSCs once with pre-warmed (37 °C) DMEM/F12.
   2. Add pre-warmed reagent B (e.g., 1 mL in a 60 mm dish) and incubate the iPSCs for 5 min at 37 °C and 5% CO₂.
   3. Flick 20 times with a fingertip on the side and bottom of the dish to detach the iPSCs. Check for the detachment of cells under the microscope.
   4. Pipette the cell suspension up and down in the dish 5 times with a 1-mL micropipette.
   5. Add 3 mL of medium A to dilute 1 mL of reagent B and collect the cell suspension in a 15-mL centrifuge tube.
   6. Gently spin down the iPSCs (500 x g) for 4 min at room temperature.
   7. Resuspend the cell pellet in 1 mL of medium B and count the cell number with a hemocytometer.
      NOTE: Be aware of using only medium B for resuspension. Avoid using medium A, as it contains a too-high concentration of bFGF, which might inhibit the differentiation.
3. Dilute the cell suspension to 4.5 x 10⁵ cells per mL in medium B supplemented with 10 µM rho-associated protein kinase inhibitor (Y-27632).
4. Add 100 µL per well in a non-adherent, v-bottom, 96-well plate.
   NOTE: Make sure the cells are equally distributed in the suspension by shaking the tube each time before taking out 100 µL portions. It is important that each well should contain an equivalent cell number in order to obtain neurospheres homogenous in size and shape (round, defined surfaces).
5. Gently spin down the plate with the cells at 500 x g and room temperature for 3 min and incubate at 37 °C and 5% CO₂.
6. Change the medium daily by removing 50 µL and adding 50 µL of fresh medium B into each well for the next 5 days.

2. Embedding Neurospheres in EHS Matrix (4 days)

1. Prepare neurosphere medium by mixing the following: 1:1 mixture of DMEM/F12 and medium C (v/v), 1:200 (v/v) supplement 1, 1:100 (v/v) supplement 2 without (w/o) Vitamin A, 1:100 L-glutamine, 0.05 mM non-essential amino acids (MEM), 100 U/mL penicillin, 100 µg/mL streptomycin, 1.6 g/L insulin, and 0.05 mM β-mercaptoethanol.
2. Collect the neurospheres with a 200 µL micropipette using a ~2 mm tip previously cut with sterile scissors.
3. Place the neurospheres approximately 5 mm away from each other on paraffin film (3 x 3 cm²) in an empty 100 mm dish and carefully remove as much of the remaining medium as possible.
4. Add a drop (7 µL) of EHS matrix onto each single neurosphere.
5. Incubate the EHS matrix drops with the neurospheres for 15 min in an incubator.
6. Wash the neurospheres carefully from the paraffin film by flushing them with neurosphere medium. To flush, use a 1 mL micropipette and a new 100 mm Petri dish containing 10 mL of neurosphere medium.
7. Incubate the neurospheres for the next 4 days and add 2 mL of fresh neurosphere medium on day 2.
   NOTE: Make sure that the shelves in the incubator are flat so that the EHS matrix-embedded neurospheres will not clump together on one side of the dish.

3. Organoids in a Rotary Suspension Culture (14 Days)

1. Prepare brain organoid medium by mixing the following: 1:1 mixture of DMEM/F12 and medium C (v/v), 1:200 (v/v) supplement 1, 1:100 (v/v) supplement 2 w/o Vitamin A, 1:100 L-glutamine, 0.05 mM MEM, 100 U/mL penicillin, 100 µg/mL streptomycin, 1.6 g/L insulin, 0.5 µM dorsomorphin, 5 µM SB431542, and 0.05 mM β-mercaptoethanol.
2. Add 100 mL of brain organoid medium to each spinner flask through its side arms and place them in an incubator for pre-warming for at least 20 min.
3. Set up a stirring program at 25 rpm, according to the manufacturer's instructions.
   NOTE: Before transferring the EHS matrix-embedded neurospheres into spinner flasks, ensure they are all separated. If two or more are connected through the EHS matrix, separate them by cutting the connecting matrix with a scalpel.
4. Carefully transfer the EHS matrix-embedded neurospheres into spinner flasks containing 100 mL of organoid medium using a 2 mL serological pipette. Use the side arms of the spinner flask to transfer the neurospheres into the flask.
5. Place the spinner flasks on a magnetic stirring platform in an incubator at 37 °C and 5% CO₂; this is day 0 of the organoid culture.
6. Change the medium once per week (or more often when there is a color change) by removing half of the medium and adding the same amount of fresh medium.
   NOTE: When taking the spinner flasks out of the incubator, wait 3-5 min to let the organoids sink down to the bottom of the flask. Remove the medium by placing the glass pipette tip (connected to a pump) on the surface of the liquid; aspirate the medium carefully through one side opening/arm of the flask. These manipulations must be done under the laminar hood.