A Technique to Generate Brain Organoids from Human Embryonic Stem Cells

1. Stem Cell Maintenance

1. Maintain H9 hESCs on a layer of growth factor reduced basement membrane matrix (see the Table of Materials, henceforth simply referred to as matrix) according to the manufacturer's instructions.
   1. To coat one 6-well plate or one 10 cm dish, combine 100 µL of matrix with 5.9 mL of ice-cold Dulbecco's modified Eagle medium (DMEM)/F12 media. Wrap plates in paraffin film and store overnight at 4 °C. Use them on the next day for passaging cells after the excess matrix/media is aspirated.
2. Culture the cells week to week at approximately a 1:12 split ratio every 7 days. Maintain the cells using mTESR-1 media in a 37 °C, low oxygen incubator (5% O₂, 5% CO₂). Refresh media daily. Weed out differentiating cells from the culture between passages using glass tools.
3. H9 cells should be passaged four to six days prior to utilizing them to produce organoids. The cells should be passaged at approximately a 1:8 ratio of cell clusters. To do this, start by rinsing the cells with Dulbecco's modified Eagle medium/nutrient mixture F-12 (DMEM/F12) media and dissociate the cells with a neutral protease (e.g., dispase, henceforth referred to simply as protease), rinse with DMEM/F-12, and plate as 30-60 cell clusters across 4 plates (6-well or 10 cm) at ~20% confluency. Two days prior to harvest, transition them to a regular incubator (21% O₂, 5% CO₂). Plates should reach ~80% confluency when starting organoid formation.

2. Dissociation of the hESCs for Organoid Culture

1. Aliquot the protease stock solution (5 U/mL).
   NOTE: We typically freeze down 1 mL aliquots at -20 °C for use over several months.
2. Dilute the protease stock solution to the working concentration by adding 1 mL of the stock solution plus 5 mL of DMEM/F12 for each 6-well or 10 cm plate of hESCs.
3. Aspirate and remove cell culture media, then cover the hESCs with the protease solution. Place plates in the incubator for 10-15 min or until the edges of the colonies round up and begin to separate from the matrix.
4. Tilt the plate, aspirate the protease solution, and gently wash the cells with DMEM/F12 three times. Use 2 mL/well for each wash when using a 6-well plate and 6 mL when using a 10 cm plate. Make sure colonies stay attached to the matrix when performing this step.
5. Add back about 1.5 mL of fresh mTESR media to each well (or 5 mL for a 10 cm plate) and flush the cells off the plate using gentle pipetting.
6. Using a 10 mL pipette, gently aspirate and dispense hESC within the plate until they reach approximately 1/30th of their original size. Colony clusters should resemble ~250-350 µm sized squares at the completion of these steps.

3. Generation of Organoids

1. Transfer cells into a single ultra-low attachment T75 flask containing 30 mL of mTESR media without basic fibroblast growth factor (bFGF).
2. The next day, tilt the flask(s) such that the live cells pool in the corner (this may take 5-10 min on the first day, but will get quicker as the clusters get larger).
   NOTE: If there are a large number of cells that have adhered to the bottom of the flask at this step or any subsequent steps, transfer the cells to a new flask. It is normal to have a high population of dead cells for the first two days. When performing media changes, be sure to remove as much of the cell debris as possible.
3. Once the cells settle, aspirate off the media and dead cells, leaving about 10 mL of media containing the live cells.
4. Add ~20 mL of low bFGF media (DMEM/F12 supplemented with 1x N2, 1x B27, 1x L-glutamine, 1x non-essential amino acid (NEAA), 0.05% bovine serum albumin (BSA), and 0.1 mM monothioglycerol (MTG) supplemented with 30 ng/mL bFGF).
5. Check the cells on day 2. If most of the cells look healthy and bright, there is no need to do anything. However, if more than a third of the cells appear dark, replace the media (using the same tilting technique as in step 3.2) with ~20 mL of low bFGF media supplemented with 20 ng/mL bFGF.
6. On day 3, replace half of the media (using the tilting technique in step 3.2) with 20 mL of low bFGF media supplemented with 10 ng/mL bFGF.
7. On day 5, replace half of the medium (using the tilting technique in step 3.2) with 20 mL of neural induction media (NIM: DMEM/F12, 1x N2 supplement, 0.1 mM MEM NEAA, 2 µg/mL heparin).
   NOTE: If there are any large clusters of cells or organoids that are much larger than the others, they should be removed from the culture. Size is estimated by appearance under the microscope; for example, using an eyepiece with reticle. The majority of organoids are similarly sized (roughly 100 ± 20 µm). We removed organoids that were approximately 2x smaller or larger than the others.
8. Replace half of the medium (~15 mL) (using the tilting technique) with NIM every other day.
9. After 3 weeks in culture, add 100x penicillin/streptomycin to the media (NIM: DMEM/F12, 1x N2 supplement, 0.1 mM MEM NEAA, 2 µg/mL heparin) at a final concentration of 1x if desired. Refresh the media every other day.
   NOTE: In this fashion, we maintained the organoids for up to 6 months in culture.