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

To generate pluripotent stem cell aggregates, when the stem cell cultures reach 70% to 90% confluency, add 500 microliters of cell dissociation reagent to one well of the six-well culture plate to detach the cells.

Induced pluripotent stem cells, which are adapted to monolayer culture conditions, are a good cell type to generate aggregates as they are less prone to stress-induced cell death during the dissociation and aggregation procedure.

After 5 to 10 minutes at 37 degrees Celsius, gently tap the plate, and use 2 milliliters of DMEM/F-12 to wash the cells from the bottom of the well. Transfer the resulting cell suspension into a 15-milliliter conical tube, and bring the volume of the cell solution up to 10 milliliters with more DMEM/F-12 medium.

After counting, transfer 4.5 times 10 to the third cells per pluripotent stem cell aggregate into a new 15-milliliter tube, and collect the cells by centrifugation. Resuspend the pellet in the appropriate volume of pluripotent stem cell medium, supplemented with 50 micromolar ROCK inhibitor to achieve a 4.5 x 10³ cells per 150 microliters of medium concentration.

Next, add 150 microliters of cells to individual wells of a 96-well low-attachment U-bottom plate, and place the plate in a 37 degrees Celsius and 5% carbon dioxide incubator. For monitoring anterior neuroectoderm induction, use a tissue culture microscope to closely observe the morphologic changes of the pluripotent stem cell aggregates every day under low magnification.

On day 1, cell aggregates with clear borders should be observed. On day 2, carefully aspirate approximately 2/3 of the medium without disturbing the cell aggregates at the bottom of each well, and replace the discarded medium with 100 microliters of fresh pluripotent stem cell medium.

Between four and six days later, when the cell aggregates reach 350 to 450 micrometers in diameter and exhibit smooth edges, use a modified 100-microliter pipette tip to transfer up to 20 aggregates into a single 6-centimeter low-attachment culture plate containing 5 milliliters of cortical induction medium.

Replace the cortical induction medium with fresh medium once every three days, monitoring the morphology of the aggregates daily under the 4x objective. After four to five days in the cortical induction medium, the edges of the cell aggregates should begin to brighten at the surface, indicating neuroectodermal differentiation, and the radial organization of a pseudostratified epithelium should emerge.

To embed the neuroectodermal aggregates in a matrix scaffold, fill a basement membrane extract on ice for two to three hours. While the extract is thawing, use sterile scissors to cut plastic paraffin film into one 4-by-4 centimeter square per 16 organoids, and place each piece of film over an empty 100-microliter micropipette tip tray.

Press the paraffin film with a gloved fingertip so that small dimples appear, and clean the film with 70% ethanol. After UV irradiation in a closed, sterile biosafety cabinet for 30 minutes, use a modified 100-microliter pipette tip with a 1 and 1/2 to 2-millimeter opening to transfer each cell aggregate into one dimple in the film.

When all of the aggregates have been transferred, use an uncut 100-microliter pipette tip to carefully aspirate the medium from each dimple, and add 40 microliters of undiluted basement membrane extract to each cell aggregate.

Be careful not to damage the organoids by rough manipulation or aspiration into the pipette tip, as this will damage the developing neuroepithelium.

Use the pipette tip to position the aggregates into the middle of each drop, and use sterile forceps to carefully transfer the plastic paraffin film sheet into a 10-centimeter Petri dish. Place the dish in the incubator for 15 to 20 minutes.

While the extract is solidifying, add 5 milliliters of fresh cortical induction medium to one six-centimeter low attachment culture dish. At the end of the incubation, flip the paraffin film sheet over, and use sterile forceps to gently squeeze up to 16 polymerized droplets into each 6-centimeter dish. Then, return the aggregates to the cell culture incubator.

The next day, transfer the organoid culture dishes onto a rocking cell culture shaker tilted at a 5-degree angle at 14 RPM within a cell culture incubator with daily light microscopy monitoring until the aggregates reach the differentiation stage of interest.