Making cerebral organoids:
Things you need at D0:
1) low FGF2 organoid start media
- iPS media
- regular FGF2 stock (1:2500)
- glutamaxx (1:100)
- Rock inhibitor (1:200)
- FBS (1:33.3)
2) Trypsin/EDTA at 0.05% (our stock is 0.25%)
- 1mL trypsin in 4mL PBS.
3) U-bottom ultra-low attachment 96wps
D0
1) Scrape colonies off in current iPS media and pipette into 15mL conical tube.
2) Let colonies settle to bottom of tube (2 minutes) and pipette off old media.
3) Wash 1X with PBS. Aspirate off PBS.
4) Add 1mL 0.05% trypsin/EDTA to each conical tube. Incubate tube at 37C for 2 minutes.
5) Pipette mixture with P1000 to break up call clumps until solution is cloudy.
5) Add 1mL MEF media (FBS will inhibit trypsin).
6) Take 10uL aliquot for cell counting.
7) Then Add 8mL low FGF2 organoid start media to each conical tube.
8) Spin in Selkoe centrifuge at 270g for 5 minutes.
9) While spinning, count cells. Make sure to calculate total number of cells per 2mL media (That was the volume from which you took the sample).
10) Calculate volume for resuspension for 9,000 cells/150uL (well).
11) Add 1mL of low FGF2 organoid start media and pipette cell pellet to resuspend.
12) Then add remainder of volume for proper cell concentration.
13) Plate 150uL/well of round bottom 96wp.
14) Place in incubator.
D1 – Observe organoids
1) Observe aggregates under microscope. Aggregates should be small with clear borders, with some dead cells around edges. No maintenance on this day.
D2 – Feed organoids
1) Aspirate 75uL media without disturbing organoid (half of well volume).
2) Add 150uL low FGF2 organoid start media to each well (same recipe as above).
D4 – Feed organoids
1) Aspirate 100uL media without disturbing organoid (~half of well volume).
2) Add 150uL iPS media + glutamaxx (1:100) and FBS (1:33.3) – no more FGF2 or ROCK inhibitor.
Things you need at D6:
1) N2 Neural Induction media + 1:100 glutamaxx
2) 24-well non-adherent plates (sterile, cell culture)
3) Small molecules and EdU if also doing treatments
D6 – Transfer organoids to non-adherent plates (24wps)
***If desired treat organoids with EdU at this time!
**** If desired treat with small molecules (DMSO/Chir/XAV) at this time!
- Use DMSO/Chir/XAV at 1:5000
1) Plate 1 organoid in 1 well of non-adherent plates (24wps)
- N2 Neural Induction media + 1:100 glutamaxx, 500uL/well
2) Transfer organoids using 5mL stripette to each well.
D8 – Feed organoids
1) Add another 500uL/well of N2 Neural Induction media (+1:100 glutamaxx).
*** Keep patterning with small molecules (Chir/XAV)
- Use DMSO/Chir/XAV at 1:2500 (Since twice as much volume on this day).
Things you need at D10:
1) Matrigel
2) Parafilm
3) Empty 200uL pipette tip rack (gray)
4) Tweezers (sprayed with ethanol)
5) 60mm culture dishes
6) Organoid Differentiation media (no Vit. A)
- For 500mL
- 240mL DMEM/F12
- 240mL Neurobasal
- 5mL N2 suppl.
- 5mL Glutamaxx
- 5mL NEAA
- 5mL Penstrp
- 5mL B27 (NO vitamin A suppl.)
- 250uL BME
- 125uL insulin
7) If patterning, Organoid Differentiation media (no Vit. A) + small molecules (DMSO/Chir/XAV)
- Use DMSO/Chir/XAV at 1:5000
D10/11 – Transfer to matrigel droplets
1) View organoids under microscope – should be brighter on outside, with smooth edges (see below).
2) If not like figures ‘c’ and looks like ‘e’, wait another day before embedding or possibly discard expt.
3) Thaw matrigel on ice for ~1-2hrs before you want to use it (500uL matrigel aliquot).
***Use 30uL matrigel per organoid, so each 500uL aliquot will allow for the embedding of 15 organoids.
4) Spray down parafilm rectangle with 70% ethanol (to fit over empty pipette tip rack) and use your findgertips to push parafilm into a 4×4 grid on the rack. Make sure to spray down hands first with ethanol.
***Each 60mm dish can hold 16 organoids (hence 4×4 grid/plate).
5) You can use a razor blade (sprayed with ethanol) to crop parafilm.
6) Use wide-orifice tips (yellow box) to transfer organoids from 24wp into each little parafilm hole. I generally try to pipette an organoid up in around 50-60uL media.
7) Remove excess medium from each tissue by carefully sucking off the fluid with an uncut 200-μl tip (once on parafilm).
8) Immediately add droplets of matrigel to each aggregate by dripping ~30 uL onto each tissue so that the droplet fills the parafilm hole.
***Add the matrigel quickly to avoid letting the tissues dry out. We typically perform embedding of 16 tissues at a time, a number that is manageable in a time frame that will not cause aggregates to dry out.
9) Position each aggregate in the center of the droplet using a 10-μl pipette tip to move the tissue within the droplet.
***This must be done immediately after adding the droplet, as the matrigel will begin solidifying once it is at room temperature.
10) Place the 60-mm dish containing droplets on parafilm back into the 37 °C incubator, and incubate it for 20–30 min to allow the matrigel to polymerize.
11) Grab the parafilm with droplets with tweezers and spray 5 ml of cerebral organoid differentiation medium without vitamin A over the droplets to dislodge them. Repeat a few times to get all the droplets, without generating too many bubbles.
***If patterning, use proper Organoid Differentiation media (no Vit. A) + small molecules (DMSO/Chir/XAV – 1:5000) for each condition.
12) You can also try to remove the Matrigel droplets from parafilm by first using sterile forceps to turn the parafilm sheet over and by agitating the dish until the droplets fall off the sheet. Any remaining droplets can be removed by using forceps to shake the Parafilm sheet in the medium more vigorously.
13) Place in incubator.
D11 – Observe organoids
Observe embedded tissues 24 h later under the microscope. Tissues should begin forming buds of more expanded neuroepithelium containing fluid-filled cavities within 1–3 days. See below.
D12 – Feed organoids
1) Feed with 5mL fresh organoid differentiation medium without vitamin A. Incubate for a further 48 h without agitation.
***If patterning, use proper Organoid Differentiation media (no Vit. A) + small molecules (DMSO/Chir/XAV – 1:5000) for each condition.
***Change the medium by tilting the dish, allowing the droplets to sink, and by carefully aspirating the medium.
Aspirate as much medium as possible without disturbing organoids. Replace with 5 ml of fresh medium.
D14/15- Feed organoids
1) If doing a DISC organoid experiment, continue feeding/patterning in 60mm dishes with above media until harvest at D19. If not, proceed to D14 – transfer organoids to spinner flasks.
Things you need at D14
1) Spinner flasks (cleaned and autoclaved)
2) Organoid Differentiation media (+Vit. A)
- For 500mL
- 240mL DMEM/F12
- 240mL Neurobasal
- 5mL N2 suppl.
- 5mL Glutamaxx
- 5mL NEAA
- 5mL Penstrp
- 5mL B27 (+vitamin A suppl.)
- 250uL BME
- 125uL insulin
D14 – Transfer organoids to spinner flasks
1) Gently pipette up organoids with a 25mL stripette and add to spinner flask with 75mL organoid differentiation media + Vitamin A (same media as above, but B27 supplement should have vitamin A supplement!!!)
2) Spin at 30rpm, forever.
3) Feed every week with 75mL organoid differentiation media + Vitamin A. Pipette off ~75mL and add fresh 75mL.