Table 2 |.

Troubleshooting table

Step	Problem	Possible reason	Possible solution
All steps	Contamination	Lapse in sterile handling	Regularly test for mycoplasma
			Check water sources for contamination and regularly clean water baths and incubator water pans
			Perform regular cleaning, maintenance, and certification of biosafety cabinets and incubators
			Ensure all users are following sterile technique (use 70% vol/vol ethanol spray before moving anything into the hood, regularly change/clean laboratory coats, be careful to not talk while facing an open incubator, proper gloving and degloving technique)
Procedure 1
All steps	EBs/organoids stick or fuse together	Too many EBs/organoids per well	Aspirate joined EBs/organoids. It is usually not possible to separate them once fused, and the mass may cause additional organoids to fuse to it if not promptly removed
			If using 4 mL of media per well, experiment with decreasing to 3 mL
			If there are too many EBs/organoids and spent media changes color too quickly, split EBs/organoids into more wells (1:2 split is most common)
			Verify that the orbital shaker remains at 95 rpm. Adjacent EBs/organoids can fuse if not under continuous agitation
		Cell death	Dying cells will leak DNA, which is sticky and may cause originally healthy neighboring cells/ organoids to stick together. Be sure to remove poor-quality organoids or dying cells during media changes. The tilt method of media changing (Step 15) allows healthy dense organoids to settle faster than the lighter dying organoids or cell debris, which can then be quickly aspirated away
	Media becomes yellow quickly	Too many EBs/organoids per well	Split EBs/organoids into more wells (1:2 split is most common)
		Contamination	Check for mycoplasma contamination
		CO2 sensor needs calibration	Check that the incubator is actually running at 5% CO2—it is possible for sensors to be off, and an incubator running at a slightly higher CO2 level can result in more acidified spent media
13 (day 1)	EBs do not form	Contamination or differentiation of iPSCs	Perform mycoplasma testing regularly
			Perform karyotyping to ensure iPSCs are karyotypically normal. Acquiring mutations that confer proliferative advantages or cause cell population drift in favor of pluripotency can both affect the differentiation potential and efficiency of iPSCs
			Check iPSCs for differentiation and other abnormalities before use for organoid generation
			Be selective when picking iPSCs, picking only iPSCs with good morphology. If there are not enough good iPSCs then continue picking/passaging until sufficient healthy colonies are obtained
		Not enough iPSCs seeded	Be selective when picking iPSCs, picking only iPSCs with good morphology. If there are not enough iPSCs then continue picking/passaging until sufficient healthy colonies are obtained
			If issues persist, try picking 600 fragments when passaging iPSCs before generating organoids. This will result in iPSCs becoming confluent quickly, but will also result in a more spread out/homogeneous plate with less large colonies (which are more likely to have dense areas of iPSCs with slower growth rate)
			Consider adding Y-27632 ROCK inhibitor to iPSC media 1 h before dissociation at 10 μM
			For sensitive iPSC lines that have genotype specific proliferation issues, EBs can be generated using forced aggregation in an AggreWell-800 plate instead of with spontaneous aggregation on the orbital shaker (Supplementary Methods)
		iPSCs were too confluent in plate at time of collection for protocol (growth inhibited)	Perform dissociation when colonies are 80–90% confluent (when they are in log phase growth)
16 (day 16 onward)	EBs/organoids disintegrate or are too small	Contamination	Perform mycoplasma testing regularly
			Check organoid cultures at every media change for signs of contamination visually (turbid media, unusual media color, yeast or fungal growth) and under inverted microscopy at higher magnification (bacterial growth that has directional motion)
		Started protocol with not enough iPSCs or iPSCs were too confluent in plate	Start over and adjust seeding density at next plating
			Use iPSCs in log phase growth. Do not use colonies that are too thin (too sparse) or too overgrown (growth inhibited)
		Being too aggressive with EB/organoid plates (excessive pipetting, splitting organoids too much)	Transfer nondisintegrating EBs/organoids to new wells and ensure sufficient number of organoids per well
			Split only when organoid mass exceeds half the diameter of the well. Splitting too often does not leave enough organoids to properly condition the media
			Perform splitting using a cut p1000 pipette tip or a 5 mL serological pipette, with gentle slow pipetting to not deform the organoids
		Reagent batch issue	Gem21, B27 and SM1 are interchangeable as the media supplement. Experiment with what works best for your iPSC line. This reagent is known to have potential lot-to-lot variability
		iPSC cell line specific issue	For sensitive iPSC lines that have genotype specific proliferation issues, EBs can be generated using forced aggregation in an AggreWell-800 plate instead of with spontaneous aggregation on the orbital shaker (Supplementary Methods)
			For difficult-to-neuralize iPSC lines, try using a different clonal line
			For difficult-to-neuralize iPSC lines, consider extending Neural Induction (Procedure 1, Step 14). Step 14 can be extended from 6 d up to 12 d
18 (days 28–30)	Rosettes not observed (Extended Data Fig. 3)	Internal rosettes are difficult to see	Validate lack of rosettes with immunohistochemistry, as rosettes may be internalized. If using brightfield, adjust microscope settings to see if the image resolves to show structures within the organoid
		Inefficient neural induction, specification, or patterning	Try to have enough organoids per well to condition the media (turn the media yellow) when it comes time to change media. This is important for maintaining the interorganoid secretion of growth factors that help the organoids effectively neuralize. This will change depending on which step of the protocol you are on, and you may not observe full media conditioning on daily media change steps
			For iPSC lines or genetic mutations that repeatedly are difficult to neuralize, consider extending the neural induction step from 6 d to up to 12 d, to ensure sufficient dual SMAD inhibition and the initiation of neural differentiation
			See previous solution for EBs not forming
		Reagent batch issue	Check efficacy of supplements, make sure supplements are not left in refrigerator for more than a week, minimize time supplements spend outside of the refrigerator or are exposed to direct light
		Incubator issue	Check CO2 and water/humidity levels in the incubator. Slightly higher CO2 levels that acidify the media quicker or a too dry incubator will adversely affect organoid differentiation and rosette formation
Procedure 2
9 (MEA)	Poor organoid attachment onto MEA plate	Problem with coating	When changing media, add 1:1,000 laminin to the media to try to help with reattachment
			In subsequent plating, Increase the concentration of laminin to 1:50 or 1:25 to help attachment
			In subsequent plating, add a preconditioning step to increase the MEA surface hydrophilicity: treat the surface with freshly prepared 0.07% Terg-a-zyme solution and/or 70% ethanol at room temperature for 1 h; then add fresh culturing media to each well and incubate for 2 d
		Organoids are too big	See Procedure 2, Step 7 and experiment with the amount of gentle trituration needed without breaking up the organoid
			If organoids are too big because later stage/older organoids are being plated, try plating closing to the 1-month-old timepoint to get better attachment, and allow them to mature in plate until desired assay timepoint
			If organoids are too big because they are created with AggreWell using forced aggregation, experiment with seeding fewer cells to create smaller starting organoids
		Organoids are unhealthy	Plate only rosetted organoids that do not exhibit cell death. In the 6-well plate that these organoids are being selected from, there should not be a large amount of cell debris, or neighboring organoids that are breaking apart in the well
18 (MEA)	Organoids show low activity	Unhealthy or not well-developed organoids	Plate only rosetted organoids that do not exhibit cell death. Using organoids with well-developed and visible rosettes is important
			Assess dead cells in the media. After initial plating there should be low cell death
			Due to the higher volume of media relative to the amount of organoids per well of an MEA plate, conditioning of the media may not produce a readily visible change in the phenol red indicator in the spent media. In addition to no or very low cell death and debris observed, operators can look for cell detachment as a negative indicator and neuronal cell outgrowth to cover the treated cell culture surface as a positive indicator of continued cell health. In long-term culture, healthy organoids should demonstrate steady or increasing measurable spontaneous electrical activity
		Poor organoid attachment onto MEA plate	Check the organoids’ position. If the organoids and their network are not in contact with any electrodes, the plate will not record any activity. Improve organoid attachment over the electrode by adding the primary and secondary coating (5–10 μL solution) only onto the active area (electrode array) of the well
			See additional solutions for poor organoid attachment above
		Other factors	If antibiotics were added, remove antibiotics from the culture media. Penicillin/streptomycin can affect neuronal development and activity
			If organoids demonstrate zero activity at 3 months old, start with a new batch
Procedure 3
20 (Calcium imaging)	Low fluorescence signal	Unhealthy cells	Check the level of health of your cells. There should not be cell debris, or cells that shrink when dye loaded (dying cells are small, round and absorb more dye, becoming abnormally bright)
		Poorly labeled cells	Prepare your dye solution fresh every time. Warm only enough dye solution for the imaging experiment. Do not re-use previously heated dye solution
			Make sure your dye has not expired
			Check the concentration of your dye solution and try to increase it
			Increase the incubation time
		Incorrect imaging settings	Make sure your optical setup is correct: check the filter settings and make sure they fit the fluorescent dye spectral properties (Oregon Green 488 BAPTA-1 is excited at 494 nm and imaged at 523 nm); check the power of your light source
			Make sure you do not bleach your dye by exposing the cells to extended and/or too intense light stimulation
Procedure 4
11 (AAV transduction)	Low expression	Unhealthy cells	Check the level of health of your organoids (e.g., is there higher than normal cell shedding, are the organoids consuming media, etc.)
		Organoid age	AAV transduction in early stage organoids (<4 weeks) can lead to poor expression levels (AAV does not integrate into the cell DNA); wait for older and more developed organoids where the number of mature neurons is higher (5 weeks and older)
		Low transgene expression	Check that vector promoter is appropriate to experimental purpose, e.g., cell specific, constitutive or ubiquitous promoters
			Check the AAV serotype is appropriate to your target cell type
			Increase dose and/or allow more time after transduction before analyzing
			Store AAV appropriately (see next solution)
		Poor AAV storage	Avoid repeated freeze–thaw by storing the AAV in single-use aliquots
			Store the AAV at −80 °C
			Keep the AAV on ice all the time once thawed