TITLE: Generation and assembly of human brain region-specific organoids
AUTHORS: Steven A. Sloan, Jimena Anderson, Anca M. Pașca, Fikri Birey, Sergiu Pasca

[DESCRIPTION]
<div class = "text-blocks"><div class = "text-block"><span style = "font-weight:bold;">The ability to generate region-specific three-dimensional (3D) models to study human brain development offers great promise for understanding the nervous system in both healthy individuals and patients. In this protocol, we describe how to generate and assemble subdomain-specific forebrain spheroids, also known as brain region–specific organoids, from human pluripotent stem cells (hPSCs). We describe how to pattern the neural spheroids toward either a dorsal forebrain or a ventral forebrain fate, establishing human cortical spheroids (hCSs) and human subpallial spheroids (hSSs), respectively. We also describe how to combine the neural spheroids in vitro to assemble forebrain assembloids that recapitulate the interactions of glutamatergic and GABAergic neurons seen in vivo. Astrocytes are also present in the human forebrain–specific spheroids, and these undergo maturation when the forebrain spheroids are cultured long term. The initial generation of neural spheroids from hPSCs occurs in <1 week, with regional patterning occurring over the subsequent 5 weeks. After the maturation stage, brain region–specific spheroids are amenable to a variety of assays, including live-cell imaging, calcium dynamics, electrophysiology, cell purification, single-cell transcriptomics, and immunohistochemistry studies. Once generated, forebrain spheroids can also be matured for >24 months in culture.</span><span></span></div><div class = "text-block"><span style = "font-weight:bold;">Introduction</span></div><div class = "text-block">Understanding the development of the human nervous system and elucidating the mechanisms that lead to brain disorders represent some of the most challenging ongoing endeavors in neurobiology. One major obstacle is the restricted access to healthy and diseased human brain tissue for functional molecular and cellular studies. As a result, experimental paradigms have been largely confined to animal models or in vitro cell culture systems that do not fully recapitulate the developmental, architectural, and species-specific aspects of the human brain. Thus, there is a great need for humanderived model systems that recapitulate features of human CNS development and allow for the study of these processes in both healthy and diseased conditions.</div></div>

[STEPS]
?. [MEF feeder seeding and initiation of hPSC cultures]
Coat 10-cm cell culture dishes with 0.1% gelatin for at least  at  .
0 Room temperature
?. [MEF feeder seeding and initiation of hPSC cultures]
Thaw one vial of MEFs (containing ~5 × 106 cells) by placing it in a water bath at  until it is 80% thawed (and for no longer than ).
37 °C
?. [MEF feeder seeding and initiation of hPSC cultures]
Briefly spin the cells at  for  at  and plate at a density of 1–2 × 106 cells per 10-cm plate.
0 Room temperature
?. [MEF feeder seeding and initiation of hPSC cultures]
The next day, assess feeder quality before use; this will determine the quality of the hPSCs.
?. [Passaging hPSCs on the MEF feeder layer]
Remove hPSC medium from the refrigerator, aliquot the required amount into a centrifuge tube (~ per 10-cm plate), wrap the container’s cap with Parafilm to prevent contamination when putting it into the water bath, and warm in a water bath for up to .
12 ml
?. [Passaging hPSCs on the MEF feeder layer]
Remove the hPSC plate from the incubator and look at it under a microscope.
?. [Passaging hPSCs on the MEF feeder layer]
After wiping with a paper towel sprayed with 70% (vol/vol) ethyl alcohol, transfer the plates to the cell culture hood.
?. [Passaging hPSCs on the MEF feeder layer]
Use a pipette tip to carefully remove demarcated differentiated colonies from the plate.
?. [Passaging hPSCs on the MEF feeder layer]
While under the microscope, mark the differentiated areas on the bottom of the plate, using a marker.
Differentiated colonies may appear to have a hole or a pit in the center of the colony or a border of differentiated cells. (See Fig. 3a of the original publication).
?. [Passaging hPSCs on the MEF feeder layer]
Aspirate the medium and add  of dispase (0.875 mg/ml in hPSC medium) to each culture plate, and place in the cell culture incubator for  or until hPSC colonies have lifted.
5 ml
?. [MEF feeder seeding and initiation of hPSC cultures]
Once the MEFs are ready for plating hPSCs (after ), wash the culture plates at least once with fresh DPBS.
?. [Passaging hPSCs on the MEF feeder layer]
Transfer the suspended cells from a single plate to a 50-ml centrifuge tube. Use  of hPSC medium with 10 ng/ml FGF2 and 10 µM Y-27632 to wash each 10-cm plate.
5 ml
?. [Passaging hPSCs on the MEF feeder layer]
Perform this step three times, adding the medium to the appropriate 50-ml centrifuge tube after each wash.
?. [Passaging hPSCs on the MEF feeder layer]
Gently swirl the 50-ml centrifuge tubes to suspend the harvested cells in  of hPSC medium.
20 ml
?. [Passaging hPSCs on the MEF feeder layer]
Transfer the hPSC medium and suspended cells to a freshly prepared feeder plate (from Step 4).
?. [Passaging hPSCs on the MEF feeder layer]
Gently swirl the plate and slide the plate in a back-and-forth motion on the shelf of the incubator to evenly disperse the cells.
?. [Passaging hPSCs on the MEF feeder layer]
Incubate the cells in the cell culture incubator. Leave the cells undisturbed for the first day after passaging, and replace the medium every day thereafter for 5–7 d.
Critical Step: Replace hPSC culture medium at approximately the same time each day.
?. [Passaging hPSCs on the MEF feeder layer]
To replace the medium, warm aliquoted fresh hPSC medium in a water bath for up to  at , aspirate old medium from the plates, and add  of fresh medium per 10-cm culture plate.
37 °C
12 ml
?. [Passaging hPSCs on the MEF feeder layer]
Monitor the hPSC colonies to determine when they are ready for spheroid formation. This will usually be ~6–7 d from the last passage (depending on the hPSC line); colonies should measure ~1.5 mm in diameter.
See Fig. 3a in original publication for examples of hPSC colonies of appropriate, and inappropriate, morphology.
?. [Passaging hPSCs on the MEF feeder layer]
When cultures are ready, proceed to the next step. Alternatively, continue to passage hPSCs every 5–7 d (depending on the growth rate of each line) by repeating Steps 1–13; however, bear in mind that, ideally, hPSCs should not be kept beyond passage 40.
Critical Step: It is essential for the colonies to be large in order to generate spheroids (i.e., just before the time at which colonies would be passaged, or 1–2 d later). If the colonies are too small, allow more time for growth before subsequent steps. Avoid using hPSC colonies that touch each other because they will detach as large clusters together with the MEF layer. Avoid hPSC colonies that have grown too large and that are differentiating in the center.
?. [Suspending hPSC colonies to form spheroids]
Remove the hPSC media from the culture plates. Detach hPSC colonies by incubating them with 0.35 mg/ml dispase (dissolved in hPSC medium;  per plate) for  to  at , 5% CO2.
5 ml
37 °C
Check the status of the detachment after   and then every   thereafter. Different lines will require different durations of dispase treatment. Plates can be gently moved (back and forth) at the end of the incubation period to facilitate the detachment of curled hPSC colonies from the feeders. Almost all feeders should remain attached to the 10-cm plate, whereas the hPSC colonies should come off. Intact colonies should curl up and lift off the plate (see Fig. 3b in original publication).
?. [Suspending hPSC colonies to form spheroids]
Gently transfer the floating colonies with a pre-wet 25-ml pipette to a 50-ml Falcon tube.
?. [Suspending hPSC colonies to form spheroids]
Once the colonies settle at the bottom of the tube, aspirate the media and gently wash the colonies two times with  of warm hPSC medium (without growth factors).
20 ml
Critical Step: Do not centrifuge. Let the colonies settle at the bottom of the Falcon tube.
?. [Suspending hPSC colonies to form spheroids]
Gently resuspend 30–50 suspended colonies in  of neural induction medium (the medium should also contain 5 µM DM, 10 µM SB, and 10 µM Y-27632) and transfer to a 10-cm ultra-lowattachment plate.
12 ml
Caution: It is important to distribute the spheroids evenly once the plate is placed in the incubator to avoid clustering and fusion of colonies at the center of the plate.
?. [Suspending hPSC colonies to form spheroids]
Denote this day as day 0. Place the plate in the incubator at, 5% CO2 for . Keep the spheroids in the incubator on day 1 without any medium change to allow the spheroids a day to recover.
37 °C
?. [Neural induction]
Critical: Minimize the overall time that the spheroids are kept outside of the incubator at all stages of differentiation.(Day 2)After  of incubation, i.e., on day 2, replace the medium with  of neural induction medium (with DM and SB but without Y-27632).
12 ml
?. [Neural induction]
To change the medium, gently transfer the spheroids to a 50-ml Falcon tube and carefully aspirate the medium once they settle. Incubate the cells for a further.
?. [Neural induction]
(Day 3)Replace the medium as described in Step 18 and incubate the cells for a further .
?. [Neural induction]
(Day 4) If you plan to differentiate hSSs (when you reach Step 22, Step 24), replace the medium as described in Step 18, but include  IWP-2 in the medium in addition. If you plan to generate hCSs, replace the medium as before. Incubate the cells for a further .
Critical Step: IWP-2 should not be thawed at RT, as it will precipitate. Frozen aliquots should be placed directly into a water bath before use.
?. [Patterning and differentiation]
From day 6 onward, replace the medium daily with NM in place of the neural induction medium. Use NM supplemented with 20 ng/ml FGF2 and 20 ng/ml EGF. In addition, include 5 µm IWP-2 in the medium if you are generating hSSs.
?. [Patterning and differentiation]
From day 12 onward, the medium requirements for the generation of hCSs and hSSs deviate further.For generation of hCSs go to step 23.For generation of hSSs go to step 24.For generation of hSS-ISRAs go to step 25.
After neural spheroids grow larger (>2–3 mm in diameter), no more than 30 spheroids should be maintained per 10-cm plate. Critical Step: Neural spheroids tend to fuse to each other as they get larger (Fig. 3d of original publication). When changing the medium, be sure to separate them by either pipetting up and down gently with a 10-ml pipette or by using a P10 pipette tip to separate them when they are bigger. If spheroids fuse during later time points (after ~day 50), it may greatly compromise the outcome.
?. [Generation of hCSs]
From days 12 to 15, continue to replace medium daily with NM supplemented with 20 ng/ml FGF2 and 20 ng/ml EGF.
?. [Generation of hCSs]
From day 16 onward, change the medium every other day until day 25. Continue to use NM supplemented with 20 ng/ml FGF2 and 20 ng/ml EGF.
?. [Generation of hCSs]
From days 25 to 43, replace FGF2 and EGF with 20 ng/ml BDNF and 20 ng/ml NT3. Perform medium changes with  -  of supplemented NM per 10-cm plate every 2–3 d.Go to step 27
14 ml
15 ml
?. [Generation of hSSs]
From days 12 to 23, supplement the NM (already containing 20 ng/ml FGF2, 20 ng/ml EGF, and 5 µm IWP-2) with  SAG. Perform medium changes with   -  of supplemented NM per 10-cm plate every day.
14 ml
15 ml
Critical Step: Note that early hSSs can be smaller in diameter in comparison with early hCSs during 12–20 d of differentiation
?. [Generation of hSSs]
From days 25 to 43, replace FGF2 and EGF with 20 ng/ml BDNF and 20 ng/ml NT3. Perform medium changes with  -  of supplemented NM per 10-cm plate every 2–3 d.Go to Step 26.
14 ml
15 ml
?. [Generation of hSS-ISRAs]
On days 12, 13, and 14, supplement the NM (with IWP-2) with 100 nM SAG. Perform medium changes with  -  of supplemented NM per 10-cm plate every day.
14 ml
15 ml
?. [Generation of hSS-ISRAs]
From days 12 to 15, add  RA (in addition to the IWP-2) to the neural induction medium (Fig. 2c of original publication).
Caution: RA is air- and light-sensitive. Minimize light exposure.
?. [Generation of hSS-ISRAs]
From days 25 to 43, replace FGF2 and EGF with 20 ng/ml BDNF and 20 ng/ml NT3. Perform medium changes with   -  of supplemented NM per 10-cm plate every 2–3 d.
14 ml
15 ml
?. [Maintenance and long-term culture]
From day 43 onward, perform media changes with   -  of NM without growth factors every 4 d. Using careful sterile technique, spheroids can be maintained for many months in culture, if desired (Fig. 3d of original publication).Carry out any additional procedures at time points determined by your experiment.  Examples of additional procedures that can be carried out on the spheroids include:
17 ml
18 ml
?. [Maintenance and long-term culture]
For assembly of forebrain spheroids, transfer one hCS and one hSS to a 1.5-ml microcentrifuge tube that is resting in a standard microtube rack (Fig. 4f of original publication). Denote the day of assembly as day 0. The size of the spheroids to be assembled should not exceed 3 mm, as they are challenging to fuse once they are too large.
Critical Step:  Once in the tube, make sure that there are no bubbles underneath the spheroids and that they rest next to each other in contact at the bottom of the tube and are not stacked up vertically.
?. [Maintenance and long-term culture]
Incubate the spheroids for at least  in  of NM, completely replacing the medium on day 2.
1 ml
Caution: Because the assembly interface is fragile at this stage, perform the medium change in the tube very carefully.
?. [Maintenance and long-term culture]
Check that fusion is complete. Fusion is considered complete when the two spheroids are inseparable with gentle shaking of the Eppendorf tube (Fig. 4g of original publication).
?. [Maintenance and long-term culture]
Add enzyme stock solution (use  per dissociation dish, which should correspond to about 15 early-stage spheroids or seven late-stage spheroids, plus an extra ) to a 50-ml Falcon tube, and bubble 5% CO2 through the solution at  until the solution turns from red to orange.
10 ml
2 ml
0 Room temperature
?. [Maintenance and long-term culture]
Once warmed and orange in color, add papain to the enzyme stock (for neural spheroids up to day 200, use 30 U/ml; after that differentiation stage, use 50 U/ml).
?. [Maintenance and long-term culture]
Add  –  of L-cysteine per  of enzyme stock solution and put the solution into a   water bath.
0.0036 g
0.0042 g
20 ml
[to 37 °C ]
?. [Maintenance and long-term culture]
Transfer the spheroids to a 6-cm Petri dish, remove any residual medium, and use a no. 10 scalpel blade to chop the spheroids into
Finer pieces are preferable (this will require at least   of chopping).
?. [Maintenance and long-term culture]
Use  of enzyme stock solution to wet a 0.22-µm filter, and then filter the  of enzyme stock solution into each Petri plate containing the finely chopped spheroids.
2 ml
10 ml
?. [Maintenance and long-term culture]
For the papain digestion, put the Petri dishes on a  heat block on top of a wet paper towel. Attach a hose from a CO2 tank to a 0.22-µm syringe filter and place the filter tip into a small hole cut into a lid of the Petri dish to flow 5% CO2 over the enzyme stock solution containing tissue pieces.
34 °C
Critical Step: The filter tip should not touch the solution to ensure that gas is not bubbling into the liquid but rather blowing over the top.
?. [Maintenance and long-term culture]
Gently shake the Petri plate every . Digest for  – .
The digestion time should be increased with the age of spheroids to a maximum of   for spheroids older than 250 d. During the digestion proceed with the next step.
?. [Maintenance and long-term culture]
During digestion, prepare low-Ovo and high-Ovo solutions by diluting 10× low-Ovo and high-Ovo solutions in inhibitor stock solution to make 1× final concentrations of each.
Trituration will require ~  of low-Ovo solution and   of high-Ovo solution per dissociation dish.
?. [Maintenance and long-term culture]
After digestion, transfer the enzyme solution with tissue pieces to a 15-ml Falcon tube. Add  to  of low-Ovo solution.
4 ml
5 ml
?. [Maintenance and long-term culture]
Add  of low-Ovo solution to the tube, and use a P1000 pipette to triturate 10–20 times. After the first round of trituration, allow chunks to settle to the bottom, and then transfer single cells (supernatant) to a new tube.
1 ml
The solution should become cloudy. Be careful to avoid introducing bubbles; do not lift the pipette out of the solution during trituration.
?. [Maintenance and long-term culture]
Repeat the trituration process with additional  aliquots of low-Ovo solution and 10–20 triturations each time.
1 ml
It may take two to three rounds until tissue chunks are dissolved. With each successive round, be more aggressive with trituration.
?. [Maintenance and long-term culture]
Once all tissue chunks have dissolved, use a P1000 pipette to carefully layer  of highOvo solution under the single-cell suspension.
1 ml
This should lead to a clear layer of liquid beneath a cloudy cell suspension.
?. [Maintenance and long-term culture]
Spin the cells down through the high-Ovo solution at  for  at .
0 Room temperature
?. [Maintenance and long-term culture]
Resuspend the cell pellet in the medium of choice.
Single-cell suspensions can be plated for cell culture, time-lapse imaging, calcium imaging, or immunohistochemistry, or can then be immunopanned or FACS-separated.
?. [Maintenance and long-term culture]
At the time point of choice, transfer the spheroids to a 1.5-ml microcentrifuge tube, using a cut P1000 tip.
Critical Step: Use the same fixation and embedding protocol when quantifying across hCSs or hSSs derived from multiple hPSC lines and differentiations. Variation in fixation and the sample-processing protocol can lead to variability in antibody staining.
?. [Maintenance and long-term culture]
After fixation, gently remove PFA and perform three  washes with DPBS at .
0 Room temperature
?. [Maintenance and long-term culture]
Add  of 30% (wt/vol) sucrose to facilitate cryoprotection. Leave the spheres in this solution at  until they sink (~ – ).
1 ml
4 °C
?. [Maintenance and long-term culture]
To embed spheres, use square disposable molds that have been filled with embedding solution.
To avoid creating bubbles while filling the mold, use a P1000 pipette, the tip of which has been slightly cut, to widen the opening.
?. [Maintenance and long-term culture]
Warm the solution mixture for at least  before proceeding to the digestion (Step 32). In the meantime, proceed to the next step.
?. [Maintenance and long-term culture]
Add  of DNase to each Petri plate and swirl the dish to mix.
200 µl
?. [Maintenance and long-term culture]
Spin at  for  at  until tissue chunks settle to the bottom. Aspirate and discard the supernatant.
0 Room temperature
?. [Maintenance and long-term culture]
Aspirate and discard the supernatant. A cell pellet should be visible at the bottom of the Falcon tube.
?. [Maintenance and long-term culture]
Gently remove the medium and briefly wash with  of DPBS.
1 ml
?. [Maintenance and long-term culture]
Remove the DPBS and add  of cold 4% (vol/vol) PFA. Leave the spheres to fix for at least  at .
500 µl
4 °C
The optimal fixation time varies depending on the antibody being used, and the age and size of the spheres. We thus recommend fixation trials be undertaken to optimize the optimal fixation required for specific antibody stainings; e.g., for membrane antigens, we have found that fixation for shorter periods of time (~ ) is optimal, whereas for nuclear antigens, optimal results are seen after a longer fixation (up to overnight).
?. [Maintenance and long-term culture]
During this time, prepare the embedding solution (1:1, OCT/30% (wt/vol) sucrose; see the ‘Reagent setup’ section).
?. [Generation of hSS-ISRAs]
From days 15 to 23, add  AlloP (in addition to the IWP-2) to the neural induction medium (Fig. 2c of original publication).
?. [Generation of hSS-ISRAs]
On day 24, replace medium with NM without additional growth factors.
?. [Generation of hSSs]
On day 24, replace medium with NM without additional growth factors.
?. [Maintenance and long-term culture]
After fusion, use a P1000 pipette with the tip cut off for a larger bore opening to transfer the assembled hCS–hSSs back to the ultra-low-attachment 10-cm plates. If fusion is carried out between days 60 and 90, when this process is most efficient, hCS–hSS assembly is usually complete after 3–4 d.
Earlier time points are also permissive. Assembly of hCS–hSSs at later stages of differentiation (after day 90) may take longer (up to 7 d).
?. [Maintenance and long-term culture]
Place the spheres into a mold using a P1000 cut pipette tip. Place four to ten spheroids into the mold, always making sure to mark their locations. If the spheres move while embedding, use a P20 tip to gently move them around the embedding solution.
Critical Step: Avoid carrying excess 30% (wt/vol) sucrose when transferring the spheres.
?. [Maintenance and long-term culture]
Place the mold on ice for  and allow the spheres to sink to the bottom.
?. [Maintenance and long-term culture]
Snap-freeze the spheres by placing the mold directly on dry ice. Once completely frozen, store the molds at −80 °C.
Pause Point: Frozen molds can be stored indefinitely at –80 °C until ready to cryosection.
?. [Maintenance and long-term culture]
When ready for sectioning, remove the mold from the –80 °C freezer and allow it to increase in temperature to  by placing it in the cryostat chamber for ~ before sectioning.
-20 °C
?. [Maintenance and long-term culture]
Remove the block from the mold, use OCT to paste it onto a specimen stage, and section using standard techniques. Use a brush to prevent crumbling of the sections.
We recommend that hCSs or hSSs be cryosectioned at a 10- to 15-μm thickness. Thicker sections are difficult to quantify because of the overlapping nuclei and high density of cells, especially in proliferative areas. However, if using a confocal microscope, thicker sections may be adequate.
?. [Maintenance and long-term culture]
Collect the sections on Superfrost Plus slides.
Pause Point: Slides can be stored for up to 1 year at –80 °C until ready to use.
?. [Maintenance and long-term culture]
For immunostaining, remove the slide from the –80 °C freezer and leave at  to thaw for  –.
0 Room temperature
?. [Maintenance and long-term culture]
Wash the slide once with DPBS to remove OCT/sucrose.
?. [Maintenance and long-term culture]
Remove the DPBS and use a hydrophobic PAP pen to draw a circle around the sections. Let the ink dry and wash the slide with DPBS again to remove excess OCT. From this step onward, add all solutions within the circle
?. [Maintenance and long-term culture]
Block for  at  with 10% NDS, 0.3% (vol/vol) Triton X-100, and DPBS. Note that the optimal concentration of NDS and Triton X-100 may vary according to the antigen of interest.
0 Room temperature
Critical Step: Perform immunostaining steps in a humidified chamber to prevent sections from drying out.
?. [Maintenance and long-term culture]
Add primary antibodies diluted in blocking solution at 4 °C overnight. Some examples of antibodies and concentrations that we have used successfully are listed in Table 1 of the original publication.
Critical Step: Some antibodies might need longer incubation periods; therefore,further optimization may be required.
?. [Maintenance and long-term culture]
Perform three  washes at  with DPBS.
0 Room temperature
?. [Maintenance and long-term culture]
Add appropriate secondary antibodies (we use species-specific anti-IgG (H+L)–Alexa Fluor–conjugated antibodies, diluted between 1:500 and 1:2,000) diluted in blocking solution for  at .
0 Room temperature
Critical Step: Protect the sections from light during this and the following steps.
?. [Maintenance and long-term culture]
Perform three  washes at  with DPBS.
0 Room temperature
?. [Maintenance and long-term culture]
Add Hoechst solution (1/10,000 in DPBS) for  –  at .
0 Room temperature
?. [Maintenance and long-term culture]
Remove excess Hoechst solution by washing with DPBS.
?. [Maintenance and long-term culture]
Remove the DPBS, let the sections air-dry for 5–10 min, and mount by adding one drop of Aquamount solution on top of the sections and placing a cover glass on top.
?. [Maintenance and long-term culture]
Use a fluorescence microscope to visualize the immunostained sections. See Fig. 4 of the original publication forrepresentative immunostainings of hCSs and hSSs with dorsal- and ventral-specific domain markers.
Critical Step: Ideally, cryosections for quantification should be sampled throughout the thickness of the spheroid. Avoid quantification of cryo-sections that are not at least 50 μm apart (five cryo-sections apart, if cutting at 10 μm) to prevent sampling the same cells. Avoid extreme edges of the hCSs or hSSs, as they are not likely to be representative, and important cell types might be missed that are usually present only in the deeper parts of the spheroids.
?. [Maintenance and long-term culture]
Incubate intact spheroids with  Fluo-4 diluted in NM for  at  and 5% CO2, followed by a  wash.
37 °C
?. [Maintenance and long-term culture]
Using an appropriate imaging system, record spontaneous calcium activity for several minutes (one frame every 8–10 s) in one 10-μm z-stack plane. Maintain the spheroids in NM during the recordings. We do not perfuse during calcium-imaging sessions, in order to minimize drift.
?. [Maintenance and long-term culture]
Export fluorescence intensity (F) as mean gray values in the ImageJ software (https://imagej.nih.gov/ij/). Signal decay should be controlled by subtracting the mean fluorescence of the background (Fb). To estimate changes in intracellular calcium, ΔF is computed as (Fell-Fb)/F0, where F0 represents the minimum F value per cell across the entire duration of recording from which Fb was subtracted.
?. [Maintenance and long-term culture]
Fix the spheroids with 4% (vol/vol) PFA for at .
4 °C
Note that this incubation time was optimized for 50- to 150-d-old hCSs, hSSs, or forebrain assembloids. However, given that the size of the spheroids after this age does not substantially change, this fixation time is probably suitable for older spheroids as well. If using younger spheroids (up to 25 d old), incubation times may be shortened. Keep in mind, however, that iDISCO clearing tends to result in a shrinking of tissue, so using early spheroids may make handling challenging.
?. [Maintenance and long-term culture]
Perform three  washes at  with DPBS.
0 Room temperature
?. [Maintenance and long-term culture]
Transfer the spheroids to 4-ml glass vials.
?. [Maintenance and long-term culture]
Dehydrate the spheroids with a day-long methanol (MeOH) dilution series at  (20%, 40%, 60%, 80%, and 100% MeOH/DPBS,  per step).
0 Room temperature
?. [Maintenance and long-term culture]
Incubate the spheroids in 5% H2O2/95% MeOH overnight at .
4 °C
?. [Maintenance and long-term culture]
The following day, rehydrate with a reverse MeOH dilution series (80% MeOH/DPBS, 80%, 60%, 40%, 20%, 0%, and 0% MeOH/0.2% Triton X-100/DPBS,  per step).
?. [Maintenance and long-term culture]
Incubate the spheroids overnight in 0.2% (vol/vol) Triton X-100, 20% DMSO, and 0.3 M glycine/DPBS at .
37 °C
?. [Maintenance and long-term culture]
Block the spheroids with 0.2% (vol/vol) Triton X-100, 10% DMSO, and 6% goat serum/ DPBS at  for .
37 °C
Critical Step: Perform this and all the following steps in closed tubes filled to the topto prevent oxidation of tissue.
?. [Maintenance and long-term culture]
Incubate the spheroids with 0.2% Tween 20 for .
?. [Maintenance and long-term culture]
Incubate the spheroids in 10 μg/ml heparin/DPBS (PTwH) for .
?. [Maintenance and long-term culture]
Incubate the spheroids with the desired primary antibodies (e.g., anti-GFAP, 1:1,000 dilution; anti-GFP, 1:1,000; and anti-SOX2, 1:500) for  in PTwH with 5% DMSO and 3% goat serum at .
37 °C
?. [Maintenance and long-term culture]
Undertake a day-long wash series with PTwH ( to  steps for at least 5 h: : PTwH; : PTwH; : PTwH; : PTwH; and : PTwH).
?. [Maintenance and long-term culture]
Dilute the secondary antibodies in PTwH and 3% goat serum and incubate for an additional  at .
37 °C
?. [Maintenance and long-term culture]
Wash off excess secondary antibody by washing for  with PTwH. Repeat three times so spheroids have been washed for at least .
?. [Maintenance and long-term culture]
Incubate the spheroids in 50% THF/H2O overnight at .
0 Room temperature
?. [Maintenance and long-term culture]
Perform a three-step THF series ( 80%/H2O,   100% THF,   100% 1016 THF) at .
0 Room temperature
?. [Maintenance and long-term culture]
Clear the spheroids by incubating in benzyl ether. Neural spheroids should be cleared within only a few minutes of incubation.
?. [Maintenance and long-term culture]
Image cleared tissue. When imaging cleared tissue, it is critical to use a chamber that is resistant to the organic solvents used in Step 42 and 43). For this purpose, we 3D-printed a chamber using semi-clear acrylic-type plastic from ProJet at the Stanford 3D Printing Core and followed the instructions that can be found at https://idisco.info/idisco-protocol/. If using this chamber, use silicone sealant to fix a rectangular coverslip on the bottom of the 3D-printed chamber. When spheres are cleared, add them to the chamber with benzyl ether.
Critical Step: Make sure the chamber is completely sealed to the cover glass to avoid any benzyl ether leaking onto the microscope. We recommend visiting https://idisco.info/ for additional troubleshooting tips.
?. [Live imaging of cell migration and calcium dynamics]
Transfer a fused hCS–hSS to a well of a 96-well plate (glass-bottom plate) in  of neural medium.
200 µl
Owing to the limited amount of medium that can be included in a 96-well plate, avoid putting more than one brain assembloid into a well.
?. [Live imaging of cell migration and calcium dynamics]
Place a 96-well plate into an inverted microscope with a motorized stage under environmentally controlled conditions (37 °C, 5% CO2).
?. [Live imaging of cell migration and calcium dynamics]
Maintain the assembled spheroids in an environmentally controlled chamber for  –  before imaging to let them acclimatize and settle. Movement in the room should be kept to a minimum.
?. [Live imaging of cell migration and calcium dynamics]
Set up the imaging positions so that up to ~8 fields with z stacks (corresponding to eight assembled hCS–hSSs) can be imaged in a sequential manner, with each position or frame being imaged at a rate of 15–20 min per frame. Our z stacks normally span a depth of 50–150 μm and have a step size of 1.5–2.5 μm. We have found that hCS–hSSs can be imaged for up to ~18 h, after which time imaging must be paused, so the medium should be carefully changed before continuing imaging. On our live-imaging system (Leica SP8 confocal microscope), we use The laser power density should be empirically determined in other imaging systems.
?. [Live imaging of cell migration and calcium dynamics]
If spheroids drift and this impacts image analysis, correct the images for drift computationally. We have found that the image registration plug-ins from ImageJ (e.g., linear stack alignment with SIFT) are generally sufficient to correct for minor-to-moderate drift. However, we discard sessions with major drift due to handling. Occasional drifting of spheroids may occur, as they are not embedded during imaging sessions.
?. [Patterning and differentiation]
Change the medium daily on days 6–11. At day 6, neural spheroids should be ~0.3–0.5 mm in diameter (Fig. 3c of original publication).
The medium can be changed as described in Step 18 or by gently tilting the plate (allowing the spheroids to settle) and removing the remaining medium.