Generation of Human Brain Organoids for Mitochondrial Disease Modeling

NOTE: The use of human iPSCs may require an ethical approval. iPSCs used in this study were derived from healthy control individuals following local ethical approval (#2019-681). All cell culture procedures must be performed under a sterile cell culture hood, carefully disinfecting all reagents and consumables before transferring under the hood. Human iPSCs used for differentiation should have a passage number below 50 to avoid potential genomic aberrations that may occur upon extensive culture. The pluripotent state of the cells should be validated before organoid generation, for example, by monitoring the expression of pluripotency-associated markers such as NANOG or OCT4. Mycoplasma tests should be conducted weekly to ensure mycoplasma-free cultures.

1. Generation of brain organoids

1. Culture of human iPSCs
   1. Culture human iPSCs under feeder-free conditions in iPSC medium (see the Table of Materials) on coated 6-well plates and keep them in a humidified tissue culture incubator at 37 °C and 5% CO₂.
      ​NOTE: The carryover of feeder cells may hamper the organoid differentiation. Passage the cells at least once in feeder-free conditions.
   2. Passage the iPSCs at 80% confluency using enzyme-free detachment medium in ratios ranging from 1:4 to 1:12. To increase cell survival, add 10 µM Rho-associated protein kinase (ROCK) inhibitor (Y27632) after each splitting.
2. Dissociate the iPSCs (80% confluency)-Day 0.
   1. Prepare Cortical Differentiation Medium I (CDMI) (Table 1). Prewarm CDMI medium at room temperature (22-25 °C) before adding it to the cells.
   2. Wash the wells containing the iPSCs with phosphate-buffered saline (PBS) to remove dead cells and debris.
   3. Add 500 µL of prewarmed Reagent A (Table of Materials) to each well and incubate for 5 min at 37 °C. Check under the microscope to ensure cell detachment.
   4. Add 1 mL of iPSC medium to dilute reagent A to neutralize its activity.
   5. Use a 1000 µL pipette to dissociate the cells by pipetting up and down and transfer the cell suspension to a 15 mL centrifuge tube.
   6. Gently centrifuge the iPSCs at 125 x g for 5 min at room temperature (22-25 °C).
   7. Carefully aspirate the supernatant to avoid disturbing the cell pellet.
   8. Resuspend the pellet with 1 mL of CDMI to obtain a single-cell suspension, and count the cell number.
   9. Prepare the seeding medium with 9,000 iPSCs per 100 µL in CDMI supplemented with 20 µM ROCK inhibitor, 3 µM WNT-catenin inhibitor (IWR1), and 5 µM SB431542.
   10. Add 100 µL of seeding medium per well to a 96-well v-bottom plate.
   11. Keep the plate in a humidified tissue culture incubator at 37 °C and 5% CO₂.
3. Neurosphere generation
   1. On day 1, observe that round cell aggregates (neurospheres) with defined smooth borders are forming. Note the dead cells around the aggregates. Continue to culture in the incubator at 37 °C and 5% CO₂.
   2. On day 3, agitate the plate by tapping on the sides three times to detach dead cells.
   3. Add 100 µL of CDMI supplemented with 20 µM ROCK inhibitor, 3 µM IWR1, and 5 µM SB431542 to each well.
   4. Return the plate to the incubator at 37 °C and 5% CO₂.
   5. On day 6, carefully remove 80 µL of the supernatant medium from each well. Avoid touching the bottom of the well.
   6. Add 100 µL of CDMI supplemented with 3 µM IWR1 and 5 µM SB431542 to each well. Return the plate to the incubator at 37 °C and 5% CO₂.
   7. Repeat steps 5 and 6 every 3 days until day 18.
4. Transfer of neurospheres
   1. On day 18, prepare Cortical Differentiation Medium II (CDMII) (Table 1) and add 10 mL to a 100 mm ultra-low attachment cell culture plate.
   2. Use a 200 µL pipette with the tip cut off to transfer the round neurospheres from the 96-well plate to the 100 mm ultra-low attachment cell culture plate.
      NOTE: Be gentle to avoid damaging the neurospheres by making sure the opening of the tip is wide enough and that the aggregates are not aspirated too quickly.
   3. Remove 5 mL of medium from the plate containing the neurospheres and add 5 mL of fresh CDMII.
      NOTE: This procedure helps to reduce the amount of CDMI medium that may have carried over from the transfer of neurospheres.
   4. Place the plate on an orbital shaker at 70 rpm inside a humidified tissue culture incubator at 37 °C and 5% CO₂.
      NOTE: Visually inspect the neurospheres the next day. Increase the speed of the orbital shaker if the neurospheres are clumped together or attached to the bottom of the plate.
   5. Every 3 days, carefully aspirate the supernatant medium and replace it with fresh CDMII. Leave a small amount of the medium to prevent the neurospheres from drying out.
   6. On day 35, prepare Cortical Differentiation Medium III (CDMIII) (Table 1).
      NOTE: The matrix component should be dissolved in cold CDMIII.
   7. Aspirate the medium from the plate and add 10 mL of cold CDMIII.
      NOTE: It is more effective to use cold medium so that the matrix component can coat the organoids without forming clumps.
   8. After changing the medium, place the plate back on an orbital shaker at 70 rpm inside a humidified tissue culture incubator at 37 °C and 5% CO₂.
   9. Change the medium every 3-5 days depending on the rate of growth, as indicated by the color of the medium.
   10. On day 70, prepare Cortical Differentiation Medium IV (CDMIV) (Table 1). Use CDMIV medium until the desired age of organoids is reached. During this period, keep the plate on an orbital shaker set at 70 rpm inside a humidified tissue culture incubator (37 °C and 5% CO₂).
   11. Change the medium every 3-5 days, depending on the growth rate.

2. Immunostaining of brain organoids

1. Tissue preparation
   1. Prepare 4% paraformaldehyde (PFA) solution, and place it under a safety hood.
      NOTE: Wear personal safety equipment when handling PFA.
   2. Collect brain organoids and gently transfer them with a blunt-tipped 3 mL plastic Pasteur pipette to a 6-well plate filled with PFA.
      NOTE: Use organoids older than 40 days to allow the visualization of structures with higher cellular complexity.
   3. Keep the organoids in the PFA solution for 1 h at room temperature.
   4. Carefully remove the PFA with a 3 mL plastic Pasteur pipette, and wash the fixed organoids three times using PBS.
   5. Store the fixed organoids at 4 °C in PBS until further use.
2. Preparation of brain organoid slices
   1. Prepare a 3% agar solution and heat slowly until liquefied.
   2. Place the mold (the cut end of a 10 mL syringe) on a piece of absorbent filter paper (smooth side up). Place a droplet of agar onto it.
   3. Quickly take a single organoid out of the 6-well plate with a spatula and remove excessive PBS with filter paper.
      NOTE: Be careful not to touch the organoid directly with filter paper.
   4. Place the organoid onto the agar droplet.
   5. Repeat this procedure with up to three organoids.
      NOTE: Work fast to avoid solidification of the agar during this step.
   6. Refill the mold with agar until all the organoids are fully covered.
   7. Wait until the agar begins to solidify, and then gently transfer the entire mold containing the organoids, including the absorbent filter paper, onto a cooling element.
      NOTE: If a cooling element is unavailable, store the organoids for a few minutes in a refrigerator at 4 °C.
   8. In the meantime, prepare for the slicing procedure: place a razor blade (cleaned with acetone and washed with double-distilled water) into the holder of the vibratome, mount on the bath, and fill it with PBS.
   9. Remove the mold from (solidified) agar and use a scalpel to trim it to form a cube.
   10. Attach the agar cube containing the organoids on the carrier plate of the vibratome with adhesive gel (see the Table of Materials), and place it in the bath containing PBS.
   11. Adjust the vibratome (see the Table of Materials) to cut slices at a thickness of 150 µm.
       NOTE: Vibratome settings (proper angle, amplitude, frequency, and velocity of the blade) may be similar to those used for slicing fixed brain tissue derived from early postnatal animals. However, the ideal settings depend strongly on the type of the vibratome and must be determined in a first step to prevent distortion or even ripping of the tissue while cutting.
   12. Start the cutting procedure. Use a glass pipette or a spatula to gently transfer each freshly cut slice into a 24-well plate filled with PBS.
   13. Store the plate containing slices at 4 °C (for up to a few days) until further processing.
   14. Transfer the slices out of the plate with a glass pipette or a spatula onto microscope slides. Use a minimum of 2 slices per slide.
   15. Carefully remove the agar and excess PBS with a syringe.
   16. Allow the slices to dry until they adhere to the slides.
       NOTE: While microscope slides can be stored in plastic chambers filled with PBS at 4 °C, they should be stained as soon as possible after the slicing procedure.
3. Immunohistochemical staining
   1. Prepare the blocking solution (Table 1).
   2. Use a PAP pen to draw a hydrophobic border around the slices on the slide to help keep all the solutions on the slides.
   3. Carefully add the blocking solution on the slide, and incubate for 1 h at room temperature (22-25 °C). To avoid destroying the tissue, do not add the solution directly on top of the slices.
   4. Aspirate the blocking solution and apply the desired primary antibody diluted in the blocking solution.
   5. Incubate the slide overnight in a humidified chamber at 4 °C.
   6. Rinse the slide three times with 1x PBS for 10 min each.
   7. Incubate the slices with the specific secondary antibody diluted in the blocking solution and perform Hoechst staining (1:2,500) for 1 h at room temperature in the dark.
      NOTE: Remember to perform negative controls to confirm there are no non-specific binding or auto-fluorescence.
   8. Rinse three times with 1x PBS for 10 min each in the dark.
   9. Add one drop of mounting medium to the slice, place a coverslip on the edge of the drop, and slowly lay the coverslip down toward the slice to avoid air bubbles.
   10. Allow the slide to rest overnight at room temperature. Apply nail polish on the border of the coverslip to further seal the slide. For long-term storage, store at 4 °C.
4. Documentation of staining
   1. To scan large images for stitching, utilize a motorized upright wide-field microscope equipped with (see the Table of Materials for details) a high-quality objective; DAPI filter set (e.g., excitation (EX): 340-380 nm, dichroic mirror (DM): 400 nm, barrier filter (BA): 435-485 nm); fluorescein isothiocyanate filter set (e.g., EX: 465-495 nm, DM: 505 nm, BA: 515-555 nm); Alexa594 filter set (e.g., ET 575/40; T 600 LPXR; HC 623/24); digital camera; high-performance acquisition software allowing for automated stitches and stack operations.
   2. For image handling, use an image processing program capable of generating 8-bit tif-files, cropping stitches, adjusting contrast and brightness, merging the channels (e.g., blue, green, and red), and adding scale bars.
   3. To scan details, use a motorized confocal laser scanning microscope equipped with a high-quality objective, a UV laser (EX: 408 nm), an Argon laser (EX: 488 nm), a Helium-Neon laser (EX: 543), imaging software for a confocal microscope.
   4. For image handling of details, use an image processing program capable of generating maximum-intensity projections of confocal z-stacks (e.g., optical sections of 0.6 µm each), generating 8-bit tif-files, adjusting contrast and brightness, merging the channels (e.g., blue, green, and red), adding scale bars.
   5. Use a graphic editor to arrange the figures.

3. Bioenergetic profiling of brain organoids

1. Preparation of organoids for bioenergetic profiling
   1. Prepare the papain and DNase solution following the manufacturer's protocol.
   2. Transfer 3-5 organoids into a 6-well plate. Wash them two times with prewarmed PBS.
   3. Add 2 mL of prewarmed activated papain solution containing DNase. Using a blade, cut the organoids into small pieces.
   4. Place the plate onto an orbital shaker set at 27 rpm inside a cell culture incubator (at 37 °C, 5% CO₂), and incubate for 15-20 min.
      NOTE: The time of incubation depends on the organoid stage. Early-stage organoids can be used as they are. For organoids older than 3 months, it is recommended to cut the organoids into 2-3 pieces before dissociation and incubate the pieces at a rocking speed set at 27 rpm for 15-20 min at 37 °C. This procedure can help remove necrotic tissue that may be present in the later-stage organoids.
   5. Collect the digested tissues into a 15 mL tube and add 5 mL of organoid culture medium CDMIV (Table 1).
   6. Triturate the tissue with a 10 mL plastic pipette by pipetting up and down 10-15 times. Let the undissociated tissue settle down to the bottom of the tube.
   7. Carefully transfer the cell suspension to a 15 mL tube, avoiding any pieces of undissociated tissue. Filter the solution through a 40 µm cell strainer (e.g., polystyrene round-bottom tubes with cell strainer caps).
   8. Pellet the cells by centrifuging at 300 x g for 5 min at room temperature.
   9. Assess the cell number and quality using trypan blue.
   10. Plate the desired number (~20,000/well) of cells onto coated 96-well microplates. Change the medium 6-8 h after plating to Neuronal Medium (Table 1).
   11. Incubate the coated 96-well microplate in a CO₂ incubator (37 °C, 5% CO₂) for 4 days.
2. Bioenergetic profiling
   1. On day 3 after replating the dissociated cells, add 200 µL of calibration solution into each well of the bottom part of the 96-well microplate, and place the top green sensor cartridge onto the hydrated microplate.
      NOTE: Place the sensor cartridge on top of the microplate in the correct orientation, and ensure that the calibrant solution covers all the sensors.
   2. Incubate the hydrated 96-well microplate in a non-CO₂ incubator at 37 °C overnight.
   3. Turn on the analyzer to allow the instrument to stabilize at 37 °C overnight.
   4. On day 4 after replating, inspect the disassociated organoid culture on the 96-well microplate under the microscope to ensure that the cells appear as a confluent monolayer.
   5. Prepare Assay Medium (Table 1).
   6. Remove Neuronal Medium from all wells with a pipette without touching the bottom of the well to prevent cell damage. Alternatively, carefully invert the whole plate and then dry it on clean paper. Work quickly to avoid cell death.
   7. Wash the cells twice with prewarmed 200 µL of Assay Medium. Add Assay Medium to a final volume of 180 µL per well. Incubate the 96-well microplate in a non-CO₂ incubator at 37 °C for 1 h.
   8. Prepare 10 µM solutions of mitochondrial inhibitors in Assay medium. Note that the final concentration after injection is 1 µM.
   9. Load the sensor cartridge placed in the hydrated microplate with 10x solutions of the mitochondrial inhibitors.
      1. Add 18 µL of mitochondrial inhibitor 1 into port A.
      2. Add 19.8 µL of mitochondrial inhibitor 2 into port B.
      3. Add 21.6 µL of mitochondrial inhibitor 2 into port C.
      4. Add 23.4 µL of mitochondrial inhibitor 3 into port D.
   10. Place the loaded cartridge in the hydrated microplate in a non-CO₂ incubator at 37 °C until the start of the assay.
   11. Set up a running protocol in the instrument's software (Table 2).
   12. Press START. Take the loaded cartridge from the non-CO₂ incubator and place it into the analyzer for calibration.
       NOTE: Make sure the plate is inserted in the correct orientation and without the lid.
   13. Once the calibration step ends, remove the calibration plate. Take the 96-well microplate from the non-CO₂ incubator and place it into the analyzer. Click on CONTINUE to start the measurements.
   14. When the run is finished, remove the 96-well cell culture microplate from the analyzer and collect the medium from all the wells without disturbing the cells.
       NOTE: The medium can be stored at -20 °C and used later for measuring the amount of lactate released by the cells in the medium using an appropriate lactate assay kit.
   15. Wash the cells with 200 µL of 1x PBS in each well.
   16. After removing the PBS, freeze the plate at -20 °C.
       NOTE: The frozen plate can be used to quantify cells, proteins, or DNA in each well of the microplate. This quantification will be needed for normalizing the obtained bioenergetic rates. Follow the manufacturer´s instructions for cell, protein, or DNA quantification assays.