Preparation and Immunostaining of Myelinating Organotypic Cerebellar Slice Cultures

All work involving animals complied with institutional policies and guidelines established by the UPMC, INSERM and the French and European Community Council Directive 86/609/EEC.

1. Preparation of Culture Medium and Culture Inserts (Hands-on Time ≈ 10–15 min)

NOTE: Perform this step in a flow culture hood under sterile conditions

1. Prepare 40 mL of culture medium, which consists of 50% BME, 25% Hank’s Balanced Salt Solution (1x), 25% heat-inactivated horse serum, supplemented with 2 mM glutamine derivative, 100 IU/mL penicillin-streptomycin and 4.5 mg/mL D-Glucose. Adjust the pH to 7.4.
2. Filter-sterilize the medium through a 0.22 μm filter adapted on a 50 mL plastic syringe.
   NOTE: Culture medium can be stored at 4 °C for at least a week.
3. For a standard litter of 6 pups, use two sterile 6-well plates. For each plate, remove the cover and add 1 mL of culture medium per well.
4. Place one culture insert into each individual well by holding the plastic edge with sterile forceps. Put the cover back on the plate.
   NOTE: The cerebellar slices collected from one pup (6-8) can be dispatched on two membranes (3-4 cerebellar slices per membrane insert).

2. Preparation of Dissection Medium (Hands-on Time ≈ 5 min)

NOTE: Perform this step in a flow culture hood under sterile conditions

1. Prepare 100 mL of dissection medium, which consists of Gey’s Balanced Salt Solution supplemented with 4.5 mg/mL D-Glucose and 1x penicillin–streptomycin (100 IU/mL each).
2. Filter-sterilize the medium through a 0.22 μm filter unit.
   NOTE: The dissection medium can be stored at 4 °C for at least a month.
3. Add 5 mL of cold dissection medium into 60 mm cell culture dishes. Prepare one 60 mm cell culture dish for each pup to be dissected.
4. Keep the dissection dishes on ice until use.

3. Preparation of dissection material (Hands-on Time ≈ 15 min)

1. Clean the bench with 100% ethanol.
2. Sterilize all dissection tools, a razor blade and the tissue chopper’s plastic platform with 100% ethanol.
3. Fix the razor blade and the plastic platform on the tissue chopper and adjust the cutting thickness to 300 µm.
4. Make sure all the ethanol has evaporated before starting the dissection.

4. CerebellumDissection and Sections Preparation (Hands-on Time ≈ 15–20 min per animal)

1. Place one of the 60 mm cell culture dishes containing the ice-cold dissection medium under the binocular microscope and remove the top plate.
2. Swab the head fur with 70% ethanol (carefully avoiding touching the eyes of the animal) and decapitate the animal using large sharp scissors, according to the approved procedure.
   NOTE: The cerebellar slices are generated from P9-P10 mice without any specific sex or strain bias.
3. Use small scissors to cut the skin along the midline of the head starting from the neck and going up to the mouse muzzle.
4. To hold the head and expose the skull, fold back the skin ventrally under the head, and pinch the two folds of skin between one’s fingers.
5. Insert the small scissors gently into the foramen magnum and cut the skull by making one lateral incision towards the side and then cut all around the head skull. Keep the tip of the scissors as parallel and close to the skull as possible while cutting, to avoid damaging the brain tissue.
6. Retrieve the dorsal part of the skull using fine-straight forceps. While carefully lifting the dorsal part of the skull, cut the adhering meninges (translucent irrigated membrane surrounding the brain tissue) if needed with small scissors to avoid damaging the brain tissue.
7. Carefully introduce fine-straight forceps (or alternatively use a spatula) between the ventral skull and the brain, gently flip out the brain and cut the optic and trigeminal nerves with small scissors.
8. Help the brain to drop by gravity into the 60 mm cell culture dish containing ice-cold dissection medium by turning the head upside down just above the dish and release the last adhesions with small scissors if needed.
9. Using fine forceps, orientate the brain with the dorsal side facing the researcher, and the ventral side lying on the bottom of the dish.
10. Under the binocular microscope, use the fine-straight forceps to immobilize the brain on the forebrain side and avoid touching the hindbrain, as it could get damaged. Separate the hindbrain from the rest of the brain (fore- and midbrain, see schematic on Figure 1Aa), using fine-straight forceps. To separate the cerebellum from the rest of the hindbrain, use the fine forceps to cut the cerebellar peduncles underneath the cerebellum (see schematic on Figure 1Aa’).
11. Once the cerebellum is isolated, carefully tear away the meninges using fine-straight forceps (see schematic on Figure 1Aa’).
12. Hold the cerebellum gently with the fine curved forceps and place it with the dorsal side up onto the plastic platform perpendicularly to the chopper razor blade (see schematic on Figure 1Bb).
13. Aspirate any excess of dissection medium around the cerebellum using a sterile thin-end pipette tip adapted on a 1 mL pipette (see schematic on Figure 1Bb).
14. Ensure that the cerebellum lays flat, but is not stretched, once placed onto the platform to get optimal sagittal slices during sectioning (see schematic on Figure 1Bb).
15. Slice 300 μm-thick sagittal sections of the cerebellum with the tissue chopper (see schematic on Figure 1Bb’). The force and speed of the blade have to be optimized on site.
16. Gently add a drop of dissection medium onto the sliced cerebellum. Using a wide-bore pipette tip placed onto a 1 mL pipette, slowly aspirate the sliced cerebellum and transfer it back into the 60 mm cell culture dish containing ice-cold dissection medium (see schematic on Figure 1Bb’’).
17. Clean the tissue chopper’s plastic platform with 100% ethanol between each cerebellum slicing. Let the ethanol evaporate before placing the next cerebellum onto the platform.
18. Use two fine-straight forceps (or alternatively titanium needles) to separate individual slices and select 6-8 slices from the vermis (see schematic on Figure 1Cc and 1Cc’).
19. Take a 6-well plate with culture inserts and transfer up to 4 selected slices onto one culture insert along with some dissection medium, using a wide-bore pipette tip adapted on a 1 mL pipette (see schematic on Figure 1Cc’).
20. Place the slices in the middle of the culture insert using the dissection medium or forceps to push and pull them gently into the right location, avoiding them to be in contact with each other (see schematic on Figure 1Cc’).
21. Remove any excess of dissection medium around the slices using a thin-end pipette tip. Ensure that the slices lay flat on the membrane (see schematic on Figure 1Cc’).
22. Place the 6-well plate into the incubator immediately after having added the slices on the membrane.

5. Slices Culture and Demyelination (Hands-on Time ≈ 15–20 min)

NOTE: Perform this step in a flow culture hood under sterile conditions

1. Replace the culture medium with 1 mL per well of pre-warmed and buffered fresh culture medium every 3 days after slice preparation.
2. For demyelination, remove all culture medium below the culture inserts after 6 days in vitro (DIV) and replace it with 1 mL per well of pre-warmed fresh culture medium containing 0.5 mg/mL LPC. Incubate for 15-17 h at 37 °C, 5% CO₂.
3. After incubation, wash the inserts by placing them in a 25 mm petri dish containing 1 mL of pre-warmed culture medium.
   NOTE: The inserts should be in contact with the medium, but not be covered by it.
4. Immediately transfer the culture inserts in a new 6-well plate containing fresh pre-warmed culture medium.
5. Replace the culture medium with 1 mL of pre-warmed fresh culture medium every 2-3 days until slices fixation.

6. Immunohistochemistry (Hands-on Time ≈ 1:30–2 h)

NOTE: Carry out the steps 6.1. to 6.3. under a fume hood. Avoid exposition to paraformaldehyde (PFA) and refer to the product safety datasheet for adequate manipulation and protection.

1. Use forceps to lift each culture inserts by holding the plastic edge and remove the culture medium beneath the membrane inserts from each well.
2. Fix the cerebellar slices for 30 min by adding 2 mL of 4% PFA in 1x PBS pH 7.4 onto the membrane inserts.
   NOTE: The time point of fixation depends on the stage of myelination studied: 4 DIV corresponds to the onset of myelination, 7 DIV to fully myelinated slices. In case of LPC treatment, 9 DIV corresponds to the peak of demyelination, 11 DIV to the onset of remyelination and 14 DIV to fully remyelinated slices.
3. Wash the inserts with the slices three times with 2 mL 1x PBS for 10 min.
4. Separate the slices from the membrane inserts under the binocular microscope using a 25x to 30x magnification, by gently pushing them using a scalpel or a brush. Alternatively, to avoid the risk of damaging the slices while detaching them from the membrane, use a scalpel to cut the membrane with the slices still attached.
5. Transfer the floating slices into the wells of a 4-well plate containing 1x PBS, using a brush.
6. Aspirate the PBS from each well and incubate the slices in pre-cooled 100% ethanol at -20 °C for 15 to 20 min. This step facilitates antibody penetration in myelinated fibers.
7. Aspirate the 100% ethanol and wash once briefly with 1x PBS, then twice in 1x PBS for 10 min at room temperature.
8. Aspirate the PBS and incubate the slices for 30 to 45 min with a solution containing 1x PBS, 5% NGS, 0.3% non-ionic detergent at room temperature to block non-specific antibody fixation sites.
9. Aspirate the blocking solution. No washing is necessary.
10. Incubate the slices overnight with the primary antibodies diluted in blocking solution at 4 °C. To visualize the myelin on cerebellar organotypic slices, use MBP (Chicken, 1/150 or Mouse IGg2b, Smi99, 1/200) or PLP (Rat, hybridoma, 1/5 to 1/10) antibodies.
    NOTE: To reveal the expression of more than one antigen in the same slice, perform a double- or triple staining procedure (see Figure 1 as an example). To perform the incubation simultaneously, ensure primary antibodies are produced in different species. Then, use their corresponding secondary antibodies coupled to different fluorophores (see Table of Materials for nodal and paranodal marker¹⁸ antibodies).
11. On the second day, wash the slices three times in 1x PBS for 10 min.
12. Incubate for 3h with the secondary antibodies diluted in blocking solution (1/500 dilution) in the dark at room temperature.
    NOTE: Optimization of antibody dilution and incubation time might be required when using other antibodies than the ones described here.
13. Wash the slices three times in 1x PBS for 10 min in the dark.
14. Under a binocular microscope, mount the slices onto a glass slide. Place 100 µL of 1x PBS on the slide and transfer the slices into the PBS using a brush. Unfold the slices if needed and flatten them on the slide. Remove any excess of PBS. In case the immunolabeling is performed with the slices still attached to membranes, mount with the slices facing the coverslip, with the membrane on the glass slide.
15. Place a drop of mounting medium directly onto the glass coverslip and gently cover the slices. Mounted slides can be stored for several months at 4 °C in the dark.