Analyzing Purkinje Cell Survival in Organotypic Cerebellar Slice Cultures

To harvest the cerebellum, use straight dressing forceps to grasp the pup head by the nose, and use straight eye scissors to cut open the scalp from the posterior end laterally to the midline. Cut open the skull in the same manner, pointing the scissor tips outward to avoid damaging the cerebellum, and use a spatula to transfer the brain to a 60-millimeter dish containing cold HBSS and 5 milligrams per milliliter of glucose.

Use the straight dressing forceps to carefully dissect out the cerebellum, and use sterile curved fine forceps to place the tissue onto a plastic disk on the cutting table of a tissue chopper. Rotate the cutting table to orient the tissue to allow the acquisition of parasagittal sections, and pull the table release knob to the right to move the cutting table until the blade is positioned at the edge of the tissue. Then, adjust the slice thickness to 350 micrometers and the blade speed to medium and start the chopper. When the whole cerebellum has been sliced, turn off the chopper.

Using sterile forceps, hold the disk over a new 60-millimeter dish. Use a transfer pipette to flush HBSS plus glucose over the disk so that the slices fall into the dish. Then, touching the samples as minimally as possible, use a microprobe to separate the slices.

Use the transfer pipette to select the sections of the cerebellum close to the vermis onto individual cell culture inserts in the six-well plate, and use the microprobe to position the slices into the center of each insert. When all of the slices have been placed, carefully aspirate any excess HBSS plus glucose and return the plate to the cell culture incubator.

For immunofluorescence staining, remove the supernatant from each well and wash the inserts with PBS. Fix the slices with 1 milliliter of cold 4% paraformaldehyde in the well of each insert, and 500 microliters on top of each insert for 1 hour.

At the end of the fixation, wash the inserts four times for 10 minutes with 1 milliliter of fresh PBS under each insert and 500 microliters of fresh PBS on top of each insert per wash on an orbital shaker. Prefill each well of a 24-well plate with 500 microliters of PBS-TB, and use a paintbrush to transfer the cerebellar slices from each cell culture insert into individual wells of a 24-well plate.

Permeabilize and block the slices at room temperature for one hour. At the end of the incubation, label the cells with 200 microliters of the primary antibody of interest diluted in PBS-TB per well overnight at 4 degrees Celsius on the orbital shaker.

The next morning, wash the slices four times for 10 minutes and 500 microliters of fresh PBS per wash on the shaker before labeling the samples with the appropriate fluorophore-conjugated secondary antibodies for two hours at room temperature on the shaker protected from light.

At the end of the incubation, counter-stain the sections with 500 microliters of an appropriate nuclear stain per well for 10 minutes at room temperature protected from light, and use a transfer pipette to mount the slices onto glass slides.

Then, let the sections air-dry completely before rehydrating with PBS and mounting the tissues with coverslips coated with approximately 80 microliters of mounting medium. Once the mounting medium has cured, the cerebellar sections are ready to be imaged.