This article describes a method to study neuronal death in an ex vivo epilepsy model using organotypic slices from rat pup brains. The process involves propidium iodide staining and immunostaining to visualize neuronal death.
Epilepsy — spontaneous and recurrent seizures in the brain — leads to neuronal cell death.
To study neuronal death in an ex vivo epilepsy model, begin by taking a rhinal cortex-hippocampus organotypic slice — prepared from a rat pup brain — in an appropriate medium.
The organotypic slice consists of the rhinal cortex and hippocampus. Under gradual serum deprivation, these regions depict evolving epileptic events, leading to seizure-induced neuronal death.
Add the required amount of propidium iodide, PI — a fluorescent dye — to the culture and incubate. Through the damaged cell membrane, PI enters the dead neurons and binds to DNA; the live neurons, with intact membrane, are not capable of PI uptake.
Place the organotypic slice on a microscope slide for immunostaining. Add permeabilization-blocking solution and incubate.
Detergent molecules in the solution solubilize membrane lipids, creating pores in the membrane of intact cells — rendering them permeable for subsequent immunostaining steps. Proteins in blocking solution passively bind to non-specific sites of the cells, reducing background staining.
Add a primary antibody solution that penetrates inside the neurons and binds to the neuron-specific nuclear antigen — NeuN — distributed in the nuclear matrix. Add fluorescently-labeled secondary antibodies that bind to the primary antibodies.
Place mounting medium on the slide to stabilize the specimen and seal with a coverslip. Under a confocal microscope, cells showing dual staining — NeuN-stained neurons which are PI-positive — indicate neuronal death.
To perform the propidium iodide uptake assay, work in a biosafety cabinet.
Lift the insert from the well, and add propidium iodide in medium to a final concentration of 2 micromolar per well. Slowly agitate the plate before placing the insert back into the well, taking care that there are no bubbles beneath the slices. When all slices have been treated, return the plate to the cell culture incubator.
For immunostaining of the slices, at the end of the propidium iodide incubation, aspirate the medium from the bottom of each well, and add 1 milliliter of 4% paraformaldehyde to the top and bottom of each insert.
After one hour at room temperature, wash the slices 2 times for 10 minutes and 1 milliliter of PBS per wash, and use a hydrophobic pen to draw two rectangles on microscope slides.
Use a sharp blade to cut the slices from the inserts and place one slice into each rectangle. Add permeabilization/blocking solution onto each slice for a three-hour incubation at room temperature. At the end of the incubation, add the primary antibodies of interest to each slice for a 4-degree Celsius incubation overnight.
The next morning, wash the slices with PBS-Tween three times for 10 minutes per wash, and incubate the slices with the appropriate secondary antibodies for four hours at room temperature. Afterwards, wash the slices as demonstrated, and add 50 microliters of Hoechst solution to each slice for a 20-minute incubation at room temperature.
Following Hoechst incubation, wash the slices again, and add 50 microliters of mounting medium to each one. Then, place a coverslip over the slices and seal with nail polish.
After allowing the slides to dry for 24 hours at room temperature, visualize the immunostaining and propidium iodide uptake by confocal microscopy.
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