This article details a protocol for immunostaining fixed cerebellar organoid sections to visualize cerebellar neurons. The method includes steps for blocking, permeabilization, and antibody incubation to ensure specific binding and accurate visualization of neuronal markers.
Take fixed cerebellar organoid sections comprising distinct types of cerebellar neurons.
Wash the sections in buffer to remove residual embedding media.
Incubate in glycine to block residual reactive sites from fixation, reducing non-specific binding during immunostaining.
Add a non-ionic detergent to permeabilize cells.
Wash with buffer, removing the excess detergent.
Now, dry the slide. Transfer it to an immunostaining dish with buffer-soaked paper to prevent tissue drying.
Apply a blocking solution that binds to non-specific sites on the neurons.
Remove the blocking solution. Incubate with primary antibodies, which bind to target antigens within cerebellar neurons.
Wash with buffer, removing unbound antibodies.
Incubate with fluorophore-bound secondary antibodies to bind antigen-bound primary antibodies.
Then, wash with buffer, removing unbound antibodies.
Add a dye to stain the cellular nuclei.
Using fluorescence microscopy, visualize markers expressed in cerebellar neurons, indicating organoid maturation.
Wash the microscope slides containing the organoid sections with 50 milliliters of 1X PBS for five minutes. Then, transfer the slides to a Coplin jar containing fresh 1X PBS. Transfer the slides to a Coplin jar containing 50 milliliters of freshly prepared glycine, and incubate for 10 minutes at room temperature.
Then, transfer the slides to a Coplin jar containing 50 milliliters of 0.1% Triton and permeabilize for 10 minutes at room temperature. Wash the slides twice with 1X PBS for five minutes each time. Prepare the immunostaining dish with three-millimeter paper soaked in 1X PBS.
Dry the slides with a tissue all around the slices and place them on the 3-millimeter paper. With a Pasteur pipette, cover each slide with 0.5 milliliters of blocking solution. After incubating for 30 minutes at room temperature, remove excess blocking solution, and dry the slides with a tissue all around the slices.
Place 50 microliters of diluted primary antibody on each slide, and cover them with coverslips. Place the slides in the previously prepared immunostaining dish and incubate them at 4 degrees Celsius.
After overnight incubation, transfer the slides to a Coplin jar with 50 milliliters of TBST, letting the coverslips fall off, then, wash the slides three times with TBST for five minutes each time. Place 50 microliters of diluted secondary antibody on each slide, and cover it with the coverslips.
Place the slides in the previously prepared immunostaining dish. Incubate for 30 minutes at room temperature protected from light. Transfer the slides to a Coplin jar again, and wash them three times with 50 milliliters of TBST for five minutes each time. Using a Pasteur pipette, add 0.5 milliliters of DAPI solution over the whole surface of each slide and incubate for five minutes at room temperature.
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