Fixation and Staining of Mouse Brain Sections

Mice were sacrificed followed by perfused with ice‐cold phosphate buffer saline (PBS), the brains were isolated and post‐fixed in 4% paraformaldehyde (PFA), then transferred brains into 30% sucrose in PBS solution to make them dehydration until they were totally saturated. Put prepared brains in a freezing microtome (Leica, German) to obtain 15 µm sequential coronal sections, subsequently, the sections were moved onto the glass slides. After permeabilized with 0.3% Triton X‐100 in PBS and blocked with 10% goat serum (BOSTER Biological Technology, China) for 30 min at room temperature, the brain sections were collected to incubate with targeted antibodies according to the previously published procedures.^[60 (link)
^] When the in vitro cultured cells were used for immunofluorescence assays, cells on coverslips were fixed with 4% paraformaldehyde, permeabilized in 0.3% Triton X‐100, and blocked with 10% goat serum. Cells were then incubated with primary antibody of ADAM10 (Abcam) for 4 °C overnight, stained with goat anti‐rabbit secondary antibody Alexa Fluor 488 (Beyotime, China) for 60 min at room temperature. Then, coverslips were mounted with DAPI Fluoromount‐G (Southern Biotech, Birmingham, Alabama, USA). Images were captured by laser scanning confocal microscope (Leica TCS SP8 X, Germany) and quantified by Image‐J software (National Institutes of Health, USA).