Brain tissue sample preparation and Western blotting

The right brain hemisphere was weighed and suspended in lysis buffer (25 mM Tris-HCl pH 6.8, 1% SDS, 5% glycerol, and 200 mM DTT), at a ratio of 1 ml of lysis buffer per 100 mg of tissue. The suspension was sonicated for approximately 30 s and centrifuged at 12000 ×g for 5 min, and the supernatant was then transferred to a fresh tube. Total protein was determined with the Bradford Protein Assay Kit (Pierce), and samples (50 μg) were analyzed by immunoblot as described previously [60]. Briefly, the samples were separated on a 8% SDS polyacrylamide gels and transferred to a PVDF membrane, blocked in 5% skim milk, and incubated overnight at 4°C with primary antibody (mouse anti-dynamin 1 antibody, 1:1000; rabbit anti-PSD-95 antibody, 1:1000; mouse anti-synaptotagmin antibody, 1:1000; mouse anti-synaptophysin antibody, 1:1000; anti-β-actin antibody, 1:1000) followed by incubation with secondary antibody. The results were visualized using ECL reagents (Pierce) and quantified using an image analyzer (Gel Doc 2000, Bio-Rad). The values were finally normalized to β-actin. For cultured cells, after treatments, the cells were washed with cold PBS and harvested with a cell scraper and sonicated for approximately 30 s in the cell lysis buffer, and the supernatant was collected by centrifugation at 12000 ×g for 5 min at 4°C. The total protein of each sample was determined using the Bradford assay, and samples (30 μg) were analyzed by immunoblotting as mentioned above.