Western blot analysis

The PC-12 cells were plated at a density of 3×10⁴ cells/well in a 6-well plate placed on cover glass and were stabilized for 24 hours. The cells were incubated and treated according to the experimental schedule for 5 days. PC-12 cells were placed on ice and washed using PBS. The cells were dislodged using a cell scraper and pipetted into microcentrifuge tubes. They were centrifuged at 1,500 rpm for 5 minutes and the supernatant was discarded. Cells were lysed in mammalian tissue lysis/extraction reagent including protease inhibitor. Proteins were quantified using the bicinchoninic acid protein assay kit. Samples were prepared in 1×sodium dodecyl sulfate (SDS) sample buffer (50 mM Tris pH 6.8, 2% SDS, 10% glycerol, 50 mM dithiothreitol, and 0.01% bromophenol blue). Proteins were separated in 12% SDS-PAGE and transferred onto a polyvinylidene fluoride membrane. After blocking the membrane with 5% skim milk in Tris-buffered saline and Tween 20 at room temperature for 60 minutes, the membrane was incubated and immunoblotted with anti-caspase-3 (1:1,000), anti-cleaved caspase-3 (1:1,000), anti-Bcl-2 (1:1,000), anti-Akt (1:1,000), anti-phospho-Akt (Ser473, 1:1,000), and anti-α-tubulin (1:1,000) at 4℃ overnight. The biotinylated anti-mouse or anti-rabbit immunoglobulin G secondary antibodies were conjugated and were applied for 1 hour at room temperature. The membrane was developed by the alkaline phosphatase-conjugated development kit (Bio-Rad, Hercules, CA, USA). Developed protein bands were quantified by Multi Gauge version 2.2 program (Fuji photo film, Tokyo, Japan). α-Tubulin served as an internal control.