To begin this procedure, transfer 400 microliters of cell suspension to a pre-chilled cuvette, then, add 20 micrograms of selected protein to reach a final concentration of 50 micrograms per milliliter. Flick the cuvette gently, about 10 times for mixing and to avoid damaging the cells. After that, dry the outside of the cuvette with a paper towel to avoid electrical arcing in the electroporator.
Place the sample in the electroporator and set it to 0.3 kilovolts for 1.5 to 1.7 milliseconds. Immediately after electroporation, flick the cuvette gently, about 10 times, to mix the sample thoroughly before proceeding to either fixation and immunofluorescence staining or affinity purification.
After four hours of recovery from electroporation, wash the cells with sterile PBS. Then, fix the cells in 100% methanol for two minutes at room temperature. Subsequently, wash the cells with sterile PBS three more times. Permeabilize the cells with 0.4% Triton X-100 in PBS for 15 minutes. Adjust the length of permeabilization and the strength of Triton X-100 according to the epitope and location of the target protein. Then, block the cells with 5% BSA in PBS for one hour at room temperature.
Afterward, wash them three times with PBS. Next, incubate them with the primary antibody in the antibody-binding solution overnight, at 4 degrees Celsius with gentle rocking.
The next day, wash the cells four more times with PBS. Then, incubate them with the appropriate fluorescently-conjugated secondary antibody in the antibody-binding solution. Add other stains as needed, such as 5 micromolar Wheat Germ Agglutinin, or WGA, conjugated to Alexa 647 or DAPI for one hour at room temperature and protected from light. Then, wash the cells with PBS five times and store at 4 degrees Celsius, protected from light until they are ready to be imaged.
