Detection of Inflammasome Activation via Fluorescence Microscopy

Begin by replacing the medium from LPS-primed bone marrow-derived macrophages seeded on glass coverslips with 290 microliters of DMEM-5 medium supplemented with 5-micromolar nigericin and 5-millimolar glycine per well. Return the 24-well plate to the cell culture incubator for 60 minutes at 37 degrees Celsius and 5% carbon dioxide, adding 10 microliters of 30X FAM-YVAD-FMK after the first 15 minutes.

At the end of the incubation, wash the cells with 1 milliliter of cold PBS per well three times for 5 minutes per wash, and fix the cells with 250 microliters of 2% paraformaldehyde per well for 30 minutes on ice, protected from light, labeling the cells with an appropriate fluorescent nuclear dye during the last five minutes.

At the end of the incubation, wash the cells three times with cold PBS as just demonstrated, and load each slide with 7 microliters of anti-fade mounting medium. Place a coverslip onto each slide, and allow the mounting medium to harden overnight before sealing the coverslips with nail polish.

To image the cells by fluorescence confocal microscopy, place the untreated control slide onto the microscope stage, and manually focus on the cells. Using the microscope lookup table settings, adjust the offset until no positive probe standing is observed in the untreated cells.

Next, using a nigericin-treated sample, locate a field that contains cells that are both positive and negative for the probe staining, and adjust the imaging plane of the probe channel to the plane that has the highest intensity of positive staining. Then, adjust the gain until the foci are visible in the cells with condensed nuclei, and obtain images of five randomly selected fields per slide at a 100X total magnification.