Detection of Inflammasome Activation via Fluorescence Microscopy

All procedures involving animal models have been reviewed by the local institutional animal care committee and the JoVE veterinary review board.

1. Harvest of Bone Marrow

1. Aseptically remove the femur and tibia from a mouse and clean the bones using sterile instruments. Place the cleaned bones in Dulbecco Modified Eagle Medium (DMEM) + 5 mM N-2-hydroxyethylpiperazine-N'-2-ethanesulfonic acid (HEPES) + 0.2 mg/mL L-glutamine + 0.05 mM 2-mercaptoethanol + 50 mg/mL gentamicin sulfate + 10,000 U/mL penicillin/streptomycin + 10% fetal bovine serum (FBS, DMEM-10 complete) and incubate the tube on ice for 15 min.
2. Remove the proximal and distal ball joints using scissors. Insert a 25 G needle attached to a 10 mL syringe filled with medium into the hollow of the bones. Flush out the bone marrow using DMEM-10 complete. Resuspend any clumps by pipetting the medium up and down gently.
3. Centrifuge the cells for 10 min at 300 x g and count the number of cells using a hemocytometer. At this point, either freeze the cells using 90% FBS + 10% dimethyl sulfoxide (DMSO) or plate the cells in a 15 cm Petri dish to differentiate macrophages using L929 conditioned medium (step 2.1).

2. Differentiation of Bone Marrow-derived Macrophages

1. Using pre-warmed differentiation medium (21 mL of DMEM-10 complete and 9 mL of L929 cell-conditioned medium as described by Swanson et al.), put bone marrow cells in a 15 cm non-tissue culture-treated Petri dish (1.2-1.5 x 10⁷ cells). Incubate the plate at 37 °C and 5% carbon dioxide, or CO₂ for 7 days. Add an additional 30 mL of differentiation medium to the plate on day 3 or 4.  
   NOTE: It is important not to use tissue culture-treated plates, as macrophages will be difficult to detach and harvest.
2. To collect macrophages, wash the plate with phosphate-buffered saline (PBS) and add 20 mL of cold PBS + 1 mM ethylenediaminetetraacetic acid (EDTA). Incubate the plate at 4 °C for 10 min. Harvest the cells by pipetting the PBS + EDTA over the cells and washing the plate with 10 mL of PBS. Combine both washes into two 15 mL conical tubes.
3. Centrifuge the cells at 300 x g for 10 min and resuspend the cells in 10 mL of phenol-red free DMEM + 5 mM HEPES + 0.2 mg/mL L-glutamine + 0.05 mM 2-mercaptoethanol + 5% FBS (DMEM-5) and count the cells using either a hemocytometer or a Coulter counter. Keep the cells on ice during the counting.
4. Seed the macrophages in tissue culture-coated plates at 2 x 10⁵ cells/mL. For microscopy experiments, seed 1 mL of cells on coverslips in 24 well plates. For lactate dehydrogenase (LDH) release assays, seed 100 µL of cells in a 96-well plate. For the LDH assay, seed 9 wells (3 wells untreated, 3 wells with 100% lysis controls, and 3 wells for the experimental stimulus).
5. Centrifuge the 96-well plate for 5 min at 300 x g to make certain the cells are equally distributed across the well.
6. Incubate the plate overnight at 37 °C + 5% CO₂ before starting the priming process.

3. Priming

1. Replace the medium with fresh medium (DMEM-5) containing 100 ng/mL lipopolysaccharide or LPS (500 μL for the 24-well plate or 50 μL for the 96-well plate).
   NOTE: There are a variety of structural variations in LPS, which affect the ability to stimulate TLR4-mediated priming. LPS from Salmonella minnesota R595 (Re) is available from multiple vendors and is recommended.
2. Incubate the plate for 3 h at 37 °C and 5% CO₂.

4. Activation of the NLRP3 Inflammasome with Nigericin and Labeling with FAM-YVAD-FMK

1. Remove the medium and replace it with 290 μL of DMEM-5 containing 5 μM nigericin and 5 mM glycine. Glycine is added to reduce the amount of cell lysis that occurs during caspase-1 activation. Incubate for 60 min at 37 °C and 5% CO₂. Use both wild-type and caspase-1 deficient macrophages to ensure that the observed labeling is specific for caspase-1.
2. During the last 45 minutes, add 10 μL of 30x FAM-YVAD-FMK, prepared according to the manufacturer's instructions.
3. After 60 min, remove the medium and wash the cells three times for 5 min each with 1 mL of cold PBS.
4. Add 250 μL of 2% paraformaldehyde to each well and incubate on ice covered with aluminum foil for 30 min.
5. During the last 5 minutes, add 4 μL of 0.2 mM far-red fluorescent nucleic acid stain to label the nuclei. 4',6-diamidino-2-phenylindole (DAPI), or other dyes can also be used to label DNA.
6. Wash the cells three times with 1 mL of cold PBS and mount the coverslips onto microscope slides using a 7 μL anti-fade mounting medium. Let the mounting medium harden overnight before sealing the coverslip to the microscope slide using nail polish.
7. Image the macrophages by confocal microscopy. Ex/Em for FAM-YVAD-FMK is 492/520nm and 642/661 for far-red fluorescent nucleic acid stain. Make sure to set up the microscope such that untreated cells do not show any background staining in the 488 channel. Otherwise, background FAM-YVAD-FMK staining will be detected and not actual active caspase-1.

5. Imaging of Labeled Macrophages by Confocal Microscopy

NOTE: The stained cells can be viewed after the coverslips have been allowed to cure overnight and have been sealed to the microscope slide with nail polish. Here, imaging using a confocal microscope is described. Cells can also be viewed by standard fluorescence microscopy.

1. Place the slide with the untreated control cells on the microscope stage and focus the microscope on the cells.
2. Using the microscope Look Up Table (LUT) settings, adjust the offset such that there is no positive FAM-YVAD-FMK staining in the untreated cells. This process can also be performed using caspase-1 deficient macrophages.
   NOTE: The offset has to be adjusted for each channel used in the experiment, but setting the correct offset is most critical for the FAM-YVAD-FMK channel and fluorescent antibody channels. The offset for the DNA channel is less critical. Once the offset has been set, do not adjust this setting for the rest of the experiment.
3. Switch the slide to the nigericin-treated sample. Find a field that contains both cells that are positive and negative for FAM-YVAD-FMK staining. Adjust the imaging plane of the FAM-YVAD-FMK channel to the plane that has the highest intensity of positive staining. This will normally be the plane where the center of the inflammasome focus is imaged.
4. Adjust the gain such that foci are visible in cells that have condensed nuclei. After setting both gain and offset, leave these settings for the rest of the imaging session.
   NOTE: One easy way to determine if a cell is pyroptotic is by looking at nuclear morphology. Cells with active caspase-1 have rounded and condensed nuclei, while nucleoli are visible in cells without caspase-1 activation and the nuclear morphology is similar to those of untreated cells.
5. Collect images of 5 randomly selected fields at 100X total magnification. This will normally result in about 40 cells per image. Repeat this process for each sample.
6. Count the number of cells in each image using image analysis software. Then count the number of cells that have positive FAM-YVAD-FMK staining. Represent the level of caspase-1 activation as the percentage of FAM-YVAD-FMK positive cells.