After preparing human macrophages and apoptotic Jurkat cells, use a hemocytometer to count the apoptotic cells. Then, transfer a sufficient quantity of apoptotic cells to a 1.5-milliliter microcentrifuge tube.
Pellet the Jurkat cells via centrifugation at 500 times g for 5 minutes. Then, resuspend the cells in 500 microliters of PBS. While the Jurkat cells are in the centrifuge, aliquot 10 microliters of DMSO into a new microcentrifuge tube.
Next, dissolve a minimal amount of NHS-biotin in the DMSO. After this, transfer 500 microliters of the apoptotic cell and PBS suspension to the tube containing DMSO and NHS-biotin. Then, dilute an appropriate cell-tracking dye according to the manufacturer's instructions.
Incubate the cell suspension at room temperature for 20 minutes in the dark. Then, add an equal volume of RPMI 1640 with 10% FBS, and incubate the suspension at room temperature in the dark for an additional 5 minutes.
After this, pellet the cells via centrifugation at 500 times g for 5 minutes. Then, discard the supernatant and resuspend the stained cells in 100 microliters of RPMI 1640 with 10% FBS. Add 100 microliters of stained cell suspension drop-wise to each well of macrophages. Then, centrifuge the plate at 200 times g for 1 minute. Incubate the plate at 37 degrees Celsius with 5% CO2 in a tissue culture incubator.
After the appropriate incubation period, remove the cells from the incubator. Then, wash the cells twice with 1 milliliter of PBS. After this, add diluted FITC-conjugated streptavidin to each well and incubate the plate for 20 minutes in the dark.
Wash the cells three times with 1 milliliter of PBS, and gently shake the samples for 5 minutes after each rinse. Then, fix the cells with 4% paraformaldehyde in PBS for 20 minutes at room temperature. Rinse the cells once with PBS to remove any excess PFA.
Mount coverslips on an imaging slide, and transfer the slide to a fluorescence microscope. Capture z-stacks of a sufficient number of cells for quantification.
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