Assessment of Leishmania Virulence within Cultured Macrophages

On day seven or eight after culture, use an aspirating tip fitted to a vacuum line to transfer four autoclave-sterilized 12-millimeter glass coverslips to the bottom of each well of a six-well culture plate without overlapping. Prepare identical plates for each time point of the experiment. Next, wash the bone marrow-derived macrophage cultures 2 times with fresh 37 degrees Celsius PBS and five milliliters of PBS without calcium and magnesium, supplemented with one millimolar EDTA to each plate.

After five minutes at 37 degrees Celsius, pool the detached cells in a 50-milliliter conical tube, rinsing each plate two times with an additional five milliliters of PBS without calcium and magnesium. After centrifugation, re-suspend the macrophages in five to 10 ml of fresh bone marrow-derived macrophage medium on ice. After counting, dilute the cells to 5 x 10⁵ per milliliter concentration in fresh bone marrow-derived macrophage medium and add two milliliters of cells per well to the 6-well plate. Then, incubate the cells overnight in the cell culture incubator.

To infect the cells with L amazonensis, dilute the parasite suspension to the appropriate experimental multiplicity of infection, and add 50 to 100 microliters of parasites to each well of all six-well plates, including all time points of the experiment. Incubate the bone marrow-derived macrophages for the appropriate infection period and temperature.

Then, wash the wells three times with two milliliters of fresh 37 degrees Celsius PBS without calcium or magnesium per well, and fix the initial time point samples with 1.5 to 2 ml of 2% paraformaldehyde for 10 minutes. For the plates corresponding to the later time points, add 2 milliliters of fresh bone marrow-derived macrophage medium to each well, and return the plates to the appropriate incubator. Then, fix, and wash the cells for each remaining time point as just demonstrated, and store the samples at 4 degrees Celsius in fresh PBS.

To stain the cells with DAPI, replace the supernatant with 1.5 ml of fresh PBS without calcium and magnesium, supplemented with 0.1% non-ionic detergent for 10 minutes at room temperature, followed by three 2-milliliter washes with PBS without calcium or magnesium. Next, stain the cells with two micrograms per milliliter of DAPI in 1 milliliter of PBS for one hour at room temperature. After three washes, place the coverslips cell-side down onto individual glass microscope slides mounted with a commercially available anti-fade mounting reagent.

Then, to quantify the number of infected cells, use the 100x immersion oil objective of a fluorescence microscope, and Emanuel counter to identify the number of smaller amastigote nuclei clustered around each large DAPI-stained macrophage nucleus.