1. Cholesterol Depletion of J774A.1 Cells with MβCD

1. In a sterile biosafety cabinet, seed 10⁵ J774A.1 macrophage-like cells per well on a 96-well cell culture plate in 200 μl of Dulbecco’s Modified Eagle Medium (DMEM) supplemented with 10% fetal bovine serum (FBS) and 1% penicillin/streptomycin (P/S). Incubate at 37 °C and 5% CO₂ O/N.
2. Remove media from the cell monolayer and wash the cells twice with 1x phosphate buffered saline (PBS) that has been filtered or autoclaved.
3. Add 200 μl of MβCD solution at the desired concentration (10 mM or 30 mM in PBS) or 1x PBS as a control and incubate for 30 min at 37 °C with shaking. Remove supernatant and reserve at RT for quantitative analysis with commercially available kit immediately following the procedure.
4. Wash the cells two to three times with 1x PBS or serum free DMEM and continue with infection or lyse cells by pipetting two to three times with deionized H₂O for analysis with thin layer chromatography or with a kit.
   NOTE: Following cholesterol depletion a commercially available cholesterol quantification kit can be used. See materials section for details. Follow manufacturer’s instructions as written.

2. Observation of Cholesterol Content by Thin Layer Chromatography (TLC)

1. Wash a TLC tank twice with acetone and once with a solution of petroleum ether:diethyl ether:acetic acid (65:30:1 by volume). Saturate the tank with the petroleum ether:diethyl ether:acetic acid (65:30:1 by volume) solution and leave O/N.
   NOTE: Organic solvents should always be used under a fume hood to prevent inhalation of vapor. Gloves and lab coat should be worn at all times. Acetic acid is a strong acid and should be used with caution.
2. In a sterile biosafety cabinet, seed 10⁶ J774A.1 macrophage-like cells per well on a 6-well cell culture plate in a volume of 5 ml of warm DMEM supplemented with 10% FBS and 1% P/S. Incubate at 37 °C, 5% CO₂ O/N.
3. Deplete macrophages of cholesterol by following steps 1.2 – 1.4, substituting 1 ml for 200 μl where applicable to account for the larger well size.
4. Add 500 μl of Trypsin-EDTA to each well, incubate for 3 min at 37 °C, and gently scrape cells with a cell scraper.
5. Transfer into a microfuge tube and add an additional 500 μl of warm DMEM supplemented with 10% FBS and 1% P/S.
6. Spin the cells for 5 min at 300 x g and remove the supernatant.
7. Add an additional 500 μl of warm DMEM supplemented with 10% FBS and 1% P/S to the cell pellet and resuspend carefully by pipetting up and down.
8. Remove 10 μl of cells and count cells on a hemocytometer. Normalize the cell concentrations from each sample and add an equal number of cells to glass tubes.
   NOTE: At this step, one may choose to combine cells from the same treatment groups to obtain a more concentrated final lipid extract.
9. Centrifuge cells at 300 x g for 5 min at RT and remove media.
10. Add 2 ml of methanol and vortex. Then, add 1 ml of chloroform and vortex. Check the phase status to make sure the solution in monophasic.
    NOTE: Tubes can be stored at 4 °C O/N.
11. Centrifuge at 1,700 x g for 10 min at RT and transfer the supernatant to a new tube
12. Add an additional 1 ml of chloroform followed by 1 ml of dH₂O. Vortex twice for 30 sec. Centrifuge at 1,700 x g for 5 min at RT.
13. Weigh a glass tube on a sensitive balance and use a glass Pasteur pipette to transfer the lower phase into the glass tube of known weight.
14. Dry down lipids in a centrifugal evaporator until dry (approximately 2 hr). Weigh tube with dried lipids and calculate the dry lipid weight.
    NOTE: Dry lipids can be stored in -20 °C until ready to perform TLC.
15. Dilute dried lipids in enough chloroform to normalize the concentration of lipid (usually 20 – 50 µl) and load 20 µl of the diluted lipid on a silica TLC plate. Load 20 μg of cholesterol diluted in 20 μl of chloroform as a standard.
16. Add dried TLC plate to the saturated TLC tank and allow solvent to migrate up to 1 cm before plate edge. Remove TLC plate from tank and allow it to dry about 5 min.
17. Visualize lipids by placing in an iodine vapor tank to check migration. Remove and allow spots to fade for about 10 – 15 min under the hood.
    NOTE: Iodine is an inhalation hazard. Always use under a fume hood.
18. Prepare a solution for neutral lipid staining by combining 60 ml of methanol with 60 ml of deionized H₂O, 4 ml of sulfuric acid, and 630 mg of manganese chloride.
19. Carefully and slowly dip the TLC plate into the neutral lipid staining solution in a tray and remove without sloughing off the silica layer.
    NOTE: The neutral lipid staining solution can be reused several times and so it can be retrieved from the tray and placed back in a bottle for later use.
20. Allow plate to dry under the hood at RT until all bubbles has disappeared. Heat the TLC plate on a heat block set to 160 °C and char to the desired color.
    NOTE: A densitometry program such as Vision Works LS can be used to quantify the charred bands to compare lipid samples.

3. Infection of Macrophages with C. neoformans (H99)

1. In a sterile biosafety cabinet, seed 10⁵ macrophage-like cells per well on a 96-well cell culture plate in 200 μl of DMEM supplemented with 10% FBS and 1% P/S. Incubate at 37 °C, 5% CO_2, 5% CO₂ O/N.
   NOTE: Infection can also be done in glass bottomed confocal dishes for easier imaging; all amounts remain the same.
2. Grow a culture of C. neoformans (H99) by inoculating 10 ml of YNB with one colony obtained from a struck plate and incubating it O/N at 30 °C with shaking.
3. Wash and count C. neoformans (H99) cells.
   1. Centrifuge C. neoformans O/N culture at 1,700 x g for 10 min at 4 °C.
   2. Remove media and discard. Wash cells with 5 ml of 1x PBS. Centrifuge at 1,700 x g for 10 min at 4 °C.
   3. Remove PBS and wash with 5 ml of filtered 1x PBS. Centrifuge at 1,700 x g for 10 min at 4 °C. Repeat this step 2 more times.
   4. Remove PBS and resuspend in 5 ml of 1x PBS.
   5. Make a serial dilution in PBS to obtain a 1:500 dilution of the washed culture.
      1. Add 100 μl of the original sample to 900 μl of 1x PBS to obtain a 1:10 dilution.
      2. Add 100 μl of 1:10 diluted sample to 900 μl of 1x PBS to obtain a 1:100 dilution.
      3. Add 200 μl of the 1:100 diluted sample to 800 μl of 1x PBS to obtain a 1:500 dilution
   6. Take 10 μl of 1:500 dilution and count on hemocytometer to calculate the number of cells.
4. Prepare working solution for activating macrophages and opsonizing C. neoformans.
   1. Dilute LPS and IFNγ 100x from stock solutions by adding 10 μl to 990 μl.
      NOTE: LPS and IFNγ are used to enhance phagocytic uptake but are not required for phagocytosis. If there is an interest in fungicidal activity of macrophages, perform activation O/N at 37 °C with shaking prior to infection.
   2. Per sample combine 7.5 μl of diluted LPS, 1.25 μl of diluted IFNγ, 1.25 μl of GXM antibody and the volume of the C. neoformans culture that gives 1.25 x 10⁵ cells. Bring volume up to 250 μl multiplied by number of samples with DMEM supplemented with 10% FBS and 1% P/S.
   3. Vortex and incubate solution for 20 min at 37 °C with shaking.
      NOTE: Cholesterol depletion (steps 1.2 – 1.4) can be done concurrently with the opsonization step. Be sure to treat macrophages prior to combining the working solution, as the opsonized cells should optimally be used no longer than 20 min following the step 3.4.3 incubation.
5. Infect Macrophages
   1. Wash macrophages twice with serum free DMEM and add 200 μl of opsonized C. neoformans working solution to each well.
   2. Incubate for 2 hr at 37 °C.
6. Fix and Stain Cells
   1. Remove media and wash cells 2 times with DMEM.
   2. Air dry the cell monolayer for 10 min and add 200 μl of ice cold methanol to fix the cells.
   3. Incubate for 15 min at RT and remove any remaining methanol.
   4. Add 200 μl of 10x Giemsa and incubate for 5 min at RT.
   5. Wash 2 – 3 times with deionized water and dry O/N with cap off.
      NOTE: Imaging can be done the next day or up to a week following staining.
7. Visualize and Count
   1. Using a microscope, count 300 cells per data point (if there are 2 of the same treatment count 150 per well) and note the number of infected macrophages and number of engulfed Cryptococcus cells.
      NOTE: To ensure even sampling per data point use 2 plates per condition, choose 3 non-overlapping areas per plate and count 50 cells per area.
   2. Calculate phagocytic index by multiplying the percentage of infected macrophages by the mean number of C. neoformans per macrophage. Normalize phagocytic index by expressing values as a percentage of the 1x PBS treated control. After several trials calculate the mean value and the standard deviation of the mean to determine trends in phagocytic index. Use student t-test to determine significance.
   3. Take micrographs of cells at 1,000X or 400X magnification.

4. Trypan Blue Assay

1. In a sterile biosafety cabinet, seed 10⁶ macrophage-like cells per well on a 6-well cell culture plate in a volume of 5 ml of Dulbecco’s minimal essential medium (DMEM) supplemented with 10% fetal bovine serum and 1% penicillin/streptomycin. Incubate at 37 °C, 5% CO₂ O/N.
2. Deplete macrophages of cholesterol by following steps 1.2 – 1.4 substituting 2 ml for 200 μl where applicable to account for the larger well size.
   NOTE: Be sure to include a control treated with 1x PBS as well as a control that is scraped prior to any treatment.
3. Add 500 μl of 1x PBS to each well and gently scrape cells with a cell scraper. Transfer into a microfuge tube and suspend the cells by gently pipetting up and down.
4. Remove 10 μl of cells and stain with 1 μl of 4% Trypan blue.
5. Count cells on a hemocytometer and calculate viability using the following equation: % viability = [1 – (Blue cells / total cells)] x 100. Normalize values to the control that was untreated. After several trials calculate the mean value and the standard deviation to determine trends in viability. Use student t-test to determine significance.