Real-Time Assessment of the Antifungal Activity of Primary Human Immune Cells

All procedures involving human participants have been performed in compliance with the institutional, national, and international guidelines for human welfare and have been reviewed by the local institutional review board.

1. A. fumigatus Culture and Conditions

1. Prepare glucose minimal agar medium as described.
   1. Adjust media to pH 6.5 using 1 M sodium hydroxide (NaOH) and autoclave at 120 °C for 20 min.
   2. Allow to cool to 65 °C.
   3. Working in a sterile flow hood, pour 30 mL of cooled media into a T75 flask and allow it to cool with the neck of the flask resting on a 25 mL pipette to create an agar slope.
2. Streak out the dsRed Af293-Aspergillus fumigatus strain on the agar and incubate for 7 days at 37 °C + 5% carbon dioxide (CO₂). Two flasks will yield approximately 10⁹ conidia in 5 mL phosphate-buffered saline (PBS) pre-biotinylation.

2. A. fumigatus Labeling and Preparation

NOTE: When performing this assay for the first time, prepare the parental Af293-A. fumigatus strain. Take samples of both strains in microcentrifuge tubes and label only one of each strain as described below. This will allow one to have control conidia with no color and conidia with a single color, either dsRed or Alexa Fluor 633 (AF633), to test the setup and equipment.

1. Harvest A. fumigatus conidia by immersing the culture with 30 mL PBS + 0.05% Tween-80. Filter the resulting suspension through a 40 µm cell strainer into a 50 mL tube to remove hyphal fragments.
2. Centrifuge at 805 x g for 10 min, remove the supernatant, and wash once in 20 mL of sterile PBS. Pool conidia from two plates of the same strain during the wash step.
3. Following the washing step, resuspend the pellet in 5 mL of PBS and transfer 1 mL aliquots of conidial suspension into microcentrifuge tubes. 1 mL of suspension of each strain is usually sufficient for live cell imaging. Wrap the remaining aliquots in foil and store at 4 °C.
4. Centrifuge aliquots of conidial suspension at 9,300 x g for 10 min at room temperature and carefully remove supernatant. Resuspend the conidial pellet in 1 mL 0.05 M sodium bicarbonate (NaHCO₃) pH 8.3.
5. Prepare 10 mg biotin/200 µL dimethyl sulfoxide (DMSO) stock solution by reconstituting 25 mg biotin in 500 µL DMSO. Add 10 µL biotin/DMSO stock solution to the microcentrifuge tube, cover it in aluminum foil, and incubate for 2 h on a rocker at 4 °C.
   1. Aliquot the remainder of the biotin/DMSO stock solution and freeze at -20 °C for use in subsequent experiments.
6. Centrifuge for 10 min at 9,300 x g and carefully remove the supernatant. Resuspend the conidial pellet in 1 mL of 100 mM Tris (hydroxymethyl) aminomethane-hydrochloride (Tris-HCl) pH 8.0 for 1 h to deactivate free-floating biotin.
7. Centrifuge for 10 min at 9,300 x g and carefully remove the supernatant. Wash the pellet twice with 1 mL of sterile PBS and resuspend the pellet in 1 mL of PBS.
8. Dissolve 1 mg of Streptavidin-AF633 in 0.5 mL of PBS to make a 2 mg/mL stock solution. Add 10 µL of 2 mg/mL streptavidin-AF633 per 1 mL of conidial suspension and incubate for 40 min at room temperature on a rocker covered in aluminum foil. The remaining Streptavidin-AF633 can be frozen in aliquots for use in subsequent experiments.
9. Centrifuge for 10 min at 9,300 x g and carefully remove supernatant. Resuspend the conidial pellet in 1 mL of PBS and count the conidia using a hemocytometer at a 1:1,000 dilution (1:100 and then 1:10). Adjust the conidial concentration to 3.6 x 10⁶/mL with CO₂-independent media, wrap in foil and store at 4 °C.
   NOTE: The conidia are now labeled with AF633 and will be referred to as FLuorescent Aspergillus REporter (FLARE) conidia.
10. Optional: If stimulation with swollen conidia is required, after counting conidia (step 2.9), dilute conidia to 7.2 x 10⁶/mL in yeast nitrogen base (YNB) medium and add 500 µL conidial suspension to 4.5 mL YNB medium in an autoclaved 50 mL Erlenmeyer flask. Cover and place the flask in a shaking incubator (200 rpm at 37 °C) for 6 h.
    1. Transfer the medium to a 15 mL tube. Centrifuge at 805 x g for 10 min at room temperature and wash once in PBS. Centrifuge again and resuspend pellet in 1 mL CO₂ independent medium, giving 3.6 x 10⁶ conidia/mL.
       NOTE: Conidia frequently clumps after they become swollen making accurate counting difficult, it is advised to calculate with the counted numbers of resting conidia used.

3. Isolation of Human Neutrophils and Monocytes

1. Draw 20 mL of venous blood from healthy volunteers into two 10 mL blood tubes containing ethylenediaminetetraacetic acid (EDTA). For each donor separately, pour 20 mL blood into a 50 mL tube and dilute with 15 mL PBS.
2. Underlay the blood with 15 mL of a lymphocyte isolation solution (density 1.077 g/mL) with a syringe and iron needle. Place the needle at the bottom of the tube and gently force the lymphocyte isolation solution out. Centrifuge at 630 x g for 20 min with no brake and low acceleration.
3. Prepare PBS, chilled on ice.        
   NOTE: Steps 3.4 and 3.5 must be followed simultaneously to generate monocytes (3.4) and neutrophils (3.5).
4. Using a pastette, harvest the peripheral blood mononuclear cells (PBMC) layer. This is the layer in the center between the yellow serum (upper) and the transparent lymphocyte isolation solution (lower) layer, and transfer into a fresh 50 mL tube.
   1. Fill the tube with the PBMC up to 50 mL with cold PBS and centrifuge at 582 x g for 10 min. Remove the supernatant and wash twice with cold PBS and resuspend in 1 mL of PBS per 20 mL of donor blood used initially.
   2. Make 1:10 dilutions for counting by adding 20 µL of the cell suspension to 160 µL of PBS and 20 µL of trypan blue. Count cells with a hemocytometer.
   3. Prepare a buffer solution by adding 26 mL of heat-inactivated fetal calf serum (FCS) and 2.1 mL of 0.5 M EDTA to 500 mL of Hank's balanced salt solution (HBSS). Centrifuge the cells for 10 min at 515 x g and resuspend in 40 µL of buffer solution per 1 x 10⁷ cells.
   4. Transfer the cell suspension to a 15 mL tube and add 10 µL of CD14 microbeads per 1 x 10⁷ cells, mix well, and incubate for 15 min at 4 °C, making sure to mix the tubes every 5 min.
   5. Wash cells by adding 1 mL of buffer solution per 1 x 10⁷ cells and centrifuge at 515 x g for 10 min.
   6. Aspirate supernatant completely and resuspend up to 1 x 10⁸ cells in 500 µL of buffer solution.
   7. Place 1 column per sample on a magnetic separator according to the manufacturer's instructions. First, wash the column once with 500 µL of buffer solution.
   8. Pipette the 500 µL of cell suspension through the column, wait for the column to dry, and then wash by repeating this three times with buffer solution.
   9. Place a new 15 mL tube underneath the column and take the column off the magnetic separator. Then flush the cells out of the column into the tube with 1 mL of buffer solution.
   10. Add 4 mL of buffer solution and centrifuge at 515 x g for 10 min, and then resuspend in 1 mL of Roswell Park Memorial Institute (RPMI) medium.
   11. Make 1:10 dilutions for counting by adding 20 µL of the cell suspension to 160 µL of PBS and 20 µL of trypan blue. Count the cells with a hemocytometer.
   12. Adjust the cell concentration to 6 x 10⁵/mL with RPMI and seed 200 µL on a 35 mm glass-based imaging dish. Incubate overnight at 37 °C with 5% CO₂.
   13. The next day, prior to imaging, remove the RPMI and add 200 µL of CO2-independent medium.
       NOTE: These monocytes can also be differentiated into various macrophage subsets prior to imaging. If this is a technique to be explored, an excellent starting point is a recent paper by Ohradanova-Repic et al.
5. After harvesting the PBMC layer, remove all of the serum and most of the lymphocyte isolation solution, but be careful not to remove the small white band on top of the red pellet as this will contain most of the neutrophils.
   1. Prepare a 10x hypotonic lysis buffer stock by adding 83 g of ammonium chloride (NH₄Cl) and 10 g of potassium bicarbonate (KHCO₃) in 1 L of sterile water. Store at 4 °C.
   2. Dilute this stock 10x with sterile water and add it to the tubes. Then, carefully invert 3 times and incubate in ice for 15 min.
   3. Centrifuge at 394 x g for 10 min and resuspend in hypotonic lysis buffer for 10 min in ice.
   4. Centrifuge at 394 x g and wash twice with cold PBS. Resuspend in 1 mL of CO₂-independent medium per donor.
   5. Make 1:10 dilutions for counting by adding 20 µL of the cell suspension to 160 µL of PBS and 20 µL of trypan blue. Count the cells with a hemocytometer.
   6. For microscopy, add 200 µL of the same cell suspension to a 35 mm glass-based imaging dish.   
      NOTE: Neutrophil imaging or flow cytometry should be initiated immediately due to their propensity to undergo apoptosis if left unstimulated. If this is not possible neutrophils should be kept on ice and used as soon as possible (within the same day).

4. Live Cell Video Microscopy

NOTE: The choice of the microscope will depend upon what is available locally, but the microscope setup will need to include an inverted stage, an environmental chamber heated to 37 °C and appropriate excitation/emission filters for dsRed (532/561 nm) and AF633 (633 nm). FLARE conidia do not work with the 488 nm laser in lieu of the 532/561 nm laser for dsRed. When performing this experiment for the first time, use the control conidia with no color and single color to test for background fluorescence and bleed-through between the dsRed and AF633 channels.

1. Turn on the microscope heater prior to the experiment and allow sufficient time for the environmental control chamber to warm to 37 °C. The time taken for chamber temperature to stabilize will vary for different microscope setups.
2. Turn on the microscope and computer, and load the imaging software. Mount the imaging dish on the microscope stage and adjust the focus to find the human neutrophils or monocytes.
3. For dsRed and AF633, set the laser power to 10% and exposure time to 1 s in the acquisition settings.
4. Remove the imaging slide and add 100 µL of 3 x 10⁶/mL FLARE conidial suspension to the appropriate wells (total volume 300 µL/well). Record the time that FLARE conidia are added to the dish.
5. Return the slide to the stage and adjust camera sensitivity for dsRed and AF633 to clearly see the conidia but not overexpose the image. Set up a stage points list if multi-point acquisition is required.
6. Commence imaging when all points are in focus, the channels are optimized and the cycle time and duration is set. Cycle time and duration of imaging depend on the experimental question. The goal is to keep the cycle time as short as possible (≤2 min) with 1 min allowing for in-depth analysis of uptake dynamics.