Assessing Anti-fungal Activity of Isolated Alveolar Macrophages by Confocal Microscopy

All procedures performed on animals in this study were approved by the Animal Care and Use Committee at Roswell Park Cancer Institute and complied with all state, federal, and National Institutes of Health regulations.

1. In vivo Macrophage Labeling

Note: PHK26 is a lipophilic dye that will be taken up by phagocytic cells both in circulation and in the tissues when administered intravenously (i.v.). PKH26 was used for in vivo labeling to minimize the amount of post-harvest manipulation of the macrophage. PKH-26 has the advantage of stably accumulating in alveolar macrophages over prolonged periods. Waiting 10 days after administration allows for clearance of all labeled cells from circulation leaving only alveolar macrophage stably labeled with PKH26^8,9. PKH26 is a red fluorochrome that has excitation (551 nm) and emission (567 nm) characteristics compatible with rhodamine or phycoerythrin detection systems. Any labeling procedure can introduce artifact, including loss of viability. However, since we use the same approach for labeling of macrophages in different mouse genotypes (e.g., WT vs. NADPH oxidase-deficient), the effects of these artifacts would be uniform.

1. Dilute stock solution of PKH26 (1 x 10^-3 M in ethanol) 1:5 in diluent C provided by manufacturer.
2. Load 100 µl of diluted PKH26 into a 1/2 ml insulin syringe (29 G x 1/2”) for each mouse.
3. Administer to mice by i.v. injection (either tail vein or retro orbital) at least 10 days prior to harvest.

2. Harvesting Alveolar Macrophages by Bronchoalveolar Lavage (BAL)

The number of mice to use for an experiment can vary. A typical yield from an unstimulated mouse is in the range of 3-5 x 10⁵ alveolar macrophages, however this number can vary greatly based on factors such as size of the mouse, experience of the person performing the lavage, and the number of instillations of lavage fluid and the volume withdrawn from lungs. In this protocol lung lavage is performed with a closed thoracic cavity which along with repeated 1 ml lavages helps to increase cell yield.

1. Euthanize mouse by administering a lethal dose of avertin anesthetic (12.5 mg) intraperitoneally (i.p.) using aseptic conditions to keep the solution sterile.
2. Spray mouse thoroughly with 70% ethanol.
3. Remove skin from the cranial end of animal exposing the thorax (Figure 1A).
   1. Make a small cut in skin just beneath the diaphragm.
   2. Insert finger into the cut and pull skin away from the cranial end of the animal (Figure 1A).
   3. Push up slightly on the head, extending the ventral side of the neck (Figure 1A).
4. Cut a small hole through the rib cage into the right side of the thorax to deflate the lungs and also allow visualization of lungs during the following steps.
5. Locate the trachea, and carefully dissect away surrounding tissue.
   1. Remove the salivary glands, sternohyoid muscles and larynx, etc. with curved forceps.
   2. Carefully lift with a curved forceps and cut away with scissors the thin tissue covering the trachea (adventitia) revealing the underlying ringed cartilage structure.
6. Insert catheter into trachea.
   1. Using curved forceps, place a suture behind (dorsal to) the trachea and pull a 10 cm length through the opening.
   2. Starting above the cricoid (the thickened ring of cartilage) carefully guide a 22 G catheter down the trachea stopping where the trachea disappears behind the sternum.
   3. Firmly tie suture around trachea and catheter for a snug fit. Remove needle from catheter.
7. Assemble 4-way stopcock.
   1. Fill one 6 ml syringe with DPBS, 1% FBS and attach to 4-way stopcock as indicated in Figure 1B.
   2. Attach an empty 6 ml syringe at the indicated position on the 4-way stopcock (Figure 1B).
   3. Attach open port on the 4-way stopcock to catheter (Figure 1B). Be careful to not dislodge catheter from trachea.
8. Collect lavage fluid from lungs.
   Note: Investigators may opt to use lower volumes of lavage fluid to minimize the possibility of damage to the tissue. It is important that the same approach be consistently applied in all mouse groups.
   1. Turn the handle on the stopcock so the empty syringe is closed and slowly inject ~1 ml DPBS, 1% FBS to inflate lungs. Observe lung inflation through the hole cut in step 2.4.
   2. Be careful NOT to overinflate the lungs.
   3. Turn the handle on the stopcock so that the full syringe is closed, and carefully withdraw the fluid from the lungs.
   4. Observe the lungs deflate through the hole cut in step 1.4. The catheter may need to be moved slightly back or forth to get a good draw. Take special precaution to keep the catheter within the trachea.
   5. Repeat steps 2.8.1-2.8.4 5-6x until all the DPBS, 1% FBS has been injected and withdrawn from the lungs. Note: recovery of lung fluid will not be 100% of injected fluid.
9. Empty syringe with lavage fluid into a 50 ml conical tube. Keep cells at room temperature.
10. Pellet cells by centrifuging at 300 x g for 5 min at room temperature and pour off supernatant. This can be saved for cytokine analysis if desired.
11. Resuspend cells in 1-5 ml RPMI 1640 with 10% FBS and count on a hemocytometer. In the unstimulated mouse, > 95% of cells harvested by BAL are expected to be macrophages, which can be confirmed by cytology.

3. Culturing and Harvesting Fungus

The Aspergillus fumigatus used in this protocol expresses GFP throughout all stages of its life cycle: spores, germlings and hyphae. This fungal strain was provided by Dr. Margo Moore, Simon Fraser University, Burnaby, BC, Canada.

1. Plate conidia of an Aspergillus fumigatus strain expressing GFP on Sabouraud brain heart infusion (BHI) agar slants with Chloramphenicol (0.05 g/L) and Gentamicin (0.5 g/L).
2. Incubate in the dark loosely capped for 7 to 10 days at room temperature.
3. Harvest conidia by washing slant with 10 ml of 0.01% Tween 20 in DPBS. Lightly scrape the fungus with the stripette to loosen. Rinse the slant several times to increase yield.
4. Pass conidial suspension through a 100 µm cell strainer into a 50 ml conical.
5. Pellet conidia by centrifuging at 400 x g for 10 min.
6. Resuspend conidia in 50 ml DPBS and count using a hemocytometer.
7. Wash twice with DPBS and dilute to desired concentration.

4. Fungal Challenge and Confocal Microscopy

To investigate the role of macrophage NADPH oxidase in host defense, the ability of isolated alveolar macrophages from mice of three genotypes to control the growth of phagocytosed Aspergillus spores was evaluated. The genotypes used in this study include; 1) wild-type mice, 2) Ncf1*/* Mn- mice with a naturally occurring Ncf1 mutation (Ncf1 encodes for p47^phox, a required component of the phagocyte NADPH oxidase) leaving them NADPH oxidase deficient, and 3) Ncf1*/* Mn+ transgenic mice (MN denotes the transgene harboring wildtype Ncf1 under the control of the human CD68 promoter) where NADPH oxidase activity has been reconstituted in the monocyte/macrophage population^10,11.

Confocal microscopy is best suited for this assay due to the presence and growth of non-phagocytosed conidia on the slides. The confocal microscope will allow for visualization of individual planes within the Z-axis, and enable differentiation between fungus growing around rather than inside macrophage. All confocal images are acquired using a 63X oil immersion lens. Z-stacks are acquired with an electronic zoom factor of 2.5. The thickness of a Z-stack is determined using DAPI and bright field images to locate the top and bottom of the field. Z-stacks ranged from 14 to 24 µm with individual slices ~1 µm thick. For quantification of macrophages, single scans of 1X electronic zoom were performed on 10 fields per genotype. Intact macrophages were identified based on PKH26 and DAPI staining of the membrane and nucleus respectively. Conidia were identified based on FITC expression and bright field image.

1. Prepare sterile coverslips and plates
   1. Under a Biological Safety Cabinet, soak 22 x 22 mm glass coverslips in 70% ethanol for 5-10 min.
   2. Rinse coverslips in sterile PBS.
   3. Using sterile forceps, gently place a single coverslip in the bottom of each well of a sterile singly packed 6-well plate.
2. Seed ~1 x 10⁶ harvested PKH26 labeled alveolar macrophages suspended in 300 µl RPMI 1640, 10% FBS onto each coverslip. The exact number may vary depending on the harvest yield.
3. Allow macrophage to adhere by incubating at 37 °C with 5% CO₂ overnight.
4. The following morning, add ~1 x 10⁷ GFP-expressing A. fumigatus conidia suspended in 100 µl pre-warmed (37 °C) RPMI 1640, 10% FBS.
5. Return plate to 37 °C incubator with 5% CO₂.
6. Cells should be fixed for a minimum of 2 hr at room temperature or overnight at 4 °C. At 3, 7, and 14 hr fix cells by adding an equal volume of 2% formaldehyde to each well (final concentration 1% formaldehyde). Place plates for the early time points at 4 °C overnight. For the final time point (14 hr), fix cells at room temperature for 2 hr.
7. After fixation, gently remove coverslips using forceps and rinse in DPBS.
8. Remove excess fluid by placing the edge of the coverslip on a folded Kimwipe.
9. Apply 6 µl fluorescence mounting medium with DAPI to a glass microscope slide.
10. Lay one edge of the coverslip on the slide and gently drop with cell side down into the mounting medium. Mounting medium will spread and form an even layer beneath the coverslip. There is no need to press on the coverslip.
11. Seal edges of coverslip with clear nail polish and allow to air dry.
12. Store prepared slides in a slide box (protected from light) at room temperature until ready to analyze by confocal microscopy.