Histological Quantification to Determine Lung Fungal Burden in Experimental Aspergillosis

All animal procedures were approved by the Animal Care and Use Committee of Indiana State University, the host campus of Indiana University School of Medicine — Terre Haute.

1. Preparation of A. fumigatus Conidia for Infection

1. While working in a well-ventilated fume hood, add 125 µL of A. fumigatus 293 stock solution to 900 µL cell culture grade water (CCGW). Then transfer the solution to malt extract agar (MEA) plate via pipette and spread evenly with an inoculating loop.
2. Store the fungal plate in the dark at 24 °C for at least 14 days and no more than 28 days.
3. Harvest conidia using glass beads as previously described⁹.
4. Pour 1.5 g of 0.5 mm glass beads onto the plate and gently tilt it back and forth until the beads are coated in order to extract conidia. Collect the bead/conidia mixture in a 15-mL conical tube. Suspend in 5 mL Dulbecco's phosphate-buffered saline (DPBS) and vortex.
5. Count the conidia in a 50x dilution of supernatant in DPBS using a hemocytometer. Count the number of conidia in the area shown in Figure 1 and use Equation 1 to calculate the concentration.
   #Conidia × Dilution Factor × 10⁴=Conidia/mL (Equation 1)
   NOTE: Additional 4 x 4 square corner areas may be counted, with the mean of these areas used as #Conidia in Equation 1.

Figure 1: Hemocytometer representative area used for conidia counting (box, arrow).

6. Dilute the conidia suspension with sterile saline (DPBS) to obtain the desired concentration, 1.0 x 10⁸ conidia/mL for infection with 5 x 10⁶ conidia/50 µL. Prepare 50 µL of conidia solution per 'n + 2' mice to be infected to account for pipetting error.
   NOTE: Standard fungus harvest and preparation/filtration may be alternatively used¹⁰. The glass bead method is best used to ensure that conidia samples with minimal small/soluble hyphal antigens are obtained for aspiration.

2. Immunosuppression and Infection of Murine Model

1. Use 7-10-week-old mice.
   NOTE: BALB/c, C57BL/6 (B6), eosinophil-deficient (ΔdblGATA, BALB/c background), and γδ T cell-deficient (TCRδ-/-, B6 background) mice were used for this study. For each experiment, age and gender-matched (both males and females) mice were used for each group.
2. Deplete neutrophils via intraperitoneal (IP) injection of 0.1 mL (0.5 mg) anti-mouse-Ly-6G antibody in sterile saline at 45° in the lateral lower right quadrant (For an experimental schedule, see Figure 2).
3. 24 h later, infect mice by aspiration with 5.0 x 10⁶A. fumigatus conidia suspended in 50 µL sterile saline (see steps 2.4-2.10) as previously described¹¹. Following infection, inject a subset of animals with antifungal agent such as 5 mg/kg caspofungin diacetate in DPBS used in this study (Daily, see Figure 2).
4. Anesthetize mice with 99.9% isoflurane using a veterinary anesthesia machine.
   1. Place a paper towel on the floor of the induction chamber to catch feces and urine.
   2. Place the mouse in the induction chamber, secure the lid, and set the vaporizer to 5% isoflurane for 2 min and 30 s.
      NOTE: Vet ointment is not used because the total time under anesthesia is less than 5 min.
   3. Reduce the vaporizer to half of the maximum setting for 30 s.
5. Confirm that the mouse is properly anesthetized by observing slowed breathing, lack of movement, and lack of response to toe-pinch stimulus.
6. Remove the mouse from the induction chamber and place on a slant board. Place the mouse with its back against the board. Ensure that its upper incisors are gently restrained with a rubber band and the lower incisors are gently restrained with a metal wire.
7. Carefully hold the tongue at full extension with small forceps and pipette 50 µL of conidia/PBS suspension at the base of the tongue.
8. Hold the tongue at full extension; listen for rapid breathing and the distinct clicking noise of aspiration. Hold for 20 s or until suspension is fully aspirated.
9. Remove the mouse from the slant board and gently place in a cage for observation.
10. Following aspiration, the mouse should regain consciousness within 1 min. Monitor the mouse until fully conscious and able to walk around the cage. The animals should be housed at 21 °C and monitored twice daily until the lungs are harvested.
11. Repeat the depletion of neutrophils (step 2.2) 24 h post-infection (Figure 2).

Figure 2: Experimental schedule. This figure was modified from a previous publication⁸.

3. Harvest and Preservation of Mouse Lungs for Histological Analysis

1. Deeply anesthetize the animal with a fatal dose of 0.1 ml of 390 mg/ml pentobarbital sodium solution IP.
2. Confirm euthanasia by observing lack of spontaneous breathing and absence of response to limb stimulation with tweezers. Once euthanized, spray the carcass with 70% ethanol to sterilize.
3. Place the carcass on a foam board covered with an absorbent pad.
   NOTE: The carcass should be placed on its back with pins securing all four limbs to the foam board.
4. Open the upper chest cavity using scissors with caution. Cut the rib cage away in order to expose the lungs and heart. Cut the inferior vena cava to allow for perfusion. Avoid cutting the blood vessels and puncturing organs.
5. Slowly perfuse with 5 mL of cold DPBS in the right ventricle of the heart at a 45° angle with a 25-G needle. Repeat the perfusion with 5 mL of 10% formalin.
6. Carefully expose the trachea and detach the surrounding fat pads. Use caution and avoid puncturing the trachea. Ensure the excess connective tissue is removed in order to fully visualize the trachea before proceeding. Once the trachea is exposed, place forceps under the trachea to elevate it.
7. Poke a small hole in the upper trachea using a 25-G needle.
   NOTE: The hole should be positioned so that the entire cannula fits in the trachea. Use caution to avoid rupturing the trachea.
8. Insert a cannula through the hole and towards the lungs. Secure the cannula with a surgical thread tied loosely around the trachea.
   NOTE: If the trachea is ruptured, the cannula may be inserted past the trachea as far as possible in the airway.
9. Using a syringe, inflate the lungs with 1 mL of 10% formalin buffer through the cannula. Ensure that the inflation of the lungs is visible. If not, adjust the catheter and repeat. Once the lungs are inflated, quickly remove the cannula with caution and tighten the thread securely to seal off the trachea.
   NOTE: If the trachea is ruptured and the lungs cannot be inflated, the lungs may still be harvested but this may prevent proper fixation.
10. Gently remove the trachea and lungs from the chest by carefully disconnecting the connective tissue of the trachea with forceps while gently pulling up on the thread. Place the lungs and trachea in a 50-mL conical tube with 15 mL of 10% formalin buffer. Place one end of the thread outside of the tube and seal the cap. Invert the tube to completely submerge the sample in fixative.
11. Store the samples at room temperature for at least 24 h or until further processing.
    CAUTION: Formalin is hazardous. Wear appropriate Personal Protective Equipment including goggles and a face mask. Work under a chemical hood when processing samples.

4. Harvest and Preservation of Mouse Lungs for DNA Fungal Burden

1. Follow steps 3.1-3.5 as described above on a separate mouse.
2. Slowly perfuse with 10 mL of cold DPBS in the right ventricle of the heart at a 45° angle with a 25-G needle.
3. Excise the lungs with scissors and place in labeled 1.5 mL tubes.
4. Store the lungs at -80 °C until further processing.
5. Use standard techniques as described (embedding and sectioning) and according to manufacturer's protocols (see Table of Materials).

5. Microscopy and Imaging

1. Open the imaging quantification program (see Table of Materials).
2. Using a light microscope, obtain at least 4 distinct fields of the GMS stained slides (10X objective). Ensure that the images are representative of the infected airways within the lung with similar tissue density.
   NOTE: If the density and distribution of hyphal foci appear markedly different between control and experimental groups, additional fields may be taken, or the GMS area of the whole lung section may be imaged (4X objective). Obtain images of the same areas of the lung (identified by equivalent tissue morphology) on a serial lung section stained with hematoxylin and eosin, if desired.
3. Add scale bars to the images where necessary and save the images to be quantified. Select, Edit > Add/Edit Calibration Marks > Menu: choose line thickness, color and scale bar size > click on the image for where the scale bar is to be placed. Close the Menu and choose, Edit > Merge > Merge Calibration Marks > choose File Save As > name the file and click on Save.

6. Using an Image Processing Program to Calculate the Fungal Burden (Figure 3)

1. Open an image processing program. Select File > folder with samples to quantify > image (Figure 3A). Select Image > Type > convert to "RGB Stack" (Figure 3B).
2. Use the right arrow key to select the second of the three given images (Figure 3B). Hit "control + shift + T" to bring up the "Threshold" menu settings adjustor (Figure 3C). Locate the original .tiff or .jpg image as a reference and open with an image viewing program (Figure 3C).
3. Pull the top slider all the way to the left and adjust the bottom slider until the area selected (in red) is representative of the microscopy image (typically ranging from 130-160 as indicated to the right of the slider (Figure 3C).
4. Once selected, press "Set" > "OK" (Figure 3D). Select "Analyze" > "Set Measurements" > check "Area, Area Fraction, Limit to Threshold, Display Label" and leave the bottom settings (the drop-down menu and decimal places) as default > "OK" (Figure 3D).
5. Significant areas of white space or background staining may be excluded by selection of the appropriate tissue with an area or polygon tool.
6. Finally select, "Analyze" > "Measure" (a window with data should appear, or a tab on the windows taskbar will appear labeled, "Results") (Figure 3E). Transfer the data to a data analysis program for graphing and statistical analysis.
7. Enter the mean of the four area percentages for each lung sample. 5-8 samples are generally sufficient to detect significant differences between groups. If comparing two experimental groups, perform an unpaired student's t-test to determine significance.

Figure 3: Representative screenshots for each step of GMS histological field quantification. (A) Select File > folder with samples to quantify > select image. (B) Select Image > Type > convert to "RGB Stack". Use the right arrow key to select the second of the three given images. (C) Hit "control + shift + T" to bring up the "Threshold" menu settings adjustor. Locate the original .tiff or .jpg image as a reference and open with photo viewer program. Pull the top slider all the way left and adjust the bottom slider until the area selected (in red) is representative of the microscopy image (typically ranging from 130-160 as indicated to the right of the slider. (D) Once selected press "Set" > "OK". Select "Analyze" > "Set Measurements" > check "Area, Area Fraction, Limit to Threshold, Display Label" and leave the bottom settings (the drop-down menu and decimal places) as default > "OK". (E) Finally select "Analyze" > "Measure" (a window with data should appear, or a tab on the windows taskbar will appear labeled "Results"). Please click here to view a larger version of this figure.

7. DNA Fungal Burden

1. Remove the lungs from -80 °C and lyophilize for at least 4 h (overnight is optimal) or until completely dried by a freeze dry system.
2. While the lungs are drying, prepare DNA extraction buffer (0.1 M NaCl, 10 mM EDTA pH 8.0, 10 mM Tris HCl pH 8.0, and 0.5% SDS).
3. Place the prepared DNA extraction buffer in a 60 °C water bath to pre-warm. Also, place phenol: chloroform: isoamyl alcohol (25:24:1) in a 50-mL conical tube and allow for separation.
4. Place the lyophilized lungs in a 2-mL screw cap tube with 0.2 mL of glass beads. Using a bead beater, grind dry lung tissue for 15-30 s until it forms a fine powder.
5. Add 0.8 mL warm DNA Extraction buffer to each tube and vortex well. Incubate each tube for 15 min in a 65 °C bead bath and then vortex before incubating for an additional 15 min. After incubating, vortex and then centrifuge tubes at 25,000 x g for 10 min.
6. Label 3 sets of 1.5 mL tubes for each of the samples and transfer the top layer of the supernatant into one set of tubes. Use caution while pipetting out the aqueous top layer to avoid disturbing the middle layer.
7. Add equal volume of phenol:chloroform:isoamyl alcohol and mix well. Then centrifuge at 25,000 x g for 15 min before pipetting out the upper layer into the second set of tubes.
8. Add equal volume of chloroform:isoamyl chloroform, mix well, and centrifuge at 25,000 x g for 10 min. Carefully pipette out the upper layer into the third set of tubes.
9. Add 500 µL of isopropanol and 50 µL NaoAc (pH 5.2), mix well, and allow samples to incubate at room temperature for 10 min. Then centrifuge at 25,000 x g for 30 min and decant the supernatant.
10. Wash the DNA pellet with 750 µL of ice cold 70% ethanol and centrifuge for 5 min at 25,000 x g. Decant the ethanol and allow the pellets to air dry in the tubes in a fume hoods for 40 min.
11. Resuspend the dry pellets in 300 µL of TE buffer.
    NOTE: The DNA may be stored at -20 °C until quantified.
12. Quantify DNA using a spectrophotometer and prepare 100 µL of 0.2 µg/µL of each DNA sample for qPCR analysis.
13. Perform a standard qPCR in triplicate as previously described with 1 µg of DNA sample, fungal 18S rDNA primer, and probe sets with a modified probe quencher (5'-/56-FAM/AGC CAG CGG/ZEN/CCC GCA AAT G/3IABkFQ/-3')^12,13.
14. Prepare a standard curve from A. fumigatus genomic DNA and calculate the concentration of fungal DNA in pg in each sample using the average reporter dye Ct values.
15. Plot the pg/µg of fungal DNA with standard error of the mean. Perform a student's t-test analysis to determine significance.
    NOTE: As a less toxic alternative to the phenol-chloroform DNA extraction used herein, column based DNA extraction kits can be used.