Modeling Asthma and Influenza Co-morbidity in C57BL/6J Mice

Ethics Statement. All experiments were approved by the Institutional Animal Use and Care Committee at St. Jude Children's Research Hospital.

Note: All described experiments in this protocol must be conducted under BSL-2 and ABSL-2 conditions since Aspergillus fumigatus and influenza A virus are BSL-2 agents.

1. Modeling Allergic Asthma in Mice (Figure 1)

1. Fungal sensitization
   1. Re-constitute A. fumigatus antigen in sterile PBS at 1 mg protein/mL. Aliquot and store at -20°C.
   2. House animals in a micro-isolator cages with autoclaved paper bedding under specific pathogen-free conditions with ad libitum food and water in a 12 hour light:dark cycle. Obtain C57BL/6J female mice at six weeks of age and acclimatize for one week. Start mice on the asthma protocol at seven weeks of age.
   3. Dilute A. fumigatus antigen to 20 µg/mL in sterile PBS and add dropwise to an equal volume of Alum while stirring. Allow antigen to be adsorbed for 30 mins at room temperature with constant stirring.
   4. Fill 1 mL luer-lock syringes with the mixture and deliver 0.1 mL intraperitoneally and 0.1 mL subcutaneously between the ears to each mouse. Administer PBS and Alum as a mock-sensitization to mice that serve as naïve controls.
   5. Two weeks after the initial sensitization, dilute A. fumigatus antigen to a concentration of 200 µg/mL in sterile PBS and deliver 20 µL intranasally to each mouse. Repeat once weekly for a total of three intranasal administrations. Administer PBS as a mock-sensitization to mice that serve as naïve controls.
2. Fungal challenge
   1. Prepare Sabouraud dextrose agar at 65 g/L per manufacturer's guidelines and sterilize in an autoclave at 121°C for 15 mins. Place in a water bath set at 50°C. to temper.
   2. Once tempered, aseptically add 10 mL to 25 cm² canted neck, vented cap cell culture flasks. Lay the flasks flat and allow agar to set. Once set, recap flasks and store at 4°C until use.
   3. Re-suspend the A. fumigatus culture aseptically by adding 5 mL of sterile PBS. Store overnight at 4°C. Prepare 60 µL aliquots for use and store at 4°C.
   4. Eight days before the final intranasal administration of A. fumigatus antigen, culture A. fumigatus on Sabouraud dextrose agar for the fungal challenge. Add one well vortexed 60 µL aliquot of fungus in each agar flask and spread using a disposable inoculating loop. Incubate flasks at 37°C for 8 days.
   5. Set up the inhalation challenge apparatus in a class II biological safety cabinet as described in detail by Schuh and Hoselton.⁷ Create holes in the fungal culture flasks, one on the rear top and one on the bottom of the flask immediately above the culture by using a cork-borer.
   6. Attach the fungal culture flask to the regulated medical grade air supply through the bottom hole and to the inhalation chamber through the top hole. Cover the inhalation ports with laboratory tape.
   7. Run medical grade air over the culture at 2 psi for 10 mins to coat the chamber. Prepare sterile anesthetic cocktail with 75 mg/kg ketamine and 25 mg/kg xylazine.
   8. Inject the drug cocktail to three mice intraperitoneally and determine anesthesia by toe pinch reflex and lack of movement. Apply a drop of lubricating ointment in each eye to prevent drying out.
   9. Turn off the air, and replace the fungal flask with a new one and open the ports of the inhalation chamber.
   10. Place the anesthetized mice supine with their noses in the ports. Turn on the air over the culture and allow the animals to inhale the liberated fungal conidia for 10 mins.
   11. Place each animal supine in a new cage and place the cage on a heating pad. Monitor animals for recovery from anesthesia marked by turning over and movement around the cage.
   12. Repeat procedure after 14 days. Anesthetize animals that serve as naïve controls and place in a new cage.

2. Modeling Influenza in Mice

NOTE: Adjust the age, gender, and strain of mice and the dose of the virus according to the experimental goals.

1. Obtain female C57BL/6J mice at 14 or 17 weeks of age and acclimatize to the facility in micro-isolator cages with autoclaved paper bedding and ad libitum food and water in a 12 hour light:dark cycle. Number animals in each cage by tail mark or ear notch. One week later, weigh each mouse and record weight with date and time.
2. Thaw a stock of A/CA/04/2009 virus and dilute in sterile PBS at log₁₀4.305 TCID50/mL in a glass vial. Anesthetize mice with 2.5% isoflurane with oxygen in an anesthesia chamber within the class II biological safety cabinet.
3. Deliver 50 µL of virus intranasally to each mouse for a final dose of 1000 TCID50 pH1N1 per mouse. Hold mouse supine until conscious and place in new micro-isolator cage. Administer PBS under anesthesia to naïve controls.
4. Record animal weights every 24 hours until experimental endpoints are reached.
   NOTE: It is good practice to use the same stock virus tube(s) to infect all animals in the study and not use previously thawed tubes as the viral titer may decrease during repeated freeze-thaw cycles.

3. Modeling Asthma and Influenza Co-morbidity in Mice (Figures 2 and 3)

1. To mimic acute asthma and influenza, infect animals one week after the second fungal challenge with pH1N1 as noted in section 2. To mimic chronic asthma and influenza, infect animals 4 weeks after the second fungal challenge.

4. Pulmonary function test

1. Weigh and record each mouse weight.
2. Perform pulmonary function test on each animal. Note: Of the various equipment available to measure pulmonary function in rodents, we have opted to use the flexiVent FX1 module and flexiware software as previously described.²²
   1. Open the software and go to "Study Definition and Planning." Click on "Create a new study" to launch the Study Definition Wizard.
   2. Follow the Wizard to input details of the study: Title and Status, Objective and Hypothesis; Protocol; Subject (animal) Groups.
   3. Create subjects by clicking on the mouse icon and add all necessary information on the animal including the Group Membership which will assign subjects to Subject Groups.
   4. Verify all information and confirm to close the wizard.
3. After concluding the "Study Definition and Planning" session, open the "Experimentation Session" in the software. Follow the steps of the pop-up wizard to set-up the instrument.
   1. Select the Study (see 4.2.1) and the Experiment Template which depends on the instrument.
   2. Add details as deemed necessary to Session Properties. The instrument will then be connected to the software.
   3. Select the instrument module attached to the software when promoted and click OK.
   4. Select a subject in the Study to Site 1 and enter the current weight of the animal when prompted.
   5. Add 0.5 mL of saline to the finemist nebulizer and click OK when prompted to Prime the nebulizer (the generated mist will be visible). Attach the nebulizer to the instrument adaptor when complete.
   6. Perform Channel Calibrations when prompted. Calibrate the Cylinder Pressure and Airway Opening Pressure following the software wizard using a manometer to apply known pressures.
   7. At the conclusion of Channel Calibrations, the Tube Calibration Wizard will be initiated; attach an 18 g cannula to the Y-tubing adaptor and select the desired perturbations. Perform closed and open calibrations and verify that Calibration Results are void of errors.
   8. Once the calibrations are complete, select whether to start recording continuous data when prompted. Click "Yes" to start the default ventilation pattern. The instrument is ready to gather data.
4. Prepare a stock acetyl-β-methylcholine (AβM) at 125 mg/mL in sterile saline; freeze aliquots at -20°C until use. Dilute the frozen stock of AβM to the desired doses in sterile saline (3.125 mg/mL, 6.25 mg/mL, 12.5 mg/mL, and 25 mg/mL).
5. Prepare anesthetics in sterile PBS before procedure. Administer 10 mg/kg body weight xylazine intraperitoneally and place mouse its own cage to rest for 5 min. Administer 70 mg/kg body weight pentobarbital intraperitoneally and allow mouse to rest for 5 min.
6. Intubate mouse as previously described.²³
   1. Briefly, place the anesthetized mouse supine on a slanted surgical board and fix in place with plastic tape over the forelimbs and tail.
   2. Make a small incision between the upper thorax and lower jaw. Separate the salivary glands and thyroid gland to expose the trachea. Gently pass curved surgical forceps underneath the trachea and pull a piece of thread underneath.
   3. Perform tracheostomy between two tracheal rings with sharp scissors taking care not to cut through the trachea.
   4. Insert the 18 g cannula that was used to calibrate the instrument. Fasten the cannula to the trachea with thread and attach the mouse to the Y-tubing of the instrument adaptor with the cannula.
7. Perform a deep inflation to recruit the airways and help the animal succumb to the ventilator. Select the "mouse inhaled dose response v7.0" script with default ventilation at 150 breaths/minute.
8. Add saline followed by each incremental dose of AβM to the nebulizer when prompted to perform a dose response curve.
9. At the conclusion of the data acquisition after the highest AβM dose, disconnect the mouse from the ventilator and humanely euthanize the animal by performing a cervical dislocation. Proceed to tissue harvest.
10. Add saline to the nebulizer and activate the nebulizer to remove any remnant high dose AβM. Change the subject in the software program and proceed with next mouse beginning with a tube calibration (step 4.1-4.3).

5. Tissue harvest

1. Spray euthanized mouse with 70% ethanol and open the thoracic cavity. Open diaphragm and perform bronchoalveolar lavage (BAL) with 1 mL of sterile PBS injected through the 18 g cannula and gently draw out repeating once for a total of 2 mL.²⁴ Place the sample in ice.
2. Harvest the cardiac and diaphragmatic lobes of the right lung for RNA work, and azygous and apical lobes for protein work, place in separate 5 mL snap-cap tubes and snap freeze in liquid nitrogen.
3. Harvest the left lobe using blunt forceps and scissors taking care not to damage/slice through the lobe. Fix the lung ex vivo with 10% normal buffered formalin with a 30 g needle. Store in 10% normal buffered formalin until histological analyses.
   NOTE: When animals are taken through the model for determination of viral burden in the lungs, harvest whole lungs after euthanasia and snap freeze in liquid nitrogen.

6. Tissue processing for Flow Cytometry

1. Centrifuge the BAL contents at 600 g for 10 mins at 4°C and save the BAL fluid for protein assays at -80°C until use.
2. Perform red blood cell lysis if necessary noting that cells are more susceptible to death after viral infection. Perform red cell lysis by adding 8 parts cold distilled water and pulse-vortexing for 30 seconds followed by a one-minute incubation at room temperature. Add one part 10 PBS, mix to neutralize the water and centrifuge as above.
3. Re-suspend the BAL cells in 200 µL of staining media (sterile PBS with 5% FBS) and enumerate cells by trypan blue dye exclusion assay to determine the cell concentration.
4. Cytospin 50,000 cells as follows:
   1. Transfer the calculated volume of cells into a tube and bring volume up to 100 µL with sterile PBS. Assemble the labeled slide and cytofunnel in the metal clip and place in the centrifuge.
   2. Add the diluted cells into the cytofunnel and spin at 55 g for 5 minutes to affix the cells to the slides.
   3. Perform differential staining by sequentially dipping the slides 20 times each in methanol, xanthene, and thiazine followed by distilled water and air-drying.
5. Add 1 µL of human gamma globulin for every 10,000 cells in the cell suspension in step 6.3 and incubate in ice for 30 mins. Adjust volume to 1 mL with staining media and centrifuge at 600 g for 10 mins at 4°C and aspirate supernatant. Re-suspend cells in 100 µL of staining media and prepare four pooled samples with 10 µL from each sample. Adjust volumes to 200 µL and centrifuge as above.
6. Re-suspend cells in 50 µL of antibody cocktail containing fluorochrome-conjugated antibodies diluted in staining media (see table of materials). Re-suspend one of the pooled samples in 50 µL of staining media, one pooled sample in 50 µL of isotype control antibodies, and the remaining pooled samples with diluted antibodies individually. Incubate samples in ice protected from light for 30 mins.
   NOTE: The use of antibodies will be based on research interests and should be optimized previously for the experiment. Each antibody used should be used individually in a pooled sample for instrument set up. The use of bone marrow cells or splenocytes for single color controls may be necessary if the cell yield is low and is acceptable if fluorochrome-conjugated antibodies are made against an abundant marker such as B220.
7. Add 450 µL of staining media and centrifuge at 600 g for 10 mins at 4°C. Aspirate supernatant and repeat.
8. Aspirate supernatant and re-suspend cells in 50 µL of freshly prepared stabilizing fixative by gentle pipetting. Add 200 µL more of stabilizing fixative and strain through a 40 µm nylon mesh into a polystyrene FACS tube for analysis by flow cytometry.
   NOTE: Whole lungs may be digested to make single cell suspensions and used to analyze cell populations by flow cytometry.

7. Tissue Processing for RNA Analysis

1. Place frozen lung lobes in ice within a fume hood, and perform standard Trizol extraction of RNA according to manufacturer's recommended protocol. After DNase treatment and purification of RNA, convert 1 µg of RNA to cDNA.
2. Using RNA-specific validated primer sets, measure changes in gene expression with quantitative real-time PCR and analyze data using the 2^–ΔΔCt method normalized to the internal housekeeping gene²⁵ validated for lungs.

8. Tissue processing for Viral Titration

1. Place frozen lungs in ice and add 1 mL of sterile PBS and mechanically homogenize tissue until entire tissue is pulsed. Centrifuge samples at 600 g for 10 mins at 4°C and store supernatants in aliquots at -80°C until use.
2. Quantify virus using standard tissue culture infectious dose 50 (TCID₅₀) method²⁶ with MDCK.2 cells.