Experimental Model to Evaluate Resolution of Pneumonia

All the animal protocols described in this study were approved by the Institutional Animal Care and Use Committee (ACUC) at the Johns Hopkins University, School of Medicine for the animal protocol MO21M160. In addition, the experiments were performed following the institutional, state, and federal regulations for animal research.

CAUTION: The replication of all the protocols described below requires to be performed in a Biosafety level 2 (BSL-2) cabinet, following all of the institutional BSL-2 guidelines under the biosafety cabinet.

1. Plating bacterial stocks from commercial loops

NOTE: This protocol can be used for growing bacterial stocks for Spn (ATCC 49619) and Kpn (ATCC 43816), starting with the culti-loops obtained from the provider (see Table of Materials for details).

1. Warm a 5% sheep blood agar plate for 15 min at 37 °C in an incubator. A cell incubator can be used for this purpose.
2. Under the hood, remove the sheath from the bacterial inoculating loop and spread it carefully, streaking into the agar plate, following a zig-zag pattern. Repeat this step using a separate plate as a duplicate. Up to five plates can be streaked with the same loop.
3. Incubate the plates overnight at 37 °C for optimal bacteria growth. Passage the bacteria daily for 3 days. Warm the blood agar plates for 15 min at 37 °C, take between 15 to 20 colonies from the first agar plate using a disposable inoculating loop, and spread them into a new, prewarmed agar plate. Label the plate appropriately.

2. Bacteria growth and storage for future use

1. After 3 days, pick up to 30 colonies from the blood agar plate with a streak loop, and place it directly into a 250 mL flask containing Todd Hewitt broth (TH broth). Incubate at 37 °C, shaking at 250 rpm, with 5% CO₂ for approximately 4 h.
2. Take aliquots every 15 min to measure the OD_620nm until it reaches 0.3, which is equivalent to approximately 3 x 10⁸ colony forming units (CFU) per mL.
3. Aliquot out 1 mL of the new bacteria stock immediately into 2 mL cryogenic vials. Flash freeze the freshly aliquoted vials in liquid nitrogen for 5 min. Store aliquots of bacteria in a -80 °C freezer (vials can be used for up to 6-8 months before they lose potency).
4. After 7 days of freezing, the aliquots can be used for animal studies. Therefore, determine the new concentration of the bacteria.

3. Bacteria thawing for intratracheal instillation

1. Warm a blood agar plate for 10 min at 37 °C in a shaker. Take one of the new stock vials from the freezer and thaw it down by gentle agitation in a 37 °C water bath for approximately 2 min. Avoid touching the O-ring or cap with the warm water.
2. Plate on a blood agar plate to manually count the bacterial colonies. Perform a 1 x 10^-6 dilution, and plate 200 µL from the last dilution in a prewarmed blood agar plate. Do this in duplicates.
3. Incubate the plates overnight at 37 °C for optimal growth of the bacteria. The next day, apply the following formula to determine the new bacterial concentration: CFU/mL = (number of colonies x dilution factor) / volume of stock plated

4. Intratracheal instillation of live bacteria

NOTE: This protocol has been optimized to instill a volume of 50 µL intratracheally. Bacterial stocks can be stored for up to 6-9 months. To ensure the bacterial CFU in each vial, make sure to plate it before each experiment, calculate the bacterial stock CFU as described above, and use TH broth for subsequent dilutions.

CAUTION: Under the biosafety cabinet, perform a rodent survival procedure using sterile surgical instruments.

1. Anesthetize the mouse by injecting intraperitoneally 100 mg/kg of ketamine and 2.5 mg/kg of acetylpromazine. Repeat for the number of mice being injected at a time.
   NOTE: Mice can be exposed to isoflurane to facilitate animal handling. However, an experienced handler who is capable of injecting intraperitoneal anesthetics without causing significant animal distress can avoid isoflurane exposure. The number of mice to inject depends on the expertise of the surgeon.
2. After all the mice are under appropriate anesthesia, confirm anesthesia by tail pinch. Place the anesthetized mouse on a clean and sterilized surface, hang the animal by the incisors, and gently tape the forelimbs (Figure 1A).
   NOTE: To ensure animal wellness, provide corneal lubrication in all the anesthetized animals. Carboxymethylcellulose eye drops can be used, applying one drop per eye.
3. Shave the neck region and disinfect the area with chlorhexidine and 70% alcohol. Using surgical scissors, make a 1 cm superficial midline neck incision to visualize the trachea (Figure 1B). One small incision should be enough, but if an abundance of fat tissue is seen, carefully dissect the adipose tissue vertically to visualize the trachea. 
   NOTE: Use sterile gloves and sterile instruments via the tips-only technique. The risk of infection is minimal if the procedure is performed under sterile conditions.
4. Using forceps, intubate each mouse. Pull the tongue outward gently and introduce a 20 G angio-catheter through the mouth, advancing the catheter into the trachea. Apply gentle pressure over the trachea to facilitate intubation.
   NOTE: Because the trachea is anterior to the mouse body, slightly bend the tip of the catheter, as this will help get the angle to successfully intubate the mouse. The neck incision is performed mainly for visualization purposes; alternatively the operator can intubate the trachea blindly.
5. After intubation, connect the mouse to a respirator to confirm intubation. The regular ventilator parameters used in this procedure are 200 µL of stroke volume and 200 strokes per minute. Turn the ventilator rate up and/or down briefly for quick and easy confirmation.
6. After confirming intubation, disconnect the mouse from the respirator and carefully instill 50 µL of the bacterial agent through the angio-catheter using a 200 µL pipet gel loading tip. For control mice, inject 50 µL of sterile TH broth.
7. After instilling the bacterial agent, connect the mouse to the respirator again to help restart breathing. Leave the mouse on the respirator for 30-60 s. After injecting all the mice, monitor their breathing. If there's a slow breathing pattern, connect the mice to respirator again.
8. Close the incision by adding one small drop of glue on the skin. Bring together the skin folds and apply gentle pressure until the glue dries.
9. Place the mice on a heating pad for recovery and closely monitor them until they regain sufficient consciousness. Transfer the mice back in the cage once fully recovered. For post-operative pain/distress management, the mice can be treated with buprenorphine, with a dose of 0.05-0.1mg/kg subcutaneously (SC).
   ​NOTE: Animals with weight loss greater than 20% or experiencing significant distress post-procedure such as lethargy, an inability to reach water or food, labored breathing, impaired response to external stimuli, or decreased mental alertness should be euthanized.

5. Bronchoalveolar lavage and lung harvest

1. Euthanize the mouse by placing it in a closed container containing 5% isoflurane. Continue isoflurane exposure for 1 min after the mouse stops breathing. Perform thoracotomy to confirm euthanasia.
   NOTE: Confirm euthanasia by visual and physical examination. The heart must have stopped beating and the mice not breathing. Mucous membranes should be white or pale.
2. Lay the mouse in a supine position on a clean surgical board and hang it by the incisors. Spray the mouse skin with 70% ethanol. Using surgical scissors, make a small superficial midline neck incision to visualize the trachea of the mouse.
3. Cannulate the trachea with a 20 G catheter. Carefully, add 1 mL of calcium-free phosphate-buffered saline (PBS) intratracheally, using a 1 mL syringe. Allow full lung expansion, and then aspirate the fluid using the same syringe. Repeat this step for a total of 2 mL.
4. Transfer the BAL into a 2 mL aliquot. Perform this step twice for a final volume of 2 mL. Do not lavage the lungs with more than 1 mL at a time, due to the high risk of alveolar space disruption.
5. Open the thoracic cavity and expose the lung, heart, and trachea using scissors and forceps. Carefully dissect the diaphragm and remove the rib cage. Make sure not to pinch the lung tissue.
6. Transect the abdominal aorta to allow exsanguination. Perfuse the lung tissue by making a small incision of approximately 2 mm in the right ventricle using scissors and inject 5 mL of cold PBS using a 20 G catheter. The PBS should perfuse the lung. If an adequate perfusion is performed, the lung tissue will turn white pale and the PBS will leave the intravascular compartment through the abdominal aorta.
7. Carefully, extract the lungs and dissect them from the trachea to perform either histology or further processing to single cell suspension.
8. If processing for histology, carefully insert a 20 G catheter to inflate the lungs up to 25 cm H₂O with formalin solution (neutral buffered 10%). Once the lungs are insufflated, pass a 3.0 suture string about 5 cm long underneath the trachea, and tie it firmly twice to ensure the formalin stays in the lung tissue. Gently, dissect out the lung from the rest of the tissues and place it in a 15 mL conical tube containing 10 mL of formalin solution.

6. Bronchoalveolar lavage processing

1. Centrifuge the BAL at 500 x g at 4 °C for 5 min. Remove the cell-free supernatant in a separate tube and store at -80 °C.
   NOTE: Further analysis can be performed in the BAL supernatant, including protein quantification (e.g., BCA assay¹³) or measurement of specific biomarkers or cytokines (ELISA assays and multiple assays with platforms such as MSD and Luminex).
2. Lyse the red blood cells by adding 100 µL of lysing buffer for 1 min. Neutralize the lysing reaction by adding 1 mL of PBS. Centrifuge the BAL at 500 x g at 4 °C for 5 min and remove the supernatant.
3. Resuspend the cells in PBS (100-300 µL; based on cell pellet size). Perform a cell count with 0.4% trypan blue stain by manual or automated cell counting. Use the remaining pellet for flow cytometry staining and/or cryopreserve (by resuspending in cryopreserving solution) in liquid nitrogen for further testing.

7. Lung processing for single cell suspension

1. Gently dissect the lung from the rest of the tissues and place it in a 15 mL conical tube containing 5 mL of cold PBS. Remove the lung from PBS and dry it using a paper towel.
2. Prepare a digestion cocktail by adding 1 mg of DNase and 5 mg of collagenase into 1 mL of low glucose Dulbecco's modified Eagle's medium (DMEM). Transfer the lung to a C-tube containing 1 mL of digestion cocktail.
3. Transfer the C-tube to the tissue dissociator and follow the standardized protocol for processing lung tissue¹⁴.
4. Add 10 mL of cold PBS into the C-tube and mix properly. Filter the single cell suspension using a 70 µm cell strainer on top of a 50 mL conical tube. Perform the filtration twice.
5. Centrifuge the suspension at 500 x g at 4 °C for 5 min. Carefully decant the supernatant and add 1 mL of lysing buffer for 1 min at room temperature. Add 10 mL of cold PBS to stop the lysing reaction and remove supernatant.
6. Centrifuge the suspension at 500 x g at 4 °C for 5 min. Carefully decant the supernatant and add 10 mL of cold PBS.
7. Perform a cell count with trypan blue stain by manual or automated cell counting. Use a cell pellet for flow cytometry staining and/or cryopreserve in liquid nitrogen for further testing.