Ex vivo Pig Lung Model of Biofilm: A Technique to Develop Lung Model to Study Bacterial Biofilms on Pig Bronchiolar Tissue

All procedures involving animal models have been reviewed by the local institutional animal care committee and the JoVE veterinary review board.

1. Dissection and infection of ex vivo pig lung (EVPL) tissue

1. Make SCFM for use with EVPL tissue, follow the recipe for 1 L modified SCFM supplied in Table 1.
2. Prior to dissection, prepare an agar plate/s of required bacterial strain/s for infection using whatever agar is standard in the lab for P. aeruginosa/S. aureus (e.g., lysogeny broth + 1.2% agar).
3. Calculate how many porcine bronchiolar tissue pieces are required for the experiment, including uninfected control tissue pieces. Multiply this number by two to repeat the experiment in two replicate lungs to confirm repeatability of results.
4. Multiply the total number of tissue pieces required by 0.5 to determine the volume of SCFM agarose (mL) needed to make agarose pads to make enough medium for 400 μL/tissue piece plus spare SCFM agar to account for any pipetting errors or evaporation during preparation.
5. Add 0.12 g of agarose to every 15 mL of SCFM required to make the desired total volume of SCFM with 0.8% weight/volume agarose.
6. Heat the SCFM agarose solution until the agarose is fully dissolved. A domestic microwave on low power is recommended. The time required depends on the wattage of the microwave. Allow the agarose to cool to approximately 50 °C (warm to the touch but comfortable to hold). Do not allow to cool any further.
7. Using a pipette, add 400 μL of the SCFM agarose to one well of a 24-well plate per tissue piece needed.
8. Sterilize the SCFM agarose-containing 24-well plate/s under ultraviolet light for 10 min.
9. Prepare three replicate washes for every intact lung being dissected using 20 mL of sterile Dulbecco's modified Eagle medium (DMEM) plus 20 mL of sterile Roswell Park Memorial Institute (RPMI) 1640 supplemented with 50 μg/mL ampicillin.
10. Make an aliquot of 40 mL SCFM as a final wash for every intact lung being dissected. All washes can be stored overnight at 4 °C or used immediately.
11. Obtain lungs from the designated source as soon as possible after slaughter, ensuring they are kept cold by transporting to the laboratory in a domestic coolbox.
    NOTE: Lungs closer to the day of slaughter show less bruising from storage, but tissue kept on cold storage for up to 4 days from slaughter can also be used. As the coolbox needs to be taken into the butcher's shop or abattoir, it must be decontaminated following local lab guidelines after each use and stored outside the microbiology lab when not in use, to reduce the risk of contamination and a breach of containment.
12. Working on a sterilized surface and under a flame, place the lungs on a clean plastic chopping board covered with autoclaved aluminum foil. Check that the bronchioles remain intact. If there has been any damage at the abattoir or during transport the lungs are not suitable for use.
13. Heat a palette knife under a flame and very briefly touch the knife to the area of the lung surrounding the bronchiole to sterilize the surface of the tissue.
14. Cut away the surface tissue surrounding the bronchiole using a sterile mounted razor blade. Make incisions parallel to the bronchiole to prevent any damage.
15. Once the bronchiole has been exposed, make a cross-sectional incision through the bronchiole at the highest point visible to free the bronchiole.
16. Using sterile forceps, lightly hold the free end of the bronchiole and cut away any remaining unwanted tissue using a sterile mounted razor blade. Make a final cross-sectional incision across the bronchiole before any branching is visible to remove the bronchiole from the lungs.
17. Place the bronchiole in the first DMEM/RPMI 1640 wash. Leave the bronchiole in the wash and repeat steps 1.12 to 1.15 to harvest additional sections of bronchiole from the same lung as required to yield sufficient tissue sections for the planned experiment.
18. Place any additional bronchiolar sections from the same lung into the wash (step 1.17). Leave in the wash for at least 2 min.
19. Remove the bronchioles from the first DMEM/RPMI 1640 wash and place the samples in a sterile Petri dish.
20. Hold each bronchiole lightly using sterile forceps, making sure not to damage the tissue. Remove as much remaining soft tissue as possible and cut the tissue into ~5 mm wide strips using sterile dissection scissors.
21. Place all of the bronchiolar tissue strips into the second DMEM/RPMI 1640 wash. Leave in the wash for at least 2 min.
22. Remove the tissue strips from the second wash using sterile forceps, taking care not to damage the tissue. Place the tissue in a clean, sterile Petri dish.
23. Remove any remaining soft tissue attached to the bronchiole and cut the strips into squares (~5 mm x 5 mm) using sterile dissection scissors.
24. Add the third DMEM/RPMI 1640 wash into the Petri dish. Lightly mix the tissue pieces in the wash by swirling the dish.
25. Pour the third wash out of the Petri dish without removing the tissue pieces.
26. Add the final SCFM wash to the tissue-containing Petri dish, ensuring that all of the tissue pieces are covered.
27. Sterilize the tissue pieces in SCFM under UV light for 5 min.
28. Use sterile forceps to transfer each sterilized bronchiolar tissue piece into individual wells of a 24-well plate/s containing SCFM agarose pads.
29. To infect each tissue piece with the desired bacterial strain, touch a colony grown on an agar plate with the tip of a 29 G needle attached to a sterile 0.5 mL insulin syringe. Then touch the colony onto the tissue piece, gently pricking the tissue surface.
    NOTE: Using an insulin syringe equipped with a 29 G needle allows the needle to be held accurately and comfortably while keeping fingers at a safe distance from the both needle and lung tissue. It is possible to perform this step using 29 G needles that are not attached to a syringe, but this requires greater dexterity and increases the risk of a needlestick injury. Insulin syringes are readily available.
30. For the uninfected controls, gently prick the surface of each of the tissue pieces with the tip of a 29 G needle attached to a sterile 0.5 mL insulin syringe.
31. Use a pipette to add 500 μL of SCFM to each well.
32. Sterilize a breathable sealing membrane for each 24-well plate under ultraviolet light for 10 min (Table of Materials).
33. Remove the lid/s from the 24-well plate/s and replace with the breathable membrane.
34. Incubate the plates at 37 °C for the desired incubation (infection) time without shaking. Check that there is no visible growth of the inoculated pathogen on the uninfected control pieces (contamination control).
    NOTE: If desired, ampicillin may be added to the SCFM agarose pads and covering SCFM in step 1.31 to a final concentration of 20 μg/mL. This will suppress the growth of most endogenous bacteria on the lungs without affecting P. aeruginosa or S. aureus growth but, as the presence of ampicillin may affect susceptibility to other antibiotics, the reader is left to make this choice depending on the strains and antibiotics they wish to test.

TABLE 1

1. Preparation of Synthetic CF Sputum Media (SCFM) for use with ex vivo pig lung model

Once made, the SCFM should be filter sterilized, and may then be stored at 4°C for up to one month.