A Mouse Model for the Transition of Streptococcus pneumoniae from Colonizer to Pathogen upon Viral Co-Infection Recapitulates Age-Exacerbated Illness

All animal studies were performed in compliance with the recommendations in the Guide for the Care and Use of Laboratory Animals. All procedures were approved by the University at Buffalo Institutional Animal Care and Use Committee.

1. Preparing chemically defined media (CDM)

1. Prepare the stocks as follows:
   1. Dissolve the mix I compounds listed in Table 1 in 100 mL of ultrapure water while stirring. Store in 200 µL aliquots at −20 °C.
   2. Dissolve the mix II compounds listed in Table 1 in 20 mL of 0.1 M NaOH while stirring. Store in 100 µL aliquots at −20 °C.
   3. Dissolve the mix III compounds listed in Table 1 in 1 mL of ultrapure water while stirring. Store in 10 µL aliquots at 4 °C.
   4. Dissolve the mix IV compound listed in Table 1 in 1 mL of ultrapure water while stirring. Store in 10 µL aliquots at −20 °C.
   5. Dissolve the compounds listed in Table 2 initially in 15 mL of ultrapure water while stirring. Adjust the pH to 7.0 with a few drops of 0.1 M NaOH and adjust the final volume to 20 mL using ultrapure water. Store in 1 mL aliquots at −20 °C.
   6. Dissolve the compounds listed in Table 3 in 90 mL of ultrapure water on a hot plate at 50 °C while stirring. Adjust the pH to 7.0 with 0.1 M NaOH, and then adjust the final volume to 100 mL using ultrapure water. Store in 5 mL aliquots at −20 °C.
2. Make the starter stock freshly every time by dissolving the compounds in Table 4 in 70 mL of ultrapure water while stirring.
3. To the fresh starter stock, add the following mix stocks in order: 200 µL of Mix I stock (Table 1), 80 µL of Mix II stock (Table 1), 10 µL of Mix III stock (Table 1), 10 µL of Mix IV stock (Table 2), 1 mL of Vitamin stock (Table 3), and 5 mL of Amino Acid stock (Table 4).
4. Once the stocks have been added, adjust the final volume to 100 mL by adding 30 mL of ultrapure water to the beaker.
5. Supplement the CDM with compounds from Table 4. Once mixed thoroughly, filter-sterilize and store at 4 °C for a maximum of 2 weeks.

2. Growing the S. pneumoniae biofilm

1. Prepare RP-10 medium by mixing 445 mL of RPMI 1640 with 50 mL of heat-inactivated fetal bovine serum (FBS) and 5 mL of penicillin/streptomycin at 10,000 U/mL and 10,000 µg/mL, respectively.
2. Grow the NCI-H292 (H292) mucoepidermoid carcinoma cell line. Add the cells from one purchased vial to 5 mL of RP-10 medium in a T-25 tissue culture-treated flask. Incubate at 37 °C/5% CO₂ for 3-5 days to reach 100% confluence.
3. Check the cells under a light microscope using 10x magnification to assess confluency.
   NOTE: When all the cells are in contact with other cells and there are no gaps in between, then the desired 100% confluency is reached.
4. Wash the cells 2x in 5 mL of room-temperature PBS. Ensure the buffer is calcium free to avoid chelating the EDTA in the following step.
5. Add 1 mL of trypsin-EDTA to the flask and incubate at 37 °C/5% CO₂ for 5-10 min until the cells detach. Neutralize with 4 mL of RP-10 medium. Gently mix by pipetting up and down and transfer to a 50 mL conical tube.
6. Add 500 µL of the cell suspension per well to a tissue culture-treated 24-well plate. From a confluent T-25 flask, expect 2 × 10⁶-4 × 10⁶ cells/mL.
7. On the following day, check the cells under a light microscope to make sure that they are confluent, as in step 2.3. If they are not, then incubate for longer.
8. Once the H292 cells are 100% confluent in the 24-well plate, gently wash the cells 3x with 1 mL of room-temperature PBS to ensure that no medium containing antibiotics or debris remains.
9. After washing the cells, add 250 µL/well of 4% paraformaldehyde to fix the cells. Incubate for either 1 h on ice or overnight at 4 °C.
10. The night prior to cell fixation, streak the S. pneumoniae strain of interest on blood agar plates and incubate overnight at 37 °C/5% CO₂.
    NOTE: The data presented here are with the following S. pneumoniae strains obtained via collaborative exchange: serotype 19F otitis media isolate EF3030²⁴, classical serotype 2 Avery strain D39²⁵, and serotype 4 bacteremia isolate TIGR4²⁶. The strains are also available from public collections referenced in the Table of Materials.
11. Prepare CDM plus oxyrase (0.15 U/mL) by adding 100 µL of oxyrase (30 U/mL) to 20 mL of CDM.
    NOTE: Oxyrase is used to eliminate oxygen to allow the efficient growth of S. pneumoniae in liquid culture²⁷.
12. Inoculate the bacteria from the plate into fresh CDM + oxyrase by washing the bacteria off the plate by adding 1 mL of the CDM + oxyrase and gently lifting the bacterial colonies using the side of a 1 mL pipette tip, being careful to not scrape the agar. Alternatively, use an inoculating loop to lift the bacteria and inoculate them into a tube containing 1 mL of the CDM + oxyrase.
13. Dilute the bacteria in CDM + oxyrase to a starting OD₆₀₀ of 0.05.
14. Grow the bacteria in a loosely capped 50 mL conical tube standing at 37 °C/5% CO₂ until an OD₆₀₀ of 0.2 is reached (this will take anywhere between 2-5 h). Check the OD₆₀₀ every hour to ensure the OD does not exceed 0.2.
15. Once the OD has reached 0.2, vortex the bacterial culture tube. Seed 0.5 mL of the bacteria on the fixed H292 cells and add another 0.5 mL of CDM + oxyrase medium per well. Add 1 mL of CDM + oxyrase to the control wells with no bacteria. Incubate the plate for 48 h at 34 °C/5% CO₂.
    NOTE: Growth at 34 °C is used to more closely mimic the lower temperature in the nasopharynx²¹.
16. Every 12 h following the initial seeding, gently remove 0.5 mL of the medium and replenish with 0.5 mL of fresh CDM + oxyrase. Be careful not to disrupt the forming biofilm. Check the bottom of the plate for biofilm and look for increasing cloudiness as time goes on due to the growth of the biofilm. To control for contamination, check the wells without bacteria to ensure that the control wells remain clear.
17. At 48 h post inoculation, remove the supernatant and wash 2x very gently with 1 mL of PBS. Resuspend in 1 mL of fresh CDM and pipette up and down vigorously to lift the biofilm. For each bacterial strain, pool the bacteria from all the wells into a 50 mL conical tube. Mix well by gently tilting the tightly capped tube up and down several times.
18. To the 50 mL conical tube, add 40% glycerol in CDM at equal volumes to achieve a bacterial suspension with a final concentration of 20% glycerol. Aliquot 1 mL into microcentrifuge tubes, flash-freeze on dry ice, and save at −80 °C.
19. Prior to use, enumerate the bacteria by thawing one aliquot on ice, spinning the tube at 1,700 × g for 5 min, removing the supernatant, resuspending the pellet in 1 mL of PBS, and plating serial dilutions on blood agar plates²⁸.
20. Grow the agar plates overnight at 37 °C/5% CO₂ and count the colonies at relevant dilutions to obtain the bacterial concentration in colony-forming units (CFU)/mL.
    ​NOTE: It is recommended to enumerate the bacteria in the stocks at least a day after freezing or later, as there is a drop in bacterial viability within the first 24 h. The stored frozen aliquots can be used for subsequent infection of mice for a maximum of 2 months.

3. Intranasal inoculation of mice with biofilm-grown S. pneumoniae

1. Purchase mice and use at the desired age.
   NOTE: Mice aged 3-4 months old are preferred to model young hosts, and mice aged 21-24 months old can be used to model elderly individuals >65 years of age²⁹. The data presented here are with C57BL/6 male mice.
2. Thaw the biofilm-grown bacterial aliquots on ice and spin at 1,700 × g for 5 min. Carefully remove and discard the supernatant without disrupting the pellet, wash the bacteria by resuspending the pellet in 1 mL of PBS, and spin again at 1,700 × g for 5 min. Remove the supernatant and resuspend the pellet in the volume needed to reach the desired concentration (aim for 5 × 10⁶ CFU/10 µL for intranasal inoculation). Confirm the amounts of bacteria administered by plating the prepared inoculum on blood agar plates as in step 2.19.
3. Inoculate the mice intranasally with 5 × 10⁶ CFU by pipetting 5 µL of the diluted inoculum into each naris. Make sure to hold the mice firmly, stabilizing the head, until the volume is inhaled (typically within seconds of pipetting the volume into the nares). Perform this step in the absence of anesthesia to prevent pulmonary aspiration of the inoculum.

4. Viral infection with influenza A virus (IAV)

1. At 48 h following intranasal inoculation with S. pneumoniae, thaw the IAV strain of interest on ice.
   NOTE: The data presented here are with a mouse-adapted strain of influenza A virus A/PR/8/34 H1N1 that was obtained via collaborative exchange³⁰.
2. Once the virus has thawed, dilute the virus in PBS to the desired concentration; aim for 20 plaque-forming units (PFU)/50 µL for intratracheal infection and 200 PFU/10 µL for intranasal infection. For mock-infected and bacteria-only groups, use PBS to inoculate the mice.
3. Place ophthalmic lubricant on the eyes of the mice prior to anesthesia. Anesthetize the mice using 5% isoflurane and confirm anesthesia by a firm toe pinch.
4. Once the animal is anesthetized, remove it from the isoflurane chamber and immediately infect the anesthetized mice with 50 µL (20 PFU) of IAV intratracheally using blunt tweezers to pull the tongue out of the mouth and pipetting the volume of liquid down the trachea.
5. Place the mice in a separate cage and monitor until complete recovery (they are able to maintain sternal recumbency [able to lay upright on the chest]).
6. Following recovery, immediately intranasally inoculate the mice with 10 µL (200 PFU) of IAV using the inoculation method in step 3.3.
7. House mice that have undergone single or dual bacterial and viral infection with the same infection group and separate them from the other groups.

5. Monitoring the mice for disease symptoms

1. Monitor the mice daily for at least 10 days and blindly score for signs of sickness as follows:
   1. Score as follows for weight loss: 0 = 5% or less; 1 = 5%-10%; 2 = 10%-15%; 3 = 20% or more. Euthanize the mice using CO₂ inhalation when the weight loss score is at 3.
   2. Score as follows for activity: 0 = normal/active; 1 = moving but slightly diminished; 2 = diminished; 3 = severely diminished/lethargic (only moves if touched), 4 = coma/immobile. Euthanize the mice when the activity score is at 3.
   3. Score as follows for posture: 0 = no hunch (normal); 1 = slightly hunched posture; 2 = severe hunch. Euthanize the mice when the posture score is at 2.
   4. Score as follows for the eyes: 0 = normal; 1 = protruding; 1 = sunken; 1 = closed; 1 = discharge. It can be a combination. Add the totals for the final eye score.
   5. Score as follows for breathing: 0 = Normal breathing; 1 = irregular or altered (higher/lower rate); 2 = labored (exaggerated effort or gasping). Euthanize the mice when the breathing score is at 2.
2. Based on the above criteria, add the individual scores for a total clinical score of healthy (0) to extremely sick (15). Consider any mouse displaying a total score above 2 to be sick. Humanely euthanize any mice displaying a total score above 9 or the indicated scores for each criterion and mark them on the survival curve.

6. Processing of infected tissues for bacterial enumeration

1. At 48 h following IAV infection, euthanize the mice.
2. Place the mouse in a supine position. Using 70% ethanol, spray the chest and abdomen of the mouse to clean the fur. Using forceps, pinch the fur and skin in the middle of the mouse and cut the fur with 4.5 in dissection scissors to expose the area from the liver up to the chest.
3. Blood collection
   1. Using dissection scissors, gently cut into the peritoneal cavity to expose the liver. Using forceps, expose the hepatic portal vein at the top of the liver near the diaphragm. Cut the hepatic portal vein using the dissection scissors. Once the blood starts to pool in the peritoneal cavity, collect 10 µL of blood using a micropipette and place into 90 µL of anticoagulant solution (50 mM EDTA solution in PBS) in a microcentrifuge tube for plating for bacterial burden.
   2. Use a P-1000 micropipette to collect the rest of the blood, place it in a blood collection tube, and centrifuge at 7,600 × g for 2 min to collect the serum. Save the sera in microcentrifuge tubes at −80 °C for subsequent analysis of any desired cytokine or metabolite.
4. Lung collection
   1. Using dissection scissors, make a cut up the sides of the exposed rib cage and gently pull the ribs up toward the head of the mouse to expose the heart. Insert a 25 G needle attached to a 10 mL syringe prefilled with PBS into the right ventricle and begin slowly perfusing. Look for bleaching of the lungs as an indicator of successful perfusion. Flush slowly to avoid breaking the pulmonary tissue.
   2. Lift the heart with the forceps and make a cut to separate the lungs and heart. Once separated, pick up all lobes of the lung with the forceps and rinse in a dish with sterile PBS to remove any residual blood. In a Petri dish, mince the lungs into small pieces and mix well. Remove half of the lung mix for determination of the bacterial CFU or viral PFU and place it in a round bottom 15 mL tube prefilled with 0.5 mL of PBS for homogenization.
      NOTE: It is important not to take different lobes of the same lung for the various assessments. Instead, all the lobes should be minced, mixed well together, and parsed out equally for the different assessments.
   3. Remove the other half of the lung for flow cytometry (section 7 below) and place it in a non-tissue culture-treated 24-well plate with each well prefilled with 0.5 mL of RP-10. Leave at room temperature until processing.
5. Nasopharynx collection
   1. At the neck, use the dissection scissors to cut away the fur, and then cut away the muscle and expose the trachea.
      NOTE: The trachea is a tube-like structure located under the muscle.
   2. Place small forceps under the trachea at a distance of 1 cm from the mouse's jaw to stabilize it. Using dissection scissors, gently make a 0.1 cm slit on the anterior portion of the trachea, avoiding cutting the trachea completely.
   3. Prepare a 1 mL syringe filled with 0.5 mL of PBS with 0.58 mm tubing attached to a 25 G needle. Collect the nasal wash by inserting the tubing into the trachea going upward toward the nasopharynx. Once resistance is felt entering the nasal cavity, place a microcentrifuge tube at the nose and slowly flush the PBS through the trachea to collect the nasal lavage.
   4. Place the mouse in a prone position. Spray the head of the mouse with ethanol. Use dissection scissors to cut the fur and mystacial pad to expose the head bone of the mouse.
   5. Using the dissection scissors, make a 1 cm cut down the sides of the mandible and between the eyes. Using forceps, slowly pull the facial bones away from the body to expose the nasal cavity.
   6. Use forceps to gently remove the nasal tissue and place it into a round bottom tube prefilled with 0.5 mL of PBS for homogenization.
6. To homogenize the collected tissue, first clean the homogenizer probe by putting it in 70% ethanol and turning on the homogenizer at 60% power for 30 s. Repeat the step in sterile water for 10 s. Homogenize each tissue for 1 min. Clean the homogenizer probe in sterile water between each sample and in a fresh tube of 70% ethanol between each organ and sample group.
7. Enumeration of bacterial numbers
   1. Once all the organs have been harvested and homogenized, plate serial dilutions on blood agar plates. To calculate the total CFU, use 10 µL to plate and note down the final volume in mL for each sample. Plate the nasopharynx samples on blood agar plates supplemented with 3 µg/mL gentamicin to select for the growth of S. pneumoniae while inhibiting the growth of other microorganisms that colonize that tissue. Incubate overnight at 37 °C/5% CO₂.
   2. To enumerate the bacterial CFU for the lung and nasopharynx, first count the colonies on the blood agar plates. Then, use equation (1) and equation (2) to calculate the amount per mL and total number.
      Amount per mL = number of colonies × dilution factor × 100   (1)
      Total number = amount per mL × total volume per sample   (2)
      NOTE: In equation (1), 100 is used to multiply since 10 µL is plated, which is a 100-fold dilution of 1 mL. The total volume per sample in equation (2) is from step 6.7.1, which results in the limit of detection of 100 per organ.
   3. To enumerate the bacterial CFU For bacteremia, first count the colonies on the blood agar plates. Then, use equation (3) to determine the amount per mL of blood.
      Amount per mL of blood = number of colonies × dilution factor × 100 × 10   (3)
      NOTE: In equation (3), 100 is used as 10 µL is plated, which is a 100-fold dilution of 1 mL, and 10 indicates a 1:10 dilution of the blood in anticoagulant. This results in the limit of detection of 1,000/mL.

7. Processing of the lung samples for flow cytometry

1. Prepare the required media as follows:
   1. Prepare RP-10 as described in step 2.1.
   2. Prepare digestion buffer by mixing RP-10 with 2 mg/mL collagenase and 30 µL/mL DNase I.
   3. Prepare lysis buffer by dissolving 8.29 g of NH₄Cl, 1 g of NaHCO₃, and 0.038 g of EDTA in 1 L of H₂O.
   4. Prepare 10x FACS buffer by mixing 450 mL of HBSS with 50 mL of heat-inactivated FBS and 5 g of sodium azide.
   5. Prepare 1x FACS buffer by diluting 50 mL of 10x FACS buffer in 450 mL of HBSS.
2. Take the lung samples from step 6.4.3 and place in a 24-well plate. Add 500 µL of digestion buffer to each well. Incubate for 45 min up to 1 h at 37 °C/5% CO₂.
3. Prefill 50 mL conical tubes for each sample with 5 mL of RP-10. When the incubation is over, place a 100 µm filter at the top of the 50 mL conical tube and wet it with 1 mL of RP-10.
4. Using a P-1000 micropipette, move the digested lungs and place them on the filter. Use the plunger of a 3 mL syringe to mash the organ. Rinse 2x with 1 mL of RP-10 each time.
5. Spin the samples at 4 °C and 327 × g for 5 min. Aspirate the supernatant and resuspend the pellet in 1 mL of lysis buffer. Leave for 3 min to allow lysis of the red blood cells. Neutralize with 5 mL of RP-10.
6. Spin the samples at 4 °C and 327 × g for 5 min. Aspirate the supernatant, resuspend the pellet in 1 mL of RP-10, and take 10 µL for counting the samples.
7. Spin the samples at 4 °C and 327 × g for 5 min. Aspirate the supernatant and resuspend the pellet in RP-10 at 2 × 10⁶-4 × 10⁶ cells/mL. Add 60 µL of each sample into a 96-well plate to stain for the desired cell types²³ listed in step 7.9, Table 5, and Table 6.
8. Spin the plate at 4 °C and 327 × g for 5 min.
9. Meanwhile, prepare the antibody master mixes, florescent minus one (FMOs), and single-stain controls with the desired antibodies. To stain for polymorphonuclear leukocytes (PMNs), macrophages, monocytes, dendritic cells, and T cells, use the antibodies and final dilutions listed in Table 5 and Table 6. Use a total volume of 100 µL/well of the antibody mix. Follow the dilutions listed in the tables for determining the appropriate volume of the master mix and the individual antibodies required.
10. When the spin is done (step 7.8), decant the supernatant, resuspend the pellets in 100 µL of the antibody mixes, FMOs, or single-stain controls, and incubate on ice for 30 min in the dark.
11. Wash the cells 2x by adding 150 µL of FACS buffer to the wells and spinning the plate at 4 °C and 327 × g for 5 min.
12. When the spin is done, decant the supernatant, resuspend the pellets in 100 µL of fixation buffer, and incubate on ice for 20 min.
13. Wash the cells 2x by adding 150 µL of FACS buffer to the wells and spinning the plate at 4 °C and 327 × g for 5 min.
14. Prepare labeled FACS tubes with 200 µL of FACS buffer. Resuspend the pellets in 150 µL of FACS buffer. Individually filter each sample into their corresponding FACS tube using a 100 µm filter. Keep on ice or at 4 °C and protected from the light until ready to analyze.
15. Analyze the cells using a flow cytometer.

8. Plaque assay for enumerating IAV

1. Prepare the required media as follows:
   1. Prepare infection medium by dissolving 2.5 g of bovine serum albumin (BSA) into 40 mL of DMEM while stirring at 37 °C for 10-20 min until dissolved. Filter-sterilize into 460 mL of DMEM.
   2. Prepare 2.4% microcrystalline cellulose by dissolving 1.2 mg of microcrystalline cellulose into 50 mL of H₂O. Autoclave on the liquid setting and store at room temperature.
   3. Prepare 5% of BSA DMEM by dissolving 2.5 g of BSA into 40 mL of DMEM while stirring at 37°C for 10-20 min. Add the remaining 10 mL of DMEM for a final volume of 50 mL. Filter-sterilize and store at 4 °C.
   4. Prepare 2x MEM/0.5% BSA by mixing 1 mL of 5% BSA DMEM with 9 mL of 2x MEM.
   5. Prepare low-viscosity overlay medium by mixing a 1:1 ratio of 2.4% microcrystalline cellulose and 2x MEM/0.5% BSA with 1 mg/mL TPCK (inhibitor of chymotrypsin) trypsin.
   6. Prepare EMEM/10% FBS by mixing 450 mL of Eagle's Minimum Essential Medium (EMEM) with 50 mL of heat-inactivated FBS.
2. Grow the Madin-Darby canine kidney (MDCK) cell line. Add the cells from one purchased vial to 5 mL of EMEM/10% FBS in a T-25 tissue culture-treated flask. Incubate for 3-5 days at 37 °C/5% CO₂ until the cells reach 100% confluence. Check for confluency as in step 2.3.
3. Remove and discard the culture medium, and rinse 2x with 5 mL of room-temperature PBS. Add 1 mL of trypsin-EDTA to the flask and incubate at 37 °C/ 5% CO₂ for 10-15 min until the cells detach. Once lifted, neutralize with 4mL of EMEM/10% FBS to obtain a cell suspension at 2 × 10⁵ cells/mL.
4. Seed the MDCK cells in a 12-well tissue culture-treated plate by adding 1 mL of resuspended cells per well (at 2 × 10⁵ cells/well) 1 day before beginning the plaque assay.
   NOTE: Make sure the cells reach 100% confluency prior to use and incubate for longer if needed to reach confluency.
5. For use as standards, make 10-fold serial dilutions (10⁶-10¹) of IAV stock (of a known titer) in the infection medium listed in step 8.1.1. Make 1.2 mL of each dilution to test in triplicate.
6. Thaw the organ homogenates on ice. Spin down on a tabletop centrifuge at 2,000 × g and collect the clear supernatant.
7. Repeat step 8.5 but with the supernatant from the samples in step 8.6.
8. Aspirate the medium from the cells and wash 2x with 1 mL of PBS to remove all the FBS.
9. Add 300 µL of each standard dilution or serially diluted sample gently along the side of each well, beginning at the highest dilution to the lowest, and do this in triplicate.
10. Place the plates in the incubator at 37 °C/5% CO₂, shaking the plate every 10 min for a total of 50 min. Make sure to place them flat in the incubator and do not stack them.
11. After the 50 min, wash the cells 2x with 1 mL of PBS.
12. Add 2 mL of the low-viscosity overlay medium into each well except the lowest-dilution and no-virus wells; to those, add infection medium and trypsin.
13. Place the plate back into the incubator at 37 °C/5% CO₂ for 2-4 days to achieve plaques that can be visualized by the naked eye.
14. Wash the plates by adding 2 mL of PBS into each well fast from the side and gently shake to suspend the settled low-viscosity overlay medium.
15. Discard the whole liquid volume in the well by gently pipetting the medium off.
16. Repeat the wash one more time with 2 mL of PBS in each well, and then discard the whole liquid volume by gentle pipetting.
17. To fix the plaques, add 500 µL of 4% paraformaldehyde into each well, shake, and let sit for 30 min.
18. Wash slowly down the side with 1 mL of PBS; then, gently discard the liquid.
19. Add 500 µL of 1% crystal violet (diluted in water) to each well to cover the cell monolayer. Incubate for 5 min.
20. Wash with 1 mL of tap water. Make sure to discard all the liquid in the well by gentle pipetting. Place the plate upside down on a diaper pad to dry overnight.
21. Count the plaques visually and save the images on any available imager.