Evaluation of Vaccine-Induced Immunity Against Bacterial Infection in a Mouse Model

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

1. Immunization of Mice

1. Prepare a master vaccine mix in a sterile physiological buffer such as saline or DPS.
   NOTE: The total volume in the mix should include 10% more than that required for immunization of the whole group. Concentrations of the vaccine composition are detailed below in steps 1.1.1 and 1.1.2. Transport the items to the vivarium on ice.
   1. For B. pertussis studies, include the following vaccine groups: acellular pertussis vaccine (aPV) and aPV + Bordetella colonization factor A (BcfA). In addition, use alum, vaccine antigens alone (filamentous hemagglutinin (FHA), and pertactin (Prn)), and naïve (non-immunized) mice as control groups.
      NOTE: aPV is 1/5^th the human dose of an FDA-approved aPV, which is composed of tetanus toxoid, reduced diphtheria toxoid, pertussis toxoid (PT), filamentous hemagglutinin (FHA), and pertactin (Prn) adsorbed to aluminum salts. One human dose of current pertussis aPV is 0.5 mL, therefore, 1/5^th of a dose is 0.1 mL. Volumes for aPV + BcfA are 0.1 mL of the aPV (1/5^th the human dose) + 0.046 mL of BcfA (30 µg) per mouse. The maximum volume that can be safely delivered to one muscle is 0.1 mL. Because the aPV + BcfA vaccine volume is greater than 0.1 mL, the vaccine must be delivered into both shoulders, divided roughly equally, in order to be administered safely.
   2. To prepare an experimental vaccine, for one dose combine 1 µg of FHA and 0.5 µg of Prn with 50 µg of aluminum hydroxide gel in sterile PBS. Allow the experimental aPV to mix on a laboratory roller for 15 min at room temperature (RT). Afterward, spin the tube at 18,000 x g for 5 min at 4 °C. Remove the supernatant and resuspend the contents in 1 mL of sterile PBS.
2. In the vivarium, set up the anesthesia machine (Figure 1).
   NOTE: Ensure there are adequate levels of oxygen and isoflurane in their respective tanks before use. Weigh the isoflurane scavenger canister and replace it when its weight increases by 50 g.
3. Thoroughly clean the biological safety cabinet (BSC) and place the clean induction chamber inside the BSC. Connect the anesthesia and scavenger lines from the anesthesia machine to the induction chamber. Open the oxygen tank via the regulator to ensure oxygen is flowing and set the level to 1.5-2 L/min.
4. Place mice of desired age (6-12 weeks) into the induction chamber. Turn on the isoflurane to 2.5-3% in order to lightly anesthetize the mice. Monitor the animals until they are not moving in the chamber and their breathing has slowed.
5. Test the level of induction by toe-pinching, observing for any reflexive movement. If there is none, then the mice are ready to inject.
   NOTE: In this experiment, the whole cage of animals was anesthetized at once. One can choose to anesthetize animals individually for injection or to inject the animals without anesthesia. Either of these methods is appropriate.
6. Meanwhile, prepare 1 mL insulin syringes (28G1/2) with 0.1 mL of vaccine each. When animals are fully induced, remove one animal from the chamber and lay the mouse ventrally on a towel or bench pad.
7. Administer the vaccine intramuscularly. With the syringe at a 45° angle and with the bevel facing up, insert the syringe ~5 mm into the deltoid of the mouse and inject the vaccine.
   NOTE: The hind quarter/flank may be used as an injection site instead of the deltoid.
8. After injection, keep the needle inserted in the muscle for ~5-10 s. Once the volume is delivered, rotate the syringe at 180° so that the bevel faces away to create a seal and prevent the leakage of the vaccine. Then, slowly pull the needle out of the injection site.
9. Return the mouse to the cage. Repeat the procedure with all animals in the designated group.
10. Turn off the isoflurane and flush the induction chamber with oxygen before anesthetizing the next cage of mice.
11. Monitor the animals until they are all alert and moving in the cage. Return the cage to the housing location.
12. Monitor animals every 12 h post-vaccination, up to 48 h, for clinical symptoms of morbidity such as rough/unkempt coat, slow gait, or labored breathing. If symptoms persist, confer with the institutional veterinarian, and remove the mice from the study if needed.
13. Four weeks after initial immunization, boost mice by anesthetizing and administering intramuscular injections into the right deltoid of the mouse with the same vaccine formulation that was previously given.
14. Again, monitor the animal for any clinical symptoms. House animals for an additional 2 weeks and then proceed with infection.

2. Murine Intranasal Infection Model

1. In the vivarium, set oxygen and anesthesia conditions as described in steps 1.3 and 1.4. As described in step 1.5, place the mice in the chamber to anesthetize. Monitor the animals until anesthesia takes effect.
2. When the mice are anesthetized, remove one animal at a time from the induction chamber. Pick up the mouse by the scruff of the dorsal thorax, behind the shoulder blades and neck. Position the mouse upright to access the nose.
3. Using a 200 µL pipette tip, slowly apply 40-50 µL of the B. pertussis bacterial inoculum to the nares of the mouse (20-25 µL in each nare). Hold the mouse until the entire inoculum has been inhaled by the animal. If some of the inoculum bubbles out, collect the bubbles and re-instill them into the nares. Deliver an equal amount of sterile PBS to one group of mice as a negative control.
4. As done in steps 1.10-1.12, return the inoculated animals to their cage and monitor the animals until they are alert and moving. Return the cage back to its original housing space on the rack. Flush the induction chamber with oxygen to remove the residual isoflurane before anesthetizing the next set of animals. Monitor the animals for 48 h post-infection.
5. In the laboratory, plate serial dilutions of the inoculum, in sterile PBS, to verify the actual colony-forming units (CFUs) delivered to the mice. Plate 0.1 mL of dilutions on 10% BG + Sm plates (Bordet-Gengou agar plates supplemented with 10% defibrinated sheep's blood and 100 μg/mL streptomycin) (Figure 2A). Place the plates in the 37 °C incubator and grow for 4 days. Count and calculate the CFUs from the inoculum delivered to the mouse (Figure 2B).

3. Harvesting of Animal Tissue after Infection

1. Prepare for mouse dissection and tissue harvest.
   1. Sterilize all tools (coarse and fine scissors, curved scissors, fine forceps, pellet pestles, and triangle spreaders) needed for processing tissues in an autoclave. Seal the tools in sterilization pouches. Wrap the glass dounce homogenizers in foil and add autoclave tape to indicate sterility.
   2. Prepare agar plates 1 or 2 days prior to the tissue harvest to ensure that plates are solidified prior to use. For B. pertussis, use 10% BG + Sm. Label plates for the lung for each mouse with the desired dilutions, determined empirically for each experiment.
   3. Fill and label CFU dilution tubes. Add 0.9 mL of sterile PBS to sterile 1.5 mL microcentrifuge tubes based the on number of organs to process and dilutions per organ.
   4. Fill and label tubes for tissue collection:
      1. Lungs: add 2 mL of sterile 1% casein in PBS to a sterile 15 mL conical tube and store at 4 °C. To harvest lungs for histology, add 2 mL of 10% neutral buffered formalin (NBF) and store it at RT.
   5. Fresh 70% ethanol should be prepared to fill beakers and spray bottles for dissection.
2. Transport all materials to the vivarium on a cart on the day of harvest. Clean the BSC and set up the anesthesia machine as in steps 1.3 and 1.4.
3. Place the mice to be dissected in the induction chamber and, with oxygen flowing, turn on the isoflurane to 5% and wait until the animals are unconscious. Remove mice one at a time, and cervically dislocate the animal as a secondary method to ensure death. Following the cervical dislocation, euthanasia should be confirmed by the absence of a respiratory rate and/or absence of the withdrawal reflex (toe pinch test).
4. Position the mouse, ventral side facing up, on the dissection board and pin the arms and legs open. Spray down the body of the mouse with 70% ethanol.
5. Using forceps, clasp the skin below the abdomen, in the center of the pelvis. Using sharp scissors, make a vertical cut up to the mandible. Then, dissect the skin away from the peritoneal wall by pushing the two layers apart, using small motions with the scissors closed. Place the skin to the sides of the carcass to create an unobstructed field for sterile organ harvest.
6. Use forceps to stabilize the rib cage at the xiphoid process and cut open the diaphragm. Lungs should deflate and fall towards the spine. Cut open the rib cage at the sides to remove the breastplate.
7. Isolate and remove the superior lobe of the right lung, cutting the pulmonary arteries and veins. Place the lobe in a 15 mL conical tube containing 10% NBF and place the tube at RT for at least 24 h to fix the tissue. Isolate the remaining lobes of the right lung and the left lung. Place lobes in a 15 mL conical tube containing 2 mL of 1% casein in PBS and place it on ice.
8. Dispose of the carcass in a biohazard bag. Before dissecting the next animal, rinse the tools with deionized water, and then place them in a beaker filled with 70% ethanol. Remove the tools, shake off excess ethanol, and then proceed with the next dissection.
9. Dissect each mouse following the steps mentioned above.
10. Process the tissues as described below.

4. Processing of Lungs

1. Transfer the lungs (in 2 mL of 1% casein in PBS) into a sterile, 15 mL Dounce homogenizer. Use a pestle to homogenize tissue. Dissociate until no large particles of tissue remain. Place the homogenized suspension back in the original 15 mL tube.
2. Remove a 0.3 mL aliquot for plating CFUs and centrifuge the homogenate at 450 x g for 5 min at 4 °C. Collect and aliquot the supernatant in 0.5 mL aliquots into pre-labeled microcentrifuge tubes. Store the samples at -20 °C until the analysis of cytokines by ELISA.
3. Prepare chosen dilutions by serially diluting 0.1 mL of the homogenized lung suspension in dilution tubes (0.9 mL of sterile PBS). Plate 0.1 mL of empirically chosen dilutions onto pre-labeled 10% BG + Sm plates and spread using a sterile triangle spreader.
4. Incubate the plates at 37 °C in room air for 4 days.
5. Count and calculate CFUs/lung (Figure 3). Briefly, multiply recovered CFU numbers by the dilution factor of the plate, then also by the total volume in which the organ was processed. The CFU calculations are then transformed into Log₁₀ values for each animal and graphed.
   1. Plates with 30-300 colonies are easily and accurately counted. Plates with less than 6 colonies should not be used for calculations since such low colony numbers introduce errors. To obtain a lower limit of detection, the total volume in which an organ was homogenized is divided by the volume used to spot onto a plate from the undiluted homogenate (e.g., 2 mL total volume divided by 0.1 mL of undiluted homogenate equals the lower limit of 20 CFUs detectable).