This article describes a method for visualizing an attenuated strain of pathogenic bacteria in a mouse model using bioluminescence. The process involves preparing bacterial cells, injecting them into mice, and monitoring the bioluminescent signal to assess infection clearance.
Begin with a suspension of an attenuated strain of a pathogenic bacterium.
The attenuated strain displays reduced pathogenicity due to targeted deletions in virulence genes.
The bacteria are chromosomally labeled with a bioluminescent operon that enables autonomous light production.
The bacteria express the bioluminescent operon, producing a luciferase enzyme and an enzyme complex that synthesizes the luciferase substrate.
Luciferase oxidizes the substrate to produce visible light.
Centrifuge the suspension and discard the supernatant.
Wash the cells with a buffer, centrifuge again, and discard the supernatant to remove residual non-cellular components.
Resuspend the cells in the buffer to achieve an optimal bacterial concentration and draw the suspension into a syringe.
Inject the suspension intraperitoneally into a mouse, then transfer it to a cage.
Visualize the bioluminescent signal at regular intervals.
The attenuated strain remains localized, and the bioluminescence gradually decreases, indicating clearance of infection.
On the morning of injections, thaw the cryovials of bacterial cells at four degrees Celsius for three to four hours. Keep the vials on ice after thawing and inject the mice within two hours. Transfer the contents of each cryovial to a new two milliliter tube and centrifuge it at 4,500 times G for 10 minutes.
Discard the supernatant and re-suspend the cell pellet in one milliliter of PBS. Repeat the centrifugation and re-suspend the cells in PBS to a final concentration of 2.5 times 10 to the ninth colony forming units per milliliter. Take three samples from the final suspension of each strain to validate concentration, genotype, and phenotype.
For each strain, aliquot 1.5 milliliters of the cell suspension into a two milliliter tube and prepare the PBS for control injections. Gather the mice and materials needed for injections in a sterile animal surgical room and wipe all surfaces with sanitizing wipes prior to starting. Wear two pairs of latex gloves to limit the risk of puncture, if bitten, as well as a lab coat, safety glasses, and face mask.
Remove the mouse from the cage and weigh it, marking the tail with permanent marker for post-injection tracking. Open a new one milliliter syringe with a 27 gauge needle and draw up 200 microliters of sterile PBS. Grab the mouse behind its ears, using the thumb and forefinger, and pinch to create a skin fold at the nape of the neck.
Then, secure the tail into the palm using the pinky to hold the mouse flat and immobile. Insert the needle at a 30 degree angle into the peritoneal cavity to the left or right of the midline. Slightly lift the needle to ensure that it was not inserted into organs.
Then, slowly inject the PBS and withdraw the needle. A bolus at the injection site is typical. Place the needle in the designated sharps disposal container and move the mouse to a separate cage.
Repeat the procedure with the next mouse and after all mice from one cage are injected, move them back to their original cage. After injecting the control group, inject the test groups using the same procedure. Once all injections are complete, return the mice to the housing room and clean the work area with sanitizing wipes.
Set the oxygen flow to 1.5 liters per minute and isoflurane to 3.5% and move the mouse to the anesthetic chamber, then to the temperature stabilized stage, following anesthesia. Position the mouse on its back with arms outstretched and fit a nose cone for administration of 2.5% isoflurane during imaging.
Close the door and take bioluminescent images and x-rays of the mouse. When imaging is complete, return the mouse to its cage and monitor it. It should regain consciousness within three to five minutes.
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