Quantifying Bacterial Load in the Mouse Heart Following Tail Vein Injection of Listeria monocytogenes

Abstract

Source: McMullen, P. D., Freitag, N. E. Assessing Bacterial Invasion of Cardiac Cells in Culture and Heart Colonization in Infected Mice Using Listeria monocytogenes. J. Vis. Exp. (2015)

This video demonstrates the injection of Listeria monocytogenes into the tail vein of a mouse to establish systemic infection. It shows bacterial circulation, organ colonization, and the quantification of bacterial load in the heart by homogenizing tissue, performing serial dilutions, and plating.

Protocol

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

1. Inoculating Listeria monocytogenes into Mice via Tail Vein Injection and Assessing Bacterial Burden within the Liver, Spleen, and Heart

NOTE: All animal work is carried out in accordance with CDC Biosafety Level 2 guidelines. Mice are typically ordered one week in advance and caged together with 5 animals per cage. Mice are allowed to acclimate to the new laboratory environment for four days before injection. These experiments used 6-8 week old female Swiss-Webster mice that were housed 5 to a cage in a barrier environment and fed a non-restricted diet.

1. Prepare one cage of mice at a time by removing the lid and food/water bins, and placing the cage under a heat lamp for five minutes. This process allows for the tail veins to dilate, making injections easier to perform.
2. Remove one animal and place it in a harness (or falcon tube with the end cut off) in order to restrain the animal during the injection.
3. Clean the injection site using an alcohol pad, and allow the alcohol to evaporate. This not only cleanses the site, but also further dilates the tail vein.
4. Using a 1 ml syringe equipped with a 27.5 gauge needle, gently inject the mouse with 200 μl of the desired culture into the tail vein.
5. Use a gauze to stop any bleeding which may occur as a result of the injection, and place the animal in a fresh cage.
6. Repeat this process for the remaining animals and strains. Be sure to label each cage of animals with the strain they have received.
7. Check on the animals daily, removing and sacrificing any animals which appear to be ill. If no animals exhibit signs of illness, allow the infections to proceed for 72 hr before sacrificing all animals.
8. To sacrifice, use CO₂ anesthesia from a bottled source until all respirations have stopped, then use cervical dislocation in order to assure that the animal is dead.
9. Following sacrifice, move the animals to a tissue culture hood for dissection.
10. Using a dissection block, pin each leg of the animal with large gauge needles and spray the animal with 70% ethanol until it’s saturated.
11. Cut the skin of the abdomen and thorax back using a Y-shaped incision that extends from the vaginal opening up to the xiphoid process, progressing then up to the axilla of each arm.
12. Pull back the skin and remove the needles from each leg to hold the dissection open.
13. Cut gently through the peritoneum, exposing the intestines, stomach, liver, and spleen.
14. By first cutting the hepatic vein and coronal ligament at the top of the liver, begin to remove the liver. Additional ligaments to be removed are found beneath the liver, connecting it to the first section of the small intestine, as well as to the musculature of the back.
15. Place the liver in 5 ml of sterile ddH₂0 in a 50 ml falcon tube.
16. Next, remove the spleen by isolating it and gently cutting away the vasculature and ligaments. The spleen will be removed more readily than the liver. Place the spleen in a separate falcon tube containing 5 ml sterile ddH₂0.
17. Locate the diaphragm and cut gently along the ribcage in order to visualize the thorax. Cut through the xiphoid process and sternum to the level of the neck in order to open the thorax, exposing the heart and lungs.
18. Using forceps, grab the heart gently by its apex, and lift it such that the aorta and pulmonary vessels are in tension. Cut these vessels to free the heart.
19. Place the heart into a 50 ml falcon tube containing 5 ml of sterile ddH₂0.
20. Discard the mouse carcass, and repeat this process for every animal, using clean falcon tubes containing 5 ml sterile ddH₂0 for each organ removed.
21. After all organs have been removed, clean the workstation and return to the bench.
22. Prepare a set of 5 tubes to be used for cleaning and sterilizing the homogenizer for each set of organs from five mice (i.e. one 5 tube set for 5 livers, 5 tubes for 5 spleens, and 5 tubes for 5 hearts). Four of each of the tubes should be filled with 30 ml sterile ddH₂0, and one should be filled with 30 ml 90% EtOH.
23. Using a TissueMaster (or equivalent homogenizer), dip the homogenizer (while running) into the tubes prepared in step 1.21 as follows to clean the probe:
24. Tube 1 (ddH₂O) 5 sec in Tube 2 (ddH₂O) 5 sec in Tube 3 (EtOH) 5 sec in Tube 4 (ddH₂O) 5 sec in Tube 5 (ddH₂O)
25. Homogenize one liver using the homogenizer for at least 2 min, or until no visible portions of the organ remain. Use tweezers to remove any large debris from the probe.
26. Repeat the cleaning process described.
27. Repeat the homogenization process for the other livers, making sure to wash the probe between each liver using the process described.
28. After all of the livers are completely homogenized, discard the wash tubes (1-5) for the liver.
29. Repeat the homogenize/clean process for both the spleen and hearts as well, making sure to use a new set of wash tubes for each series of organs. If at any point the wash tubes become turbid, discard them and replace with new tubes to finish the series.
30. After all of the organs have been homogenized, do one last cleaning cycle on the homogenizer and dry it well for storage.
31. Place approximately 200 μl of each organ into an individual well of a 96-well plate.
32. Perform serial dilutions on the organ samples by diluting 1:10 in a series up to a 1:10,000 dilution (liver and spleen) or up to a 1:1,000 dilution (heart).
33. Spot plate the dilution series onto pre-warmed LB agar. Also remove 20 ul of each undiluted organ and plate onto separate LB agar plates in order to enumerate the CFU from poorly colonized organs.
34. Discard the falcon tubes containing the homogenized organs, and wrap the 96-well plates containing the dilution series with parafilm and place it in a freezer box at -80 °C.
35. Allow the spots to dry. This process is quickened by placing the plates in the hood and removing the lids.
36. After the spots have dried, place the plates in a 37 °C incubator overnight.
37. Count the number of colonies in each spot the following morning and use the dilution series to calculate the total number of bacteria per organ.

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
Difco	Agar	214510	2 kg of Granulated Agar
In vitrogen	LB Broth Base	12975-084	2 kg of Luria Broth Base Powder (without agar)
Falcon	96-well U-bottom plate	35-3077	Non-tissue culture treated 96-well plate