Assessing Bacterial Invasion of Cardiac Cells in Culture and Heart Colonization in Infected Mice Using Listeria monocytogenes

1. Storage and Culture Conditions for L. monocytogenes Strains

1. Prepare solid media by autoclaving brain-heart infusion agar (BHI) and pouring the molten media into petri plates. Allow the plates to dry overnight at room temperature, then overnight again at 37 °C.
2. Using a sterile loop, obtain a small sample of L. monocytogenes from either previously made freezer stocks (bacteria suspended in 20% glycerol in BHI liquid media, stored at -80 °C) or from an agar plate containing colonies previously struck for isolation.
3. Using aseptic technique, streak the sample for single colony isolation. Be sure to sterilize the loop between cross streaks to prevent excess carry-over and ensure the isolation of individual colonies of the strain being assessed. Incubate the plate(s) overnight at 37 °C.
4. Using a sterile loop, remove one colony of the isolate and inoculate 2 ml of BHI broth (without agar) in a 14 ml polystyrene tube.
5. For assays and infections, incubate the broth culture statically (without shaking) overnight in a 37 °C incubator. For stocking of cultured isolates, incubate the broth culture at 37 °C with shaking at 180-200 rpm overnight. Cultures can be stocked by adding glycerol to a final concentration of 20% and storing sealed tubes at -80 °C indefinitely.

2. Storage and Culturing Conditions of H9c2 Cardiac Myoblast-like Cells

1. Purchase or obtain a frozen stock of H9c2 cells, as well as Dubelco’s Modified Eagle’s Medium (DMEM) containing high glucose and pyruvate, Fetal Bovine Serum (FBS), L-glutamine (Glut), and a Penicillin-Streptomycin-Glutamine mixtures (PSG).  FBS, Glut, and PSG should be stocked at -20 °C, while DMEM can be stored at 4 °C. It is helpful to aliquot FBS into 50 ml conical tubes prior to freezing.
2. In a laminar flow hood, thaw two aliquots of FBS. Add one aliquot to one 500 ml bottle of DMEM, making a 10% FBS/DMEM mixture. To this, add 6 ml of Glut and stock the resulting solution at 4 °C. This solution can be labeled “DMEM without Antibiotics.” (DMEM –Ab)
3. To a second 500 ml bottle of DMEM, add the other 50 ml aliquot of FBS. To this solution, add 6 ml of PSG and stock at 4 °C. This solution can be labeled “DMEM with Antibiotics.” (DMEM +Ab)
4. While in the laminar hood, remove 10 ml of DMEM +Ab and place in a 25 ml tissue culture flask.
5. Leaving the flask in the laminar hood, remove a frozen aliquot of H9c2 cells from storage in liquid nitrogen and quickly place the tube in a 37 °C water bath until the culture has thawed.
6. Spray the tube with 70% ethanol to sanitize, then return to the hood. Working quickly to minimize cell time in concentrated DMSO, add the full aliquot of cells to the 10 ml of DMEM +Ab.  This dilutes the DMSO sufficiently for overnight cell viability and adherence.
7. Place the flask in a tissue-culture incubator at 37 °C, 5% C0₂, and 95% humidity overnight.
8. The following morning, check the cell layer to ensure adherence. Cells will likely be between 80-100% confluent by this time.
9. In the tissue-culture hood, remove the media contained within the flask using the vacuum, leaving only the cells within the flask.
10. Add approximately 2 ml of 0.25% Trypsin-EDTA to the cells and rock gently for approximately 3 sec.
11. Invert the flask such that the trypsin solution is no longer in contact with the monolayer, and remove the trypsin by vacuum.
12. Add a second 2 ml aliquot of trypsin to the cells, and move the flask to an inverted microscope. Observe the cells while gently rocking the flask to ensure the monolayer disperses.
13. Once the monolayer is dispersed, immediately return to the tissue culture hood and add 4 ml of DMEM +Ab to the cell suspension.
14. Remove a 2 ml portion of the cell suspension, and add it to a 50 ml tissue culture flask containing 23 ml of DMEM +Ab. Rock the flask gently, then place it in the tissue-culture incubator in the conditions listed in step 2.5.
15. Check the cells daily until they reach approximately 80% confluence (approximately 2.0-4.0 x 10⁵ cells per ml). It is very important that cells do not become too confluent, as they may differentiate into skeletal muscle myotubes when starved of FBS for prolonged periods of time¹⁴ and this differentiation can alter bacterial invasion.  Monitor the cells carefully and start with a new tube of cells if a culture becomes confluent.
16. Once the cells reach 80% confluence, they can either be passaged into another 50 ml flask as just described, or prepared for invasion assay.

3. Preparing H9c2 Cells for Invasion Assay

1. In the tissue culture hood, remove the media from a 50 ml flask of H9c2 cells that have reached 80% confluence using vacuum aspiration.
2. Add 4 ml of 0.25% Trypsin-EDTA solution to the monolayer and remove immediately using vacuum aspiration.
3. Add another 4 ml portion of 0.25% Trypsin-EDTA to the flask and move the flask to an inverted microscope to observe the dispersion of the monolayer.
4. Once the cells have dispersed, return to the hood and add 4 ml of DMEM –Ab to the cells. Pipetting the solution up and down ensures complete removal of cells from the bottom of the flask.
5. Following resuspension of the monolayer, remove all of the suspension and place it in a 50 ml conical tube.
6. Remove a 20 ul portion of this suspension and count the number of cells per milliliter using a hemocytometer.
7. Once the concentration of cells in the solution is determined, adjust the concentration to 2.25 x 10⁴ cells/ml by adding the appropriate volume of cell solution to fresh DMEM –Ab. The total volume of the adjusted solution should be 25 ml.
8. While in the hood, sterilize glass coverslips by dipping them in 90% Ethanol and briefly passing each coverslip through a flame. Add each coverslip to an individual well of a 24-well tissue culture treated plate directly after it is sterilized.
9. After all wells have an individual coverslip, use a 25 ml pipet to add 1 ml of the diluted cell solution to each of the wells in the plate, and push each coverslip down with a sterile needle.
10. Check each well using an inverted microscope to ensure similar amounts of cells are in each well. Put the 24 well plate in the tissue culture incubator overnight at 37 °C in 5% CO₂ and 95% humidity.
11. The following morning, view each well to ensure that the coverslips have comparable amounts of adherent myoblasts and that the coverslips are not floating in the well. 

4. Preparing Strains of L. monocytogenes for Invasion Assays

NOTE: All laboratory work is carried out in accordance with CDC Biosafety Level 2 guidelines.

1. After preparing the 24-well plate for assay, use a sterilized loop to remove single colonies of the strains to be examined for bacterial invasion and inoculate each into individual 14 ml tubes containing 2 ml of BHI broth.
2. Place the inoculated tubes into a 37 °C incubator tilted at 45° and incubate statically (without shaking) overnight.
3. The following morning, remove the culture tube from the incubator and vortex briefly to ensure uniform suspension of the bacteria.
4. Measure the optical density of the culture using a spectrophotometer at 600 nm wavelength. Be sure to zero the spectrophotometer using sterile BHI broth before reading the optical density.
5. For L. monocytogenes, the optical density is directly related to the concentration of bacteria per ml, such that a density of 1.000 = 1.0 x 10⁹ CFU per ml.
6. Calculate the amount of culture needed to infect 2.25 x 10⁴ cells with 2.25 x 10⁶ (MOI = 100).
7. Remove the amount of culture needed and place it in a 1.5 ml centrifuge tube.
8. Spin the culture at 19,000 x g for 3 min, then discard the supernatant. Tap the open end of the tube against a paper towel in order to remove excess media stuck to the lip of the tube.
9. Resuspend the bacteria in 1 ml of sterile PBS, and vortex to ensure sufficient mixing. This mixture should contain approximately 2.25 x 10⁶ CFU per 20 ul (1.125 x 10⁸ CFU/ml).

5. Performing the Invasion Assay

1. The day before the invasion assay, prepare 24-well plates of H9c2 cells as described in Section 3, and also inoculate sterile BHI broth with the desired strain(s) of Listeria as described in Section 4.
2. Using the protocol in Section 4, prepare PBS-resuspended cultures of the strains to be assessed for invasion. Note: Infectious titer can be assessed at this point by plating dilutions of each sample onto solid media and enumerating colonies after overnight incubation.
3. Load at least 20 ul of the PBS-bacteria solution into individual wells of the 24-well plate. Note that at least three individual wells are needed for each strain in order to obtain results in triplicate, so up to 8 strains can be assessed in tandem.
4. After loading the wells with bacteria, gently rock the plate to encourage homogenous spread of each sample within the well, and place the 24-well plate back in the incubator for 45 min of incubation at 37 °C and 95% humidity, with 5% CO₂.
5. During this incubation, prepare an aliquot of DMEM – Ab (Section 2) by simply removing 25 ml of DMEM –Ab and placing it in a 50 ml falcon tube (or equivalent).
6. Add gentamicin to the 25 ml DMEM –Ab aliquot to a concentration of 15 ug/ml (7.5 ul of a 50 mg/ml stock solution). Place the DMEM –Ab +Gent solution in a 37 °C waterbath for the duration of the 45 min incubation.
7. Aliquot 25 ml of PBS into a 50 ml falcon tube and place in the 37 °C waterbath during the 45 min incubation.
8. After the 45 min incubation is complete, remove the 24-well plate and place it in the hood. Remove the media containing bacteria from each well by vacuum aspiration, changing glass pipet tips between different strains.
9. Add approximately 1 ml of PBS to each well in order to wash loosely-adherent bacteria from the surface of the cells, then remove the PBS using vacuum aspiration.
10. Following complete removal of PBS, add 1 ml of DMEM –Ab +Gent to each well. The gentamicin will only kill those bacteria that remain outside of the cells, thus those which are intracellular will remain viable.
11. Place the 24-well plate back into the incubator and allow to incubate for an additional hour.
12. During the hour long incubation, prepare 14 ml polypropylene tubes by adding 1 ml of sterile ddH₂0 to each tube. One tube will be required for each coverslip, so for one 24-well plate, prepare 24 tubes total.
13. Following the hour long incubation, remove the 24-well plate from the incubator and place it in the hood, along with the tubes prepared in step 5.12.
14. Using sterile tweezers, remove each coverslip from its respective well and place it immediately in an individual tube. Be sure to dip the tweezers in ethanol and flame them between each coverslip in order to prevent contamination.
15. After all of the coverslips have been removed and placed into individual tubes, discard the 24-well plate.
16. Return to the bench and vortex each tube for 5-10 sec.
17. Remove a portion from each tube and place each sample in an individual well of a 96-well plate, then serially dilute each sample using 1:10 dilutions up to a 1:1,000 dilution.
18. Using prewarmed and dry LB plates, spot plate each of the dilution series onto the agar plates. A multichannel pipet can be used to plate 5-10 μl spots from each sample’s dilution series.
19. Also remove 20 μl samples from each undiluted well and spot plate this amount directly onto LB agar on a separate plate from the dilution series. (This larger volume can be used to enumerate poorly invasive strains).
20. After all of the samples have been spot plated, discard the 14 ml tubes containing the coverslips. Parafilm the 96-well plate and place it in a freezer box at -80 °C for replating, if necessary.
21. Allow the spots to dry completely. This process is hastened greatly by placing the plates in the hood and removing the lids.
22. After the spots have dried, place the plates in a 37 °C incubator overnight.
23. The following morning, count the number of colonies in each spot of the dilution series for which colony numbers can be easily assessed (between approximately 5 and 50 colonies) (Figure 1), and use the series to calculate the number of bacterial per coverslip for each strain assessed. 

6. Preparing Strains of L. monocytogenes for Mouse Infections

1. The night before infection, use a sterilized loop to remove single colonies of the strains desired and inoculate each into individual 14 ml tubes containing 2 ml of BHI broth.
2. Place the inoculated tubes into a 37 ˚C incubator tilted at 45˚ and incubate statically (without shaking) overnight.
3. The following morning, remove the culture tube from the incubator and vortex briefly to ensure a uniform suspension of the bacteria.
4. Measure the optical density of the culture using a spectrophotometer at 600 nm wavelength. Be sure to zero the spectrophotometer using sterile BHI broth before reading the optical density. (For L. monocytogenes, the optical density is directly related to the concentration of bacteria per ml, such that a density of 1.000 = 1.0 x 10⁹ CFU per ml)
5. Calculate the amount of culture needed to infect each animal. Injections typically contain 200 μl of PBS with 10,000 CFU of an individual strain, which translates to a desired concentration of 5.00 x 10⁴ CFU/ml.
6. Remove a 5 μl aliquot of each final dilution and spread-plate it directly onto LB agar.  Incubate overnight at 37 °C to confirm the concentration used for inoculation.
7. Inject each CFU suspension within 1 hr of diluting (see below).

7. Inoculating L. 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 in 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 7.21 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 organ remain. Use tweezers to remove any large debris from the probe.
26. Repeat the cleaning process described in step 7.22
27. Repeat the homogenization process for the other livers, making sure to wash the probe between each liver using the process described in step 7.22.
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.