To visualize internalized bacteria, first, aseptically transfer autoclaved glass coverslips inside each well of a 12-well plate. Plate five times 10 to the four HUVECs per well on the coverslip, and incubate for 24 hours. To label bacteria, centrifuge mid-log phase bacteria, and resuspend the pellet with anaerobic PBS. Centrifuge again, and repeat the washing step.
Add 20 microliters of 0.2 millimolar BCECF-AM to two milliliters of anaerobic PBS containing five to seven times 10 to the eighth bacteria per milliliter. Incubate at 37 degrees Celsius for 30 minutes in the dark.
The fluorescent dye we use can stain most anaerobic bacteria. Thus, no other reagents, such as a specific antibody, are required.
Centrifuge the bacterial suspension at 5,000 times g for 10 minutes to remove the unbound labeling reagent, and resuspend the bacteria in VEGF medium. Then, infect the HUVEC cells at a ratio of 100 to 1, bacteria to cells, and incubate at 37 degrees Celsius for 30 minutes. Following incubation, remove the infection medium, and wash the cells three times with PBS. Then, add 4% paraformaldehyde, and incubate at room temperature for 10 minutes to fix the cells in the anaerobic chamber.
Remove the fixed cells from the chamber, and then, wash the coverslips with PBS three times. Incubate the coverslips in one milliliter of 0.2% Triton X-100 for 10 minutes to permeabilize the cells. Then, wash the coverslips with PBS three times. Now, add 50 microliters of 50 micrograms per milliliter of TRITC phalloidin to each coverslip, and incubate for 45 minutes at room temperature to stain cellular oligomeric actin.
After incubation, wash the coverslip with PBS three times. Invert the coverslip on a glass slide with soft-set mounting medium containing one microgram per milliliter DAPI for nuclear staining. Then, seal the periphery of the coverslip with nail polish, and observe under confocal microscopy.
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