Fluorescence-Based Viability Assessment of Oral Anaerobic Bacteria

Begin with a sealed tube containing a nutrient-rich broth maintained under anaerobic conditions.

Using a syringe, transfer a small amount of the broth into another tube.

Take a blood agar plate containing colonies of an anaerobic oral bacterial strain. Pick a single colony and resuspend it in the broth.

Return the suspension to the sealed tube and incubate to promote bacterial growth.

Dilute the culture in a buffer to achieve an optimal cell concentration for cell viability assessment.

Add a solution containing a membrane-permeable dye and a membrane-impermeable dye, then incubate.

The permeable dye enters all cells and binds to intracellular nucleic acids.

The impermeable dye penetrates only dead cells with damaged cell envelopes and binds to their nucleic acids.

Analyze the sample using flow cytometry.

Dead cells exhibit dominant red fluorescence, whereas live cells show only green fluorescence, allowing assessment of bacterial viability.

To begin, autoclave the medium, and then add 5 microgram per milliliter of hemin and 1 microgram per milliliter of menadione. Dispense 9 milliliters of the medium into Hungate tubes, and close the tubes with a rubber stopper and an aluminum cap. Then flush the tubes with a mixture of 90% nitrogen and 10% carbon dioxide.

Use the syringe to retrieve 1 milliliter of anaerobic medium, and transfer it to a 1.5 milliliter microcentrifuge tube. Pick single colonies of each bacterial species and resuspend them in the medium. Then transfer the colonies into the anaerobic modified BHI, and incubate them at 37 degrees Celsius for 48 hours.

Following the incubation, use filter sterilized PBS to dilute the cultures to ten to the fifth cells per milliliter. Then use flow cytometry with the SYBR green/Propidium Iodide stain to measure the cell number. Evaluating the microbiome community composition and viability is essential for avoiding bias in our experiment.