A Method for Sample Preparation to Visualize Biofilm-Forming Bacteria on Fungal Hyphae

After growing the bacteria overnight, centrifuge the culture at 5,000 times g for three minutes, and then suspend the pellet in 25 milliliters of sterile 0.1 molar potassium phosphate buffer. After repeating the centrifugation, use the same buffer to adjust the final bacterial concentration to 10 to the ninth cells per milliliter.

Fill a 6-well microplate with 5 milliliters of the bacterial suspension. Use a sterile razor blade to cut the cellophane membrane of the fungal culture into squares, with a single colony on each membrane. Then with forceps, carefully remove cellophane squares containing hyphae from the solid medium, and transfer them to individual wells of the bacterial suspension.

Gently shake the microplate until the fungal colonies are detached from the cellophane. Then, remove the cellophane sheets leaving the fungal colonies in the plate. Incubate the microplate with gentle agitation at 20 degrees Celsius for a time depending on the strains used and the stage to be analyzed.

For P. fluorescens BBc6/L. bicolor, incubate the cultures for 30 minutes to get early-stage biofilms and for up to 16 hours for mature biofilms. To remove planktonic bacteria and bacteria electrostatically attached to the hyphae, rinse the fungus by transferring it to a new 6-well microplate filled with strong salt solution. Gently shake the plate for 1 minute.

Transfer the fungus to a new 6-well microplate containing 5 milliliters of sterile 0.1 molar potassium phosphate buffer. Gently shake the plate for two minutes, and then transfer the fungus to fresh buffer.

While keeping the fungal colony in buffer, use a scalpel to cut the colony approximately in half. Transfer half of the colony to be stained to a Petri dish containing 1 milliliter of sterile water supplemented with an appropriate fluorescent dye to visualize the fungal network, such as wheat germ agglutinin conjugated to Alexa Fluor633. Then, incubate the sample in the dark.

After staining, rinse the sample by transferring it to a Petri dish lid containing 10 milliliters of sterile 0.1 molar potassium phosphate buffer, and gently shake for 1 minute. Half submerge a slide in the Petri dish lid and delicately position the cut section to float above the slide. Then, slowly lift the side from the buffer solution allowing the sample to gently settle on the slide.

The most delicate aspect is the positioning of the sample on the slide. To avoid biofilm disruption, the sample and the slide must be immersed while positioning the sample, and the sample must not be moved anymore before imaging.

Finally, add 10 microliters of anti-fading mounting medium to the sample and cover it with a glass coverslip.