Bacterial biofilms are important for human health, protecting their bacterial residents from environmental insults and anti-microbial agents. The goal of this procedure was to develop a model system to evaluate the effects of small molecule inhibitors on biofilm formation. These methods can help to establish a tool-set essential for the biofilm field to select small molecule inhibitors that specifically target biofilm formation.
The main advantage of these methodologies is that they start from a consistent, robust experimental framework, and give quantitative and qualitative insights into the mechanism of action of biofilm-specific inhibitors. Though these methods can provide insight into Bacillus subtilis biofilms, they can also be applied to other biofilm-forming bacteria, such as Pseudomonas aeruginosa or the plant pathogen Xanthomonas citri. Scanning electron microscopy can be essential to analyze the effect of small molecules on biofilm formation, because it allows observation of the extra-cellular matrix, and gives single-cell resolution.
To begin, first select an appropriate single colony, and transfer it into three-milliliters of LB broth. Place this starter culture into a shaking incubator for four hours at 37 degrees Celsius. After incubation, take 1.5 milliliters of the starter culture, and centrifuge for four minutes.
Carefully remove the supernatant, and then re-suspend the pellet in 1.5 milliliters of MSgg medium. To grow pellicles, prepare a 12-well cell culture plate by adding three milliliters of MSgg to each well. To some of these wells, add MSgg with a small molecule inhibitor in a concentration range, distributing the location of different concentrations around the dish, to avoid edge effects.
Measure the optical density at 600 nanometers of the re-suspended starter culture. The culture should be between 0.6 and one. This is critical for the robustness of the system.
Innoculate each well of the culture plate with 3 microliters of the re-suspended starter culture. Then, incubate the plate at 23 degrees Celsius for three days under static conditions. Afterwards, remove the plate from the incubator, and observe the pellicles.
To grow biofilms, use a template and symmetrically spot four separate drops of 1.5 microliters of unwashed starter culture onto an 8.5 centimeter dried 1.5%Agar MSgg plate. Let the drops absorb into the medium before moving the plate. Incubate the plates at 30 degrees Celsius for three days.
Using a binocular with homogeneous exposure of lighting, check that biofilm colonies have developed and formed a three-dimensional wrinkled structure. First, take a clean razor blade and cut the biofilm colonies into two equal parts with the help of the template. Carefully lift one half of the colony with a clean spatula, and transfer it to a 1.5-milliliter micro-centrifuge tube, containing 500 microliters of PBS.
Take the second half of the colony, and transfer it to a micro-centrifuge tube containing 500 microliters of 50%ethanol. These will be used to assess resistance to sterilizing agents. Afterwards, similarly take the first half of the D-lysine treated colony, and place it in PBS.
Then, take the second half of this colony, and transfer it to ethanol. Incubate all the tubes containing the biofilm halves for 10 minutes at room temperature. And then, centrifuge them for five minutes at 18, 000 times g.
Using a pipette, carefully remove the supernatant. Add 300 microliters of PBS, and then sonicate the cells at a low setting, with a microtip sonicator. Add an additional 700 microliters of PBS for a final volume of one milliliter.
Next, perform a serial dilution of 10 to the negative seven in PBS. Take 100 microliters of one of the dilutions, and innoculate a 1.5%Agar LB plate. Repeat this step for two other dilutions per sample.
Spread the innoculum using glass beads, and then incubate the plates overnight at 30 degrees Celsius. Examine the plates and count the colony-forming units, or CFU. After calculating the CFU per milliliter value, calculate the percentage of survivors in the PBS versus ethanol treatments.
To begin sample fixation, first prepare enough fresh fixative solution for the desired number of biofilm colonies. Carefully add five milliliters of fixative to each petri dish, and avoid pipetting directly onto colonies. Colonies will begin to detach from the Agar and float.
Carefully seal the plates with parafilm, and incubate on a rotary shaker for two hours at room temperature. Transfer the plates to four degrees Celsius storage overnight. Gently remove the fixative liquid with a Pasteur pipette, connected to a vacuum.
Add 10 milliliters of 100 nanomolar sodium cacodylate, five millimolar calcium chloride buffer to wash the biofilm, and incubate for five minutes. Carefully detach the center of the biofilm colony from the Agar plate with a Pasteur pipette. To air-dry colonies, cut cellulose filter paper into quarters, and then submerge one section in 100%ethanol.
Carefully transfer one floating biofilm colony onto the paper. Place the paper in a Petri dish lined with dry filter paper. Then, cover the dish, and leave it in a chemical hood to dry overnight.
In order to preserve the morphology of the biofilm colony, it is important to underlay the floating biofilm completely with the cellulose paper. Once on the paper, the biofilm cannot be readjusted. Coat an electron microscopy stub with carbon tape, and then use tweezers to carefully transfer the biofilm colonies onto the stub.
After connecting each colony to the stub, by adding a thin bridge of carbon tape, store the stubs in a desiccator for 24 hours or until needed. This step requires a steady and precise hand, because at this stage the biofilms are very fragile and easily fractured. The challenge is to mount a substantial part of the biofilm without cracks.
On the day of the examination, place the colonies into a gold-palladium sputter coater. Coat the samples for two minutes at a 60 degree angle. Repeat this step twice, rotating samples 120 degrees in between.
Finally, coat the samples once for three minutes from the top. Samples are now ready for imaging on the SEM. These images show pellical formation of B.subtilis grown in biofilm-inducing MSgg medium.
With the addition of the small molecule inhibitor D-lysine, pellicle development is reduced. And as the concentration of the inhibitor increases, pellicle formation shows a correlating decrease. This graph shows the effect of exposure to ethanol on the survival of cells within biofilm colonies, treated with a D-lysine inhibitor, or left untreated.
Colonies treated with D-lysine, and exposed to 50%ethanol for 10 minutes, showed a dramatic reduction in survival compared to the untreated fraction. Top-down binocular images show that colonies grown in the presence of D-lysine were smaller overall, and formed less pronounced wrinkles than those left untreated. Scanning electron microscope images of critical point dried biofilm colonies reveal yet further the altered biofilm development.
The extra-cellular polymeric substances matrix, or EPS, appeared more like a spider web in untreated samples, with treated samples showing less organization. Finally, examination at the resolution of individual bacterial cells shows that treated cells are elongated in appearance, less covered with EPS, and not as tightly connected to their neighbors as the untreated cells. While attempting this procedure, it's important to remember that setting a consistent framework to study biofilm inhibitors is essential for reproducible results.
Once mastered, a small molecule screening for biofilm inhibition can be done in less than a week if it's performed properly. Following this procedure, other methods, such as investigation of gene expression in single biofilm cells, can be performed in order to obtain additional insights into the target of the small molecule inhibitor. After watching this video, you should have a good understanding of how to analyze the overall effect of specific biofilm inhibitors on biofilm development and antibiotic resistance.
