Methodologies for Studying B. subtilis Biofilms as a Model for Characterizing Small Molecule Biofilm Inhibitors

1. Assessing the Effect of Small Molecule Inhibitors on Pellicle and Biofilm Colony Formation

1. Prepare a 2x solution of defined biofilm-inducing MSgg medium²⁵ without the calcium chloride and iron(III) chloride hexahydrate. After filter sterilization, add the calcium chloride. The medium is ready to use directly or it can be stored at 4 °C in the dark.
2. Prepare the 1x MSgg dilution on the day of the experiment.
   1. Dilute the 2x MSgg medium to 1x with sterile distilled water (pellicles) or sterile 3% hot (80 °C) agar (biofilms) and add iron(III) chloride hexahydrate to a final concentration of 50 μM (pellicles) or 250 µM (biofilms). Add antibiotics or small molecule inhibitors to the desired concentration and mix well. For example, to obtain a final concentration of 0.5 mM D-leucine in 30 ml to establish pellicles or biofilms, add 196.6 µl of a 76.3 mM (10 mg/ml) D-leucine stock solution.
      NOTE: The final 1x MSgg composition is described in Table 1. Compared to the original recipe²⁵, the medium contained 50 µg/ml threonine and the iron concentration to grow biofilm colonies on solid MSgg medium was increased 2.5x to optimize the wrinkled colony morphology.
3. After solidification of the agar, dry the solid MSgg plates in a biological hood for 30-45 min prior to the inoculation.
4. To select specific inhibitors that interfere with the mechanisms of pellicle formation (Figure 1), rule out that the concentrations used affect planktonic and static growth.
   1. Determine planktonic growth (increase in optical density over time in liquid culture) in a simple growth curve by measuring the optical density at 600 nm every hour until the stationary growth phase.
   2. To confirm that the measured culture turbidity represents live cell counts, determine the numbers of colony forming units (CFU) of cells in the planktonic growth phase from a shaking culture after several time points.
   3. To assess the effect of small molecule inhibitors on static pellicle growth, harvest cells at the end of a 3-day incubation at 23 °C from a 24-well cell-culture well, inoculated under the same conditions as described in sections 1.7-1.9 and determine the CFU. For this control, use a pellicle-deficient strain that lacks the operons encoding for the extracellular matrix components (i.e., B. subtilis ΔepsH, ΔtasA).
      NOTE: This strain is capable of growing under static conditions, but in contrast to a pellicle-forming wild type, it is deficient in the ability to float to the liquid-air interface, where growth is favored due to increased oxygen levels²⁶. Thus, this extracellular matrix- and pellicle-deficient strain is a recommended reference strain to assess growth under static conditions.
      NOTE: For the specific example of the non-canonical D-amino acid D-leucine described below, an effect on planktonic and static growth at concentrations that interfered with pellicle formation was ruled out^12,17. The methods to determine planktonic and static growth are described in detail¹⁷.

Figure 1. Conceptual overview for the identification of a robust experimental setup to assess the specific inhibition of biofilm formation. Selection criteria for small molecule inhibitors that indicate specific interference with biofilm formation without pronounced effect on planktonic growth. Please click here to view a larger version of this figure.

5. Streak out B. subtilis from a -80 °C stock (LB culture of 10⁹ cells/ml frozen in 20% glycerol) to isolate single colonies on a LB-1.5% agar plate with a sterile tip or applicator stick.
6. Grow overnight at 30 °C.
7. CRITICAL STEP: For a robust pellicle inhibition by the non-canonical D-amino acids such as D-leucine, grow a single colony picked from the LB-1.5% agar plate in 3 ml LB broth at 37 °C for 4 hr in a shaking incubator (shaking speed 200 rpm). Replace the LB broth with biofilm-inducing MSgg medium prior to inoculation by centrifuging 1.5 ml starter culture for 4 min at 6,000 x g, carefully removing the supernatant and re-suspending the pellet in 1.5 ml MSgg medium. The rest of the culture can be discarded.
   Important: To ensure the robustness of the system, the optical density at 600 nm (OD₆₀₀) of the washed starter culture should be between 0.6 and 1.
8. During the growth of the starter culture, prepare a 12-well cell-culture multidish plate containing 3 ml of MSgg medium without or with a concentration range of the small molecule inhibitor (e.g., 0.3, 0.5, 1 mM D-leucine¹⁷). To rule out edge effects, distribute the location of the different concentrations across the multidish plate. Alternatively, use 24-well cell-culture multidish plates containing 1.5 ml of MSgg medium.
9. Inoculate the wells of the 12-well cell-culture multidish plate with 3 µl of the washed starter culture (1:1,000 dilution).
   NOTE: A lower dilution ratio, i.e., 1:500 can be used. This decreases the development time of the pellicles.
10. Grow the pellicles at 23 °C under static conditions for three days. Do not move the pellicles during this time, as it can affect the final surface morphology of the pellicle.
11. Acquire pictures with a binocular and homogenous exposure of lightning. Alternatively, take a picture of the pellicles with a high resolution camera. To avoid artefacts caused by inconsistent light angles and shadows, take top-down pictures with the camera fixed on a tripod and use a soft and large light source at 45° from both sides.
    NOTE: An alternative method to study B. subtilis multicellularity is the biofilm colony assay on solid, biofilm-inducing MSgg medium. Like pellicles, this assay allows the study of spatiotemporal processes. Once the active range of small molecule inhibitors is determined, their effect on biofilm colony formation can be studied.
12. To grow biofilm colonies, symmetrically spot 1.5 µl of the unwashed pre-culture (Step 1.7) on the dried MSgg 1.5% agar plate with the help of a template — 4 drops per Petri dish of 8.5 cm diameter. Let the drops adsorb to the plate before moving them.
    NOTE: The template helps to get an equal distribution of the biofilm colonies within the area where the cells are grown. To prepare the template, draw the total area of growth at original scales, divide it to equal sectors and mark the center. For a round Petri dish of 8.5 cm diameter, this assigns 14 cm² to one biofilm colony.
13. Incubate the plates at 30 °C for three days. During this time, biofilm colonies develop and form a three-dimensional, wrinkled structure.
14. Take pictures as in step 1.11.

2. Ethanol Resistance Assay

1. Grow biofilms as described in steps 1.1-1.7 and 1.12-1.14.
2. After 68 hr of growth at 30 °C, cut the biofilm colonies into two equal parts with the help of a razor blade and the template.

Figure 2. Example experimental design to assess the resistance of biofilm colony cells to sterilizing agents. (A) Template used for the equal distribution of biofilm colonies across a Petri dish and for cutting. (B) Top-down images of untreated wild-type biofilm grown for 68 hr on solid, defined biofilm-inducing MSgg medium at 30 °C. The enlargement shows how a biofilm colony can be cut in two equal halves. (C) The two equal biofilm halves are treated equally (control, PBS) or with either PBS or sterilizing agent and processed as described. Scale bar: 1 cm. Please click here to view a larger version of this figure.

3. Carefully lift each half of the biofilm colony from the agar plate with a small spatula and move it to a 1.5 ml microcentrifuge tube containing 500 µl of phosphate-buffered saline (PBS). If necessary, scrape the remaining cells from the plate and transfer them to the microcentrifuge tube as well.
   NOTE: The second half of the biofilm colony is treated differentially, depending on whether it is the control or to test the resistance to sterilizing agents.
4. For the control, incubate the second half of the biofilm colony in 500 µl PBS as in step 2.3. To assess resistance to sterilizing agents, transfer the second half of the biofilm colony to 500 µl 50% (v/v) ethanol.
   NOTE: Alternative sterilizing agents such as sodium hypochlorite can be used. For all sterilizing agents used, determine the active concentration and incubation time in a preliminary experiment.
5. Incubate the biofilm colonies for 10 min on the bench-top at room temperature.
6. Centrifuge the biofilm colonies for 5 min at 18,000 x g and carefully remove the supernatant with a pipette. Add 300 µl of PBS.
7. Sonicate the cells mildly (amplitude 10%, pulse 5 sec) with the microtip of a sonicator.
   NOTE: The sonication energy must be sufficient to separate biofilm aggregates. However, too harsh sonication may lyse the cells. Confirm in advance by light microscopy that the sonication energy used does not lyse the cells and that all aggregates are dissolved.
8. Add 700 µl of PBS to a final volume of 1 ml. Perform a serial dilution (to 10^-7) in PBS and spread 100 µl of 3 dilutions on a LB-1.5% agar plate using sterile glass beads.
   NOTE: The optimal dilutions to be plated should be determined in a preliminary experiment, as this depends on the amount of cells in the biofilm colony of interest and the survival rate of the cells in response to the sterilizing agent.
9. Incubate the plates overnight at 30 °C, count the CFU and determine CFU/ml. From the final CFU/ml of each the half biofilm colonies, calculate the percentage of survivors.
   NOTE: When performed and analyzed as described, the two halves of the control biofilm colony and the untreated versus ethanol-treated half of the untreated biofilm colony should yield differences below 10% in viable cell counts, verifying the symmetry or the resistance of the colony, respectively. Alternatively, the results can be represented in total CFU. The cell counts of the control and untreated biofilm colony should remain in the same order of magnitude. In contrast, the cell counts of the ethanol-treated half of a small molecule-treated biofilm colony are expected to drop by a minimum of two orders of magnitude to claim for an increased sensitivity to the sterilizing agent.

3. Biofilm Colony Sample Preparation for Scanning Electron Microscopy

1. Grow biofilm colonies as described in steps 1.1-1.7 and 1.12-1.14.
2. Prepare a fresh batch of 2% (v/v) glutaraldehyde, 3% (v/v) paraformaldehyde solution in 100 mM sodium cacodylate, 5 mM calcium chloride buffer, pH 7.3. Prepare 5 ml of fixative for every Petri dish of 8.5 cm diameter.
   CAUTION: Glutaraldehyde and paraformaldehyde are hazardous. Handle them with safety equipment inside a chemical hood. Discard the solutions and the contaminated materials to the hazardous waste.
3. Carefully add the fixative to the biofilm colonies, without dispensing directly on top of the biofilms.
   NOTE: Due the hydrophobic character of the biofilm colony, the colonies slowly detach from the agar and start to float.
4. Carefully seal the plates with a strip of Parafilm. Incubate on a rotary shaker for 2 hr at room temperature and subsequently transfer the plates to 4 °C for overnight.
5. The next day, carefully remove the liquid with a Pasteur glass pipette connected to a vacuum pump.
6. Carefully add 10 ml 100 mM sodium cacodylate, 5 mM calcium chloride buffer to wash the biofilm and incubate for 5 min. Gently remove the liquid with the Pasteur glass pipette from the corner of the plate to avoid damaging the biofilm and add fresh washing solution by gentle pipetting. Repeat this step once.
7. For the dehydration of the biofilm colonies, proceed with the following steps: 2x 5 min in ddH₂O; 2x 20 min in 30% ethanol; 2x 20 min in 50% ethanol; 2x 20 min in 70% ethanol; 2x 20 min in 96% ethanol; 2x 30 min in 100% ethanol.
   1. Add 15 ml of liquid for every Petri dish of 8.5 cm diameter in each step and remove the liquid carefully after each incubation.
8. Use one of two different methods for drying the samples from ethanol.
   1. For air-drying from ethanol:
      1. Cut a cellulose filter paper (diameter of 9 cm) in quarters. Briefly submerge one quarter in 100% ethanol, and then carefully transfer one floating biofilm colony onto it. Put the wet filter paper in a Petri dish lined with a filter paper. Cover the Petri dish and let the biofilm colonies dry overnight in a chemical hood.
   2. For critical point (CP)-drying using carbon dioxide (CO₂) as the transition fluid:
      1. Fill 75% of the critical point drying machine chamber with 100% ethanol. Transfer the samples into a holder, each sample into its own chamber. If necessary, cut the biofilm with scissors into smaller pieces. Leave the samples submerged in ethanol during all the handling. Then, transfer the holder into the chamber and close the chamber tightly.
      2. Cool the chamber to 7 °C and start stirring. Fill the chamber completely with liquid CO₂. During a 7 min incubation time, let the ethanol mix with the CO₂. Then, discharge 25% of the solution.
         NOTE: Do not empty the chamber below the level of the sample.
      3. Repeat step 3.8.2.2 four times.
      4. Repeat step 3.8.2.2 five times with an incubation time of 5 min only. Finally, the ethanol should be completely replaced by CO₂.
      5. During the last round, empty only 5% of the chamber. Turn off the stirring and the cooling. Start heating the chamber to 42 °C. At a temperature of 31.1 °C and a pressure of 73.9 bar, the liquid CO₂ reaches its critical point, the state where the gaseous phase has the same density as the liquid phase of the solvent²⁷. Once the temperature reaches 42 °C, incubate for 10 min. At 42 °C, the CO₂ in the chamber exists as supercritical gas.
         NOTE: Constantly check the pressure of the chamber. The pressure should not exceed 120 bar at 42 °C.
      6. Start to slowly release the gas with continuous heating. This keeps the samples in the CO₂-gas phase and prevents the deformation of the sample morphology through the liquid surface tension. Set the flowmeter to 5 L/hr by fine tuning the metering valve controlling the flowmeter. Wait until the all the pressure in the chamber is released. Now open the chamber and remove the samples carefully from the holder.
9. Coat an electron microscopy stub with carbon tape. With the help of tweezers, carefully transfer the biofilm colonies onto the stub. Connect each colony to the stub by adding a thin bridge from carbon tape, which is crucial for charge elimination under the electron beam. At this stage, handle the biofilm colonies with care as they are very fragile. Store the samples in a desiccator for at least 24 hr or until examination.
10. The day of the examination with the scanning electron microscope, sputter-coat the biofilm colonies for 2 min in a 60° angle in a gold-palladium sputter-coater. Repeat this step twice and rotate the samples by 120° in between. At the end, sputter-coat the samples once for 3 min from the top. The 20 nm thin layer of gold-palladium improves the conductivity and enhances the contrast of the sample for the imaging in the SEM.
11. Store the samples in a desiccator to avoid the rehydration of the sample²⁸ until imaging with a scanning electron microscope^29,30.