Monitoring Extracellular pH in Cross-Kingdom Biofilms using Confocal Microscopy

The protocol presents an easy-to-perform method for the cultivation of cross-kingdom biofilms consisting of StreptococcuS. mutans and Candida albicans. Subsequent confocal microscopy-based pH ratiometry allows for accurate monitoring of pH developments inside the extracellular matrix of those biofilms.

The careful choice of image acquisition parameters is of utmost importance to obtain image data with sufficient contrast for subsequent analysis. pH ratiometry is a rapid, precise, and inexpensive method to study both horizontal and vertical pH gradients in cross-kingdom biofilms in real time. pH ratiometry can also be performed in purely fungal and bacterial biofilms.

The method contributes to increase our understanding of microbial metabolism in biofilms. Demonstrating the procedure for biofilm growth will be Anette Aakjaer Thomsen, a technician from my laboratory. Grow S.mutans and C.albicans on blood agar plates at 37 degrees Celsius under aerobic conditions.

Then transfer single colonies of each organism to test tubes filled with five milliliters of brain heart infusion and grow them for an additional 18 hours. On the next day, centrifuge the cultures at 1, 200 times g for five minutes and discard the supernatant. Resuspend the cells in physiological saline and adjust the OD 550 to 0.5 for both C.albicans and S.mutans.

Then dilute the S.mutans suspension one to 10 to achieve equivalent concentrations. Pipette 50 microliters of sterile salivary solution into the wells of an optical bottom 96-well plate for microscopy. Incubate the plate for 30 minutes at 37 degrees Celsius, then wash the plate three times with 100 microliters of sterile physiological saline and empty the wells.

Add 100 microliters of C.albicans suspension to each well, incubate the plate at 37 degrees Celsius for 90 minutes, then wash the well three times with saline. Next, add 100 microliters of heat-inactivated fetal bovine serum to each well. Incubate the plate for two hours, then wash it three times with saline.

Empty the wells but leave a 20 microliter reservoir to avoid excessive shear forces. Add 100 microliters of S.mutans suspension and 150 microliters of BHI with 5%sucrose to each well. Then incubate the plate at 37 degrees Celsius for 24 hours or longer.

When cultivating older biofilms, replace the medium daily. At the end of the cross-kingdom growth phase, wash the plate five times with sterile physiological saline. For ratiometric pH imaging, use an inverted confocal laser scanning microscope with a 63X oil or water immersion lens, a 543 nanometer laser line, and a spectral imaging system.

Use an incubator to warm the microscope stage to 35 degrees Celsius. Set the detector for simultaneous detection of green fluorescence from 576 to 608 nanometers and red fluorescence from 629 to 661 nanometers. Then choose an appropriate laser power and gain to avoid over and underexposure.

Set the pinhole size to one air unit or an optical slice of about 0.8 micrometers. Then set the image size to 512 by 512 pixels and the scan speed to two. Choose a line average of two using the mean option.

Prepare 100 microliters of sterile physiological saline with 0.4%glucose titrated to pH seven. Then make a stock solution of C-SNARF-4 and add the dye to a final concentration of 30 micromolar. Empty one of the wells with the cross-kingdom biofilm leaving a 20 microliter reservoir and add the saline with glucose and ratiometric dye.

Place the plate on the microscope stage and start imaging. Prepare a series of 50 millimolar MES buffer titrated to pH four to 7.8 in the increments of 0.2 pH units at 35 degrees Celsius. Pipette 150 microliters of each buffer solution into the wells of an optical bottom 96-well plate.

Add the dye to the buffer-filled wells at a concentration of 30 micromolar and let it equilibrate for five minutes. Warm the microscope stage to 35 degrees Celsius and choose the same settings as for ratiometric pH imaging previously described. Place the 96-well plate on the microscope stage and focus on the bottom of the wells.

Acquire green and red channel images for all buffer solutions. At regular intervals, take images with the laser turned off to correct for detector offset. Perform the calibration experiment in triplicate and export all images as TIFF files.

Store the green and red images in separate folders and rename both series of files with sequential numbers. Import the images into dedicated image analysis software such as ImageJ. In the images taken with the laser off, click analyze and histogram to determine the average fluorescence intensity.

Subtract this value from the biofilm images by clicking process, math, and subtract. Then import the two image series into daime and perform a threshold-based segmentation of the red channel images by clicking segment, automatic segmentation, and custom threshold. Set the low threshold above the fluorescence intensity of the fungal cytoplasm and the high threshold below the intensity of the fungal cell walls and the bacteria.

Then transfer the object layer of the segmented image series to the green channel image series by clicking segment and transfer object layer. Delete non-object pixels in the red and green channel series. Now the biofilm images are cleared from bacterial and fungal cells and can be exported as TIFF files.

Import the image series back into ImageJ and divide the red image series by itself. Then multiply the result by the original image series. The resulting image series is identical to the original except all zero intensity pixels are removed.

Repeat the process with the green image series. Next, use the mean filter on both image series to compensate for detector noise. Divide the green by the red image series which then yields the green-to-red ratio for all object pixels.

Robust cross-kingdom biofilms developed in the well plates after 24 and 48 hours. Single cells and chains of S.mutans grouped around fungal hyphae and large intracellular spaces indicated the presence of a voluminous matrix. The pH in the extracellular space was visualized using a lookup table.

The extracellular pH in the biofilms dropped quickly in the first five minutes after exposure to glucose. Thereafter, acidification slowed down typically reaching values of 5.5 to 5.8 after 15 minutes. Due to the local pH changes, fluorescence intensity in the biofilms changed over time.

During image segmentation, high and low thresholds were chosen to adequately eliminate all areas covered by bacterial and fungal cells. The blue and green areas were eliminated by the low and high thresholds respectively. The careful choice of image acquisition parameters is crucial to obtain a good contrast between bacterial cells, fungal cells, and the biofilm matrix.

If cross-kingdom biofilms are grown in flow cells, pH developments can be monitored under flow condition mimicking those in the oral cavity. pH ratiometry has been employed to identify local areas in dental biofilms with particularly low pH, so called acidogenic hotspots.