In order to allow measurement of cell wall deformation using surface-enhanced fluorescence, two green fluorescent bacterial strains were selected. S. mutans UA159^GFP (Deng et al., 2007) and S. aureus ATCC 12600^GFP (Li et al., 2011) were grown on Todd Hewitt Broth (THB) (Oxoid, Basingstoke, United Kingdom) and Tryptone Soya Broth (TSB) (Oxoid, Basingstoke, United Kingdom) agar plates, respectively. THB agar plates were supplemented with 10 μg/ml erythromycin (Sigma-Aldrich, St Louis, MO, United States) and TSB agar plates were supplemented with 10 μg/ml tetracycline (Sigma-Aldrich, St. Louis, MO, United States). One colony of S. mutans was inoculated in THB supplemented with 10 μg/ml erythromycin and similarly for S. aureus in TSB supplemented with 10 μg/ml tetracycline. S. mutans UA159^GFP was grown at 37°C with 5% CO₂ and S. aureus was grown under aerobic conditions. After 24 h, these precultures were inoculated in 200 ml (1:20) THB for S. mutans and TSB for S. aureus without antibiotics and cultured for 16 h at 37°C, yielding early-stationary phase bacteria.

Bacterial cultures were harvested by centrifugation (5 min, 5,000 g, 10°C) and washed twice with a phosphate buffer (2 mM potassium phosphate, 50 mM KCl, and 1 mM CaCl₂, pH 7.0). After washing, bacteria were resuspended in phosphate buffer with THB or TSB (30:1) to maintain metabolic activity. The bacterial suspension was sonicated (3 × 10 s, 30 W) in an ice-water bath (Vibra Cell Model 375, Sonics and Materials Incorporation, Danbury, CT, United States) and the bacteria were enumerated using a Bürker-Türk counting chamber and suspensions (3 × 10⁸/ml) were diluted in phosphate buffer supplemented with THB or TSB (30:1) to a bacterial concentration, appropriate for the different experiments. All the experiments were done in triplicate with different bacterial cultures.

A commercial chlorhexidine mouthrinse (Curasept ADS 212, Curasept S.p.A., Saronno, Italy) containing 0.12% (1,200 μg/ml) chlorhexidine digluconate was used in this study. Bacteria were exposed to 2-fold serial dilutions (i.e., 9.6, 4.8, 2.4, 1.2, 0.6, and 0.3 μg/ml) of the mouthrinse in sterile water. 100 μl of different dilutions of the mouthrinse were added to 100 μl of a bacterial suspension (2 × 10⁵/ml) in the appropriate growth medium in 96-well plates and incubated at 37°C for 24 h. After incubation, the minimal inhibitory concentration (MIC) was taken as the lowest chlorhexidine concentration at which no visible growth was observed.

For the development of chlorhexidine-resistant bacterial strains, precultures of S. mutans UA159^GFP and S. aureus ATCC 12600^GFP were diluted 1:100 in appropriate growth media and grown with chlorhexidine added at its MIC. Bacteria were subcultured daily at the same chlorhexidine concentration over 4 consecutive days. Culturable bacteria that remained present after 4 days were subsequently grown over another sequence of 4 consecutive days at a 0.3 μg/ml higher chlorhexidine concentration than in the previous sequence. MIC was measured daily during each sequence of 4 consecutive days. Subculturing was discontinued when a culture did not demonstrate any bacterial growth by visual inspection of the culture turbidity. In order to check whether prolonged culturing under chlorhexidine exposure affected bacterial fluorescence, fluorescence was monitored regularly on agar plates using an In Vivo Imaging System (IVIS, Lumina II, Caliper LifeScience, Hopkinton, MA, United States), with an excitation wavelength of 465 nm and emission in a range from 515 to 575 nm. Cultures after repeated growth in presence of chlorhexidine were stored in a −80°C freezer, with 7% dimethyl sulfoxide added.

Molecular fluorescence is enhanced when fluorescent molecules are in the close vicinity (<30 nm) of a metal surface (Lee et al., 2011). This phenomenon, called surface-enhanced fluorescence, can be applied to measure the small deformation that adhering fluorescent bacteria experience in response to the adhesion forces arising from a substratum surface (see Figure 1 for schematics). As a major advantage over atomic force microscopy, surface-enhanced fluorescence is more sensitive and measures over a larger population of bacteria, depending on the number of bacteria adhering to the metal surface.

For surface-enhanced fluorescence, bacteria suspended in phosphate buffer supplemented with growth medium (30:1) were injected in a parallel plate flow chamber, with a glass top-plate and a polished, stainless steel bottom-plate. Fluorescence was measured using a bio-optical imaging system (see above). Images had a field of view of 7.5 cm × 7.5 cm, while exposure time was set at 10 s, employing a focal ratio of 1. Temperature throughout an experiment was maintained at 20°C. With the Living Image software package 3.1 (Caliper LifeScience), a user-defined region of interest was constructed in each image of 4.0 cm × 1.0 cm to calculate the average fluorescence radiance (photons s^–1 cm^–2 steradian^–1).

Background fluorescence in the region of interest was measured in a flow chamber filled with phosphate buffer, supplemented with growth medium (30:1) in the absence of suspended bacteria. This background fluorescence radiance was subtracted from all the fluorescent radiances measured. Next, bacteria were injected into the flow chamber. Sedimentation of suspended bacteria and their adhesion was allowed in the absence of flow under the influence of gravity, while acquiring images every 15 min for 3 h, previously found sufficient for complete sedimentation of all the suspended bacteria (Li et al., 2014). Subsequently, the flow chamber was filled with 5 ml of the commercial chlorhexidine mouthrinse at different dilutions. Fluorescence image acquisition continued every 15 min for an additional 2 h. Phosphate buffer supplemented with growth medium (30:1) was used as a control.

Assuming even intracellular distribution of green fluorescent proteins, the increase in fluorescence radiance due to adhering bacteria relative to planktonic bacteria was expressed as total fluorescence enhancement (Li et al., 2014), according to:

Where, R(t) is the fluorescence radiance at time t, R(0) is the fluorescence radiance measured for a planktonic suspension, and R₀ is the fluorescence radiance of the background.

To determine the percentage of damage upon chlorhexidine exposure to the cell membrane, as an integral part of the cell wall, bacteria were stained with red fluorescent propidium iodide (LIVE/DEAD BacLight Bacterial Viability, Thermo Fisher Scientific, Waltham, MA, United States). Propidium iodide is only able to enter bacteria with a damaged membrane (Lehtinen et al., 2004). Membrane damage was inflicted according to a similar protocol as in surface-enhanced fluorescence. After incubation with chlorhexidine, 15 μl of propidium iodide (20 mM) was added to each well. Fluorescence was imaged using a Leica DM4000B fluorescence microscope with a 40× water objective by taking three images of each well. While closing the red channel, green fluorescent bacteria were enumerated as a measure of the total number of bacteria in an image, while those bacteria also displaying red fluorescence were taken as membrane damaged (Lehtinen et al., 2004).

The influence of the different concentrations of chlorhexidine on bacterial metabolic activity was determined with 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT) reduction assay (Krom et al., 2007). After exposure to chlorhexidine according to a similar protocol as in surface-enhanced fluorescence, MTT solution (0.5 mg/ml thiazolyl blue tetrazolium bromide in phosphate buffer, 10 mg/ml glucose, and 0.1 mM menadione) was added to each well and left for 30 min at 37°C in the dark. After 30 min, wells were washed once with water and mixed with acid isopropanol (5% 1 M HCl in isopropanol) for 15 min. After 15 min, 100 μl of the suspension was removed and added to a new 96-wells plate and absorptions were measured at 560 nm with the FLUOstar Optima Microplate Reader (BMG Labtech, Offenburg, Germany). All the experiments were performed in triplicate with different bacterial cultures. The data were normalized with respect to the metabolic activity of the bacterial strains after 3 h adhesion, i.e., before chlorhexidine exposure.

To determine whether the physicochemical bacterial cell surface properties were affected by chlorhexidine exposure, microbial adhesion to hydrocarbons (kinetic MATH assay) and zeta potentials were measured as a function of exposure time of the bacteria to chlorhexidine dissolved in phosphate buffer. MATH was carried out as previously described (Lichtenberg et al., 1985). Briefly, bacteria were resuspended in 3 ml phosphate buffer (pH 7.0) supplemented with chlorhexidine and containing 20:1 hexadecane to an initial absorbance at 600 nm between 0.4 and 0.6. Initial absorbance at time zero ([A0]) was measured photospectrometrically (Spectronic 20 Genesys, Spectronic Instruments, Rochester, NY, United States). After vortexing, the suspension for 10 s, and settling of the bacteria for 10 min, absorbance was measured again at time t ([At]). This procedure was repeated five more times, to allow calculation of the initial rate of bacterial removal from the aqueous phase according to:

where, t is vortexing time.

The charge properties of the bacterial surfaces were determined by measuring the electrophoretic mobility using a bacterial suspension (3 × 10⁸ bacteria/ml) in phosphate buffer (pH 7.0) supplemented with chlorhexidine. Particulate microelectrophoresis was carried out on a Zetasizer Nano ZS (Malvern Instruments, Worcestershire, United Kingdom). Electrophoretic mobilities measured were converted into zeta potentials, employing the Helmholtz–Smoluchowski equation (Van der Wal et al., 1997).

Data were statistically analyzed using the paired and two-tailed Student’s t-tests with Microsoft Excel 2010. Significance was established at p < 0.05.

Prior to chlorhexidine exposure, the MIC of both the S. mutans UA159^GFP and S. aureus ATCC 12600^GFP was 1.2 μg/ml. In order to induce chlorhexidine resistance, planktonic cultures of S. mutans UA159^GFP and S. aureus ATCC 12600^GFP were grown in medium, supplemented with chlorhexidine. After daily subculturing at increasing chlorhexidine concentrations starting at sub-MIC over 4 consecutive days, no culturable S. mutans was observed when the chlorhexidine concentration exceeded MIC (Supplementary Figure 1). For S. aureus, culturable staphylococci remained present after 13 repetitions of 4 day sequences of growth at stepwise increasing chlorhexidine concentrations, acquiring a MIC of 4.8 μg/ml, i.e., four times the starting MIC of S. aureus ATCC 12600^GFP (Supplementary Figure 1). This resistant variant of S. aureus ATCC 12600^GFP was subsequently used in further experiments.

Both the chlorhexidine-susceptible S. aureus strain (Figure 2A) and its chlorhexidine-resistant variant (Figure 2B) demonstrated a gradual increase in total fluorescence enhancement during the first 3 h of sedimentation and adhesion from phosphate buffer to a stainless steel surface, indicative of cell wall deformation. After 3 h, total fluorescence enhancement remained constant for at least 6 h upon continued exposure to buffer. However, the resistant variant (Figure 2B) deformed faster and more extensively than the chlorhexidine-susceptible strain (Figure 2A). Total fluorescence enhancement of S. mutans during sedimentation and adhesion followed by continued exposure to phosphate buffer hovered around 1 for at least 6 h, indicating absence of cell wall deformation upon adhesion (Figure 2C). Continued exposure to different concentrations of chlorhexidine after sedimentation and adhesion in phosphate buffer led to a different response in staphylococci than in streptococci. Both the chlorhexidine-susceptible (Figure 2A) and chlorhexidine-resistant S. aureus (Figure 2B) demonstrated gradual increase in total fluorescence enhancement that was independent of chlorhexidine concentration. These increases in total fluorescence enhancement remained to increase for at least 6 h, indicative of permanent cell wall deformation. Cell wall deformation was significantly (p < 0.05, Student’s t-test) larger for the already weakened cell wall of the resistant variant (Figure 2B) as compared with its chlorhexidine-susceptible parent strain (Figure 2A). S. mutans, on the other hand, demonstrated a fast increase in total fluorescence enhancement upon chlorhexidine exposure (Figure 2C). The increase in total fluorescence enhancement was dependent on the chlorhexidine concentration during exposure and decreased gradually over time to the level observed in phosphate buffer without chlorhexidine added.

The physicochemical surface properties, bacterial zeta potentials (Figure 3A), and hydrophobicities (Figure 3B) relevant for adhesion did not change upon chlorhexidine exposure for all the three strains.

Cell wall weakening was further studied by assessing cell wall damage based on cell membrane permeability to red fluorescent propidium iodide. Damage to the membrane, as an integral part of the cell wall, can be demonstrated at microscopic level and over a large population of bacteria by staining with red fluorescent propidium iodide, entering only damaged cell walls with increased membrane permeability.

Exposure of adhering bacteria to different concentrations of chlorhexidine caused nearly 50% of S. aureus to become membrane damaged, irrespective of concentration (Figure 4A). In the chlorhexidine-resistant S. aureus variant, exposure to chlorhexidine led to significantly fewer bacteria with membrane damage (<25%) (Figure 4B). Very few S. mutans became membrane damaged upon exposure to chlorhexidine (Figure 4C). Collectively, these data suggest that S. mutans suffers considerably less cell wall damage upon exposure to chlorhexidine than the susceptible S. aureus strain (p < 0.05) and even less than a chlorhexidine-resistant S. aureus variant although initially upon exposure to chlorhexidine it deforms equally as staphylococci do.

In order to verify the suggestion of self-repair, cell wall damage was inflicted upon both the strains by a short-term, 15 min exposure to 1.2 μg/ml chlorhexidine (MIC) and compared with the cell wall damages inflicted by longer-term, 2 h exposure. Short-term exposure to chlorhexidine yielded only 10% of the S. aureus bacteria to become cell wall damaged, while 99% of S. mutans was already cell wall damaged, allowing penetration of red fluorescent propidium iodide (Figure 5). Upon continued exposure to chlorhexidine up to 2 h, however, 38% of the S. aureus appeared cell wall damaged than upon short-term exposure, but almost the entire population (96%) of S. mutans within a microscopic field of view had fully self-repaired the damage over time. A sub-MIC concentration of chlorhexidine showed similar self-repair as observed at MIC (Supplementary Figure 2). This confirms the presence of a self-repair mechanism against cell wall damage inflicted by chlorhexidine in S. mutans that is absent in S. aureus. However, at chlorhexidine concentration far above MIC (600 μg/ml), self-repair in S. mutans was no longer observed (see also Supplementary Figure 2).

Initially, short-term (15 min) exposure to 1.2 μg/ml chlorhexidine did not show an increase in metabolic activity (Figure 6) in the resistant S. aureus and in S. mutans, but a small increase in the susceptible S. aureus. Exposure to the lower chlorhexidine concentrations shows a small, but not significant increase in metabolic activity for the S. aureus strains as compared to buffer exposure. After 2 h exposure to chlorhexidine, the metabolic activity of both the S. aureus strains at 0.6 and 1.2 μg/ml was significantly (p < 0.05, Student’s t-test) higher than of the S. mutans strain. S. mutans exhibited no significant increased metabolic activity upon self-repair of chlorhexidine-inflicted cell wall damage (Figure 6).