Research Article
22 July 2016

New Insight into Daptomycin Bioavailability and Localization in Staphylococcus aureus Biofilms by Dynamic Fluorescence Imaging

ABSTRACT

Staphylococcus aureus is one of the most frequent pathogens responsible for biofilm-associated infections (BAI), and the choice of antibiotics to treat these infections remains a challenge for the medical community. In particular, daptomycin has been reported to fail against implant-associated S. aureus infections in clinical practice, while its association with rifampin remains a good candidate for BAI treatment. To improve our understanding of such resistance/tolerance toward daptomycin, we took advantage of the dynamic fluorescence imaging tools (time-lapse imaging and fluorescence recovery after photobleaching [FRAP]) to locally and accurately assess the antibiotic diffusion reaction in methicillin-susceptible and methicillin-resistant S. aureus biofilms. To provide a realistic representation of daptomycin action, we optimized an in vitro model built on the basis of our recently published in vivo mouse model of prosthetic vascular graft infections. We demonstrated that at therapeutic concentrations, daptomycin was inefficient in eradicating biofilms, while the matrix was not a shield to antibiotic diffusion and to its interaction with its bacterial target. In the presence of rifampin, daptomycin was still present in the vicinity of the bacterial cells, allowing prevention of the emergence of rifampin-resistant mutants. Conclusions derived from this study strongly suggest that S. aureus biofilm resistance/tolerance toward daptomycin may be more likely to be related to a physiological change involving structural modifications of the membrane, which is a strain-dependent process.

INTRODUCTION

Staphylococcus aureus is a Gram-positive bacterial species shown to be the most frequent cause of biofilm-associated infections (BAI) (1) and one of the major causes of morbidity and mortality in hospitals and communities (2). Unlike planktonic cells, biofilms exhibit specific phenotypic traits allowing them to resist host defenses and antibiotic treatments (3), which frequently leads to chronic infections such as endocarditis, sinusitis, and osteomyelitis and also to implant-associated infections (4).
Among the most recent clinically used antibiotics, daptomycin is a cyclic lipopeptide approved for the treatment of serious staphylococcal infections such as bacteremia and implant-related infections (5). Daptomycin is a calcium-dependent antibiotic that acts by insertion into the Gram-positive cytoplasmic membranes where it forms oligomeric pores, causing potassium ion leakage and subsequent membrane depolarization, leading ultimately to cell death (6). As is the case for many antibiotics, daptomycin has been shown to exhibit a significant bactericidal activity against planktonic cells (79). However, the eradication of adherent bacteria is rarely achieved despite the large number of in vitro and animal studies in which daptomycin activity was evaluated (1016). Besides the results of the literature that appear controversial (17), direct comparison between studies is not directly possible, since the biofilm growth and treatment protocols used differ greatly.
To obtain a realistic representation of daptomycin action against S. aureus biofilms, we developed an in vitro model built on the basis of our recently published in vivo study on S. aureus prosthetic vascular graft infections using the same strains (18). The interest of this approach was the possibility to use fluorescence imaging techniques that cannot be employed in vivo (confocal laser scanning microscopy [CLSM], time-lapse microscopy, and fluorescence recovery after photobleaching [FRAP]) to examine the penetration, diffusion, bioavailability, and localization of the fluorescently labeled antibiotic inside the biofilms. To validate this approach, we monitored for 72 h the activity of daptomycin against biofilms formed by methicillin-susceptible and methicillin-resistant clinical and collection strains. In addition, we enriched the culture medium with proteins and calcium to mimic the in vivo physiological conditions of our mouse model (18). The same experiments were performed in the presence of rifampin, an antibiotic that can be combined with daptomycin for recalcitrant S. aureus BAI.

MATERIALS AND METHODS

Bacterial strains.

Four S. aureus strains were tested in the present study: two were collection strains (methicillin-susceptible S. aureus [MSSA] ATCC 27217 and methicillin-resistant S. aureus [MRSA] ATCC 33591), and two others were isolated from patients with S. aureus bloodstream infections (MSSA 176 and MRSA BCB8). All strains were kept at −80°C in tryptic soy broth (TSB) (bioMérieux, France) containing 20% (vol/vol) glycerol. The frozen cells were subcultured twice in TSB (one 8-h culture, followed by an overnight culture) to constitute the stock cultures from which aliquots were kept at −20°C. Bacterial growth and experiments were both conducted at 37°C.

Antimicrobial agents and medium.

Daptomycin and rifampin were both purchased from Sigma (France). The fluorescently labeled antibiotic BODIPY-FL-labeled daptomycin (BODIPY-FL-daptomycin) (BODIPY-FL is fluorescently labeled boron-dipyrromethene) was a kind gift from Cubist Pharmaceuticals (MA, USA), and BODIPY-FL was purchased from Invitrogen (France). According to the manufacturer's instructions, the stock solutions were prepared by diluting daptomycin and BODIPY-FL-daptomycin in dimethyl sulfoxide (1 mg/ml) and by diluting rifampin in sterile water (2 mg/ml), which were then kept at −20°C. Before the solutions were used, they were diluted in TSB enriched with proteins (bovine serum albumin[BSA], 36 g/liter; Sigma, France) and calcium (CaCl2·2H2O, 50 mg/liter; Sigma, France) to mimic in vivo physiological levels. It has been determined that, under these conditions, the final concentration of dimethyl sulfoxide was noncytotoxic for the bacteria. Clinically meaningful concentrations were used in this study: 20 μg/ml for both daptomycin and rifampin. When combined, the antibiotics were mixed together before application to the biofilm surface.

Susceptibility testing.

The MICs of daptomycin and rifampin were determined by the broth microdilution method in cation-adjusted Mueller-Hinton broth (CAMHB), according to the European Committee on Antimicrobial Susceptibility Testing (EUCAST). Media were supplemented with 50 mg/liter Ca2+ for daptomycin.

Characterization of molecular interactions between daptomycin and rifampin by absorption and fluorescence spectroscopies.

Absorption spectra of antibiotics alone and in combination were measured with a Varian Cary 300 spectrophotometer (Agilent Technologies, France). The corresponding fluorescence emission spectra were recorded using a Fluorolog-3 (Jobin Yvon, Inc., France) fluorescence spectrophotometer mounted with front-face detection geometry by exciting daptomycin at 360 nm. The measurements were made five times on each sample.

In vitro biofilm preparation and antibiotic activities.

Biofilms were studied in a polystyrene microtiter plate-based assay since it has been shown that this material has physicochemical properties close to that of biomaterial surfaces such as polyethylene terephthalate, which is used in vascular grafts (19, 20). For the preparation of S. aureus biofilms, 250-μl portions of an overnight subculture adjusted to an optical density at 600 nm of 0.02 (corresponding to ∼108 CFU/ml) were added to 96-well microplates (μClear; Greiner Bio-One, France). After a 1.5-h adhesion period at 37°C, the wells were rinsed with sterile physiological water (150 mM NaCl) in order to eliminate nonadherent cells, refilled with sterile TSB enriched with proteins and calcium (TSBpc) and then incubated for 24 h at 37°C to allow biofilm growth.
To assess antibiotic activities, the 24-h biofilms were rinsed with a 150 mM NaCl aqueous solution before adding the antibiotic solutions diluted in TSBpc as described previously. Viable culturable bacteria were then counted at time points at regular intervals: 0 h (when antibiotics are added), 24, 48, and 72 h after antibiotic injection. For each time point, bacterial cultures were centrifuged 10 min at 7,000 × g in order to eliminate excess antibiotic. The bacterial pellet was suspended in a 150 mM NaCl sterile saline solution, centrifuged again, and suspended in the same conditions. Successive decimal dilutions were then performed. For each dilution, 6 drops (10 μl) were deposited on tryptic soy agar (TSA) plates (bioMérieux, France) and incubated at 37°C during 24 h. CFU were counted and averaged for each dilution at each time point. The detection limit of viable culturable cells was 100 CFU/ml.

Percentages of rifampin-resistant mutants in biofilms.

To determine the percentages of rifampin-resistant mutants, the antibiotic solutions (rifampin alone or combined with daptomycin) were added to 24-h biofilms. CFU were counted on TSA plates or TSA plates containing rifampin (20 μg/ml) at 24 and 48 h after antibiotic injections. The percentages were obtained by calculating the ratio between the number of CFU grown on rifampin-containing TSA plates and the number of CFU grown on rifampin-free TSA plates.

Statistical analysis.

The mean log10 CFU/milliliter and biovolume for each therapy were compared with each other by analysis of variance (ANOVA). They were performed using the Statgraphics software (Manugistics, Rockville, MD, USA). Statistical significance was defined as a P value of less than 0.05 by a Fisher test.

Confocal laser scanning microscopy. (i) Visualization of antibiotic activities against biofilms using LIVE/DEAD staining.

Biofilms grown for 24 h prepared as described previously were observed 24, 48, and 72 h after antibiotic addition using a Leica TCS SP5 confocal laser scanning microscope (Leica Microsystems, France) at the Centre de Photonique Biomédicale (CPBM) (Orsay, France). Prior to each observation, bacteria were stained with 2.5 μM Syto9 (Life Technologies, France), a green cell-permeant nucleic acid dye, and 30 μM propidium iodide (PI) (Life Technologies, France), a red nucleic acid dye that can penetrate cells with compromised membranes (dead cells) only. Syto9 and PI were sequentially excited at 488 nm and 543 nm, respectively, and their fluorescence emissions were collected between 500 and 600 nm for Syto9 and between 640 and 750 nm for PI. Images were acquired using a 63× oil immersion objective with a 1.4 numerical aperture. The size of the confocal images was 512 by 512 pixels (82 by 82 μm2), recorded with a z-step of 1 μm. For each biofilm, four typical regions were analyzed. Images were reconstructed in three dimensions (3D) using ICY software.

(ii) Quantification of biovolumes and maximum biofilm thickness.

Maximum biofilm thickness (in micrometers) was measured directly from xyz stacks. Biovolumes (in cubic micrometers) were calculated by binarizing images with a java script executed by ICY software as described previously (21). The biovolume was then defined as the overall volume of cells in the observation field. The percentage of dead cells corresponds to the ratio between biovolumes of PI-stained bacteria and Syto9-stained bacteria.

(iii) Antibiotic penetration and localization inside biofilms.

To study the penetration of BODIPY-FL-daptomycin alone and combined with rifampin within 24-h biofilms, we employed time-lapse microscopy, as described before (22), using the same Leica TCS SP5 confocal microscope. Briefly, the fluorescence intensity evolution over time was observed in a defined focal plane (5 μm above the substratum surface). As soon as the TSBpc-diluted solutions of BODIPY-FL-daptomycin alone or combined with rifampin were added to the biofilm, fluorescence intensity images were acquired every second for 15 min. Simultaneously, transmission images were acquired to ensure that no structural alteration of the biofilm occurred during this process. The labeled antibiotic was excited with a continuous argon laser line at 488 nm through a 63× oil immersion objective, and the emitted fluorescence was recorded within 500 and 600 nm.
The corresponding diffusive penetration coefficients (Dp) through the biofilms were determined according to the relationship previously described by Stewart (23):
Dp=1.03×L2/t90
(1)
where L is the biofilm thickness and t90 is the time required to attain 90% of the equilibrium staining intensity at the deeper layers of the biofilm.
To observe the localization of the fluorescently labeled daptomycin within biofilms, bacteria were counterstained with the FM4-64 dye (Life Technologies, France): this dye was also excited at 488 nm, but its fluorescence emission was collected within the range 640 to 750 nm. The images (512 by 512 pixels) of both fluorophores were simultaneously recorded with a z-step of 1 μm.

(iv) Antibiotic diffusion and bioavailability inside biofilms using FRAP experiments.

Image-based fluorescence recovery after photobleaching (FRAP) measurements was used to assess local diffusion and bioavailability of the fluorescently labeled daptomycin. Briefly, FRAP is based on a brief excitation of fluorescent molecules by a very intense light source in a user-defined region to irreversibly photobleach their fluorescence. Fluorescence recovery is then probed over time at a low light power in the same photobleached region (22, 24). All time-resolved measurements were obtained using the same confocal microscope. The time course of fluorescence intensity recovery was analyzed with mathematical models, giving us the quantitative mobility of the fluorescent molecules and allowing us to determine the diffusion coefficients. For all FRAP experiments, the fluorescence intensity image size was fixed to 512 by 128 pixels with an 80-nm pixel size and recorded using a 12-bit resolution. The line scan rate was fixed at 1,400 Hz, corresponding to a total time between frames of ∼265 ms. As determined previously, the full widths at half maximum in xy and z (along the optical axis) of the bleached profile were 0.8 μm and 14 μm, respectively, allowing us to neglect diffusion along the axial/vertical axis and thus to consider only two-dimensional diffusion. Each FRAP experiment started with the acquisition of 50 images at 7% of laser maximum intensity (7 μW) followed by a 200-ms single bleached spot at 100% laser intensity. A series of 450 single-section images was then collected with the laser power attenuated to its initial value (7% of the bleach intensity). The first image was recorded 365 ms after the beginning of bleaching.

RESULTS

Susceptibility testing.

The MICs obtained for daptomycin and rifampin against the four S. aureus strains are presented in Table 1. All isolates in planktonic conditions were susceptible to daptomycin and rifampin. The breakpoint values of drugs according to the European Committee on Antimicrobial Susceptibility Testing (EUCAST) in 2016 are as follows: 2 mg/liter for vancomycin; 1 mg/liter for daptomycin; for rifampin, sensitivity of ≤0.06 mg/liter and resistance of >0.5 mg/liter.
TABLE 1
TABLE 1 MICs of daptomycin and rifampin against the four S. aureus strainsa
S. aureus strainMIC (mg/líter) of:
DaptomycinRifampin
MSSA ATCC 272170.25<0.06
MSSA 1760.50.015
MRSA ATCC 335910.250.0075
MRSA BCB80.125<0.06
a
The standard deviations of the values are ±5%.

Spectroscopic characterization of the interaction between rifampin and daptomycin.

Neither the photonic absorption properties nor the fluorescence emission spectrum of daptomycin were significantly influenced by the addition of rifampin (Fig. 1), revealing the absence of cross-reaction between the two antibiotics.
FIG 1
FIG 1 Absorption and fluorescence spectra of daptomycin (20 μg/ml) alone and combined with rifampin (20 μg/ml). The excitation wavelength was 360 nm.

BODIPY-FL-daptomycin penetration, diffusion, bioavailability, and localization inside S. aureus biofilms. (i) Time-lapse imaging.

Time-lapse experiments were performed to visualize in situ penetration of fluorescently labeled daptomycin alone and combined with rifampin throughout the deepest layers of S. aureus biofilms. By setting the focal plane 5 μm above the substratum surface, we demonstrated that BODIPY-FL-daptomycin penetrated the biofilms (∼30-μm thickness) within a few minutes: fluorescence intensity was measured a few seconds after antibiotic addition and increased rapidly to reach 90% of the maximal intensity in 9 min (see Fig. S1 and Movie S4 in the supplemental material). The penetration coefficients obtained from equation 1 ranged from 2.5 μm2/s ± 0.7 μm2/s for S. aureus ATCC 27217, 176, and ATCC 33591 to 4.9 μm2/s ± 0.7 μm2/s for S. aureus BCB8. These values are both of the same order by comparison with BODIPY-FL alone for which the penetration coefficient was higher at 140 μm2/s (22). The coefficients were not statistically different in comparison with those of daptomycin in the presence of rifampin.

(ii) FRAP imaging.

FRAP was used to measure the local diffusion of BODIPY-FL-daptomycin and its interaction with bacteria within biofilms. According to the FRAP principle (22, 24, 25), if the fluorescently labeled daptomycin molecules are allowed to move freely in the sample, total fluorescence recovery is observed, meaning that the fluorescence is redistributed in the defined region. Conversely, if the fluorescence recovery is not total after the photobleaching, it means that a fraction of molecules is not diffusing freely and thus interacts with its local environment. The other fraction diffuses and is thus bioavailable.
First, we checked that no bacterial movement occurred during image acquisition by representing kymograms, two-dimensional graphs of fluorescence intensity measured along a line (here a straight line drawn on the full width of the images) for each image of the FRAP series (Fig. 2a). A typical FRAP curve of BODIPY-FL-daptomycin in S. aureus biofilms is presented in Fig. 2b. Whatever the bacterial strains or the treatments used (daptomycin alone or in combination with rifampin), we demonstrated here that the fluorescence recovery was not total after photobleaching: only 20% of BODIPY-FL-daptomycin molecules interacted with the environment, meaning that a large excess of molecules (80%) were diffusing freely in the defined regions and thus bioavailable (mean local diffusion coefficient, 7.1 ± 0.6 μm2/s).
FIG 2
FIG 2 FRAP acquisitions for BODIPY-FL-daptomycin inside S. aureus biofilms. (a) Kymogram representation (x,t) of FRAP acquisitions. The line along which the kymogram was done is 38 μm. (b) Typical fluorescence recovery curves representative of six different zones for each condition: BODIPY-FL-daptomycin inside biofilms (black) and inside TSBpc without biofilm (gray). The kymogram and fluorescence recovery curve presented here are the ones obtained for MSSA ATCC 27217 biofilms, since they were representative of the data obtained for the other strains in the presence or absence of rifampin.

(iii) Localization of the fluorescently labeled daptomycin alone or combined with rifampin depending on the surrounding environment.

As daptomycin is known to be highly bound to serum proteins (90 to 93%) (26), we addressed the question of whether there could be a different localization of the fluorescently labeled daptomycin in a protein-enriched medium. As shown in Fig. 3, fluorescence intensity images were significantly different depending on the surrounding medium: regardless of the strain tested, the antibiotic appeared to be colocalized with the FM4-64 dye at the bacterial site when the surrounding medium was an aqueous NaCl solution supplemented with calcium, while the antibiotic localization appeared to be mainly extracellular when the medium was enriched with proteins (TSBpc). The addition of rifampin did not affect daptomycin localization in the biofilm.
FIG 3
FIG 3 Fluorescence imaging of BODIPY-FL-daptomycin (green channel) and FM4-64 (red channel) in S. aureus biofilms. Merged images are also shown. In the top panels (Without proteins), the surrounding medium is an aqueous NaCl (150 mM) solution supplemented with calcium ions (50 mg/liter). In the bottom panels (With proteins), the surrounding medium is TSB enriched with proteins (36 g/liter) and calcium ions (50 mg/liter). Only images of MSSA ATCC 27217 biofilms are represented, since they were representative of all biofilms visualized for other strains in the presence or absence of rifampin.

Combining 3D fluorescence imaging and time-kill studies to assess S. aureus biofilm inactivation in the presence of daptomycin alone and in association with rifampin.

We used confocal microscopy to describe the three-dimensional structures of biofilms and the temporal distribution of both live and dead cells throughout the biofilm thickness.
Fluorescence intensity images showed that the biofilms formed by the four strains yielded similar compact structures (controls in Fig. 4a and controls in Fig. S2 in the supplemental material). Their thicknesses were not significantly variable from one strain to another (25 to 29 μm; P > 0.05) or from one time point to another time point (24 h and 72 h). This is a reasonable result given that the culture medium (TSBpc) was renewed before the first observation but not over time: thus, biofilm development mainly occurred during the first 24 h.
FIG 4
FIG 4 (a) Visualization of MSSA ATCC 27217 and MRSA BCB8 biofilms using 3D reconstruction to observe biofilm thickness. Images were collected without any drug exposure (control) and after 72-h exposure to unlabeled daptomycin (20 μg/ml) alone and in association with rifampin (20 μg/ml). Dead cells were stained red with propidium iodide, and all bacteria were stained green with Syto9. The acquisition was performed on the whole biofilm thickness with an axial displacement of 1 μm. The dimensions of the images are 82 by 82 μm2. The mean thickness values of the biofilms over time (from 24 to 72 h) are written in white on each image. Bar, 20 μm. (b) Percentage of dead cells over time calculated from three series of biofilm images. The biofilms were not treated with daptomycin (control) or were treated with daptomycin (20 μg/ml) alone or combined with rifampin (20 μg/ml). The values are means ± standard deviations (error bars).
When treated with daptomycin, biofilms exhibited more areas free of cells compared to the controls (Fig. 4a; see Fig. S2 in the supplemental material). This can be related to significant decreases in biofilm thicknesses (19 to 21 μm in the presence of daptomycin; P < 0.05) compared to the control biofilms. Moreover, in the presence of daptomycin, no statistically significant change in the proportion of cell death over time was quantified (10 to 30%; P > 0.05) (Fig. 4b). However, the MRSA clinical isolate (BCB8) was more susceptible to daptomycin: ∼60% of cell death was quantified at 24 h, a value that decreased beyond 24 h (40% at 72 h; P < 0.05) due to cell regrowth (see below).
Compared to the monotherapy treatment, the structures and thicknesses of biofilms were not affected when they were treated with daptomycin in combination with rifampin. However, under these conditions, a significantly higher proportion of cell death was observed, achieving 85% at 72 h (P < 0.01) (Fig. 4b). For the two MSSA strains (ATCC 27217 and strain 176) and the MRSA collection strain (ATCC 33591), the antibiotic association activity (daptomycin plus rifampin) gradually increased over time (Fig. 4), while the maximum activity against the MRSA clinical strain (BCB8) was reached within 24 h.
Another observation of interest (Fig. 4a) is that upon daptomycin exposure, dead cells were observed over the whole biofilm depth, including the basal layer of cells in contact with the substratum, providing further evidence of the antibiotic penetration throughout the deepest layers of the biofilms. This result is even more pronounced for the BCB8 strain because of the greater proportion of dead cells involved by the action of daptomycin. This process was also observed when daptomycin was used in association with rifampin.
These data from fluorescence imaging were supported by CFU counts of suspended biofilms (Fig. 5). The results confirm that daptomycin alone was ineffective against biofilms formed by the two MSSA strains (ATCC 27217 and strain 176) and the MRSA collection strain (ATCC 33591). For the MRSA clinical isolate (BCB8), reduction in bacterial counts of only ∼1 to 2 log units was measured at 24 h before regrowth was observed to reach the same values as for the MSSA biofilms.
FIG 5
FIG 5 Time-kill curves of daptomycin (20 μg/ml) alone or combined or with rifampin (20 μg/ml) against MSSA ATCC 27217 and MRSA BCB8 biofilms. The values are means ± standard deviations (error bars).
In accordance with fluorescence imaging data, the activity of the combination of antibiotics was much greater than that of monotherapy. The sensitivity of the microbiological method allowed us to determine that the cell population decreased by ∼4 log units after 72 h of treatment (P < 0.05). The emergence of rifampin-resistant mutants was verified. As presented in Fig. 6, daptomycin dramatically prevented the emergence of rifampin-resistant mutants when the combination of daptomycin and rifampin was used.
FIG 6
FIG 6 Number of rifampin-resistant mutants determined in MSSA ATCC 27217 biofilms counted on rifampin-containing TSA plates. The biofilms were treated with rifampin alone (20 μg/ml) or with the daptomycin-rifampin combination (20 μg/ml for both antibiotics). Above each bar is shown the percentage of rifampin-resistant mutants in S. aureus biofilms among the total bacterial population. Error bars represent the standard deviations.

DISCUSSION

The choice of antibiotics to treat S. aureus BAI remains a challenge for the medical community. In this context, the ambivalence of the published results on daptomycin activity is a relevant example. Despite increasing data about daptomycin as an option to treat implant-associated S. aureus infections, as many failures (18, 27) as successes (79, 12) have been reported both in clinical practice and in laboratory models. This highlights that S. aureus BAI resistance/tolerance mechanisms to antimicrobials deserve more attention.
The biofilm-associated exopolymeric matrix may be considered to act as a shield to the antimicrobial diffusion reaction (2832) by delaying its penetration and/or reducing its bioavailability. To verify this hypothesis noninvasively, we took advantage of dynamic fluorescence imaging methods: confocal microscopy, time-lapse imaging, and FRAP. For our biofilm model and whatever the bacterial strain, no failure of daptomycin penetrability or bioavailability was observed. The opposite finding described by Siala et al. (31) may be related to the conditions of fluorescence acquisition that were not well adapted to BODIPY-FL fluorescence. In this study, time-lapse fluorescence imaging experiments demonstrated that daptomycin rapidly reached the biofilm's deepest layers, while section views of fluorescence intensity images presented in Fig. 3 ascertain the presence of the fluorescently labeled antibiotic through the whole biofilm structure. Furthermore, FRAP results ascertained that only ∼20% of the antibiotic molecules were immobilized. Thus, the majority of the antibiotic molecules were in free movement and could be bioavailable through the biomass (∼80% of nonimmobilized molecules).
We further addressed the question of whether or not daptomycin reached its bacterial target. Fluorescence intensity images provided interesting information, showing that the majority of fluorescently labeled antibiotic was distributed in the extracellular matrix rather than in the bacterial cell membranes (Fig. 3). This is in agreement with the well-known property of daptomycin to have a very high degree of protein binding, especially with serum albumin (90 to 93%) (26, 33) which is naturally present in physiological conditions. Nevertheless, the fluorescence recovery curves obtained by FRAP experiments in free medium and in the biofilms strongly suggested the reversibility of daptomycin protein binding (33, 34): the equilibrium between the bound and unbound states may conserve the apparent mobility of the antibiotic. Additional experiments were performed in a protein-free medium (a saline solution supplemented with calcium ions). In this case, bacterial cell membranes appeared as hot spots on fluorescence images, consistently with the described antibiotic interaction with its target (6). Surprisingly, whether the medium was protein-free or not, daptomycin exhibited the same lack of effectiveness, as revealed by time-kill studies performed by fluorescent LIVE/DEAD staining and conventional plating on agar (data in the absence of proteins not shown). Thus, the interaction with the matrix components cannot explain biofilm tolerance to the antibiotic.
Thus, the particular physiology of embedded bacteria should be considered, and more specifically, cells with low metabolic activity should be investigated. Previous studies using a bromodeoxyuridine (BrdU) immunofluorescent labeling technique demonstrated that the large majority of staphylococcal cells in a biofilm were actually in a low metabolic state (35, 36). Additionally, in the present study, the comparison of cell viability results obtained by CFU counts and fluorescence imaging highlighted a significant proportion of viable cells detected by LIVE/DEAD staining but not by CFU measurements. This subpopulation may be considered viable but nonculturable (VBNC), a subpopulation known to have a slow metabolism (3739). Moreover, it has been demonstrated that daptomycin is poorly effective against bacteria in stationary stage (7, 27). One can thus reasonably suggest that for bacteria with low metabolic activity, the cell membrane dysregulation induced by daptomycin may be slower and/or more difficult to attain due to structural modifications of the cell membrane. This assumption is supported by the reported data revealing that daptomycin displays a concentration-dependent bactericidal activity against dormant cells (7, 9, 12). In the present study, we tested a higher concentration (double) of daptomycin (40 μg/ml) on the different S. aureus biofilms, almost leading to bactericidal effects after 24 h of drug exposure (see Fig. S3 in the supplemental material) and showing no cell regrowth over time. However, biofilm clearance was not reached. This achievement was reported to occur at a daptomycin concentration equal to or greater than 100 μg/ml but may not be relevant in clinical practice (7, 9, 12).
Two distinctive findings in this study concern the BCB8 clinical isolate, which discriminated itself by a twofold-higher penetration coefficient compared to the other strains tested and a greater susceptibility as revealed by the observation of a larger proportion of dead cells over the whole biofilm thickness, including the basal layer in contact with the substratum. These results are in line with those obtained in vivo (18), which demonstrated a strain-dependent activity of daptomycin against S. aureus biofilms. In view of the antibiotic mechanism of action which is supposed to target the plasma membrane, the observed variable response depending on the bacterial strain may be due to a change in membrane composition or conformation from a strain to another.
Facing the lack of daptomycin efficiency in treating recalcitrant S. aureus BAI, the addition of rifampin has raised great interest (1013, 15, 4042). In this study, we demonstrated that the combined therapy was indeed highly efficient against S. aureus biofilms but did not allow total bacterial clearance. Both antibiotics have been shown not to cross-react with each other, as evidenced by steady-state fluorescence spectroscopy. Moreover, the penetration, diffusion, and localization of the fluorescently labeled daptomycin were not affected by the presence of rifampin. We also proved here that rifampin-resistant mutants emerged when biofilms were treated with rifampin alone, but not when treated with the antibiotic combination. Altogether, the data presented here confirm that daptomycin prevents the emergence of rifampin-resistant mutants, allowing the bactericidal activity of rifampin to occur quickly, regardless of the cell physiological state.
In conclusion, consistently with the previous in vivo study aiming at evaluating the antibiotic efficacy in S. aureus prosthetic vascular graft infections (18), we demonstrated in the present in vitro model a strain-dependent lack of daptomycin activity toward biofilms. Dynamic fluorescence microscopy allowed discarding a lack of antibiotic availability and interaction with bacteria. Given the mode of action of daptomycin, these observations suggest a membrane-dependent factor of tolerance in such biofilms. Therefore, to provide a better understanding of the reduced activity of daptomycin against biofilms, the composition of the membrane should be analyzed.

ACKNOWLEDGMENTS

We thank the Centre de Photonique Biomédicale (CPBM) of the Centre Laser de l'Université Paris-Sud (CLUPS/LUMAT FR2764, Orsay, France) for allowing us to use the confocal microscope and L2 microbiology facilities, Rachel Méallet-Renault for allowing us to use the spectrofluorimeter facilities at the Ecole Normale Supérieure (ENS Cachan), and Jared Silverman (Cubist Pharmaceuticals) for providing BODIPY-FL-labeled daptomycin.
This work was supported by a grant from the Ministère de l'Education Nationale, de l'Enseignement Supérieur et de la Recherche, Université Paris-Sud, for Rym Boudjemaa's Ph.D. thesis (grant 2014-172).

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cover image Antimicrobial Agents and Chemotherapy
Antimicrobial Agents and Chemotherapy
Volume 60Number 8August 2016
Pages: 4983 - 4990
PubMed: 27297479

History

Received: 31 March 2016
Returned for modification: 1 May 2016
Accepted: 3 June 2016
Published online: 22 July 2016

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Authors

Rym Boudjemaa
Institut des Sciences Moléculaires d'Orsay (ISMO), CNRS, Université Paris-Sud, Université Paris-Saclay, Orsay, France
Romain Briandet
Micalis Institute, INRA, AgroParisTech, Université Paris-Saclay, Jouy-en-Josas, France
Matthieu Revest
CHU Rennes, Rennes, France
Université de Nantes, Faculté de Médecine, UPRES EA 3826, Nantes, France
Cédric Jacqueline
Université de Nantes, Faculté de Médecine, UPRES EA 3826, Nantes, France
Jocelyne Caillon
Université de Nantes, Faculté de Médecine, UPRES EA 3826, Nantes, France
Marie-Pierre Fontaine-Aupart
Institut des Sciences Moléculaires d'Orsay (ISMO), CNRS, Université Paris-Sud, Université Paris-Saclay, Orsay, France
Karine Steenkeste
Institut des Sciences Moléculaires d'Orsay (ISMO), CNRS, Université Paris-Sud, Université Paris-Saclay, Orsay, France

Notes

Address correspondence to Rym Boudjemaa, [email protected].

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