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. 2013 Nov;57(11):5572–5579. doi: 10.1128/AAC.00888-13

Effects of Bacteriocins on Methicillin-Resistant Staphylococcus aureus Biofilm

Ken-ichi Okuda a,✉, Takeshi Zendo b, Shinya Sugimoto a, Tadayuki Iwase a, Akiko Tajima a, Satomi Yamada a, Kenji Sonomoto b, Yoshimitsu Mizunoe a,✉
PMCID: PMC3811281  PMID: 23979748

Abstract

Control of biofilms formed by microbial pathogens is an important subject for medical researchers, since the development of biofilms on foreign-body surfaces often causes biofilm-associated infections in patients with indwelling medical devices. The present study examined the effects of different kinds of bacteriocins, which are ribosomally synthesized antimicrobial peptides produced by certain bacteria, on biofilms formed by a clinical isolate of methicillin-resistant Staphylococcus aureus (MRSA). The activities and modes of action of three bacteriocins with different structures (nisin A, lacticin Q, and nukacin ISK-1) were evaluated. Vancomycin, a glycopeptide antibiotic used in the treatment of MRSA infections, showed bactericidal activity against planktonic cells but not against biofilm cells. Among the tested bacteriocins, nisin A showed the highest bactericidal activity against both planktonic cells and biofilm cells. Lacticin Q also showed bactericidal activity against both planktonic cells and biofilm cells, but its activity against biofilm cells was significantly lower than that of nisin A. Nukacin ISK-1 showed bacteriostatic activity against planktonic cells and did not show bactericidal activity against biofilm cells. Mode-of-action studies indicated that pore formation leading to ATP efflux is important for the bactericidal activity against biofilm cells. Our results suggest that bacteriocins that form stable pores on biofilm cells are highly potent for the treatment of MRSA biofilm infections.

INTRODUCTION

A biofilm is a community of microorganisms that adheres to biotic and abiotic surfaces. In a biofilm, cells form a cluster embedded in a self-produced extracellular matrix consisting of biomolecules, such as polysaccharides, proteins, nucleic acids, and lipids (1, 2). Bacterial cells in a biofilm exhibit resistance to the host's immune system and antibiotic treatments (3–6). Therefore, once biofilms form on surfaces of medical devices, such as indwelling vascular catheters, prosthetic joints, and cardiac pacemakers, they can be extremely difficult to eradicate, and invasive procedures such as removal of the infected device may be required (7). Biofilm-associated infections are often chronic or relapsing; therefore, successful prevention and treatment of these infections are significant issues.

Bacteriocins are bacterially produced peptides or proteins that show bacteriostatic and/or bactericidal activity against other bacteria. It has been suggested that bacteriocins are valuable alternatives to antibiotics because they have many properties (e.g., significant potency, high stability, low toxicity, and both broad and narrow spectra of activity) that make them suitable for clinical applications (8). Bacteriocins produced by Gram-positive bacteria are classified into two distinct categories: the lanthionine-containing bacteriocins (class I) and the non-lanthionine-containing bacteriocins (class II) (9). The class I bacteriocins, termed lantibiotics, are characterized by the presence of unusual amino acids, such as lanthionine, 3-methyllanthionine, dehydroalanine, and dehydrobutyrine (10, 11). The lanthionine residue consists of two alanine residues cross-linked via a thioether linkage; therefore, lantibiotics are typically characterized by cyclic structures. The unusual amino acids are introduced by posttranslational modification and contribute to the high stability of the lantibiotics against many stresses (10, 11). The class II bacteriocins are small heat-stable peptides without unusual amino acids. Cotter et al. proposed a classification of class II bacteriocins into 4 groups: IIa (pediocin-like bacteriocins), IIb (two-peptide bacteriocins), IIc (cyclic bacteriocins), and IId (nonpediocin single linear peptides) (9). To date, many bacteriocins have been identified, and their structures, modes of action, and biosynthetic mechanisms have been studied. Nisin A, a lantibiotic produced by Lactococcus lactis (12), uses lipid II, which is a membrane-bound precursor of the cell wall, as a docking molecule (13–15). Nisin A is known to possess a dual mode of action: inhibition of cell wall synthesis by masking lipid II (bacteriostatic) and membrane insertion followed by pore formation that rapidly kills cells (bactericidal). Nukacin ISK-1, a lantibiotic produced by Staphylococcus warneri ISK-1 (16), is also known to bind lipid II (17). Unlike nisin A, nukacin ISK-1 has only a bacteriostatic mode of action, suggesting that nukacin ISK-1 prevents cell wall synthesis but does not form pores leading to cell death (18). Nisin A and nukacin ISK-1 belong to distinct subgroups of lantibiotics on the basis of their structures (Fig. 1); nisin A belongs to type AI lantibiotics (linear), and nukacin ISK-1 belongs to type AII lantibiotics (linear N terminus and globular C terminus) (10, 11). Lacticin Q, a class IId bacteriocin produced by Lactococcus lactis QU 5 (19), forms huge toroidal pores, causing leakage of protein molecules from target cells with lipid flip-flop (20), thus following a bactericidal mode of action. Importantly, lacticin Q does not require a specific receptor, such as lipid II, for its activity (21).

Fig 1.

Fig 1

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Structures of bacteriocins. The shaded residues indicate modified amino acids. Ala-S-Ala, lanthionine; Abu-S-Ala, 3-methyllanthionine; Dha, dehydroalanine; Dhb, dehydrobutyrine; fMet, formylmethionine.

The emergence of methicillin-resistant Staphylococcus aureus (MRSA) is a significant public health problem worldwide (22). Recently, we reported that S. aureus MR23, a clinically isolated MRSA strain, has the potential to form a robust biofilm with a matrix primarily consisting of proteins (2). In this study, we investigated the antibacterial effects of three bacteriocins (nisin A, lacticin Q, and nukacin ISK-1; Fig. 1) and vancomycin against planktonic and biofilm cells of S. aureus MR23 and several staphylococcal biofilms. In addition, mode-of-action studies of these bacteriocins and vancomycin against biofilm cells were carried out by measuring ATP efflux and membrane potential. Our data suggest that bacteriocins that form stable pores on the membrane of target cells are effective against MRSA biofilms and have the potential to prevent or cure biofilm-associated infections.

MATERIALS AND METHODS

Bacterial strains and culture conditions.

S. aureus and S. epidermidis strains were cultured in brain heart infusion (BHI; BD Biosciences, Franklin Lakes, NJ) medium at 37°C with shaking. L. lactis QU 5 was cultured in MRS (Oxoid, Basingstoke, United Kingdom) medium at 30°C under static conditions. S. warneri ISK-1 was cultured in tryptic soy broth (BD Biosciences) at 37°C with shaking.

Bacteriocin purification.

Nisin A was purified from a commercial preparation (Sigma-Aldrich, St. Louis, MO) as described previously (23). Nukacin ISK-1 was purified from an overnight culture supernatant of S. warneri ISK-1 as described previously (23). Lacticin Q was purified from an overnight culture supernatant of L. lactis QU 5 by using the same procedure used for purification of nukacin ISK-1.

MIC determination.

S. aureus and S. epidermidis strains were precultured in BHI medium overnight, inoculated into fresh BHI medium, and incubated until an optical density of 0.05 at 595 nm was attained. Aliquots (20 μl) of serially diluted peptide solutions (nisin A, lacticin Q, nukacin ISK-1, or vancomycin) were added to the wells of a 96-well flat-bottomed polystyrene plate (Corning, Corning, NY) containing 80 μl of the bacterial culture. After culturing for 24 h at 37°C with shaking, the absorbance at 595 nm was measured using an Infinite 200 PRO microplate reader (Tecan, Mannedorf, Switzerland). The MICs were defined as the lowest concentrations of the peptide that completely inhibited bacterial growth.

Bactericidal activity against S. aureus MR23 planktonic cells.

S. aureus MR23 was precultured in BHI medium overnight, inoculated into fresh BHI medium, and incubated until an optical density of 0.4 at 595 nm was attained. Peptide solutions (50 μl) were added to 150 μl of the bacterial culture in a 96-well flat-bottomed polystyrene plate to a final concentration of 4× MIC and incubated at 37°C with shaking. The numbers of CFU on BHI agar were enumerated at different time points (0, 1, 4, 8, 16, and 24 h).

Bactericidal activity against S. aureus MR23 biofilm cells.

S. aureus MR23 was precultured in BHI medium overnight and diluted 1,000-fold with BHI medium supplemented with 1% glucose (BHI-G). Aliquots (200 μl) of this bacterial suspension were added to the wells of a 96-well flat-bottomed polystyrene plate and incubated for 24 h at 37°C. The supernatants were removed, and the biofilms formed on the bottoms of the wells were washed twice with phosphate-buffered saline (PBS). Biofilms were incubated with 50 μl of the peptide solutions in PBS at a concentration of 4× MIC for 1 and 24 h at room temperature. After removal of the supernatants, the biofilms were washed twice and then resuspended in 100 μl of PBS. The suspensions were sonicated for 30 s, and the numbers of CFU on BHI agar were counted.

Dose dependencies of bacteriocins and vancomycin against S. aureus MR23 biofilm.

Biofilms were formed on a 96-well flat-bottomed polystyrene plate as described above and incubated with 50 μl of the peptide solutions in PBS at a concentration of 4× MIC for 1 h at room temperature. After removal of the supernatants, the biofilms were washed twice with PBS. Living and dead cells in the biofilms were stained using a FilmTracer LIVE/DEAD biofilm viability kit (Invitrogen, Carlsbad, CA) according to the manufacturer's protocol. After two washes with PBS, the fluorescence derived from propidium iodide (PI) was measured with an Infinite 200 PRO microplate reader at excitation and emission wavelengths of 485 and 635 nm, respectively.

Three-dimensional live/dead imaging of S. aureus and S. epidermidis biofilms.

S. aureus and S. epidermidis strains were precultured in BHI medium overnight and diluted 1,000-fold with BHI-G medium. An aliquot of 2 ml of this bacterial suspension was added to a 35-mm glass-bottomed dish (Iwaki, Tokyo, Japan) and incubated for 24 h at 37°C. The supernatant was removed, and the biofilm formed at the bottom of the dish was washed twice with PBS. The biofilm was incubated with 100 μl of peptide solution in PBS at a concentration of 4× MIC for 1 h at room temperature. After removal of the supernatant, the biofilm was washed with PBS and stained using a FilmTracer LIVE/DEAD biofilm viability kit according to the manufacturer's protocol. After washing with PBS, the biofilm was observed by fluorescence microscopy (BZ9000; Keyence, Tokyo, Japan). A three-dimensional image was constructed by stacking multiple images with different z-foci.

ATP efflux from S. aureus MR23 biofilm.

Biofilms were formed on a 96-well flat-bottomed polystyrene plate as described above and incubated with 50 μl of the peptide solutions in PBS at a concentration of 4× MIC for 1 h at room temperature. The supernatants were filtered using a 0.22-μm-pore-size centrifugal filter (Millipore, Billerica, MA) and diluted 10-fold with water. The ATP concentrations of the filtrates were measured using the BacTiter-Glo reagent (Promega, Madison, WI) according to the manufacturer's protocol. The bioluminescence response in relative light units was detected with an Infinite 200 PRO microplate reader. The ATP concentration of each sample was determined using standard ATP solutions.

Measurement of membrane potential of S. aureus MR23 biofilm.

Biofilms were formed on a 96-well flat-bottomed polystyrene plate as described above. The membrane potentials of the biofilm cells were measured using a membrane-potential-sensitive probe, bis-(1,3-dibutylbarbituric acid)trimethine oxonol [DiBAC4(3); Invitrogen]. The biofilms were preincubated with 100 μl of 5 μM DiBAC4(3) in PBS for 30 min. After removal of the supernatant, the biofilms were incubated with 50 μl of solution containing 5 μM DiBAC4(3) and peptides at a concentration of 4× MIC in PBS for 1 h at room temperature. After two washes with PBS, the fluorescence derived from DiBAC4(3) was measured with an Infinite 200 PRO microplate reader at excitation and emission wavelengths of 485 and 535 nm, respectively.

Binding of fluorescently labeled vancomycin to S. aureus MR23 biofilm.

Biofilms were formed on a 96-well flat-bottomed polystyrene plate as described above. After two washes with PBS, the biofilms were incubated with 50 μl of 40 μM BODIPY FL and vancomycin (BODIPY-vancomycin; Invitrogen) dissolved in PBS for 0, 5, 15, 30, and 60 min at room temperature. After two washes with PBS, the fluorescence derived from BODIPY-vancomycin was measured with an Infinite 200 PRO microplate reader at excitation and emission wavelengths of 485 and 535 nm, respectively. After a 60-min incubation, the biofilms were washed with PBS to remove unbound BODIPY-vancomycin and resuspended, and the localization of BODIPY-vancomycin was analyzed by fluorescence microscopy using a green fluorescent protein filter (Eclipse E600; Nikon, Tokyo, Japan).

RESULTS

Susceptibility of S. aureus MR23 to bacteriocins and vancomycin under planktonic growth conditions.

At first, the MICs of nisin A, lacticin Q, nukacin ISK-1, and vancomycin against S. aureus MR23 were determined by the micro-liquid dilution method (Table 1). After addition of 4× MIC peptides to S. aureus MR23 cells adjusted to 108 CFU/ml in the culture medium, the numbers of CFU were counted following different incubation periods (Fig. 2). After the addition of nisin A and lacticin Q, which have a bactericidal mode of action, the cells were killed quickly. Incubation times for complete bactericidal action (i.e., no colony formation on BHI agar) were 1 and 4 h for nisin A and lacticin Q, respectively. Nukacin ISK-1 showed bacteriostatic activity against S. aureus MR23, consistent with its previously reported mode of action (18). Vancomycin showed bactericidal activity, but the number of viable cells after a 4-h incubation was 107 CFU/ml (Fig. 2), indicating that its killing rate is lower than that of nisin A and lacticin Q.

Table 1.

MICs of antimicrobial peptides

Bacterial straina MIC (μM) of antimicrobial peptide
Nisin A Lacticin Q Nukacin ISK-1 Vancomycin BODIPY-vancomycin
MR23 2.5 5.0 12 10 10
MR10 2.5 5.0 10 5 NTb
MR11 2.5 5.0 20 10 NT
SH1000 1.25 5.0 20 5 NT
SE4 1.25 >20 20 5 NT
SE21 1.25 >20 10 10 NT
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a

MR23, S. aureus MR23; MR10, S. aureus MR10; MR11, S. aureus MR11; SH1000, S. aureus SH1000; SE4, S. epidermidis SE4; SE21, S. epidermidis SE21.

b

NT, not tested.

Fig 2.

Fig 2

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Bactericidal activities of bacteriocins and vancomycin against planktonic cells. Peptides at final concentrations of 4× MIC were incubated with S. aureus MR23 in BHI medium and further incubated at 37°C with shaking. The CFU were enumerated following different incubation periods (0, 1, 4, and 24 h). Data points represent the means and standard deviations of triplicate determinations.

Antibacterial activities of bacteriocins and vancomycin against S. aureus MR23 biofilm cells.

Biofilms of S. aureus MR23 formed on a 96-well plate were incubated with the peptides at a concentration of 4× MIC for 1 and 24 h; subsequently, the CFU of the biofilms suspended in PBS were enumerated. Nisin A and lacticin Q showed bactericidal activities against biofilm cells, but complete killing (as observed for planktonic cells) was not achieved (Fig. 3). After a 24-h incubation, the bactericidal activity of nisin A was about 10 times higher than that of lacticin Q. Nukacin ISK-1, which showed bacteriostatic activity against planktonic cells, did not show bactericidal activity against biofilm cells. Vancomycin showed bactericidal activity against planktonic cells, but it did not show bactericidal activity against biofilm cells under the experimental conditions used.

Fig 3.

Fig 3

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Bactericidal activities of bacteriocins and vancomycin against biofilm cells. Peptide solutions at a concentration of 4× MIC were incubated with preformed biofilms of S. aureus MR23 at room temperature. The numbers of CFU of biofilm cells were measured after 1-h (white bars) and 24-h (black bars) incubations. The means and standard deviations of triplicate determinations are presented.

Diffusion of bacteriocins into S. aureus MR23 biofilm.

After treatment with bacteriocins or vancomycin, the biofilms were stained with SYTO9 and PI (components of the FilmTracer LIVE/DEAD biofilm viability kit), which stain the living and dead cells, respectively. Subsequently, three-dimensional images of the stained biofilms were observed using fluorescence microscopy (Fig. 4). After incubation of the biofilm with PBS for 60 min, living cells constituted the majority of the biofilm (Fig. 4A). After addition of nisin A, the proportion of dead cells was significantly higher, and it increased with the incubation time (Fig. 4B and C). Lacticin Q also increased the proportion of dead cells, but not as much as nisin A (Fig. 4D). Addition of vancomycin or nukacin ISK-1 had no effect on the viability of the biofilm cells (Fig. 4E and F). These results are consistent with the CFU counts of the biofilms after treatment with the peptides. It should be noted that dead cells in the biofilms treated with nisin A and lacticin Q seemed to be uniformly distributed throughout the biofilm (Fig. 4B′ to D′). If the biofilm matrix blocked the penetration of the bacteriocins, dead cells would be found primarily at the top of the biofilm. Therefore, it is suggested that both nisin A and lacticin Q penetrated the biofilm matrix and reached the deepest part of the biofilm.

Fig 4.

Fig 4

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Three-dimensional images of S. aureus MR23 biofilms after treatment with bacteriocins and vancomycin. Biofilms formed on 35-mm glass-bottomed dishes were incubated with peptide solutions at a concentration of 4× MIC for 1 h at room temperature. After washing, the living and dead cells in the biofilm were stained by SYTO9 (green) and PI (red), respectively. (A) PBS (60 min); (B) nisin A (5 min); (C) nisin A (60 min); (D) lacticin Q (60 min); (E) nukacin ISK-1 (60 min); (F) vancomycin (60 min); (A′ to D′) enlarged views of x-z sections of panels A to D, respectively. Bars, 5 μm.

Modes of action of bacteriocins against S. aureus MR23 biofilm.

To determine the effective concentrations of the peptides against biofilm cells, serially diluted peptides were added to the biofilms and the fluorescence derived from dead cells stained by PI was measured with a fluorometer (Fig. 5). The fluorescence intensity of cells treated with nisin A was higher than that of untreated cells, and it increased with the nisin A concentration. The increase in fluorescence was observed even when 0.25× MIC (0.625 μM) nisin A was added. Addition of lacticin Q also increased the fluorescence intensity of the biofilm in a dose-dependent manner, but the intensity was significantly lower than that of the biofilm treated with nisin A. Nukacin ISK-1 and vancomycin did not increase the fluorescence even at 8× MIC (96 and 80 μM, respectively). None of the peptides reduced the amount of biofilm, even at 8× MIC (data not shown), indicating that they do not have destructive activity against the established S. aureus MR23 biofilm. Next, we analyzed the efflux of ATP molecules from the biofilm cells by luciferase assay (Fig. 6). The amount of ATP leaked from biofilm cells increased in a dose-dependent manner following treatment with nisin A or lacticin Q. Nisin A caused ATP efflux at 0.25× MIC (0.625 μM), whereas the minimum concentration of lacticin Q required for ATP efflux was 1× MIC (5.0 μM). Nukacin ISK-1 and vancomycin had no effect on ATP efflux even at 8× MIC (96 and 80 μM, respectively). In addition, to analyze the effects of the peptides on membrane integrity, we measured the membrane potentials of the biofilm cells by using DiBAC4(3), a membrane-potential-sensitive probe (24) (Fig. 7). As might be expected from the results of ATP efflux measurements, nisin A and lacticin Q disrupted the membrane potentials of biofilm cells, and vancomycin did not. Interestingly, nukacin ISK-1 disrupted the membrane potential of the biofilm cells, but it did not cause ATP efflux.

Fig 5.

Fig 5

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Dose-dependent effects of bacteriocins and vancomycin against S. aureus MR23 biofilm. Peptide solutions at a concentration of 4× MIC were incubated with preformed S. aureus MR23 biofilms for 1 h at room temperature. Subsequently, biofilms were stained using a FilmTracer LIVE/DEAD biofilm viability kit. Fluorescence derived from PI was measured at excitation and emission wavelengths of 485 and 635 nm, respectively. The means and standard deviations of triplicate determinations are presented. A.U., arbitrary units.

Fig 6.

Fig 6

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ATP efflux from S. aureus MR23 biofilm cells. After incubation of S. aureus MR23 biofilm cells with bacteriocins and vancomycin at a concentration of 4× MIC for 1 h at room temperature, the amount of ATP in the supernatants was measured by luciferase assay. ATP concentrations were determined using standard ATP solutions. The means and standard deviations of triplicate determinations are presented.

Fig 7.

Fig 7

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Membrane potential of S. aureus MR23 biofilm after incubation with bacteriocins and vancomycin. Fluorescence derived from DiBAC4(3), a membrane-potential-sensitive probe, was measured at excitation and emission wavelengths of 485 and 535 nm, respectively. The means and standard deviations of triplicate determinations are presented. A.U., arbitrary units.

Localization of fluorescently labeled vancomycin in S. aureus MR23 biofilm.

Vancomycin showed no bactericidal activity or membrane depolarization activity against biofilm cells. A possible explanation is that the biofilm matrix physically inhibited vancomycin penetration, so that vancomycin was not able to attack the biofilm cells. To test this hypothesis, we analyzed the localization of BODIPY-vancomycin (fluorescently labeled vancomycin) in the S. aureus MR23 biofilm. BODIPY labeling seemed to have no effect on the activity of vancomycin, since the MIC of BODIPY-vancomycin against S. aureus MR23 was identical to that of vancomycin (Table 1). After treatment of biofilm cells with BODIPY-vancomycin, the fluorescence derived from BODIPY-vancomycin was measured at different incubation times. The fluorescence increased with the incubation time and peaked after 30 min (Fig. 8A). After 60 min, the biofilm was suspended in PBS and observed by fluorescence microscopy. Fluorescence was observed at the surface of virtually every cell, and in some cells, the division septa were also stained (Fig. 8B). These results suggest that vancomycin can penetrate the S. aureus MR23 biofilm and bind to lipid II, a target molecule for vancomycin located at the division septum.

Fig 8.

Fig 8

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Localization analysis of BODIPY-vancomycin. (A) S. aureus MR23 biofilm was incubated with BODIPY-vancomycin at a concentration of 4× MIC for 0, 5, 15, 30, and 60 min at room temperature. Fluorescence derived from BODIPY-vancomycin was measured at excitation and emission wavelengths of 485 and 535 nm, respectively. The means and standard deviations of triplicate determinations are presented. van, vancomycin; A.U., arbitrary units. (B) After a 60-min incubation with BODIPY-vancomycin, the biofilm resuspended in PBS was observed by fluorescence microscopy. Arrows, division septa of the cells. (Inset) Enlargement of boxed area. Bar, 1 μm.

Effects of bacteriocins against other staphylococcal biofilms.

The activities of bacteriocins and vancomycin against biofilms of several clinical isolates of MRSA (S. aureus MR10 and MR11) and S. epidermidis (S. epidermidis SE4 and SE21) (F. Sato et al., unpublished data) and S. aureus SH1000 (25), a laboratory strain frequently used for biofilm assays, were investigated. The MICs of the peptides are shown in Table 1. After treatments of the biofilms with peptides at a concentration of 4× MIC for 1 h, followed by staining with SYTO9 and PI, live/dead images were observed. By way of exception, biofilms of SE4 and SE21 were treated with 20 μM lacticin Q, since the MICs of lacticin Q against these two strains were not determined at the highest concentration (Table 1). The x-y sections of the three-dimensional images are shown in Fig. 9. Among the tested peptides, nisin A showed the highest activity against all biofilms, and dead cells constituted the majority of the biofilms after treatments with nisin A. Nukacin ISK-1 and vancomycin did not affect the viability of the biofilm cells, as observed in assays using the MR23 biofilm. The MR10 biofilm was less sensitive to lacticin Q than the MR11 and SH1000 biofilms, even though the MICs of lacticin Q against the three strains were identical. Interestingly, lacticin Q was slightly active against the SE4 biofilm at 20 μM, which is a concentration below the MIC. These results suggest that biofilm formation affects susceptibility to lacticin Q and/or the antimicrobial property of lacticin Q.

Fig 9.

Fig 9

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Live/dead images of staphylococcal biofilms after treatment with bacteriocins and vancomycin. Biofilms formed on 35-mm glass-bottomed dishes were incubated with peptide solutions at a concentration of 4× MIC for 1 h at room temperature. By way of exception, biofilms of SE4 and SE21 were treated with 20 μM lacticin Q. After washing, the living and dead cells in the biofilm were stained by SYTO9 (green) and PI (red), respectively. x-z sections of three-dimensional images are shown. MR10, S. aureus MR10; MR11, S. aureus MR11; SH1000, S. aureus SH1000; SE4, S. epidermidis SE4; SE21, S. epidermidis SE21. Bar, 5 μm.

DISCUSSION

In this study, we investigated the effects of bacteriocins with different structures against a clinical isolate of MRSA. Among the tested bacteriocins, nisin A showed the highest bactericidal activities against both planktonic cells and biofilm cells. In the case of growing planktonic cells, nisin A and lacticin Q completely killed 108 cells/ml at 4× MIC (10 and 20 μM, respectively) after a 4-h incubation. However, 4× MIC nisin A and lacticin Q could not completely kill biofilm cells after a 24-h incubation. These results suggest that S. aureus MR23 is less susceptible to bacteriocins when growing as a biofilm than when growing under planktonic conditions. Although the mechanisms underlying the bacteriocin resistance of biofilm cells are not understood, nonspecific interactions between bacteriocin molecules and the components of the biofilm matrix and/or the physiological state of the biofilm cells might affect bacteriocin activity. Notably, the bactericidal activity of nisin A against MR23 biofilm cells was about 10-fold higher than that of lacticin Q (Fig. 3). Additionally, nisin A was significantly more effective than lacticin Q against biofilms of several strains of S. aureus and S. epidermidis (Fig. 9). These results suggest that nisin A attacks biofilm cells more effectively than lacticin Q. What is the key factor influencing the superior bactericidal activity of nisin A against biofilm cells? Nisin A and lacticin Q represent different classes of bacteriocins (class I and class IId, respectively) and have different structures, molecular weights, and modes of action. One possibility is that lipid II, a docking molecule for nisin A, contributes to the high activity of nisin A against biofilm cells. Nisin A is able to act against an anionic model membrane lacking lipid II (26–28). However, the presence of lipid II dramatically enhances the efficiency of nisin A, making nisin A active against susceptible cells at nanomolar concentrations (13). Mode-of-action studies with living cells and artificial membranes demonstrated that lipid II is not required for the activity of lacticin Q (21). Lacticin Q forms toroidal pores on the target membranes in the absence of a specific receptor (20), and the pore-forming activity depends on the physicochemical features of the membrane components (29). In the biofilm, the cells are surrounded by a matrix consisting of various kinds of macromolecules bonded by weak interactions, such as hydrogen bonds; hydrophobic, electrostatic, and van der Waals interactions; and divalent cation bridges (1). In such environments, specific docking molecules like lipid II may contribute to specific binding to target cells.

It has been reported that nukacin ISK-1, a bacteriocin belonging to class I, is bacteriostatic against planktonic cells and uses lipid II as a docking molecule (17, 18). In this study, nukacin ISK-1 did not show bactericidal activity against biofilm cells, although it disrupted the membrane potential of biofilm cells (Fig. 7). According to a previous report, SA-FF22, a bacteriocin produced by Streptococcus pyogenes FF22 with a structure similar to that of nukacin ISK-1 (30), disrupts the membrane potential by forming small short-lived pores (from 0.5 to 0.6 nm in diameter and with life spans of a few milliseconds) (31). Therefore, nukacin ISK-1 may have penetrated the biofilm, forming small short-lived pores that allow small ions to pass. The levels of ATP efflux from biofilms were well correlated with the bactericidal activity of each bacteriocin (Fig. 3 to 5). For effective killing of biofilm cells, a pore size large enough for ATP to pass (>1.5 nm in diameter) seems to be required. Indeed, nisin A forms large stable pores (from 2 to 2.5 nm in diameter and with life spans of a few seconds) on membranes containing lipid II (32), and lacticin Q forms huge toroidal pores (from 4.6 to 6.6 nm in diameter) on membranes without lipid II (20).

Vancomycin is widely used as a first-line therapy for serious MRSA infections. We have shown that vancomycin exhibits bactericidal activity against growing planktonic cells of S. aureus MR23 but not against biofilm cells. It has been reported that vancomycin inhibits cell wall synthesis by binding to the N-acyl–d-Ala–d-Ala moiety of lipid II and prevents cross-linking of peptidoglycan chains, thereby leading to bacterial cell death by osmotic lysis (33). Although the detailed molecular mechanisms underlying antibiotic resistance of biofilms remain unclear, one of the possibilities is that the matrix inhibits penetration of antimicrobial agents (3, 34). On the other hand, a recent study that analyzed the diffusion of BODIPY-vancomycin into an S. aureus biofilm by using time-resolved fluorescence microscopy suggested that the biofilm matrix is not an obstacle to the diffusion of molecules the size of antibiotics (35). The authors demonstrated that BODIPY-vancomycin penetrates into the deepest layer of the biofilm within minutes. In an earlier study, another group reported that penetration of BODIPY-vancomycin to the deepest layer took more than 1 h (36). The difference in the penetration time in the two studies might be due to differences between the components of the biofilm matrix produced by the S. aureus strains used in the studies. We estimated the penetration of BODIPY-vancomycin by measuring fluorescence intensity. We found that fluorescence reached a steady state after 30 min of incubation of the biofilm with BODIPY-vancomycin (Fig. 8A). Notably, BODIPY-vancomycin localized not only at cell surfaces but also at division septa (Fig. 8B). Since the division septum is the only site of active cell wall biosynthesis of S. aureus, antibiotics that inhibit the late steps of cell wall synthesis must be able to access lipid II at the septum (37, 38). Our results suggest that vancomycin penetrated the biofilm matrix and functionally bound to its molecular target. The lower susceptibility to vancomycin of cells in biofilms may be explained by the low growth rate of cells (3, 7, 39–41).

In conclusion, we have demonstrated that nisin A is significantly effective against MRSA and other staphylococcal biofilms. To prevent biofilm-associated infections in patients using central venous catheters, an antibiotic-lock technique has been attempted whereby the lumen of the catheter is flushed and then filled with an antibiotic solution. It has been demonstrated that an antibiotic lock of central venous catheters employing a vancomycin-containing solution prevents endoluminal catheter-related infection (42–44). In such clinical applications, pore-forming bacteriocins which have modes of actions similar to the mode of action of nisin A might be more effective than antibiotics for the prevention and treatment of biofilm-associated infections.

ACKNOWLEDGMENTS

This work was supported by the MEXT-supported Program for the Strategic Research Foundation at Private Universities, 2012 to 2016, and The Jikei University Research Fund.

We acknowledge Naruhiko Sawa (Faculty of Agriculture, Kyushu University) for technical support during bacteriocin purification. We also thank Yumi Tokoro for secretarial assistance.

Footnotes

Published ahead of print 26 August 2013

REFERENCES

  • 1.Flemming HC, Wingender J. 2010. The biofilm matrix. Nat. Rev. Microbiol. 8:623–633 [DOI] [PubMed] [Google Scholar]
  • 2.Sugimoto S, Iwamoto T, Takada K, Okuda K, Tajima A, Iwase T, Mizunoe Y. 2013. Staphylococcus epidermidis Esp degrades specific proteins associated with Staphylococcus aureus biofilm formation and host-pathogen interaction. J. Bacteriol. 195:1645–1655 [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 3.Costerton JW, Stewart PS, Greenberg EP. 1999. Bacterial biofilms: a common cause of persistent infections. Science 284:1318–1322 [DOI] [PubMed] [Google Scholar]
  • 4.Melchior MB, Fink-Gremmels J, Gaastra W. 2006. Comparative assessment of the antimicrobial susceptibility of Staphylococcus aureus isolates from bovine mastitis in biofilm versus planktonic culture. J. Vet. Med. B Infect. Dis. Vet. Public Health 53:326–332 [DOI] [PubMed] [Google Scholar]
  • 5.Vlastarakos PV, Nikolopoulos TP, Maragoudakis P, Tzagaroulakis A, Ferekidis E. 2007. Biofilms in ear, nose, and throat infections: how important are they? Laryngoscope 117:668–673 [DOI] [PubMed] [Google Scholar]
  • 6.Kania RE, Lamers GE, Vonk MJ, Dorpmans E, Struik J, Tran Ba Huy P, Hiemstra P, Bloemberg GV, Grote JJ. 2008. Characterization of mucosal biofilms on human adenoid tissues. Laryngoscope 118:128–134 [DOI] [PubMed] [Google Scholar]
  • 7.Donlan RM. 2001. Biofilm formation: a clinically relevant microbiological process. Clin. Infect. Dis. 33:1387–1392 [DOI] [PubMed] [Google Scholar]
  • 8.Cotter PD, Ross RP, Hill C. 2013. Bacteriocins—a viable alternative to antibiotics? Nat. Rev. Microbiol. 11:95–105 [DOI] [PubMed] [Google Scholar]
  • 9.Cotter PD, Hill C, Ross RP. 2005. Bacteriocins: developing innate immunity for food. Nat. Rev. Microbiol. 3:777–788 [DOI] [PubMed] [Google Scholar]
  • 10.Chatterjee C, Paul M, Xie L, van der Donk WA. 2005. Biosynthesis and mode of action of lantibiotics. Chem. Rev. 105:633–684 [DOI] [PubMed] [Google Scholar]
  • 11.Nagao J, Asaduzzaman SM, Aso Y, Okuda K, Nakayama J, Sonomoto K. 2006. Lantibiotics: insight and foresight for new paradigm. J. Biosci. Bioeng. 102:139–149 [DOI] [PubMed] [Google Scholar]
  • 12.Rogers LA. 1928. The inhibiting effect of Streptococcus lactis on Lactobacillus bulgaricus. J. Bacteriol. 16:321–325 [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 13.Breukink E, Wiedemann I, van Kraaij C, Kuipers OP, Sahl HG, de Kruijff B. 1999. Use of the cell wall precursor lipid II by a pore-forming peptide antibiotic. Science 286:2361–2364 [DOI] [PubMed] [Google Scholar]
  • 14.Breukink E, van Heusden HE, Vollmerhaus PJ, Swiezewska E, Brunner L, Walker S, Heck AJ, de Kruijff B. 2003. Lipid II is an intrinsic component of the pore induced by nisin in bacterial membranes. J. Biol. Chem. 278:19898–19903 [DOI] [PubMed] [Google Scholar]
  • 15.Hsu ST, Breukink E, Tischenko E, Lutters MA, de Kruijff B, Kaptein R, Bonvin AM, van Nuland NA. 2004. The nisin-lipid II complex reveals a pyrophosphate cage that provides a blueprint for novel antibiotics. Nat. Struct. Mol. Biol. 11:963–967 [DOI] [PubMed] [Google Scholar]
  • 16.Sashihara T, Kimura H, Higuchi T, Adachi A, Matsusaki H, Sonomoto K, Ishizaki A. 2000. A novel lantibiotic, nukacin ISK-1, of Staphylococcus warneri ISK-1: cloning of the structural gene and identification of the structure. Biosci. Biotechnol. Biochem. 64:2420–2428 [DOI] [PubMed] [Google Scholar]
  • 17.Islam MR, Nishie M, Nagao J, Zendo T, Keller S, Nakayama J, Kohda D, Sahl HG, Sonomoto K. 2012. Ring A of nukacin ISK-1: a lipid II-binding motif for type-A(II) lantibiotic. J. Am. Chem. Soc. 134:3687–3690 [DOI] [PubMed] [Google Scholar]
  • 18.Asaduzzaman SM, Nagao J, Iida H, Zendo T, Nakayama J, Sonomoto K. 2009. Nukacin ISK-1, a bacteriostatic lantibiotic. Antimicrob. Agents Chemother. 53:3595–3598 [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 19.Fujita K, Ichimasa S, Zendo T, Koga S, Yoneyama F, Nakayama J, Sonomoto K. 2007. Structural analysis and characterization of lacticin Q, a novel bacteriocin belonging to a new family of unmodified bacteriocins of gram-positive bacteria. Appl. Environ. Microbiol. 73:2871–2877 [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 20.Yoneyama F, Imura Y, Ohno K, Zendo T, Nakayama J, Matsuzaki K, Sonomoto K. 2009. Peptide-lipid huge toroidal pore, a new antimicrobial mechanism mediated by a lactococcal bacteriocin, lacticin Q. Antimicrob. Agents Chemother. 53:3211–3217 [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 21.Yoneyama F, Imura Y, Ichimasa S, Fujita K, Zendo T, Nakayama J, Matsuzaki K, Sonomoto K. 2009. Lacticin Q, a lactococcal bacteriocin, causes high-level membrane permeability in the absence of specific receptors. Appl. Environ. Microbiol. 75:538–541 [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 22.Zetola N, Francis JS, Nuermberger EL, Bishai WR. 2005. Community-acquired methicillin-resistant Staphylococcus aureus: an emerging threat. Lancet Infect. Dis. 5:275–286 [DOI] [PubMed] [Google Scholar]
  • 23.Aso Y, Okuda K, Nagao J, Kanemasa Y, Thi Bich Phuong N, Koga H, Shioya K, Sashihara T, Nakayama J, Sonomoto K. 2005. A novel type of immunity protein, NukH, for the lantibiotic nukacin ISK-1 produced by Staphylococcus warneri ISK-1. Biosci. Biotechnol. Biochem. 69:1403–1410 [DOI] [PubMed] [Google Scholar]
  • 24.Pag U, Oedenkoven M, Papo N, Oren Z, Shai Y, Sahl HG. 2004. In vitro activity and mode of action of diastereomeric antimicrobial peptides against bacterial clinical isolates. J. Antimicrob. Chemother. 53:230–239 [DOI] [PubMed] [Google Scholar]
  • 25.Horsburgh MJ, Aish JL, White IJ, Shaw L, Lithgow JK, Foster SJ. 2002. SigmaB modulates virulence determinant expression and stress resistance: characterization of a functional rsbU strain derived from Staphylococcus aureus 8325-4. J. Bacteriol. 184:5457–5467 [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 26.Ruhr E, Sahl HG. 1985. Mode of action of the peptide antibiotic nisin and influence on the membrane potential of whole cells and on cytoplasmic and artificial membrane vesicles. Antimicrob. Agents Chemother. 27:841–845 [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 27.Demel RA, Peelen T, Siezen RJ, De Kruijff B, Kuipers OP. 1996. Nisin Z, mutant nisin Z and lacticin 481 interactions with anionic lipids correlate with antimicrobial activity. A monolayer study. Eur. J. Biochem. 235:267–274 [DOI] [PubMed] [Google Scholar]
  • 28.van Kraaij C, Breukink E, Noordermeer MA, Demel RA, Siezen RJ, Kuipers OP, de Kruijff B. 1998. Pore formation by nisin involves translocation of its C-terminal part across the membrane. Biochemistry 37:16033–16040 [DOI] [PubMed] [Google Scholar]
  • 29.Yoneyama F, Ohno K, Imura Y, Li M, Zendo T, Nakayama J, Matsuzaki K, Sonomoto K. 2011. Lacticin Q-mediated selective toxicity depending on physicochemical features of membrane components. Antimicrob. Agents Chemother. 55:2446–2450 [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 30.Dufour A, Hindre T, Haras D, Le Pennec JP. 2007. The biology of lantibiotics from the lacticin 481 group is coming of age. FEMS Microbiol. Rev. 31:134–167 [DOI] [PubMed] [Google Scholar]
  • 31.Jack R, Benz R, Tagg J, Sahl HG. 1994. The mode of action of SA-FF22, a lantibiotic isolated from Streptococcus pyogenes strain FF22. Eur. J. Biochem. 219:699–705 [DOI] [PubMed] [Google Scholar]
  • 32.Wiedemann I, Benz R, Sahl HG. 2004. Lipid II-mediated pore formation by the peptide antibiotic nisin: a black lipid membrane study. J. Bacteriol. 186:3259–3261 [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 33.Barna JC, Williams DH. 1984. The structure and mode of action of glycopeptide antibiotics of the vancomycin group. Annu. Rev. Microbiol. 38:339–357 [DOI] [PubMed] [Google Scholar]
  • 34.Stewart PS. 1998. A review of experimental measurements of effective diffusive permeabilities and effective diffusion coefficients in biofilms. Biotechnol. Bioeng. 59:261–272 [DOI] [PubMed] [Google Scholar]
  • 35.Daddi Oubekka S, Briandet R, Fontaine-Aupart MP, Steenkeste K. 2012. Correlative time-resolved fluorescence microscopy to assess antibiotic diffusion-reaction in biofilms. Antimicrob. Agents Chemother. 56:3349–3358 [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 36.Jefferson KK, Goldmann DA, Pier GB. 2005. Use of confocal microscopy to analyze the rate of vancomycin penetration through Staphylococcus aureus biofilms. Antimicrob. Agents Chemother. 49:2467–2473 [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 37.Pinho MG, Errington J. 2003. Dispersed mode of Staphylococcus aureus cell wall synthesis in the absence of the division machinery. Mol. Microbiol. 50:871–881 [DOI] [PubMed] [Google Scholar]
  • 38.Lunde CS, Rexer CH, Hartouni SR, Axt S, Benton BM. 2010. Fluorescence microscopy demonstrates enhanced targeting of telavancin to the division septum of Staphylococcus aureus. Antimicrob. Agents Chemother. 54:2198–2200 [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 39.Brown MR, Allison DG, Gilbert P. 1988. Resistance of bacterial biofilms to antibiotics: a growth-rate related effect? J. Antimicrob. Chemother. 22:777–780 [DOI] [PubMed] [Google Scholar]
  • 40.Duguid IG, Evans E, Brown MR, Gilbert P. 1992. Effect of biofilm culture upon the susceptibility of Staphylococcus epidermidis to tobramycin. J. Antimicrob. Chemother. 30:803–810 [DOI] [PubMed] [Google Scholar]
  • 41.Duguid IG, Evans E, Brown MR, Gilbert P. 1992. Growth-rate-independent killing by ciprofloxacin of biofilm-derived Staphylococcus epidermidis; evidence for cell-cycle dependency. J. Antimicrob. Chemother. 30:791–802 [DOI] [PubMed] [Google Scholar]
  • 42.Schwartz C, Henrickson KJ, Roghmann K, Powell K. 1990. Prevention of bacteremia attributed to luminal colonization of tunneled central venous catheters with vancomycin-susceptible organisms. J. Clin. Oncol. 8:1591–1597 [DOI] [PubMed] [Google Scholar]
  • 43.Carratala J, Niubo J, Fernandez-Sevilla A, Juve E, Castellsague X, Berlanga J, Linares J, Gudiol F. 1999. Randomized, double-blind trial of an antibiotic-lock technique for prevention of gram-positive central venous catheter-related infection in neutropenic patients with cancer. Antimicrob. Agents Chemother. 43:2200–2204 [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 44.Henrickson KJ, Axtell RA, Hoover SM, Kuhn SM, Pritchett J, Kehl SC, Klein JP. 2000. Prevention of central venous catheter-related infections and thrombotic events in immunocompromised children by the use of vancomycin/ciprofloxacin/heparin flush solution: a randomized, multicenter, double-blind trial. J. Clin. Oncol. 18:1269–1278 [DOI] [PubMed] [Google Scholar]

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