Lysostaphin Disrupts Staphylococcus aureus and Staphylococcus epidermidis Biofilms on Artificial Surfaces

Staphylococci often form biofilms, sessile communities of microcoloniesencased in an extracellular matrix that adhere to biomedicalimplants or damaged tissue. Infections associated with biofilmsare difficult to treat, and it is estimated that sessile bacteriain biofilms are 1,000 to 1,500 times more resistant to antibioticsthan their planktonic counterparts. This antibiotic resistanceof biofilms often leads to the failure of conventional antibiotictherapy and necessitates the removal of infected devices. Lysostaphinis a glycylglycine endopeptidase which specifically cleavesthe pentaglycine cross bridges found in the staphylococcal peptidoglycan.Lysostaphin kills Staphylococcus aureus within minutes (MICat which 90% of the strains are inhibited [MIC₉₀], 0.001 to0.064 µg/ml) and is also effective against Staphylococcusepidermidis at higher concentrations (MIC₉₀, 12.5 to 64 µg/ml).The activity of lysostaphin against staphylococci present inbiofilms compared to those of other antibiotics was, however,never explored. Surprisingly, lysostaphin not only killed S.aureus in biofilms but also disrupted the extracellular matrixof S. aureus biofilms in vitro on plastic and glass surfacesat concentrations as low as 1 µg/ml. Scanning electronmicroscopy confirmed that lysostaphin eradicated both the sessilecells and the extracellular matrix of the biofilm. This disruptionof S. aureus biofilms was specific for lysostaphin-sensitiveS. aureus, as biofilms of lysostaphin-resistant S. aureus werenot affected. High concentrations of oxacillin (400 µg/ml),vancomycin (800 µg/ml), and clindamycin (800 µg/ml)had no effect on the established S. aureus biofilms in thissystem, even after 24 h. Higher concentrations of lysostaphinalso disrupted S. epidermidis biofilms.

INTRODUCTION

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Introduction

Staphylococcal infections of both Staphylococcus aureus andStaphylococcus epidermidis continue to be a major problem inhospital settings, especially among immunocompromised and immunosuppressedpatients, particularly those with indwelling devices (29). Inthe Centers for Disease Control and Prevention's national emerginginfectious disease plan entitled Preventing Emerging InfectiousDiseases: a Strategy for the 21st Century (4), one of the specifiedgoals is to determine how to control biofilms on medical devicesand tubing in medical settings. In 1998, an estimated 200 millioncatheters, including urinary catheters and central venous catheters,were implanted in North American patients alone (14). Staphylococcicause a large percentage of catheter infections, and like manyother pathogens, rather than living as free planktonic cellswithin the host, they tend to form a multilayered communityof sessile bacteria cells known as a biofilm on medical implantsor damaged tissue (9, 10, 15, 17). Biofilm infections are difficultto treat due to their inherent antibiotic resistance (9, 15,17).

Once a staphylococcal biofilm has formed on an implanted medicaldevice or damaged tissue, it is difficult to disrupt. A biofilm-infectedimplant often must be removed and replaced, placing the patientat increased risk for complications due to these additionalprocedures (5, 32). Current antimicrobial therapies for biofilmshave largely proven unsuccessful (15), and the exact explanationfor these treatment failures is still unclear (9). Additionalstrategies for the clearance of biofilm-associated infectionsare needed.

Lysostaphin is an antibacterial enzyme which is specificallycapable of cleaving the cross-linking pentaglycine bridges inthe cell walls of staphylococci (38). S. aureus cell walls containhigh proportions of pentaglycine, making lysostaphin a highlyeffective agent against both actively growing and quiescentbacteria. Lysostaphin has also been shown to be effective againstS. epidermidis, albeit at higher concentrations of the enzyme(25, 42). Lysostaphin has gained renewed interest as an antistaphylococcaltherapeutic agent (12, 24, 30) because of the growing emergenceof antibiotic-resistant S. aureus. Antibiotic-resistant S. aureusorganisms (including both methicillin-resistant S. aureus [MRSA]and intermediately vancomycin-susceptible S. aureus) are susceptibleto lysostaphin action (21). Despite the effectiveness of lysostaphinagainst S. aureus both in vitro and in various animal models,it has yet to be determined whether lysostaphin would proveany more effective than any other antibiotic against staphylococciin biofilms. In this paper, we demonstrate the efficacy of lysostaphinagainst S. aureus and S. epidermidis biofilms in vitro by useof a modification of previous biofilm assays (6, 13, 19).

MATERIALS AND METHODS

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Materials and Methods

Bacterial strains. Six S. aureus and three S. epidermidis strains were used inthese studies and are listed in Table 1. Stocks were maintainedat -70°C in tryptic soy broth (TSB) before their use inthe biofilm assays. Strain ATCC 35556 (SA113) was selected becauseit is considered a benchmark strain among the biofilm-producingS. aureus species (11, 19), and S. epidermidis strain ATCC 35984was used for the same reason (34). Pseudomonas aeruginosa strainATCC 15692 was also examined in some experiments.

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TABLE 1. Strains of staphylococci used in these studies

Antimicrobial agents. Recombinant homogenous lysostaphin was purchased from Nutrition21, Inc. (Purchase, N.Y.), or produced by Biosynexus Incorporated.Oxacillin, vancomycin, and clindamycin were purchased from SigmaChemical Co. (St. Louis, Mo.).

Biofilm plate assays with lysostaphin treatment. The biofilm plate assay used was a modification of several previousbiofilm plate assays (6, 13, 19). Briefly, S. aureus from ablood agar plate (Remel, Lenexa, Kans.) was grown in 5 ml ofTSB (BBL, Sparks, Md.) supplemented with 0.25% D-(+)-glucose(Sigma). The overnight bacterial culture was diluted 1:50 inTSB plus 0.25% glucose to a final volume of 200 µl ineach well of a 96-well tissue culture plate (Costar Inc., Corning,N.Y.), a final volume of 500 µl in each chamber of a Lab-TekII chamber slide (Nalge-Nunc International, Naperville, Ill.),or a final volume of 1 ml in polycarbonate transwell tissueculture inserts (10-mm-diameter tissue culture inserts, 0.02-µm-pore-sizeAnapore membrane; Nalge-Nunc International) placed in a 24-welltissue culture plate (Nunclon surface multidish with 24 wells;Nalge-Nunc International).

The various plates were incubated at 37°C with shaking at100 rpm for 24 h; in some experiments, plates were removed fromthe shaker and supplemented with either 50 µl (96-wellplates) or 200 µl (tissue culture inserts) of additionalfresh TSB plus 0.25% glucose to compensate for medium evaporation.These plates were placed in the stationary incubator at 37°C,and growth was allowed to continue for an additional 24 h.

After 24 to 48 h of growth, depending on the experiment, thevarious wells were washed twice with 200 µl of phosphate-bufferedsaline (PBS) per well (96-well plates), 500 µl of PBSper chamber (chamber slides), or 1 ml of double-distilled water(ddH₂O) per insert (transwells) to remove nonadherent bacteria.The biofilm-containing wells were treated either with variousconcentrations of lysostaphin, namely, 0.8 to 200 µg/ml(96-well plates), 25 to 200 µg/ml (chamber slides), or50 to 100 µg/ml (tissue culture inserts), diluted in PBSor TSB plus 0.25% glucose, or with PBS or TSB plus 0.25% glucosealone (control wells) for up to 24 h with gentle shaking atambient temperature. The treatment buffer was gently aspiratedat the end of treatment, and the wells were washed once withvolumes of PBS equivalent to the treatment volume. The assayplates were inverted and allowed to dry at 37°C for approximately2 h. The wells of the 96-well plate and the tissue culture insertswere stained with 200 µl of enhanced Gram safranin (Remel)for 1 min. The stain was removed, and the wells were gentlywashed twice with 200 µl of PBS (microtiter plates) or500 µl of ddH₂O (transwells). The assay plates were observedfor the degree and intensity of staining in the wells. Followingincubations and washing, partitions on chamber slides were removed,and the slides were heat fixed and Gram stained (Remel).

A biofilm of P. aeruginosa was formed by a method similar tothat used for staphylococci. A plastic chamber slide containingTSB plus 0.25% glucose was inoculated with an overnight cultureof P. aeruginosa, and a biofilm was allowed to form for 48 h.The established P. aeruginosa biofilm was treated with 200 µgof lysostaphin/ml for 5 h in PBS. Following treatment, the P.aeruginosa biofilm was washed with PBS and Gram stained.

To prepare samples for scanning electron microscopy (SEM), theprocedures of the biofilm plate assay in transwell inserts (describedabove) were followed. After the inserts were air dried, theywere fixed in a 4% glutaraldehyde solution (16, 39) in ddH₂O(pH 7.0). SEM was performed at the Carnegie Institute of Washington(Department of Embryology, Baltimore, Md.).

The SEM samples were stored at 4°C in TSB for several daysprior to being processed. The tissue culture inserts were dehydratedin the following ethanol series: 35% ethanol (three times for10 min), 50% ethanol (10 min), 75% ethanol (10 min), 95% ethanol(10 min), and 100% ethanol (three times for 10 min). The sampleswere placed in hexamethyldisilazane twice for 30 min each timeand then were dried and sputter coated.

Comparison of lysostaphin to antibiotics in a 96-well plate assay. To directly compare the kinetics of lysostaphin's effect onS. aureus biofilms to those of other antibiotics, biofilms wereprepared in 96-well plate assays as described above. The plateswere washed twice with PBS, and serial dilutions of lysostaphinor antibiotics, as noted in the legend to Fig. 5 and Results,were added to each well in either PBS or TSB plus 0.25% glucosein a total volume of 200 µl per well. The wells were agitatedat room temperature for 24 h. The absorbance at 650 nm of bacteriain biofilm was measured every 20 min for the first 3 h aftertreatment and then again at 24 h using a microtiter plate reader.After treatment was complete, the wells were washed with PBSand then dried. Wells were stained with safranin and then washedtwice with PBS to remove excess stain.

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FIG. 5. Oxacillin or vancomycin had no visible effect on S. aureus biofilms in PBS or bacterial media after incubation for 24 h. Polystyrene 96-well tissue culture wells were inoculated with, per ml, $~$ 10⁸ CFU of S. aureus ATCC 35556 (MIC of lysostaphin, 0.004 µg/ml; MIC of oxacillin, 0.125 µg/ml; and MIC of vancomycin, 1 µg/ml). Following 24 h of biofilm formation, wells were washed twice with PBS and then treated for 24 h in either PBS (A) (this panel shows one of the plates from the experiment reported in Fig. 4) or TSB plus 0.25% glucose bacterial medium (B) with either no added antibiotic (first column of wells in each plate) or serial twofold dilutions of lysostaphin (first two rows; 0.8 to 200 µg/ml), oxacillin (third and fourth rows; 1.6 µg/ml to 400 µg/ml), or vancomycin (bottom two rows; 3.2 to 800 µg/ml) as indicated on the figure. Following treatment, the wells were washed with PBS and then stained with safranin. The darkly staining wells indicate the presence of biofilm following treatment. Lysostaphin in PBS cleared the biofilm at 0.8 µg/ml (A), while lysostaphin in TSB plus 0.25% glucose cleared the biofilm at 12.5 µg/ml (B).

RESULTS

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Results

Lysostaphin specifically disrupts S. aureus biofilms on abiotic surfaces. The biofilm-forming capacities of the various S. aureus strainswere observed by cultivating the biofilms on polycarbonate,polystyrene, or glass surfaces. The six S. aureus strains (Table1) were inoculated in the wells of tissue culture-treated microtiterplates, polycarbonate transwells, or chambers of chamber slidesand allowed to form biofilms over a 24- to 48-h period. Theresulting S. aureus biofilms in tissue culture wells (Fig. 1and 2) and inserts (data not shown) were stained with safranin,while the chamber slides (data not shown and see Fig. 6) wereGram stained to examine the biofilms on the surface of the slide.As previously reported (13, 19), the dltA-negative mutant ofstrain ATCC 35556 did not form a biofilm in our system (datanot shown); however, all other strains of S. aureus examinedformed biofilms on the various surfaces.

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FIG. 1. Lysostaphin-disrupted biofilms of MRSA strains Col and MBT 5040. Ninety-six-well polystyrene tissue culture wells (four for each sample) were inoculated with $~$ 10⁸ CFU of S. aureus strain Col or MBT 5040 (as indicated). After we allowed 48 h for biofilm formation, the wells were washed twice and then incubated with (+) or without (-) 50 µg of lysostaphin/ml in PBS for 3 h. Following incubation, the wells were washed again and stained with safranin to visualize biofilms.

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FIG. 2. Lysostaphin did not disrupt biofilms formed by lysostaphin-resistant S. aureus variants. Tissue culture wells (three for each sample) of a microtiter plate were inoculated with $~$ 10⁸ CFU of S. aureus strain SA113, SA5 LysoR, MBT 5040, or MBT 5040 LysoR (as indicated). After we allowed 48 h for biofilm formation, the wells were washed twice and then incubated with (+) or without (-) 50 µg of lysostaphin/ml in PBS for 3 h. Following incubation, the wells were washed again and stained with safranin to visualize biofilms.

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FIG. 6. Lysostaphin-disrupted S. epidermidis biofilms. Glass chamber slide wells were inoculated with $~$ 5 x 10⁷ CFU of either S. aureus strain SA113 as a control (A), S. epidermidis strain Hay (B), S. epidermidis strain ATCC 35984 (C), or S. epidermidis strain SE1175 (D) per ml. Biofilms were allowed to form for 24 h, and then wells were washed twice with PBS. Established biofilms were treated with PBS alone or with lysostaphin in PBS (200 µg/ml) for 3 h. Following treatment, the wells were washed with PBS followed by ddH₂O and then Gram stained. The biofilms in the PBS-treated row (-) stained gram positive, while no gram-positive cells were seen in lysostaphin-treated (+) S. epidermidis wells. Only the residual extracellular glycocalyx in the corners of the wells stained gram negative in the lysostaphin-treated wells. The two enlarged sections reveal the multilayered biofilm of S. epidermidis strain ATCC 35984 (top) and the residual glycocalyx of the same strain with no intact staphylococci following lysostaphin treatment (bottom).

The capacity of lysostaphin to disrupt the biofilms of the variousS. aureus strains was determined by visually comparing the differencesof safranin stain or Gram stain intensities between lysostaphin-treatedand untreated biofilms. Figure 1 shows 48-h biofilms of MRSAstrains Col and MBT 5040 treated for 3 h with lysostaphin (50µg/ml) or PBS. The darkly stained, PBS-treated Col andMBT 5040 biofilms contrasted with the lysostaphin-treated wellsthat had light or no staining, which revealed that lysostaphinsuccessfully disrupted the biofilms of both MRSA strains. Uponmicroscopic observation of chamber slides of similar experiments,the residual staining in the lysostaphin-treated wells was determinedto be debris and contained no intact staphylococci (data notshown).

When biofilms were treated with 25 µg of lysostaphin/ml,the biofilms of lysostaphin-sensitive S. aureus strains MBT5040 and SA113 were disrupted; however, when biofilms of invitro-isolated, lysostaphin-resistant variants of two S. aureusstrains, MBT 5040 LysoR and SA5 LysoR, were treated with thesame concentration of lysostaphin, the biofilms were not affectedby the lysostaphin treatment (Fig. 2). Microscopic examinationof S. aureus biofilms on chamber slides treated with 25 µgof lysostaphin/ml revealed that there were no remaining intactbacteria associated with the slide, and in most cases, the biofilmglycocalyx was not evident on the treated slides (data not shown).

The capacity of lysostaphin to disrupt the biofilms of anotherprolific biofilm producer, P. aeruginosa, was also evaluated.P. aeruginosa biofilms grown in plastic chamber slides weretreated with lysostaphin at concentrations as high as 200 µg/mlfor 5 h. This treatment had no visible effect on the biofilms;i.e., the staining intensity was the same with or without lysostaphintreatment, and microscopic examination of the P. aeruginosabiofilm revealed no change in the biofilm following lysostaphintreatment (data not shown).

SEM reveals that lysostaphin eradicates S. aureus biofilms. To further assess the extent of lysostaphin's effects on theS. aureus biofilms, biofilm assays were conducted in polycarbonatetranswells to allow SEM observation. Transwells were found tocultivate confluent biofilms and were the best substrate onwhich to observe these biofilms by SEM, as they were small andflat enough to view under an SEM without damaging the biofilm.SEM observation revealed that S. aureus biofilms grew as prolificallyon a polycarbonate surface as on a polystyrene surface (Fig.3A and B) and that 100 µg of lysostaphin/ml eradicatedthe S. aureus biofilm (Fig. 3C and D), releasing both sessilecells and the extracellular matrix from the polycarbonate transwells.Similar observations were made for six separate wells of eachsample in two separate experiments. In similar polycarbonatetranswell experiments observed by SEM, the biofilms producedby S. aureus strains MBT 5040, Col, and ATCC 35556 were alsofound to be eradicated; i.e., no sessile cells or glycocalyxremained after treatment with 100 µg of lysostaphin/mlin all wells examined (data not shown).

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FIG. 3. SEM revealed that lysostaphin eradicates S. aureus biofilms, removing both the sessile cells and the extracellular matrix. Polycarbonate transwells were inoculated with 5 x 10⁸ CFU of S. aureus ATCC 49521 per ml. Following 48 h of biofilm formation, transwells were washed and then treated with either PBS (A and B) or 100 µg of lysostaphin/ml in PBS (C and D) for 3 h. The treated wells were washed again and then fixed with gluteraldehyde prior to SEM. The results shown are representative of the entire well. Magnifications, x1,800 (A and C) and x5,940 (B and D).

Lysostaphin begins to disrupt S. aureus biofilms immediately and does so more effectively than other antibiotics. Oxacillin and vancomycin have often been used in antibioticsusceptibility studies of S. aureus biofilms (1, 18, 20, 31,41). These antibiotics were compared to lysostaphin to determinewhether lysostaphin was more effective in disrupting S. aureusstrain ATCC 35556 biofilms than the conventional antibiotics.Twenty-four-hour biofilms in polystyrene 96-well tissue cultureplates were treated with serial dilutions of lysostaphin, oxacillin,and vancomycin (Fig. 4 and 5).

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FIG. 4. Lysostaphin caused an immediate and continuous drop in the absorbance of S. aureus biofilms, which continued over time. Polystyrene 96-well tissue culture wells were inoculated, per ml, with $~$ 10⁸ CFU of S. aureus ATCC 35556 (MIC of lysostaphin, 0.004 µg/ml; MIC of oxacillin, 0.125 µg/ml; and MIC of vancomycin, 1 µg/ml). Following 24 h of biofilm formation, wells were washed twice with PBS and then treated with either lysostaphin (6.25 µg/ml), oxacillin (400 µg/ml), or vancomycin (800 µg/ml) in PBS. The absorbance at 650 nm following treatment was monitored by a microtiter plate reader every 20 min for 3 h and then again at 24 h. The data are the means of results for six samples from three separate experiments ± the standard deviations.

In order to examine the kinetic effects of lysostaphin, oxacillin,and vancomycin on biofilms, the absorbances at 650 nm of establishedbiofilms in a 96-well tissue culture plate were measured overtime (0 to 3 h and 24 h). Tissue culture wells containing biofilmsof S. aureus SA113 were incubated with serial dilutions of lysostaphin(0.8 to 200 µg/ml), oxacillin (1.6 to 400 µg/ml),or vancomycin (3.2 to 800 µg/ml) for 24 h. The absorbanceof the lysostaphin-treated biofilms dropped from approximately0.35 at time zero to 0.125 after 3 h of treatment and droppedto near baseline (0.04) by 24 h when the biofilms were treatedwith a dose of lysostaphin of 6.25 µg/ml in PBS (Fig.4). The absorbances of the biofilms treated with oxacillin orvancomycin for 24 h showed minimal change, remaining around0.325, despite the fact that the biofilms were treated withas much as 400 µg of oxacillin/ml or 800 µg of vancomycin/mlin PBS (Fig. 4). Since antimicrobials like oxacillin and vancomycinare known to be more effective against actively metabolizingbacteria, a similar experiment was conducted but with incubationof the biofilms with the three antimicrobials in bacterial medium(TSB). Very similar results were found when the assay was conductedwith TSB rather than PBS. Lysostaphin reduced the absorbanceof biofilms to near background by 24 h, while oxacillin andvancomycin had little or no effect even after 24 h of incubation(data not shown).

The capacity of the three agents to disrupt S. aureus biofilmsin polystyrene wells could be visualized by comparing the stainingintensities of treated wells with those of control (buffer-treated)wells. Biofilms from the above-described kinetics experimentthat were treated for 24 h stained darkly on the bottom of thewells (Fig. 5), while wells cleared of biofilms did not stainwith safranin. Lysostaphin concentrations as low as 0.8 µg/mlin PBS (Fig. 5A) and 12.5 µg/ml in TSB plus 0.25% glucose(Fig. 5B) appeared to clear biofilms from the transwells, while400 µg of oxacillin/ml or 800 µg of vancomycin/mlin PBS or TSB had no obvious effect on established biofilmseven after 24 h of treatment (Fig. 5). The antibiotic clindamycinhas also been shown to be somewhat effective for the disruptionof some biofilms (27, 35). When this antibiotic was tested ina similar 96-well plate assay, concentrations of clindamycinas high as 800 µg/ml had little or no effect on the S.aureus biofilms even after 24 h of treatment (data not shown).

Lysostaphin disrupts S. epidermidis biofilms. While lysostaphin demonstrated activity against S. aureus biofilms,it was of interest to explore whether biofilms of S. epidermidis,known to be less sensitive to lysostaphin (25, 42), were alsosensitive to the biofilm-disrupting effect of lysostaphin. ThreeS. epidermidis strains with various capacities for glycocalyx(slime) production were examined, including S. epidermidis strainHay (a low slime producer), S. epidermidis strain SE1175 (amoderate slime producer), and S. epidermidis ATCC 35984 (34)(a high slime producer). All three of these S. epidermidis strainsproduced biofilms on a glass chamber slide (Fig. 6), with ATCC35984 producing the thickest and most darkly staining biofilm,as expected. Incubation of these S. epidermidis biofilms with200 µg of lysostaphin/ml for 3 h disrupted the biofilmsof all three strains of S. epidermidis (Fig. 6). S. aureus strainSA113 was included in this experiment as a control. Microscopicexamination of the disrupted biofilms revealed that there wereno intact bacteria left on the artificial surface (Fig. 6, inset).The stained material visible in lysostaphin-treated wells wasextracellular glycocalyx, which stained pink by safranin andcontained no intact gram-positive S. epidermidis cells, onlycellular debris.

DISCUSSION

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Discussion

Lysostaphin's activity against planktonic staphylococci hasbeen well documented (3, 7, 8, 25, 26, 42; J. F. Kokai-Kun,T. Chanturiya, and J. J. Mond, Abstr. 102nd Gen. Meet. Am. Soc.Microbiol. abstr. A-62, p. 12, 2002); it was not known, however,whether lysostaphin would be any more effective in disruptingstaphylococcal biofilms than conventional antibiotics. We havenow demonstrated that, in vitro, lysostaphin disrupted S. aureusbiofilms on polystyrene, polycarbonate, and glass surfaces andalso disrupted S. epidermidis biofilms on glass. When S. aureusbiofilms were visualized by SEM, lysostaphin eradicated boththe sessile S. aureus cells of the biofilm and the extracellularmatrices (Fig. 3).

The exact mechanism of lysostaphin's action against S. aureusbiofilms remains unclear. It has been shown that, even if anantimicrobial agent can kill some of the bacteria embedded inthe biofilm, the biofilm is not disrupted unless the extracellularmatrix is also destroyed (1, 18, 20, 36, 41). Lysostaphin may,therefore, be able to target and disrupt both the individualsessile cells of the S. aureus biofilms and the extracellularmatrix. While there is no evidence that lysostaphin acts directlyon the extracellular matrices of staphylococcal biofilms, itis possible that lysostaphin may target some components of thesematrices. Since lysostaphin's primary enzymatic substrate isthe pentaglycine cross bridges of the peptidoglycan of staphylococci(42), the extracellular staphylococcal biofilm matrix may containcomponents of cell walls, including polyglycines, that mightbe affected by lysostaphin, thus disrupting the extracellularbiofilm matrix. Indeed, other macromolecules have been foundin the extracellular matrix of biofilms; extracellular DNA isrequired for P. aeruginosa biofilm formation (40). A more likelyexplanation, however, is that the disruption of staphylococcalbiofilms by lysostaphin occurs through the rapid lysis of thesessile staphylococci, which may be sufficient to destabilizethe entire biofilm matrix in such a manner as to allow detachmentfrom artificial surfaces. This explanation was supported bythe finding that lysostaphin's capacity to disrupt S. aureusbiofilms appeared to be specific for lysostaphin-sensitive S.aureus (Fig. 2), since biofilms formed by lysostaphin-resistantS. aureus were not disrupted by lysostaphin. The finding thatbiofilms from three strains of S. epidermidis, each with a differentcapacity to form slime, could also be disrupted by lysostaphinfurther supported the theory that lysostaphin attacks the actualcells of the biofilm, since the glycocalyxes of biofilms ofvarious S. epidermidis strains are thought to have differentchemical compositions (17).

The resistance to antibiotics of bacteria growing in biofilmsremains an incompletely understood process and is an area ofactive research. The antibiotic resistance of bacteria in establishedbiofilms may be due to a number of factors, including the multilayerstructure of biofilms (33) and/or the unique genetic characteristicsof bacteria in biofilms compared to those of planktonic cells(9). Antibiotics used to treat biofilm infections in hospitalsalso have to overcome the potential emergence of antibioticresistance as well as the increased risk to the patient of developingallergies to the antibiotic therapies or other problems associatedwith antibiotics, such as diarrhea, as higher doses and longercourses of antibiotic treatments are used to try to eradicatebiofilms (37). Antibiotic efficacy also decreases as treatmenttime increases (1). Thus, the recommended course of action fortreatment of staphylococcal biofilm infections is to removethe infected device, treat the patient with rigorous antibiotictherapy, and reinsert a new device (2, 5). Despite this treatmentregimen, the recurrence of infection is high (23, 41). Biofilmbacteria can usually survive antibiotics at concentrations 1,000to 1,500 times higher than antibiotic concentrations used totreat bactericidal planktonic bacteria (9).

Using lysostaphin to treat staphylococcal-biofilm-associatedinfections may prove to be preferable to using antibiotics suchas oxacillin, vancomycin, and clindamycin to treat these infections.It may be possible to administer lysostaphin at relatively lowdoses and disrupt a staphylococcal biofilm (Fig. 5), obviatingthe need for surgical removal of the infected device. Lysostaphinalso has the advantage of a relatively short response time interms of disrupting the biofilm; within 20 min of applicationthere is measurable disruption of the biofilm (Fig. 4), anddepending on the concentration of lysostaphin used, this disruptionmay be completed within hours. This time to biofilm disruptionis relatively short compared to those with oxacillin and vancomycin,both of which have been reported to require approximately 24h to disrupt biofilms (1, 28), an exposure time which had verylittle effect on S. aureus biofilms in this study (Fig. 4 and5). The specific activity of lysostaphin for staphylococci hasthe added benefit of not disrupting normal flora, thus minimizingthe occurrence of antibiotic-associated diarrheas or selectionof antibiotic resistance in other opportunistic pathogens. Whetherlysostaphin proves to be an effective therapy for staphylococcal-biofilminfections, however, will depend on results of preclinical testingthat is under way (J. F. Kokai-Kun, T. Chanturiya, A. Shah,S. M. Walsh, and J. J. Mond, Abstr. 103rd Gen. Meet. Am. Soc.Microbiol., abstr. A-026, p. 6, 2003) and clinical testing whichis planned for the future.

Lysostaphin is not the only enzyme to have been examined forits capacity to disrupt biofilms. Various combinations of enzymeshave been studied for the removal and disinfection of bacterialbiofilms in various systems (22). This process, however, requiresa minimum of two enzymes, one enzyme for removal of the adherentbacteria of the biofilms and a second enzyme with bactericidalactivity. Lysostaphin bridges this two-step process by bothdisrupting the S. aureus and S. epidermidis biofilms and killingthe released bacteria.

It has been shown that coating medical implants with antimicrobialsmay effectively prevent the initial adherence of staphylococcalbiofilms to the implants (29, 32, 37). Coating biomedical materialswith lysostaphin may also prove successful in preventing earlyadherence of staphylococci to the implants, thus averting biofilmformation. Work is under way to determine whether lysostaphin-coatedmedical implants, such as catheters, will prevent staphylococcaladherence and the formation of biofilms (A. Shah and S. M. Walsh,unpublished data).

Bacterial biofilms in medical systems are thought to be virtuallyidentical to biofilms in any other aquatic system (9). Our datademonstrate that lysostaphin disrupted both S. aureus and S.epidermidis biofilms in vitro and suggest that lysostaphin maybe effective against staphylococcal-biofilm infections in vivo.We are currently examining the effectiveness of lysostaphinfor the in vivo clearance of staphylococcal-biofilm-associatedcatheter infections by using a catheterized mouse model (Kokai-Kunet al., Abstr. 103rd Gen. Meet. Am. Soc. Microbiol.).

ACKNOWLEDGMENTS

We thank Mike Sepanski for conducting SEM and Andreas Peschelfor providing various strains of S. aureus.

J. A. Wu and C. Kusuma contributed equally to this work.

All of the authors of the manuscript are, or formerly were,employees of Biosynexus Incorporated. Biosynexus Incorporatedis currently developing commercial applications of lysostaphin.

FOOTNOTES

* Corresponding author. Mailing address: Biosynexus Incorporated, 9119 Gaither Rd., Gaithersburg, MD 20877. Phone: (301) 987-1172. Fax: (301) 990-4990. E-mail: johnkun{at}biosynexus.com.

Present address: Department of Molecular and Cell Biology, UniformedServices University of the Health Sciences, Bethesda, MD 20814.

REFERENCES

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