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Antimicrobial Agents and Chemotherapy, March 2009, p. 918-925, Vol. 53, No. 3
0066-4804/09/$08.00+0     doi:10.1128/AAC.00766-08
Copyright © 2009, American Society for Microbiology. All Rights Reserved.

Oritavancin Kills Stationary-Phase and Biofilm Staphylococcus aureus Cells In Vitro {triangledown}

Adam Belley,1 Eve Neesham-Grenon,1 Geoffrey McKay,1 Francis F. Arhin,1 Robert Harris,2,3 Terry Beveridge,2,4 Thomas R. Parr Jr.,1 and Gregory Moeck1*

Targanta Therapeutics Incorporated, 7170 Frederick Banting, St. Laurent, Quebec H4S 2A1, Canada,1 MicroTEM Inc., P.O. Box 1107, 101 Chalmers St., Elora, Ontario N0B 1S0, Canada,2 Guelph Regional Integrated Imaging Facility, New Science Complex, 488 Gordon St., University of Guelph, Guelph, Ontario N1G 2W1, Canada,3 Department of Molecular and Cellular Biology, New Science Complex, 488 Gordon St., University of Guelph, Guelph, Ontario N1G 2W1, Canada4

Received 12 June 2008/ Returned for modification 19 October 2008/ Accepted 11 December 2008


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ABSTRACT
 
Slow-growing bacteria and biofilms are notoriously tolerant to antibiotics. Oritavancin is a lipoglycopeptide with multiple mechanisms of action that contribute to its bactericidal action against exponentially growing gram-positive pathogens, including the inhibition of cell wall synthesis and perturbation of membrane barrier function. We sought to determine whether oritavancin could eradicate cells known to be tolerant to many antimicrobial agents, that is, stationary-phase and biofilm cultures of Staphylococcus aureus in vitro. Oritavancin exhibited concentration-dependent bactericidal activity against stationary-phase inocula of methicillin-susceptible S. aureus (MSSA) ATCC 29213, methicillin-resistant S. aureus (MRSA) ATCC 33591, and vancomycin-resistant S. aureus (VRSA) VRS5 inoculated into nutrient-depleted cation-adjusted Mueller-Hinton broth. As has been described for exponential-phase cells, oritavancin induced membrane depolarization, increased membrane permeability, and caused ultrastructural defects including a loss of nascent septal cross walls in stationary-phase MSSA. Furthermore, oritavancin sterilized biofilms of MSSA, MRSA, and VRSA at minimal biofilm eradication concentrations (MBECs) of between 0.5 and 8 µg/ml. Importantly, MBECs for oritavancin were within 1 doubling dilution of their respective planktonic broth MICs, highlighting the potency of oritavancin against biofilms. These results demonstrate a significant activity of oritavancin against S. aureus in phases of growth that exhibit tolerance to other antimicrobial agents.


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INTRODUCTION
 
Infections in which bacteria are either slow growing, dormant, or in a biofilm pose a serious clinical challenge for therapy because cells in these states exhibit tolerance to the activity of antimicrobial agents (15). Osteomyelitis, infective endocarditis, chronic wounds, and infections related to indwelling devices are examples of infections that harbor tolerant cells (10, 16). Antimicrobial therapies for these infections are not optimal and thus require protracted treatment times. A model theory has been proposed to explain biofilm recalcitrance to chemotherapy (27): the diversity of the growth phases of the biofilm community (44) and the composition of the slime matrix act to limit the effectiveness of otherwise useful antimicrobial agents. It is believed that a population of slow-growing, stationary-phase, or "persister," cells within the biofilm can tolerate the killing action of antibacterial agents. This has been demonstrated with the fluoroquinolone antibiotic ofloxacin, for which a small population of cells within a biofilm was not killed by this agent (43). Furthermore, it is thought that these tolerant cells are protected from immune clearance in vivo by the biofilm slime matrix and ultimately give rise to relapse infections by reseeding the biofilm once drug levels drop below their antibacterial concentration (27).

Oritavancin is a semisynthetic lipoglycopeptide in clinical development for the treatment of serious gram-positive infections. It exerts activity against both susceptible and methicillin-resistant Staphylococcus aureus (MRSA) and vancomycin-resistant enterococci. The rapidity of its bactericidal activity against exponentially growing S. aureus cells (≥3-log reduction within 15 min to 2 h against methicillin-sensitive S. aureus [MSSA], MRSA, and vancomycin-resistant S. aureus [VRSA]) is one feature that distinguishes it from the prototypic glycopeptide vancomycin (31). Recent work demonstrated that oritavancin has multiple mechanisms of action that can contribute to the cell death of exponentially growing S. aureus cells, including the inhibition of cell wall synthesis by both substrate-dependent and -independent mechanisms (2, 4, 48), disruption of membrane potential and increasing membrane permeability (32), and inhibition of RNA synthesis (4). The ability of oritavancin but not vancomycin to interact with the cell membrane, leading to a loss of membrane integrity and collapse of transmembrane potential, correlates with the rapidity of oritavancin bactericidal activity (32). Mechanisms of action beyond substrate-dependent cell wall synthesis inhibition have not been described to date for vancomycin; consequently, vancomycin typically requires 24 h and actively dividing cells to exert bactericidal activity (7, 31). With this in mind, we sought to characterize oritavancin activity in vitro against S. aureus in slow-growing and biofilm states.

(Part of this work was presented at the 47th Interscience Conference on Antimicrobial Agents and Chemotherapy, Chicago, IL, 17 to 20 September 2007 [7].)


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MATERIALS AND METHODS
 
Bacterial strains. The strains used in this study were MSSA reference strain ATCC 29213 (susceptible to daptomycin, linezolid, rifampin, and vancomycin as determined by CLSI broth microdilution guidelines) (11), MRSA isolates ATCC 33591 and ATCC 43300 (both strains susceptible to daptomycin, linezolid, rifampin, and vancomycin), and VRSA isolate VRS5 (Network on Antimicrobial Resistance in Staphylococcus aureus) (resistant to vancomycin and susceptible to daptomycin, linezolid, and rifampin). MSSA strain ATCC 29213 and MRSA strains ATCC 33591 and ATCC 43300 were grown overnight to stationary phase in cation-adjusted Mueller-Hinton broth (CAMHB; Becton, Dickinson, and Company, Sparks, MD) at 37°C with rotation at 225 rpm. VRSA VRS5 was grown overnight to stationary phase at 37°C with rotation at 225 rpm in brain heart infusion broth (Becton, Dickinson, and Company) containing 4 µg/ml of vancomycin.

Nutrient-depleted CAMHB. Nutrient-depleted CAMHB (29) from each respective strain was prepared by three rounds of inoculation of CAMHB with exponential-phase bacteria, incubation overnight at 37°C with rotation (225 rpm), and centrifugation (8,000 x g for 30 min) to remove bacteria. After the final round of inoculation, growth, and centrifugation, the pH of the nutrient-depleted CAMHB was adjusted to pH 7.0, and the spent medium was filter sterilized using a 0.22-µm membrane (Corning Incorporated, Corning, NY).

Antibacterial agents and concentrations. Antibacterial agents were tested at pharmacologically achievable concentrations that have been determined from clinical studies: concentrations were chosen to approximate free peak (fCmax) and free trough levels in plasma following administration of standard doses in humans. For oritavancin, fCmax and free trough levels from a standard dose of 200 mg (47) as well as an additional concentration that approximates the fCmax following a single 800-mg dose (fCmax800) in humans were used (18). Oritavancin diphosphate powder (Targanta Therapeutics, Cambridge, MA) was dissolved in water containing 0.002% (vol/vol) polysorbate 80; polysorbate 80 was also maintained at 0.002% in assays to minimize oritavancin loss to the surface of vessels (3), except where indicated. Concentrations that approximate the fCmax and free trough levels in plasma when administered at standard dosages for the prototypical glycopeptide vancomycin, the oxazolidinone linezolid, and the lipopeptide daptomycin were determined from pharmacokinetic data and protein binding values reported in their respective package inserts (vancomycin, Vancocin; linezolid, Zyvox; daptomycin, Cubicin). The approximation of the rifampin fCmax was derived from data reported previously by Burman et al. (8).

Time-kill studies. Nutrient-depleted CAMHB containing diluted antimicrobial agents was inoculated with stationary-phase bacteria from cultures of S. aureus strains grown overnight at approximately 107 CFU/ml. Other experiments compared the killing of stationary- and exponential-phase S. aureus ATCC 29213 cells when inoculated into nutrient-depleted CAMHB containing the test agents. For assays involving daptomycin, nutrient-depleted CAMHB was supplemented with 50 µg/ml CaCl2 (11). All time-kill studies were performed using 96-well deep-well plates at 37°C with rotation (225 rpm) in a total volume of 750 µl. To prevent drug carryover during serial dilution plating, aliquots of the drug-challenged culture were added to an equal volume of activated charcoal suspension (25 mg/ml) (6). Bactericidal activities of the antimicrobial agents were defined as a reduction in viable cell counts of ≥3 log at 24 h relative to cell counts in the starting inoculum (35). Experiments were repeated at least three times and produced similar results; results from one experiment are presented.

Short-duration (2-h) time-kill studies were performed using membrane assay buffer (see below) to characterize the killing of S. aureus ATCC 29213 cells under conditions used in the membrane depolarization and permeability assays. Exponential- and stationary-phase S. aureus ATCC 29213 cells were diluted to an optical density at 600 nm [OD600] of 0.005 (approximately 106 CFU/ml) in membrane assay buffer (10 mM HEPES-Cl [pH 7.5], 50 µg/ml CaCl2) with and without 5 mM glucose, respectively (glucose was included or omitted to prevent the de-energization or energization of exponential- or stationary-phase cell membranes, respectively). Experiments were initiated by the addition of antimicrobial agents at the indicated concentrations, and bacteria were enumerated by serial dilution plating. Testing with oritavancin was done in the absence of 0.002% polysorbate 80 to reflect conditions used in membrane depolarization and permeability assays. Experiments were repeated three times and produced similar results; results from one assay are presented.

Measurements of membrane depolarization and permeability. Membrane depolarization was monitored using the fluorescent probe 3,3'-dipropylthiadicarbocyanine iodide [DiSC3(5)] (Invitrogen Corporation, Carlsbad, CA), which partitions into the plasma membrane in proportion to the membrane potential. Dissipation of the membrane potential releases the probe, leading to an increase in fluorescence. Previous studies with the glycopeptide telavancin (19) and the lipopeptide daptomycin (42) have used this probe to demonstrate the membrane-perturbing activity of these drugs against exponential-phase cells. S. aureus ATCC 29213 was chosen for testing in membrane studies. Bacteria were grown overnight in CAMHB and subcultured the following day in CAMHB to exponential phase (OD600 {approx} 0.25). Exponential- and stationary-phase cells were washed in membrane assay buffer with and without 5 mM glucose, respectively, and resuspended at an OD600 of 0.25. DiSC3(5) was added to a final concentration of 1.5 µM, and the solution was incubated in the dark at ambient temperature for 30 min to allow the loading of the fluorescent dye into cell membranes. After the loading period, cells were diluted 50-fold (OD600 of 0.005) in depolarization buffer with or without glucose for exponential- or stationary-phase cells, respectively. Assays were initiated by the addition of antimicrobial agents over a range of concentrations and were monitored in real time by fluorescence spectroscopy (excitation wavelength of 612 nm and emission wavelength of 665 nm) for a period of 30 min. Note that 0.002% polysorbate 80 was found to interfere with fluorescence in these assays and was therefore omitted from the assay. Experiments were repeated three times and produced similar results; results from one assay are presented.

Changes in bacterial membrane permeability were quantified using the fluorescent dye pair Syto 9 and propidium iodide: bacterial membrane damage (increased permeability) allows the otherwise membrane-impermeable dye propidium iodide to enter the cell and displace the permeative dye Syto 9, leading to a loss of fluorescence. Bacteria were prepared as described above for the membrane depolarization assay, but Syto 9 and propidium iodide (Invitrogen Corporation) were added at 5 µM and 30 µM, respectively (32). Fluorescence spectroscopy (excitation wavelength of 485 nm and emission wavelength of 535 nm) was monitored for 30 min following the addition of antimicrobial agents. As mentioned above, 0.002% polysorbate 80 was omitted from the assay to prevent interference with fluorescence determinations. Experiments were repeated three times and produced similar results; results from one assay are presented.

Determination of ultrastructural effects of oritavancin and vancomycin on stationary-phase cells by transmission electron microscopy. Stationary-phase MRSA ATCC 43300 cells (5 x 107 CFU/ml) were exposed to 1 µg/ml oritavancin (2x its broth microdilution MIC in the absence of 0.002% polysorbate 80) or 16 µg/ml vancomycin (16x its broth microdilution MIC) in nutrient-depleted CAMHB for 3 h. Bacteria were fixed in 2.5% glutaraldehyde to cross-link proteins and help preserve morphological structure. Prior to embedding, the samples were treated with fresh 2.5% (vol/vol) glutaraldehyde in HEPES buffer (pH 6.8) for 2 h. The samples were then postfixed in 2.0% (wt/vol) osmium tetroxide, followed by en bloc staining with 2.0% (wt/vol) uranyl acetate, as a heavy-metal stain, to add contrast to the cells. The cells were then dehydrated through a series of ethanol washes and then embedded in LR White resin. Once polymerized by curing, each culture sample was thin sectioned and stained by uranyl acetate and lead citrate to view the internal cellular constituents and the juxtaposition of cell envelope layers such as the plasma membrane and cell wall. Transmission electron microscopy was used to view the thin sections using a Philips CM10 microscope under standard operating conditions at 100 kV.

Determination of MBEC. In vitro biofilms were established using the MBEC Physiology & Genetics Assay plate (Innovotech; Edmonton, AB, Canada) according to the manufacturer's protocol (20). The MBEC system also allows the determination of the MIC of the test agent for planktonic cells shed from the biofilm as well as the minimum biofilm eradication concentration (MBEC), the concentration of antimicrobial agent required to sterilize the biofilm after 24 h of exposure. Briefly, 150 µl of bacterial inocula at 107 CFU/ml in tryptic soy broth was aliquoted into each well of an MBEC plate. Biofilms were established on the MBEC peg lid for 24 h in a rotary incubator at 37°C and 150 rpm. For experiments involving 72-h biofilms, MBEC peg lids were transferred each day into 96-well plates containing 150 µl/well of fresh tryptic soy broth and incubated another 24 h. MBEC peg lids with established biofilms were washed once in sterile saline (200 µl/well) and then placed onto plates containing antimicrobial agents diluted in CAMHB (200 µl/well). Antimicrobial agents were serially diluted in CAMHB in 96-well plates, and MBEC peg lids were exposed for 24 h or for the indicated times. Note that 0.002% polysorbate 80 was found to adversely affect biofilm cell numbers and was therefore omitted from MBEC determinations for oritavancin. Following antimicrobial challenge and determination of the planktonic MICs, MBEC peg lids were washed once in sterile saline and then placed into recovery plates containing CAMHB (200 µl/well). The recovery plates were sonicated for 5 min in an ultrasonic sonicating bath (VWR Aquasonic model 550D) at the maximum setting and then incubated for 24 h, and the MBECs were recorded. MBECs were determined from at least three independent experiments; results represent the ranges of MBECs obtained. To enumerate the biofilm CFU on individual control pegs, pegs were broken off the MBEC peg lid using sterile forceps placed into 1 ml of sterile saline, sonicated for 5 min, and vortexed for 1 min at the highest setting. Bacteria were then enumerated by serial dilution plating. CFU/peg counts were determined from at least three independent experiments; results presented are the averages ± standard deviations.


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RESULTS
 
Oritavancin retains activity against stationary-phase S. aureus cells. Time-kill studies were performed using nutrient-depleted CAMHB to determine whether growth phase affects the antibacterial action of oritavancin and comparator agents under conditions of slow growth. In the first experiment, the killing of stationary-phase MSSA ATCC 29213 (Fig. 1A) was compared to the killing of an exponential-phase inoculum (Fig. 1B). Nutrient-depleted CAMHB limited the growth of the bacterial inocula over a 24-h period (stationary-phase ATCC 29213 increased by approximately 0.6 logs, whereas exponential-phase cells increased by approximately 1.3 logs by 24 h) (Fig. 1A and B). Estimated fCmax levels of oritavancin, vancomycin, and daptomycin were bactericidal against exponential-phase MSSA ATCC 29213 in nutrient-depleted CAMHB at the 24-h time point (Fig. 1B). In contrast, only oritavancin exerted bactericidal activity against stationary-phase MSSA strain ATCC 29213 in nutrient-depleted CAMHB (Fig. 1A), with vancomycin being the most affected by growth phase (approximately 2.4-log-less killing activity against stationary-phase cells than against exponential-phase cells at the 24-h time point). The estimated free trough concentration derived from a 200-mg dose of oritavancin in humans (0.5 µg/ml) nearly achieved bactericidal levels (approximately 2.9 logs) (Fig. 1A) against stationary-phase MSSA at the 24-h time point, whereas free trough concentrations of vancomycin and daptomycin had negligible effects on cell numbers (data not shown).


Figure 1
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  		FIG. 1. Time-kill kinetics of stationary- and exponential-phase MSSA ATCC 29213 in nutrient-depleted CAMHB. Viability was enumerated at the indicated time points by serial dilution plating. Each point represents the mean of duplicate determinations. The limit of detection is indicated as a dashed line. (A) Stationary-phase inocula challenged with estimated free trough, fCmax, fCmax800 of oritavancin, and fCmax of comparators. (B) Exponential-phase inocula challenged with estimated free trough, fCmax, and fCmax800 of oritavancin and comparators. *, untreated control; Figure 1, 0.5 µg/ml oritavancin; {circ}, 4 µg/ml oritavancin; •, 16 µg/ml oritavancin; {blacksquare}, 16 µg/ml vancomycin; {square}, 4 µg/ml daptomycin; {blacktriangleup}, 8 µg/ml linezolid; {triangledown}, 2 µg/ml rifampin.

In a follow-up experiment, the killing of stationary-phase inocula of MRSA and VRSA in nutrient-depleted CAMHB were determined (Fig. 2). As was seen with MSSA, limited growth of MRSA ATCC 33591 (approximately a 0.6-log increase) (Fig. 2A) and VRSA VRS5 (approximately a 0.1-log increase) (Fig. 2B) occurred in nutrient-depleted CAMHB over 24 h. Oritavancin exhibited concentration-dependent bactericidal activity against the MRSA and VRSA strains: oritavancin at its fCmax and fCmax800 was bactericidal at the 24-h time point (Fig. 2), with the exception of the fCmax against VRS5 (Fig. 2B), which nearly achieved bactericidal killing (approximately a 2.2-log decrease). Vancomycin exhibited some killing activity against the stationary-phase inocula of MRSA (approximately a 1.4-log reduction) and no killing against VRSA in nutrient-depleted CAMHB after 24 h (Fig. 2). Daptomycin and rifampin exhibited bactericidal activity at their respective fCmax values against the stationary-phase inoculum of MRSA (Fig. 2A) but were not bactericidal against the VRSA strain (Fig. 2B). The bacteriostatic agent linezolid (33) had no effect on the cell numbers of both strains over the 24-h incubation period. The estimated free trough concentration of oritavancin (0.5 µg/ml) reduced cell counts by approximately 0.4 logs against the MRSA strain (Fig. 2A) and approximately 1.2 logs against the VRSA strain (Fig. 2B), whereas the estimated free trough concentrations of vancomycin and daptomycin had no effect on the cell counts of these isolates (data not shown).


Figure 2
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  		FIG. 2. Time-kill kinetics of stationary-phase MRSA ATCC 33591 and VRSA VRS5 by oritavancin and comparators in nutrient-depleted CAMHB. Viability was enumerated at the indicated time points by serial dilution plating. Each point represents the mean of duplicate determinations. The limit of detection is indicated as a dashed line. (A) MRSA ATCC 33591 challenged with estimated free trough, fCmax and fCmax800 of oritavancin, and fCmax of comparators. (B) VRSA VRS5 challenged with estimated free trough, fCmax and fCmax800 of oritavancin, and fCmax of comparators. *, growth control; Figure 2, 0.5 µg/ml oritavancin; {circ}, 4 µg/ml oritavancin; •, 16 µg/ml oritavancin; {blacksquare}, 16 µg/ml vancomycin; {square}, 4 µg/ml daptomycin; {blacktriangleup}, 8 µg/ml linezolid; {triangledown}, 2 µg/ml rifampin.

Oritavancin perturbs membrane integrity of stationary-phase S. aureus cells. Recent studies demonstrated that the rapid bactericidal activity of oritavancin against exponential-phase S. aureus cells is temporally correlated with membrane depolarization and increased membrane permeability (32). To determine whether oritavancin also affects the membrane energetics of stationary-phase cells, we explored the effects of oritavancin and comparator agents on membrane potential and permeability in stationary-phase MSSA ATCC 29213 using fluorescent probes. A dissipation of membrane potential, measured as increased fluorescence resulting from the release of DiSC3(5) from stationary- and exponential-phase cell membranes, occurred in a concentration-dependent manner in response to oritavancin (Fig. 3A); however, the rate of release of the dye from stationary-phase cell membranes was less than that from exponential-phase cells. The addition of vancomycin had no effect on membrane potential, as indicated by the unchanged fluorescence signals that were comparable to those of the untreated control cells (Fig. 3B). Under the conditions of this in vitro system, daptomycin slightly reduced the membrane potential of exponential-phase cells and did not effect changes on the stationary-phase inoculum within the time frame (30 min) and concentrations (4 µg/ml) used in the experiment (Fig. 3B).


Figure 3
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  		FIG. 3. Measurement of oritavancin effects on membrane depolarization, permeability, and killing of MSSA ATCC 29213. (A and B) Membrane depolarization was monitored by measuring DiSC3(5) fluorescence. (C and D) Permeabilization of the cell membranes was monitored by measuring Syto 9 and propidium iodide fluorescence. Note that in D, the curve for vancomycin versus stationary-phase cells overlaps the curve for daptomycin versus stationary-phase cells. RFU, relative fluorescence units. (E and F) Killing kinetics of stationary- and exponential-phase inocula in membrane assay buffer. Glucose was omitted from the membrane assay buffer for stationary-phase cells and included at 5 mM for exponential-phase cells. The limit of detection is indicated as a dashed line. For A, C, and E, symbols are as follows: {circ}, 4 µg/ml oritavancin versus exponential-phase cells; •, 4 µg/ml oritavancin versus stationary-phase cells; {square}, untreated exponential-phase cells; {blacksquare}, untreated stationary-phase cells. For B, D, and F, symbols are as follows: •, 16 µg/ml oritavancin versus stationary-phase cells; {circ}, 16 µg/ml oritavancin versus exponential-phase cells; {blacksquare}, 16 µg/ml vancomycin versus stationary-phase cells; {square}, 16 µg/ml vancomycin versus exponential-phase cells; {blacktriangleup}, 4 µg/ml daptomycin versus stationary-phase cells; {triangleup}, 4 µg/ml daptomycin versus exponential-phase cells.

In the membrane permeability assay, a quantitative difference in initial fluorescence was observed in stationary-phase cells compared to exponential-phase cells (Fig. 3C). This finding may reflect that stationary-phase cells either have a lower uptake of Syto 9 or are initially more permeable to propidium iodide. Oritavancin increased the membrane permeability of stationary-phase cells in a concentration-dependent manner (data not shown), as evidenced by decreases in Syto 9 fluorescence (Fig. 3C); however, the rate of loss of Syto 9 fluorescence from stationary-phase cells was less than that from exponential-phase cells (Fig. 3C). Vancomycin had no effect on fluorescence within the time frame of the assay (30 min) (Fig. 3D), and daptomycin exposure caused a loss of fluorescence from exponential-phase cells only and not from stationary-phase cells (Fig. 3D).

Time-kill studies using membrane assay buffer over a short duration of exposure also showed that the rate of killing of stationary-phase MSSA by oritavancin was decreased compared to that of the exponential-phase inoculum (Fig. 3E). The rapid bactericidal activity of oritavancin against exponential-phase cells was exemplified by a 3.2-log reduction in CFU within 15 min when tested at 16 µg/ml, its predicted fCmax800. Bacterial killing was also seen with the fCmax of oritavancin (4 µg/ml; 2.9-log reduction) and daptomycin (4 µg/ml; 3.4-log reduction) within 2 h of exposure (Fig. 3E and F). In contrast, stationary-phase cells exhibited approximately 1.5-log and 0.9-log reductions in CFU within the 2-h time period following exposure to oritavancin at the fCmax800 and fCmax, respectively. Daptomycin activity was similarly reduced, as it exerted a 0.7-log reduction in CFU at its fCmax (Fig. 3F). Vancomycin did not effect any change on bacterial counts of either inoculum over the short exposure time of the assay (Fig. 3F).

Oritavancin targets the septum of stationary-phase MRSA ATCC 43300 cells. We recently examined the effect of oritavancin on the ultrastructure of exponential-phase MRSA ATCC 43300 cells by transmission electron microscopy and observed septal deformations and a loss of staining of the nascent septal cross wall, the "midline" (30), in exposed cells (5). These effects were not seen following vancomycin exposure. In this study, qualitative differences were evident upon examination of the stationary-phase culture compared to exponential-phase cells: cell ghosts were present but at a low frequency (data not shown), and septa appeared broader (Fig. 4A) than in exponential-phase cells (Fig. 4B). Furthermore, an electron-dense material was present throughout the extracellular space and attached to the surface of stationary-phase cells (Fig. 4A, C, and D). Septa of oritavancin-treated cells were also broad, but staining of the midline was conspicuously absent (Fig. 4C), corroborating observations of exponential-phase cells (5). The midline was evident in vancomycin-treated cells (Fig. 4D), which overall had an ultrastructural appearance similar to that of the untreated cells.


Figure 4
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  		FIG. 4. Ultrastructural analysis of stationary-phase MRSA ATCC 43300 by transmission electron microscopy of thin sections. (A) Untreated control cells. The arrow indicates the septal midline. (B) Exponential-phase MRSA ATCC 43300 is shown for comparison. The septum is not as broad as in stationary-phase cells (compare to A). The arrow indicates the septal midline. (C) Cells exposed to 1 µg/ml oritavancin for 3 h. Note the absence of a well-defined midline. (D) Cells exposed to 16 µg/ml vancomycin for 3 h. The arrow indicates the septal midline.

Oritavancin is active against in vitro S. aureus biofilms. The commercially available MBEC system (20) was used to establish in vitro biofilms of S. aureus and to determine MBECs of oritavancin and comparator antimicrobial agents. To prevent the loss of oritavancin due to binding to vessel surfaces (3), the Clinical and Laboratory Standards Institute (CLSI; formerly NCCLS) recommends the inclusion of 0.002% polysorbate 80 for oritavancin broth microdilution MIC determinations (12). Initial experiments determined that the inclusion of polysorbate 80 during S. aureus biofilm establishment or antimicrobial challenge caused a significant reduction in CFU/peg densities (data not shown). Therefore, polysorbate 80 was omitted from the planktonic MIC and MBEC determinations in the MBEC system. This omission is predicted to result in an underestimation of oritavancin potency.

The capacity of each strain to form a biofilm on the pegs of the MBEC plate was determined by enumerating the CFU attached to the peg surface (CFU/peg). Biofilm cell densities on the pegs for each strain were (2.9 ± 2.4) x 105 CFU/peg for MSSA ATCC 29213, (2.2 ± 1.7) x 105 CFU/peg for MRSA ATCC 33591, and (2.6 ± 1.1) x 105 CFU/peg for VRSA VRS5 after 24 h of incubation. Planktonic MICs determined for comparator antimicrobial agents in the MBEC assay were within the CLSI quality control ranges (Table 1). Although the testing methodologies were not identical to the conventional broth microdilution MIC method based on CLSI guidelines (10), oritavancin planktonic MICs were also within the quality control range (0.5 to 2 µg/ml) for MSSA ATCC 29213 as determined in the absence of 0.002% polysorbate 80 (13). The growth of S. aureus in a biofilm resulted in dramatic decreases in the antimicrobial activities of vancomycin and linezolid as measured by the concentration of antimicrobial agent needed to sterilize the 24-hour biofilm (MBEC) compared to their respective planktonic MICs (Table 1): MBECs for both agents were >128 µg/ml against all three strains. In contrast, oritavancin MBECs ranged from 2 to 8 µg/ml against the S. aureus strains (Table 1) and were within 1 doubling dilution of their respective planktonic MICs in each experiment.


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  		TABLE 1. Oritavancin exhibits antibiofilm activity in vitro against S. aureus strains of different resistance phenotypesa

The time required for oritavancin to sterilize 24-h biofilms of MSSA ATCC 29213 was determined by measuring the MBECs after shorter exposure times. Oritavancin sterilized the biofilm after a 1-h exposure at an MBEC of 4 µg/ml. As expected, MBECs for the comparator agents were >128 µg/ml at this exposure time. To further test the ability of oritavancin to eradicate biofilm-associated S. aureus in vitro, biofilms of MSSA ATCC 29213 were grown for 72 h to increase the cellular density of the peg biofilms. Indeed, CFU/peg increased to (4.6 ± 1.3) x 106 CFU/peg (approximately a 1.2-log increase compared to the 24-h biofilm cellular density), and oritavancin planktonic MIC and MBEC values were concomitantly affected, ranging from 4 to 32 µg/ml and 8 to 32 µg/ml, respectively. Importantly, within each experiment, oritavancin MBECs were no more than 1 doubling dilution higher than their respective planktonic MICs. Planktonic MICs for vancomycin and linezolid were also affected by the increased cellular peg density and were 2 to 16 µg/ml and 8 to 16 µg/ml, respectively. MBECs for both agents were >128 µg/ml.


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DISCUSSION
 
Oritavancin has been defined as a lipoglycopeptide, distinguishing it structurally from vancomycin (12). Its 4'-chlorobiphenylmethyl group confers the ability to disrupt membrane potential and to increase the permeability of exponential-phase gram-positive pathogens (32). Because membrane barrier function is essential to the viability of all cells, we hypothesized that oritavancin would be active against slowly dividing cells and biofilms. The finding that oritavancin exerted bactericidal activity in vitro against stationary-phase cells, as shown in the present study, suggests that its additional mechanism of action relative to vancomycin, namely, its effect on membrane integrity, is its principal antibacterial mechanism against bacteria in these states. The membrane-perturbing antimicrobial agent daptomycin (1, 42) also exhibited antibacterial activity against stationary-phase cells in our study, confirming findings from a previous report (29). In contrast, the reduced activity of vancomycin against stationary-phase S. aureus, reported here and in other studies (34, 45), suggests the dispensability of cell wall synthesis for viability in this test system and is likely a function of its single mode of action: vancomycin inhibits cell wall synthesis, which occurs in S. aureus only during septum formation in dividing cells (41). Thus, infections in which cell growth is limited by nutrient limitation, quorum sensing, or anaerobiosis may render cells tolerant to the killing action of bactericidal antimicrobial agents, but these cells remain vulnerable to membrane-perturbing agents such as oritavancin and daptomycin.

A reduced rate of killing of stationary-phase cells by oritavancin was observed in either nutrient-depleted CAMHB or membrane assay buffer and was concomitant with a reduced rate of depolarization and permeabilization of cell membranes. Although we cannot exclude the possibility that oritavancin targets are less abundant in stationary-phase cells, these findings are likely an indication that the bactericidal mechanism of action of oritavancin is linked to the energized state of the cell: membrane potential is known to be greater in exponential-phase S. aureus cells (–167 mV) than in stationary-phase cells (–123 mV) (23). Membrane potential and the protonated state of the cell wall are thought to regulate cell wall hydrolase (autolysin) activity (24), and oritavancin-induced changes in the membrane potential of stationary-phase cells may account for the observed loss of the septal midline: the decreased staining intensity of the septal midline in stationary-phase S. aureus cells may have reasonably resulted from a loss of the chemically reactive sites that are exposed (and that bind the heavy metal stain) when cell wall hydrolases cleave septal polymers of the nascent cross walls during division (30, 46). Such a loss of the septal midline in stationary-phase cells corroborates previously reported observations of exponential-phase cells (5). Another possibility is that the inhibition of nascent cross-wall synthesis could also account for the loss of the midline: oritavancin inhibits cell wall synthesis (4) and can directly inhibit the transglycosylase activity of S. aureus penicillin-binding protein 2 (48), an essential transglycosylase-transpeptidase that is localized to the division septum during cell division (36). Thus, while a full understanding of the means by which oritavancin causes cell death requires further investigation, we show in the current work that it likely involves a common mechanism against stationary- and exponential-phase cells.

The potency of oritavancin against S. aureus in vitro biofilms is highlighted by (i) MBEC values within 1 doubling dilution of their planktonic MICs, (ii) sterilization of MSSA biofilms within 1 h, and (iii) sterilization of 72-h biofilms of MSSA that also had increased cellular densities at MBECs within 1 doubling dilution of its planktonic MIC against the test strain. As was seen against stationary-phase cells, vancomycin and linezolid did not sterilize the biofilms, confirming previously reported findings of greatly reduced activities of these agents in in vitro biofilm models (9, 17, 21, 39). Thus, in the in vitro biofilms described here, a tolerant population of cells was present. The finding that the stratification of DNA synthesis activity in an in vitro colony biofilm model in which the vast majority of cells synthesizing DNA (and, therefore, actively dividing) was at the air interface (37) indicates that biofilms are composed of a mixed population of cells with different levels of metabolic activity. Furthermore, another study reported that only a subpopulation of cells in a biofilm were readily killed by the fluoroquinolone ofloxacin (43). Regardless of whether the in vitro (24- or 72-h) biofilms of S. aureus described here were composed of cells of different metabolic activities, oritavancin sterilized the biofilms at a concentration that was 1 doubling dilution higher than that needed to kill planktonic cells. Although rifampin exhibited reduced activity against the in vitro S. aureus biofilms, its MBEC (4 µg/ml) approached pharmacologically achievable concentrations (fCmax {approx} 2 µg/ml) (8). However, development of rifampin resistance has been observed when this agent is used alone in in vitro biofilm models (32) and in vivo (22, 28), which therefore should restrict its use to combination therapy.

The development of antimicrobial agents and therapies that are active against biofilms and tolerant cells would benefit the treatment of infections that harbor cells in these states (14). Unfortunately, these types of infections are often complicated by ischemia (i.e., sequestra of osteomyelitis and chronic wound of diabetic foot ulcers), compromising antimicrobial exposure and activity at the infection site. Thus, new agents and therapies must overcome significant hurdles. The present study shows that oritavancin exerts bactericidal activity against stationary-phase S. aureus cells, likely by its capacity to disrupt the membrane integrity of susceptible bacteria. Furthermore, its potency against the in vitro biofilm was remarkable in light of the observed significant tolerance of the biofilm to comparator antimicrobial agents. To date, oritavancin has shown efficacy in infections that likely harbor tolerant cells, including rat central venous catheter (38), rat granuloma pouch (26), and rabbit endocarditis (25, 40) models. Studies to examine the efficacy of oritavancin in in vivo models of biofilm infections are warranted to confirm its promising activity against cells in a tolerant state in vitro.


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ACKNOWLEDGMENTS
 
We thank the scientific support personnel at Targanta Therapeutics for their contributions to this research. We also thank Norris Allen for helpful discussions and critical review of the manuscript.

With the exception of R.H and T.B., all authors of the manuscript are Targanta Therapeutics employees.


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FOOTNOTES
 
* Corresponding author. Mailing address: Targanta Therapeutics Incorporated, 7170 Frederick Banting, St. Laurent, Quebec H4S 2A1, Canada. Phone: (514) 332-1008, ext. 232. Fax: (514) 332-6033. E-mail: gmoeck{at}targanta.com Back

{triangledown} Published ahead of print on 22 December 2008. Back


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Antimicrobial Agents and Chemotherapy, March 2009, p. 918-925, Vol. 53, No. 3
0066-4804/09/$08.00+0     doi:10.1128/AAC.00766-08
Copyright © 2009, American Society for Microbiology. All Rights Reserved.



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